The FASEB Journal • Research Communication
Amyloid  peptide stimulates platelet activation through RhoA-dependent modulation of actomyosin organization Vijay K. Sonkar, Paresh P. Kulkarni,1 and Debabrata Dash2 Department of Biochemistry, Institute of Medical Sciences, Banaras Hindu University, Varanasi, Uttar Pradesh, India Platelets contribute to 95% of circulating amyloid precursor protein in the body and have widely been employed as a “peripheral” model of neurons in Alzheimer’s disease. We sought to analyze the effects of amyloid  (A) on platelets and to understand the underlying molecular mechanism. The A active fragment containing amino acid sequence 25–35 (A25–35; 10 –20 M) was found to induce strong aggregation of human platelets, granule release, and integrin activation, similar to that elicited by physiological agonists. Platelets exposed to A25–35 retracted fibrin clot and displayed augmented adhesion to collagen under arterial shear, reflective of a switch to prothrombotic phenotype. Exposure of platelets to A peptide (20 M) resulted in a 4.2- and 2.3-fold increase in phosphorylation of myosin light chain (MLC) and MLC phosphatase, respectively, which was reversed by Y27632, an inhibitor of Rho-associated coiled-coil protein kinase (ROCK). A25–35induced platelet aggregation and clot retraction were also significantly attenuated by Y27632. Consistent with these findings, A25–35 elicited a significant rise in the level of RhoA-GTP in platelets. Platelets pretreated with reverse-sequenced A fragment (A35–25) and untreated resting platelets served as controls. We conclude that A induces cellular activation through RhoA-dependent modulation of actomyosin, and hence, RhoA could be a potential therapeutic target in Alzheimer’s disease and cerebral amyloid angiopathy.—Sonkar, V. K., Kulkarni, P. P., Dash, D. Amyloid  peptide stimulates platelet activation through RhoA-dependent modulation of actoABSTRACT
Abbreviations: A, amyloid ; A25–35, amyloid  active fragment containing amino acid sequence 25–35; A, amyloid ; A35–25, amyloid  active fragment containing amino acid sequence 35–25; A40, amyloid  peptide ending at residue 40; A42, amyloid  peptide ending at residue 42; AD, Alzheimer’s disease; APP, amyloid precursor protein; CAA, cerebral amyloid angiopathy; ECL, enhanced chemiluminescence; EGTA, ethylene glycol tetraacetic acid; H&E, hematoxylin and eosin; HRP, horseradish peroxidase; Hylite-555A42, Hylite-555 Fluor-labeled amyloid  peptide ending at residue 42; MLC, myosin light chain; MYPT1, MLC phosphatase; oxphos, oxidative phosphorylation; PVDF, polyvinylidene fluoride; PKC, protein kinase C; RP, resting platelets; ROCK Rho-associated coiled-coil protein kinase; vWF, von Willebrand factor 0892-6638/14/0028-1819 © FASEB
myosin organization. FASEB J. 28, 1819 –1829 (2014). www.fasebj.org Key Words: platelet adhesion 䡠 clot retraction 䡠 mitochondrial respiration 䡠 myosin light chain 䡠 thromboembolism Alzheimer’s disease (AD) is the most common form of dementia affecting the elderly that is characterized by gradually but relentlessly progressive cognitive impairment. Amyloid plaques and neurofibrillary tangles are the two hallmark pathological features of the disease (1). They are composed of aggregated amyloid  (A) peptide (2) and hyperphosphorylated Tau proteins (3), respectively. Accumulation of A is considered central to development and progression of cognitive decline and neuronal death. However, the precise signaling pathways mediating the effects of A on neurons remain to be elucidated (4). A is also cytotoxic to cerebral endothelial cells (5) and vascular smooth muscle cells (6). A peptide ending at residue 40 (A40), in particular, can accumulate in the cerebral blood vessels, leading to cerebral amyloid angiopathy (CAA; ref. 7). Further, patients with AD and CAA also present with vascular and hemostatic abnormalities (8), the etiology of which remains largely obscure. Platelets are small disk-shaped circulating blood cells that play a central role in hemostasis and thrombosis. Amyloid processing in platelets closely reflects that occurring in neurons (9). Platelets contain ⬎90% of the circulating amyloid precursor protein (APP; ref. 10), which, on proteolytic cleavage by - and ␥-secretases, yields A40 (11). Hence, platelets have been evaluated both as an easily accessible model to study A generation, as well as a source for peripheral markers of AD (12). Platelets generate A40 on stimulation with physiological agonists like thrombin or collagen in a 1 Current address: Department of Biochemistry, Bharati Vidyapeeth University Medical College, Pune 411043, Maharashtra, India. 2 Correspondence: Department of Biochemistry, Institute of Medical Sciences, Banaras Hindu University, Varanasi 221005, Uttar Pradesh, India. E-mail:
[email protected] doi: 10.1096/fj.13-243691 This article includes supplemental data. Please visit http:// www.fasebj.org to obtain this information.
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manner regulated by protein kinase C (PKC; ref. 13). Platelets are also known to contain preformed A40 in their cytosol, which is released extracellularly on activation (14, 15). However, the latter process is PKC independent (13). Hence, platelets could potentially contribute to circulating A40 and possibly also to deposits in the cerebral vessel walls, while there have been suggestions to link A with platelet activity (14, 16). In the present study, we employed the active fragment of A containing amino acid residues 25–35 (A25–35), which demonstrates more expedient solubilization (17) while exhibiting effects equivalent to full-length A (14, 16 –18). Platelets have been used as peripheral model to understand the influence of A on neuronal cells and pathogenesis of AD. Here, we aim to analyze the effects of A25–35 on platelets and unveil the underlying molecular mechanisms to throw light on vascular abnormalities associated with AD and CAA.
MATERIALS AND METHODS
Platelet aggregation and dense granule secretion Pretreated washed human platelets were stirred (1200 rpm) at 37°C in an optical lumi-aggregometer (Chrono-log model 700-2; Wheecon Instruments, New Delhi, India) for 1 min, after which thrombin (1 U/ml) or varying concentration of A25–35 was added, and transmittance was recorded. Aggregation was measured as percentage change in light transmission, where 100% refers to transmittance through blank solution (19). ATP secretion was assessed simultaneously by recording luminescence in the same instrument using Chrono-lume luciferin-luciferase reagent following manufacturer’s instructions (21). A35–25 was used as a negative control.
Measurement of PAC-1 binding
A25–35 (GSNKGAIIGLM), A35–25 (MLGIIAGKNSG), apyrase, ethylene glycol tetraacetic acid (EGTA), ethylene diamine tetraacetic acid (EDTA), sodium orthovanadate, acetylsalicylic acid, antimycin A (respiratory chain complex III inhibitor), oligomycin (ATP synthase inhibitor), thrombin, fibrinogen, avertin (2,2,2-tribromoethanol), and bovine serum albumin were purchased from Sigma (St. Louis, MO, USA). Y27632 was from Calbiochem (Merck, Darmstadt, Germany), and reagents for electrophoresis were from Merck. Polyvinylidene fluoride (PVDF) membranes and enhanced chemiluminescence (ECL) detection kit were from Millipore (Billerica, MA, USA). Rabbit polyclonal anti-phospho-myosin light chain (MLC) and anti-phospho-MLC phosphatase (MYPT1) were procured from Cell Signaling Technology (Boston, MA, USA). FACSFlow sheath fluid, FITC-labeled PAC-1, and anti-CD62P antibodies were from BD Biosciences (San Jose, CA, USA). Horseradish peroxidase (HRP)-labeled anti-mouse IgG and anti-rabbit IgG secondary antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA) and Bangalore Genei (Bangalore, India), respectively. Collagen, epinephrine, and chrono-lume luciferin-luciferase reagent were procured from Chrono-log (Havertown, PA, USA), and RhoA activation assay Biochem kit was from Cytoskeleton (Denver, CO, USA). Hylite-555 Fluor-labeled A42 (Hylite-555-A42) was purchased from Anaspec (Fremont, CA, USA). All other reagents were of analytical grade. Milli-Q grade deionized water (Millipore) was used for preparation of solutions. Preparation of A25–35 and A35–25 solutions Working stocks (1 mM each) of A25–35 and A35–25 (the reverse-sequenced peptide as negative control) were prepared in ultrapure water (Milli-Q) and stored at ⫺20°C in aliquots. A25–35 and A35–25 were activated before the experiments by incubating solutions at 25°C for 4 h. Hylite-555A42 was reconstituted in buffer containing 50 mM Tris (pH 7.4) and 0.1% NH4OH and stored at ⫺80°C till use at 0.5 mg/ml working concentration. Vol. 28
Platelets were isolated by differential centrifugation of fresh peripheral venous blood drawn from young (20 – 45 yr) healthy male and female volunteers after obtaining informed consent, as already described (19) Mouse platelets were isolated by standard procedure from blood drawn out of Swiss albino mice of either sex aged 8 –12 wk (20).
Flow cytometry
Materials
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Agonist-induced conformational switch in integrin ␣IIb3 results in high-affinity binding of fibrinogen to platelet surface (22). The effect of A25–35 on integrin activation was studied by flow cytometry using PAC-1 antibody, which specifically recognizes the open conformation of ␣IIb3 (22). Washed human platelets (2⫻108 cells) were incubated at 37°C for 5 min without stirring in the presence of either thrombin (1 U/ml) or A25–35 (10 –20 M). FITC-labeled PAC-1 antibody (10 l) was then added to each sample and incubated for 30 min in dark at room temperature. Samples were fixed for 30 min with equal volume of 4% paraformaldehyde. Cells were washed twice in phosphate-buffered saline (PBS; pH 7.4) and finally resuspended in sheath fluid and analyzed on a flow cytometer (FACSCalibur; Becton-Dickinson, Gurgaon, India). An amorphous gate was drawn to encompass platelets separate from noise and multiplatelet particles. All fluorescence data were collected using 4-quadrant logarithmic amplification for 10,000 events in the platelet gate from each sample and analyzed using CellQuest Pro software (Becton-Dickinson; ref. 19). The reverse peptide A35–25 was used in experiments as a negative control. Surface expression of P-selectin Platelet ␣-granule secretion was evaluated by quantifying surface expression of P-selectin. Washed human platelets (2⫻108 cells) were incubated at 37°C for 10 min without stirring in the presence of either thrombin (1 U/ml) or A25–35 (10 –30 M) and fixed for 30 min with equal volume of 4% paraformaldehyde. Cells were washed twice in PBS, following which 5 l FITC-labeled anti-CD62P antibody was added to each sample and incubated for 30 min in the dark at room temperature. Cells were washed, resuspended in sheath fluid, and analyzed by flow cytometry as described above (19). A-platelet interaction Washed platelets were incubated with 5 M Hylite-555-A42 for 15 min, washed with PBS, and resuspended in sheath
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SONKAR ET AL.
fluid. Fluorescence data were collected in FL2 channel in a flow cytometer using 4-quadrant logarithmic amplification of 10,000 events as described above. Untreated platelets served as control. For confocal microscopy, Hylite-555-A42-treated platelets were fixed overnight with 2% paraformaldehyde. Cells were examined under a Zeiss LSM 700 laser scanning confocal microscope (Carl Zeiss, Bangalore, India) with ⫻63 oil objective and 1 AU pinhole size. Images were acquired and analyzed using ZEN imaging software (Carl Zeiss). Platelet adhesion and spreading on immobilized fibrinogen Platelets (1⫻106 cells) were pretreated for 10 min with either thrombin (1 U/ml) or A25–35 (20 M) and charged onto slides coated with fibrinogen (100 g/ml). After incubation for either 7.5 or 15 min at room temperature, cells were fixed with 4% formaldehyde for 30 min. Slides were washed 3 times with PBS, and adhered cells were observed under a fluorescence microscope with phase-contrast attachment (Leica model DM LB2; Labindia Instruments, Maharashtra, India) at ⫻100 in oil. Images were captured by Leica DFC 320 CCD camera and analyzed using Leica IM50 software (19). In vitro model of thrombosis by dynamic flow chamber assay Washed human platelets were rendered fluorescent by incubation with calcein-AM (2 g/ml) at 37°C for 15 min and resuspended in buffer B containing von Willebrand factor (vWF; 10 g/ml) and calcium chloride (2 mM). Type I collagen-coated glass coverslips were assembled in a parallel plate flow chamber (GlycoTech, Raleigh, NC, USA). The chamber was mounted on the stage of an inverted epifluorescence videomicroscope (Nikon model Eclipse Ti-E; Towa Optics, New Delhi, India) equipped with a monochrome CCD cooled camera. Platelets with or without stimulation by A25–35 (20 M) were perfused with help of a syringe pump (Pump 22 infusion/withdrawal with standard syringe holder; Harvard Apparatus, Holliston, MA, USA) through the chamber at a constant flow rate to yield a wall shear rate of 1500 s⫺1 (15 dyn/cm2). Images were digitalized with DS-Qi1MC digital camera using NIS-Elements AR imaging software (Nikon, Tokyo, Japan). Movie data were converted into sequential photo images, and thrombus growth was evaluated in 2 dimensions by measuring the percentage of area covered by adherent fluorescent platelets at 15 min using NIS-Elements AR (23). High-resolution respirometry in intact human platelets Measurement of mitochondrial respiration was performed in a high-resolution respirometer (Oxygraph-2k; Oroboros Instruments, Innsbruck, Austria; ref. 24) at 37°C under stirring at 750 rpm. A 2-ml suspension of washed human platelets (1.5⫻108/ml in buffer B containing 5.5 mM glucose) was transferred into each oxygraph chamber. Respiration was first allowed to stabilize without any addition at the routine state, i.e., in the physiological coupling state controlled by cellular energy demands on oxidative phosphorylation (oxphos). Then, platelets were treated with varying concentrations of A25–35, and the corresponding oxygen flux was recorded. In other experiments, cells were pretreated either with oligomycin (1 g/ml) or antimycin A (1 g/ml) to study leak respiration or residual oxygen consumption, respectively, followed by addition of A25–35 (20 M). Calibration at air saturation was performed each day before starting experiments by letting Millipore water/buffer B stir with air in the oxygraph chamber until equilibration and a stable signal was AMYLOID  ACTIVATES PLATELETS THROUGH RHOA
obtained. All experiments were performed at an oxygen concentration in the range of 100 –205 M O2. Data were recorded with DatLab 5.1 software (Oroboros Instruments) with sampling rate set to 2 s. Clot retraction studies Washed platelets (2⫻106 cells) were treated either with ADP (10 M) or A25–35 (20 M) in the presence of calcium chloride (1 mM) for 5 min at room temperature, followed by addition of fibrinogen (1.5 mg/ml) and atroxin (0.1 g/ml). Contents were incubated at 37°C. Clot retraction kinetics was observed, and images were captured at different time intervals and analyzed as described previously (25). To study the involvement of Rho-associated coiled-coil-containing protein kinase (ROCK), platelets were preincubated with Y27632 (10 M) for 15 min before treatment with A25–35 (20 M). Immunoblotting Platelet proteins were separated on 10% SDS-PAGE gels and electrophoretically transferred to PVDF membrane by using the TE 77 PWR semidry system (GE Healthcare, Bangalore, India). Membranes were blocked with 5% nonfat dry milk in Tris-buffered saline (10 mM Tris-HCl and 150 mM NaCl, pH 8.0) containing 0.05% Tween-20 (TBST) for 1 h at room temperature. Blots were incubated overnight with respective primary antibody, followed by 3 washings with TBST for 5 min each. Membranes were then placed in HRP-labeled anti-IgG secondary antibody diluted in blocking buffer or TBST for 1 h. Blots were similarly washed, and antibody binding was detected using the ECL detection kit. Images were acquired on a multispectral imaging system (BioSpectrum 800 Imaging system; UVP; Medispec, New Delhi, India) and quantified using VisionWorks LS software (UVP). RhoA-GTP pulldown assay The assay was carried out using a kit (Cytoskeleton) and following manufacturer’s instructions as described previously (26). Briefly, 500 l washed platelets (1.5⫻108/ml), pretreated with either A25–35 (20 M) or thrombin (1 U/ml), were lysed. Supernatants were incubated with 15 l RhotekinRho binding domain (Rhotekin-RBD) beads at 4°C for 1 h. Beads were sedimented, washed, and boiled with Laemmli sample buffer. Samples were subjected to SDS-PAGE, Western blotted, and probed with mouse anti-human RhoA antibody (1:500), followed by goat anti-mouse anti-IgG (1:20.000). Pulmonary thromboembolism model Pulmonary thromboembolism was induced in 8- to 12-wk-old Swiss albino mice of either sex as described previously (19). Thrombosis was initiated by injecting a mixture of collagen (1000 g/kg) plus epinephrine (10 g/kg) or collagen (1000 g/kg) plus epinephrine (10 g/kg) plus A25–35 (2.5 mg/ kg) into the tail vein. The mouse was euthanized after 15 min by administering an overdose of anesthesia. Lungs were perfused with cold saline, dissected out, and immediately fixed in 10% formalin for 24 h. Histological sections were drawn from paraffin-embedded lung tissue, stained with hematoxylin and eosin (H&E), and observed under a light microscope (Nikon Eclipse Ti-E) for the presence of thrombi in pulmonary vessels. Mice injected with normal saline served as negative control. At least 10 low-power fields (⫻10) were observed 1821
from each specimen to obtain a count of thrombus-occluded vessels.
RESULTS
Tail bleeding assay
A25–35 and not A35–25 induces platelet activation
Mice of either sex (age 8 –12 wk) were injected with either saline or A25–35 (2.5 mg/kg) and anesthetized after 15 min. A 1-mm segment of tail tip was cut with a sharp scalpel. Bleeding was monitored by gently dabbing the tip with a Whatman filter paper (Whatman, Maidstone, UK) at 20-s intervals, without touching the wound site, till the cessation of bleeding. The experiment was stopped after 20 min even when bleeding continued to occur (23). Animal studies were carried out as per the recommendations of the Laboratory Animals Division, Central Drug Research Institute (Lucknow, India), and the Laboratory Animal Welfare Committee of the Institute of Medical Sciences, Banaras Hindu University.
Platelet aggregation was induced by A25–35 under stirring at 37°C in a concentration-dependent manner. Amplitude of aggregation elicited by 20 M peptide was 63.81 ⫾ 5.37% (n⫽10), which was comparable to that evoked by a strong physiological agonist like thrombin (1 U/ml; Fig. 1A), while 17.5, 15, 12.5, and 10 M peptide stimulated 69.3 ⫾ 4.04, 59.3 ⫾ 8.14, 46.7 ⫾ 10.4, and 20 ⫾ 8.66% aggregation of platelets, respectively (see Supplemental Fig. S1). Aggregation (32⫾8.1%, n⫽3) was also induced by A25–35 (20 M) in platelets isolated from mice. A25–35-induced aggregation of washed human platelets was inhibited by pretreatment with RGDS (1 mM) or EGTA (5 mM), indicating that the response was integrin dependent (see Supplemental Fig. S2). Release of ATP from platelet dense granules was monitored concurrently with aggregation by luminometry employing luciferinluciferase reagent. A25–35 (20 M) triggered a signifi-
Statistical methods All data are presented as means ⫾ sd of ⱖ3 individual experiments. Two-tailed Student’s t test was used for evaluation of significance and values of P ⬍ 0.05 were considered significant.
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Figure 1. A25–35 induces platelet activation. A) Traces 1 and 2 represent percentage platelet aggregation induced by thrombin (1 U/ml) or A25–35 (20 M), respectively, while traces 1= and 2= correspondingly represent dense granule secretion. B) Corresponding bar diagram represents mean aggregation and secretion induced by either thrombin (1 U/ml) or A25–35 (20 M) (n⫽10). C, D) Histograms represent P-selectin exposure (C) and PAC-1 binding (D) in resting platelets (RP) and thrombin (1 U/ml)- or A25–35-treated platelets, as indicated. Figures are representative of ⱖ5 independent experiments. 1822
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cant release from dense granules of aggregating platelets (Fig. 1A, B). Externalization of P-selectin (CD62P) was studied by flow cytometry using fluorescent-labeled anti-CD62P antibody. Treatment of platelets with 10 and 20 M A25–35 brought ⬃1.1- and 1.24-fold increase, respectively, in surface P-selectin expression as compared with the resting cells (Fig. 1C). Platelets exposed to 10 and 20 M A25–35 exhibited 1.3- and 1.8-fold increase, respectively, in PAC-1 binding as opposed to untreated platelets (Fig. 1D). Thrombin was used as positive control in all above experiments. Remarkably, the reverse-sequenced peptide A35–25 induced neither platelet aggregation nor integrin activation, even at concentrations as high as 40 M (see Supplemental Fig. S3). Since aggregation involved high-affinity binding of platelet integrin ␣IIb3 to soluble fibrinogen, we next studied static adhesion and spreading of platelets on fibrinogen immobilized on matrix (27). Pretreatment of cells with A25–35 (20 M) substantially augmented spreading of platelets, characterized by extension of filopodia/lamellopodia, which was particularly distinct at 7.5 min after adhesion compared with control (Fig. 2A). To examine dynamic adhesion of platelets under conditions simulating physiological arterial flow, platelets suspended in the presence of vWF were allowed to run over immobilized collagen at arterial shear (1500 s⫺1). A25–35-pretreated cells adhered remarkably better and more stably than untreated control (Fig. 2B).
A
Possible effects of A25–35 on outside-in signaling were investigated by studying platelet-mediated retraction of fibrin clot. Platelets were activated by addition of either ADP (10 M) or A25–35 (20 M). Clot formation was induced by addition of atroxin (0.1 g/ml) to solution of fibrinogen (1.5 mg/ml) in the presence of activated platelets. Contractile forces originating in the cytoskeleton of A25–35-pretreated platelets progressively compacted bulk of the fibrin clot through integrin ␣IIb3fibrin interaction. This led to reduction in area occupied by clot by 80.3% in 30 min, which was comparable to that induced by ADP (83.2%; see Fig. 4F, G). A25–35 spikes mitochondrial respiration in platelets Platelet activation responses elicited by A25–35, namely platelet aggregation, granule release, and retraction of fibrin clot, are highly energy demanding. As oxphos is the major source of ATP in resting platelets (RPs; ref. 28), we examined influence of A25–35 on mitochondrial respiration by high-resolution respirometry. Platelets suspended in buffer containing 5.5 mM glucose exhibited routine (endogenous) respiration at 16.43 ⫾ 1.7 pmol O2/s/108 cells, which represented resting cellular energy demands through oxphos and was consistent with an earlier report (29). Strikingly, A25–35 (20 M) brought ⬃65.7 ⫾ 6.02% rise in respiration in 1 min (Fig. 3A, top panel). Stimulation of respiration was transient, and its magnitude was depen-
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Figure 2. A) Phase-contrast microscopy images of control and A25–35 (20 M)-treated platelets undergoing shape change and spreading on adhesion to immobilized fibrinogen, observed at 7.5 min (top panels) and 15 min (bottom panels) after platelet adhesion to matrix. B) Fluorescent microscopy images of control (top panel) and A25–35 (20 M)-treated (bottom panel) calcein-labeled platelets adhered to immobilized collagen under flow at 1500 s⫺1, acquired after 6 min. Figures are representative of 3 independent experiments. AMYLOID  ACTIVATES PLATELETS THROUGH RHOA
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concentration of the peptide (Fig. 3B). To the component of coupled respiration indeof ADP phosphorylation (leak state), respirarecorded in the presence of oligomycin. Ex-
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pectedly, leak respiration in platelets was low (3.61⫾ 0.18 pmol O2/s/108 platelets). However, subsequent addition of A25–35 (20 M) led to 2.8-fold augmentation in leak respiration (Fig. 3A, middle panel). To
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Figure 4. A) Immunoblots representing MYPT1 and MLC phosphorylation in washed human platelets stimulated with either thrombin (1 U/ml) or A25–35 (20 M) in presence or absence of Y27632 (10 M). B, C) Corresponding bar diagrams represent the densitometry analyses of immunoblots for p-MYPT1 (B) and p-MLC (C), normalized with respect to -actin expression (n⫽3). D) Traces 1 and 3 represent percentage platelet aggregation induced by A25–35 (20 M) or thrombin (1 U/ml), respectively; traces 2 and 4 represent percentage platelet aggregation induced by A25–35 (20 M) or thrombin (1 U/ml), respectively, in the presence of Y27632 (10 M). Traces 1=, 2=, 3=, and 4= represent corresponding dense granule secretion. E) Corresponding bar diagram represents mean aggregation and dense granule secretion (n⫽3). F) Photographs show retraction of fibrin clot by washed human platelets pretreated with ADP (10 M), A25–35 (20 M) or A25–35 (20 M) plus Y27632 (10 M), as indicated. G) Corresponding graphic representation of mean reduction in clot size (n⫽3) at different time points in the presence of reagents as indicated. RP, resting (untreated) platelets. Data are presented as means ⫾ sd. *P ⬍ 0.05 vs. control platelets; #P ⬍ 0.05 vs. A25–35-treated platelets.
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examine input of extramitochondrial oxygen consumption, platelets were treated with antimycin A, which dropped oxygen consumption in cells to 1.87 ⫾ 0.1 pmol O2/s/108 platelets. Subsequent addition of A25–35 had no significant effect on residual oxygen consumption (Fig. 3A, bottom panel). A25–35 modulates platelet cytoskeletal reorganization through ROCK Platelet activation by physiological agonists is associated with extensive reorganization of cytoskeleton, brought about by actomyosin contraction. Exposure of platelets to A25–35 (20 M) resulted in 4.2- and 2.3-fold increase in phosphorylation of MLC and its phosphatase (MYPT1), respectively, which was reversed by Y27632 (10 M), an inhibitor of ROCK (Fig. 4A–C), thus implicating RhoROCK-MLC/MYPT1 axis in A25–35-mediated signaling. In support of this, A25–35-induced platelet physiological responses such as aggregation, secretion, and retraction of fibrin clot were also significantly attenuated in the presence of Y27632 (Fig. 4D–G). Thrombin was employed as positive control in these experiments, which significantly elevated phosphorylation of MLC and MYPT1 in a Y27632-sensitive manner. A25–35 promotes RhoA activity As earlier results were consistent with activation of ROCK-MLC/MYPT1 axis in A25–35-stimulated platelets, we evaluated the activity of RhoA in A-treated cells by pulldown assay. A25–35 (20 M) induced a significant rise in RhoA-GTP expression in platelets as compared with untreated cells (Fig. 5). There was a progressive increment in RhoA-GTP level by 21.95, 61.19, and 73.35% with respect to untreated cells measured at 2, 5, and 10 min after A addition, respectively. Thrombin (1 U/ml) was employed as positive control in these experiments. Exogenous A associates with platelets and accumulates intracellularly To understand the nature of interaction, platelets were preincubated with Hylite-555-A42, followed by flow
cytometry. Platelets were strongly fluorescent when examined in FL2 channel, which was consistent with close interaction between the peptide and cells (Fig. 6A). This finding was further corraborated when fixed platelets pretreated with fluorescently labeled A42 were examined under a confocal microscope. Overlay images, as well as z scanning (0.41-m steps), clearly demonstrated cytosolic localization of red fluorescence, suggestive of intracellular accumulation of the peptide (Fig. 6B, C). The fluorescence density of Hylite555-A42 was maximum within the cytosol of platelets in optical sections between 2.86 and 4.90 m. A25–35 potentiates thrombus formation in vivo We studied the effect of A25–35 in a mouse model of pulmonary thromboembolism to establish its influence on thrombus formation in vivo. H&E-stained lung sections obtained from mice euthanized 15 min after intravenous administration of collagen plus epinephrine, collagen plus epinephrine plus A25–35, or normal saline were examined for thrombi in pulmonary vessels. A25–35treated mice had considerably greater numbers of pulmonary vessels associated with thrombi than those administered collagen plus epinephrine alone (8.6/low-power field vs. 5/low power field; n⫽3), while saline-injected mice did not exhibit thromboembolism (Fig. 7). We also examined the effect of A25–35 on hemostasis by tail bleeding assay. A25–35 (2.5 mg/kg)-pretreated mice had significantly shorter bleeding times (9.5⫾2.9 vs. 6.5⫾0.7, n⫽3, P⫽0.04) than saline-treated controls.
DISCUSSION The present study demonstrates that the A-derived peptide A25–35 elicits significant activation of human platelets independent of known physiological agonists. This response was associated with high-affinity binding of integrin ␣IIb3 to fibrinogen and concomitant release of granule contents. Predictably, A25–35 substantially augmented spreading of platelets on immobilized fibrinogen, as well as retraction of fibrin clot, processes that are critically dependent on integrin-fibrinogen
Aβ (20μM)
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*
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*
1.5 1 0.5 0
RP
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5 min
10 min
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Figure 5. A) Level of RhoA-GTP in washed human platelets stimulated with A25–35 (20 M) or thrombin (1 U/ml) for different time points. B) Corresponding bar diagram represents densitometry analyses of ⱖ3 independent immunoblots normalized with respect to total RhoA expression. Data are presented as means ⫾ sd. *P ⬍ 0.05 vs. control platelets. 1826
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(A)
(B) Control
Fluorescence
Aβ
(0.00%)
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(83.92%)
(4.43%)
(99.85%)
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Figure 6. A) FL1 and FL2 fluorescence of 10,000 platelet events acquired by flow cytometry, representing interaction between fluorescently labeled A42 and platelets. Number within parentheses in each quadrant represents percentage of total gated events in the respective quadrant. B) Overlay of confocal fluorescence and DIC images of platelets incubated with Hylite-555 (red fluorescence)-labeled A42. C) Confocal fluorescence images obtained from z scanning (with 0.41-m steps) shows intracellular localization of fluorescently labeled peptide.
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A
B
C
Figure 7. Light microscopy images of H&E-stained lung sections from mice administered normal saline (A), mixture of collagen (1000 g/kg) plus epinephrine (10 g/kg) (B), and mixture of collagen (1000 g/kg) plus epinephrine (10 g/kg) plus A25–35 (2.5 mg/kg) (C). Arrows indicate thrombi within the lumen of pulmonary vessels.
interaction and subsequent outside-in signaling. The peptide remarkably enhanced stable adhesion of platelets to immobilized collagen under flow at arterial shear (1500 s⫺1), suggesting that A peptide could favor stable thrombus formation. Since events triggered by platelet stimulation are energy intensive, and oxphos contributes to nearly 85% of energy requirements in RPs (28), we examined the influence of A25–35 on mitochondrial respiration. Highresolution respirometry demonstrated that A25–35 evoked a sharp albeit transient rise in mitochondrial respiration in intact platelets, primarily attributable to ADP phosphorylation and partly to leak. This spurt in oxphos could help sustain the platelet responses succeeding A25–35-induced platelet activation. Augmented respiration may be explained from enhanced nutrient catabolism generating endogenous substrates for mitochondrial respiration in A25–35-stimulated platelets. Platelet activation induces actomyosin contractility leading to extensive cytoskeletal reorganization, which underpins activation-specific responses, such as aggregation of cells, shape change, granule release, and clot retraction. Phosphorylation and inhibition of MYPT1 by ROCK and subsequent phosphorylation of MLC facilitates actomyosin complex formation and contraction in activated platelets (30). A peptide (20 M) increased phosphorylation of MLC and MYPT1 in platelets that was reversed by Y27632 (ROCK inhibitor). Furthermore, A-induced platelet aggregation, secretion, and clot retraction were also at least partially abrogated by Y27632, thus implicating the RhoAROCK-MYPT1-MLC axis in A25–35-mediated signaling. Consistent with these results, RhoA activity was potentiated in A-stimulated platelets. Platelet agonists acting through Gq and G13 coupling are known to induce RhoA activation that mediates different platelet responses through actomyosin contraction (30). Thus, A25–35 could be effecting RhoA activation by interaction with G13- or Gq-coupled receptors on the platelet surface. However, the possibility of A25–35 directly influencing RhoA activity cannot be ruled out (31). We examined the nature of interaction between A and platelets by employing fluorescently labeled peptide. 1828
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A was found to strongly associate with platelets, as well as localize in their cytoplasm. The consequence of A-stimulated platelet activation on thrombus formation in vivo was evaluated in a mouse model of pulmonary thromboembolism. A25–35 was found to potentiate collagen-epinephrine-initiated thrombosis in Swiss albino mice. However, A is also known to bind fibrinogen, induce its polymerization, and protect the fibrin clot from plasmin-mediated lysis (32). Therefore, it remains to be determined whether the observed prothrombotic effect of A25–35 in vivo is a consequence of direct stimulation of platelets or its interaction with fibrinogen. In summary, this study highlights the ability of A to induce platelet activation independently in the absence of physiological agonists, which could have consequences in understanding hemostatic abnormalities observed in AD as well as in pathogenesis of CAA. Although the plasma level of A is much lower than that required for stimulating platelets, high local concentration may be achieved in plasma (32) by release from stimulated platelets at the site of thrombus formation (13, 33) and also possibly at the site of CAA (34) or atherosclerotic plaques (35), thus initiating a vicious cycle of platelet stimulation and A release. We have for the first time demonstrated that effects of A are mediated through activation of small-GTPase RhoA, leading to cytoskeletal reorganization and activation-specific cellular responses, thus positioning RhoA as an essential signaling component downstream of A in cells. Pharmacological inhibition of Rho kinase abrogates the effects of A on cell functions that could therefore be potential therapeutic targets in AD and CAA. This research was supported by grants received by D.D. from the Department of Science and Technology (DST) and the Department of Biotechnology (DBT), Government of India; the Indian Council of Medical Research (ICMR); and the Council of Scientific and Industrial Research (CSIR). D.D. thankfully acknowledges a Tata Innovation Fellowship grant received from the DBT. V.K.S. is the recipient of a research fellowship from ICMR. P.P.K. was supported by residency program of the Institute of Medical Sciences (IMS), Banaras Hindu University (BHU). The authors declare no
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SONKAR ET AL.
conflicts of interest. The authors sincerely thank Dr. Mohan Kumar (Department of Pathology, IMS, BHU) for helping with the lung histology study.
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