Research ArticleCOAGULATION

Impaired hemostatic activity of healthy transfused platelets in inherited and acquired platelet disorders: Mechanisms and implications

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Science Translational Medicine  11 Dec 2019:
Vol. 11, Issue 522, eaay0203
DOI: 10.1126/scitranslmed.aay0203

Strength in numbers (of platelets)

Platelet transfusions are effective at restoring hemostasis in patients with low platelet counts, but this tactic does not necessarily work in patients who have normal numbers of platelets but impaired platelet function resulting from genetic or drug-induced defects. Using mouse models and platelets from human patients, Lee et al. demonstrated that dysfunctional platelets can interfere with the formation and/or consolidation of a platelet plug at the site of injury, but this impairment can be overcome by providing wild-type platelets at a sufficiently high ratio.


Platelet transfusions can fail to prevent bleeding in patients with inherited platelet function disorders (IPDs), such as Glanzmann’s thrombasthenia (GT; integrin αIIbβ3 dysfunction), Bernard-Soulier syndrome [BSS; glycoprotein (GP) Ib/V/IX dysfunction], and the more recently identified nonsyndromic RASGRP2 variants. Here, we used IPD mouse models and real-time imaging of hemostatic plug formation to investigate whether dysfunctional platelets impair the hemostatic function of healthy donor [wild-type (WT)] platelets. In Rasgrp2−/− mice or mice with platelet-specific deficiency in the integrin adaptor protein TALIN1 (“GT-like”), WT platelet transfusion was ineffective unless the ratio between mutant and WT platelets was ~2:1. In contrast, thrombocytopenic mice or mice lacking the extracellular domain of GPIbα (“BSS-like”) required very few transfused WT platelets to normalize hemostasis. Both Rasgrp2−/− and GT-like, but not BSS-like, platelets effectively localized to the injury site. Mechanistic studies identified at least two mechanisms of interference by dysfunctional platelets in IPDs: (i) delayed adhesion of WT donor platelets due to reduced access to GPIbα ligands exposed at sites of vascular injury and (ii) impaired consolidation of the hemostatic plug. We also investigated the hemostatic activity of transfused platelets in the setting of dual antiplatelet therapy (DAPT), an acquired platelet function disorder (APD). “DAPT” platelets did not prolong the time to initial hemostasis, but plugs were unstable and frequent rebleeding was observed. Thus, we propose that the endogenous platelet count and the ratio of transfused versus endogenous platelets should be considered when treating select IPD and APD patients with platelet transfusions.


Platelets are the essential cellular mediators of primary hemostasis. After blood vessel damage, platelets make transient contacts with von Willebrand factor (vWF) exposed in the extracellular matrix (ECM) (1), mediated by the glycoprotein (GP) Ib/V/IX complex. Reduced platelet velocity then allows for platelet activation via engagement of immune-type and G protein–coupled receptors (2, 3). Activation of these receptors triggers a rapid increase in cytosolic Ca2+ concentrations, leading to the activation of calcium and diacylglycerol-regulated guanine nucleotide exchange factor (CalDAG-GEFI; gene name RASGRP2) and, in turn, the small guanosine triphosphatase RAP1. Active RAP1 engages the cytoskeletal adapter protein TALIN to induce a conformational change in αIIbβ3 integrin, resulting in the high-affinity state required for fibrinogen binding and platelet aggregation (4, 5).

Both clinical and experimental observations have shown that loss of expression or function of any of these molecular components leads to impaired platelet-mediated hemostasis and bleeding. Bernard-Soulier syndrome (BSS), a rare but severe bleeding diathesis, results from molecular pathology in genes encoding the subunits of GPIb/V/IX, leading to loss of expression or function of the major platelet receptor for vWF, and is often associated with macrothrombocytopenia (6). Glanzmann’s thrombasthenia (GT) is caused by mutations to either subunit of the αIIbβ3 integrin, leading to loss of function/expression of the complex, abolished platelet aggregation, and severe bleeding (7, 8). More recently, patients have been identified with mutations in RASGRP2 (917). In agreement with the mouse model (18), human platelets lacking CalDAG-GEFI function exhibit a markedly impaired integrin activation response to various agonists. As a result, patients with RASGRP2 mutations present with a moderate to severe bleeding diathesis, including bleeding after tooth extraction, excessive menstrual bleeding, and bleeding complications after pregnancy.

To prevent or reverse bleeding in patients with RASGRP2 mutations, a number of treatments have been used, with varying success. The two major treatment options are administration of recombinant active FVII (rFVIIa; NovoSeven) (19) and/or platelet transfusions. Unlike patients with BSS or GT, patients with RASGRP2 mutations have normal surface GP expression (15) and are unlikely to generate antiplatelet antibodies against the major platelet GPs after transfusions; because of this, and the high cost of recombinant coagulation factors, platelet transfusion is a desired treatment option. To date, however, transfusion is reported to be ineffective in some patients. For example, a recent study of a family with a truncation deletion in the C-terminal regulatory domain of RASGRP2 (c.1490delT) demonstrated that perioperative platelet transfusion was unsuccessful in controlling bleeding in two homozygous family members. Cessation of bleeding occurred only after administration of rFVIIa (15).

Besides inherited disorders, patients on antiplatelet therapy may also require platelet transfusion. Platelet inhibitors are widely used in the prevention of thrombotic events but carry an inherent risk of bleeding. The current gold standard in the prevention of secondary cardiovascular events is dual antiplatelet therapy (DAPT) with low-dose aspirin and a P2Y12 inhibitor (typically clopidogrel, prasugrel, or ticagrelor) (20, 21). Patients on DAPT who require emergency surgical intervention often receive platelet transfusion ahead of the procedure to prevent bleeding (22), but the effectiveness of transfusion in these patients is not entirely understood (23). The inability to control peri- and postoperative bleeding is a major concern in patients with platelet function disorders (24). Therefore, it is of great clinical relevance to understand why certain patients with platelet dysfunction do not benefit from platelet transfusion.

In this work, we used a mouse model system to investigate the interaction of dysfunctional platelets with healthy [wild-type (WT)] platelets at sites of vascular injury. To analyze transfusion effectiveness in vivo, we used two experimental approaches: (i) transfusion of WT platelets into murine models of inherited platelet disorders or (ii) recipient mice rendered thrombocytopenic (TP) and transfused with specific ratios of WT and dysfunctional platelets. Our studies suggest that platelets with impaired integrin activation compete with WT platelets for GPIbα binding sites on vWF or other GPIb ligands at sites of injury and that this competition negatively affects the hemostatic activity of transfused WT platelets. Spinning disk confocal (SDC) imaging of hemostatic plugs further identified negative effects of dysfunctional platelets on the composition of the plugs. In addition, in mice cotransfused with WT and “DAPT” platelets, hemostatic plugs formed rapidly but demonstrated instability and reopening. We conclude that the endogenous platelet count and the ratio of transfused versus endogenous platelets would be useful in predicting the success of platelet transfusion therapy in patients with platelet integrin signaling defects.


Healthy transfused platelets have reduced hemostatic efficacy in the presence of Rasgrp2−/− platelets

Deficiency in CalDAG-GEFI causes severe bleeding in mice (18) and moderate to severe bleeding in humans, often requiring platelet transfusions (17). In patients with RASGRP2 mutations, transfused healthy platelets need to function in the presence of a normal number of dysfunctional platelets to promote hemostasis. We sought to examine the potential competition between transfused and endogenous platelets during hemostasis. To investigate aspects of hemostatic plug formation in vivo, we performed top-down penetrating laser injury of the murine saphenous vein coupled with intravital microscopy (25). This model allows for the quantification of bleeding time (time to hemostasis) with simultaneous visualization of fluorescently labeled platelets and other blood components. We first determined the hemostatic impact of WT platelets transfused into Rasgrp2−/− mice (Fig. 1A). Similar to previous studies, WT mice formed a hemostatic plug in 20 to 30 s (Fig. 1, B and C, purple, and data file S1) (25), whereas Rasgrp2−/− mice typically bled for the entire observation period (270 s) (26) with only few injuries reaching hemostasis (Fig. 1, B and C, red). As expected, platelet-depleted mice (TP IL-4R/GPIb-transgenic mice; TP Tg) also bled for the entire observation period (Fig. 1, B and C, black). Consistent with recent studies demonstrating that a low peripheral platelet count (5 to 10% of normal) is sufficient for hemostasis in mice (27), we were able to prevent bleeding in TP Tg mice by transfusing WT platelets to achieve a peripheral count of ~5 × 107/ml (Fig. 1, B and C, open black circles/dashed line). Transfusing twice that number of WT platelets into nondepleted Rasgrp2−/− mice [resulting in a ~10:1 ratio of mutant (endogenous) to WT (transfused) platelets] had only a minimal effect on the bleeding time after laser injury (Fig. 1, B and C, blue). To secure hemostasis, Rasgrp2−/− mice had to be transfused with considerably more WT platelets than were required in TP mice (Fig. 1, B and C, orange). A peripheral count of 3 × 108/ml WT platelets, which results in a ratio of ~3:1 Rasgrp2−/−:WT platelets, was established as the threshold required to normalize hemostasis when transfusing WT platelets into Rasgrp2−/− mice.

Fig. 1 Rasgrp2−/− mice require a large number of transfused WT platelets for hemostasis.

(A) Model depicting platelet (plt) transfusion scheme in Rasgrp2−/− mice. Endogenous platelets were labeled by injection of anti–GPIX–Alexa Fluor 647 antibody (Ab) before transfusion of GPIX–Alexa Fluor 488–labeled WT platelets. (B) Comparison of bleeding times in the saphenous vein laser ablation model. Rasgrp2−/− mice were transfused, or not, with a low (1 × 108/ml) or high (3 × 108/ml) number of WT platelets and compared with WT mice or platelet-depleted IL4R-GPIb-Tg (TP Tg) mice transfused, or not, with a very low (5 × 107/ml) number of WT platelets. Each dot represents the average time to hemostasis for four to six individual injuries in one mouse; n = 3 to 6 mice per group; data shown as means ± SEM. P < 0.05: † versus WT, ‡ versus Rasgrp2−/− (no transfusion), # versus TP Tg (no transfusion), $ versus TP Tg (+5 × 107/ml WT platelets), and & versus Rasgrp2−/− (+1 × 108/ml WT platelets). (C) Kaplan-Meier curve representation of bleeding time data, including all individual injury sites. Note that >50% of injuries are still bleeding after 270 s in Rasgrp2−/− mice receiving 1 × 108/ml WT platelets.

We next performed adoptive transfer studies with differentially labeled WT and Rasgrp2−/− platelets in TP Tg recipient mice (Fig. 2A) (28). Flow cytometry dot plots show depletion and reconstitution of Tg recipient mice, and both the WT and Rasgrp2−/− [knockout (KO)] platelet populations were clearly distinguishable by their fluorescence signals with no double-staining of platelets (Fig. 2B). Furthermore, WT and Rasgrp2−/− platelets in whole blood from transfused mice demonstrated similar integrin activation profiles to endogenous platelets from WT and Rasgrp2−/− mice, suggesting that platelet function was minimally affected by the labeling and/or transfusion process (fig. S1). TP Tg mice transfused with WT platelets alone rapidly achieved hemostasis after laser injury (Fig. 2, C and D, purple; data file S1; and movie S1), whereas mice receiving Rasgrp2−/− platelets alone bled for almost the entirety of the observation period (Fig. 2, C and D, red, and movie S2). Guided by our findings from WT platelet transfusions into Rasgrp2−/− mice (Fig. 1), we transfused mutant and WT platelets at a ratio of either 5:1 or 2:1 into TP Tg mice (with the absolute number of WT platelets kept constant). In agreement with the results obtained in Rasgrp2−/− mice, a ratio of 2:1 Rasgrp2−/−:WT platelets was considerably more effective than a 5:1 ratio in normalizing time to hemostasis (Fig. 2, C and D, blue and orange, and movies S3 and S4, respectively). These results demonstrated that increasing numbers of Rasgrp2−/− platelets led to prolonged bleeding times in mice transfused with the same absolute number of WT platelets, strongly suggesting that Rasgrp2−/− platelets impair the hemostatic efficacy of WT platelets.

Fig. 2 Cotransfusion of 2:1 Rasgrp2−/−:WT platelets is required for normal hemostasis.

(A) Model depicting platelet transfusion scheme. IL4R-GPIb-Tg mice were depleted of endogenous platelets (TP Tg) and transfused with labeled WT and/or Rasgrp2−/− platelets at the desired ratios, before undergoing the laser ablation hemostasis model. (B) Flow cytometry dot plots of whole blood from IL4R-GPIb-Tg mice before and after endogenous platelet depletion, and after WT/Rasgrp2−/− platelet transfusion. In blood collected after transfusion, the anti-CD41 antibody MWReg30 was used to label all transfused platelets, and distinct populations of differentially labeled WT(488) and Rasgrp2−/−(647) platelets were observed. (C) TP Tg mice were transfused with WT or Rasgrp2−/− platelets only or the indicated ratios of Rasgrp2−/−:WT platelets, and time to hemostasis was determined in the saphenous vein laser ablation model (movies S1 to S4). n = 3 to 10 per group; data shown as means ± SEM. P < 0.05: † versus TP Tg (+WT platelets only), ‡ versus TP Tg (+Rasgrp2−/− platelets only), and # versus TP Tg (+Rasgrp2−/−:WT platelets 5:1). (D) Kaplan-Meier curve representation of bleeding time data, including all individual injury sites. (E) Still frame images from epifluorescence videos at ~60 s after laser injury in TP Tg mice transfused with WT(647):WT(488) (top) or Rasgrp2−/−(647):WT(488) platelets (bottom) at a ratio of 2:1. Scale bars, 25 μm. (F) Quantification of WT(488) platelet adhesion (sum fluorescence intensity) in WT:WT or Rasgrp2−/−:WT platelet-transfused TP Tg mice after laser injury. Red boxes in (E) define the areas where 488 intensity was measured. Sum intensity was normalized to the maximum intensity at time to hemostasis for each injury. n = 3, data graphed as means ± SEM.

To substantiate this conclusion, we quantified the adhesion of anti–GPIX–Alexa Fluor 488 antibody–labeled WT platelets in TP Tg mice cotransfused with Rasgrp2−/−(AF647):WT(AF488) platelets at a ratio of 2:1 and, as a control, TP Tg mice cotransfused with WT(647):WT(488) platelets at the same ratio. Representative still frame images from epifluorescence videos at ~60 s after laser injury demonstrate successful hemostatic plug formation in both groups of recipient mice (Fig. 2E). However, WT(488) platelets showed markedly delayed adhesion to the site of laser injury in the presence of Rasgrp2−/−(647) platelets compared to cotransfusion with WT(647) platelets (Fig. 2F). In addition, Rasgrp2−/− platelets adhered to the site of injury at a similar rate to WT platelets when cotransfused (fig. S2), suggesting that WT platelets can also enhance the recruitment of dysfunctional platelets with impaired adhesive function (26, 29) to the site of injury.

Dysfunctional platelets alter hemostatic plug structure and integrity

At the conclusion of time-lapse epifluorescence imaging, SDC microscopy was used to acquire Z-stacks of hemostatic plugs. Epifluorescence imaging established that both WT and Rasgrp2−/− platelets rapidly accumulated at the site of vascular injury when cotransfused into TP Tg mice, and both platelet populations seemed randomly distributed throughout the plug immediately after hemostasis was achieved (Fig. 2E). However, SDC Z-stacks revealed a distinctive pattern whereby WT platelets consolidated into discrete areas within the plug, and Rasgrp2−/− platelets filled areas devoid of WT platelets (Fig. 3A, top, and movie S5). Segregation between differentially labeled populations of platelets was not observed in TP Tg mice transfused with WT platelets only (Fig. 3A, middle, and movie S6). In addition, when WT platelets were in excess (TP Tg mice transfused with 1:3 Rasgrp2−/−:WT platelets), the WT platelets did not segregate and formed a normal hemostatic plug (Fig. 3A, bottom).

Fig. 3 Rasgrp2−/− platelets disrupt hemostatic plug architecture.

(A) TP Tg recipient mice were transfused with differentially labeled WT and Rasgrp2−/− platelets, in the combinations and ratios noted to the left of the image panels. Laser injury was performed and hemostatic plugs allowed to form. Spinning disk confocal (SDC) Z-stacks were then acquired at the conclusion of time-lapse imaging. Shown here are representative single Z planes from the center of the hemostatic plug (see cartoon in upper right), with individual channels shown in grayscale along with a colored merged image (see movies S5 and S6 for Z-stack series). Scale bars, 25 μm. (B) Still frames from time-lapse SDC imaging in TP Tg mice transfused with either WT:WT or Rasgrp2−/−:WT platelets at a 2:1 ratio. Colored arrows indicate several instances of loosely adhered platelets being shed from the edges of the Rasgrp2−/−:WT hemostatic plug (movie S7), whereas the WT:WT plug retains its structure with little to no platelet shedding (movie S8). Scale bars, 10 μm.

To monitor platelet dynamics during plug formation at single platelet resolution, we performed real-time SDC imaging in a single plane immediately after laser ablation. Despite achieving hemostasis within 60 to 90 s, plugs containing 2:1 Rasgrp2−/−:WT platelets were loosely packed, with single platelets shedding off from the edges and constant plug remodeling occurring (Fig. 3B, top, and movie S7). Enrichment of WT and mutant platelets in distinct areas of the plug was observed. In contrast, differentially labeled WT platelets integrated randomly and firmly throughout the plug; no shedding of platelets or plug remodeling was observed (Fig. 3B, bottom, and movie S8).

Platelets with impaired integrin activation compete with healthy platelets for GPIbα ligand-binding sites

CalDAG-GEFI and RAP1 are critically important for αIIbβ3 activation and firm platelet adhesion, but not for GPIbα-mediated transient adhesion to vWF (30). Thus, we hypothesized that Rasgrp2−/− platelets interfere with the function of healthy transfused platelets by competing for GPIbα binding sites in the damaged vascular wall. To test this hypothesis, we evaluated the hemostatic efficacy of WT platelets in two other mouse models of inherited platelet disorders: a model of BSS with mice lacking the extracellular domain of GPIbα (IL-4R/GPIb-Tg, “BSS-like”) (31) and a model of GT with mice lacking the integrin adapter protein, TALIN1, specifically in platelets (Talin1f/f x Pf4-Cre+, “GT-like”) (Fig. 4A) (32). Both BSS-like and GT-like mice were previously shown to exhibit a severe hemostatic defect in the saphenous vein laser injury model (25). Transfusion of a low number of WT platelets, yielding a peripheral count of 1 × 108/ml, normalized time to hemostasis in BSS-like, but not GT-like, mice (Fig. 4B). Consistent with our hypothesis, BSS-like platelets showed very poor incorporation into the hemostatic plug (Fig. 4C, top, and movie S9). Similar results were obtained when we cotransfused normal WT platelets with WT platelets treated ex vivo with O-sialoglycoprotein endopeptidase (OSGE) to cleave the 45-kDa N terminus of GPIbα (movie S10) (33). In contrast, GT-like platelets incorporated into the growing plug together with WT platelets (Fig. 4C, bottom, and movie S11). Similar to our findings with Rasgrp2−/− platelets, GT-like platelets also affected the composition of the hemostatic plug, leading to shedding of platelets and segregation of WT and GT-like platelets (movie S12).

Fig. 4 Interference by dysfunctional platelets is dependent on GPIbα.

(A) Model depicting platelet transfusion scheme. Endogenous platelets in GT-like or BSS-like mice were labeled by injection of anti–GPIX–Alexa Fluor 647 antibody before transfusion of GPIX–Alexa Fluor 488–labeled WT platelets. (B) GT-like or BSS-like mice were transfused with WT platelets to achieve a circulating count of 1 × 108/ml, for a ratio of ~7:1 endogenous:transfused platelets in both mouse models. Time to hemostasis was then assessed in the laser injury model. n = 3 per group. (C) Mice were transfused with WT platelets to reach a ratio of ~2:1 endogenous:transfused platelets, to allow for hemostatic plug formation in GT-like mice. Still frame images from epifluorescence videos (movies S9 and S11) demonstrate incorporation of GT-like (bottom), but not BSS-like (top), platelets within the hemostatic plug. Some BSS-like platelets can be seen adhered to the luminal side of the WT plug. Scale bars, 25 μm. (D) Flow chamber assay was performed on collagen-coated (200 μg/ml) coverslips at a shear rate of 1600 s−1. WT blood was flowed alone or after mixing at a 3:1 mutant:WT ratio with Rasgrp2−/− or BSS-like blood for 5 min over the collagen surface and visualized with a 100× oil objective. Platelets were labeled with anti-GPIX antibody before mixing. Representative still frame images are shown. White arrows denote areas of WT thrombus formation. Scale bars, 5 μm.

To validate our findings ex vivo, we studied platelet adhesion to a collagen surface under arterial flow conditions in a microfluidics chamber system. Although our bleeding model uses the venous system, the shear rate substantially increases where blood flows from a penetrating injury (34), which increases dependence on GPIb/vWF. Heparinized whole blood was obtained from WT, Rasgrp2−/−, and BSS-like mice, differentially labeled with Alexa Fluor 488– and Alexa Fluor 647–conjugated antiplatelet antibodies and mixed at defined ratios before perfusion at a shear rate of 1600 s−1. Thrombus formation observed in WT blood (Fig. 4D, left) was markedly impaired in the presence of an excess of Rasgrp2−/− (Fig. 4D, middle), but not BSS-like, platelets (Fig. 4D, right). Consistent with our in vivo studies, Rasgrp2−/− platelets were also successfully recruited to adherent WT platelets. These results demonstrate a mechanism by which platelets with dysfunctional integrin activation compete for GPIbα-mediated binding to reduce the adhesion and hemostatic efficacy of healthy platelets.

Human GT platelets interfere with healthy platelets

To expand our findings to the human clinical situation, we performed ex vivo competition studies with human platelets from healthy volunteers and GT patients. Platelet function was assessed using the Impact-R method, where shear stress is applied to blood in a polystyrene well and platelet surface coverage is measured. Platelet adhesion in this assay is dependent on both GPIbα/vWF and αIIbβ3/fibrinogen interactions (35). Healthy whole blood was supplemented with 33 or 66% (by volume) platelet concentrate from another healthy donor or from a GT patient. In accordance with our findings in the GT-like mouse model, increasing numbers of GT platelets impaired the function of normal platelets in whole blood in a ratio-dependent manner (Fig. 5). These results demonstrate that competition between healthy and dysfunctional platelets also occurs in humans.

Fig. 5 GT patient platelets interfere with the function of healthy donor platelets.

Blood collected from healthy donors was supplemented with platelet concentrates from other healthy donors or from GT patients. Platelet function was then tested using the Impact-R cone and plate analyzer. Platelet surface coverage was determined by light microscopy after May-Grünwald staining of the well. n = 6 to 7 per condition; data shown as means ± SEM. ***P < 0.001 compared to 0% added platelets. Red/black checkered bar represents control for both healthy and GT platelets added, where healthy blood was run with no added platelets.

DAPT platelets impair the hemostatic activity of WT platelets

The use of platelet transfusions in patients on antiplatelet drugs is not well standardized, and the efficacy is not completely understood (23). To investigate whether platelets with acquired dysfunction also impair the hemostatic activity of fully functional platelets, we cotransfused TP Tg mice with WT and aspirin-treated P2ry12−/− (DAPT) platelets (Fig. 6A) and then monitored the hemostatic plug formation after laser injury. At a ratio of 2:1 DAPT:WT platelets, the adhesion kinetics of WT platelets and the initial time to hemostasis were comparable to mice receiving 2:1 WT:WT platelets (Fig. 6, B and C). However, hemostatic plugs containing DAPT platelets showed instability at the center of the plug, leading to visible remodeling and rebleeding (Fig. 6, D and E, and movie S13). Thus, platelets with an acquired function defect, such as seen with antiplatelet therapy, also interfere with the hemostatic efficacy of transfused WT platelets, at least in the mouse system.

Fig. 6 DAPT platelets impair plug stability in the presence of WT platelets.

(A) Model depicting platelet transfusion scheme. P2ry12−/− platelets were treated with 2 mM acetylsalicylic acid (DAPT) ex vivo before transfusion. (B) Time to hemostasis in TP Tg mice transfused with 2:1 DAPT:WT platelets or 2:1 WT:WT platelets was determined after laser injury. (C) Adhesion of WT-488 platelets was quantified in WT:WT or DAPT:WT platelet-transfused TP Tg mice. Data shown as means ± SEM. (D) Rebleeding events (%) at individual injury sites during RT SDC imaging. Five to eight injuries from two to three mice per group. (E) Still frame images from RT SDC video showing hemostasis and rebleeding at the same injury site (movie S13). White arrow indicates site of blood flow from hemostatic plug. Scale bars, 25 μm.


Inherited and acquired platelet function disorders (IPDs and APDs, respectively) are often associated with bleeding, which necessitates transfusion of blood products and administration of procoagulant and/or antifibrinolytic agents (36). However, platelet transfusions can fail to correct hemostasis in these patients. In this study, we used various murine models of both IPDs and APDs in combination with in vivo, real-time imaging of hemostatic plug formation to establish the underlying mechanism(s) for the limited hemostatic activity of fully functional (normal) transfused platelets in these situations. The main conclusions from our work are as follows: (i) normal platelets are less hemostatically active in the presence of dysfunctional platelets, (ii) dysfunctional platelets compete with normal platelets for adhesion to GPIbα ligands in the vessel wall and the growing thrombus, (iii) dysfunctional platelets disrupt the architecture and integrity of the plug, and (iv) the posttransfusion ratio between normal and dysfunctional platelets determines whether platelet transfusion will correct hemostasis. Our findings have important clinical implications, because they provide a basis for guidelines to optimize platelet transfusions for patients with platelet dysfunction but normal platelet counts.

The clinical usefulness of platelet transfusions for the treatment or prevention of bleeding in TP patients is well established (37). Largely based on clinical experience, it is believed that platelet counts greater than 10 × 109 to 20 × 109/liter are sufficient to prevent spontaneous bleeding, whereas counts greater than 50 × 109/liter are required to prevent bleeding after major surgery (3840). Prophylactic platelet transfusions are recommended when platelet counts drop below 10 × 109/liter (37). The corrected count increment (CCI), which compares the posttransfusion to pretransfusion platelet counts, provides a fairly reliable method to estimate posttransfusion platelet survival in patients (41). With the help of the CCI and other methods, it was established that the transfusion of 3 × 1011 platelets, the equivalent of one unit of either apheresis platelets or pooled whole blood-derived platelets, increases the peripheral platelet count in a patient by ~30 × 109/liter. Thus, transfusion of one apheresis or whole blood-derived pooled platelet unit is recommended to prevent bleeding in patients suffering from severe thrombocytopenia (37). However, the efficacy of platelet transfusion in patients with IPDs/APDs is less clear. Compared to TP patients, peripheral platelet counts in IPD and APD patients are generally in the normal range. Whether or not these patients require the same number of transfused platelets as TP patients to restore hemostasis is not known. A case report by Jennings et al. (42) describes findings in a GT patient that suggest that platelet transfusion protocols may need to be adjusted in patients with a qualitative platelet function disorder. In a first, unsuccessful attempt to control bleeding associated with tonsillectomy and adenoidectomy, the GT patient received apheresis platelets from a single donor, yielding a posttransfusion GT:normal (endogenous:transfused) platelet ratio of ~5:1. Hemostasis was restored after an additional transfusion with apheresis platelets from four donors, leading to a posttransfusion GT:normal platelet ratio of ~1:1 (42). Another indication that transfusion requirements are different in the presence of dysfunctional platelets comes from recent reports of patients with mutations in RASGRP2, who show a poor response to platelet transfusion therapy (15, 16, 43). Similar to GT, patients with RASGRP2 mutations exhibit marked defects in integrin-mediated platelet adhesion and hemostatic plug formation. To date, 19 unrelated pedigrees with mutations in RASGRP2 have been reported (17). All of these patients present with moderate to severe bleeding symptoms, which often require combinatorial treatments including platelets, desmopressin, and rFVIIa (15, 16). Of note, the prohemostatic mechanism of rFVIIa in platelet function disorders (most commonly GT and RASGRP2 patients) is not entirely understood. Although platelets from these patients have impaired aggregation, they are still able to adhere and likely provide some surface for coagulation factor complex formation. Therefore, rFVIIa may function by enhancing platelet surface thrombin generation in vivo (44). Moreover, RASGRP2 mutant platelets typically have normal or only minimally reduced response to thrombin (11, 13) and therefore may benefit from increased thrombin generation not only due to fibrin formation (45) but also via thrombin receptor signaling. Future studies should address the mechanism by which procoagulants improve hemostatic plug formation in vivo in the setting of platelet dysfunction.

Our study investigated in vivo whether and how endogenous (dysfunctional) platelets affect the hemostatic activity of transfused platelets. We first investigated the platelet transfusion requirements for correcting hemostasis in Rasgrp2−/− mice. Similarly to patients with RASGRP2 mutations, the KO mice are characterized by a markedly impaired platelet aggregation response, caused by a defect in the near-immediate activation of integrin receptors (46). Our studies demonstrate that, compared to TP mice, hemostasis in Rasgrp2−/− mice requires considerably higher numbers of transfused WT (healthy) platelets. Consistent with findings in the GT patient reported by Jennings et al. (42), the time to hemostasis after vascular injury was markedly improved at posttransfusion Rasgrp2−/−:WT ratios of 2:1 but not 5:1. We showed that both WT and mutant platelets localized to the injury site with similar kinetics. However, high-resolution SDC imaging revealed that their distribution within the plug was not random, because we observed areas rich in either WT platelets or Rasgrp2−/− platelets. At this point, it is not clear why WT and Rasgrp2−/− platelets segregate into different areas within the hemostatic plug. Impaired platelet-platelet cohesion and contraction, mediated by activated integrin receptors, seem to be the most likely explanation. Segregation of procoagulant platelet subpopulations within clots has recently been demonstrated (47) and thus could be an alternative explanation for our findings. However, we have shown that CalDAG-GEFI–deficient platelets have a reduced procoagulant response (48) and procoagulant platelets are enriched on the surface of the plug, whereas Rasgrp2−/− platelets surround clusters of WT platelets within the plug.

Consistent with an integrin-centered explanation, we also found platelet segregation within hemostatic plugs consisting of WT and TALIN1-deficient platelets. TALIN1 is crucial for integrin signaling, and thus, Talin1f/f x Pf4-Cre+ mice (GT-like) show a platelet phenotype similar to that described for GT patients (32). Talin1f/f x Pf4-Cre+ platelets were recruited into the plug and markedly impaired the hemostatic activity of WT platelets. GT is the most severe form of IPD, causing major bleeds in 50% of deliveries (49). In several patients, platelet transfusion failed to prevent these pregnancy-related bleeds. We confirmed the relevance of our in vivo findings for human patients, because we observed that platelets from GT patients impaired the adhesion of healthy donor platelets in whole blood when assessed by the Impact-R method (35). Similar findings were recently reported with the PFA-100 method (50). We acknowledge that our studies in mice are limited in that we exclusively focused on the saphenous vein hemostasis model, with the translational aspect coming from a single ex vivo assay using human platelets. However, the general principles of hemostatic plug formation are conserved between various vessel types and models (51, 52), suggesting that our mechanistic conclusions are transferrable to other vascular beds, although the required transfusion ratio between dysfunctional and functional platelets may slightly vary depending on the vessel and type of injury. Validation of these concepts in patients will require additional empirical data from the clinic, and they may also be studied in mouse models of human platelet transfusion (53).

Using a mouse model for BSS (IL4R/GPIb-Tg, BSS like) (31), we further demonstrated that the recruitment of dysfunctional platelets (Rasgrp2−/− or GT like) to the forming hemostatic plug is likely mediated by GPIbα binding to its main ligands (most likely vWF), in the ECM, and potentially on the surface of WT platelets within the growing plug (54). Platelets lacking GPIbα did not interfere with transfused WT platelets during hemostatic plug formation, even when the ratio of mutant:WT platelets exceeded 5:1. A similar mechanism was previously described in arterial thrombosis in mice; platelets lacking the GPIbα extracellular domain or the 45-kDa N-terminal domain were entirely unable to incorporate into WT arterial thrombi (55). In BSS and GT, platelet transfusions are reserved for patients with serious bleeding due to the potential risk of alloimmunization and platelet refractoriness after repeated exposure to transfused blood products (56). Unlike for GT, however, there are no reports in the literature that platelet transfusion was ineffective in BSS patients. Given that BSS patients also exhibit a marked macrothrombocytopenia, a phenotype that is not seen in patients with GT or RASGRP2 mutations, it is possible that the better hemostatic efficacy of transfused healthy platelets in BSS is, at least in part, a reflection of a lower ratio between dysfunctional and healthy platelets. However, our studies in BSS-like and GT-like mice, which have similar peripheral platelet counts, argue in favor of a mechanistic explanation.

Whereas patients with IPDs are comparatively rare, the use of antiplatelet drugs and the associated bleeding complications are common in the clinic setting. The efficacy and use of platelet transfusion for the acute reversal of DAPT is not well understood (23, 57), and the AABB (formerly called American Association of Blood Banks) has not found convincing evidence supporting the usefulness of platelet transfusion for patients receiving antiplatelet therapy who suffer from intracranial hemorrhage (37). The consensus thinking is that, in patients on aspirin and clopidogrel/prasugrel, irreversible inhibitors of cyclooxygenase-1 and the P2Y12 receptor, platelet transfusion might be beneficial once the drugs are eliminated from circulation. However, by cotransfusing healthy and dual-inhibited platelets and by excluding circulating inhibitors, we provided evidence that DAPT platelets themselves also impair the hemostatic activity of fully functional platelets by markedly destabilizing the hemostatic plug. Unlike Rasgrp2−/− or GT-like platelets, which delayed WT platelet adhesion and hemostasis, DAPT platelets did not affect the initial adhesion of WT platelets and the time to hemostasis. Consistent with the key role of thromboxane A2 (TxA2) and adenosine diphosphate (ADP) in sustained platelet activation (58, 59), however, hemostatic plugs consisting of WT and DAPT platelets were unstable and frequently reopened. Patients on antiplatelet therapy typically have platelet counts within a normal range, similar to those with RASGRP2 mutations, increasing the dysfunctional:healthy platelet ratio.

An important conclusion of our study is that hemostasis can be achieved in IPDs associated with integrin dysfunction if the posttransfusion ratio of dysfunctional/normal platelets is ≤2:1, at least in the saphenous vein laser injury hemostasis model. Obviously, studies in patients will be required to validate this number, but given the case report by Jennings et al. (42), we believe that our number is a good starting point to guide future studies. An important variable to be considered, however, is the age of the platelet product. Our experiments used only freshly washed WT platelets, which were transfused within several hours of blood collection and platelet washing. An extended platelet storage time, however, reduces the hemostatic capacity of donor platelets in humans (60). Therefore, the ratio of dysfunctional:normal platelets required for hemostasis could be altered when transfusing platelets stored for several days. Another important variable is the nature of the platelet function defect. Our studies used mice with severe defects in platelet integrin signaling. Additional studies will be required to determine whether healthy platelets are equally impaired in their hemostatic function in other IPDs and APDs. Mouse models will provide a great tool for highly controlled studies. In humans, an improved understanding of the transfusion efficacy will require more detailed reporting practices, such as the inclusion of the CCI and its correlation with the success of platelet transfusion in individual patients. In disorders like GT and BSS, where the expression of surface GPs is altered, flow cytometry has been used to more accurately determine the ratio between endogenous and transfused platelets (61, 62). For other patients, such as those with mutations in RASGRP2, whose platelets express normal numbers of surface GPs (1113), distinguishing endogenous from transfused platelets would require either platelet prelabeling (63) or an intracellular staining protocol for flow cytometry, the latter of which could be standardized for use in the clinical laboratory. In addition, future studies will be necessary to determine whether the required ratio would be affected by transfusing platelets prophylactically versus therapeutically. Our experimental design modeled prophylactic transfusion, but we speculate that therapeutic transfusion may be less effective because the dysfunctional platelets would have the first opportunity to absorb vWF binding sites.

In summary, we present systematic in vivo analysis of the interaction between endogenous dysfunctional and healthy transfused platelets in murine models of IPDs and APDs. Our studies provide evidence that dysfunctional platelets can interfere with the hemostatic activity of transfused platelets by at least two mechanisms: (i) early interference by competition for GPIbα ligands at the site of injury and (ii) late interference by impaired consolidation of the hemostatic plug. Our finding that a critical ratio of ~2:1 endogenous (dysfunctional) to transfused (healthy) platelets is required to prevent prolonged injury-induced bleeding suggests that patients with select qualitative platelet function disorders will require greater numbers of transfused platelets than TP patients to achieve successful hemostasis. However, current guidelines, which are mostly based on clinical experience in TP patients, suggest the transfusion of up to one apheresis unit (or equivalent) into patients with bleeding complications (37). We propose that multiple apheresis units (or equivalents) are required to control bleeding in patients with an IPD or APD and that the pretransfusion platelet count and the posttransfusion platelet ratio are important variables for determining the required number of platelet concentrates and the predicted effect on hemostasis. Future studies should be directed toward the development of simple clinical tests that help establish the posttransfusion platelet ratio.


Study design

Our experimental design aimed to determine the effectiveness of healthy (WT) platelet transfusion in the setting of IPDs and APDs. Bleeding time was assessed using the saphenous vein laser injury model. The translational relevance of this model is that it is a penetrating injury and is sensitive to both platelet and coagulation defects. The hemostatic efficacy of WT platelets was tested in two ways: transfusion of WT platelets into mouse models of inherited platelet disorders (representative of the clinical situation) or cotransfusion of specific amounts/ratios of WT/dysfunctional platelets into TP recipient mice. For WT and Rasgrp2−/− platelet mixing, n ≥ 3 mice per group were included for statistical analysis. Each data point represents the average time to hemostasis for at least three individual injuries along the saphenous vein for a single mouse. The cutoff for bleeding time in this study was set at 270 s, similar to the duration of the first of three consecutive injuries at the same injury site described in the first publication of this model (25). Only injuries ~50 to 70 μm in diameter were included. In the laser injury model, the researcher typically performed the platelet transfusion and laser injury model in one mouse per day and was not blinded to the condition. Post-experiment determination of bleeding time, assessed by visual cessation of blood flow from the site of laser ablation, was confirmed by two independent but unblinded assessors. For proof-of-principle experiments, where WT platelets were transfused into GT-like or BSS-like mice, n = 2 to 3 per condition. For cotransfusion of WT/WT or WT/DAPT platelets, n ≥ 4. Images of flow chamber assay were representative of three independent experiments. For the Impact-R assay using human platelets, n = 6 to 7 samples were tested at least in duplicate.


Low–molecular weight heparin (enoxaparin sodium, Fresenius Kabi) and isoflurane (Piramal Critical Care) were purchased from UNC Hospitals pharmacy. Antibodies recognizing GPIX (clone Xia.B4), GPIbα [clone Xia.G5, phycoerythrin (PE)–conjugated], αIIbβ3 integrin (clone MWReg30, PE-conjugated), and activated αIIbβ3 integrin (JON/A-PE, clone Leo.H4) were from emfret ANALYTICS. Anti-GPIX antibodies were conjugated with Alexa Fluor dyes (488 and 647) from Life Technologies. Anti–human interleukin-4 receptor (IL-4R) antibody was from R&D Systems. Convulxin was purchased from K. Clemetson (Theodor Kocher Institute, University of Bern, Switzerland). PAR4-activating peptide (PAR4p) was from GL Biochem Inc. ADP and acetylsalicylic acid were from Sigma-Aldrich.

Human subjects

GT patients had been enrolled within the aims of the “Inherited Platelet Disorders Project,” Hemorrhagic Diathesis Working Group, Spanish Society for Thrombosis and Hemostasis (SETH). Investigations in this project abided by the Declaration of Helsinki and were approved by the Local Ethics Committees of Hospital Universitario Reina Sofía (Murcia, Spain). All patients and healthy subjects gave their written informed consent.


The following mouse mutant strains were used in this study: Rasgrp2−/−, Talin1f/f x Pf4-Cre+, hIL-4Rα/GPIbα-Tg, and P2ry12−/−, all on a C57BL/6 background and bred in-house. Both male and female mice were used at 8 to 16 weeks old (20 to 25 g). All experimental protocols were approved by the University of North Carolina Institutional Animal Care and Use Committee.

Platelet isolation

Whole blood was obtained via the retro-orbital plexus using heparinized glass capillaries (VWR) and collected into tubes containing low–molecular weight heparin (30 IU/ml). Platelet-rich plasma was obtained by two rounds of centrifugation of whole blood at 130g and collecting the supernatant with some erythrocytes, followed by two rounds of centrifugation at 100g to pellet erythrocytes. Platelets were then pelleted by centrifugation at 700g in the presence of prostacyclin (PGI2) (1 μg/ml), and the pellet was resuspended in modified Tyrode’s buffer [137 mM NaCl, 12 mM NaHCO3, 2.0 mM KCl, 0.3 mM Na2HPO4, 1 mM MgCl2, 5 mM N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid, and 5 mM glucose (pH 7.3)]. Platelets in Tyrode’s buffer were allowed to rest in a 37°C water bath for 20 to 30 min before transfusion.

Platelet transfusion

For non-TP recipients, endogenous platelets were labeled by intravenous injection of anti–GPIX–Alexa Fluor 488/647 (2.5 μg/mouse) at least 4 hours before platelet transfusion. Platelet counts in Rasgrp2−/− mice were similar to WT mice (1 × 109/ml, ±1 × 108). Recipient mice received transfusions of WT platelets to achieve circulating counts of 1 × 108 or 3 × 108/ml, which were slightly adjusted if the baseline platelet count of the recipient mouse was greater or less than 1 × 109/ml. This transfusion scheme resulted in approximate ratios of 10:1 or 3:1, respectively. For TP recipients, hIL-4Rα/GPIbα-Tg mice were depleted of endogenous platelets via intravenous injection of anti–human IL-4Rα antibody (2 μg/g body weight) at least 4 hours before platelet transfusion (28). Thrombocytopenia was confirmed shortly after injection by flow cytometric analysis of whole blood sample. Donor platelets were washed as described above and labeled with anti–GPIX–Alexa Fluor 488 or anti–GPIX–Alexa Fluor 647 (2.5 to 5 μg/ml) for 15 min at room temperature, washed again, and injected into recipient mice via the retro-orbital plexus. In some experiments, platelets were treated ex vivo with OSGE (250 μg/ml, 60 min, 37°C) before labeling (33). Cleavage of the 45-kDa N-terminal of GPIbα was confirmed by flow cytometry using an anti-GPIbα antibody (clone Xia.G5) recognizing the N terminus. For DAPT experiments, P2ry12−/− platelets (64) were treated ex vivo with acetylsalicylic acid (2 mM) for 10 min at room temperature before labeling. In mice receiving two populations of platelets, the two populations were washed after labeling to remove residual antibody and then combined for a single injection. To achieve desired circulating platelet counts, the platelet count/ml was multiplied by 2.5 to estimate the total number of platelets for transfusion (~2-ml blood volume, +50% to account for platelet loss/sequestration). After transfusion, circulating platelet counts/ratios were confirmed by flow cytometry before starting saphenous vein laser injury experiments.

Saphenous vein laser ablation hemostasis model

Saphenous vein laser injury was performed as previously described (25), with modifications. Mice (8 to 16 weeks of age) were anesthetized by isoflurane inhalation (Veterinary Anesthesia Vaporizer, DRE Veterinary). Hair removal was performed using Nair, the saphenous vein was exposed, and the mouse was placed on the stage on a heating pad. A perfusion drip on the exposed saphenous vein was started and maintained at 37°C via a Sloflo In-line solution heater (SF-28) and a single-channel heater controller (model TC-324B, Warner Instruments) [physiologic salt solution containing 132 mM NaCl, 4.7 mM KCl, 1.2 mM MgSO4, 2 mM CaCl2, and 18 mM NaHCO3 (pH 7.4), which was bubbled with 5% CO2/95% N2 for 15 min before the start of the experiment]. Baseline fluorescence was recorded for 30 s using a Zeiss Axio Examiner Z1 upright microscope (Intelligent Imaging Innovations) equipped with a 20×/1 numerical aperture (NA) water immersion objective lens and a Hamamatsu Orca Flash 4.0 camera (Hamamatsu Photonics). Injury to the endothelium was initiated using an Ablate photoablation system equipped with an attenuable 532-nm pulse laser (Intelligent Imaging Innovations), and fluorescence intensity was recorded for 270 s (SPECTRA X light engine, Lumencor). All data were recorded and analyzed using SlideBook 6.0 software (Intelligent Imaging Innovations). Time to hemostasis was defined as the time from laser ablation to the visual cessation of blood flow from the injury site. Platelet adhesion at the injury site was determined by calculating the sum of fluorescence intensity in the 488 and 647 channels. To directly compare fluorescence intensities between experiments, all data points were normalized to the intensity at time to hemostasis for each individual injury, to account for variations in intensity of platelet labeling. For some conditions, hemostatic plug formation after laser ablation was imaged in real time using SDC microscopy [Yokogawa CSU-W1, equipped with a LaserStack (Intelligent Imaging Innovations) with 488 and 647 laser lines], essentially as described for epifluorescence imaging. Laser ablation was initiated, and real-time SDC microscopy was performed in a single plane at or superior to the position of the endothelium for up to 10 min. For each mouse, up to 10 injuries were performed along the saphenous vein, beginning proximal (downstream) and moving distal (upstream) so that subsequent injuries were not affected by embolizations from previous injuries. After real-time imaging, mice were euthanized by isoflurane overdose, and SDC Z-stacks of hemostatic plugs were acquired, from the most intraluminal point of the plug to the most extravascular.

Flow chamber assay

Flow chamber assay was performed essentially as previously described (65). Platelets in heparinized whole blood were labeled with Alexa Fluor–conjugated anti-GPIX antibody (2.5 μg/ml) and mixed at desired ratios just before starting the experiment. No double labeling of platelets was observed after mixing, as confirmed by flow cytometry. Collagen (200 μg/ml) was deposited on a glass slide, and the polydimethylsiloxane microfluidic device was oriented perpendicularly to the collagen strip. Blood was flowed at arterial shear rates (1600 s−1) for 5 min. Platelet adhesion was monitored using a Nikon TE300 inverted microscope equipped with a Hamamatsu Orca Flash 4.0 camera and a 100× oil objective and analyzed with NIS Elements software.

Flow cytometry

Flow cytometry was performed essentially as previously described (65). Platelet counts were determined in diluted whole blood using a BD Accuri C6 flow cytometer (BD Biosciences). In samples where platelets were not prelabeled, blood samples were first incubated with anti-GPIX antibody (2 μg/ml) for 10 min. For blood samples from transfused TP Tg mice, all platelets in blood were labeled by incubating with MWReg30-PE (2 μg/ml) for 10 min before analyzing the sample. Transfused platelets were gated on MWReg30-PE intensity, and WT or mutant populations were distinguished by anti–GPIX–Alexa Fluor 488 or 647 intensity. For determination of platelet integrin activation, diluted whole blood was incubated in Tyrode’s buffer with agonist (convulxin, PAR4p, or ADP) for 10 min in the presence of 1 mM Ca2+ and JON/A-PE (2 μg/ml). After activation, samples were diluted in phosphate-buffered saline (PBS) and immediately analyzed. Platelet populations were gated on anti–GPIX–Alexa Fluor 488 or 647 intensity, and JON/A-PE intensity was analyzed.


Venous blood was drawn from six healthy volunteers and seven patients previously diagnosed with GT (66), into buffered 3.2% sodium citrate. Aliquots of these citrated blood samples were saved for in vitro transfusion assays, and the rest of the volume was used for preparation of platelet concentrates using the buffy coat procedure. Blood samples from controls and GT patients were centrifuged at 1000g for 15 min, and the plasma supernatant and the buffy coat layer separated into different tubes. After resting for 15 min, the buffy coat was diluted 1:4 with autologous plasma and centrifuged at 150g for 7 min, and the upper phase containing the majority of platelets was separated. Platelet counts in these platelet concentrates from control (PC-C) or GT (PC-GT) (1 × 1012 to 2 × 1012/liter) were determined using a Sysmex hematological analyzer (Sysmex Corporation).

We performed in vitro transfusion by adding variable volumes of PC-C and PC-GT to the nonmanipulated control blood. The mixtures were then tested on the Impact-R cone and plate analyzer (Impact-R platelet analyzer 47600, Matis Medical Inc.), essentially as originally described (35). In brief, 130 μl of blood mixture, at least in duplicate, was placed into a polystyrene well, and a Teflon cone was placed on top and rotated (2050/s) to induce a shear stress that promotes platelet adhesion to the plastic. After washing the well and staining with May-Grünwald solution, the plate surface covered with platelets (% SC) was quantified using an inverted light microscope connected to a camera (Aptina MT9M0001 digital image sensor) and image analyzing software (Image Analysis software version 1.21) that calculates median values from seven images from each well.


Results are reported as means ± SEM, and statistical significance was assessed by analysis of variance (ANOVA) with post hoc analysis when appropriate, unless otherwise indicated. A P value less than 0.05 was considered significant. Original data are provided in data file S1.


Fig. S1. Integrin activation is similar between normal and transfused platelets.

Fig. S2. WT and Rasgrp2−/− platelets adhere at similar rates to the site of injury.

Data file S1. Raw data.

Movie S1. TP Tg mouse WT platelets only epifluorescence.

Movie S2. TP Tg mouse Rasgrp2−/− platelets only epifluorescence.

Movie S3. TP Tg mouse Rasgrp2−/−:WT platelets 5:1 epifluorescence.

Movie S4. TP Tg mouse Rasgrp2−/−:WT platelets 2:1 epifluorescence.

Movie S5. TP Tg mouse Rasgrp2−/−:WT platelets 2:1 SDC Z-stack.

Movie S6. TP Tg mouse WT:WT platelets 2:1 SDC Z-stack.

Movie S7. TP Tg mouse Rasgrp2−/−:WT platelets 2:1 RT SDC.

Movie S8. TP Tg mouse WT:WT platelets 2:1 RT SDC.

Movie S9. BSS-like mouse WT platelets epifluorescence.

Movie S10. TP Tg mouse WT:WT-OSGE platelets epifluorescence.

Movie S11. GT-like mouse WT platelets epifluorescence.

Movie S12. GT-like mouse WT platelets RT SDC.

Movie S13. TP Tg DAPT:WT platelets RT SDC.


Acknowledgments: We acknowledge D. Paul for expert technical assistance with SDC imaging experiments, K. Poe for assistance with mouse husbandry, M. Diaz-Ricart for assistance with human platelet experiments, and especially The C.T. and Nancy Owens Fund for a generous contribution made toward the purchase of the SDC confocal system. M.L.L. and J.R. are the coordinators of the multicentric project “Functional and Molecular Characterization of Patients with Inherited Platelet Disorders” Spanish Society of Thrombosis and Haemostasis [Hemorrhagic Diathesis Working Group, Spanish Society for Hemostasis and Thrombosis (SETH)]. Funding: This work was supported by NIH grants 1R35 HL144976-01 to W.B. and T32HL007149 to R.H.L. M.L.L. and J.R. were supported by grants from Instituto de Salud Carlos III and Feder (PI17/01311 and CB15/00055) and Fundación Séneca (19873/GERM/15). V.P.-B. was supported by a CIBERER fellowship. Author contributions: R.H.L. designed and performed experiments, analyzed data, performed statistical analysis, and wrote the manuscript. R.P. designed and performed experiments. A.D. designed and performed experiments. M.L.L. designed and performed experiments and analyzed data. V.P.-B. designed and performed experiments. J.R. designed experiments, analyzed data, and edited the manuscript. W.B. designed and supervised experiments, analyzed data, and wrote the manuscript, which was approved by all authors. Competing interests: All authors declare that they have no competing interests. Data and materials availability: All data associated with this study are available in the main text or the Supplementary Materials. Rasgrp2−/− mice are available from A. Graybiel under a material agreement with the Massachusetts Institute of Technology. Talin1f/f mice are available from M. Ginsberg under a material agreement with the University of California at San Diego. P2ry12−/− mice are available from P. Conley under a material agreement with Portola Pharmaceuticals. hIL-4Rα/GPIbα mice are available from J. Ware under a material agreement with the University of Arkansas at Little Rock.

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