Skip to main content
NCBI home page
Transfusion Medicine and Hemotherapy logoLink to Transfusion Medicine and Hemotherapy
. 2019 May 7;46(4):267–275. doi: 10.1159/000499958

Assessment of Time-Dependent Platelet Activation Using Extracellular Vesicles, CD62P Exposure, and Soluble Glycoprotein V Content of Platelet Concentrates with Two Different Platelet Additive Solutions

Sami Valkonen a,b, Birte Mallas b, Ulla Impola b, Anne Valkeajärvi b, Juha Eronen b, Kaija Javela b, Pia R-M Siljander a, Saara Laitinen b,*
PMCID: PMC6739716  PMID: 31700509

Abstract

Novel analytical measures are needed to accurately monitor the properties of platelet concentrates (PCs). Since activated platelets produce platelet-derived extracellular vesicles (EVs), analyzing EVs of PCs may provide additional information about the condition of platelets. The prospect of using EVs as an auxiliary measure of platelet activation state was investigated by examining the effect of platelet additive solutions (PASs) on EV formation and platelet activation during PC storage. The time-dependent activation of platelets in PCs with PAS-B or with the further developed PAS-E was compared by measuring the exposure of CD62P by flow cytometry and the content of soluble glycoprotein V (sGPV) of PCs by an immunoassay. Changes in the concentration and size distribution of EVs were determined using nanoparticle tracking analysis. A time-dependent increase in platelet activation in PCs was demonstrated by increased CD62P ex­posure, sGPV content, and EV concentration. Using these strongly correlating parameters, PAS-B platelets were shown to be more activated compared to PAS-E platelets. Since the EV concentration correlated well with the established platelet activation markers CD62P and sGPV, it could potentially be used as a complementary parameter for platelet activation for PCs. More detailed characterization of the resulting EVs could help to understand how the PC components contribute the functional effects of transfused PCs.

Keywords: Platelet concentrate, Extracellular vesicle, Platelet activation, Platelet additive solution, CD62P, Soluble glycoprotein V

Introduction

Platelet concentrates (PC) are manufactured for patients with hemostatic problems, e.g., excessive bleeding or thrombocytopenia in cancer. The average life span of a platelet in blood circulation is 10 days [1], and the storage time of PC is typically 5–7 days [2] mainly due to an increased risk of bacterial contamination during extended storage at room temperature (RT) [3]. Efforts are made to increase the storage time of PCs by developing platelet additive solutions (PASs), improving protocols in PC preparation, pathogen inactivation, and by more detailed quality control [4, 5, 6].

A crucial aspect of the quality control of PCs is the determination of platelet activation state. One widely used marker of platelet activation is P-selectin (CD62P) exposure of platelets. CD62P is transferred to the platelet plasma membrane through the fusion of α-granules upon activation [7]. Another platelet-specific marker is the soluble form of glycoprotein V (sGPV), which is released from activated platelets through proteolytic cleavage [8]. The transmembrane form of glycoprotein V is located on the platelet surface as part of a complex with glycoproteins Ib and IX, which is the major receptor for von Willebrand factor [9], and it also participates in thrombin [10] and collagen [11] binding. Other current quality control assays of PCs include the quantification of platelet metabolites (glucose, lactate, pH) and dissolved gases (pO2, pCO2) as well as the determination of platelet function using the extent of shape change and hypotonic shock response. Also, different platelet parameters, such as the mean platelet volume and platelet count, are commonly monitored to examine the quality of platelets [6].

Besides the platelets' main role in regulation of hemostasis, they have been shown to influence immune responses [7], which is an aspect to consider when PCs are administered to patients. During storage of PCs, platelets liberate a large variety of bioactive components that have been proposed to relate to adverse proinflammatory effects observed in storage lesion [12, 13]. Although several different markers for measuring storage lesion have been suggested, a gold standard to evaluate the usability of PCs for transfusion has not yet been established [6]. Therefore, novel markers are needed to assess the condition of platelets in more detail.

Besides platelets, the PCs contains extracellular vesicles (EVs), which in their majority are derived from platelets, but which also originate from red blood cells and leukocytes residually present in the plasma fraction of PCs [14]. One of the first functions in which platelet-derived EVs were shown to participate was hemostasis [15, 16], implemented by the interaction of coagulation factors on the phosphatidylserine surface of the EVs [17]. Additionally, EVs are considered to be biomarkers of thrombotic and inflammatory diseases as well as cancer [18, 19], and in general EVs have already been shown to mediate several (patho)physiological processes [20, 21]. Furthermore, EVs have been suggested to contribute to adverse transfusion-related reactions [22], underscoring the need to understand the possible effector functions of EVs in transfusion. In addition to the possible effects of the transfused EVs in patients, EV generation during the storage of PCs could be considered as auxiliary parameter to monitor the activation state of platelets, since the generation of platelet-derived EVs is dependent on the aging and activation status of platelets [23, 24].

In the current study, EVs of PC were quantified to see whether the EV content could be utilized as a parameter of the activation state of platelets in PCs together with the recognized platelet activation markers, CD62P exposure of platelets, and sGPV content. This was investigated by examining the platelet activation state in aging PCs with different PASs.

Materials and Methods

Sample Collection

Standard leukocyte-reduced PCs, each derived from buffy coats of four ABO RhD-matched whole blood donations with PAS-B or PAS-E, were obtained from the Finnish Red Cross Blood Service (Helsinki, Finland) and handled anonymously, as accepted by the Finnish Supervisory Authority for Welfare and Health (Valvira, Helsinki, Finland). The exact compositions of PAS-B and PAS-E, also known as PAS-2 and modified PAS-3 [25], respectively, have been reported elsewhere [26].

Sterile sampling was done using 50-mL syringes (Henke-Sass; Wolf GmbH, Tuttlingen, Germany) and 18-gauge needles (Terumo, Tokyo, Japan). Before sampling, the contents of the storage bag's tube were emptied into the storage bag and the PC was mixed by gently turning it from side to side five times. This procedure was repeated three times to obtain a representative sample. After extracting 20 mL of sample via the storage bag's tube, the tube was resealed. The sampling days (d) were d1, d2, d5, and d8 counting from the blood donation (d0), d1 being the production day of PC. The d1 sampling was performed within 2 h once the PCs were available from the production line, approximately at 3 p.m., whereas the d2–d8 samplings were performed at 9 a.m. PCs were stored at 22°C under constant horizontal agitation.

Determination of CD62P Exposure of Platelets

The CD62P expression on the platelet surface was determined by flow cytometry using 1 mL of PC sample. PC samples were diluted 1:100 (to approximately 1 × 107/mL) using a diluent consisting of the same PAS used for PC production (either PAS-B or PAS-E; SSP or SSP+ [Macopharma, Tourcoing, France]) with 0.5% w/v bovine serum albumin (Sigma-Aldrich, St. Louis, MO, USA). Fifty microliters of diluted PC sample were labeled using 2 µL of fluorescein isothiocyanate (FITC)-coupled anti-CD41 (FITC mouse anti-human CD41, clone HIP8 [Becton Dickinson, Franklin Lakes, NJ, USA]) and 5 µL of phycoerythrin-cyanine 5 (PE-Cy5)-coupled anti-CD62P (PE-Cy5 mouse anti-human CD62P, clone AK-4 [Becton Dickinson]). For each sample an isotype control sample (50 µL of diluted PC sample labeled with 5 µL of PE-Cy5 mouse IgG1 κ isotype control, clone MOPC-21 [Becton Dickinson]) and a thrombin-activated positive control sample with maximum CD62P expression (50 µL of diluted PC sample labeled with 2 µL of CD41-FITC and 5 µL of CD62P-PE-Cy5 and activated with 1 IU/mL thrombin [Roche, Basel, Switzerland]) were prepared. Samples and controls were labeled and analyzed in BD TruCount tubes (Becton Dickinson) containing a known number of fluorescent beads. Samples were analyzed in singlicates since previously the CD62P measurements had been found well repeatable [27]. After labeling samples were mixed, incubated for 20 min at RT in the dark, further diluted with 500 µL of diluent, and stored in the dark until analysis.

The samples and controls were analyzed with a Navios flow cytometer (Beckman Coulter, Brea, CA, USA) at “high-flow” speed. The forward (FS) and side scatter detectors' (SS) volt and gain settings had been adjusted such that the platelet population was centered in the FS-SS dot plot (at around 101) and the fluorescence detectors' FL1 (FITC) and FL4 (PE-Cy5) settings such that the detected fluorescence signals were well within the displayed ranges for all samples. The TruCount beads were gated based on their fluorescence in FL1, FL2, and FL3 channels and a platelet gate had been defined in the FL1-SS dot plot. For each sample 5,000 bead events were acquired, corresponding to about 60,000 platelet events. The gated platelet population was used to calculate the percentage of CD62P-positive platelets, defined as:

  • Based on the isotype control, a threshold was set to include 1% of all events with the highest fluorescence in the FL4 channel. All events with FL4 fluorescence above this threshold were defined as CD62P-positive in comparison to the isotype control.

  • Based on the positive control, a threshold was defined to include 95% of the thrombin-activated platelet population with the highest fluorescence. All platelets with FL4 fluorescence above this threshold were considered CD62P-positive in comparison to the positive control.

Platelet activation state was expressed in relation to both isotype and positive control as percentage of gated platelet population above the respective CD62P positivity threshold.

Quantification of sGPV

The quantification of sGPV was performed as reported previously [6]. Briefly, 1 mL of the PC was centrifuged (Biofuge 13; He­raeus Sepatech, Hanau, Germany), first at 3,600 g at RT for 15 min and the supernatant again at 11,000 g at RT for 5 min (Biofuge 13). The supernatant was transferred to new tubes in 500-µL aliquots and stored at −70°C until sGPV quantification with a commercial kit (Asserachrom; Diagnostica Stago, Asnières sur Seine, France). For the measurement, samples were diluted 1:80–1:640 using phosphate buffer provided with the kit, and the amount of sGPV was expressed as pmol/109 platelets.

Isolation of EVs

A total of 17 mL of PC was used for EV isolation. To prevent platelet activation, Anticoagulant Citrate Dextrose Solution pH Eur Solution A (Terumo BCT, Lakewood, CO, USA) and Apyrase (Sigma-Aldrich) were added to the final concentrations of 4.25% v/v and 2 U/mL, respectively, and the PC was diluted 1:4 with phosphate-buffered saline (PBS) (Thermo Fisher, Waltham, MA, USA). The diluted PC was centrifuged at 650 g at RT for 7 min (Eppendorf centrifuge 5810R; Eppendorf, Hamburg, Germany) without brake, and the supernatant was centrifuged at 1,560 g at RT for 20 min (Eppendorf centrifuge 5810R). The residual platelet content of the supernatant was reduced to 1 × 106 platelets/mL, as confirmed with Coulter Cell Counter T-540 (Beckman Coulter). To extract the whole EV population from the PC, the supernatant was ultracentrifuged at 100,000 g at 4°C for 1 h (MLA-50 rotor, k-factor 92; Beckman Coulter). The supernatant was carefully decanted and remaining supernatant was removed with a pipette, after which the EV sample was resuspended into 200 µL of PBS and stored in Protein LoBind tubes (Eppendorf) at −70°C until analysis.

Quantification and Size Determination of Particles in EV Samples

The concentration and size distribution of particles in EV samples was determined using nanoparticle tracking analysis. The LM14C model used was equipped with a 70-mW violet (405 nm) laser (Malvern Instruments Ltd., Malvern, UK) and a sCMOS camera (Hamamatsu Photonics K.K., Hamamatsu, Japan). Data were captured using camera level 14, and three videos of 90 s were recorded, manually mixing the sample between measurements. EV samples from PAS-B PCs were diluted 1:1,000, 1:2,000, 1:5,000, and 1:10,000 with filtered (0.2 µm) PBS on d1, d2, d5, and d8 samples, respectively, and EV samples from PAS-E PCs were diluted 1:1,000, 1:2,000, 1:5,000, and 1:5,000–1:10,000 on d1, d2, d5, and d8 samples, respectively. Data analysis was performed with threshold 5 and gain 10. Data were recorded and analyzed with NanoSight software version 3.0 (Malvern Instruments Ltd.). The data were reported as EV concentration of the PC on the sampling day by calculating the particle content of the EV sample using the determined particle concentration and taking into account that the particles were isolated from a 17-mL sample, considered as a representative sample of PC.

Staining and Characterization of EV Samples on ImageStream®X Mark II

EV samples from PAS-E were labeled with Alexa Fluor 488 C5 maleimide (Invitrogen, Carlsbad, CA, USA) for 60 min at RT as described previously [28]. Excess maleimide was removed by using exosome resin spin columns (Invitrogen) which were prepared according to the manufacturer's instructions. Maleimide labeling without EVs was performed in a parallel fashion to confirm the dye retention by columns and to get “mock” controls for the experiments.

Different fluorescent stains for further characterization of EVs were used according to the manufacturer's instructions. Antibodies were: CD41a AF647 (clone HIP8; BioLegend, San Diego, CA, USA), CD45 PerCP-Cy5.5 (clone 2DI; BioLegend), CD63 BV510 (clone H5C6; BD BioScience, San Jose, CA, USA), and CD235a Pacific Blue (clone HI264; BioLegend). Apolipoprotein A contamination was surveyed with ApoA1 PerCP (BioSite, Täby, Sweden). Antibodies were incubated for 30 min at RT in the dark in PBS.

Maleimide 488 and fluorescent-positive EVs were detected using a 12-channel Amnis® ImageStream®X Mark II (EMD Millipore, Burlington, MA, USA) imaging flow cytometer. Samples were acquired at 60× magnification with low flow rate/high sensitivity. The integrated software INSPIRE® (EMD Millipore) was used for data collection. The instrument and INSPIRE software were set up as follows: excitation lasers 488, 642, and 785 and channel 01 (Ch01) and Ch09 (bright field), Ch06 (scattering channel), plus fluorescence channels Ch02, Ch05, Ch07, Ch08, and Ch011 were activated for signal detections.

At least 10,000 events for each sample were acquired. Positive events for maleimide 488 were gated based on the intensity values and used for further analysis. Single-color controls were used for compensation and unlabeled EVs were used to determine the autofluorescence. Buffers with and without antibody/maleimide 488 molecules were used to determine the background noise. Compensated data files were analyzed using image-based algorithms available in the IDEAS® statistical analysis software package (version 6.2.188.0).

Statistical Analysis

Kruskal-Wallis test together with Dunn's multiple comparison test to take into account the effect of multiple testing was used to determine the significance of the results within PASs, and p values <0.05 were considered significant. To determine the significance between PASs on d5 sample Mann-Whitney test together with Bonferroni correction was used. Spearman correlation coefficient and related p value together with R2 value of the standard curve was used to determine the correlation between the different platelet activation parameters. All statistical analyses were performed using GraphPad Prism v. 6.07 (GraphPad Software, Inc., La Jolla, CA, USA).

Results

CD62P Exposure of Platelets

A statistically significant, time-dependent increase in CD62P exposure was observed during the 8-day storage period. The exposure of CD62P in the PAS-B stored platelets increased from 8.5 to 78% (p = 0.0226 and p = 0.0002 on d5 and d8, respectively) when compared to the positive, thrombin-activated control (Fig. 1A). In PAS-E platelets, the average CD62P exposure increased from 3.6 to 71% (p = 0.0028 on d8) compared to the positive control (Fig. 1A).

Fig. 1.

Fig. 1

Open in a new tab

Quality control markers used for the evaluation of platelet activation during storage of PCs with PAS-B and PAS-E. A, B Time-dependent changes in the CD62P exposure of platelets when compared to a positive (A) or an isotype control (B). C sGPV production of platelets. D, E Concentration (D) and size distribution (E) of particles in the EV samples isolated from PCs. Statistical difference within a given PAS is indicated with black and grey asterisks for PAS-B and PAS-E PCs, respectively. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001 compared to d1 using Kruskal-Wallis test with Dunn's multiple comparison. Statistical difference between PAS-B and PAS-E PCs on d5 (Bonferroni-adjusted p < 0.05 using Mann-Whitney test with Bonferroni correction is indicated with a red asterisk). Bars represent mean with standard deviation in A–D; columns present mean and bars standard deviation in E. Data were acquired in three independent experimental settings, n = 4–5 (PAS-B in all figures, PAS-E in A and B) or n = 10 (PAS-E in C–E). d, day; EV, extracellular vesicle; PAS, platelet additive solution; PCs, platelet concentrates; sGPV, soluble glycoprotein V.

When CD62P exposure was determined by comparison to the isotype control, the CD62P exposure of PAS-B platelets increased from 36 to 70% (p = 0.0177 and p = 0.0005 on d5 and d8, respectively) and from 24 to 53% (p = 0.0117 on d8) in the PAS-E platelets (Fig. 1B).

Regardless of the CD62P exposure determination method, on d5, the last day when the PC can be transfused to patients, platelets in PAS-B PCs exposed more CD62P than platelets in PAS-E PCs (Bonferroni-adjusted p = 0.0016 for both) (Fig. 1A, B).

sGPV Content in PCs

The sGPV content of PAS-B PCs increased from the average of 2.1 to 24.5 pmol/109 platelets during storage (p = 0.0225 and p = 0.0002 on d5 and d8, respectively) (Fig. 1C). Also in the PAS-E PCs, the increase in sGPV was statistically significant, but more subtle, as the sGPV concentration increased from 1.8 to 12.1 pmol/109 platelets (p = 0.0224 and p = 0.0002 on d5 and d8, respectively) during the 8-day storage (Fig. 1C). The sGPV content of PCs was significantly higher in PAS-B PCs than PAS-E PCs on d5 (Bonferroni-adjusted p = 0.0158) (Fig. 1C).

Concentration and Size Distribution of Particles in EV Samples

The particle concentration in EV samples of PAS-B PCs significantly increased during the 8-day storage period from 1.1 × 1010 particles/mL on d1 to 1.3 × 1011 particles/mL on d8 (p = 0.0292 and p = 0.0021 on d5 and d8, respectively) and in the PAS-E PCs from 7.9 × 109 particles/mL on d1 to 3.7 × 1010 particles/mL on d8 (p = 0.0019 and p < 0.0001 on d5 and d8, respectively) (Fig. 1D). Both the PAS-B and PAS-E PCs initially had similar particle concentration in EV samples, but from d2 onwards the particle concentration in EV samples of PAS-B PCs was higher compared to PAS-E PCs (Bonferroni-adjusted p = 0.016 on d5) (Fig. 1D).

The size distribution of particles in EV samples changed significantly during aging only in the PAS-B PCs (Fig. 1E). Initially, 61% of the particles were <100 nm in diameter, but after 8 days of storage, the percentage of particles <100 nm had decreased to 27% (p = 0.0070). Consequently, the percentage of particles with a diameter of 101–200 nm was initially 31%, which increased to 56% on d8 (p = 0.0484). In contrast to PAS-B, no significant alteration in the size distribution of particles in EV samples was observed in PAS-E PCs (Fig. 1E).

Characteristics of EVs Isolated from PCs

From all the maleimide-positive particles in the EV samples, the majority expressed CD41, indicating that the EVs isolated from the PCs are mainly derived from platelets. Also, the EV marker CD63 was abundantly present, and the number of CD41- and CD63-positive particles increased during the 7-day storage. Besides the platelet-derived particles, minute amounts of leukocyte- and erythrocyte-derived particles and ApoA1 were detectable in the EV samples (Table 1).

Table 1.

Frequency of surface markers identified from maleimidepositive particles of extracellular vesicle samples isolated from platelet concentrate on days 1 and 8

Day 1 Day 8
ApoA1 0.8% 1.4%
CD41 68.7% 88.0%
CD45 0.6% 2.6%
CD63 29.0% 57.7%
CD235a 5.1% 2.1%
Open in a new tab

Correlation of EV Sample Particle Concentration with CD62P Exposure and sGPV Content

A strong positive correlation was observed between the particle concentration of EV samples and the sGPV content of PCs (R2 = 0.7639, Spearman r = 0.7906 with p < 0.0001) (Fig. 2A), and a very strong positive correlation was observed between the particle concentration of EV samples and the CD62P exposure of platelets (R2 = 0.6626, Spearman r = 0.8269 with p < 0.0001) (Fig. 2B) and between the sGPV content of PCs and the CD62P exposure of platelets (R2 = 0.8816, Spearman r = 0.9253 with p < 0.0001) (Fig. 2C).

Fig. 2.

Fig. 2

Open in a new tab

Correlation analysis of the three different markers for platelet activation. A Particle concentration in the EV samples and sGPV production of platelets. B Particle concentration in the EV samples and CD62P exposure of platelets when compared to a positive control. C sGPV production and CD62P exposure of platelets when compared to a positive control. The figure was compiled using data from both PAS-B and PAS-E PCs acquired from three independent experimental settings. EV, extracellular vesicle; PAS, platelet additive solution; PCs, platelet concentrates; sGPV, soluble glycoprotein V.

Discussion

For a few decades, PCs have been prepared with PAS together with some plasma. Initially only plasma-containing PCs were favored due to better functionality (estimated by corrected count increments and bleeding) compared to only PAS-containing PCs [29]. The disadvantages of plasma-containing PCs include increased incidence of adverse transfusion-related reactions, mainly allergic reactions, but possibly also transfusion-related acute lung injury and ABO-mismatched hemolysis [26]. Currently, approximately 30% of the volume of PAS-containing PCs still contains plasma to maintain platelet functionality [30], but the development of PASs has resulted in a notable improvement of platelet quality and functionality, leading to experimentations with decreased content of plasma in PCs with PASs [31]. At the moment, PAS-E is considered to be the best PAS developed, having similar platelet functionality to the PCs with plasma in terms of corrected count increment [29], and it has even been hypothesized whether with the addition of further components such as glucose [30], the advantage of plasma could be surpassed in favor of the PAS-only PCs. The driving force behind reducing the plasma content in PCs is the potential decline in the incidence of adverse transfusion-related reactions. Additionally, the leftover plasma could be utilized for fractionation to produce other transfusable products [26].

All common PASs contain NaCl and acetate [26]. NaCl is added in varying amounts to adjust PC osmolarity, and acetate is added for two reasons: firstly, to provide an alternative energy source in addition to glucose, as it reduces lactate production and consequently influences the pH of the PC; secondly, during the enzymatic processing of acetate, carbon dioxide is formed, which further reacts with water to form bicarbonate providing increased buffer capacity to PCs [32]. Most PASs also contain citrate as an anticoagulant, which provides yet another energy source and added buffer capacity [26, 33]. Furthermore, PAS-E contains phosphate, potas­sium, and magnesium, whereas PAS-B does not [26]. Phosphate is added to PCs for improved buffer capacity and to stimulate platelet glycolysis [34]. PASs with different compositions have been extensively tested, and addition of potassium and magnesium has been connected to decreased cytokine [35] and lactate [36, 37] production as well as decreased CD62P [37, 38] and phosphatidylserine [38] exposure of platelets. Our results on time-dependent platelet activation are in line with these previous findings as based on the three assessed platelet activation markers (CD62P exposure of platelets together with sGPV and EV content of PCs), platelets in PAS-B PCs were more activated than PAS-E PC platelets on d5, the last day when PC could be transfused to patients. A significant increase in the activation state of the PAS-B platelets was detected with all three activation markers at d5 sampling. For PAS-E PCs a significant increase in sGPV content and EV particle concentration was observed on d5 as well, whereas a significant increase in CD62P exposure was observed only on d8. Although it is unclear how altered PAS composition affects platelet activation, the mechanism might involve membrane potassium movement and permeability [39]. Similarly to CD62P and sGPV, a time-dependent increase in EV concentration in PCs was observed, in line with previously published results [40, 41]. Based on the current data, EV concentration correlated well with the sGPV content and the CD62P exposure of platelets, indicating that EV concentration could be used to indicate platelet activation in PCs.

As shown previously, determination of both the CD62P exposure of platelets and the sGPV content in PCs were sensitive and reproducible methods to detect platelet activation [6, 27]. As an additional advantage of these methods, the maximum extent for both parameters can be determined, which helps to estimate the platelet activation state by giving either a relative or an absolute [42] boundary value (for CD62P and sGPV, respectively). Contrary to CD62P and sGPV measurements, it is not possible to generate an accurate control for a maximum EV production as different agonists produce varying amounts of platelet-derived EVs [23]. Although current PC manufacturing processes ensure minimal cell contamination, EVs from erythrocytes, platelets, and leukocytes are present in PCs already due to the plasma component of the PCs, as shown in this study and by others [14]. It is difficult to determine whether the platelet-derived EVs are produced due to aging-related platelet activation, as a result of interaction of buffy coat components during storage, or even due to an apoptosis-like process [43]. The interaction of buffy coat EVs and platelets might explain the platelet activation to some degree [44] and consequently the high variation in the particle concentration of EV samples from PCs especially seen in the d8 samples. Additionally, considering the variance in donors [45, 46], current EV sample preparation methods [47], and the lack of standardized and accurate EV quantification methods [48, 49, 50], it must be stressed that although EV concentration seems to be a potential marker of platelet activation, significant development and standardization will be needed before the current methods can be replaced to determine platelet activation state in PCs. The authors would like to underline that EVs could still be used as a complementary platelet activation marker to CD62P and sGPV.

In addition to EV concentration being a marker for platelet activation similarly to CD62P or sGPV, EVs could also provide qualitative information of PC aging and possibly even functionality. Besides influencing the size [23, 51] and the molecular cargo [23, 24, 52] of produced EVs, platelet activating conditions have been shown to affect the subsequent function of produced EVs [53], and future studies could concentrate on the qualitative information provided by PC-derived EVs. In the current study, we observed a time-dependent increase in the size distribution of particles in the EV samples from platelets stored in PAS-B. Since platelet activation was influenced by PAS composition, it may also have an influence on the produced EVs. Another possible explanation for the altered size could be an artefactual clumping of EVs, which has been shown previously [54]. However, the effect was only subtle compared to the size of for instance EV doublets and without a corresponding decline in the particle concentration, so formation of stickier EVs is unlikely to explain the current results. A change in the EV population in the PC, reflected by size change, could also have functional effects upon transfusion [55]. To understand the potential effects of EVs in storage lesion, adverse transfusion-related reactions, or immunomodulatory functions in general, it will be necessary to carefully examine the molecular composition of EV populations by lipidomics, proteomics, or metabolomics [56, 57, 58]. Moreover, characterizing the PC-derived EVs could also be a step towards personalized transfusion treatments, where patients could be targeted to receive PCs that would best suit their needs [45, 46], e.g., PCs that have more procoagulant potency in the case of severe bleeding. By doing this, the utilization of PCs could be optimized, leading to less wasted PCs and hopefully to transfusions with less adverse transfusion-related reactions.

In conclusion, EVs may be a useful tool in the quality control of PCs in the future, and the molecular characterization of EVs could provide more information about the state and usability of the PCs, ultimately benefitting patients receiving transfusions.

Statement of Ethics

The authors have no ethical conflicts to disclose.

Disclosure Statement

The authors have no conflicts of interest to declare.

Funding Sources

Part of this work was funded by SalWe Research Program Personalized Diagnostics and Care (GET IT DONE); Tekes – the Finnish Funding Agency for Technology and Innovation grant Dno 3986/31/2013 (S. Valkonen, P.R.-M. Siljander, S. Laitinen); the Academy of Finland program grant No. 287089 (S. Valkonen, P.R.-M. Siljander); and Magnus Ehrnrooth Foundation (S. Valkonen, P.R.-M. Siljander).

Acknowledgments

The authors would like to thank Lotta Sankkila for assistance in sample preparation and Reija Soukka for assistance in the sampling procedure. Mikko Arvas is also thanked for guidance and support in statistical analyses.

References

  • 1.Harker LA, Roskos LK, Marzec UM, Carter RA, Cherry JK, Sundell B, et al. Effects of megakaryocyte growth and development factor on platelet production, platelet life span, and platelet function in healthy human volunteers. Blood. 2000 Apr;95((8)):2514–22. [PubMed] [Google Scholar]
  • 2.Slichter SJ, Bolgiano D, Corson J, Jones MK, Christoffel T, Bailey SL, et al. Extended storage of buffy coat platelet concentrates in plasma or a platelet additive solution. Transfusion. 2014 Sep;54((9)):2283–91. doi: 10.1111/trf.12634. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3.Braine HG, Kickler TS, Charache P, Ness PM, Davis J, Reichart C, et al. Bacterial sepsis secondary to platelet transfusion: an adverse effect of extended storage at room temperature. Transfusion. 1986 Jul-Aug;26((4)):391–3. doi: 10.1046/j.1537-2995.1986.26486262752.x. [DOI] [PubMed] [Google Scholar]
  • 4.Mohanty D. Current concepts in platelet transfusion. Asian J Transfus Sci. 2009 Jan;3((1)):18–21. doi: 10.4103/0973-6247.45257. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Magron A, Laugier J, Provost P, Boilard E. Pathogen reduction technologies: the pros and cons for platelet transfusion. Platelets. 2017;•••:1–7. doi: 10.1080/09537104.2017.1306046. [DOI] [PubMed] [Google Scholar]
  • 6.Kiminkinen LK, Krusius T, Javela KM. Evaluation of soluble glycoprotein V as an in vitro quality marker for platelet concentrates: a correlation study between in vitro platelet quality markers and the effect of storage medium. Vox Sang. 2016 Aug;111((2)):120–6. doi: 10.1111/vox.12402. [DOI] [PubMed] [Google Scholar]
  • 7.Herter JM, Rossaint J, Zarbock A. Platelets in inflammation and immunity. J Thromb Haemost. 2014 Nov;12((11)):1764–75. doi: 10.1111/jth.12730. [DOI] [PubMed] [Google Scholar]
  • 8.Phillips DR, Agin PP. Platelet plasma membrane glycoproteins. Identification of a proteolytic substrate for thrombin. Biochem Biophys Res Commun. 1977 Apr;75((4)):940–7. doi: 10.1016/0006-291x(77)91473-5. [DOI] [PubMed] [Google Scholar]
  • 9.Canobbio I, Balduini C, Torti M. Signalling through the platelet glycoprotein Ib-V-IX complex. Cell Signal. 2004 Dec;16((12)):1329–44. doi: 10.1016/j.cellsig.2004.05.008. [DOI] [PubMed] [Google Scholar]
  • 10.Dong JF, Sae-Tung G, López JA. Role of glycoprotein V in the formation of the platelet high-affinity thrombin-binding site. Blood. 1997 Jun;89((12)):4355–63. [PubMed] [Google Scholar]
  • 11.Moog S, Mangin P, Lenain N, Strassel C, Ravanat C, Schuhler S, et al. Platelet glycoprotein V binds to collagen and participates in platelet adhesion and aggregation. Blood. 2001 Aug;98((4)):1038–46. doi: 10.1182/blood.v98.4.1038. [DOI] [PubMed] [Google Scholar]
  • 12.Garraud O, Cognasse F, Tissot JD, Chavarin P, Laperche S, Morel P, et al. Improving platelet transfusion safety: biomedical and technical considerations. Blood Transfus. 2016 Mar;14((2)):109–22. doi: 10.2450/2015.0042-15. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13.Kreuger AL, Caram-Deelder C, Jacobse J, Kerkhoffs JL, van der Bom JG, Middelburg RA. Effect of storage time of platelet products on clinical outcomes after transfusion: a systematic review and meta-analyses. Vox Sang. 2017 May;112((4)):291–300. doi: 10.1111/vox.12494. [DOI] [PubMed] [Google Scholar]
  • 14.Nollet KE, Saito S, Ono T, Ngoma A, Ohto H. Microparticle formation in apheresis platelets is not affected by three leukoreduction filters. Transfusion. 2013 Oct;53((10)):2293–8. doi: 10.1111/trf.12088. [DOI] [PubMed] [Google Scholar]
  • 15.Chargaff E, West R. The biological significance of the thromboplastic protein of blood. J Biol Chem. 1946 Nov;166((1)):189–97. [PubMed] [Google Scholar]
  • 16.Wolf P. The nature and significance of platelet products in human plasma. Br J Haematol. 1967 May;13((3)):269–88. doi: 10.1111/j.1365-2141.1967.tb08741.x. [DOI] [PubMed] [Google Scholar]
  • 17.Lentz BR. Exposure of platelet membrane phosphatidylserine regulates blood coagulation. Prog Lipid Res. 2003 Sep;42((5)):423–38. doi: 10.1016/s0163-7827(03)00025-0. [DOI] [PubMed] [Google Scholar]
  • 18.van der Pol E, Harrison P. From platelet dust to gold dust: physiological importance and detection of platelet microvesicles. Platelets. 2017 May;28((3)):211–3. doi: 10.1080/09537104.2017.1282781. [DOI] [PubMed] [Google Scholar]
  • 19.Aatonen M, Grönholm M, Siljander PR. Platelet-derived microvesicles: multitalented participants in intercellular communication. Semin Thromb Hemost. 2012 Feb;38((1)):102–13. doi: 10.1055/s-0031-1300956. [DOI] [PubMed] [Google Scholar]
  • 20.Yáñez-Mó M, Siljander PR, Andreu Z, Zavec AB, Borràs FE, Buzas EI, et al. Biological properties of extracellular vesicles and their physiological functions. J Extracell Vesicles. 2015 May;4((1)):27066. doi: 10.3402/jev.v4.27066. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21.van der Pol E, Böing AN, Harrison P, Sturk A, Nieuwland R. Classification, functions, and clinical relevance of extracellular vesicles. Pharmacol Rev. 2012 Jul;64((3)):676–705. doi: 10.1124/pr.112.005983. [DOI] [PubMed] [Google Scholar]
  • 22.Boudreau LH, Marcoux G, Boilard E. Platelet microparticles in transfusion. ISBT Sci Ser. 2015;10(S1):305–8. [Google Scholar]
  • 23.Aatonen MT, Öhman T, Nyman TA, Laitinen S, Grönholm M, Siljander PR. Isolation and characterization of platelet-derived extracellular vesicles. J Extracell Vesicles. 2014;3 doi: 10.3402/jev.v3.24692. eCollection 2014. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24.Milioli M, Ibáñez-Vea M, Sidoli S, Palmisano G, Careri M, Larsen MR. Quantitative proteomics analysis of platelet-derived microparticles reveals distinct protein signatures when stimulated by different physiological agonists. J Proteomics. 2015 May;121:56–66. doi: 10.1016/j.jprot.2015.03.013. [DOI] [PubMed] [Google Scholar]
  • 25.Ashford P, Gulliksson H, Georgsen J, Distler P. Standard terminology for platelet additive solutions. Vox Sang. 2010 May;98((4)):577–8. doi: 10.1111/j.1423-0410.2009.01298.x. [DOI] [PubMed] [Google Scholar]
  • 26.Alhumaidan H, Sweeney J. Current status of additive solutions for platelets. J Clin Apher. 2012;27((2)):93–8. doi: 10.1002/jca.21207. [DOI] [PubMed] [Google Scholar]
  • 27.Curvers J, de Wildt-Eggen J, Heeremans J, Scharenberg J, de Korte D, van der Meer PF. Flow cytometric measurement of CD62P (P-selectin) expression on platelets: a multicenter optimization and standardization effort. Transfusion. 2008 Jul;48((7)):1439–46. doi: 10.1111/j.1537-2995.2008.01738.x. [DOI] [PubMed] [Google Scholar]
  • 28.Roberts-Dalton HD, Cocks A, Falcon-Perez JM, Sayers EJ, Webber JP, Watson P, et al. Fluorescence labelling of extracellular vesicles using a novel thiol-based strategy for quantitative analysis of cellular delivery and intracellular traffic. Nanoscale. 2017 Sep;9((36)):13693–706. doi: 10.1039/c7nr04128d. [DOI] [PubMed] [Google Scholar]
  • 29.van der Meer PF, de Korte D. Platelet Additive Solutions: A Review of the Latest Developments and Their Clinical Implications. Transfus Med Hemother. 2018 Apr;45((2)):98–102. doi: 10.1159/000487513. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30.van der Meer PF. PAS or plasma for storage of platelets? A concise review. Transfus Med. 2016 Oct;26((5)):339–42. doi: 10.1111/tme.12325. [DOI] [PubMed] [Google Scholar]
  • 31.Sandgren P, Mayaudon V, Payrat JM, Sjödin A, Gulliksson H. Storage of buffy-coat-derived platelets in additive solutions: in vitro effects on platelets stored in reformulated PAS supplied by a 20% plasma carry-over. Vox Sang. 2010 Apr;98((3 Pt 2)):415–22. doi: 10.1111/j.1423-0410.2009.01255.x. [DOI] [PubMed] [Google Scholar]
  • 32.Shimizu T, Murphy S. Roles of acetate and phosphate in the successful storage of platelet concentrates prepared with an acetate-containing additive solution. Transfusion. 1993 Apr;33((4)):304–10. doi: 10.1046/j.1537-2995.1993.33493242637.x. [DOI] [PubMed] [Google Scholar]
  • 33.Ringwald J, Zimmermann R, Eckstein R. The new generation of platelet additive solution for storage at 22 ° C: development and current experience. Transfus Med Rev. 2006 Apr;20((2)):158–64. doi: 10.1016/j.tmrv.2005.11.003. [DOI] [PubMed] [Google Scholar]
  • 34.Gulliksson H, Larsson S, Kumlien G, Shanwell A. Storage of platelets in additive solutions: effects of phosphate. Vox Sang. 2000;78((3)):176–84. doi: 10.1159/000031177. [DOI] [PubMed] [Google Scholar]
  • 35.Shanwell A, Falker C, Gulliksson H. Storage of platelets in additive solutions: the effects of magnesium and potassium on the release of RANTES, β-thromboglobulin, platelet factor 4 and interleukin-7, during storage. Vox Sang. 2003 Oct;85((3)):206–12. doi: 10.1046/j.1423-0410.2003.00359.x. [DOI] [PubMed] [Google Scholar]
  • 36.Gulliksson H, AuBuchon JP, Vesterinen M, Sandgren P, Larsson S, Pickard CA, et al. Biomedical Excellence for Safer Transfusion Working Party of the International Society of Blood Transfusion Storage of platelets in additive solutions: a pilot in vitro study of the effects of potassium and magnesium. Vox Sang. 2002 Apr;82((3)):131–6. doi: 10.1046/j.1423-0410.2002.drfgv158.x. [DOI] [PubMed] [Google Scholar]
  • 37.de Wildt-Eggen J, Schrijver JG, Bins M, Gulliksson H. Storage of platelets in additive solutions: effects of magnesium and/or potassium. Transfusion. 2002 Jan;42((1)):76–80. doi: 10.1046/j.1537-2995.2002.00012.x. [DOI] [PubMed] [Google Scholar]
  • 38.van der Meer PF, Kerkhoffs JL, Curvers J, Scharenberg J, de Korte D, Brand A, et al. In vitro comparison of platelet storage in plasma and in four platelet additive solutions, and the effect of pathogen reduction: a proposal for an in vitro rating system. Vox Sang. 2010 May;98((4)):517–24. doi: 10.1111/j.1423-0410.2009.01283.x. [DOI] [PubMed] [Google Scholar]
  • 39.Weis-Fogh US. The effect of citrate, calcium, and magnesium ions on the potassium movement across the human platelet membrane. Transfusion. 1985 Jul-Aug;25((4)):339–42. doi: 10.1046/j.1537-2995.1985.25485273813.x. [DOI] [PubMed] [Google Scholar]
  • 40.Black A, Pienimaeki-Roemer A, Kenyon O, Orsó E, Schmitz G. Platelet-derived extracellular vesicles in plateletpheresis concentrates as a quality control approach. Transfusion. 2015 Sep;55((9)):2184–96. doi: 10.1111/trf.13128. [DOI] [PubMed] [Google Scholar]
  • 41.Black A, Orsó E, Kelsch R, Pereira M, Kamhieh-Milz J, Salama A, et al. Analysis of platelet-derived extracellular vesicles in plateletpheresis concentrates: a multicenter study. Transfusion. 2017 Jun;57((6)):1459–69. doi: 10.1111/trf.14109. [DOI] [PubMed] [Google Scholar]
  • 42.Azorsa DO, Moog S, Ravanat C, Schuhler S, Folléa G, Cazenave JP, et al. Measurement of GPV released by activated platelets using a sensitive immunocapture ELISA—its use to follow platelet storage in transfusion. Thromb Haemost. 1999 Jan;81((1)):131–8. [PubMed] [Google Scholar]
  • 43.Nieuwland R, van der Pol E, Gardiner C, Sturk A. Platelets. Elsevier; 2013. Platelet-Derived Microparticles; pp. pp. 453–67. [Google Scholar]
  • 44.Kohli S, Ranjan S, Hoffmann J, Kashif M, Daniel EA, Al-Dabet MM, et al. Maternal extracellular vesicles and platelets promote preeclampsia via inflammasome activation in trophoblasts. Blood. 2016 Oct;128((17)):2153–64. doi: 10.1182/blood-2016-03-705434. [DOI] [PubMed] [Google Scholar]
  • 45.Maurer-Spurej E, Larsen R, Labrie A, Heaton A, Chipperfield K. Microparticle content of platelet concentrates is predicted by donor microparticles and is altered by production methods and stress. Transfus Apheresis Sci. 2016 Aug;55((1)):35–43. doi: 10.1016/j.transci.2016.07.010. [DOI] [PubMed] [Google Scholar]
  • 46.Maurer-Spurej E, Chipperfield K. Could Microparticles Be the Universal Quality Indicator for Platelet Viability and Function? J Blood Transfus. 2016;2016:6140239. doi: 10.1155/2016/6140239. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 47.Coumans FA, Brisson AR, Buzas EI, Dignat-George F, Drees EE, El-Andaloussi S, et al. Methodological Guidelines to Study Extracellular Vesicles. Circ Res. 2017 May;120((10)):1632–48. doi: 10.1161/CIRCRESAHA.117.309417. [DOI] [PubMed] [Google Scholar]
  • 48.Varga Z, Yuana Y, Grootemaat AE, van der Pol E, Gollwitzer C, Krumrey M, et al. Towards traceable size determination of extracellular vesicles. J Extracell Vesicles. 2014;3 doi: 10.3402/jev.v3.23298. eCollection 2014. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 49.Valkonen S, van der Pol E, Böing A, Yuana Y, Yliperttula M, Nieuwland R, et al. Biological reference materials for extracellular vesicle studies. Eur J Pharm Sci. 2017 Feb;98:4–16. doi: 10.1016/j.ejps.2016.09.008. [DOI] [PubMed] [Google Scholar]
  • 50.Nicolet A, Meli F, van der Pol E, Yuana Y, Gollwitzer C, Krumrey M, et al. Inter-laboratory comparison on the size and stability of monodisperse and bimodal synthetic reference particles for standardization of extracellular vesicle measurements. Meas Sci Technol. 2016;27((3)):35701. [Google Scholar]
  • 51.Ponomareva AA, Nevzorova TA, Mordakhanova ER, Andrianova IA, Rauova L, Litvinov RI, et al. Intracellular origin and ultrastructure of platelet-derived microparticles. J Thromb Haemost. 2017 Aug;15((8)):1655–67. doi: 10.1111/jth.13745. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 52.De Paoli SH, Tegegn TZ, Elhelu OK, Strader MB, Patel M, Diduch LL, et al. Dissecting the biochemical architecture and morphological release pathways of the human platelet extracellular vesiculome. Cell Mol Life Sci. 2018 Oct;75((20)):3781–801. doi: 10.1007/s00018-018-2771-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 53.Vasina E, Heemskerk JW, Weber C, Koenen RR. Platelets and platelet-derived microparticles in vascular inflammatory disease. Inflamm Allergy Drug Targets. 2010 Dec;9((5)):346–54. doi: 10.2174/187152810793938008. [DOI] [PubMed] [Google Scholar]
  • 54.Yuana Y, Böing AN, Grootemaat AE, van der Pol E, Hau CM, Cizmar P, et al. Handling and storage of human body fluids for analysis of extracellular vesicles. J Extracell Vesicles. 2015 Nov;4((1)):29260. doi: 10.3402/jev.v4.29260. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 55.Redman CW, Tannetta DS, Dragovic RA, Gardiner C, Southcombe JH, Collett GP, et al. Review: does size matter? Placental debris and the pathophysiology of pre-eclampsia. Placenta. 2012 Feb;33(Suppl):S48–54. doi: 10.1016/j.placenta.2011.12.006. [DOI] [PubMed] [Google Scholar]
  • 56.Pienimaeki-Roemer A, Kuhlmann K, Böttcher A, Konovalova T, Black A, Orsó E, et al. Lipidomic and proteomic characterization of platelet extracellular vesicle subfractions from senescent platelets. Transfusion. 2015 Mar;55((3)):507–21. doi: 10.1111/trf.12874. [DOI] [PubMed] [Google Scholar]
  • 57.Altadill T, Campoy I, Lanau L, Gill K, Rigau M, Gil-Moreno A, et al. Enabling Metabolomics Based Biomarker Discovery Studies Using Molecular Phenotyping of Exosome-Like Vesicles. PLoS One. 2016 Mar;11((3)):e0151339. doi: 10.1371/journal.pone.0151339. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 58.Puhka M, Takatalo M, Nordberg ME, Valkonen S, Nandania J, Aatonen M, et al. Metabolomic Profiling of Extracellular Vesicles and Alternative Normalization Methods Reveal Enriched Metabolites and Strategies to Study Prostate Cancer-Related Changes. Theranostics. 2017 Aug;7((16)):3824–41. doi: 10.7150/thno.19890. [DOI] [PMC free article] [PubMed] [Google Scholar]

Articles from Transfusion Medicine and Hemotherapy are provided here courtesy of Karger Publishers

RESOURCES