Platelet function testing in apheresis products: flow cytometric, resonance thrombographic (RTG) and rotational thrombelastographic (roTEG) analyses

Transfusion and Apheresis Science 26 (2002) 147–155 www.elsevier.com/locate/transci

Platelet function testing in apheresis products: flow cytometric, resonance thrombographic (RTG) and rotational thrombelastographic (roTEG) analyses q K. Gutensohn a

a,* ,

K. Geidel a, N. Kroeger b, B. Eifrig c, N. Crespeigne d, P. Kuehnl a

Department of Transfusion Medicine/Transplantation Immunology, University Hospital Eppendorf, University of Hamburg, Martinistrasse 52, 20246 Hamburg, Germany b Department of Bone Marrow Transplantation, University Hospital Eppendorf, University of Hamburg, Martinistrasse 52, 20246 Hamburg, Germany c Department of Oncology/Hematology, University Hospital Eppendorf, University of Hamburg, Martinistrasse 52, 20246 Hamburg, Germany d Baxter Research and Development Europe, Fenwal Division, Nivelles, Belgium Received 21 September 2001; accepted 24 September 2001

Abstract During storage of platelet concentrates, quality control of the units is mandatory. This includes the important testing of the hemostatic function of platelets. So far, mostly platelet aggregation analyses have been performed. In this study, new approaches were tested to evaluate the applicability of modern techniques for quality monitoring. Plateletpheresis was performed with two different cell separators (AMICUSe cell separator, Fenwal, Baxter Healthcare, Deerfield, USA; COBE Spectrae, COBE BCT, Lakewood, USA). In each procedure split products ðn ¼ 22Þ were prepared and stored for 1–2 days ðn ¼ 22Þ or 3–5 days ðn ¼ 22Þ. Platelet hemostatic capacity was tested by applying flow cytometry, platelet aggregation (platelet-rich-plasma [PRP] + agonist), resonance thrombography (RTG; PRP, no agonist) and rotational thrombelastography (roTEG; PRP+agonist). Flow cytometric analyses did not reveal significant changes in structural (CD41a, CD42b) or activation-dependent antigens (CD62p, CD63, LIBS, RIBS). Also, differences in the data from the flow cytometric reactivity tests were not significant between the two groups. In platelet aggregation assays, shape change ðp ¼ 0:8Þ, maximum aggregation ðp ¼ 0:4Þ, and maximum gradient ðp ¼ 0:8Þ did not show significant differences between the two groups. In the RTG test, differences between r-time (reaction time; p ¼ 0:4), and f time (clot formation time [fibrin influence]; p ¼ 0:3), and in roTEG r-time (coagulation time; p ¼ 0:1) and k-time (clot formation time; p ¼ 1:0) were not significant. P-time (clot formation time [platelet influence]) and M (maximum amplitude) in RTG, and k-time and MA (maximum amplitude) in roTEG showed a slight decrease in platelet function ðp 6 0:05Þ. We conclude that platelet function is well maintained during storage. This is reflected by the results of immunological and platelet function assays. Rotational thrombelastography (in the case of PRP) and especially

q

This study was supported by a grant from Baxter Deutschland GmbH, Biotech Group Europe, Munich, Germany. Corresponding author. Tel.: +49-40-42803-5340; fax: +49-40-42803-5213. E-mail address: [email protected] (K. Gutensohn).

*

1473-0502/02/$ - see front matter Ó 2002 Elsevier Science Ltd. All rights reserved. PII: S 1 4 7 3 - 0 5 0 2 ( 0 2 ) 0 0 0 0 7 - 1

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resonance thrombography represent promising methods for quality control of platelet concentrates and rapidly provide information about the status of platelet function and the whole clotting process. Ó 2002 Elsevier Science Ltd. All rights reserved. Keywords: Platelets; Storage; Flow cytometry; Resonance thrombography; Rotational thrombelastography

1. Introduction

2. Materials and methods

The application of platelet concentrates has steadily increased over the last few years [1–3]. Many products are not transfused immediately and therefore have to be stored. During this period platelets continuously deteriorate, a process that is also referred to as platelet storage lesion. The progressive reduction of quality is reflected by morphological and functional changes [4–6]. For this reason, and to ensure a product with an adequate hemostatic capacity at the time of transfusion, quality control of the platelet units is performed; this is recommended in several guidelines [7,8]. Many different approaches exist to measure the condition of platelets during storage [9]. They include methods such as morphological scores, the swirling effect, platelet count, analyses of release products like LDH, PF-4, serotonin and bthromboglobulin, and analyses of metabolic changes like pH or lactate production. Several suggestions have been made and discussed [10]. Apart from tests that are applicable for routine testing, further methods have been discussed to achieve a more detailed insight into the platelet condition. For this purpose analyses like electron microscopy and flow cytometry may be applied. Also of importance are tests that reflect the hemostatic function of platelets; the most common of these up to now have been platelet aggregation analyses [10,11]. In this study, the expression of platelet antigens and the binding of fibrinogen were monitored by flow cytometry in concentrates obtained by two different cell separators. Furthermore, platelet function testing was performed using flow cytometry, platelet aggregation, resonance thrombography and rotational thrombelastography to evaluate the applicability of these tests for quality control of platelet concentrates during storage.

2.1. Cytapheresis Plateletpheresis was performed with two blood cell separators (AMICUSe cell separator, Fenwal, Baxter Healthcare, Deerfield, USA; COBE Spectrae, COBE BCT, Lakewood, USA). A doubleneedle continuous-flow procedure was performed with both devices, and in each procedure split products were prepared. For the AMICUSe blood cell separator (software version 2.37) the yield predictor was used to determine the whole blood volume to be processed, the number of platelets and the plasma volume to be collected to reach the targeted yield of 5:8  1011 platelets ð2:9  1011 platelets for each split product). For the COBE Spectrae apheresis system the software-controlled collection mode (5.1 LRSe) was applied to reach the same target yield. For all procedures ACD-A (Baxter Healthcare) was used in a ratio of 1:9 (1:8–1:9 for COBE Spectrae). All donors fulfilled the requirements for blood donation, were healthy non-smokers, and had not taken any type of drug for at least four weeks prior to the apheresis procedure [7]. Cytapheresis was performed two times in each donor. Donors were randomly allocated to either the Baxter or the COBE device and vice versa in the second procedure following a time interval of at least two weeks.

2.2. Storage of platelet concentrates One of the split products was stored for a short time (1–2 days; n ¼ 22), and the other for a period of 3–5 days ðn ¼ 22Þ. Following apheresis, platelet concentrates (PC) obtained with the AMICUSe device were stored in polyolefin-based plastic containers (PL-2410, Baxter). Following apheresis,

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AMICUSe products had to rest for a minimum of 2 h before being agitated. The PCs obtained with the COBE Spectrae were stored in polyvinylchloride containers (COBE). COBE platelet concentrates were agitated immediately after collection. All PCs were stored on a horizontal flatbed agitator (LPR1, Melco Engineering, Glendale USA) with an agitation rate of 60 shakes/min and a constant temperature of 22  2°C (ICE 800, Memmert, Schwabach, Germany). In this study, the day of apheresis was regarded as day 0.

2.3. Sample collection For in vitro testing, samples were taken from the PCs. The initial 2 ml were discarded to avoid the collection of activated platelets [12]. For hematological analyses K2-EDTA, for hemostasis tests citrate (0.106 mol/l) were used as anticoagulants in ready-to-use containers (Sarstedt, Nuembrecht, Germany). Hematological analyses were performed on a Coulter counter (STKRâ , Beckman–Coulter, Glendale, USA). For flow cytometry platelet-rich-plasma was directly collected into polypropylene tubes (Greiner, Hamburg, Germany) without further centrifugation steps. Samples were fixed and stabilized immediately as described previously [13]. Briefly, fixation was performed with 0.15 M phosphate-buffered saline (PBS; Gibco BRL, Eggenstein, Germany) containing 0.2% w/v glyoxal (Merck, Darmstadt, Germany) and 0.4% w/v paraformaldehyde (Merck). After an incubation time of 10 min at room temperature, the samples were stabilized [14]. For this purpose, a 1:10 dilution with PBS containing 0.2% w/v glycine (Serva, Heidelberg, Germany) was used. 2.4. Flow cytometry An aliquot of 100 ll platelet-rich-plasma was directly labeled with monoclonal antibodies. For analyses, CD41a, CD42b, CD62p (P-selectin), and CD63 were used in saturating concentrations (Beckman–Coulter). IgG isotype controls were applied to detect non-specific staining (Beckman– Coulter). PAC-1 was applied to analyze the ligand-

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induced binding site (LIBS) of GPIIb–IIIa (Becton Dickinson, Mountain View, USA) [15]. To detect fibrinogen bound to GPIIb–IIIa, an FITC-labeled antibody and an FITC–chicken–anti–human IgG control were used (biopool, Ume
a, Sweden) [16]. CD41a was used as tagging antibody to detect CD62p- and CD63-positive platelets, and CD42b for the detection of PAC-1 and fibrinogen binding. After incubation (30 min,20  2 °C) the samples were resuspended in 1 ml of PBS and stored at þ4 °C for a maximum of 120 min prior to flow cytometric analysis. For flow cytometric reactivity testing 10 ll of ADP (final concentration 1 lmol=l; Boehringer, Mannheim, Germany) were added to 500 ll platelet-rich-plasma (PRP; 250,000 plt/ll) from the sample and incubated for 2 min before fixation and stabilization of the sample. Samples were analyzed on a FACScanâ cytometer (Becton Dickinson). Fluorescent beads were applied daily to ensure the stability of the system (CaliBRITEe, Becton Dickinson). Following the setting of the appropriate threshold in the FSC, 10,000 events were acquired in a life-gate. Listmode data were acquired and analyzed using CELLQuestâ software (Becton Dickinson). The results were expressed in arbitrary units as mean channel fluorescence intensity (MCFI), or percentage of antibody-positive cells. For MCFI, positivity for a specific antibody was defined as fluorescence higher than that of the isotype control, calculated by subtracting the MCFI of the non-specific IgG isotype control from specific MCFI [12]. Percentage of antibody-positive cells was defined as cells with specific fluorescence higher than the isotype and autofluorescence samples. 2.5. Platelet aggregation assays Aliquots of 2 ml of the PC were centrifuged (700g) for 10 min at room temperature. Platelet count was adjusted by dilution with platelet-poorplasma (PPP) to 250,000 plt/ll. Cartridges for single-use (Laborger€ate GmbH, Hamburg, Germany) were placed into the aggregometer (APACT Aggregometer, Laborger€ate GmbH) and filled with 500 ll of the sample each. 500 ll of (PPP) were used as negative control. Then, 10 ll 1 lM

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ADP (Boehringer) were added to each cartridge and analyses were started. Automatic evaluation of the measurement was performed with a software program (APACT Software 1.0, PASS Engineering, Bonn, Germany). 2.6. Resonance thrombography (RTG) An RTG device (CS-3; Heinrich Amelung, Lemgo, Germany) was used for platelet function testing. PRP was diluted with NaCl (0.9%) to 250,000 plt=ll. 250 ll of the sample and 150 ll CaCl2 (0.025 mol/l; Sigma–Aldrich, Deisenhofen, Germany) were pipetted into the cups before the automatically controlled analysis was started. Thereafter, data evaluation was performed with the RTG software (Heinrich Amelung). The parameters used for statistical evaluation were the r-time (reaction time), f -time (clot formation time [fibrin influence]), p-time (clot formation time [platelet influence]) and M (maximum amplitude). With this analysis the elasticity of the thrombus is measured. The result of the orbitometer is a graphic output, a curve divided into different time periods and the maximum amplitude of the curve. The analysis starts with the r-time, during which contact activation takes place and the coagulation process starts. The f -time reflects the growing of the fibrin structures and the clot resulting in the largest amplitude of the analysis curve, the maximum clot strength of the clot or M. Hereafter, the p-time follows and mainly demonstrates the platelet function and retraction of the clot. The shorter the p-time, the better the platelet function. 2.7. Rotational thrombelastography (roTEG) For rotational thrombelastography, a roTEG Coagulation Analyzer (Dynabyte, Munich, Germany) was used. 300 ll (280 ll of PPP and 20 ll of PRP) were mixed with 20 ll CaCl2 (0.2 M; Dynabyte). Additionally, 20 ll of an agonist were added (InTEG: Dapptin, kaolin/sulfatide–phospholipid, Immuno, Heidelberg, Germany). Parameters used in roTEG analyses were r-time (coagulation time), k-time (clot formation time),

and MA (maximum amplitude). Measurements and data analysis were performed using the integrated software (Dynabyte). With this test the motion of a pin fixed on a rotating shaft is monitored. The pin is surrounded by an outer single-use cup containing the blood sample. When clotting starts, the displacement of the pin is being registered. The rigidity of the clot, comprised of platelet aggregates and fibrin strands, is measured over a period of time. Hereby, an insight into the whole clotting process becomes possible. Like in RTG, the initial time from the onset of the analysis until the initiation of the coagulation process is considered as the reaction time. The time measured from the onset of the coagulation until the time point where an amplitude angle of 20 mm is reached is the coagulation time (k-time). Finally, the firmness of the clot is reflected by the maximum amplitude (MA). Regarding the analysis of platelet concentrates a short r-time, a short k-time and a high MA demonstrate a good reaction of platelets, their capability to initiate a clot, and also their ability to form a firm clot.

3. Statistical analysis For statistical analyses SAS software version 6.12 (SAS, Cary, USA), Statviewâ 4.5 (Abacus Concepts, Berkeley, CA, USA), Excelâ (Microsoft, Redmond, WA, USA), and SPSSâ software (SPSS, Chicago, IL, USA) were used. Data cannot throughout be regarded as being normally distributed (Kolmogorov–Smirnov-test). Therefore, non-parametric tests were constantly applied. For comparison of paired data (dependent observations) the Wilcoxon-signed-rank-test was used. Analyses of data in independent groups were made using the Wilcoxon–Mann–Whitney-test. Descriptive statistics are shown as meanSD, or as range (min, max). P -values were based on the exact permutational distribution of the test statistics rather than using a normal approximation. A pvalue of 6 0.05 was considered to be statistically significant. Since the underlying investigations are of an explorative nature, any kind of alpha adjustment was omitted.

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4. Results 4.1. Plateletpheresis The four female donors ranged in age from 31 to 49 years (mean: 40.5 years), the seven male donors from 29 to 54 years (mean: 40.9 years). No sideeffects occurred, and a total of 22 completed runs could be evaluated. Donation time with the AMICUSe cell separator ðn ¼ 11Þ was on average 64 min (range 43–85 min), whole blood flow rate 63.9 ml/min (range 50–70 ml/min), whole blood volume processed 3614.9 ml (range 2220.0–4354.0 ml), and anticoagulant used 379.8 ml (range 272.0–497.0 ml). For the COBE Spectrae ðn ¼ 11Þ donation time was on average 83 min (range 68–100 min), whole blood flow rate was 56.1 ml/min (range 47.1– 66.7 ml/min), whole blood volume processed 4299.6 ml (range 3510.0–5260.0 ml), and anticoagulant used 406.2 ml (range 330.0–495.0 ml). 4.2. Platelet concentrates and storage A total of 44 PCs could be evaluated (22 split products). For the platelet concentrates stored for

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1–2 days (1:1  0:2 days [mean  1SD]) the weight was 212:1  13:7 g, platelet content 1391  279  103 =ll; MPV 6:9  0:9, and pH 7:1  0:1 (before transfusion). For the platelet concentrates stored for 3–5 days (3:5  0:7; p 6 0:05 [p-value compared to PCs stored for 1–2 days]) the weight was 209:0  13:0 mg ðp ¼ 0:3Þ, platelet content 1462  276  103 =ll ðp ¼ 0:1Þ; MPV 6:7  0:9; p ¼ 0:02Þ, and pH 7:5  0:1 (before transfusion ½p ¼ 0:03). 4.3. Flow cytometric analyses during storage The results for the activation-dependent epitopes CD62p and CD63 were higher in the PCs stored for the longer period (MCFI and %). Fibrinogen binding and PAC-1 (%) were also higher in this group. However, differences between the two groups were not significant (Table 1). The potential of platelets to react to an agonist was slightly decreased in the PCs with the longer storage time when flow cytometric reactivity testing was performed. This trend was seen for the release reaction as well as the conformational changes of GPIIb–IIIa. For these analyses, too, no

Table 1 Parameter

PC f

PC s

p-value PC f vs PC s

Flow cytometry CD41a (MCFI) CD42b (MCFI) CD62p (MCFI) CD62p (%) CD63 (MCFI) CD63 (%) Anti-fibrinogen (MCFI) Anti-fibrinogen (%) PAC-1 (MCFI) PAC-1 (%)

256.6  85.3 13.5  3.1 23.2  6.5 4.1  5.6 21.2  7.4 3.2  4.4 14.7  5.2 1.2  0.1 16.8  5.0 2.1  2.3

234.2  64.8 15.0  4.5 25.1  9.2 5.9  8.4 23.4  7.5 4.8  6.8 16.5  5.6 4.6  10.4 15.5  3.9 2.8  3.2

0.8 0.1 0.1 0.09 0.3 0.2 0.3 0.3 0.2 0.8

Flow cytometric platelet reactivity testing CD62p (MCFI) + ADP 37.8  18.3 CD62p (%) + ADP 8.0  7.0 CD63 (MCFI) + ADP 37.5  13.9 CD63 (%) + ADP 8.4  7.5 Anti-fibrinogen (MCFI) + ADP 19.3  9.3 Anti-fibrinogen (%) + ADP 5.8  7.6 PAC-1 (MCFI) + ADP 27.7  17.4 PAC-1 (%) + ADP 8.4  8.1

36.3  18.1 9.3  9.2 33.7  11.9 8.2  7.6 22.3  11.9 8.3  10.8 23.1  11.0 6.9  6.6

0.6 0.3 0.5 0.6 0.4 0.4 0.3 0.1

PC ¼ platelet concentrate; f ¼ PC stored for 1–2 days; s ¼ PC stored for 3–5 days.

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significant differences were found when data were compared with the results from the PCs that were stored for only a short time period (Table 1). 4.4. Platelet aggregation, resonance thrombography, rotational thrombelastography The results of the platelet aggregation assays showed that the shape change in the group of PCs stored for 1–2 days was very high due to one outlying value (Table 2). However, even when data from this test (for shape change, max. aggregation and max. gradient) were eliminated no significant changes were detected. Also, for RTG and roTEG analyses the comparison of the data of both groups was not marked (Table 2). For p-time and M (RTG), and MA (roTEG) p-values were statistically different. However, RTG analyses roTEG results showed an excellent maintenance of the capacity of platelets to aggregate and to form a firm clot.

5. Discussion The present study was performed to analyze the degree of antigenic and functional alterations in platelets during storage of platelet products prepared by apheresis. Our results demonstrate that only minor immunological changes and also only

a minimal reduction of the functional capacity of platelets occurred. Platelets are essential in staunching and preventing hemorrhage. Over the last decades, their use in transfusion practice has risen considerably, mainly as a consequence of an increasingly intensive chemo- and radiotherapeutic treatment of patients with hematological and oncological malignancies [17,18]. Until transfusion, platelet concentrates have to be stored. During this period various changes occur in the platelets, collectively referred to as the platelet storage lesion [4]. Over the past few years, there has been a lot of progress in the routine preparation and storage conditions of platelets, aimed at preserving their proper hemostatic potential [5,6]. However, there are still biochemical and physical factors that influence platelets resulting in functional, immunological, and morphological changes [5,6]. Until the end of the storage period, the platelet unit has to be maintained in a condition in which it is capable of effective hemostasis. Many different approaches have been established to monitor the quality of the platelet concentrates during storage. However, an in vitro method that accurately allows the prediction of in vivo efficacy is still lacking [19]. By using monoclonal antibodies against specific membrane epitopes, flow cytometry has been shown to be a useful technique to investigate alterations in antigenic determinants [20]. One of

Table 2 Platelet function analyses Parameter

PC f

Platelet aggregation ADP/shape change ADP/max. aggregation ADP/max. gradient

PC s

176.0  422.5 53.3  23.1 107.6  48.0

40.5  25.2 28.2  18.5 98.1  33.4

0.8 0.4 0.8

RTG r-time f -time p-time M

406.6  74.6 80.3  18.0 101.6  35.6 36.4  6.5

409.3  180.3 107.4  70.6 84.5  25.4 29.6  6.9

0.4 0.3 0.03 0.02

roTEG r (InTEG) k (InTEG) MA (InTEG)

108.1  24.7 89.8  23.3 53.4  5.9

99.2  35.3 85.6  21.0 56.6  4.0

0.1 1.0 0.04

PC ¼ platelet concentrate; f ¼ PC stored for 1–2 days; s ¼ PC stored for 3–5 days.

p-value PC f vs PC s

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the major advantages of flow cytometry is that this multiparameter technique allows analysis of platelet antigens at the single cell level, associated changes, and the sensitive detection of subpopulations [21,22]. Measurement of antigens is of interest for monitoring quality of the product during storage and is also of physiological interest as these structures are important for the proper in vivo function of platelets [23,24]. It has been shown that the expression of P-selectin is associated with a reduction of in vivo circulation of platelets after transfusion [25,26]. The sequestration is probably mediated by binding of activated platelets to white blood cells and subsequent clearance, or by entrapment in the microcirculation [27,28]. GPIb–V–IX represents the binding site for von Willebrand factor, collagen, and other mediators, thereby promoting platelet adhesion, while the heterodimer GPIIb–IIIa mediates cell– matrix and cell–cell interactions [23,29,30]. In this study structural antigens and also activation-dependent epitopes were analyzed. Although a trend towards activation is apparent statistically no significant differences could be found between the two groups of platelet concentrates. This may be due to the fact that the allowable maximum storage time of five days was not completely exploited. On the other hand, the lack of significant changes may also reflect the stable conditions of PCs during storage in newer storage containers. This could be shown in our last study in the case of polyolefin bags [31]. This form of container is highly permeable to oxygen and carbon dioxide, an important requirement to reduce the platelet storage lesion [32,33]. In addition, it is essentially free of plasticizers which may leach into the blood and activate platelets [34–36]. The results of the regular antigenic studies are reflected by the tests to evaluate platelet reactivity by flow cytometry. In these investigations, too, no major deterioration of platelets can be detected during storage in terms of their reaction to addition of an agonist. Platelet aggregation represents a classical in vitro approach to test platelet function. A reduced degree of reactivity can be detected following production, further steps of preparation and during storage [11]. In our experiments a deterioration of platelet function can be assumed. However, the in-

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terpretation of the results of aggregation assays must be looked upon critically as such measurements have major limitations [12,37]. These analyses only reflect the overall behavior of platelets. However, it is the single cell that reacts interactively to a variety of different stimuli in vitro. Also, the addition of an agonist is not a desirable approach for testing platelets stored ex vivo; in our opinion any unnecessary manipulation such as the addition of agonists should be avoided. An additional problem is that the reaction is dependent on the concentration of the agonist(s) and on their combination. The limitations of aggregation assays have been discussed by several authors [38– 41]. In addition, previous studies have not shown that platelet aggregation assay results obtained in vitro reflect the in vivo behavior of transfused platelets [42,43]. A diminished reaction in vitro is thought to be due to factors such as a partial refractoriness of receptors, predominantly of GPIIb–IIIa [44]. This could not be detected in our study, as platelets were still able to express ligandand receptor-induced binding sites following storage as measured by flow cytometry. In addition, following transfusion, a recovery of the platelets in vivo seems to occur [38,44]. Aggregation assays are therefore of limited use in quality control. Another approach to test fluid phase coagulation in combination with platelet function is thrombelastography. This technique was first developed by Hartert [45]. Recently, an approach towards standardization of the method has been introduced with an improvement of the technique, single-use cups and ready-to-use reagents. So far thrombelastography represented more a form of pattern recognition than a quantitative technique but with the new analyzer quantitation of the results has been achieved. The TEG is a global assay. During this analysis the interaction of platelets, coagulation factors, fibrin, fibrinolytic and further factors is analyzed and continuously registered. Thus, a detailed insight into the coagulation kinetic and the entire process of clot formation, stabilization and retraction is provided. Although several parameters are acquired, platelet function is believed to be reflected predominantly by the maximum amplitude [46]. However, the influence of the fibrinogen concentration has also to be

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considered when looking at this complex parameter. The results of our study demonstrate that the MA is well maintained in the stored platelets and that the reaction times to form the clot are comparable in both groups. Thus, the results reflect the preservation of platelet integrity and confirm the results of the flow cytometric approach. Although in this test, as for aggregation assays, a manipulation of the platelets occurs by adding an agonist to the platelet-rich-plasma, thrombelastography is easier to perform than aggregation assays. A further advantage is the global assessment of hemostasis. With resonance thrombelastography, more extended information is obtained. The curves allow a detailed insight into the clotting process for contact activation, for the fibrin formation, the phase of platelet function and retraction and the firmness of the clot [47]. Like in RTG the whole clotting process is monitored starting with the prephase of coagulation, continuing with information on the clot formation, the demonstration of the maximum firmness of the clot, the platelet capacity to stabilize the clot until the retraction. In this test, no manipulation of the platelet-rich-plasma occurs as no agonists are added to the sample. Our results demonstrate that platelet function is slightly reduced in platelets stored for 3–5 days. However, the results still indicate that the ability to form a plug is not lost and that hemostatic capacity is well preserved. We conclude that platelet function is well maintained during storage. This is reflected by the results of immunological and platelet function assays. Flow cytometry represents one of the most sensitive and specific methods to monitor quality during storage [4,48,49]. It may therefore be applied for the detection of antigenic alterations during storage [5,6,36]. However, although functional analysis is possible with this approach, other tests may be advantageous, easier to perform and easier to standardize for routine screening. Rotational thrombography and, even more so, resonance thrombography seem to represent promising methods for quality monitoring during storage of platelet concentrates and rapidly provide information about the status of platelet function and the whole clotting process.

Acknowledgements The authors are grateful for the scientific support of Dr. J. Schulz (Laborger€ate Corp.) and Dr. A. Calatzis (Dynabyte Corp.).

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