|Year : 2020 | Volume
| Issue : 2 | Page : 51-58
Impact of storage, leukofiltration, and ascorbic acid fortification on red cell-derived microparticles in stored packed red blood cells: A flow cytometric and procoagulant study
Raghda Fouda1, Azza A Aboul Enein1, Nermeen A El-Desoukey1, Randa M Abo Elfetouh2, Ahmed M. A Abdel Hafez1
1 Department of Clinical and Chemical Pathology, Faculty of Medicine, Cairo University, Cairo, Egypt
2 Department of Clinical and Chemical Pathology, National Cancer Institute, Cairo University, Cairo, Egypt
|Date of Submission||11-Nov-2019|
|Date of Decision||20-Feb-2020|
|Date of Acceptance||23-Feb-2020|
|Date of Web Publication||28-Jul-2020|
Dr. Raghda Fouda
2621 Verano, Irvine, CA 92617
Source of Support: None, Conflict of Interest: None
INTRODUCTION: During blood storage, red blood cells (RBCs) undergo changes termed “storage lesion” including shape modification and microparticles (MPs) production, as well as oxidative injury to lipids and proteins.
OBJECTIVE: We studied the impact of storage on RBCs by quantification of red cell MPs and their potential procoagulant activity (PCA). The antioxidant effect of ascorbic acid (AA) was assessed, as a proposed additive, to improve stored RBC quality.
METHODS: Forty randomly selected packed red cell units (filtered and nonfiltered) were sampled weekly over 35 days of storage. Flow cytometric and coagulation tests were performed on MPs isolated from erythrocyte concentrates. RBCs were either in citrate phosphate dextrose adenine-1 (CPDA1) only (standard storage) or CPDA1 with AA (Vitamin C) in three concentrations.
RESULTS: Red cell MPs progressively increased over 35 days of storage. Filtration efficiently reduced the production of MPs (P < 0.001). PCA increased significantly on storage assessed by the shortened coagulation time (P < 0.001). A strong inverse correlation between coagulation time and Annexin V positive MPs was found, (r = −0.723, P < 0.001), and the dual-positive population. This highlights the strong correlation between the functional assay (PCA) and the direct quantitative assay (flow cytometry). The red cells fortified with AA demonstrated a reduction in the accumulation of RBC MPs by almost 50% compared to standard storage in CPDA-1 alone in an evident dose-dependent manner.
CONCLUSION: We concluded that MPs increase on storage with increased PCA. AA reduced MPs production which can be possible additive for further research in the future
Keywords: Ascorbic acid, flow cytometry, microparticles, storage lesions
|How to cite this article:|
Fouda R, Aboul Enein AA, El-Desoukey NA, Abo Elfetouh RM, Abdel Hafez AM. Impact of storage, leukofiltration, and ascorbic acid fortification on red cell-derived microparticles in stored packed red blood cells: A flow cytometric and procoagulant study. J Appl Hematol 2020;11:51-8
|How to cite this URL:|
Fouda R, Aboul Enein AA, El-Desoukey NA, Abo Elfetouh RM, Abdel Hafez AM. Impact of storage, leukofiltration, and ascorbic acid fortification on red cell-derived microparticles in stored packed red blood cells: A flow cytometric and procoagulant study. J Appl Hematol [serial online] 2020 [cited 2020 Aug 12];11:51-8. Available from: http://www.jahjournal.org/text.asp?2020/11/2/51/290964
| Introduction|| |
Over years, storage techniques have been under continuous development allowing better conservation of the integrity of red blood cells (RBCs). Questions regarding the safety of stored RBCs are as old as the systems for storing them. Citrate phosphate dextrose adenine-1 (CPDA1) is one of the commonly used anticoagulants in developing countries that preserve RBCs up to 35 days. On the other hand, more developed countries use the current generation of additive solutions (ASs), e.g., AS-1, AS-3, and AS-5 in the United States, and saline, adenine, glucose, and mannitol (SAGM) in Europe which extended the storage to 42 days.
The storage of blood under blood banking conditions causes biochemical and biomechanical changes (storage lesions), which in turn affects optimal functioning and survival. Storage lesions include accumulation of free hemoglobin, lipids, microparticles (MPs), and a pH reduction. In addition, RBCs lose the ability to cope with ongoing oxidative stress and production of reactive oxygen species throughout storage, which subsequently leads to oxidation of RBC lipids and proteins. Such alterations negatively affect red cell functions.
Methods to improve the quality of RBCs during storage are currently investigated. Recently, antioxidants have been suggested for use in stored RBC units as a result of evidence of oxidative damage to protein and lipid components of RBCs. Modern storage ASs generally do not contain ingredients specifically targeted at the inhibition and/or detoxification of reactive oxidative species.
During storage, the donor leukocytes release cytokines and lipid factors, e.g., tumor necrosis factor, interleukin (IL)-1, and IL-6. The cytokine content reflects the initial amounts of leukocytes and increases with prolonged duration. Cytokines are capable of inducing a systemic inflammatory response and transfusion reactions in the recipient. The increase in inflammatory soluble markers is reduced by leukocyte reduction. Leukoreduction may significantly decrease febrile nonhemolytic transfusion reactions and may decrease cardiopulmonary transfusion reactions.
The accumulation of MPs has been recognized during red cell storage. MPs are identified as membrane-derived vesicles which are smaller than 1 μ. These particles are generated by outward budding of the plasma membrane. Their release is initiated upon cellular activation or apoptosis. MPs contain numerous components from their cell of origin, including cytoplasmic and membrane proteins and also nucleic acids. These elements can be delivered to other cells by various mechanisms. MPs participate in many biological activities, such as inflammation, angiogenesis, and transfer of surface proteins.
Many of these vesicles expose phosphatidylserine (PS), negatively charged phospholipid on their outer surface. As they bear negatively charged membranes, red cell-derived MPs (RMPs) could be involved in thrombin generation through participating in tenase and prothrombinase complex formation. It is important to establish new approaches for monitoring and precise quantification of the release of MPs in RBC units and to evaluate their potential impact on blood transfusion.
Many reports have shown that the microvesicle (MV) count in RBC concentrates gradually increases over time and suggests that MPs could be responsible for the adverse clinical outcome of transfusions., However, CPDA1 was not the commonly used storage medium in these studies. Further studies to fill this gap in the literature and correlate the results with previous studies are needed. We aimed at monitoring the production of RBCs' MPs in packed RBCs using flow cytometry. We performed coagulation tests to correlate the MPs production with procoagulant activity. We also investigated the impact of leukofiltration, which is not routinely performed in our institute, on MPs' production and procoagulant activity (PCA). The antioxidant effect of ascorbic acid (AA) was tested as a proposed additive to improve stored RBCs quality.
| Material and Methods|| |
Forty (n = 40) randomly selected packed RBC units were obtained from the Central Blood Bank of Cairo University Hospital from January to September 2017. Twenty units (n = 20) were nonleukoreduced and preserved in CPDA1, which is standard at the Cairo University Blood Bank. Additional 20 leukoreduced units (n = 20) were studied for comparison. All packed red cell units included in this study were meeting the quality control requirements as stated in the National Standards for Blood Transfusion Services.
Preparation of the red cell concentrates
The whole blood (450 ml) was collected in JMS CPDA1 bags which preserve RBCs up to 35 days. Within 6 h of blood collection, whole blood was centrifuged at light spin 1500 g for 10 min at 4°C temperature. Platelet-rich plasma was expressed off the RBCs and separated in a satellite bag. Twenty units were filtered within 30 min of collection to remove leukocytes (log5) and platelets (Macropharma, France). Packed RBC units recovered from each donor (volume ≈250 ml/unit, Hct ≈5%) were further split equally into five satellite bags (volume ≈35 ml/unit). Five satellite bags for each donor were kept under two storage modifications: (i) CDPA only which is the standard condition (2 satellite bags) and (ii) CPDA with the addition of AA in 3 doses (2.06, 4.13, and 6.19 mg/mL) each in dose in a bag. In this way, each donated unit acted as its own control.
CPDA units were aseptically sampled weekly over the total period of storage starting from day 1 of storage, i.e., 1, 7, 14, 21, 28, and 35 days, to study the red cell MP production over storage and the PCA of RBCs MPs. The AA fortified samples were sampled on day 35 (end of storage) and tested for MP production by flow cytometry compared to the corresponding CPDA1 stored unit. Sterility tests (i.e., culture at 4°C, 22°C, and 37°C) were performed as part of the routine quality control procedures.
Ascorbic acid preparation
An AA dose–response study was carried out using AA (Sigma-Aldrich, Product Number A5960) that was prepared in saline solution to a final concentration 2.06, 4.13, and 6.19 mg/mL titrated with sodium hydroxide to reach a pH of 7.1. The solution was then added to store RBCs in a 1:9 vol/vol ratio. The concentrations were selected in reference to the pilot study previously performed.
Sample preparation (centrifugation protocol)
To obtain the supernatant for MPs' quantification (MP-SN), we followed recommendations of the International Society on Thrombosis and Haemostasis. Samples were processed by two sequential centrifugations at 3000 ×g for 15 min at 4°C. MPs were freshly prepared for flow cytometric analysis.
Quantification of microparticles by flow cytometry
Flow cytometric analysis was performed with a FACS CANTO flow cytometer (BD Biosciences). Size events were defined using size reference kit (Invitrogen™ Catalog number: F13839) containing beads of 0.5, 1, and 2 μm in diameter.
50 μl of MP supernatant was stained with 2 μL phycoerythrin anti-human glycophorin A (GPA) (CD235a) antibody (eBioscience catalog Number 12-9987) and 5 μL annexin V-FITC (eBioscience BMS147FI). The optimal concentration was experimentally determined for each antibody by titration experiments. Counting beads (123count eBeads supplied from eBioscience) were used in the absolute counting of the MPs [Figure 1]. All samples were read within 20 min from preparation.
|Figure 1: Flow cytometry gating strategy: (a) Forward scatter and side scatter defining size beads of 0.5, 1, and 2 μm in diameter to establish the microparticle gate between the 0.5 and 1 um. (b) The forward scatter versus side scatter plot showing the position of microparticle gate (red). (c) The forward scatter (annexin V) versus side scatter (glycophorin A) plot showing all events. The microparticles gate (red) and the counting beads (green color)|
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An unstained sample was used as a negative control to detect the auto-fluorescence. Compensation adjustment was done to overcome spectral overlap. Collected data were analyzed by Diva software BD FACSDiva™ software version 8.0 (BD Biosciences, San Jose, CA, USA. MPs were defined by relative size on forward scatter channel (FSC) and side scatter channel (SSC) using a linear scale. The number of MPs per microliter was calculated as follows:
Sample preparation for coagulation assay needed an extra centrifugation step to concentrate the MPs. The supernatant was collected as mentioned for flow cytometric analysis, and then, another centrifugation at 18,000 g for 30 min at 4°C was performed. The supernatant was removed, and the pellet was re-suspended in 100 μl 0.9% NaCl solution.
The plasma to which MPs were added also needed preparation to remove the naturally occurring MPs in them. MP-free human plasma was prepared by three sequential centrifugations (2 × 3000 g and 1 × 18,000 g). AB-pooled plasma screened negative for antiphospholipid antibodies and with normal coagulation profile. This pooled plasma was tested by flow cytometry to ensure there are no detectable GPA-positive MPs.
The PCA of MPs was assessed by re-calcification–time assays. 50 μl MPs suspension was incubated with 50 μl of MP-free human plasma at 37°C. After 180 s, 50 μl of prewarmed 1.5 mM CaCl2 was added to start the reaction and the coagulation time was recorded. The coagulation time test was performed on the coagulation analyzer (CoaDATA 504), which uses turbodensitometric principle for clotting assays [Figure 2].
Data were coded and entered using the Statistical Package SPSS version 24 (SPSS Statistics for Windows, Version 24.0. Armonk, NY: IBM Corp.). Comparison between groups was performed using the unpaired t-test (normally distributed quantitative variables) and nonparametric Mann–Whitney test (nonnormally distributed quantitative variables). For comparison of serial measurements within each group r, ANOVA and Friedman test and Wilcoxon signed-rank test for normally distributed quantitative variables and nonnormally distributed quantitative variables were used, respectively. Correlations between quantitative variables were done using Spearman correlation coefficient. P <0.05 was considered statistically significant.
| Results|| |
Effect of storage and filtration on red cell microparticle production
MPs region was identified in the SSC and FSC parameters by relative size; the upper limit is 1 μ while the lower limit is 0.5 μ. MPs were further identified by the expression of GPA (GPA + MPs), PS which binds to annexin V (PS + MPs), as well as the dual-positive population (GPA + PS + MPS).
The flow cytometric analysis in all samples (filtered and nonfiltered samples) revealed a progressive increase in the number of red cell MPs (GPA + MPs) [Table 1]. The first statistically significant difference from day 1 of storage was reported on day 14. Annexin V-positive (PS+) population and dual-positive population (GPA + MPs) showed a much more delayed significant increase starting on day 21. This finding suggests that filtration has a positive impact on red cell survival [Figure 3].
|Table 1: Flowcytometery and procoagulant assessment of red cell microparticles on storage|
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|Figure 3: Flow cytometry experiment. The above figure shows example of one of the samples analyzed by flow cytometry through the storage duration 35 days. The dual expression of the annexin V and glycophorin A microparticles in quadrant Q2-displayed in biexponential scale|
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Filtration significantly reduced MP production. The filtered samples showed lower levels in the GPA-positive, the total PS-positive, as well as the dual-positive population (P: 0.04, P: 0.001, and P: 0.002, respectively) when compared to nonfiltered samples [Figure 4]. This was evident after 1 week of storage and all through the duration of storage [Table 2].
|Table 2: The effect of ascorbic acid on red cell-derived microparticles on day 35 in filtered samples|
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Coagulation assays of microparticles produced on storage
In both filtered and nonfiltered samples, MPs coagulation time was significantly (P < 0.001) shorter starting from day 7 and the following samples in comparison to day 1 [Table 1].
Correlation between functional and quantitative assay
A strong inverse correlation between coagulation time and MPs was found as when MPs increased in number, the coagulation time was shorter denoting a higher PCA. This was clear with GPA-positive population (r = −0.689, P < 0.001), annexin V-positive MPs (r = −0.723, P < 0.001), and the dual-positive population (r = −0.747, P < 0.001). This highlights the strong correlation between the functional assay (PCA) and direct quantitative assay (flow cytometry).
Ascorbic acid protective effect on stored red blood cells
The red cells stored in AA demonstrated a decrease in the accumulation of RBC MPs by 50% compared to standard storage CPDA-1 alone and that reduction was statistically significant (P < 0.001). The addition of AA at 6.19 mg/ml significantly reduced GPA + MPs values compared to day 35 (P < 0.001), 4.13 mg/ml samples (P: 0.029), and 2.06 mg/ml (P < 0.001). Annexin V expression on MPs was clearly affected by AA showing 82% reduction, with evident dose-dependent manner. The filtered samples in our study were also fortified with the same doses of AA; the result came in agreement with the nonfiltered samples as MP production decreased with increasing the concentration of AA [Table 2].
| Discussion|| |
The aim of blood preservation is to provide functional blood components. A better comprehension of the biochemical and functional changes that occur during the storage may provide better preserving strategies and thus improve the safety of blood products.
In this study, we analyzed red cell MP developed during the storage of packed red cell product as a sign of the storage lesions. Results from this study were consistent with the observation that red cells were subjected to changes during storage which were associated with fragmentation and microvesiculation. The number of RMPs/μl gradually increased during the storage period which was clearly significant around day 14 of storage. Some observational studies that investigated the clinical impact of storage lesions showed that storage for 14 days or longer had greater adverse effects on mortality and/or morbidity, although other studies did not report an effect of longer storage duration.
Jy et al. stated that RMPs showed little increase up to day 10 but thereafter increased steadily with time. However, other studies reported a delayed onset of red cell MP increase. This could be attributed to the use of different procedures in blood processing and storage bags, e.g., leukoreduced CPD-SAGM RBC units., It is also noted that the number of MPs produced varied from donor to donor. The reason for such a variation is unknown, but factors such as ABO blood group, age, fasting, or sex of blood donor may have a role and should be investigated.
The choice of the isolation procedure and preanalytical variables are a major source of variability and potential artifacts in MP analysis. Furthermore, some discrepancies exist in the literature regarding phenotypic characterization. In agreement with our results, the existence of MPs without PS externalization was observed;,, thus, MPs count might be underestimated using annexin V as an identifying marker. Some reports suggested that lactadherin is a more sensitive marker of PS exposure. We observed GPA-negative as well as annexin-positive population that increased over storage. This could be platelet-derived MPs as described by other authors. Further studies using more identifying markers are needed to identify this GPA-negative population.
Leukoreduction has well-established benefit in reducing immune-related transfusion reactions. Recently, more benefits have been introduced as removal of leukocytes provides more nutrients to the RBCs improving the storage conditions and thus enhancing the vitality of RBCs. Our results showed that the number of RMPs in the leukoreduced packed RBCs units was less than that in the nonleukoreduced units. This finding was evident even after 7 days of storage. This observation was also reported by other researchers.,,, Leukoreduced samples showed more delayed onset of annexin V expression; this might signify a possible value of leukoreduction in the preservation of RBCs quality/vitality for a longer time.
The PCA showed a strong correlation with the quantitative assay by flow cytometry. These results support previous observations.,,, The PCA is attributed either to PS externalization or recently to tissue factor expression. The MPs in transfused blood could increase the risk of adverse reactions, by generating a hypercoagulable state, resulting in thromboembolic complications. Inversely, in other situations, a hypercoagulable state may be useful to diminish or even help to stop the bleeding.
Recently, antioxidants have been suggested for the use in stored RBC units as a result of evidence of oxidative damage to protein and lipid components of RBCs. Modern storage ASs do not contain ingredients aiming at the inhibition and/or detoxification of reactive oxidative species. The RBCs stored in AA demonstrated a decrease in MPs production with evident dose-dependent effect. RBC MPs were almost 50% lower in AA fortified samples compared to standard storage. The higher concentration of AA (6.19 mg/ml) showed a significant reduction of RMPs when compared to other lower concentrations. It has been reported that adding AA solution led to enhanced RBC recovery, decreased MP formation, and reduced alloimmunization. In the same notion, AA has been shown to benefit RBCs survival by reducing the hemolysis and mechanical fragility index. However, Vani et al. showed that Vitamin C could not sufficiently attenuate oxidative stress on RBC storage. It is worth noting that the latter study was performed on male Wistar rats, not human RBCs, and stored for 25 days at 4°C. Rat RBCs have a shorter half-life than those of humans. The murine RBCs undergo the same aging processes under storage but with more accelerated deterioration.
It is possible that AA exerts its effect on red cells by interacting with reactive oxygen species produced in the liquid component of the packed RBCs by residual WBCs and/or PLTs. AA might also be acting in close to or directly upon the RBC membrane. AA does not passively enter RBCs; thus, to affect the internal environment of the RBC, AA would have to be actively transported across the membrane. This was postulated to occur through high-affinity AA receptors (SVCT1 and SVCT2). However, other investigators have reported that the SVCT proteins are lost during maturation of RBCs in the bone marrow. Instead, mature RBCs can use the GLUT1 transporter.
| Conclusion|| |
The field of MP study is rapidly expanding and can be implicated in the assessment of blood products by careful monitoring and quantification of their release during RBC storage time. This study showed the increased procoagulant activity of stored blood that correlated strongly with the MPs count. RBCs in the leukoreduced samples showed lower MPs production. Our work is significant since it uses a naturally occurring vitamin to extend the shelf life of stored RBCs without incurring any significant changes in current blood banking practices. However, these results should be carried out on wider scale and testing various parameters of RBCs vitality and safe transfusion.
We would like to acknowledge the soul of Prof. Dr. Ali Shams El-Din, for his substantive contributions to the field of clinical pathology in Egypt.
Financial support and sponsorship
Cairo University supported the study.
Conflicts of interest
There are no conflicts of interest.
| References|| |
Antwi-Baffour S, Danso S, Adjei J, Kyeremeh R, Addae M. Prolonged Storage of Red Blood Cells for Transfusion in Citrate Phosphate Dextrose Adenine-1 Affects Their Viability. Open Access Library Journal 2015;2:1-7. doi: 10.4236/oalib.1101908.
Antonelou MH, Kriebardis AG, Stamoulis KE, Economou-Petersen E, Margaritis LH, Papassideri IS. Red blood cell aging markers during storage in citrate-phosphate-dextrose-saline-adenine-glucose-mannitol. Transfusion 2010;50:376-89.
Rajashekharaiah V, Koshy AA, Koushik AK, Kaur H, Kumari K, Agrawal M, et al
. The efficacy of erythrocytes isolated from blood stored under blood bank conditions. Transfus Apher Sci 2012;47:359-64.
Chaudhary R, Katharia R. Oxidative injury as contributory factor for red cells storage lesion during twenty eight days of storage. Blood Transfus 2012;10:59-62.
Kriebardis AG, Antonelou MH, Stamoulis KE, Economou-Petersen E, Margaritis LH, Papassideri IS. Progressive oxidation of cytoskeletal proteins and accumulation of denatured hemoglobin in stored red cells. J Cell Mol Med 2007;11:148-55.
Straat M, Böing AN, Tuip-De Boer A, Nieuwland R, Juffermans NP. Extracellular vesicles from red blood cell products induce a strong pro-inflammatory host response, dependent on both numbers and storage duration. Transfus Med Hemother 2016;43:302-5.
Osteikoetxea X, Németh A, Sódar BW, Vukman KV, Buzás EI. Extracellular vesicles in cardiovascular disease: Are they Jedi or Sith? J Physiol 2016;594:2881-94.
Yáñez-Mó M, Siljander PR, Andreu Z, et al
. Biological properties of extracellular vesicles and their physiological functions. J Extracell Vesicles. 2015; 4:27066. Published 2015 May 14. doi:10.3402/jev. v4.27066.
van Beers EJ, Schaap MC, Berckmans RJ, Nieuwland R, Sturk A, van Doormaal FF, et al
. Circulating erythrocyte-derived microparticles are associated with coagulation activation in sickle cell disease. Haematologica 2009;94:1513-9.
van der Pol E, Böing AN, Harrison P, Sturk A, Nieuwland R. Classification, functions, and clinical relevance of extracellular vesicles. Pharmacol Rev 2012;64:676-705.
Maślanka K, Uhrynowska M, Łopacz P, Wróbel A, Smoleńska-Sym G, Guz K, et al
. Analysis of leucocyte antibodies, cytokines, lysophospholipids and cell microparticles in blood components implicated in post-transfusion reactions with dyspnoea. Vox Sang 2015;108:27-36. doi: 10.1111/vox.12190. Epub 2014 Aug 18. PMID: 25134637.
Fontes J. Biopreservation: Extension of the ex vivo
Life Span of Stored Human Erythrocytes by the Addition of Ascorbic Acid to Additive Solutions in Modern Blood Banking. Electronic Thesis or Dissertation. Ohio State University, 2014. OhioLINK Electronic Theses and Dissertations Center. 17 Mar 2020.
Gamonet C, Mourey G, Aupet S, Biichle S, Petitjean R, Vidal C, et al
. How to quantify microparticles in RBCs? A validated flow cytometry method allows the detection of an increase in microparticles during storage. Transfusion 2017;57:504-16.
Zhao L, Bi Y, Kou J, Shi J, Piao D. Phosphatidylserine exposing-platelets and microparticles promote procoagulant activity in colon cancer patients. J Exp Clin Cancer Res 2016;35:54.
Wang D, Sun J, Solomon SB, Klein HG, Natanson C. Transfusion of older stored blood and risk of death: A meta-analysis. Transfusion 2012;52:1184-95.
McQuilten ZK, French CJ, Nichol A, Higgins A, Cooper DJ. Effect of age of red cells for transfusion on patient outcomes: A systematic review and meta-analysis. Transfus Med Rev 2018;32:77-88.
Jy W, Ricci M, Shariatmadar S, Gomez-Marin O, Horstman LH, Ahn YS. Microparticles in stored red blood cells as potential mediators of transfusion complications. Transfusion 2011;51:886-93.
Almizraq R, Tchir JD, Holovati JL, Acker JP. Storage of red blood cells affects membrane composition, microvesiculation, andin vitro
quality. Transfusion 2013;53:2258-67.
Rubin O, Crettaz D, Canellini G, Tissot JD, Lion N. Microparticles in stored red blood cells: An approach using flow cytometry and proteomic tools. Vox Sang 2008;95:288-97.
Revenfeld AL, Bæk R, Nielsen MH, Stensballe A, Varming K, Jørgensen M. Diagnostic and prognostic potential of extracellular vesicles in peripheral blood. Clin Ther 2014;36:830-46.
Xiong Z, Oriss TB, Cavaretta JP, Rosengart MR, Lee JS. Red cell microparticle enumeration: Validation of a flow cytometric approach. Vox Sang 2012;103:42-8.
Connor DE, Exner T, Ma DD, Joseph JE. The majority of circulating platelet-derived microparticles fail to bind annexin V, lack phospholipid-dependent procoagulant activity and demonstrate greater expression of glycoprotein Ib. Thromb Haemost 2010;103:1044-52.
Tissot JD, Rubin O, Canellini G. Analysis and clinical relevance of microparticles from red blood cells. Curr Opin Hematol 2010;17:571-7.
Sparrow RL, Chan KS. Microparticle content of plasma for transfusion is influenced by the whole blood hold conditions: Pre-analytical considerations for proteomic investigations. J Proteomics 2012;76:211-9.
Pertinhez TA, Casali E, Baroni F, Berni P, Baricchi R, Spisni A. A comparative study of the effect of leukoreduction and pre-storage leukodepletion on red blood cells during storage. Front Mol Biosci 2016;3:13.
Saito S, Nollet KE, Ngoma AM, Ono T, Ohto H. Platelet-, leucocyte- and red cell-derived microparticles in stored whole blood, with and without leucofiltration, with and without ionising radiation. Blood Transfus 2018;16:145-53.
Crompot E, Van Damme M, Duvillier H, Pieters K, Vermeesch M, Perez-Morga D, et al
. Avoiding false positive antigen detection by flow cytometry on blood cell derived microparticles: The importance of an appropriate negative control. PLoS One 2015;10:e0127209.
Kriebardis A, Antonelou M, Stamoulis K, Papassideri I. Cell-derived microparticles in stored blood products: Innocent-bystanders or effective mediators of post-transfusion reactions? Blood Transfus 2012;10 Suppl 2:s25-38.
Sugawara A, Nollet KE, Yajima K, Saito S, Ohto H. Preventing platelet-derived microparticle formation – And possible side effects-with prestorage leukofiltration of whole blood. Arch Pathol Lab Med 2010;134:771-5.
Piccin A, Van Schilfgaarde M, Smith O. The importance of studying red blood cells microparticles. Blood Transfus 2015;13:172-3.
Ayers L, Harrison P, Kohler M, Ferry B. Procoagulant and platelet-derived microvesicle absolute counts determined by flow cytometry correlates with a measurement of their functional capacity. J Extracell Vesicles. 2014; 3:10.3402/jev.v3.25348. Published 2014 Sep 30. doi:10.3402/jev.
Leroyer AS, Anfosso F, Lacroix R, Sabatier F, Simoncini S, Njock SM, et al
. Endothelial-derived microparticles: Biological conveyors at the crossroad of inflammation, thrombosis and angiogenesis. Thromb Haemost 2010;104:456-63.
Rubin O, Crettaz D, Tissot JD, Lion N. Microparticles in stored red blood cells: Submicron clotting bombs? Blood Transfus 2010;8 Suppl 3:s31-8.
Stowell SR, Smith NH, Zimring JC, Fu X, Palmer AF, Fontes J, et al
. Addition of ascorbic acid solution to stored murine red blood cells increases posttransfusion recovery and decreases microparticles and alloimmunization. Transfusion 2013;53:2248-57.
Vani R, Soumya R, Carl H, Chandni VA, Neha K, Pankhuri B, et al
. Prospects of Vitamin C as an Additive in Plasma of Stored Blood. Adv Hematol. 2015;2015:961049. doi: 10.1155/2015/961049. Epub 2015 Aug 9. PMID: 26345502; PMCID: PMC4546735.
Makley AT, Goodman MD, Friend LA, Johannigman JA, Dorlac WC, Lentsch AB, et al
. Murine blood banking: Characterization and comparisons to human blood. Shock 2010;34:40-5.
Savini I, Rossi A, Pierro C, Avigliano L, Catani MV. SVCT1 and SVCT2: Key proteins for vitamin C uptake. Amino Acids 2008;34:347-55.
May JM, Qu ZC, Qiao H, Koury MJ. Maturational loss of the vitamin C transporter in erythrocytes. Biochem Biophys Res Commun 2007;360:295-8.
Montel-Hagen A, Kinet S, Manel N, Mongellaz C, Prohaska R, Battini JL, et al
. Erythrocyte Glut1 triggers dehydroascorbic acid uptake in mammals unable to synthesize vitamin C. Cell 2008;132:1039-48.
[Figure 1], [Figure 2], [Figure 3], [Figure 4]
[Table 1], [Table 2]