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Differences in levels of platelet-derived microparticles in platelet components prepared using the platelet rich plasma, buffy coat, and apheresis procedures
ARTICLE IN PRESS Transfusion and Apheresis Science ■■ (2016) ■■–■■
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Transfusion and Apheresis Science j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / t r a n s c i
Differences in levels of platelet-derived microparticles in platelet components prepared using the platelet rich plasma, buffy coat, and apheresis procedures Egarit Noulsri a,*, Prapaporn Udomwinijsilp b, Surada Lerdwana c, Viroje Chongkolwatana b, Parichart Permpikul b,** a
Research Division, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand Department of Transfusion Medicine, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand c Department of Research and Development, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand b
A R T I C L E
I N F O
Article history: Received 2 August 2016 Received in revised form 7 October 2016 Accepted 21 October 2016 Keywords: Platelet-derived microparticle Platelet component Buffy coat Platelet rich plasma Apheresis
A B S T R A C T
Background: There has been an increased interest in platelet-derived microparticles (PMPs) in transfusion medicine. Little is known about PMP status during the preparation of platelet concentrates for transfusion. Aim: The aim of this study is to compare the PMP levels in platelet components prepared using the buffy coat (BC), platelet-rich plasma platelet concentrate (PRP-PC), and apheresis (AP) processes. Methods: Platelet components were prepared using the PRP-PC and BC processes. Apheresis platelets were prepared using the Trima Accel and Amicus instruments. The samples were incubated with annexin A5-FITC, CD41-PE, and CD62P-APC. At day 1 after processing, the PMPs and activated platelets were determined using flow cytometry. Results: Both the percentage and number of PMPs were higher in platelet components prepared using the Amicus instrument (2.6 ± 1.8, 32802 ± 19036 particles/μL) than in platelet components prepared using the Trima Accel instrument (0.5 ± 0.4, 7568 ± 5298 particles/ μL), BC (1.2 ± 0.6, 12,920 ± 6426 particles/μL), and PRP-PC (0.9 ± 0.6, 10731 ± 5514 particles/ μL). Both the percentage and number of activated platelets were higher in platelet components prepared using the Amicus instrument (33.2 ± 13.9, 427553 ± 196965 cells/ μL) than in platelet components prepared using the Trima Accel instrument (16.2 ± 6.1, 211209 ± 87706 cells/μL), BC (12.9 ± 3.2, 140624 ± 41003 cells/μL), and PRP-PC (21.1 ± 6.3, 265210 ± 86257 cells/μL). Conclusions: The study suggests high variability of PMPs and activated platelets in platelet components prepared using different processes. This result may be important in validating the instruments involved in platelet blood collection and processing. © 2016 Elsevier Ltd. All rights reserved.
1. Introduction * Corresponding author. Research Division, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand. Fax: + 66-2-411-0175. E-mail address: [email protected] (E. Noulsri). ** Corresponding author. Department of Transfusion Medicine, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand. Fax: + 66-2-412-8419. E-mail address: [email protected] (P. Permpikul).
In recent years, there has been an increased interest in cell-derived microparticles (MPs) in transfusion medicine. The MPs in blood circulation can be generated by red blood cells, endothelial cells, white blood cells (WBCs), and platelets. Upon activation or apoptosis, the platelets release
http://dx.doi.org/10.1016/j.transci.2016.10.006 1473-0502/© 2016 Elsevier Ltd. All rights reserved.
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small plasma membranes, 0.1–1 μm, or platelet-derived microparticles (PMPs). These PMPs have been shown to be the most-abundant MPs in the blood circulation. Studies have suggested that higher levels of PMPs in the blood circulation are associated with thrombosis complications and inflammation [1–3]. Upon being shed from the activated platelets, the PMPs express the phosphatidylserine (PS) that is on their surface. Negatively charged phospholipid can further activate the coagulation factor, resulting in thrombus formation. PMPs also exhibit the tissue factor on their surfaces. Upon binding with FVIIa, the TF can initiate coagulation [4]. In addition, an in-vitro study using the thrombin generation test demonstrated the coagulation activity of the PMP [5]. Several studies have documented the potential role of PMPs in cardiovascular disease. For example, Siljander et al. demonstrated that PMPs are associated with fibril formation [6]. In addition, a significant increase in PMP was reported in patients with acute pulmonary embolism [7]. Furthermore, a number of studies have suggested the role of PMPs in tumor progression [8–11], including chemotaxis activity, angiogenesis activity, and the promotion of tumor metastasis. Therefore, to minimize recipients’ complications, it is necessary to monitor the PMP in the platelet-preparation process in transfusion medicine. Platelet blood products can be prepared by various methods, including the buffy coat (BC), platelet-rich plasma platelet concentrate (PRP-PC), and apheresis (AP) methods. In the AP method, platelets are collected from a donor, and the remaining components are returned to the donor, a process mediated by AP instruments. In contrast, platelet products collected by the BC and PRP-PC processes are pooled from multiple donors. Since each cell in the blood has its own specific gravity, centrifugation is applied to separate each blood component for removal. International guidelines recommend that the half-life of platelet concentrates in currently available plateletstorage bags is 5 days at 22 ± 2 °C with continuous agitation. A number of studies have shown that platelets undergo various changes during collection, processing, and storage [12]. PMPs can be detected in platelet components prepared using either whole-blood processes or AP instruments [13–16]. Higher levels of PMPs in the blood product may result from the harvesting conditions and the storage lesion. However, the effect of AP on PMP counts remains controversial. Recent studies have suggested that some AP procedures may generate more PMPs than others [17,18]. In contrast, other studies have shown increased levels of PMPs in stored platelet concentrate and AP [19,20]. However, no studies have investigated the effects of various platelet-concentrate preparation procedures on PMP counts. Therefore, the present study aimed to determine whether various platelet-component preparation procedures affected the PMP counts. The platelet components were prepared using the AP, BC, and PRP-PC processes under routine protocol. The PMPs and activated platelets in these platelet concentrates were determined using flow cytometry. The study’s results are important to understanding which bloodproduct manufacturing steps might trigger the release of PMPs.
2. Materials and methods 2.1. Materials Fluorescein isothiocyanate conjugated annexin A5 (annexin A5-FITC), phycoerythrin conjugated CD41a (CD41aPE), allophycocyanin conjugated CD62P (CD62P-APC), and 10 × annexin A5 binding buffer were purchased from BD (San Jose, CA, USA). CountBright™ counting beads were obtained from Invitrogen (Carlsbad, CA, USA). Blank calibration particles of 1.09 μm were purchased from Spherotech (Lake Forest, IL, USA). 2.2. Subjects This study was approved by the Institutional Review Board of Siriraj Hospital, Mahidol University School of Medicine, Bangkok, Thailand (COA no. Si349/2015). Written consent of the donors was received after the procedure was explained to them in detail, including the time the study would take and its possible hazards and benefits. After obtaining this informed consent, blood samples were collected using the standard procedure for collecting donated blood. Before the platelet-component preparation procedure, ABO/Rh typing and testing for infectious disease markers (HIV, HBV, HCV, and syphilis) were conducted. The study included healthy subjects aged 18–65 without known blood dyscrasia. The exclusion criteria included a medical history of blood-derived illness or taking any medication known to affect platelet or bone marrow function or concentration within two weeks prior to the test. In addition, the study excluded donors who had taken aspirin or other nonsteroidal anti-inflammatory drugs (NSAIDS), which are likely to affect platelet function, within two weeks prior to the test. 2.3. Buffy coat platelet concentrates (BC-PC) Platelet concentrates were prepared from buffy coats derived from whole blood using a standard procedure described previously [21,22]. Briefly, 450 mL of whole blood from healthy donors was collected in 450-mL quadruple bags containing 63 mL of citrate-phosphate dextrose (CPD) as an anticoagulant. Then, the blood was centrifuged (Heraeus 6000i; Germany) at 3380 rpm at 22 °C with acceleration and deceleration curves of 3 and 5, respectively. After 10 min, the whole blood was separated into components. The top layer, 150–200 mL, was the platelet poor plasma (PPP). The middle layer was the buffy coat, containing approximately 90% of the platelets, 70% of the WBCs, and 10% of the red blood cells. The bottom layer was the pack red-blood cells. The PPP supernatant and buffy coat were transferred into other satellite bags. A solution of saline, adenine, glucose, and mannitol (SAGM) (TERUMO PENPOL, Ltd.; Puliyarakonam, Trivandrum, India) was added to the red cells as an additive. The bags containing the red cells and plasma were removed, and the red cells were stored at 4 °C in a cold room. The PPP was stored as fresh frozen plasma (FFP) in a freezer at −20 °C. Four bags of the buffy coat were gently mixed with plasma (200–250 mL) from the same blood group as the donors’ and then centrifuged at 1600 rpm for
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6 min at 22 °C along with one empty satellite bag. The supernatant platelet rich plasma (PRP) was transferred into an empty platelet-storage bag and the tubing was sealed.
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instrument was confirmed each day using two levels of controls (Sysmex e-Check Hematology Control for Sysmex X-Series Analyzers; Sysmex; USA) according to the manufacturer’s recommendations.
2.4. Platelet rich plasma-platelet concentrates (PRP-PC) 2.7. Flow cytometry analysis of PMPs and activated platelets First, 450 ml of whole blood was collected in a 450-ml triple bag containing 63 mL of CPD anticoagulant and red blood cell preservative (TERUMO PENPOL, Ltd. Puliyarakonam, Trivandrum, India). The PRP was separated from the whole blood by centrifugation (Heraeus 6000i, Germany) at 3000 rpm at 21 °C for 10 min, with acceleration and deceleration curves of 5 and 4, respectively. Then, the platelets were concentrated by centrifugation at 3940 rpm for 5 min at 21 °C, with acceleration and deceleration curves of 9 and 5, respectively, and the supernatant plasma was removed. The platelet-concentrate bag was left at room temperature for approximately 2 h before being stored at or below 22 ± 2 °C with reciprocal agitation. 2.5. Apheresis platelet (AP) For platelet apheresis, the entire process was performed in a closed system. A disposable kit was installed onto the continuous-blood cell separator before the machine was primed. The donors were prepared by cleaning the venipuncture sites at the antecubital areas, and then phlebotomy was performed with minimal trauma to the donors. For all Trima instrument collections, Gambro Trima Accel software version 5.1 (Terumo BCT; Lakewood, CO, USA) was used, with the following settings: AC management: 2; AC ratio: 1:10; minimum donor post-HCT: 32%; minimum donor post-platelet count: 100,000; maximum draw flow: moderate; draw management: 2; return management: 1; predicted yield: 4.0–9.5 × 1011/μL; final concentration: 1200– 1400 × 103/μL; and collection volume: 285–316 mL. For all Amicus instrument collections, Amicus software version 3.1 (Fresenius-Kabi Company; Lake Zurich, IL, USA) was used, with the following settings: AC infusion rate: 1.25 mg/kg/ min; AC ratio: 10:1; maximum return rate: 120 mL/min; maximum inlet rate: 100 mL/min; and maximum cycle volume: 150–300 mL. Draw and return rates were determined by the instrument unless adjustments were needed to address access or donor-comfort issues. Plateletconcentrate products were stored at 22 ± 2 °C with continuous agitation. 2.6. Measuring pH, WBCs, and platelet counts The pH of platelet products was determined using a Mettler-Toledo Delta 320 pH meter (Mettler-Toledo; Greifensee, Switzerland), and the measurement was carried out at room temperature. The pH meter’s electrode was placed in the platelet product and swirled. The results were recorded when the pH reading stabilized. The WBC and platelet counts were determined using an automated hematology analyzer, XS-800i (Sysmex Corporation; Kobe, Japan), after control setup and calibration as recommended by the manufacturer. Samples were analyzed within 30 min of collection. Calibration of the
To investigate the expression of phosphatidylserine and CD62P on MPs and platelets, 20 μL of platelet suspensions diluted at 1:1000 was incubated with 2 μl of annexin A5FITC, 2 μL of CD62P-APC, and 2 μl of CD41a-PE in the dark at room temperature. HEPES buffer without calcium was used as a negative control for annexin A5 binding. After 15 min, 25 μL of counting beads and 300 μL of 1 × annexin A5 binding buffer were added to the stained samples, which were then acquired and analyzed using FACSDiva™ software on a FACSCanto flow cytometer (BD; San Jose, CA, USA). Fig. 1 summarizes the gating strategies for flow cytometry analysis. The forward scatter (FSC), side scatter (SSC), and fluorescence (FL) parameters were set as logarithmic scale. Instrument performance was verified using CaliBRITE™ beads (BD; San Jose, CA, USA). Initially, an FSC vs. SSC dotplot was used to identify platelets, counting beads, and MPs (Fig. 1A). MP gating was performed according to 1-μm standard beads. The gated MPs were further analyzed for their percentage and number on the annexin A5 vs. SSC dotplot (Fig. 1B). To determine the percentage and number of PMPs, a CD41 vs. CD62P dot-plot was used (Fig. 1C). Likewise, the percentage and number of activated platelets from the gated platelet population were analyzed using CD41 vs. SSC (Fig. 1D) and annexin A5 vs. CD62P dot-plots (Fig. 1E). To enable reliable analysis, data were collected for at least 1300 events of counting beads at medium speed. The absolute number was calculated using the following formula: A = ((B × C)/(D × E)) × F, where A is the concentration of the cell population (cells/μL), B is the events of the cell population, C is the number of the counting beads per test (beads/ 25 μL), D is the events of the counting beads, E is the volume of the samples (μL), and F is the dilution factor. 2.8. Statistical analysis Data analysis and graphing were performed using GraphPad Prism software version 5.0 (GraphPad; San Diego, CA, USA). Results were expressed as mean and standard deviation (SD). The Mann–Whitney U test was used to determine the mean difference between the groups. A p ≤ 0.05 was considered statistically significant. 3. Results 3.1. Number of samples, donor age, collection time, and platelet-component volume in the three processes This study analyzed a total of 97 platelet components, with 19, 17, and 61 being prepared using the PRP-PC, BC, and AP processes, respectively. Of the 61 AP samples, 46 were prepared using the Trima Accel instrument and the other 15 using the Amicus instrument. The donors’ ages for the PRP-PC, BC, and AP processes had means ± SD (range) of
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Fig. 1. Flow cytometry gating strategy for analyzing PMPs and activated platelets. (A) FSC vs. SSC dot-plot represents the population of MPs based on the standard 1-μm bead, platelets, and counting beads in the defined gate. (B) Annexin A5 vs. SSC dot-plot distinguishes the annexin A5-positive MPs in the P1 region from the gated MPs. (C) CD41a vs. CD62P dot-plot distinguishes the PMPs in quadrants Q2 and Q4 from the previous P1 region. (D) CD41a vs. SSC dot-plot distinguished CD41a-positive population in the region from the gated platelets. (E) Annexin A5 vs. CD62P dot-plot distinguishes the activated platelets in quadrants Q1-1 and Q2-1 from the CD41-positive population in the previous region. The plot also shows the percentage of cells present in each quadrant. Data are representative of three independent experiments.
38.5 ± 10.8 (21–55), 36.9 ± 11.2 (20–62), and 47.6 ± 15.8 (24– 67), respectively. The collection times (min) for the three procedures were 7.1 ± 19.3 (5–12), 6.6 ± 2 (4–12), and 93.5 ± 15.8 (51–136), respectively. 3.2. Optimizing flow cytometry methods to determine PMP and activated platelet levels The first experiment aimed to optimize the PMP gating strategy. The platelet products were left at room temperature without treatment for 7 days. The levels of PMP and activated platelets were determined by flow cytometry. A 1.9-fold increase of PMPs compared to day 1 was found in stored platelet components. Similarly, a 3.3-fold increase compared to day 1 was found in CD62P-positive platelets in stored platelet components. In addition, a 16-fold increase compared to day 1 was found in the percentage of annexin A5-positive platelets in stored platelet components. Examining the activated platelets revealed that the percentage of CD62P increased time-dependently, and the highest values were found on day 6, suggesting that flow cytometry gating is optimal for future experiments.
3.3. PMPs and activated platelets in platelet components prepared using AP, BC, and PRP-PC processes Next, the levels of PMPs and activated platelets were determined in platelet components prepared using the AP, BC, and PRP-PC processes. Because the AP was prepared using two different instruments, we categorized the AP into Amicus and Trima Accel and compared the results with those of the BC and PRP-PC processes. As shown in Fig. 2A, in platelet components prepared using the Amicus instrument (2.6 ± 1.8), the percentage of PMPs at day 1 was significantly higher than the percentage in platelet components prepared using the Trima Accel instrument (0.5 ± 0.4), BC (1.2 ± 0.6), and PRP-PC (0.9 ± 0.6). Although the percentage of PMPs in platelet components prepared using BC was slightly higher than the percentage in platelet components prepared using PRP-PC, this difference was not statistically significant. Further experiments revealed that the number of PMPs in platelet components prepared using the Amicus instrument (32,802 ± 19,036 particles/μL) was significantly higher than the number in platelet components prepared using the Trima Accel instrument
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Fig. 2. Comparison of PMPs and activated platelets in platelet concentrate prepared using various procedures. The scatter plots show the percentage (A) and the number (B) of the PMPs and the percentage (C) and the number (D) of the activated platelets in platelet components prepared using the Amicus, Trima Accel, BC, and PRP-PC processes. A p value indicates significant differences among the groups.
(7568 ± 5298 particles/μL), BC (12,920 ± 6426 particles/ μL), and PRP-PC (10,731 ± 5514 particles/μL) (Fig. 2B). Similar to the percentage, the number of PMPs in the platelet components prepared using BC was slightly higher than the number in platelet components prepared using PRP-PC, but this difference was not statistically significant. Examining the activated platelets showed a significant increase in the percentage of CD62P-positive platelets in components prepared using the Amicus instrument (33.2 ± 13.9) compared to the percentage in platelet components prepared using the Trima Accel instrument (16.2 ± 6.1), BC (12.9 ± 3.2), and PRP-PC (21.1 ± 6.3) (Fig. 2C). Similarly, the number of CD62Ppositive platelets in platelet components prepared using the Amicus instrument (427,553 ± 196,965 cells/μL) was higher than the number in platelet components prepared using the Trima instrument (211,209 ± 87,706 cells/μL), BC (140,624 ± 41,003 cells/μL), and PRP-PC (265,210 ± 86,257 cells/μL) (Fig. 2D).
3.4. Comparing platelet count, leukocyte count, volume, and pH of platelet components prepared using AP, BC, and PRPPC processes Recent experiments have shown high variability in PMP levels in platelet components prepared using the AP process. We checked whether the platelet and leukocyte count and pH might affect the level of PMP formation. As shown in Fig. 3A, the number of platelets in platelet components prepared using the Amicus instrument (1581 × 103 ± 122 cells/ μL) did not differ from the number in platelet components prepared using the Trima Accel instrument (1595 × 103 ± 191 cells/μL). The highest number of platelets was found in platelet components prepared using PRP-PC (1671 × 103 ± 452 cells/μL), but the difference was not statistically significant when compared to platelet components prepared using BC (1409 × 103 ± 215 cells/μL). The pH in the platelet components at day 1 after preparation was measured, and the
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Fig. 3. The platelet, pH, and leukocyte numbers in platelet concentrate prepared using various procedures. The scatter plots represent the number of platelets (A), pH (B), and the number of leukocytes (C) in platelet component prepared using the Amicus, Trima Accel, BC, and PRP-PC processes. A p value indicates significant differences among the groups.
results showed that the pH in the platelet components prepared using the Amicus instrument (7.1 ± 0.2) was significantly lower than the pH in the platelet components prepared using the Trima Accel instrument (7.5 ± 0.1), BC (7.2 ± 0.1), and PRP-PC (7.5 ± 0.1) (Fig. 3B). In addition, the results showed that the number of leukocytes in platelet components prepared using PRP-PC (1077 ± 615 cells/ μL) was significantly higher than the number in platelet components prepared using the Amicus instrument (96 ± 52 cells/μL), the Trima Accel instrument (20 ± 34 cells/μL), and BC (209 ± 99 cells/μL) (Fig. 3C). 4. Discussion Platelet concentrate is crucial to correcting bleeding tendencies in transfusion medicine patients. Platelet components can be prepared by various processes, including AP, BC, and PRP-PC. Recent studies have suggested that the preparation method may affect the quality of the platelets, as some studies have found increased levels of activated platelets and PMPs [14,16]. Higher PMP levels have been shown to be associated with thrombosis complications, tumor progression, and metastasis [1–3]. To minimize these complications in recipients, it is essential to validate the platelet-concentrate preparation processes in individual transfusion laboratories. The present study determined the PMP levels in platelet concentrates prepared using the AP, BC, and PRP-PC procedures, clearly demonstrating that the AP instrument used affects the PMP count, as shown by the significant increase in the number of PMPs in the platelet component prepared using various AP instruments compared to the numbers in platelet components prepared using the BC and PRP-PC processes. In addition, both the percentage and number of PMPs showed a greater range of values in the AP prepared using various AP instruments than in those prepared using the BC and PRP-PC processes. Further experiments conducted during the present study supported this result by showing increases in both PMPs and activated platelets in AP prepared using the Amicus instrument compared to that prepared using the Trima Accel
instrument. These results agree with those of previous studies that have shown differences in AP product prepared using the Amicus and Trima Accel instruments. Those studies found higher amounts of CD62P-positive platelets in AP prepared using the Amicus instrument than in that prepared using the Trima Accel instrument [17]. This is likely due to several reasons [19,23]. When preparing AP using the Amicus instrument, the platelets directly contact the chamber wall. In contrast, when AP is prepared using the Trima Accel instrument, the buffy coat prevents platelet contact with the chamber wall. In addition, in the final Amicus process, plasma is drawn into the bag containing the platelets, which may cause platelet activation via shear stress. The present study’s findings are important for the following reasons. To meet the expectations of transfusion medicine, it is essential to consider the quality of the platelet products when evaluating automated blood-processing systems or platelet-preparation procedures. Since there are a number of AP instruments on the market, laboratory tests must be established to assess the quality of these instruments. PMP enumeration by flow cytometry is recommended to test the quality of the screening of AP batches. Several studies have reported correlations between increased PMP levels in blood products and the quality of those products [12,24–27]. Studies of platelet recovery and survival have shown low post-transfusion platelet recovery and short platelet survival in platelet products with high P-selectin expression. Therefore, the levels of PMPs and activated platelets correlate with the quality of the AP, and monitoring these levels may help assess the quality of the AP equipment used. However, the possible clinical repercussions that may be caused by differences in recovery and survival characteristics remain unknown. Further studies should address these post-transfusion characteristics associated with platelet products. Results of the present study showed differences in the pH, platelet number, and leukocyte contamination. However, the differences in these parameters were within the ranges allowed by recommended guidelines, and several studies
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have found that such differences have no effect on platelet quality [28,29]. In addition, the present study’s findings agreed with the results of previous studies on the quality of platelets in platelet components [14,20]. These differences in pH, platelet, and leukocyte numbers may be explained by differences in centrifugation speed, storage bags, and storage conditions among transfusion laboratories [20]. One might suggest that the differences observed in the present study resulted from the storage lesion of the platelets. The present study compared the levels of PMPs and activated platelet components on day 1 after processing. However, previous studies have suggested that the storage lesion of the blood component begins after day 5 [20]. Therefore, the storage lesion cannot be the explanation for the difference noted in the present study. 5. Conclusions The present study found various levels of PMPs and activated platelets in platelet components prepared using the AP, BC, and PRP-PC procedures. In addition, the results documented the effects of the various instruments used in the AP method on the levels of PMPs and activated platelets. This data on PMPs in platelet components may be useful to assessing platelet quality and preparation procedures. Acknowledgments The authors thank the Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand, for supporting this research project. References [1] Solum NO. Procoagulant expression in platelets and defects leading to clinical disorders. Arterioscler Thromb Vasc Biol 1999;19:2841–6. [2] Jy W, Mao WW, Horstman L, Tao J, Ahn YS. Platelet microparticles bind, activate and aggregate neutrophils in vitro. Blood Cells Mol Dis 1995;21:217–31, discussion 31a. [3] Herring JM, McMichael MA, Smith SA. Microparticles in health and disease. J Vet Intern Med 2013;27:1020–33. [4] Puddu P, Puddu GM, Cravero E, Muscari S, Muscari A. The involvement of circulating microparticles in inflammation, coagulation and cardiovascular diseases. Can J Cardiol 2010;26:140–5. [5] Diamant M, Tushuizen ME, Sturk A, Nieuwland R. Cellular microparticles: new players in the field of vascular disease? Eur J Clin Invest 2004;34:392–401. [6] Siljander P, Carpen O, Lassila R. Platelet-derived microparticles associate with fibrin during thrombosis. Blood 1996;87:4651–63. [7] Bal L, Ederhy S, Di Angelantonio E, Toti F, Zobairi F, Dufaitre G, et al. Circulating procoagulant microparticles in acute pulmonary embolism: a case-control study. Int J Cardiol 2010;145:321–2. [8] Goubran H, Sabry W, Kotb R, Seghatchian J, Burnouf T. Platelet microparticles and cancer: an intimate cross-talk. Transfus Apher Sci 2015;53:168–72. [9] Aharon A, Brenner B. Microparticles, thrombosis and cancer. Best Pract Res Clin Haematol 2009;22:61–9.
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[10] Varon D, Hayon Y, Dashevsky O, Shai E. Involvement of platelet derived microparticles in tumor metastasis and tissue regeneration. Thromb Res 2012;130(Suppl. 1):S98–9. [11] Falanga A, Tartari CJ, Marchetti M. Microparticles in tumor progression. Thromb Res 2012;129(Suppl. 1):S132–6. [12] Singh RP, Marwaha N, Malhotra P, Dash S. Quality assessment of platelet concentrates prepared by platelet rich plasma-platelet concentrate, buffy coat poor-platelet concentrate (BC-PC) and apheresis-PC methods. Asian J Transfus Sci 2009;3:86–94. [13] Gyulkhandanyan AV, Mutlu A, Freedman J, Leytin V. Markers of platelet apoptosis: methodology and applications. J Thromb Thrombolysis 2012;33:397–411. [14] Burnouf T, Chou ML, Goubran H, Cognasse F, Garraud O, Seghatchian J. An overview of the role of microparticles/microvesicles in blood components: are they clinically beneficial or harmful? Transfus Apher Sci 2015;53:137–45. [15] Flaumenhaft R, Dilks JR, Richardson J, Alden E, Patel-Hett SR, Battinelli E, et al. Megakaryocyte-derived microparticles: direct visualization and distinction from platelet-derived microparticles. Blood 2009; 113:1112–21. [16] Cognasse F, Hamzeh-Cognasse H, Laradi S, Chou ML, Seghatchian J, Burnouf T, et al. The role of microparticles in inflammation and transfusion: a concise review. Transfus Apher Sci 2015;53:159–67. [17] Hagberg IA, Akkok CA, Lyberg T, Kjeldsen-Kragh J. Apheresis-induced platelet activation: comparison of three types of cell separators. Transfusion 2000;40:182–92. [18] Rank A, Nieuwland R, Liebhardt S, Iberer M, Grutzner S, Toth B, et al. Apheresis platelet concentrates contain platelet-derived and endothelial cell-derived microparticles. Vox Sang 2011;100:179–86. [19] Holme S, Sweeney JD, Sawyer S, Elfath MD. The expression of p-selectin during collection, processing, and storage of platelet concentrates: relationship to loss of in vivo viability. Transfusion 1997;37:12–7. [20] 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. [21] Pietersz RN, Loos JA, Reesink HW. Platelet concentrates stored in plasma for 72 hours at 22 degrees C prepared from buffycoats of citrate-phosphate-dextrose blood collected in a quadruple-bag saline-adenine-glucose-mannitol system. Vox Sang 1985;49:81–5. [22] Pietersz RN, de Korte D, Reesink HW, van den Ende A, Dekker WJ, Roos D. Preparation of leukocyte-poor platelet concentrates from buffy coats. III. Effect of leukocyte contamination on storage conditions. Vox Sang 1988;55:14–20. [23] Skripchenko A, Kurtz J, Moroff G, Wagner SJ. Platelet products prepared by different methods of sedimentation undergo platelet activation differently during storage. Transfusion 2008;48:1469–77. [24] Rinder HM, Ault KA. Platelet activation and its detection during the preparation of platelets for transfusion. Transfus Med Rev 1998; 12:271–87. [25] Bock M, Rahrig S, Kunz D, Lutze G, Heim MU. Platelet concentrates derived from buffy coat and apheresis: biochemical and functional differences. Transfus Med 2002;12:317–24. [26] Fijnheer R, Pietersz RN, de Korte D, Gouwerok CW, Dekker WJ, Reesink HW, et al. Platelet activation during preparation of platelet concentrates: a comparison of the platelet-rich plasma and the buffy coat methods. Transfusion 1990;30:634–8. [27] Mallhi RS, Kumar S, Philip J. A comparative assessment of quality of platelet concentrates prepared by buffy coat poor platelet concentrate method and apheresis derived platelet concentrate method. Indian J Hematol Blood Transfus 2015;31:453–9. [28] Kaufman RM, Djulbegovic B, Gernsheimer T, Kleinman S, Tinmouth AT, Capocelli KE, et al. Platelet transfusion: a clinical practice guideline from the AABB. Ann Intern Med 2015;162:205–13. [29] British Committee for Standards in Haematology, Blood Transfusion Task Force. Guidelines for the use of platelet transfusions. Br J Haematol 2003;122:10–23.
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Transfusion and Apheresis Science j o u r n a l h o m e p a g e : w w w. e l s e v i e r. c o m / l o c a t e / t r a n s c i
Differences in levels of platelet-derived microparticles in platelet components prepared using the platelet rich plasma, buffy coat, and apheresis procedures Egarit Noulsri a,*, Prapaporn Udomwinijsilp b, Surada Lerdwana c, Viroje Chongkolwatana b, Parichart Permpikul b,** a
Research Division, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand Department of Transfusion Medicine, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand c Department of Research and Development, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand b
A R T I C L E
I N F O
Article history: Received 2 August 2016 Received in revised form 7 October 2016 Accepted 21 October 2016 Keywords: Platelet-derived microparticle Platelet component Buffy coat Platelet rich plasma Apheresis
A B S T R A C T
Background: There has been an increased interest in platelet-derived microparticles (PMPs) in transfusion medicine. Little is known about PMP status during the preparation of platelet concentrates for transfusion. Aim: The aim of this study is to compare the PMP levels in platelet components prepared using the buffy coat (BC), platelet-rich plasma platelet concentrate (PRP-PC), and apheresis (AP) processes. Methods: Platelet components were prepared using the PRP-PC and BC processes. Apheresis platelets were prepared using the Trima Accel and Amicus instruments. The samples were incubated with annexin A5-FITC, CD41-PE, and CD62P-APC. At day 1 after processing, the PMPs and activated platelets were determined using flow cytometry. Results: Both the percentage and number of PMPs were higher in platelet components prepared using the Amicus instrument (2.6 ± 1.8, 32802 ± 19036 particles/μL) than in platelet components prepared using the Trima Accel instrument (0.5 ± 0.4, 7568 ± 5298 particles/ μL), BC (1.2 ± 0.6, 12,920 ± 6426 particles/μL), and PRP-PC (0.9 ± 0.6, 10731 ± 5514 particles/ μL). Both the percentage and number of activated platelets were higher in platelet components prepared using the Amicus instrument (33.2 ± 13.9, 427553 ± 196965 cells/ μL) than in platelet components prepared using the Trima Accel instrument (16.2 ± 6.1, 211209 ± 87706 cells/μL), BC (12.9 ± 3.2, 140624 ± 41003 cells/μL), and PRP-PC (21.1 ± 6.3, 265210 ± 86257 cells/μL). Conclusions: The study suggests high variability of PMPs and activated platelets in platelet components prepared using different processes. This result may be important in validating the instruments involved in platelet blood collection and processing. © 2016 Elsevier Ltd. All rights reserved.
1. Introduction * Corresponding author. Research Division, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand. Fax: + 66-2-411-0175. E-mail address: [email protected] (E. Noulsri). ** Corresponding author. Department of Transfusion Medicine, Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand. Fax: + 66-2-412-8419. E-mail address: [email protected] (P. Permpikul).
In recent years, there has been an increased interest in cell-derived microparticles (MPs) in transfusion medicine. The MPs in blood circulation can be generated by red blood cells, endothelial cells, white blood cells (WBCs), and platelets. Upon activation or apoptosis, the platelets release
http://dx.doi.org/10.1016/j.transci.2016.10.006 1473-0502/© 2016 Elsevier Ltd. All rights reserved.
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small plasma membranes, 0.1–1 μm, or platelet-derived microparticles (PMPs). These PMPs have been shown to be the most-abundant MPs in the blood circulation. Studies have suggested that higher levels of PMPs in the blood circulation are associated with thrombosis complications and inflammation [1–3]. Upon being shed from the activated platelets, the PMPs express the phosphatidylserine (PS) that is on their surface. Negatively charged phospholipid can further activate the coagulation factor, resulting in thrombus formation. PMPs also exhibit the tissue factor on their surfaces. Upon binding with FVIIa, the TF can initiate coagulation [4]. In addition, an in-vitro study using the thrombin generation test demonstrated the coagulation activity of the PMP [5]. Several studies have documented the potential role of PMPs in cardiovascular disease. For example, Siljander et al. demonstrated that PMPs are associated with fibril formation [6]. In addition, a significant increase in PMP was reported in patients with acute pulmonary embolism [7]. Furthermore, a number of studies have suggested the role of PMPs in tumor progression [8–11], including chemotaxis activity, angiogenesis activity, and the promotion of tumor metastasis. Therefore, to minimize recipients’ complications, it is necessary to monitor the PMP in the platelet-preparation process in transfusion medicine. Platelet blood products can be prepared by various methods, including the buffy coat (BC), platelet-rich plasma platelet concentrate (PRP-PC), and apheresis (AP) methods. In the AP method, platelets are collected from a donor, and the remaining components are returned to the donor, a process mediated by AP instruments. In contrast, platelet products collected by the BC and PRP-PC processes are pooled from multiple donors. Since each cell in the blood has its own specific gravity, centrifugation is applied to separate each blood component for removal. International guidelines recommend that the half-life of platelet concentrates in currently available plateletstorage bags is 5 days at 22 ± 2 °C with continuous agitation. A number of studies have shown that platelets undergo various changes during collection, processing, and storage [12]. PMPs can be detected in platelet components prepared using either whole-blood processes or AP instruments [13–16]. Higher levels of PMPs in the blood product may result from the harvesting conditions and the storage lesion. However, the effect of AP on PMP counts remains controversial. Recent studies have suggested that some AP procedures may generate more PMPs than others [17,18]. In contrast, other studies have shown increased levels of PMPs in stored platelet concentrate and AP [19,20]. However, no studies have investigated the effects of various platelet-concentrate preparation procedures on PMP counts. Therefore, the present study aimed to determine whether various platelet-component preparation procedures affected the PMP counts. The platelet components were prepared using the AP, BC, and PRP-PC processes under routine protocol. The PMPs and activated platelets in these platelet concentrates were determined using flow cytometry. The study’s results are important to understanding which bloodproduct manufacturing steps might trigger the release of PMPs.
2. Materials and methods 2.1. Materials Fluorescein isothiocyanate conjugated annexin A5 (annexin A5-FITC), phycoerythrin conjugated CD41a (CD41aPE), allophycocyanin conjugated CD62P (CD62P-APC), and 10 × annexin A5 binding buffer were purchased from BD (San Jose, CA, USA). CountBright™ counting beads were obtained from Invitrogen (Carlsbad, CA, USA). Blank calibration particles of 1.09 μm were purchased from Spherotech (Lake Forest, IL, USA). 2.2. Subjects This study was approved by the Institutional Review Board of Siriraj Hospital, Mahidol University School of Medicine, Bangkok, Thailand (COA no. Si349/2015). Written consent of the donors was received after the procedure was explained to them in detail, including the time the study would take and its possible hazards and benefits. After obtaining this informed consent, blood samples were collected using the standard procedure for collecting donated blood. Before the platelet-component preparation procedure, ABO/Rh typing and testing for infectious disease markers (HIV, HBV, HCV, and syphilis) were conducted. The study included healthy subjects aged 18–65 without known blood dyscrasia. The exclusion criteria included a medical history of blood-derived illness or taking any medication known to affect platelet or bone marrow function or concentration within two weeks prior to the test. In addition, the study excluded donors who had taken aspirin or other nonsteroidal anti-inflammatory drugs (NSAIDS), which are likely to affect platelet function, within two weeks prior to the test. 2.3. Buffy coat platelet concentrates (BC-PC) Platelet concentrates were prepared from buffy coats derived from whole blood using a standard procedure described previously [21,22]. Briefly, 450 mL of whole blood from healthy donors was collected in 450-mL quadruple bags containing 63 mL of citrate-phosphate dextrose (CPD) as an anticoagulant. Then, the blood was centrifuged (Heraeus 6000i; Germany) at 3380 rpm at 22 °C with acceleration and deceleration curves of 3 and 5, respectively. After 10 min, the whole blood was separated into components. The top layer, 150–200 mL, was the platelet poor plasma (PPP). The middle layer was the buffy coat, containing approximately 90% of the platelets, 70% of the WBCs, and 10% of the red blood cells. The bottom layer was the pack red-blood cells. The PPP supernatant and buffy coat were transferred into other satellite bags. A solution of saline, adenine, glucose, and mannitol (SAGM) (TERUMO PENPOL, Ltd.; Puliyarakonam, Trivandrum, India) was added to the red cells as an additive. The bags containing the red cells and plasma were removed, and the red cells were stored at 4 °C in a cold room. The PPP was stored as fresh frozen plasma (FFP) in a freezer at −20 °C. Four bags of the buffy coat were gently mixed with plasma (200–250 mL) from the same blood group as the donors’ and then centrifuged at 1600 rpm for
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6 min at 22 °C along with one empty satellite bag. The supernatant platelet rich plasma (PRP) was transferred into an empty platelet-storage bag and the tubing was sealed.
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instrument was confirmed each day using two levels of controls (Sysmex e-Check Hematology Control for Sysmex X-Series Analyzers; Sysmex; USA) according to the manufacturer’s recommendations.
2.4. Platelet rich plasma-platelet concentrates (PRP-PC) 2.7. Flow cytometry analysis of PMPs and activated platelets First, 450 ml of whole blood was collected in a 450-ml triple bag containing 63 mL of CPD anticoagulant and red blood cell preservative (TERUMO PENPOL, Ltd. Puliyarakonam, Trivandrum, India). The PRP was separated from the whole blood by centrifugation (Heraeus 6000i, Germany) at 3000 rpm at 21 °C for 10 min, with acceleration and deceleration curves of 5 and 4, respectively. Then, the platelets were concentrated by centrifugation at 3940 rpm for 5 min at 21 °C, with acceleration and deceleration curves of 9 and 5, respectively, and the supernatant plasma was removed. The platelet-concentrate bag was left at room temperature for approximately 2 h before being stored at or below 22 ± 2 °C with reciprocal agitation. 2.5. Apheresis platelet (AP) For platelet apheresis, the entire process was performed in a closed system. A disposable kit was installed onto the continuous-blood cell separator before the machine was primed. The donors were prepared by cleaning the venipuncture sites at the antecubital areas, and then phlebotomy was performed with minimal trauma to the donors. For all Trima instrument collections, Gambro Trima Accel software version 5.1 (Terumo BCT; Lakewood, CO, USA) was used, with the following settings: AC management: 2; AC ratio: 1:10; minimum donor post-HCT: 32%; minimum donor post-platelet count: 100,000; maximum draw flow: moderate; draw management: 2; return management: 1; predicted yield: 4.0–9.5 × 1011/μL; final concentration: 1200– 1400 × 103/μL; and collection volume: 285–316 mL. For all Amicus instrument collections, Amicus software version 3.1 (Fresenius-Kabi Company; Lake Zurich, IL, USA) was used, with the following settings: AC infusion rate: 1.25 mg/kg/ min; AC ratio: 10:1; maximum return rate: 120 mL/min; maximum inlet rate: 100 mL/min; and maximum cycle volume: 150–300 mL. Draw and return rates were determined by the instrument unless adjustments were needed to address access or donor-comfort issues. Plateletconcentrate products were stored at 22 ± 2 °C with continuous agitation. 2.6. Measuring pH, WBCs, and platelet counts The pH of platelet products was determined using a Mettler-Toledo Delta 320 pH meter (Mettler-Toledo; Greifensee, Switzerland), and the measurement was carried out at room temperature. The pH meter’s electrode was placed in the platelet product and swirled. The results were recorded when the pH reading stabilized. The WBC and platelet counts were determined using an automated hematology analyzer, XS-800i (Sysmex Corporation; Kobe, Japan), after control setup and calibration as recommended by the manufacturer. Samples were analyzed within 30 min of collection. Calibration of the
To investigate the expression of phosphatidylserine and CD62P on MPs and platelets, 20 μL of platelet suspensions diluted at 1:1000 was incubated with 2 μl of annexin A5FITC, 2 μL of CD62P-APC, and 2 μl of CD41a-PE in the dark at room temperature. HEPES buffer without calcium was used as a negative control for annexin A5 binding. After 15 min, 25 μL of counting beads and 300 μL of 1 × annexin A5 binding buffer were added to the stained samples, which were then acquired and analyzed using FACSDiva™ software on a FACSCanto flow cytometer (BD; San Jose, CA, USA). Fig. 1 summarizes the gating strategies for flow cytometry analysis. The forward scatter (FSC), side scatter (SSC), and fluorescence (FL) parameters were set as logarithmic scale. Instrument performance was verified using CaliBRITE™ beads (BD; San Jose, CA, USA). Initially, an FSC vs. SSC dotplot was used to identify platelets, counting beads, and MPs (Fig. 1A). MP gating was performed according to 1-μm standard beads. The gated MPs were further analyzed for their percentage and number on the annexin A5 vs. SSC dotplot (Fig. 1B). To determine the percentage and number of PMPs, a CD41 vs. CD62P dot-plot was used (Fig. 1C). Likewise, the percentage and number of activated platelets from the gated platelet population were analyzed using CD41 vs. SSC (Fig. 1D) and annexin A5 vs. CD62P dot-plots (Fig. 1E). To enable reliable analysis, data were collected for at least 1300 events of counting beads at medium speed. The absolute number was calculated using the following formula: A = ((B × C)/(D × E)) × F, where A is the concentration of the cell population (cells/μL), B is the events of the cell population, C is the number of the counting beads per test (beads/ 25 μL), D is the events of the counting beads, E is the volume of the samples (μL), and F is the dilution factor. 2.8. Statistical analysis Data analysis and graphing were performed using GraphPad Prism software version 5.0 (GraphPad; San Diego, CA, USA). Results were expressed as mean and standard deviation (SD). The Mann–Whitney U test was used to determine the mean difference between the groups. A p ≤ 0.05 was considered statistically significant. 3. Results 3.1. Number of samples, donor age, collection time, and platelet-component volume in the three processes This study analyzed a total of 97 platelet components, with 19, 17, and 61 being prepared using the PRP-PC, BC, and AP processes, respectively. Of the 61 AP samples, 46 were prepared using the Trima Accel instrument and the other 15 using the Amicus instrument. The donors’ ages for the PRP-PC, BC, and AP processes had means ± SD (range) of
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Fig. 1. Flow cytometry gating strategy for analyzing PMPs and activated platelets. (A) FSC vs. SSC dot-plot represents the population of MPs based on the standard 1-μm bead, platelets, and counting beads in the defined gate. (B) Annexin A5 vs. SSC dot-plot distinguishes the annexin A5-positive MPs in the P1 region from the gated MPs. (C) CD41a vs. CD62P dot-plot distinguishes the PMPs in quadrants Q2 and Q4 from the previous P1 region. (D) CD41a vs. SSC dot-plot distinguished CD41a-positive population in the region from the gated platelets. (E) Annexin A5 vs. CD62P dot-plot distinguishes the activated platelets in quadrants Q1-1 and Q2-1 from the CD41-positive population in the previous region. The plot also shows the percentage of cells present in each quadrant. Data are representative of three independent experiments.
38.5 ± 10.8 (21–55), 36.9 ± 11.2 (20–62), and 47.6 ± 15.8 (24– 67), respectively. The collection times (min) for the three procedures were 7.1 ± 19.3 (5–12), 6.6 ± 2 (4–12), and 93.5 ± 15.8 (51–136), respectively. 3.2. Optimizing flow cytometry methods to determine PMP and activated platelet levels The first experiment aimed to optimize the PMP gating strategy. The platelet products were left at room temperature without treatment for 7 days. The levels of PMP and activated platelets were determined by flow cytometry. A 1.9-fold increase of PMPs compared to day 1 was found in stored platelet components. Similarly, a 3.3-fold increase compared to day 1 was found in CD62P-positive platelets in stored platelet components. In addition, a 16-fold increase compared to day 1 was found in the percentage of annexin A5-positive platelets in stored platelet components. Examining the activated platelets revealed that the percentage of CD62P increased time-dependently, and the highest values were found on day 6, suggesting that flow cytometry gating is optimal for future experiments.
3.3. PMPs and activated platelets in platelet components prepared using AP, BC, and PRP-PC processes Next, the levels of PMPs and activated platelets were determined in platelet components prepared using the AP, BC, and PRP-PC processes. Because the AP was prepared using two different instruments, we categorized the AP into Amicus and Trima Accel and compared the results with those of the BC and PRP-PC processes. As shown in Fig. 2A, in platelet components prepared using the Amicus instrument (2.6 ± 1.8), the percentage of PMPs at day 1 was significantly higher than the percentage in platelet components prepared using the Trima Accel instrument (0.5 ± 0.4), BC (1.2 ± 0.6), and PRP-PC (0.9 ± 0.6). Although the percentage of PMPs in platelet components prepared using BC was slightly higher than the percentage in platelet components prepared using PRP-PC, this difference was not statistically significant. Further experiments revealed that the number of PMPs in platelet components prepared using the Amicus instrument (32,802 ± 19,036 particles/μL) was significantly higher than the number in platelet components prepared using the Trima Accel instrument
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Fig. 2. Comparison of PMPs and activated platelets in platelet concentrate prepared using various procedures. The scatter plots show the percentage (A) and the number (B) of the PMPs and the percentage (C) and the number (D) of the activated platelets in platelet components prepared using the Amicus, Trima Accel, BC, and PRP-PC processes. A p value indicates significant differences among the groups.
(7568 ± 5298 particles/μL), BC (12,920 ± 6426 particles/ μL), and PRP-PC (10,731 ± 5514 particles/μL) (Fig. 2B). Similar to the percentage, the number of PMPs in the platelet components prepared using BC was slightly higher than the number in platelet components prepared using PRP-PC, but this difference was not statistically significant. Examining the activated platelets showed a significant increase in the percentage of CD62P-positive platelets in components prepared using the Amicus instrument (33.2 ± 13.9) compared to the percentage in platelet components prepared using the Trima Accel instrument (16.2 ± 6.1), BC (12.9 ± 3.2), and PRP-PC (21.1 ± 6.3) (Fig. 2C). Similarly, the number of CD62Ppositive platelets in platelet components prepared using the Amicus instrument (427,553 ± 196,965 cells/μL) was higher than the number in platelet components prepared using the Trima instrument (211,209 ± 87,706 cells/μL), BC (140,624 ± 41,003 cells/μL), and PRP-PC (265,210 ± 86,257 cells/μL) (Fig. 2D).
3.4. Comparing platelet count, leukocyte count, volume, and pH of platelet components prepared using AP, BC, and PRPPC processes Recent experiments have shown high variability in PMP levels in platelet components prepared using the AP process. We checked whether the platelet and leukocyte count and pH might affect the level of PMP formation. As shown in Fig. 3A, the number of platelets in platelet components prepared using the Amicus instrument (1581 × 103 ± 122 cells/ μL) did not differ from the number in platelet components prepared using the Trima Accel instrument (1595 × 103 ± 191 cells/μL). The highest number of platelets was found in platelet components prepared using PRP-PC (1671 × 103 ± 452 cells/μL), but the difference was not statistically significant when compared to platelet components prepared using BC (1409 × 103 ± 215 cells/μL). The pH in the platelet components at day 1 after preparation was measured, and the
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Fig. 3. The platelet, pH, and leukocyte numbers in platelet concentrate prepared using various procedures. The scatter plots represent the number of platelets (A), pH (B), and the number of leukocytes (C) in platelet component prepared using the Amicus, Trima Accel, BC, and PRP-PC processes. A p value indicates significant differences among the groups.
results showed that the pH in the platelet components prepared using the Amicus instrument (7.1 ± 0.2) was significantly lower than the pH in the platelet components prepared using the Trima Accel instrument (7.5 ± 0.1), BC (7.2 ± 0.1), and PRP-PC (7.5 ± 0.1) (Fig. 3B). In addition, the results showed that the number of leukocytes in platelet components prepared using PRP-PC (1077 ± 615 cells/ μL) was significantly higher than the number in platelet components prepared using the Amicus instrument (96 ± 52 cells/μL), the Trima Accel instrument (20 ± 34 cells/μL), and BC (209 ± 99 cells/μL) (Fig. 3C). 4. Discussion Platelet concentrate is crucial to correcting bleeding tendencies in transfusion medicine patients. Platelet components can be prepared by various processes, including AP, BC, and PRP-PC. Recent studies have suggested that the preparation method may affect the quality of the platelets, as some studies have found increased levels of activated platelets and PMPs [14,16]. Higher PMP levels have been shown to be associated with thrombosis complications, tumor progression, and metastasis [1–3]. To minimize these complications in recipients, it is essential to validate the platelet-concentrate preparation processes in individual transfusion laboratories. The present study determined the PMP levels in platelet concentrates prepared using the AP, BC, and PRP-PC procedures, clearly demonstrating that the AP instrument used affects the PMP count, as shown by the significant increase in the number of PMPs in the platelet component prepared using various AP instruments compared to the numbers in platelet components prepared using the BC and PRP-PC processes. In addition, both the percentage and number of PMPs showed a greater range of values in the AP prepared using various AP instruments than in those prepared using the BC and PRP-PC processes. Further experiments conducted during the present study supported this result by showing increases in both PMPs and activated platelets in AP prepared using the Amicus instrument compared to that prepared using the Trima Accel
instrument. These results agree with those of previous studies that have shown differences in AP product prepared using the Amicus and Trima Accel instruments. Those studies found higher amounts of CD62P-positive platelets in AP prepared using the Amicus instrument than in that prepared using the Trima Accel instrument [17]. This is likely due to several reasons [19,23]. When preparing AP using the Amicus instrument, the platelets directly contact the chamber wall. In contrast, when AP is prepared using the Trima Accel instrument, the buffy coat prevents platelet contact with the chamber wall. In addition, in the final Amicus process, plasma is drawn into the bag containing the platelets, which may cause platelet activation via shear stress. The present study’s findings are important for the following reasons. To meet the expectations of transfusion medicine, it is essential to consider the quality of the platelet products when evaluating automated blood-processing systems or platelet-preparation procedures. Since there are a number of AP instruments on the market, laboratory tests must be established to assess the quality of these instruments. PMP enumeration by flow cytometry is recommended to test the quality of the screening of AP batches. Several studies have reported correlations between increased PMP levels in blood products and the quality of those products [12,24–27]. Studies of platelet recovery and survival have shown low post-transfusion platelet recovery and short platelet survival in platelet products with high P-selectin expression. Therefore, the levels of PMPs and activated platelets correlate with the quality of the AP, and monitoring these levels may help assess the quality of the AP equipment used. However, the possible clinical repercussions that may be caused by differences in recovery and survival characteristics remain unknown. Further studies should address these post-transfusion characteristics associated with platelet products. Results of the present study showed differences in the pH, platelet number, and leukocyte contamination. However, the differences in these parameters were within the ranges allowed by recommended guidelines, and several studies
Please cite this article in press as: Egarit Noulsri, Prapaporn Udomwinijsilp, Surada Lerdwana, Viroje Chongkolwatana, Parichart Permpikul, Differences in levels of platelet-derived microparticles in platelet components prepared using the platelet rich plasma, buffy coat, and apheresis procedures, Transfusion and Apheresis Science (2016), doi: 10.1016/j.transci.2016.10.006
ARTICLE IN PRESS E. Noulsri et al. / Transfusion and Apheresis Science ■■ (2016) ■■–■■
have found that such differences have no effect on platelet quality [28,29]. In addition, the present study’s findings agreed with the results of previous studies on the quality of platelets in platelet components [14,20]. These differences in pH, platelet, and leukocyte numbers may be explained by differences in centrifugation speed, storage bags, and storage conditions among transfusion laboratories [20]. One might suggest that the differences observed in the present study resulted from the storage lesion of the platelets. The present study compared the levels of PMPs and activated platelet components on day 1 after processing. However, previous studies have suggested that the storage lesion of the blood component begins after day 5 [20]. Therefore, the storage lesion cannot be the explanation for the difference noted in the present study. 5. Conclusions The present study found various levels of PMPs and activated platelets in platelet components prepared using the AP, BC, and PRP-PC procedures. In addition, the results documented the effects of the various instruments used in the AP method on the levels of PMPs and activated platelets. This data on PMPs in platelet components may be useful to assessing platelet quality and preparation procedures. Acknowledgments The authors thank the Faculty of Medicine Siriraj Hospital, Mahidol University, Bangkok, Thailand, for supporting this research project. References [1] Solum NO. Procoagulant expression in platelets and defects leading to clinical disorders. Arterioscler Thromb Vasc Biol 1999;19:2841–6. [2] Jy W, Mao WW, Horstman L, Tao J, Ahn YS. Platelet microparticles bind, activate and aggregate neutrophils in vitro. Blood Cells Mol Dis 1995;21:217–31, discussion 31a. [3] Herring JM, McMichael MA, Smith SA. Microparticles in health and disease. J Vet Intern Med 2013;27:1020–33. [4] Puddu P, Puddu GM, Cravero E, Muscari S, Muscari A. The involvement of circulating microparticles in inflammation, coagulation and cardiovascular diseases. Can J Cardiol 2010;26:140–5. [5] Diamant M, Tushuizen ME, Sturk A, Nieuwland R. Cellular microparticles: new players in the field of vascular disease? Eur J Clin Invest 2004;34:392–401. [6] Siljander P, Carpen O, Lassila R. Platelet-derived microparticles associate with fibrin during thrombosis. Blood 1996;87:4651–63. [7] Bal L, Ederhy S, Di Angelantonio E, Toti F, Zobairi F, Dufaitre G, et al. Circulating procoagulant microparticles in acute pulmonary embolism: a case-control study. Int J Cardiol 2010;145:321–2. [8] Goubran H, Sabry W, Kotb R, Seghatchian J, Burnouf T. Platelet microparticles and cancer: an intimate cross-talk. Transfus Apher Sci 2015;53:168–72. [9] Aharon A, Brenner B. Microparticles, thrombosis and cancer. Best Pract Res Clin Haematol 2009;22:61–9.
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Please cite this article in press as: Egarit Noulsri, Prapaporn Udomwinijsilp, Surada Lerdwana, Viroje Chongkolwatana, Parichart Permpikul, Differences in levels of platelet-derived microparticles in platelet components prepared using the platelet rich plasma, buffy coat, and apheresis procedures, Transfusion and Apheresis Science (2016), doi: 10.1016/j.transci.2016.10.006