The protocol follows the guidelines and was approved by the University Hospital of Bordeaux Human Research Ethics Committee. Blood samples were obtained from healthy volunteers who provided informed consent, and these samples were processed according to institutional protocols. Donors who had taken any substances that could affect platelet function were excluded if such substances were taken within 10 days preceding the experiments. Informed consent was obtained from healthy volunteers to collect their blood and publish their data. The details of the reagents and the equipment used are listed in the Table of Materials.
1. Blood collection and preparation of washed platelets
<ol>
	<li>Collect peripheral venous blood in collection tubes containing the active anticoagulant Tri-sodium Citrate with Citric Acid and Dextrose (ACD) after discarding the first 3 mL.</li>
	<li>After collection, gently invert the tube to mix the blood with ACD and allow the mixture to rest at room temperature for 15 min before centrifugation.</li>
	<li>Count platelets using an automated cell counter before centrifugation to determine the correct speed for centrifugation (Table 1). Prepare platelet-rich plasma (PRP) by centrifugation at 250 x g for 10 min at room temperature (RT) without applying the brake to optimize platelet recovery. 
	NOTE: This step will produce three layers in the sample: the upper layer containing plasma, platelets, and a small fraction of white blood cells; a middle layer enriched with white blood cells; and a bottom layer consisting of red blood cells. Carefully transfer the PRP from the upper layer into a new 50 mL conical tube to avoid contamination with red and white blood cells.</li>
	<li>To the PRP, add 1 mL of ACD-A (see Table 2) and 6.25 &#181;L of apyrase per 9 mL. 
	NOTE: Apyrase, at a concentration of 0.02 U/mL, degrades traces of ATP or ADP secreted by the platelets, thereby preventing desensitization of platelet ADP receptors and maintaining platelet shape25. A prostacyclin PGI2 inhibitor (0.5 &#181;M) can also be included with apyrase to prevent platelet activation.</li>
	<li>Prepare the platelet pellet by centrifugation at RT at 1100 x g for 10 min. Aspirate the supernatant containing Platelet Poor Plasma (PPP) using a gentle method, such as a Pasteur pipette, and resuspend the pellet in 1 mL of washing buffer (see Table 3 and Table 4). Homogenize gently with a pipette, then add an additional 3-4 mL of washing buffer. 
	NOTE: The calcium level is reduced to prevent coagulation with clot factors present in the plasma. At this stage, the platelet-poor plasma has been completely removed, preventing clot formation and platelet activation by thrombin production.</li>
	<li>Centrifuge again at RT at 1100 x g for 10 min. Aspirate the supernatant and resuspend the pellet in 1 mL of washing buffer. Transfer 150 &#181;L of the platelet suspension to a 1.5 mL tube for cell counting using an automated cell counter. Then, resuspend the pellet with 3-4 mL of washing buffer.</li>
	<li>Perform a final centrifugation at 1100 x g for 10 min (at RT). Aspirate the supernatant and resuspend the platelet pellet in a volume of reaction buffer (see Table 5 and Table 6) to achieve a final concentration of 5 x 1011 platelets/L.</li>
	<li>Allow the washed platelets to rest for at least 30 min before experiments to allow residual inhibitors to wear off and for the platelets to acclimatize to the buffer.</li>
</ol>
2. Assay preparation
<ol>
	<li>Prepare all agonists and fluorochromes in the reaction buffer. Add 5 &#181;L of thrombin to 45 &#181;L of buffer (dilution 1:10) and 5 &#181;L of ionophore to 495 &#181;L of buffer (dilution 1:500).
	<ol>
		<li>For internal assays, benchmark non-activated platelets against those treated with single and dual agonists. Add, in different aliquots, 100 &#181;L of washed platelets at 50 x 109 platelets/L to: (i) 10 &#181;L of reaction buffer, (ii) 3 &#181;L of non-diluted collagen (final concentration of 30 &#181;g/mL), (iii) 10 &#181;L of pre-diluted thrombin (0.5 IU/mL), (iv) 3 &#181;L of non-diluted collagen + 10 &#181;L of pre-diluted thrombin, and (v) 10 &#181;L of pre-diluted ionophore (final concentration of 2 &#181;M), which is known to directly increase intracellular calcium levels as previously noted26. 
		NOTE: Treatment with a calcium ionophore, such as A23187 or ionomycin, is critical as it induces extensive phospholipid membrane scrambling and enhanced PS externalization.</li>
		<li>Gently shake each aliquot by hand and incubate at 37 &#176;C for 5 min. 
		NOTE: Longer agonist incubation (30 min) may allow better differentiation between platelet populations during the flow cytometry analysis process.</li>
	</ol>
	</li>
	<li>Add 5 &#181;L of non-diluted Annexin-V FITC to each microtube and incubate at room temperature for 10 min in the dark. 
	NOTE: A monoclonal antibody against one of the platelet-specific receptors, GPIX (CD42b) or &#945;IIb integrin (CD41, GPIIb), could also be included to better identify platelet cells.</li>
	<li>Add 500 &#181;L of reaction buffer to stop the process and proceed with the analysis of the sample using a flow cytometer.</li>
</ol>
3. Characterization of procoagulant platelets and microvesicles by Flow Cytometry (FC)
NOTE: For reliable platelet function analysis, utilize a flow cytometry (FC) instrument configured according to established standards27. The instrument must be capable of detecting forward scatter (FSC) and at least one fluorescence signal. Set the light scatter and fluorescence detectors to logarithmic gain. Dilute the samples suitably for FC to ensure only individual platelets are counted at a reduced data acquisition speed.
<ol>
	<li>Set the flow cytometer to a &#39;slow&#39; flow rate with a threshold of at least 10,000 platelet events. Monitor fluorescence emission using a pass filter for FITC.</li>
	<li>Gate the platelet population using forward scatter (FSC). Examine platelets and released microvesicles according to their sizes. Use density plots to identify platelets and released microvesicles for both resting and stimulated platelets; analyze each gated population independently.</li>
	<li>Distinguish procoagulant platelets according to the FL1 axis. Place the negative cut-off at the far right of the platelet population in the resting state (i.e., in the presence of reaction buffer). After platelet activation, classify all events localized to the right of this cut-off as platelets exposing phosphatidylserine.</li>
	<li>Differentiate microvesicles, being smaller than platelets, by their unique light scattering characteristics. Set the cut-off at the lowest FSC value observed in the resting platelet population. After activation, classify any phosphatidylserine-positive events falling below this threshold as microvesicles. 
	NOTE: Account for individual variation in platelet size by adjusting the cut-off if necessary. Ensure that less than 1% of microvesicles are present in the resting state.</li>
</ol>

Analyzing Platelet Phosphatidylserine and Microvesicle Release

The authors declare having no conflict of interest.

Recent studies on procoagulant platelets have highlighted their changes during several diseases1,28,29, underscoring the importance of their detailed analysis and characterization30,31,32. While current clinical tests for assessing platelet procoagulant formation are limited, there has been a significant rise in clinical interest over the past two decades. For example, the calcium-dependent scramblase TMEM16F (ANO6) is involved in procoagulant phosphatidylserine (PS) exposure33. Variants of TMEM16F are associated with Scott syndrome33,34,35, a rare bleeding disorder characterized by impaired phospholipid scrambling and reduced thrombin generation in activated platelets36. Moreover, elevated procoagulant platelet levels are linked with thrombotic strokes and coronary artery disease, sparking interest in therapies to curb platelet procoagulant activity37. Of the methods currently available to evaluate procoagulant platelet formation, assessment of PS exposure by Annexin-V binding and microvesicle release is one of the most commonly used38,39.
The protocol outlined here enables the isolation of human-purified platelets and the characterization of procoagulant platelets by measuring PS exposure and microvesicle release using flow cytometry. It is crucial to advise healthy volunteers to abstain from medications that may impact platelet function for at least 10 days before blood sampling. Blood collection requires care to avoid vein trauma or excessively slow flow, as these can induce thrombin generation and platelet activation. Sodium citrate-anticoagulated blood is not recommended for preparing washed platelets, as residual calcium can enable thrombin generation and platelet activation during centrifugation17. ACD-anticoagulated blood has a pH of 6.5 and a higher citrate concentration of 22 mmol/L compared to citrate-anticoagulated blood&#39;s pH of 7.5 and 13 mmol/L citrate concentration17. The acidic pH and lowered extracellular calcium concentration in ACD-anticoagulated blood prevent platelet aggregation24. Minimizing thrombin generation during the collection process is essential. A non-traumatic venipuncture using at least a 21 G needle is recommended to prevent venous stasis or excessive pressure. Additionally, the first few milliliters of blood should be discarded to avoid contamination with tissue factors and trace amounts of thrombin17. Minimizing the time between venipuncture and sample processing is also essential, as prolonged blood storage may lead to platelet activation, thereby increasing PS exposure and microvesicle release40. Specific attention must be paid during the entire platelet preparation procedure to prevent spontaneous platelet preactivation, which can occur during the preparation process itself41,17. Moreover, isolating purified platelets necessitates careful avoidance of contamination by other blood cells17. This involves thoroughly aspirating the supernatant after centrifuging the platelets to ensure a clean preparation. As the pellet is typically visible and firmly adhered to the tube wall, the supernatant can be readily aspirated using a pipette tip.
During centrifugation wash steps, platelets are resuspended in a buffer that simulates physiological conditions. Using washed platelets in a physiological buffer isolates them from anticoagulants and plasma, thereby preventing thrombin-mediated artifacts. While platelet-rich plasma (PRP) is a rapid method to obtain high platelet concentrations, its use for functional studies is limited by anticoagulant-induced pH changes and the plasma&#39;s extracellular calcium and protein content, which can influence platelet activation17. Furthermore, high levels of PS generated during platelet activation would be sufficient to promote the assembly of plasma coagulation complexes, subsequent thrombin burst, and clot formation in PRP, rendering experimental results uninterpretable. In contrast, resuspension in a buffer containing apyrase is recommended to maintain platelet functionality by preventing desensitization of purinergic receptors through the degradation of ATP and ADP42,43. Although repeated centrifugation and washing steps require more time, suspensions of washed platelets prepared by this method offer the advantage of greater stability (5-8 h) compared with citrated PRP preparations, which are stable for no more than 1-3 h23.
This study demonstrates the feasibility and simplicity of quantifying microvesicles by flow cytometry. However, microvesicle production can get lost in the debris and noise of the machine. Therefore, it is critical to ensure that the flow cytometer has undergone extensive cleaning and that the buffers used do not contain any contaminating particles that might lead to a high background during flow cytometry analysis. Alternatively, specialized flow cytometers with small particle detection capabilities can be used to identify microvesicle formation. The binding of fluorescein-labeled Annexin-V as a function of membrane PS content was also studied, highlighting the critical role of sufficient calcium concentrations in the buffer for Annexin-V binding and procoagulant platelet response44.
Data supporting the stability of platelet samples over time are currently scarce. Several laboratories have addressed this issue by stabilizing samples through the addition of a fixative solution, such as a formaldehyde-containing buffer. Adding a fixative to freshly collected platelet samples has the advantage of preserving samples for later analysis. Nevertheless, formaldehyde fixation can cause artifactually increased Annexin-V binding. Rochat et al. demonstrated that platelet fixation with a low-concentration, calcium-free formaldehyde solution does not alter the proportion of Annexin-V positive platelets45. However, researchers who require fixation should optimize their specific protocol to ensure that it does not cause artifactually altered results. Otherwise, samples should be processed immediately.
Another pathway that may also lead to PS exposure involves intrinsic apoptosis, a process known to regulate platelet lifespan in circulation46. Activating pro-apoptotic BAK and BAX proteins results in caspase activation, which promotes the externalization of PS, leading to the in vivo clearance of platelets. Recently, it has been suggested that another platelet surface marker, P-selectin/CD62P, in addition to Annexin-V, be used to distinguish procoagulants from apoptotic platelets47. Procoagulant platelets are expected to be positive for both PS and P-selectin markers, while apoptotic platelets are positive for Annexin-V but negative for P-selectin. However, time is also a differentiating factor between procoagulant and apoptotic platelets47. Generating procoagulant platelets in vitro is a rapid process, whereas intrinsic apoptosis is a slower process, taking hours48. In this context, the use of additional markers to assess procoagulant platelets is not necessary, simplifying the method. Sustained increases in intracellular Ca2+ and procoagulant platelet formation occur within a few minutes with very strong agonists, such as ionophore or combined activation with collagen and thrombin, whereas a BCL-XL inhibitor (or BH3 mimetic) triggers the formation of apoptotic platelets by activating pro-apoptotic BAK and BAX, causing mitochondrial outer membrane permeabilization, cytochrome C release, and caspase activation49.
In conclusion, the protocol described here facilitates the isolation of purified platelets and their subsequent characterization for procoagulant activity using standard laboratory equipment and flow cytometry. This procedure paves the way for future research on procoagulant platelets in patient blood samples, with potential applications in clinical practice, such as the diagnosis of the rare Scott syndrome50.

Platelet procoagulant formation is crucial to maintaining hemostasis1,2,3. This process involves agonist-induced expression of phospholipids on the platelet membrane, which is essential for the assembly of tenase and prothrombinase complexes1,3,4. After platelet activation, platelet microvesicles are also continuously released5,6. Microvesicles (50 nm to over 1 &#181;m in diameter) encapsulate both the membrane structure and cytoplasmic constituents of the originating cells7,8. Microvesicles are important mediators of intercellular signaling9,10 and also repositories of growth factors that promote wound healing and inflammatory modulation11,12. Microvesicles have also been suggested to be 50- to 100-fold more procoagulant than activated platelets13. Notably, recent evidence suggests that stored platelets become activated, leading to the production of microvesicles-this has potential ramifications for the practice of platelet transfusion therapy. Microvesicles&#39; extended storage duration and heightened procoagulant activity render them a promising substitute for platelets in transfusion applications14.
Methodological innovations have been made to, directly and indirectly, assess platelet procoagulant formation in both health and disease conditions4,14,15. Assessments of phosphatidylserine exposure and microvesicle release using purified platelets have provided new data on how platelet procoagulant responses are perturbed in diverse human diseases1,4. This growing field of research also opens avenues for therapeutic interventions1,4. However, the isolation and analysis of purified platelets from blood are time-consuming, necessitate specialized laboratory equipment, and are, thus, not currently adaptable for routine clinical diagnostics16,17. Analyzing phosphatidylserine exposure and microvesicle release is also challenging due to their small size and the number of positive events for markers of interest18,19.
Flow cytometry, leveraging fluorescent Annexin-V binding, has been a cornerstone in the evaluation of PS expression on platelets and microvesicles since its inception two decades ago20. It has gained widespread acceptance in the examination of procoagulant platelets and microvesicles21,22. Therefore, an optimized protocol is presented here that can be used for the isolation of purified platelets from blood samples using the technique developed by Cazenave&#39;s group23, and the subsequent characterization of phosphatidylserine exposure and microvesicle release by flow cytometry after platelet activation24. This protocol will facilitate further study and in-depth characterization of procoagulant platelets in clinical populations of interest.

<table><tbody><tr><td>(CD42b, GPIX) APC</td><td>Beckman Coulter</td><td>B13980</td><td></td></tr><tr><td>ACD-A blood collection tubes</td><td>BD Vacutainer</td><td>366645</td><td></td></tr><tr><td>Annexin-V FITC</td><td>BD Pharmingen</td><td>560931</td><td></td></tr><tr><td>Apyrase Grade VII</td><td>Sigma-Aldrich</td><td>A6410</td><td></td></tr><tr><td>Bovine Serum Albumin 30%</td><td>Sigma-Aldrich</td><td>A9576</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2, &lt;/sub&gt;0.25M</td><td>Sigma-Aldrich</td><td>C3881</td><td></td></tr><tr><td>Citric acid monohydrate</td><td>Merck</td><td>5949-29-1</td><td></td></tr><tr><td>Collagen&amp;nbsp;</td><td>Stago</td><td>86924</td><td></td></tr><tr><td>Glucose&amp;nbsp;</td><td>Sigma-Aldrich</td><td>G8270</td><td></td></tr><tr><td>Hepes&amp;nbsp;</td><td>Sigma-Aldrich</td><td>H3375</td><td></td></tr><tr><td>Ionophore&amp;nbsp;</td><td>Calbiochem/VWR</td><td>100105</td><td></td></tr><tr><td>KCl&amp;nbsp;</td><td>Merck</td><td>7447-40-7</td><td></td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;</td><td>Sigma-Aldrich</td><td>M0250</td><td></td></tr><tr><td>NaCl&amp;nbsp;</td><td>VWR</td><td>27810.295</td><td></td></tr><tr><td>NaOH&amp;nbsp;</td><td>Merck</td><td>1.06498</td><td></td></tr><tr><td>Sodium hydrogen carbonate</td><td>Merck</td><td>6329</td><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td></tr><tr><td>Thrombin&amp;nbsp;</td><td>Hyphen Biomed</td><td>EZ 006 A</td><td></td></tr><tr><td>Trisodium citrate dihydrate&amp;nbsp;</td><td>Sigma-Aldrich</td><td>G8270</td><td>Na&lt;sub&gt;3&lt;/sub&gt;C&lt;sub&gt;6&lt;/sub&gt;H&lt;sub&gt;5&lt;/sub&gt;O&lt;sub&gt;7*&lt;/sub&gt;2H&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>&amp;alpha;IIb integrin (CD41, GPIIb) PC7</td><td>Beckman Coulter</td><td>6607115</td><td></td></tr></tbody></table>

procoagulant platelet characterization by measuring phosphatidylserine exposure and microvesicle release from human purified platelets

Activated platelets promote coagulation primarily by exposing the procoagulant phospholipid phosphatidylserine (PS) on their outer membrane surfaces and releasing PS-expressing microvesicles that retain the original membrane architecture and cytoplasmic components of their originating cells. The accessibility of phosphatidylserine facilitates the binding of major coagulation factors, significantly amplifying the catalytic efficiency of coagulation enzymes, while microvesicle release acts as a pivotal mediator of intercellular signaling. Procoagulant platelets play a crucial role in clot stabilization during hemostasis, and their increased proportion in the bloodstream correlates with an increased risk of thrombosis. It has also been shown that platelet microvesicles are rich in growth factors that promote wound healing and inflammatory modulation. Analyzing phosphatidylserine exposure and microvesicle release using flow cytometry poses significant challenges due to their small size and the limited number of positive events for markers of interest. Despite considerable advances in the last decade, methods for assessing phosphatidylserine exposure and microvesicle release remain a work in progress. Unfortunately, no single universally applicable protocol exists, and several factors must be evaluated to determine the most appropriate methodology for each specific application. Here, we describe a detailed protocol for isolating washed platelets from human blood, followed by collagen and/or thrombin activation, to measure the exposure of phosphatidylserine and microvesicle release that characterize procoagulant platelets. This protocol is designed to facilitate the initial preparation of platelet-rich plasma and the isolation of washed platelets. Finally, phosphatidylserine exposure and microvesicle release are quantified by flow cytometry, enabling the identification of procoagulant platelets.

Quantification of procoagulant platelets and microvesicles is achieved using Annexin-V staining, with at least 50,000 events recorded per sample. As stated in step 2.2, baseline platelet measurements were taken from samples incubated with reaction buffer, as depicted in Figure 1A. Platelet sizes were gauged using the median forward scatter (FSC) value. While FSC is influenced by various factors, it remains a common proxy for cell size estimation, including that of platelets. Density plots of area (Figure 1A) were used to identify and gate resting platelets and released microvesicles.
In the case of non-activated platelets, less than 2.9% displayed phosphatidylserine (PS) as determined by Annexin-V staining. Upon activation of washed platelets with either collagen alone or in conjunction with thrombin or calcium ionophore, there was an observable rise in PS exposure. Collagen alone prompted a 26.7% increase in PS-positive platelets. Thrombin had minimal effects on PS exposure (9.1%), but when associated with collagen, it increased the percentage of PS-exposing platelets to 36.2%. For reference, treatment with 2 &#181;M ionophore resulted in 49.6% of platelets becoming procoagulant.
Figure 1 illustrates that the baseline proportion of microvesicles from washed platelets in a healthy individual was minimal (0.3%). While thrombin and collagen minimally increased microvesicle formation, it was only with the combination of collagen and thrombin that microvesicle formation significantly increased (Figure 1D; 11.4%). Adding 2 &#181;M ionophore resulted in 44.0% of the microvesicles formed by control platelets being PS-exposing.
Figure 2 provides information regarding the variation, repeatability, and reproducibility to be expected from this process.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67042/67042fig01.jpg" /> 
Figure 1: PS exposure and microvesicle formation in resting and activated control platelets. Platelets were either activated or unactivated&#160;(Resting) (A), with 30 &#181;g/mL collagen&#160;(B), with 0.5 U/mL thrombin&#160;(C), with combinations of collagen and thrombin&#160;(D)&#160;with 2 &#181;M ionophore. Representative dot plots display platelet size (forward scatter (FSC log)) versus Annexin-V binding. Horizontal and vertical lines&#160;(E) delineate the platelet-negative population. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/67042/67042fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67042/67042fig02.jpg" /> 
Figure 2: Repeatability and reproducibility of PS exposure and microvesicle formation after incubation with ionophore. Analysis of PS exposure and microvesicle formation after incubation with either 2 &#181;M or 5 &#181;M ionophore in washed platelets, treated or untreated with citrate (a calcium chelator): (A) from 4 different healthy controls; (B) from one healthy control repeated 5 times; and (C) from two healthy controls prepared by two different technicians. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/67042/67042fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Platelet count in whole blood (109 per mL)</td>
			<td>Relative centrifugal forces (g)</td>
		</tr>
		<tr>
			<td>&#60;30</td>
			<td>70</td>
		</tr>
		<tr>
			<td>30-50</td>
			<td>100</td>
		</tr>
		<tr>
			<td>51-100</td>
			<td>130</td>
		</tr>
		<tr>
			<td>101-150</td>
			<td>200</td>
		</tr>
		<tr>
			<td>151-300</td>
			<td>250</td>
		</tr>
		<tr>
			<td>301-600</td>
			<td>290</td>
		</tr>
		<tr>
			<td>&#62;600</td>
			<td>340</td>
		</tr>
	</tbody>
</table>
Table 1: Relative centrifugal forces for platelet-rich plasma preparation in relation to platelet count in whole blood.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>Amount</td>
		</tr>
		<tr>
			<td>Citric acid monohydrate (C6H8O7.H2O, 38 mM)</td>
			<td>8 g</td>
		</tr>
		<tr>
			<td>Trisodium citrate dihydrate (Na3C6H5O7.2 H2O, 61 mM)</td>
			<td>180 g</td>
		</tr>
		<tr>
			<td>Glucose (C6H12O6, 136 mM)</td>
			<td>24.6 g</td>
		</tr>
		<tr>
			<td colspan="2">The pH of the solution should not be adjusted. Keep the solution at 4 &#176;C.</td>
		</tr>
	</tbody>
</table>
Table 2: Preparation of a 1 L aqueous solution of acid-citrate-dextrose (ACD).
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>Amount</td>
		</tr>
		<tr>
			<td>Citric acid monohydrate (C6H8O7.H2O, 36 mM)</td>
			<td>7.56 g</td>
		</tr>
		<tr>
			<td>NaCl (103 mM)</td>
			<td>6 g</td>
		</tr>
		<tr>
			<td>NaOH</td>
			<td>4 g</td>
		</tr>
		<tr>
			<td>&#160;MgCl2.6H2O (1 mM)&#160;</td>
			<td>0.203 g</td>
		</tr>
		<tr>
			<td colspan="2">Adjust the pH at 6.5. Keep the solution at 4 &#176;C.&#160;</td>
		</tr>
	</tbody>
</table>
Table 3: Preparation of stock solution of Washing Buffer.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>Amount</td>
		</tr>
		<tr>
			<td>30% Bovine Serum Albumin (BSA)</td>
			<td>583 &#181;L</td>
		</tr>
		<tr>
			<td>CaCl2, 0.25 M</td>
			<td>400 &#181;L</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>40 mg</td>
		</tr>
		<tr>
			<td>Apyrase</td>
			<td>31.5 &#181;L&#160;</td>
		</tr>
	</tbody>
</table>
Table 4: Preparation of a 50 mL extemporaneous solution of Washing Buffer from the stock solution.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>Amount</td>
		</tr>
		<tr>
			<td>NaCl (137 mM)</td>
			<td>8 g</td>
		</tr>
		<tr>
			<td>KCl (2.7 mM)</td>
			<td>0.2 g</td>
		</tr>
		<tr>
			<td>MgCl2.6H2O (1 mM)</td>
			<td>0.203 g</td>
		</tr>
		<tr>
			<td>HEPES (5 mM)&#160;</td>
			<td>1.19 g</td>
		</tr>
		<tr>
			<td>NaHCO3 (12 mM)&#160;</td>
			<td>1 g</td>
		</tr>
		<tr>
			<td>NaH2PO4.2H2O; 0.3 mM</td>
			<td>0.05 g</td>
		</tr>
		<tr>
			<td colspan="2">Adjust the pH at 7.4. Keep the solution at 4 &#176;C.&#160;</td>
		</tr>
	</tbody>
</table>
Table 5: Preparation of a stock solution of Reaction Buffer.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>Amount</td>
		</tr>
		<tr>
			<td>30% Bovine Serum Albumin (BSA)</td>
			<td>167 &#181;L</td>
		</tr>
		<tr>
			<td>CaCl2, 0.25 M</td>
			<td>400 &#181;L&#160;</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>50 mg</td>
		</tr>
	</tbody>
</table>
Table 6: Preparation of a 50 mL extemporaneous solution of Reaction Buffer from the stock solution.

Procoagulant Platelet Characterization by Measuring Phosphatidylserine Exposure and Microvesicle Release from Human Purified Platelets

Procoagulant platelet formation has been correlated with an increased risk of thrombosis. Presented here is a precise protocol for isolating washed platelets from human blood, intended to quantify the exposure of phosphatidylserine and microvesicle release, which are distinctive features of procoagulant platelets.

Activated platelets promote coagulation primarily by exposing the procoagulant phospholipid phosphatidylserine (PS) on their outer membrane surfaces and ...

procoagulant-platelet-characterization-measuring-phosphatidylserine

Universidad San Sebastian

Research

JoVE Journal

Biology

478 Views.  University Hospital of Bordeaux. Procoagulant platelet formation has been correlated with an increased risk of thrombosis. Presented here is a precise protocol for isolating washed platelets from human blood, intended to quantify the exposure of phosphatidylserine and microvesicle release, which are distinctive features of procoagulant platelets.

Video: Procoagulant Platelet Characterization by Measuring Phosphatidylserine Exposure and Microvesicle Release from Human Purified Platelets

Analyzing the Interaction of Fluorescent-Labeled Proteins with Artificial Phospholipid Microvesicles using Quantitative Flow Cytometry

In the human body, most of the major physiologic reactions involved in the immune response and blood coagulation proceed on the membranes of cells. An important first step in any membrane-dependent reaction is binding of protein on the phospholipid membrane. An approach to studying protein interaction with lipid membranes has been developed using fluorescently labeled proteins and flow cytometry. This method allows the study of protein-membrane interactions using live cells and natural or artificial phospholipid vesicles. The advantage of this method is the simplicity and availability of reagents and equipment. In this method, proteins are labeled using fluorescent dyes. However, both self-made and commercially available, fluorescently labeled proteins can be used. After conjugation with a fluorescent dye, the proteins are incubated with a source of the phospholipid membrane (microvesicles or cells), and the samples are analyzed by flow cytometry. The obtained data can be used to calculate the kinetic constants and equilibrium Kd. In addition, it is possible to estimate the approximate number of protein binding sites on the phospholipid membrane using special calibration beads.

Here, we describe a set of methods for characterizing the interaction of proteins with membranes of cells or microvesicles.

In the human body, most of the major physiologic reactions involved in the immune response and blood coagulation proceed on the membranes of cells. An ...

Biochemistry

A Microfluidic Flow Chamber Model for Platelet Transfusion and Hemostasis Measures Platelet Deposition and Fibrin Formation in Real-time

Microfluidic models of hemostasis assess platelet function under conditions of hydrodynamic shear, but in the presence of anticoagulants, this analysis is restricted to platelet deposition only. The intricate relationship between Ca2+-dependent coagulation and platelet function requires careful and controlled recalcification of blood prior to analysis. Our setup uses a Y-shaped mixing channel, which supplies concentrated Ca2+/Mg2+ buffer to flowing blood just prior to perfusion, enabling rapid recalcification without sample stasis. A ten-fold difference in flow velocity between both reservoirs minimizes dilution. The recalcified blood is then perfused in a collagen-coated analysis chamber, and differential labeling permits real-time imaging of both platelet and fibrin deposition using fluorescence video microscopy. The system uses only commercially available tools, increasing the chances of standardization. Reconstitution of thrombocytopenic blood with platelets from banked concentrates furthermore models platelet transfusion, proving its use in this research domain. Exemplary data demonstrated that coagulation onset and fibrin deposition were linearly dependent on the platelet concentration, confirming the relationship between primary and secondary hemostasis in our model. In a timeframe of 16 perfusion min, contact activation did not take place, despite recalcification to normal Ca2+ and Mg2+ levels. When coagulation factor XIIa was inhibited by corn trypsin inhibitor, this time frame was even longer, indicating a considerable dynamic range in which the changes in the procoagulant nature of the platelets can be assessed. Co-immobilization of tissue factor with collagen significantly reduced the time to onset of coagulation, but not its rate. The option to study the tissue factor and/or the contact pathway increases the versatility and utility of the assay.

This paper describes an experimental model of hemostasis that simultaneously measures platelet function and coagulation. Platelet and fibrin fluorescence is measured in real-time, and platelet adhesion rate, coagulation rate, and onset of coagulation are determined. The model is used to determine platelet procoagulant properties under flow in concentrates for transfusion.

Microfluidic models of hemostasis assess platelet function under conditions of hydrodynamic shear, but in the presence of anticoagulants, this analysis is ...

Bioengineering

Turbidimetry on Human Washed Platelets: The Effect of the Pannexin1-inhibitor Brilliant Blue FCF on Collagen-induced Aggregation

Turbidimetry is a laboratory technique that is applied to measure the aggregation of platelets suspended in either plasma (platelet-rich plasma, PRP) or in buffer (washed platelets), by the use of one or a combination of agonists. The use of washed platelets separated from their plasma environment and in the absence of anticoagulants allows for studying intrinsic platelet properties. Among the large panel of agonists, arachidonic acid (AA), adenosine di-phosphate (ADP), thrombin and collagen are the most frequently used. The aggregation response is quantified by measuring the relative optical density (OD) over time of platelet suspension under continuous stirring. Platelets in homogeneous suspension limit the passage of light after the addition of an agonist, platelet shape change occurs producing a small transitory increase in OD. Following this initial activation step, platelet clots form gradually, allowing the passage of light through the suspension as a result of decreased OD. The aggregation process is ultimately expressed as a percentage, compared to the OD of platelet-poor plasma or buffer. Rigorous calibration is thus essential at the beginning of each experiment. As a general rule: calibration to 0% is set by measuring the OD of a non-stimulated platelet suspension while measuring the OD of the suspension medium containing no platelets represents a value of 100%. Platelet aggregation is generally visualized as a real-time aggregation curve. Turbidimetry is one of the most commonly used laboratory techniques for the investigation of platelet function and is considered as the historical gold standard and used for the development of new pharmaceutical agents aimed at inhibiting platelet aggregation. Here, we describe detailed protocols for 1) preparation of human washed platelets and 2) turbidimetric analysis of collagen-induced aggregation of human washed platelets pretreated with the food dye Brilliant Blue FCF that was recently identified as an inhibitor of Pannexin1 (Panx1) channels.

We describe a straightforward method for the isolation of washed platelets from human blood followed by agonist-induced platelet aggregation measurements by turbidimetry. As an example we apply this method for studying the aggregation response of human platelets to collagen after a pre-incubation with the Pannexin1 channel inhibitor Brilliant Blue FCF.

Turbidimetry is a laboratory technique that is applied to measure the aggregation of platelets suspended in either plasma (platelet-rich plasma, PRP) or ...

Medicine

Routine Screening Method for Microparticles in Platelet Transfusions

Platelet inventory management based on screening microparticle content in platelet concentrates is a new quality improvement initiative for hospital blood banks. Cells fragment off microparticles (MP) when they are stressed. Blood and blood components may contain cellular fragments from a variety of cells, most notably from activated platelets. When performing their roles as innate immune cells and major players in coagulation and hemostasis, platelets change shape and generate microparticles. With dynamic light scattering (DLS)-based microparticle detection, it is possible to differentiate activated (high microparticle) from non-activated (low microparticle) platelets in transfusions, and optimize the use of this scarce blood product. Previous research suggests that providing non-activated platelets for prophylactic use in hematology-oncology patients could reduce their risk of becoming refractory and improve patient care. The goal of this screening method is to routinely differentiate activated from non-activated platelets. The method described here outlines the steps to be performed for routine platelet inventory management in a hospital blood bank: obtaining a sample from a platelet transfusion, loading the sample into the capillary for DLS measurement, performing the DLS test to identify microparticles, and using the reported microparticle content to identify activated platelets.

Platelet inventory management based on screening microparticle content in platelet concentrates is a new quality improvement initiative in hospital blood banks. The goal is to differentiate activated from non-activated platelets to optimize the use of platelets. Providing non-activated platelets to hematology-oncology patients might reduce their high risk to become refractory.

Platelet inventory management based on screening microparticle content in platelet concentrates is a new quality improvement initiative for hospital blood ...

Live-cell Imaging of Platelet Degranulation and Secretion Under Flow

Blood platelets are essential players in hemostasis, the formation of thrombi to seal vascular breaches. They are also involved in thrombosis, the formation of thrombi that occlude the vasculature and injure organs, with life-threatening consequences. This motivates scientific research on platelet function and the development of methods to track cell-biological processes as they occur under flow conditions.
A variety of flow models are available for the study of platelet adhesion and aggregation, two key phenomena in platelet biology. This work describes a method to study real-time platelet degranulation under flow during activation. The method makes use of a flow chamber coupled to a syringe-pump setup that is placed under a wide-field, inverted, LED-based fluorescence microscope. The setup described here allows for the simultaneous excitation of multiple fluorophores that are delivered by fluorescently labeled antibodies or fluorescent dyes. After live-cell imaging experiments, the cover glasses can be further processed and analyzed using static microscopy (i.e., confocal microscopy or scanning electron microscopy).

This work describes a fluorescence microscopy-based method for the study of platelet adhesion, spreading, and secretion under flow. This versatile platform enables the investigation of platelet function for mechanistic research on thrombosis and hemostasis.

Blood platelets are essential players in hemostasis, the formation of thrombi to seal vascular breaches. They are also involved in thrombosis, the ...

A Uniform Shear Assay for Human Platelet and Cell Surface Receptors via Cone-plate Viscometry

Many biological cells/tissues sense the mechanical properties of their local environments via mechanoreceptors, proteins that can respond to forces like pressure or mechanical perturbations. Mechanoreceptors detect their stimuli and transmit signals via a great diversity of mechanisms. Some of the most common roles for mechanoreceptors are in neuronal responses, like touch and pain, or hair cells which function in balance and hearing. Mechanosensation is also important for cell types which are regularly exposed to shear stress such as endothelial cells, which line blood vessels, or blood cells which experience shear in normal circulation. Viscometers are devices that detect the viscosity of fluids. Rotational viscometers may also be used to apply a known shear force to fluids. The ability of these instruments to introduce uniform shear to fluids has been exploited to study many biological fluids including blood and plasma. Viscometry may also be used to apply shear to the cells in a solution, and to test the effects of shear on specific ligand-receptor pairs. Here, we utilize cone-plate viscometry to test the effects of endogenous levels of shear stress on platelets treated with antibodies against the platelet mechanosensory receptor complex GPIb-IX.

We describe an in-solution method to apply uniform shear to platelet surface receptors using cone-plate viscometry. This method may also be used more broadly to apply shear to other cell types and cell-fragments and need not target a specific ligand-receptor pair.

Many biological cells/tissues sense the mechanical properties of their local environments via mechanoreceptors, proteins that can respond to forces like ...

Purification of Platelets from Mouse Blood

Platelets are purified from whole blood to study their functional properties, which should be free from red blood cells (RBC), white blood cells (WBC), and plasma proteins. We describe here purification of platelets from mouse blood using three-fold more iohexol gradient medium relative to blood sample volume and centrifugation in a swinging bucket rotor at 400 x g for 20 min at 20 &#176;C. The recovery/yield of the purified platelets were 18.2-38.5%, and the purified platelets were in a resting state, which did not contain any significant number of RBC and WBC. The purified platelets treated with thrombin showed up to 93% activation, indicating their viability. We confirmed that the purified platelets are sufficiently pure using flow cytometric and microscopic evaluation. These platelets can be used for gene expression, activation, granule release, aggregation, and adhesion assays. This method can be used for purification of platelets from the blood of other species as well.

Here, mouse blood was collected in the presence of an anti-coagulant. The platelets were purified by iohexol gradient medium using low speed centrifugation. The platelets were activated with thrombin to investigate if they were viable. The quality of the purified platelets was analyzed by flow cytometry and microscopy.

Platelets are purified from whole blood to study their functional properties, which should be free from red blood cells (RBC), white blood cells (WBC), ...

Platform for Assessing Platelet Function in Trauma and Transfusion

Microfluidics in Assessing Platelet Function

Microfluidics incorporate physiologically relevant substrates and flows that mimic the vasculature and are, therefore, a valuable tool for studying aspects of thrombosis and hemostasis. At high-shear environments simulating arterial flow, a microfluidic assay facilitates the study of platelet function, as platelet-rich thrombi form in a localized stenotic region of a flow channel. Utilizing devices that allow for small sample volume can additionally aid in evaluating platelet function under flow from volume-limited patient samples or animal models. Studying trauma patient samples or samples following platelet product transfusion may aid in directing therapeutic strategies for patient populations in which platelet function is critical. Effects of platelet inhibition via pharmacological agents can also be studied in this model. The objective of this protocol is to establish a microfluidic platform that incorporates physiologic flow, biological surfaces, and relevant hemostatic mechanisms to assess platelet function with implications for the study of trauma induced coagulopathy and transfusion medicine.

Platelet function under flow can be assessed and simulated hemostatic resuscitation can be modeled using a microfluidic device, which has applications in trauma and transfusion medicine.

Microfluidics incorporate physiologically relevant substrates and flows that mimic the vasculature and are, therefore, a valuable tool for studying ...

An In Vitro Method for the Differentiation of Megakaryocytes and Platelet Formation

Source: Perdomo, J., et al. <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/56420">Megakaryocyte Differentiation and Platelet Formation from Human Cord Blood-derived CD34+ Cells.</a> J. Vis. Exp. (2017)
This video demonstrates a technique for the differentiation of megakaryocytes and the formation of platelets. Hematopoietic stem progenitor cells &#8212; isolated from human umbilical cord blood &#8212; differentiate into mature megakaryocytes in the presence of thrombopoietin. The mature megakaryocytes produce cytoplasmic extensions termed proplatelets, which give rise to individual platelets.

Metoda in vitro do różnicowania megakariocytów i tworzenia płytek krwi

Source: Perdomo, J., et al. Megakaryocyte Differentiation and Platelet Formation from Human Cord Blood-derived CD34+ Cells. J. Vis. Exp. (2017)
This video ...

JoVE Encyclopedia of Experiments

Immunology

Immune Systems and Components

Cell Differentiation Techniques

Purification and Activation of Platelets from Murine Whole Blood

Source: Narciso, M. G., et al. <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/59803">Purification of Platelets from Mouse Blood.</a> J. Vis. Exp. (2019).
In this video, we describe a protocol for purifying platelets from murine whole blood and their subsequent activation with the platelet agonist thrombin. When analyzed with flow cytometry, the purified resting platelets are characterized based on the expression of their cell-surface marker CD41, whereas purified activated platelets co-express both the surface markers CD41 and CD62P.

Oczyszczanie i aktywacja płytek krwi z mysiej krwi pełnej

Source: Narciso, M. G., et al. Purification of Platelets from Mouse Blood. J. Vis. Exp. (2019).
In this video, we describe a protocol for purifying ...