The risk of bleeding complications is high in patients with bone marrow failure and severe thrombocytopenia. Although supportive evidence is scarce,1 prophylactic platelet transfusion is routinely performed in these patients before vascular and surgical interventions.2–5
The threshold platelet count and the timing of transfusion remain controversial among clinical studies.6–8 Moreover, many patients suffer from bleeding despite prophylactic platelet transfusion. Therefore, platelet function testing could be important in assessing the therapeutic responses to platelet transfusions. A few studies have addressed this issue,9–12 but to our knowledge, no study has comprehensively evaluated the time-dependence of platelet function after transfusion.
It seems obvious that platelet transfusions enhance both primary and secondary hemostasis by increasing the number of platelets. However, some studies indicate that platelet function is diminished in patients with hematological diseases,13–15 and that enhanced coagulation observed after platelet transfusion are only partly explained by increases in platelet count.9 To further clarify the impact of platelet transfusion, we performed a prospective observational study on patients with bone marrow failure who were scheduled for prophylactic platelet transfusion before the insertion of a central venous catheter (CVC). The aim was to measure the coagulation enhancement at 1 and 4 hours after transfusion using thromboelastometry (ROTEM®; EXTEM® and FIBTEM®), multiple electrode aggregometry (MEA) (Multiplate®; adenosine diphosphate [ADP], collagen [COL] and thrombin receptor agonist peptide [TRAP], and flow cytometry.)
This study was approved by the Regional Ethical Review Board in Lund (registration number 2011/626) and included 39 consecutive adult patients at the Department for Intensive and Perioperative Care, Skåne University Hospital, Lund, Sweden from September 2011 to March 2013. Written informed consent was obtained from all patients before inclusion.
The inclusion criteria were thrombocytopenia <50 × 109/L due to bone marrow failure (chemotherapy, malignancy, or both) and scheduled platelet transfusion for planned CVC insertion. Exclusion criteria included patients with hepatic or renal failure, heparin-induced thrombocytopenia, idiopathic thrombocytopenic purpura, and those receiving mechanical ventilation or anticoagulant therapy (except for thromboprophylactic low molecular weight and unfractionated heparin [subcutaneous]).
The indication for CVC insertion was chemotherapy in all patients. The CVC was placed either in the subclavian or internal jugular vein at the discretion of the performing physician. All patients received 1 therapeutic unit of platelets, containing approximately 260 to 270 × 109/L platelets. One therapeutic unit of platelets could be either whole blood pooled buffy coat platelets from 4 donors or apheresis platelets from 1 donor (Table 1). One therapeutic unit contained approximately 300 mL of which 75 mL was plasma in the blood pooled buffy-coat platelets, and 100 mL was plasma in the apheresis platelets. Samples for bacterial culture were taken from each therapeutic unit on the first day after collection and were proven negative before the unit was transfused in a patient. All transfused platelets were ≤7 days old and leukocyte reduced to 0.001 × 109 leukocytes per unit.
Blood sampling was performed using a vacutainer system (BD, Plymouth, UK) before the platelet transfusion, 1 hour, and 4 hours after the completion of transfusion. The 1-hour sampling point was chosen according to the recommended Hemovigilance protocol,16 and the sampling at 4 hours after transfusion was the extended time limit to perform analyses during office hours. Pretransfusion blood samples were collected from a peripheral vein within 4 hours before the platelet transfusion. Posttransfusion blood samples were collected from the CVC after the pertinent catheter was flushed with normal saline, and after 10 mL blood was discarded. No heparin or fibrinolytic drug was administered through CVCs.
Blood samples were analyzed at each time point by conventional coagulation tests; prothrombin time (PT)/international normalized ratio (INR), activated partial thromboplastin time (aPTT), platelet count, and fibrinogen along with ROTEM and Multiplate. The flow cytometry analysis was performed in the last 17 of 39 patients. Blood samples for ROTEM and flow cytometry analysis were collected in a 4.5-mL tube containing 0.109 M citrate (BD, Plymouth, UK). Blood for Multiplate analysis was collected in a 3.0-mL tube containing recombinant hirudin (Dynabyte GmbH, Munich, Germany).
Conventional Hematological Tests
PT/INR, aPTT, fibrinogen and platelet count analyses were performed at the accredited hospital laboratory. PT/INR was performed using a combined thromboplastin reagent (Stago prothrombin complex assay, SPA+, Stago). The Owren PT assay was calibrated using INR calibrators certified by the Swedish external quality assessment organization (Equalis, Uppsala, Sweden). The reference range for PT/INR is 0.9 to 1.2.
APTT was analyzed with an aPTT reagent from Actin FSL (Siemens Healthcare Diagnostics) Plasma fibrinogen concentration was measured using the Dade Thrombin reagent (Siemens Healthcare Diagnostics. CS-5100). The reference range for aPTT has been established locally to 26 to 33 seconds and for fibrinogen to 2-4 g/L.
Platelet counts were measured using the Sysmex XE 5000 cell counter (Sysmex Corp., Kobe, Japan). The locally determined reference range for platelets is 165 to 387 × 109/L for adult women and 145 to 348 × 109/L for adult men.
(ROTEM; Pentapharm, Munich, Germany) was used for viscoelastic evaluation of clot formation in the recalcified whole blood at 37°C. The test was performed within 2 hours from blood sampling. Tissue factor was used to trigger coagulation for EXTEM and FIBTEM. Thrombin-mediated platelet activation and fibrin polymerization are reflected on EXTEM, while fibrin polymerization is selectively shown on FIBTEM by inhibiting platelet–fibrin interactions using cytochalasin D.17 The latter correlates with plasma fibrinogen levels.18 The limited number of glycoprotein IIb/IIIa receptors due to thrombocytopenia results in decreased platelet–fibrin interactions, and reduced clot formation time (CFT, seconds), α angle (°), maximum clot firmness (MCF, mm).19,20
Multiple Electrode Aggregometry
Multiplate (MEA, Roche, Rotkreuz, Switzerland) was used to measure an agonist-induced platelet aggregation. Hirudin-anticoagulated whole blood was stored at room temperature before analysis within 0.5 to 3.0 hours of blood sampling. The analysis was performed in duplicate at 37°C. The extent of platelet aggregation was measured by resistance (impedance) changes between 2 electrodes and is depicted as a graph. The area under the curve (AUC) is the best measure of platelet function. Three test assays were performed: ADP test (platelet aggregation in response to adenosine-5’-diphosphate), COL test (platelet aggregation in response to collagen), and TRAP test (platelet aggregation in response to thrombin receptor agonist peptide).
Flow Cytometry Analysis
Platelet function analysis using MEA is dependent not only on platelet function but also on platelet count.21–23 Therefore, increased platelet aggregation in response to agonists in blood from thrombocytopenic patients could be due to either improved platelet function, increased platelet count, or both. To address this, we performed flow cytometry that assessed the functional status of individual platelets independent of platelet count.
Flow cytometry analyses were performed within 2 hours after blood samples. Platelet-rich plasma (PRP) was obtained by centrifugation at 140g for 10 minutes. PRP 50 μL was incubated with 5 μL CD41a-PerCPCy5.5 (BD, cat no 333148) or 5 μL CD42b-PE (Dako, cat no R7014) for 15 minutes at room temperature. To activate the platelets, 10 μL labeled PRP was incubated with 40 μL ADP (final concentration (FC) 5 μM), CRP-XL (cross-linked collagen reactive peptide; FC 0.2 μg/mL), TRAP (2.5 μM) or HEPES-bovine serum albumin (BSA) buffer (20 mM Hepes, 137 mM NaCl, 2.7 mM KCl, 1 mM MgCl2, and 1% BSA) (resting platelets) for 10 minutes at room temperature. To detect activation, PRP labeled with CD41a-PerCPCy5.5 was subsequently incubated with 5 μL CD62P-PE (Beckman Coulter, cat no IM1759) and PRP labeled with CD42b-PE was incubated with 5 μL PAC-1-FITC (BD cat no 340507) for 15 minutes at room temperature. As control 5 μL MsIgM-FITC and 5 μL MsIgG1-PE was incubated with 10 μL of resting PRP in 40 μL HEPES-BSA buffer. The samples were subsequently diluted in 2 mL Isoflow (Beckman Coulter, Bromma, Sweden) and run directly on flow cytometer, Gallios (Bio-Rad Laboratories Ltd, Hemel Hempstead, UK). In the Gallios, the laser setting W2 was used. Thresholds for forward scatter and side scatter were set to 2 and with live gate on CD41a-PerCPCy5.5 or CD42b-PE positive events. Forward scatter, side scatter, and fluorescence channels were set at logarithmic gain and live gate on CD41a-PerCPCy5.5 or CD42b-PE positive events were used. For MsIg-controls, no live gate was used, and platelets were defined according to their forward scatter/side scatter characteristics. The flow cytometry data were analyzed using the software Kaluza Flow Cytometry Software. Activation was expressed as a percentage of positive platelets, where a positive gate was set using the MsIg control antibodies (2% false positive).
Local hematomas were evaluated daily by trained staff on the hematological ward and documented on the electronic charts under set headings. Bleeding was classified according to the Common Terminology Criteria for Adverse Events (Version 4.0),24 where grade 1 bleeding is characterized by mild symptoms. Grade 2 bleeding may require minimally invasive evacuation or aspiration. In grade 3 bleeding, transfusion, radiologic, endoscopic, or elective operative intervention are indicated. Grade 4 is life-threatening bleeding, and grade 5 is death.
A power analysis was performed using ROTEM-CFT and MCF changes from the previously published data in the same type of patients at our institution.10 The sample size requirement was lower with the MCF-based calculation, and thus, CFT was used for the final sample size. Based on the change of CFT of EXTEM (CFTEXT) from 181.5 to 123 seconds after platelet transfusion with a SD of 144 seconds, the required minimal sample size was calculated to be 35 with an α level of 5% and a β level of 50%. Differences between laboratory results before, 1, and 4 hours after transfusion were calculated using 2-tailed, Wilcoxon matched pairs signed test. Correlation coefficients were calculated using Spearman rank correlation method. All variables were considered nonparametric (Gaussian distribution not assumed), and all distributions are summarized using the median with 25th and 75th percentiles Q2 (Q1 and Q3). P values < 0.01 were considered significant.
To address the risk that the increased values seen after platelet transfusion are caused by regression to the mean, we performed 3 different tests on platelet count (for details see Supplemental Digital Content, http://links.lww.com/AA/A878).
- Regression analysis baseline versus 1 and 4 hours after transfusion.
- Modified Bland-Altman plot. Baseline versus Δplatelet count 1 and 4 hours after transfusion.
- Median split analysis.
Statistical analyses were performed using GraphPad Prism version 6.03 for Windows, GraphPad Software, La Jolla, California.
Demographic data of the subjects are shown in Table 1. In terms of types of transfused platelets, 2 of 39 platelet transfusions were blood pooled buffy-coat platelets, and 31 of 39 platelet transfusions were ABO-matched (Table 1).
Because gender differences did not affect the results after the preliminary analysis, all data are presented in aggregate form. All patients received a single-lumen CVC. Twenty-six CVCs were inserted in the subclavian vein and 13 in the internal jugular vein. Four grade 1 bleeding events occurred, but no grade 2 to 4 bleeding events were detected.
Conventional Hematological Tests
Platelet count increased from 24 × 109/L (18–32) at baseline to 42 × 109/L (31–50) at 1 hour after transfusion (P < 0.0001). The count was 40 × 109/L (29–50) 4 hours after transfusion (P = 0.047 vs 1 hour after transfusion) (Table 2 and Fig. 1). PT-INR, aPTT, and fibrinogen were unchanged after transfusion.
MCFEXT was increased from 38 (32–45 mm) before to 46 (41–52 mm) at 1 hour after transfusion (P < 0.0001) and did not change 4 hours after transfusion (P = 0.06). The increase in platelet count after 1 hour (ΔPLC) correlated statistically with the increase in MCFEXT (r = 0.56, P < 0.0002) (Table 2 and Fig. 2).
Clotting time EXTEM (CTEXT) was decreased from 58.5 (50–78 s) before to 53 (45–61 s) at 1 hour after transfusion (P = 0.0006) and was 57 (52–70 s) 4 hours posttransfusion (P = 0.025, vs 1 hour after transfusion). CFT EXTEM (CFTEXT) was changed in a similar manner. The α-angleEXT was not significantly changed at any time (both P > 0.04). Similar to plasma fibrinogen values, FIBTEM results were not affected by platelet transfusion (both P > 0.74). Fibrinogen level correlated well with MCFFIBTEM (MCFFIB) (r = 0.84, P < 0.0001) (Table 2 and Fig. 2).
All Multiplate analyses were significantly increased after 1 hour and were not diminished at 4 hours after the transfusion (Table 2 and Fig. 1).
There was no change in the expression of CD62P in any of the samples. The binding of PAC-1 after ADP-stimulation showed a small but significant decrease 1 hour after transfusion. After stimulation with CRP, the binding of PAC-1 did not change after transfusion compared to before transfusion (Table 2 and Fig. 3).
This prospective observational study in thrombocytopenic patients with bone marrow failure evaluated the duration and mechanisms of changes in coagulation induced by prophylactic platelet transfusion. We found that platelet transfusions increased platelet count significantly by 74% at 1 hour after transfusion, and that this increase persisted for 4 hours after transfusion. When evaluating coagulation using thromboelastometry (ROTEM), we found that MCFEXT increased 1 hour after transfusion, and that this increase persisted for 4 hours. Platelet transfusions also accelerated the onset of tissue factor-induced coagulation (CTEXT). CTEXT is a sensitive parameter to detect the initiation of the coagulation cascade. One therapeutic unit of platelet transfusion contains about 100 mL plasma, which may also accelerate CTEXT onset. This effect may partially explain the transient decrease in CTEXT observed 1 hour after transfusion. Platelet aggregation in response to 3 different agonists on Multiplate also demonstrated steady increases in aggregation throughout the entire time of observation. These results indicate that hemostatic effects of platelet transfusion last up to at least 4 hours and suggest that an optimal window for performing an invasive procedure in patients with bone marrow failure would be 4 hours. Although the hemostatic improvement may last longer, we were limited by office hours and sampling volumes for further measurements.
Flow cytometry analyses showed that after platelet activation, expression of the granule protein CD62P on the surface of the platelets was unchanged after stimulation with different platelet activators (see Methods). Given that almost every other platelet in the blood samples after transfusion was a transfused platelet, this finding suggests that transfused platelets were functioning as well as the patient’s own platelets.
Still, after platelet activation with ADP, we observed a slight decrease in the expression of PAC-1. Overall, the nonsignificant changes in platelet function from flow cytometry indicate that the increase in hemostasis after transfusion is not explained by better functioning platelets but rather by an increase in the platelet count.
We found no significant bleeding complications but did identify 4 cases of grade I bleeding events. These findings suggest that the carefully designed clinical practice at our institution is safe. However, our study was not sufficiently powered to make a conclusive statement on this clinical end point.
Patients in our study had fibrinogen levels in the upper part of the reference range, well correlated with MCFFIBTEM (Fig. 2). Lang et al.25 suggest that low clot firmness in thrombocytopenic patients, measured with ROTEM, may be restored with high fibrinogen levels. Velik-Salichner et al.26 performed a study on thrombocytopenic pigs and showed that high-dose fibrinogen administration improved clot firmness more than platelet transfusion and that fibrinogen concentrate stopped induced bleeding faster than did platelet transfusion. Whether the high fibrinogen levels noted in our study resulted from an inflammatory process, or if they were a compensatory mechanism for low platelets, is unclear. Further clinical studies are needed to verify whether platelet transfusions can be avoided in thrombocytopenia if fibrinogen levels are high. There was a statistically nonsignificant increase of plasma fibrinogen (3.7 g/L) at 4 hours compared with the baseline and 1 hour after transfusion (both, 3.3 g/L; Table 2). This increase could have been an inflammatory reaction secondary to the CVC procedure in itself or a reaction to the transfused platelets. Because the effect was more pronounced after 4 hours as compared with 1 hour after platelet transfusion, it was unlikely to have been caused by plasma contained in platelet concentrate.
A limitation with the present study is that analyses were not performed immediately after completion of the platelet transfusion. The coagulation enhancement immediately after transfusion might have been even greater than the improvement that we observed 1 hour after transfusion. However, current recommendations state that if refractoriness to platelet transfusion is suspected, a new platelet count should be measured 10 to 60 minutes after completion of the transfusion.3,16 Our study complied with this recommendation. Another limitation of this study is that platelet function assays were done under minimal shear so that platelet function under higher flow conditions, which is influenced by hematocrit and von Willebrand factor, could be assessed.27
Neither ABO-matching nor the age of the platelet transfusions affected the coagulation enhancement in this study. This relationship has been shown in other studies,3,28 but our study was underpowered and not designed to detect these effects.
The present study is an important reappraisal of hemostatic effects of platelet transfusion in thrombocytopenic patients. There is a paucity of evidence for prophylactic platelet transfusion in the threshold for bleeding and the dose for hemostasis.3–6 Furthermore, it is unclear whether prophylactic platelet transfusion before an intervention is superior to a strategy of therapeutic transfusion only.7,8 The threshold platelet count of ≥ 50 × 109 at our institution was safe in a recent retrospective analysis29 but is arbitrary and not based on large randomized trials. Another retrospective study30 showed that a platelet count as low as 20 × 109/L did not increase bleeding complications during CVC insertions in patients with acute leukemia. On the contrary, 2 independent studies, 1 prospective audit,31 and 1 retrospective analysis32 indicated that a platelet count <50 × 109/L increased the risk for hemorrhagic complications. Furthermore, the site of CVC insertion is important. If the bleeding site is compressible (e.g., the internal or external jugular vein), the need for normal hemostasis is not as high as if the insertion site were noncompressible (e.g., the subclavian vein). Other factors of proven importance are the caliber of the inserted catheter, the experience of the operator, and the use of real-time ultrasound guidance.33 Taken together, prospective studies are needed to define optimal transfusion trigger(s) for platelet concentrates in thrombocytopenic patients due for a CVC.
This study adds information about how a platelet transfusion contributes to the coagulation system, how its effect may be measured, and the duration of this measured coagulation enhancement.
Prophylactic platelet transfusions given to thrombocytopenic patients with bone marrow failure improve hemostatic variables by increasing the number of platelets, and not via enhancement of platelet function. Improved clotting parameters as measured by thromboelastometry, and platelet aggregation on MEA appear to persist between 1 and 4 hours after transfusion.
Name: Thomas Kander, MD.
Contribution: Thomas Kander contributed with design and conduct of the study, data collection, data analysis, and manuscript preparation.
Attestation: Thomas Kander approved the final manuscript and also attests to the integrity of the original data and the analysis reported in this manuscript and is the archival author.
Conflicts of Interest: The author has no conflicts of interest to declare.
Name: Kenichi A. Tanaka, MD, Msc.
Contribution: Kenichi A. Tanaka provided help with data analysis and manuscript preparation.
Attestation: Kenichi A. Tanaka approved the final manuscript and also attests to the integrity of the original data and the analysis reported in this manuscript.
Conflicts of Interest: Kenichi A. Tanaka has served on the advisory board for the TEM International, Munich, Germany (the company was not involved in the manuscript preparation).
Name: Eva Norström, MD, PhD.
Contribution: Eva Norström contributed with design and conduct of the study, data analysis, and manuscript preparation.
Attestation: Eva Norström approved the final manuscript.
Conflicts of Interest: The author has no conflicts of interest to declare.
Name: Johan Persson, MD, PhD.
Contribution: Johan Persson contributed with study design and manuscript preparation.
Attestation: Johan Persson approved the final manuscript.
Conflicts of Interest: The author has no conflicts of interest to declare.
Name: Ulf Schött, MD, PhD.
Contribution: Ulf Schött contributed with study design, data analysis, and manuscript preparation.
Attestation: Ulf Schött approved the final manuscript and also attests to the integrity of the original data and the analysis reported in this manuscript.
Conflicts of Interest: The author has no conflicts of interest to declare.
This manuscript was handled by: Avery Tung, MD.
We would like to thank the assistant nurse, Lena Åkesson, who contributed with the collection of samples. We also like to thank 2 technicians, Eva Johansson and Anette Ingemarsson, who performed the flow cytometry analysis as well as Olof Axler, MD, PhD, and assistant professor, Sven Björnsson, for useful discussions.
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