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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

The generation of superoxide anion is essential for the stimulation of platelets and, if dysregulated, critical for thrombotic diseases. Here, we present three protocols for the selective detection of superoxide anions and the study of redox-dependent platelet regulation.

Abstract

Reactive oxygen species (ROS) are highly unstable oxygen-containing molecules. Their chemical instability makes them extremely reactive and gives them the ability to react with important biological molecules such as proteins, nucleic acids, and lipids. Superoxide anions are important ROS generated by the reduction of molecular oxygen reduction (i.e., acquisition of one electron). Despite their initial implication exclusively in aging, degenerative, and pathogenic processes, their participation in important physiological responses has recently become apparent. In the vascular system, superoxide anions have been shown to modulate the differentiation and function of vascular smooth muscle cells, the proliferation and migration of vascular endothelial cells in angiogenesis, the immune response, and the activation of platelets in hemostasis. The role of superoxide anions is particularly important in the dysregulation of platelets and the cardiovascular complications associated with a plethora of conditions, including cancer, infection, inflammation, diabetes, and obesity. It has, therefore, become extremely relevant in cardiovascular research to be able to effectively measure the generation of superoxide anions by human platelets, understand the redox-dependent mechanisms regulating the balance between hemostasis and thrombosis and, eventually, identify novel pharmacological tools for the modulation of platelet responses leading to thrombosis and cardiovascular complications. This study presents three experimental protocols successfully adopted for the detection of superoxide anions in platelets and the study of the redox-dependent mechanisms regulating hemostasis and thrombosis: 1) dihydroethidium (DHE)-based superoxide anion detection by flow cytometry; 2) DHE-based superoxide anion visualization and analysis by single platelet imaging; and 3) spin probe-based quantification of superoxide anion output in platelets by electron paramagnetic resonance (EPR).

Introduction

The superoxide anion (O2•-) is the most functionally relevant ROS generated in platelets1. O2•- is the product of the reduction of molecular oxygen and the precursor of many different ROS 2. The dismutation of O2•- leads to the generation of hydrogen peroxide (H2O2) via spontaneous reactions in aqueous solution or reactions catalyzed by superoxide dismutases (SODs3). Although different enzymatic sources have been suggested (e.g., xanthine oxidase4, lipoxygenase5, cyclooxygenase6, and nitric oxide synthase7), mitochondrial respiration8,9 and nicotinamide adenine dinucleotide phosphate-oxidases (NOXs)10 are the most prominent sources of superoxide anion in eukaryotic cells. This also seems to be the case in platelets, where electron leakage from mitochondrial respiration11,12 and the enzymatic activity of NOXs13,14 have been described as the main contributors to the superoxide anion output.

Although several studies have focused on the regulation of platelets by O2•-, there is no consensus regarding the molecular mechanisms responsible. The modulation of surface receptor activity via direct oxidation and disulfide bond formation has been proposed for different platelet receptors. The positive regulation of integrin αIIbβ3 by ROS via direct oxidation of cysteine residues has been suggested15,16,17. Similarly, since platelet responses to collagen depend on disulfide-dependent dimerization and consequent dimerization of the glycoprotein VI (GPVI)18, receptor activity potentiation by ROS-dependent oxidation has been proposed19, although not fully proven experimentally. Finally, ROS-induced oxidation of the sulfhydryl groups of glycoprotein Ib (GPIb) was shown to promote platelet adhesion and platelet-leukocyte interaction during inflammation20. Conversely, as a possible consequence of decreased sulfhydryl group oxidation and receptor activation, the shedding of the ectodomain of both GPVI and GPIb is diminished by reducing conditions21.

Modes of action independently of a direct oxidation of platelet surface receptors have also been proposed. ROS, including O2•-, have been shown to positively modulate the collagen receptor GPVI by attenuating the activity of the Src homology region 2-containing protein tyrosine phosphatase 2 (SHP-2), which negatively regulates the signaling cascade of this receptor22. Moreover, O2•- can generate ONOO- (peroxynitrite) by rapid reaction with nitric oxide (NO), which normally inhibits platelets through the NO-sensitive guanylyl cyclase (NO-GC) and the generation of the negative platelet regulator cyclic GMP (cGMP)23,24. The resulting decrease in NO levels can lead to platelet potentiation. Alternatively, the generation of O2•- by NOX2 has been suggested to contribute to lipid peroxidation and isoprostane formation, which is essential for platelet activation and adhesion25. Finally, mitogen-activated protein kinase (MAPK) extracellular signal-regulated kinase 5 (ERK5), a protein kinase proposed as a redox stress sensor in platelets26, is activated by O2•- and induces a procoagulant phenotype in platelets (as estimated by flow cytometry-based measurement of phosphatidylserine externalization)27.

The dysregulation of O2•- and other ROS generation in platelets has been associated with the exaggerated hemostatic response leading to thrombotic cardiovascular complications associated with atherosclerosis, diabetes mellitus, hypertension, obesity, and cancer28,29. In these pathological settings, the ROS output by platelets is increased, which leads to a potentiation of their adhesive and aggregatory responses. In addition to the effect on platelet responses, the free radical output of platelets may have consequences on other blood cells and vascular structures, which is a poorly understood and underinvestigated area of cardiovascular health30. Despite our limited understanding of the molecular mechanisms linking oxidative stress to thrombotic conditions, the clinical relevance of antioxidants for the protection against cardiovascular disease has received considerable attention. Plasma antioxidant levels have been shown to inversely correlate with the risk of developing cardiovascular conditions, and dietary antioxidant consumption has been shown to protect against coronary artery disease31,32. Consequently, the use of dietary antioxidants has been advocated as a promising approach for cardiovascular disease prevention33,34,35. Amongst the effects of ROS generation in platelets, the increase in apoptosis can have important pathophysiological effects36,37. Overall, reliable protocols to detect and quantify the O2•- output by platelets are increasingly relevant in cardiovascular research.

Currently, available techniques for the detection of ROS have important limitations of specificity (i.e., the chemical nature of the oxidant molecules detected is unknown) and reliability (i.e., the unwanted interaction with biological molecules and experimental reagents leads to biased non-physiological results)38,39. The most commonly used approach for the detection of ROS in platelets is based on the use of dichlorodihydrofluorescein diacetate (DCFDA), which is converted to dichlorodihydrofluorescein (DCFH) by intracellular esterases and consequently to the highly fluorescent dichlorofluorescein (DCF) by cellular oxidants, including hydroxyl radicals and peroxidase-H2O2 intermediates40,41. Despite its wide use, serious questions have been raised regarding the reliability of this approach for the measurement of intracellular ROS38. The oxidation of DCFH to DCF can be, in fact, induced by transition metal ions (e.g., Fe2+) or heme-containing enzymes (e.g., cytochromes) instead of ROS42. Moreover, DCFDA is converted by cell peroxidases to its semiquinone free radical form (DCF•-), which is in turn oxidized to DCF by reaction with molecular oxygen (O2) with the release of O2•-, which leads to the artificial amplification of oxidative responses41,43,44. Therefore, the detection of intracellular ROS by DCFDA is useful for obtaining initial insights but requires cautious consideration and extensive experimental controls38,39.

This study presents three alternative techniques for the detection and measurement of the key regulator of platelet function O2•-1. The first technique is the detection using DHE and flow cytometry, which offers advantages of reliability and specificity over DCFDA. The second technique proposed here also utilizes DHE, but the detection method is live-platelet fluorescence imaging, which allows the study of the generation of O2•- upon platelet signaling with fast kinetics and single-cell resolution. Finally, a protocol based on the use of the hydroxylamine spin probe 1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine (CMH) in EPR resonance experiments offers the possibility of quantifying the rate of O2•- generation by platelets and comparing it in different conditions.

Protocol

The collection of peripheral blood from consenting volunteers is approved by the local Ethics Committee and the National Health Service Health Research Authority (REC reference: 21/SC/0215; IRAS ID: 283854).

1. Method 1: Superoxide anion detection using DHE by flow cytometry

  1. Pre-warm (37 °C) sodium citrate solution (4% w/v) and use it as an anti-coagulant by adding it directly to the blood at the time of venepuncture at a ratio of 1:7 (0.5% w/v final).
  2. Draw peripheral human blood from healthy volunteers by median cubital vein venepuncture.
  3. Obtain platelet-rich plasma (PRP) from whole blood by an initial centrifugation step at 200 x g for 20 min.
  4. Centrifuge PRP at 500 x g for 10 min with the reversible platelet inhibitors indomethacin (10 µM) and PGE1 (40 ng/mL).
  5. Resuspend the resulting platelet pellet in modified Tyrode's buffer (145 mM NaCl, 2.9 mM KCl, 10 mM 4-(2-hydroxyethyl)-1-piperazineethane sulfonic acid (HEPES), 1 mM MgCl2, 5 mM glucose, pH 7.3) (37 °C) at a density of 2 x 108 platelets/mL.
  6. After isolation, rest platelets for 30 min at 37 °C.
  7. Prepare DHE in dimethyl sulfoxide (DMSO) at a stock concentration of 5 mM.
  8. Add DHE to the platelet suspension at 5 µM final concentration (dilute the stock solution 1 to 1,000, with final DMSO concentration 0.1% v/v) and incubate for 15 min at 37 °C.
  9. Treat platelets with pharmacological agents and/or ROS scavengers for 15 min before stimulation with the desired physiological stimuli and concentrations (e.g., 0.1 unit/mL thrombin or 3 µg/mL collagen).
  10. After stimulation, dilute platelet suspensions 1 to 10 in ice-cold modified HEPES buffer.
  11. For the flow cytometry, set side scattering (SSC) and forward scattering (FSC) gain at 40 mV and 220 mV, respectively, and use logarithmic scale scatter plots to visualize the particle population in the suspension (example in Figure 1A).
  12. (OPTIONAL) Immunostain one aliquot of platelet suspension using an anti-CD41 antibody to identify the population of events corresponding to platelets (example in Figure 1B).
  13. Analyse samples by flow cytometry using excitation at 405 nm (violet laser) and emission at 580 nm wavelength, which selectively detects the superoxide anion-specific product 2-hydroxy-ethidium (2OH-Et+) (Figure 2).
    NOTE: Fixation with paraformaldehyde and long-term sample storage are not advisable.

2. Method 2: Superoxide anion detection using DHE by single-platelet fluorescence imaging

  1. Coat an 8-well µ-slide on the day of the experiment with 0.1 mg/mL fibrillar collagen I from equine tendons (Horm collagen), 0.2 mg/mL fibrinogen or 1 mg/mL poly-L-lysine (PLL) in modified Tyrode's buffer for 2 h at 37 °C.
  2. Wash twice with modified Tyrode's buffer (10 min each).
  3. Quench non-specific adhesion by incubation in modified Tyrode's buffer plus 5 mg/mL bovine serum albumin (BSA) for a minimum of 30 min (or until the experiment).
  4. Pre-warm (37 °C) sodium citrate solution (4% w/v) and use it as an anti-coagulant by adding it directly to the blood at the time of venepuncture at a ratio of 1:7 (0.5% w/v final).
  5. Draw peripheral human blood from healthy volunteers by median cubital vein venepuncture.
  6. Obtain platelet-rich plasma (PRP) from whole blood by an initial centrifugation step at 200 x g for 20 min.
  7. Centrifuge PRP at 500 x g for 10 min with the reversible platelet inhibitors indomethacin (10 µM) and PGE1 (40 ng/mL).
  8. Resuspend the resulting platelet pellet in modified Tyrode's buffer (145 mM NaCl, 2.9 mM KCl, 10 mM HEPES, 1 mM MgCl2, 5 mM glucose, pH 7.3) (37 °C) at a density of 4 x 107 platelets/mL.
  9. After isolation, rest platelets for 30 min at 37 °C.
  10. Prepare DHE in dimethyl sulfoxide (DMSO) at a stock concentration of 10 mM.
  11. Add DHE at 10 µM final concentration to the platelet suspension (dilute the stock solution 1 to 1,000, with final DMSO concentration 0.1% v/v) and incubate for 1 min (37 °C).
  12. After the 1 min of incubation with DHE, remove the blocking solution from the chosen wells, position the µ-slide on the microscope stage ready for imaging, and gently dispense platelets.
  13. Monitor DHE conversion to 2OH-Et+ by inverted confocal imaging for 10 min (405/580 nm ex/em), with images collected every 10 s with a 40x oil immersion lens.
  14. (OPTIONAL) For wells coated with PLL, monitor the superoxide anion generation in response to a soluble agonist (e.g., 0.1 unit/mL thrombin or 3 µg/mL collagen) by adding the agonist 10 min after platelet dispensing and fluorescence image collection for a further 10 min, as described in step 2.13.
  15. (OPTIONAL) If required, add superoxide anion scavengers or selective inhibitors (e.g., NOX inhibitor) by dispensing gently to the side of the well during the time course and collect fluorescence images as described in 2.13 for a further 10 min.
  16. Quantify single-cell fluorescence analysis by selecting the Region of Interest (ROI) containing representative single platelets and analyzing fluorescence intensity in different frames with the Measure tool of imaging suite ImageJ 1.53t.

3. Method 3: Detection of superoxide anion by platelets using electron paramagnetic resonance (EPR)

  1. Pre-warm (37 °C) sodium citrate solution (4% w/v) and use it as an anti-coagulant by adding it directly to the blood at the time of venepuncture at a ratio of 1:7 (0.5% w/v final).
  2. Draw peripheral human blood from healthy volunteers by median cubital vein venepuncture.
  3. Obtain platelet-rich plasma (PRP) from whole blood by an initial centrifugation step at 200 x g for 20 min.
  4. Centrifuge PRP at 500 x g for 10 min with the reversible platelet inhibitors indomethacin (10 µM) and PGE1 (40 ng/mL).
  5. Resuspend the resulting platelet pellet in modified Tyrode's buffer (145 mM NaCl, 2.9 mM KCl, 10 mM HEPES, 1 mM MgCl2, 5 mM glucose, pH 7.3) (37 °C) at a density of 2 x 108 platelets/mL.
  6. After isolation, rest platelets for 30 min at 37 °C.
  7. Add 5 µM diethyldithiocarbamate (DETC) and 25 µM deferoxamine to the platelet suspension.
  8. Without incubation, add 200 µM 1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine (CMH).
  9. Load samples onto the aggregometer within aggregation cuvettes (including Teflon-coated stirring magnets).
  10. After 1 min, add the stimuli at the desired concentration (e.g., 1 unit/mL thrombin or 3 µg/mL collagen). Incubate for 10 min.
  11. (OPTIONAL) Obtain superoxide anion generation at different time points by proceeding to step 3.12 at the desired time.
  12. Transfer the platelet suspension from the aggregation cuvette to a microcentrifuge tube and quickly spin down at 6,000 x g for 10 s.
  13. Load 50 µL of the supernatant in capillary micropipettes and seal with EPR sealing wax.
  14. Transfer samples to the EPR scanner.
  15. Set the EPR scanner as follows: center field 3,492.5 G, field sweep 60 G, modulation amplitude 2 G, sweep time 10 s, number of scans 10, microwave frequency 9.39 GHz, microwave power 20 mW, conversion time 327.68 ms, time constant 5242.88 ms.
  16. Estimate CMH oxidation to CM● as the area under the curve (AUC) of the EPR peaks recorded using the above parameters.
  17. Build a calibration curve plotting the EPR intensity measured as described in step 3.16 on the y-axis and the concentrations of the commercially available CM● on the x-axis (e.g., 0, 0.3, 1, 3, 10, and 30 µM).
  18. From the CM● concentration in the samples, obtain the amount of superoxide anion generated by the platelets in the samples using the formulas described in the Representative Results section.

Results

For flow cytometry detection of DHE fluorescence, we show representative results for platelets either resting (Figure 3A) or stimulated with 0.1 unit/mL thrombin (Figure 3B). The O2•- output was quantified as platelet mean fluorescence intensity (MFI), as shown for stimulation with 0.1 unit/mL thrombin (Figure 3C) or 3 µg/mL collagen-related peptide (CRP) (Figure 3D

Discussion

In this manuscript, we present three different techniques with the potential to advance the capability to investigate the redox-dependent regulation of platelet function via the selective detection of O2•-. The first two methods are an improvement on existing techniques because of the redox probe utilized (DHE instead of the more common but less reliable DCFDA). These techniques are, therefore, easily accessible, and most laboratories can adopt them effectively without particular eq...

Disclosures

The authors have nothing to disclose.

Acknowledgements

This work was funded by the British Heart Foundation (PG/15/40/31522), Alzheimer Research UK (ARUK-PG2017A-3), and European Research Council (#10102507) grants to G. Pula.

Materials

NameCompanyCatalog NumberComments
1-hydroxy-3-methoxycarbonyl-2,2,5,5-tetramethylpyrrolidine (CMH)Noxygen Science trasfer and Diagnostics GmbHNOX-02.1-50mgReagent for EPR (spin probe)
BD FACSAria IIIBD Biosciences NAFlow cytometer
Bovine Serum AlbuminMerck/SigmaA7030For μ-slide coating
Bruker E-scan M (Noxyscan)Noxygen Science trasfer and Diagnostics GmbH NOX-E.11-BES EPR spectrometer
Catalase–polyethylene glycol (PEG-Cat.)Merck/SigmaC4963Hydrogen peroxide scavenger (specificity control)
ChronoLog Model 490+4Labmedics/ChronologNAAggregometer
CM radicalNoxygen Science trasfer and Diagnostics GmbHNOX-20.1-100mg Reagent for EPR (calibration control)
deferoxamine Noxygen Science trasfer and Diagnostics GmbHNOX-09.1-100mg Reagent for EPR
diethyldithiocarbamate (DETC) Noxygen Science trasfer and Diagnostics GmbHNOX-10.1-1g Reagent for EPR
DihydroethidiumThermo Fisher ScientificsD11347Superoxide anion probe
Dimethyl sulfoxideMerck/Sigma34869For stock solution preparation 
EPR sealing wax platesNoxygen Science trasfer and Diagnostics GmbHNOX-A.3-VPMConsumable for EPR
EPR-grade waterNoxygen Science trasfer and Diagnostics GmbHNOX-07.7.1-0.5L Reagent for EPR
Fibrinogen from human plasmaMerck/SigmaF4883For μ-slide coating
FITC anti-human CD41 AntibodyBioLegend303704Platelet-specific staining for flow cytometry
Glass cuvettes Labmedics/ChronologP/N 312Consumable for incubation in aggregometer
Horm CollagenLabmedics/ChronologP/N 385For platelet stimulation
ImageJ National Institutes of Health (NIH)NAImageJ 1.53t (Wayne Rasband)
IndomethacinMerck/SigmaI7378For platelet isolation
Micropipettes DURAN 50µlNoxygen Science trasfer and Diagnostics GmbHNOX-G.6.1-50µLConsumable for EPR
Poly-L-lysine hydrochlorideMerck/SigmaP2658For μ-slide coating
Prostaglandin E1 (PGE1)Merck/SigmaP5515For platelet isolation
Sodium citrate (4% w/v solution)Merck/SigmaS5770For platelet isolation
Stirring bars (Teflon-coated)Labmedics/ChronologP/N 313Consumable for incubation in aggregometer
Superoxide dismutase–polyethylene glycol (PEG-SOD)Merck/SigmaS9549Superoxide anion scavenger (specificity control)
Thrombin from human plasmaMerck/SigmaT6884For platelet stimulation and μ-slide coating
VAS2870Enzo Life ScienceBML-EI395NOX inhibitor
Zeiss 510 LSM confocal microscopeZeissNAConfocal microscope

References

  1. Vara, D., Cifuentes-Pagano, E., Pagano, P. J., Pula, G. A novel combinatorial technique for simultaneous quantification of oxygen radicals and aggregation reveals unexpected redox patterns in the activation of platelets by different physiopathological stimuli. Haematologica. 104 (9), 1879-1891 (2019).
  2. Fridovich, I. Superoxide radical: An endogenous toxicant. Annu Rev Pharmacol Toxicol. 23, 239-257 (1983).
  3. Keele, B. B., Mccord, J. M., Fridovich, I. Superoxide dismutase from Escherichia coli B. A new manganese-containing enzyme. J Biol Chem. 245 (22), 6176-6181 (1970).
  4. Berry, C. E., Hare, J. M. Xanthine oxidoreductase and cardiovascular disease: Molecular mechanisms and pathophysiological implications. J Physiol. 555, 589-606 (2004).
  5. Kim, C., Kim, J. Y., Kim, J. H. Cytosolic phospholipase a(2), lipoxygenase metabolites, and reactive oxygen species. BMB Rep. 41 (2), 555-559 (2008).
  6. Armstead, W. M., Mirro, R., Busija, D. W., Leffler, C. W. Postischemic generation of superoxide anion by newborn pig brain. Am J Physiol. 255 (2), H401-H403 (1988).
  7. Mayer, B., et al. Nitric oxide synthase-catalyzed activation of oxygen and reduction of cytochromes: Reaction mechanisms and possible physiological implications. J Cardiovasc Pharmacol. 20, S54-S56 (1992).
  8. Du, G., Mouithys-Mickalad, A., Sluse, F. E. Generation of superoxide anion by mitochondria and impairment of their functions during anoxia and reoxygenation in vitro. Free Radic Biol Med. 25 (9), 1066-1074 (1998).
  9. Turrens, J. F. Mitochondrial formation of reactive oxygen species. J Physiol. 552, 335-344 (2003).
  10. Jiang, F., Zhang, Y., Dusting, G. J. Nadph oxidase-mediated redox signaling: Roles in cellular stress response, stress tolerance, and tissue repair. Pharmacol Rev. 63 (1), 218-242 (2011).
  11. Wang, Z., et al. The role of mitochondria-derived reactive oxygen species in hyperthermia-induced platelet apoptosis. PLoS One. 8 (9), e75044 (2013).
  12. Wachowicz, B., Olas, B., Zbikowska, H. M., Buczynski, A. Generation of reactive oxygen species in blood platelets. Platelets. 13 (3), 175-182 (2002).
  13. Vara, D., et al. Nadph oxidases are required for full platelet activation in vitro and thrombosis in vivo but dispensable for plasma coagulation and hemostasis. Arterioscler Thromb Vasc Biol. 41 (2), 683-697 (2021).
  14. Violi, F., Pignatelli, P. Platelet nox, a novel target for anti-thrombotic treatment. Thromb Haemost. 111 (5), 817-823 (2014).
  15. Begonja, A. J., et al. Platelet NAD(P)H-oxidase-generated ros production regulates alphaiibbeta3-integrin activation independent of the NO/CGMP pathway. Blood. 106 (8), 2757-2760 (2005).
  16. Vara, D. S., et al. Autocrine amplification of integrin αIIbβ3 activation and platelet adhesive responses by deoxyribose-1-phosphate. Thromb Haemost. 109 (6), 1108-1119 (2013).
  17. Essex, D. W. The role of thiols and disulfides in platelet function. Antioxid Redox Signal. 6 (4), 736-746 (2004).
  18. Arthur, J. F., et al. Ligand binding rapidly induces disulfide-dependent dimerization of glycoprotein vi on the platelet plasma membrane. J Biol Chem. 282 (42), 30434-30441 (2007).
  19. Arthur, J. F., Gardiner, E. E., Kenny, D., Andrews, R. K., Berndt, M. C. Platelet receptor redox regulation. Platelets. 19 (1), 1-8 (2008).
  20. Kim, K., Li, J., Tseng, A., Andrews, R. K., Cho, J. Nox2 is critical for heterotypic neutrophil-platelet interactions during vascular inflammation. Blood. 126 (16), 1952-1964 (2015).
  21. Hosseini, E., Solouki, A., Roudsari, Z. O., Kargar, F., Ghasemzadeh, M. Reducing state attenuates ectodomain shedding of GPVI while restoring adhesion capacities of stored platelets: Evidence addressing the controversy around the effects of redox condition on thrombosis. J Thromb Thrombolysis. 50 (1), 123-134 (2020).
  22. Jang, J. Y., et al. Reactive oxygen species play a critical role in collagen-induced platelet activation via SHP-2 oxidation. Antioxid Redox Signal. 20 (16), 2528-2540 (2014).
  23. Beckman, J. S., Koppenol, W. H. Nitric oxide, superoxide, and peroxynitrite: The good, the bad, and ugly. Am J Physiol. 271 (5), C1424-C1437 (1996).
  24. Koppenol, W. H., Kissner, R., Beckman, J. S. Syntheses of peroxynitrite: To go with the flow or on solid grounds. Methods Enzymol. 269, 296-302 (1996).
  25. Cammisotto, V., et al. Nox2-mediated platelet activation by glycoprotein (GP) VI: Effect of rivaroxaban alone and in combination with aspirin. Biochem Pharmacol. 163, 111-118 (2019).
  26. Yang, M., et al. Platelet CD36 promotes thrombosis by activating redox sensor ERK5 in hyperlipidemic conditions. Blood. 129 (21), 2917-2927 (2017).
  27. Yang, M., et al. Platelet CD36 signaling through ERK5 promotes caspase-dependent procoagulant activity and fibrin deposition in vivo. Blood Adv. 2 (21), 2848-2861 (2018).
  28. Freedman, J. E. Oxidative stress and platelets. Arterioscler Thromb Vasc Biol. 28 (3), s11-s16 (2008).
  29. Masselli, E., et al. ROS in platelet biology: Functional aspects and methodological insights. Int J Mol Sci. 21 (14), 4866 (2020).
  30. Madamanchi, N. R., Vendrov, A., Runge, M. S. Oxidative stress and vascular disease. Arterioscler Thromb Vasc Biol. 25 (1), 29-38 (2005).
  31. Riemersma, R. A., et al. Risk of angina pectoris and plasma concentrations of vitamins a, c, and e and carotene. Lancet. 337 (8732), 1-5 (1991).
  32. Rimm, E. B., et al. Vitamin E consumption and the risk of coronary heart disease in men. N Engl J Med. 328 (20), 1450-1456 (1993).
  33. Zhang, W., Zheng, Y., Yan, F., Dong, M., Ren, Y. Research progress of quercetin in cardiovascular disease. Front Cardiovasc Med. 10, 1203713 (2023).
  34. Russell, C., Keshavamurthy, S., Saha, S. Nutraceuticals in the management of cardiovascular risk factors: Where is the evidence. Cardiovasc Hematol Disord Drug Targets. 21 (3), 150-161 (2021).
  35. Rotariu, D., et al. Oxidative stress - complex pathological issues concerning the hallmark of cardiovascular and metabolic disorders. Biomed Pharmacother. 152, 113238 (2022).
  36. Hosseini, E., Ghasemzadeh, M., Atashibarg, M., Haghshenas, M. R. O. S. scavenger, n-acetyl-l-cysteine and NOX specific inhibitor, VAS2870 reduce platelets apoptosis while enhancing their viability during storage. Transfusion. 59 (4), 1333-1343 (2019).
  37. Hosseini, E., Nodeh, F. K., Ghasemzadeh, M. Gamma irradiation induces a pro-apoptotic state in longer stored platelets, without progressing to an overt apoptosis by day 7 of storage. Apoptosis. 28 (7-8), 1141-1153 (2023).
  38. Griendling, K. K., et al. Measurement of reactive oxygen species, reactive nitrogen species, and redox-dependent signaling in the cardiovascular system: A scientific statement from the American heart association. Circ Res. 119 (5), e39-e75 (2016).
  39. Murphy, M. P., et al. Guidelines for measuring reactive oxygen species and oxidative damage in cells and in vivo. Nat Metab. 4 (6), 651-662 (2022).
  40. Hempel, S. L., Buettner, G. R., O'malley, Y. Q., Wessels, D. A., Flaherty, D. M. Dihydrofluorescein diacetate is superior for detecting intracellular oxidants: Comparison with 2',7'-dichlorodihydrofluorescein diacetate, 5(and 6)-carboxy-2',7'-dichlorodihydrofluorescein diacetate, and dihydrorhodamine 123. Free Radic Biol Med. 27 (1-2), 146-159 (1999).
  41. Kalyanaraman, B., et al. Measuring reactive oxygen and nitrogen species with fluorescent probes: Challenges and limitations. Free Radic Biol Med. 52 (1), 1-6 (2012).
  42. Burkitt, M. J., Wardman, P. Cytochrome c is a potent catalyst of dichlorofluorescin oxidation: Implications for the role of reactive oxygen species in apoptosis. Biochem Biophys Res Commun. 282 (1), 329-333 (2001).
  43. Rota, C., Fann, Y. C., Mason, R. P. Phenoxyl free radical formation during the oxidation of the fluorescent dye 2',7'-dichlorofluorescein by horseradish peroxidase. Possible consequences for oxidative stress measurements. J Biol Chem. 274 (40), 28161-28168 (1999).
  44. Rota, C., Chignell, C. F., Mason, R. P. Evidence for free radical formation during the oxidation of 2'-7'-dichlorofluorescin to the fluorescent dye 2'-7'-dichlorofluorescein by horseradish peroxidase: Possible implications for oxidative stress measurements. Free Radic Biol Med. 27 (7-8), 873-881 (1999).
  45. Rodig, S. J. Attaching suspension cells to slides for staining. Cold Spring Harb Protoc. 2020 (12), (2020).
  46. Abubaker, A. A., Vara, D., Eggleston, I., Canobbio, I., Pula, G. A novel flow cytometry assay using dihydroethidium as redox-sensitive probe reveals NADPH oxidase-dependent generation of superoxide anion in human platelets exposed to amyloid peptide beta. Platelets. 30 (2), 181-189 (2019).
  47. Gaspar, R. S., et al. Protein disulphide isomerase and NADPH oxidase 1 cooperate to control platelet function and are associated with cardiometabolic disease risk factors. Antioxidants (Basel). 10 (3), 497 (2021).
  48. Gaspar, R. S., Ferreira, P. M., Mitchell, J. L., Pula, G., Gibbins, J. M. Platelet-derived extracellular vesicles express NADPH oxidase-1 (NOX-1), generate superoxide and modulate platelet function. Free Radic Biol Med. 165, 395-400 (2021).
  49. Vara, D., et al. NADPH oxidase 1 is a novel pharmacological target for the development of an antiplatelet drug without bleeding side effects. FASEB J. 34 (10), 13959-13977 (2020).
  50. Lopes-Pires, M. E., et al. Zinc regulates reactive oxygen species generation in platelets. Platelets. 32 (3), 368-377 (2021).
  51. Zielonka, J., Kalyanaraman, B. Hydroethidine- and mitoSOX-derived red fluorescence is not a reliable indicator of intracellular superoxide formation: Another inconvenient truth. Free Radic Biol Med. 48 (8), 983-1001 (2010).
  52. Linden, M. D. Platelet flow cytometry. Methods Mol Biol. 992, 241-262 (2013).
  53. Kauffman, M. E., et al. MitoSOX-based flow cytometry for detecting mitochondrial ROS. React Oxyg Species (Apex). 2 (5), 361-370 (2016).
  54. Sonkar, V. K., et al. NOX2 NADPH oxidase is dispensable for platelet activation or arterial thrombosis in mice. Blood Adv. 3 (8), 1272-1284 (2019).
  55. Dikalov, S. I., Kirilyuk, I. A., Voinov, M., Grigor'ev, I. A. EPR detection of cellular and mitochondrial superoxide using cyclic hydroxylamines. Free Radic Res. 45 (4), 417-430 (2011).
  56. Dikalov, S. I., Polienko, Y. F., Kirilyuk, I. Electron paramagnetic resonance measurements of reactive oxygen species by cyclic hydroxylamine spin probes. Antioxid Redox Signal. 28 (15), 1433-1443 (2018).

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