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Articles

RAP1-RHO small GTPase cross-talk mediates integrindependent and -independent platelet procoagulant response

Department of Biochemistry and Biophysics, University of North Carolina at Chapel Hill, NC, USA; Blood Research Center, University of North Carolina at Chapel Hill, NC
Department of Biochemistry and Biophysics, University of North Carolina at Chapel Hill, NC, USA; Blood Research Center, University of North Carolina at Chapel Hill, NC
Department of Biochemistry and Biophysics, University of North Carolina at Chapel Hill, NC, USA; Blood Research Center, University of North Carolina at Chapel Hill, NC
Department of Medicine, University of California San Diego, CA
Department of Pharmacology, Lineberger Comprehensive Cancer Center, University of North Carolina at Chapel Hill, NC
Department of Biochemistry and Biophysics, University of North Carolina at Chapel Hill, NC, USA; Blood Research Center, University of North Carolina at Chapel Hill, NC
Department of Biochemistry and Biophysics, University of North Carolina at Chapel Hill, NC, USA; Blood Research Center, University of North Carolina at Chapel Hill, NC
Haematologica Early view Mar 5, 2026 https://doi.org/10.3324/haematol.2025.288459

Abstract

Platelet adhesion and procoagulant activity are critical for primary and secondary hemostasis, respectively. The small GTPase RAP1 is a central regulator of platelet aggregation as it controls αIIbβ3 integrin activation through direct interaction with the integrin adapter protein, TALIN1 (TLN1). In addition to their aggregation defect, activated platelets lacking RAP1 (Rap1mKO) exhibited a marked impairment in surface exposure of phosphatidylserine (PtdSer), a negatively charged phospholipid with procoagulant activity. However, the mechanisms by which RAP1 regulates PtdSer exposure are unclear. Here we investigated the hypothesis that RAP1 regulates platelet PtdSer exposure through cross-talk with small GTPases of the RHO family. Consistent with their defect in PtdSer exposure, Rap1mKO platelets showed reduced procoagulant activity in vitro and in vivo when compared to controls. Stimulated Rap1mKO platelets exhibited elevated RHOA-GTP levels, and inhibition of the RHOA effector, Rho associated coiled-coil kinase (ROCK), partially restored PtdSer exposure in these cells. A milder defect in PtdSer exposure was observed for platelets from Tln1mR35/118E mice, i.e., mice with impaired RAP1-TLN1 interaction but otherwise intact RAP1 signaling. ROCK inhibition fully restored PtdSer exposure in Tln1mR35/118E platelets. Opening of the mitochondrial permeability transition pore, a cellular response critical to PtdSer exposure, was impaired in Rap- 1mKO platelets and restored by pretreatment of cells with the ROCK inhibitor. Our study provides first evidence that platelet RAP1 signaling affects hemostatic plug formation independent of its key role in platelet adhesion. Additionally, our studies strongly suggest that RAP1 regulates PtdSer exposure and procoagulant activity in a RHOA/integrin-dependent and -independent manner.

Introduction

Within a hemostatic plug there are two classifications of activated platelets: proadhesive and procoagulant.1 Proadhesive platelets are characterized by high affinity integrin receptors, such as αIIbβ3, which mediate platelet adhesion to the site of injury and platelet aggregation via binding of fibrinogen and other ligands.2 Integrin-mediated platelet adhesion is essential to formation of the initial platelet plug to cease bleeding. Procoagulant platelets facilitate thrombin generation and thus enhance the formation of fibrin, a fibrous protein critical for hemostatic plug stability.

Characteristic to procoagulant platelets is exposure of the negatively charged phospholipid, phosphatidylserine (Ptd-Ser), on the cell surface.3 Procoagulant platelet formation is dependent on sustained high cytosolic calcium levels and mitochondrial depolarization, mediated by the opening of the mitochondrial permeability transition pore (MPTP) via its essential adaptor protein cyclophilin D (CypD).4,5 Depolarization of the mitochondria ultimately allows the scramblase transmembrane protein 16F (TMEM16F) to flip PtdSer to the outer plasma membrane leaftlet.6,7

Platelet activation is a tightly controlled process with small GTPases playing a central role.8-10 Small GTPases are molecular switches which are active in their GTP-bound state and inactive in their GDP-bound state. When GTP bound, small GTPases undergo conformational changes to interact with effector proteins.11 The role of small GTPases in αIIbβ3 integrin activation is well-defined, while their activity in procoagulant platelet formation remains less explored. The Ras family GTPase, RAP1, is the most abundant small GTPase in platelets.9 RAP1 is a central regulator of platelet adhesion/aggregation as loss of both isoforms (RAP1a and RAP1b) markedly impairs integrin activation, resulting in significantly prolonged bleeding times.12 Activation of αIIbβ3 integrin requires direct interaction between RAP1 and its effector protein TLN1.13-15 Disruption of the interaction between RAP1 and TLN1 (Tln1mR35/118E) results in loss of βIIbβ3 integrin activation while retaining other RAP1 signaling responses.13 Ligand binding to βIIbβ3 integrin induces outside-in signaling, an important mechanism to potentiate platelet activation.16,17 Loss of RAP1 also results in impaired platelet procoagulant response,13 although the mechanism of RAP1-mediated PtdSer exposure is unclear. The RHO family GTPase, RHOA, is primarily known for its role in cytoskeletal rearrangements.10,18 Activation of RHOA occurs downstream of G13-coupled receptors, whereas inhibition of RHOA results from integrin outside-in signaling.19,20 In contrast to RAP1, inhibition of RHOA-ROCK (Rho associated coiled-coil kinase) signaling leads to increased PtdSer exposure.21 Crosstalk between RAP1 and RHOA was demonstrated for other cell types;22,23 however, the potential role of RAP1-RHOA crosstalk in platelet procoagulant response remains unexplored.

In the present study, we investigated the mechanism of RAP1-mediated PtdSer exposure in platelets. Our studies demonstrate that RAP1-mediated PtdSer exposure occurs through integrin-dependent and -independent mechanisms, and that the integrin-dependent mechanism occurs through a connection to RHOA.

Methods

Mice

Generation of Rap1mKO,12, Tln1mR35/118E,13, IL4Rβ-GPIbβ-tg24 and CypD/25 mice has been previously described. Experimental procedures were approved by the Institutional Animal Care and Uses Committee.

Four-dimensional saphenous vein laser injury model

Adoptive transfer of platelets into thrombocytopenic mice was performed as previously described.26,27 In brief, blood was collected in phospate-buffered saline with heparin and platelets were washed.26 Platelets were depleted with β-hIL4R antibody (2.5 μg/g body weight) in IL4Rα/GPIbαtg mice. Washed platelets were labeled with AlexaFluor (AF)488 or AF647-labeled antibodies to GPIX and administered via retroorbital injection into platelet-depleted IL4Rα/GPIbα-tg mice to a final circulating concentration of 2-5x108 platelets/mL. Four-dimensional (4D) imaging of saphenous vein laser injury was performed as recently described.27 In brief, the saphenous vein was exposed and relevant fluorescently labeled antibodies were administered via retroorbital injection. Laser ablation was used to create a perforating injury in the vein and spinning disk confocal imaging was performed on a Zeiss Axio Examiner Z1 inverted spinning disk confocal microscope (4x4 binning, 7.5 µm step, 150 µm total travel). Images were acquired with SLIDEBOOK 6.0 software (Intelligent Imaging Innovations). Image analysis was performed using ImageTank software (Visual Data Tools, Inc) as previously described.28

Human blood collection

Whole blood was collected from healthy subjects (male and female subjects between the ages of 20 and 50 years who had not taken aspirin/non-steroidal anti-inflammatory drugs within 2 weeks) using a 21-gauge needle vacutainer butterfly into 3.2% citrate tubes (BD). Blood collection from healthy donors was performed with informed consent in accordance with a protocol approved by the Institutional Review Board at the University of North Carolina at Chapel Hill.

Flow cytometry

Platelets were washed as previously described12 and activated with the indicated concentrations of convulxin (CVX; purchased from Kenneth Clemetson, Theodor Kocher Institute, University of Bern, Bern, Switzerland) and PAR4p (GL Biochem) in the presence of 2 µg/mL JON/A-PE (clone Leo. H4, Emfret Analytics), α-P-selectin-FITC (clone RB40.34; BD Biosciences), and Annexin V-AF647 (generous gift from Sriram Krishnaswamy, Children’s Hospital of Philadelphia). Where indicated, samples were incubated with ROCK inhibitor (20 μM Y-27632, Tocris) for 10 minutes (min) prior to activation. After 15 min of incubation, samples were diluted and analyzed via flow cytometry (Accuri C6 Plus flow cytometer; BD Biosciences).

RAP1 and RHOA activation assays

Washed platelets (260 μL samples at 8x108/mL) were stimulated with 50 ng/mL CVX and 250 μm PAR4p in aggregometry. Platelets were lysed with cold 2x lysis buffer (100 mmol/L Tris/HCl pH 7.4, 400 mmol/L NaCl, 5 mmol/L MgCl2, 2% Nonidet P-40, 20% glycerol and protease inhibitor cocktail lacking ethylenediaminetetraacetic acid; Roche). Lysates were incubated with RalGDS-RBD beads for RAP1-GTP (Millipore, Billerica, MA) or Rhotekin-RBD beads for RHOA-GTP (Cytoskeleton) for 1 hour at 4°C. Beads were washed three times then resuspended in 2x Laemmli buffer for detection of RAP1-GTP or RHOA-GTP via standard western blotting procedure. For loading controls, 50 μL of the platelet sample was combined with 50 μL 2x Laemmli buffer (75 mmol/L Tris/HCl, pH 6.8, 2% sodium dodecyl sulfate, 10% glycerol, 5% 2-mercaptoethanol, 0.002% bromophenol blue). Antibodies to RAP1 (clone 121; Santa Cruz, cat # sc-65) or RHOA (clone 55; Sigma, cat # 05-778) were used for detection of GTP-bound and total RAP1 or RHOA, respectively.

Thrombin generation assay

Thrombin generation assay was performed in a 96-well plate; 0.5 pM TF (Innovin) and 200 ng/mL CVX were added, and wells were brought to volume (30 μL) with Tyrodes buffer containing 1 mM CaCl2. Platelet-poor plasma (PPP) or platelet-rich plasma (PRP, platelet count 5x108/mL) was added to initiate the reaction. Each sample was calibrated with α2-macroglobulin-thrombin complex calibrator (Diagnostica Stago Inc Fluca Kit). Fluorescence was measured on a Fluoroskan Ascent fluorometer (Thermo Fisher Scientific, Waltham, MA) with the Ascent Software (version 2.6, Thermo Fisher Scientific) at 37°C.

JC-1 mitochondrial depolarization assay

Platelets were washed and adjusted to 7.5x108/mL. JC-1 dye (1 μL; Invitrogen,) and washed platelets (20 μL) were added to 180 μL of Tyrode’s buffer with bovine-serum albumin and incubated at 37°C for 10 min; 30 μL of labeled platelets were transferred to 70 μL of Tyrode’s buffer containing 2 mM CaCl2. Platelets were activated with 100 μL of 2x agonist and 2.5 μg/mL Annexin V-AF647 in Tyrode’s buffer containing 2 mM CaCl2, and fluorescence intensities were recorded on a BD Accuri C6 Flow Cytometer. Analysis was completed using FlowJo (FlowJo LLC).

Fluo-4 calcium mobilization assay

Platelets were washed and adjusted to 1x109/mL. Fluo-4 dye (Life Technologies) was diluted to a concentration of 0.5 mM in dimethyl sulfoxide (DMSO). Platelets (1x108/mL in 200 μl Tyrode’s buffer) were labeled with 1 μL of Fluo-4 dye for 30 min at 37°C. After incubation, 800 μL of Tyrode’s buffer was added to labeling reaction. Labeled platelets were diluted (1:1) in Tyrode’s buffer with 4 mM CaCl2. Cellular stimulation was induced by addition of a 2x agonist solution. Fluorescence intensity (Fl1) was recorded on a BD Accuri C6 Flow Cytometer. Analysis was completed using FlowJo (FlowJo LLC).

Statistics

Results are shown as mean +/- standard error of the mean (SEM). Unless otherwise indicated, statistical significance was analyzed via Welch’s t test.

Results

Decreased phosphatidylserine exposure and thrombin generation potential in RAP1-deficient platelets in vitro

Dual-agonist stimulation of platelet glycoprotein (GP)VI and protease-activated receptor (PAR)4 results in robust PtdSer exposure in vitro.3 Therefore, procoagulant response was stimulated with convulxin (CVX; GPVI) and PAR4-activating peptide (PAR4p) and measured via Annexin V binding using flow cytometry (% PtdSer+ events). Compared to controls, exposure of PtdSer was significantly impaired in platelets lacking RAP1 (Rap1mKO) activated with CVX/PAR4p (Figure 1A). Using a modified thrombin generation assay29 in which activated procoagulant platelets in PRP provide the negatively charged phospholipid surface required for coagulation factor assembly, we evaluated the contribution of RAP1-mediated PtdSer exposure to thrombin generation in vitro. Thrombin generation was minimal in PPP when compared to PRP samples (Figure 1B, C). Congruent with the defect in PtdSer exposure, thrombin generation, defined as peak thrombin (nM), in Rap1mKO PRP was significantly reduced when compared to control PRP (Figure 1B, C); however, there was no significant difference in time to peak (Figure 1D).

Figure 1.RAP1-mediated phosphatidylserine exposure contributes to thrombin generation in vitro. (A) Flow cytometry analysis of procoagulant response (percent [%] phosphatidylserine-positive [PtdSer+] events [Annexin V binding]) in control or Rap1mKOplatelets stimulated with 50 ng/mL CVX + 250 μM Par4p; N=9. (B-D) Thrombin generation. (B) Peak thrombin (nM) levels for control platelet-rich plasma (PRP), Rap1mKO PRP, and platelet-poor plasma (PPP). (C) Representative curves for indicated samples. (D) Time to peak thrombin generation for control and Rap1mKO PRP; N=7-8. ***P<0.001; ****P<0.0001; NS: not significant; mKO: megakarycyte/ platelet-specific knockout.

Impaired procoagulant activity of RAP1-deficient platelets in vivo

We recently developed a new imaging model to quantify platelet procoagulant activity during hemostatic plug formation in vivo.27 In this model, perforating injuries ~50 µm in diameter are induced to the saphenous vein by laser injury, and the three-dimensional accumulation of platelets and fibrin is monitored in real time by spinning disk confocal microscopy. Using this model, we were able to demonstrate that fibrin accumulation is significantly impaired in mice lacking CypD in platelets only, demonstrating that platelets are the main procoagulant cellular surface during hemostatic plug formation.27 One limitation of the model is difficulties visualizing and measuring hemostatic plug components in mice with excessive bleeding, including Rap1mKO mice. To circumvent this limitation and determine the contribution of RAP1 to platelet procoagulant activity in vivo, we used an adoptive platelet transfer strategy to generate mice with specific defects in platelet function without excess bleeding: (i) mice that received a mixture of CypD-/- and wild-type (WT) platelets at a ratio of three to one, and (ii) mice that received a mixture of CypD-/- and Rap1mKO platelets at a ratio of three to one. CypD-/- platelets were co-transfused to facilitate hemostatic plug formation in the context of RAP1-deficiency (Online Supplementary Figure S1). CypD-/- platelets, however, have minimal procoagulant activity in this model.27 Thus, fibrin accumulation at sites of injury in co-transfused mice would depend on the procoagulant activity of WT or Rap1mKO platelets. As shown in Figure 2, fibrin accumulation was significantly reduced in mice transfused with CypD-/-/Rap1mKO platelets when compared to mice transfused with CypD-/-/WT platelets. Together, these studies demonstrate a critical role for RAP1 in platelet procoagulant activity in vitro and in vivo.

Integrin-dependent and -independent mechanisms of RAP1-mediated phosphatidylserine exposure

Given the documented role of integrin outside-in signaling in PtdSer exposure,30 we next determined whether the defect in platelet procoagulant response observed in Rap1mKO platelets is secondary to their defect in αIIbβ3 integrin activation. We compared the response to dual-agonist stimulation in Rap1mKO platelets to platelets from mice with impaired RAP1-TLN1 interaction (Tln1mR35/118E). Consistent with previous studies with single agonists, both Rap1mKO and Tln1mR35/118E platelets exhibited a marked defect in αIIbβ3 integrin activation (JON/A-PE binding) in response to dual-agonist stimulation (Figure 3A, D), while granule secretion (CD62P surface expression) was not impaired (Figure 3B, E). Dual agonist-induced PtdSer exposure was also significantly impaired in Rap1mKO and Tln1mR35/118E platelets (Figure 3C, F). Interestingly, PtdSer exposure was reduced by 85% in Rap1mKO platelets, while only a 48% reduction in PtdSer-positive events was observed for Tln1mR35/118E platelets compared to controls. Thus, these studies suggested that RAP1 facilitates platelet PtdSer exposure by integrin-dependent and -independent mechanisms.

Integrin outside-in signaling negatively regulates the activation state of the small GTPase RHOA,31 and previous work demonstrated that inhibition of RHOA signaling leads to increased PtdSer exposure in platelets.21 Thus, we hypothesized that impaired PtdSer exposure in Rap1mKO platelets results, at least in part, from elevated RHOA activity. Consistent with this hypothesis, we observed significantly increased RHOA-GTP levels in Rap1mKO and Tln1mR35/118E platelets at 2 min after addition of agonists (Figure 4A, B). We next studied dual agonist-induced platelet activation in Rap1mKO and Tln1mR35/118E platelets pretreated with an inhibitor of the RHOA effector protein, ROCK.21 Treatment with the ROCK inhibitor (Y-27632) did not significantly affect granule secretion or αIIbβ3 activation in controls, Rap1mKO, or Tln1mR35/118E platelets (Online Supplementary Figure S2). However, inhibition of ROCK partially recovered PtdSer exposure in Rap1mKO platelets (Figure 4C). Importantly, ROCK inhibition fully restored PtdSer exposure in Tln1mR35/118E platelets (Figure 4D). Previous studies demonstrated that sustained RAP1 activation in human and murine platelets depends on feedback activation via the ADP receptor, P2Y12.9 We thus studied PtdSer exposure in platelets pretreated with the P2Y12 inhibitor, 2-MeSAMP. Following dual agonist stimulation, PtdSer exposure was significantly reduced in murine (Online Supplementary Figure S3A) and human (Online Supplemental Figure S3B) platelets pretreated with 2-MeSAMP when compared to vehicle treated controls. The defect in PtdSer exposure induced by 2-MesAMP was reversed by preincubation of platelets with Y-27632.

Figure 2.Reduced procoagulant activity of Rap1mKO platelets in vivo. (A) Fibrin accumulation at sites of laser injury in mice transfused with CypD-/- and Rap1mKO platelets (N=21 injuries; 3 mice) compared to mice transfused with CypD-/- and wild-type platelets N=20; 3 mice). Data are shown as sum fluorescence intensity +/- standard error of the mean; *P<0.05. (B) Representative images f hemostatic plugs in mice transfused with CypD-/- (purple) and wild-type (grey) or CypD-/- (purple) and Rap1mKO (red) platelets. ibrin is shown in cyan. Images are presented with side and bottom views at indicated time points. min: minutes; mKO: mega-karyocyte/platelet-specific knockout.

Calcium mobilization is not affected in Rap1mKO platelets

One of the key factors to successful platelet procoagulant response is a robust and sustained increase in cytosolic calcium levels, which is required to trigger mitochondrial depolarization.3,5 To determine cytosolic calcium levels, control or Rap1mKO platelets were labeled with the calcium-sensitive dye, Fluo-4,5 and activated in the presence or absence of ROCK inhibitor. Dual-agonist stimulation resulted in sustained high calcium levels in control and Rap1mKO platelets, both in the presence and absence of ROCK inhibitor (Figure 5A). The integrated calcium signal (area under the curve) was comparable between Rap1mKO and control platelets. Slightly increased calcium levels were observed for both control and Rap1mKO platelets activated in the presence of ROCK inhibitor (Figure 5B). Under these experimental conditions, Rap1mKO platelets had decreased PtdSer exposure compared to controls; and inhibition of RHOA/ROCK signaling led to a significant increase in Ptd-Ser-positivity for control and Rap1mKO platelets, similar to Figure 4C. Together, these studies suggest that altered calcium mobilization does not account for the observed differences in PtdSer exposure observed in Rap1mKO platelets.

Mitochondrial depolarization is decreased in Rap1mKO platelets and partially recovered by inhibition of RHOA/ ROCK signaling

Opening of the MPTP and mitochondrial depolarization are critical events for platelet procoagulant response.4,32 We next studied mitochondrial depolarization in dual agonist-stimulated platelets using the JC-1 flow cytometry-based assay.33 Mitochondrial depolarization was significantly impaired in Rap1mKO platelets when compared to controls (Figure 6A, B). Pretreatment with ROCK inhibitor led to markedly increased mitochondrial depolarization in both control and Rap1mKO platelets (Figure 6A, B).

Figure 3.RAP1 affects platelet phosphatidylserine exposure via integrin independent and dependent mechanisms. (A-C) Flow cytometry analysis of αIIbβ3 activation (JON/A-PE MFI normalized to platelet size) (A), granule secretion (α-CD62P-FITC mean fluorescense intensity (MFI) normalized to platelet size) (B), and phosphatidylserine (PtdSer) exposure (Annexin V-AF647, % positive events) (C) for control or Rap1mKO platelets stimulated with 50 ng/mL CVX + 250 μM Par4p (N=6). (D-F) Flow cytometry analysis of αIIbβ3 activation (JON/APE) (D), granule secretion (α-CD62P-FITC) (E), and PtdSer exposure Annexin V-AF647 (F) for control or Tln1mR35/118E platelets stimulated with 50 ng/mL CVX + 250 [xM Par4p (N=8). *P<0.05; **P<0.01; **P<0.001; NS: not significant; mKO: megakaryocyte/platelet-specific knockout.

Discussion

Procoagulant platelets, i.e., platelets exposing PtdSer on their outer plasma membrane, are critical to thrombin/ fibrin generation and hemostatic plug stability. Small GT-Pases of the RAS (RAP1) and RHO (RHOA, RAC1) families are known regulators of PtdSer exposure in platelets. Our study provides first evidence for RAP1-RHO cross-talk required for platelet PtdSer exposure and platelet-dependent thrombin generation, both in vitro and in vivo (Figure 7). RAP1 is an essential regulator of αIIbβ3 integrin-mediated platelet adhesion and hemostatic plug formation.12,13 While no patients with loss-of-function mutations in RAP1 have been identified, mutations in CalDAG-GEFI (RASGRP2), an important regulator of RAP1 activation, are known. These mutations cause moderate to severe bleeding in humans,34-36 similar to what was shown for mice deficient in RAP1 or CalDAG-GEFI.12,37 As outlined above, RAP1 signaling is also important for procoagulant platelet formation. Importantly, bleeding is also observed in patients with Scott syndrome, i.e., in patients with a defect in procoagulant platelet formation but not αIIbβ3 integrin activation.38 Scott syndrome is attributed to mutations in TMEM16F, a phospholipid scramblase that mediates PtdSer translocation to the outer membrane layer.7,39 TMEM16F is highly expressed in platelets but is also found in other cell types including erythrocytes and endothelial cells (EC).40-42 The similar bleeding phenotype between TMEM16F germline and megakaryocyte/platelet-specific knockout mice suggests that impaired procoagulant activity in platelets is the main reason for bleeding observed in Scott syndrome patients.6,7,40 Furthermore, we recently demonstrated a key role of procoagulant platelets, but not EC for thrombin/fibrin formation during hemostatic plug formation.27 Here we provide the first evidence that RAP1-mediated platelet PtdSer exposure contributes to thrombin/fibrin generation in vitro and in vivo. These studies suggest that bleeding in patients with impaired RAP1 signaling may result in part from reduced thrombin/fibrin formation at sites of vascular injury.

Figure 4.Integrin-dependent RAP1-RHOA connection regulates phosphatidylserine exposure. (A) Active (GTP-bound) RHOA over total RHOA in control (black bars) or Rap1mKO platelets (red bars) activated for the indicated times with 50 ng/mL CVX + 250 μM Par4p (N=5). Percentages normalized to highest value. (B) Active (GTP-bound) RHOA over total RHOA in control (black bars) or Tln-1mR35/118E platelets (blue bars) activated for the indicated times with 50 ng/mL CVX + 250 μM Par4p (N=5). Percentages normalized to highest value. (C) Flow cytometry analysis of Annexin V binding % PtdSer-positive events on control or Rap1mKO platelets pretreated with ROCK inhibitor (ROCKi, 20 μM Y-27632) and then stimulated with 50 ng/mL CVX + 250 μM Par4p (N=6). (D) Flow cytometry analysis of Annexin V binding (% PtdSer+ events) on control or Tln1mR35/118E platelets pretreated with ROCKi and then stimulated with 50 ng/mL CVX + 250 μM Par4p (N=8). *P<0.05; **P<0.01; ***P<0.001; NS: not significant. Data shown for untreated samples in panels (C) and (D) are the same as those in Figures 3C and F, respectively. They were included in Figure 4 to better illustrate the effect of ROCKi on PtdSer exposure. sec: seconds; min: minutes; mKO: megakaryocyte/platelet-specific knockout.

Figure 5.Calcium mobilization is not affected in Rap1mKO platelets. (A) Representative curves for Fluo-4 fluorescence (FITC-H) in control (black and grey curves) or Rap1mKO platelets (red and teal curves) activated in the absence or presence of ROCK inhibitor (ROCKi, Y-27632) with 50 ng/mL CVX + Par4p 150 μM (added at 30 seconds; indicated by arrow). (B) Area under the curve analysis for Fluo-4 fluorescence traces described in (A) (N=5). **P<0.01; NS: not significant; mKO: megakaryocyte/platelet-specific knockout.

Figure 6.Mitochondrial depolarization is decreased in Rap1mKO platelets and partially recovered with ROCK inhibitor. (A) JC-1 fluorescence ratio (FITC-H/PE-H) for control (black and grey curves) or Rap1mKO platelets (red and teal curves) activated in the absence or presence of ROCK inhibitor (ROCKi, Y-27632) with 50 ng/mL CVX + Par4p 150 μM (added at 10 seconds [sec]; indicated by arrow). (B) Area under the curve (AUC) analysis (N=5). *P<0.05; **P<0.01; ***P<0.001; NS: not significant. min: minutes; mKO: megakaryocyte/platelet-specific knockout.

Figure 7.Summary diagram illustrating the crosstalk between signaling pathways discussed in this manuscript. PAR4, P2Y12, GPVI: agonist receptors; RAP1, RHOA, RAC1: small GTPases; Y-27632: ROCK inhibitor; 2MeSAMP: P2Y12 inhibitor; CypD: cyclophilinD; PtdSer: phosphatidylserine; ADP: adenosine diphosphate; Rap1mKO: megakaryocyte/platelet-specific knockout of Rap1a and Rap1b isoforms; Tln1mR35/118E: mice expressing Talin1 variant in megakaryocytes/platelets only. This figure was created in BioRender.

The RHO family of GTPases plays an important role in platelet procoagulant response. Kunzelmann et al. showed that inhibition of RHOA reduces store-operated calcium entry and PtdSer exposure.43 As discussed in their paper, this effect of RHOA inhibition is through the reorganization of actin cytoskeleton, but not through the ROCK pathway. Dasgupta et al. described increased procoagulant function for platelets deficient in ROCK1, but no alterations in other cellular functions, including calcium response and shape change. Thus, the RHOA/ROCK pathway inhibits PtdSer exposure21,30,44 and needs to be downregulated by integrin outside-in signaling during platelet procoagulant response.30,31 We observed prolonged RHOA activation in Rap1mKO and Tln1mR35/118E platelets, i.e., platelets with markedly impaired integrin inside-out activation. Our studies further established that inhibition of RHOA/ROCK signaling fully restored PtdSer exposure in Tln1mR35/118E platelets, which are defective in RAP1-TLN1 interaction. However, only a partial recovery of PtdSer exposure by inhibition of RHOA/ROCK signaling was observed in Rap1mKO platelets, a finding that suggests a TLN1/integrin/RHOA-independent contribution of RAP1 to platelet procoagulant response. We previously showed that in platelets RAP1 positively regulates the activity of RAC1,12,45 another RHO GTPase with a documented role in PtdSer exposure.21 Importantly, RAC1 and RHOA are also known to negatively regulate each other’s activation state.46 Together, these studies suggest an important role for cross-talk between RAP1 and RHO GTPases in integrin-dependent and -independent PtdSer exposure.

High sustained cytosolic calcium levels and mitochondrial depolarization are two critical steps in procoagulant platelet formation. Inhibition of RHOA/ROCK signaling led to a small but significant increase in cytosolic calcium and a marked increase in mitochondrial depolarization in both control and Rap1mKO dual agonist-stimulated platelets. Compared to controls, a significant decrease in mitochondrial depolarization but not calcium mobilization was observed for dual agonist-stimulated RAP1-deficient platelets. Given that calcium mobilization was not altered in Rap1mKO platelets, another mechanism must account for the decreased mitochondrial depolarization. Reactive oxygen species also affect mitochondrial depolarization, and both RHOA and RAC1 have been shown to regulate ROS production.47 Whether impaired mitochondrial depolarization and PtdSer exposure in Rap1mKO platelets are caused by dysregulated ROS production will be a topic of future studies.

In conclusion, our studies demonstrate that impaired RAP1 signaling leads to decreased platelet procoagulant response and thrombin generation in vitro and in vivo. RAP1 affects PtdSer exposure via integrin-dependent and -independent mechanisms, which likely involve cross-talk with RHO GTPases and the depolarization of the MPTP. This study improves our understanding of the role of small GTPases in platelet procoagulant response and thus may have important implications for the development of better therapies to prevent bleeding or thrombosis.

Footnotes

  • Received July 8, 2025
  • Accepted February 25, 2026

Correspondence

°Current address NZ: Institute of Experimental Biomedicine I, University Hospital and Rudolf Virchow Center Würzburg, University of Würzburg, Würzburg, Germany

W. Bergmeier bergmeie@email.unc.edu

Disclosures

No conflicts of interest to disclose.

Contributions

AB-K designed research, performed research, analyzed data and wrote manuscript. NZ and WS performed research and analyzed data. MHG contributed vital regents. AS designed and performed research. RHL designed research, performed research and analyzed data. WB designed research and wrote manuscript.

Funding

This work was supported by the National Institutes of Health, National Heart, Lung, and Blood Institute (grants R35 HL144976 [to WB], F31 HL165935 [to AB-K], P01 HL151433 [to MHG and WB])

Acknowledgments

The authors thank David Paul and Summer Jones (University of North Carolina at Chapel Hill) for valuable technical support and valuable expertise.

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DOI
https://doi.org/10.3324/haematol.2025.288459
Pubmed
41784022
 
Published
2026-03-05
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Ferrata Storti Foundation, Pavia, Italy
Print ISSN
0390-6078
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1592-8721
 
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