Role for ADAP in shear flow–induced platelet mechanotransduction

During hemostasis, platelets must mount a strong and rapid response under a variety of hydrodynamic shear stresses.¹  Bidirectional signaling involving integrin αIIbβ3 is particularly important for hemostasis.^1,2  Once platelets contact the damaged vessel wall, the ligand binding function of αIIbβ3 is activated by “inside-out” signals that stabilize adhesion and initiate platelet aggregation. In turn the binding of multivalent adhesive ligands, such as fibrinogen, to αIIbβ3 triggers “outside-in” signals that promote platelet cytoskeletal rearrangements, spreading, and optimal thrombus formation. Although the conversion of force into biochemical signals (mechanotransduction) in response to hydrodynamic shear stresses has been extensively studied in endothelial cells and demonstrated to elicit activation of ion channels, extracellular signal-regulated kinases, and rho GTPases,³  there is still debate as to whether and how force transmission occurs via adhesion receptors and cytoskeletal elements in other adherent cells, including platelets.^4,5  A few careful studies of platelets have pointed to shear-dependent roles for proteins, such as phosphoinositide 3-kinase,⁶  P2Y₁,⁷  and α-actinin⁸  at high arterial or pathologic shear. Overall, however, mechanotransduction in platelets, and the role of αIIbβ3 in this process in particular, remain poorly understood.

Adhesion- and degranulation-promoting adapter protein (ADAP) is a hematopoietic-specific protein that promotes cytokine production, proliferation, and integrin-mediated adhesion after stimulation of T lymphocytes through the T-cell receptor.^9,10  In these cells, ADAP forms a signaling module by binding to the SH3 domain of SKAP-55¹¹  and can bind this domain within the SKAP-55 homologue, SKAP-HOM, as well.¹²  Of note, SKAP-55 interacts with RIAM, a Rap1 effector, to promote increases in integrin affinity.¹³  ADAP also possesses binding sites for the interaction domains of several other proteins, among them the SH2 domains of SLP-76 and Fyn, the EVH1 domain of VASP, and the MAGUK region of CARMA1,^10,14  and ADAP can bind phosphoinositols through 2 helically extended SH3 domains.¹⁵  In mouse platelets, genetic deletion of ADAP reduces but does not eliminate inside-out activation of αIIbβ3 in response to von Willebrand factor (VWF) binding to GP Ib-IX-V or ADP/thrombin binding to G protein–coupled receptors.¹⁶  ADAP localizes to the periphery of human platelets spread on fibrinogen.¹⁷  In the present study, we provide evidence that ADAP is a critical component of αIIbβ3-mediated outside-in signaling by virtue of its regulation of the platelet actin cytoskeleton in the face of hemodynamic shear stresses.

Rhodamine phalloidin was from Molecular Probes/Invitrogen. Rabbit polyclonal antibodies against Vav1 pTyr-174 were from Abcam and Santa Cruz Biotechnology. Rabbit polyclonal antibodies against VASP, c-Src pTyr-418, and SLP-76 were from Alexis Biochemicals, Biosource/Invitrogen, and Cell Signaling Technology, respectively. Mouse monoclonal antibody against vinculin was from Sigma-Aldrich. Rat monoclonal antibody against GP IX was from Emfret Analytics. A polyclonal sheep antibody against murine ADAP was a kind gift from Gary Koretzky (University of Pennsylvania). Dimeric, murine A1A2 VWF (dmA1A2 VWF) was prepared as described.¹⁶  SuperSignal WestPico reagent was from Pierce Chemicals. All other reagents were from Sigma-Aldrich.

Mice deficient in ADAP, SKAP-HOM, or VASP have been described^18-21  and were obtained from Gary Koretzky, Ben Neel, (Ontario Cancer Institute), and Alexander Clowes (University of Washington), respectively. ADAP^+/+, SKAP-HOM^+/+, and VASP^+/+ mice represent littermate controls. All mouse studies were conducted with Institutional Animal Care and Use Committee approval from the University of California, San Diego, including in vivo thrombosis studies and ex vivo studies with mouse blood.

Thrombus formation in the mouse carotid artery was induced with 3.75% anhydrous ferric chloride (FeCl₃) as described.²²  The carotid artery was subsequently excised, fixed in 4% formaldehyde for 4 hours at 4°C, transferred to 70% ethanol, and paraffin embedded, and sections were mounted on slides and stained with hematoxylin and eosin. Histology was examined by the use of a Leica DMLS microscope fitted with a Leica Plan 20×/0.4 numeric aperture (NA) objective, and SPOT software (Diagnostic Instruments).

Mouse blood preparation is described in the supplemental Methods (available on the Blood website; see the Supplemental Materials link at the top of the online article). Whole blood was diluted 1:1 with Walsh buffer to permit quantitative studies of platelets on the matrix surface, and 10 μg/mL mepacrine was added to label platelets for adhesion studies. Glass coverslips were coated overnight at 4°C, with either 50 μg/mL fibrinogen or 10 μg/mL dmA1A2 VWF domain fragment, matrices that are engaged primarily by adhesion receptors αIIbβ3 and GPIb-IX-V, respectively. After rinsing with HEPES buffer (20mM HEPES [N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid], 150mM NaCl, pH 7.4), coverslips were exposed to flowing blood in a parallel plate flow chamber as described.²³  Images were acquired at 2 separate preset positions, recorded with a Sony SVO-9500MD camera at 30 fps, and data were stored on a Terabyte server. The evolution of close cell-surface contact area in platelets captured from flow was monitored by reflection interference contrast microscopy (RICM).^24-26  Major advantages of this technique are that no exogenous reagents are required for cell detection, object visualization is limited to a maximum range of 30 to 100 nm from the cell-surface interface, and distance is depicted as a quantifiable gradient of gray intensities.²⁷  Images were recorded in real time at a minimum of 3 preset positions during 5 time cycles by use of a Sony DXC 390 CCD camera (Sony Corp).

To show that major differences exist in the dynamics of adhesion and spreading as a direct consequence of exposure to shear stress and to exclude differences in methodology as the basis for the defects in ADAP spreading under flow, we investigated platelet surface interactions by RICM in real time by using identical blood preparations and experimental setup under both static and flow conditions (supplemental Figure 1). To test the effect of soluble agonist stimulation on platelet spreading, a platelet monolayer was generated during the course of 1.5 minutes at 500 s⁻¹, and then platelets were perfused with buffer containing 0.25mM PAR4 receptor-activating peptide, 0.5mM MnCl₂, or vehicle. When inhibitors were used, they were added to blood 15 minutes before perfusion through the device.

After monitoring live platelet attachment and spreading kinetics, surface-interacting platelets were fixed with 3.7% formaldehyde^28,29  without cessation of flow (perfusion fixation) to maintain hydrodynamic fluid shear stress during fixation. Although this step was sufficient to immediately freeze motion and preserve morphological characteristics, incubation with formaldehyde was continued for another 5 minutes. Then coverslips were carefully removed from the flow chamber, rinsed, permeabilized, blocked with 10% goat serum, and stained with antibodies against proteins of interest and appropriate secondary antibodies and/or fluorescently labeled phalloidin to stain F-actin.²⁸  Western blots³⁰  were performed to confirm that all antibodies used detected a single protein band of expected molecular weight in platelet lysates. Images of postflow, fixed, and stained platelets were captured with a Deltavision deconvolution microscope (Applied Precision Inc) and Nikon 100×/1.4 NA oil objective with the use of a Photometrics Sony Coolsnap HQ camera and Softworks acquisition software (Applied Precision Inc) at the University of California, San Diego Cancer Center Core Microscopy Facility. Identical acquisition settings were used for all slides derived under equivalent experimental conditions. Deconvolved images were saved as TIF files and analyzed for protein phosphorylation and localization.

Further details of image analyses are described in the supplemental Methods.

To investigate adhesion under static conditions, we tested the ability of ADAP^+/+ and ADAP^−/− platelets to adhere at a range of input fibrinogen coating concentrations. Adhesion was quantified with an alkaline phosphatase assay.³¹  Nonspecific binding was determined by subtracting residual fluorescence in bovine serum albumin-coated wells. To examine platelet spreading under static conditions, washed ADAP^+/+ and ADAP^−/− platelets resuspended at 2 × 10⁷/mL in Walsh buffer were added to 18-mm coverslips coated with 100 μg/mL fibrinogen and incubated for 1 hour at 37°C in the presence or absence of PAR4 receptor-activation peptide or MnCl₂. Alternatively, blood suspensions prepared for flow studies were applied to fibrinogen-coated coverslips in the flow chamber without perfusion, and spreading under static conditions was monitored by RICM. Adherent platelets were stained with rhodamine-phalloidin for F-actin, and images were acquired as described previously.

Analysis of variance was performed by the use of the Student t test for unpaired samples.

Hemostasis appears compromised in ADAP^−/− mice, as evidenced by rebleeding upon tail-vein transection.¹⁶  Therefore, we assessed the capacity of ADAP^−/− mice to form occlusive thrombi after carotid artery injury by FeCl₃, a thrombosis model dependent on platelets as well as fibrin deposition. The average blood flow before FeCl₃ injury was similar in ADAP^+/+ and ADAP^−/− mice (1.1 ± .05 mL/min). At 5% to 7% FeCl₃ concentration, thrombus formation was comparable for ADAP^−/− and ADAP^+/+ littermates. To test whether less severe vessel injury would uncover a role for ADAP in thrombus formation, we decreased the FeCl₃ concentration to 3.75%, which reproducibly induced vessel occlusion in wild-type mice. In ADAP^−/− mice, however, vessel occlusion was significantly decreased at this concentration.

As illustrated in Figure 1A, carotid artery blood flow tracings from all ADAP^+/+ mice demonstrated reduced blood flow and complete occlusion within an average of 10 minutes after application of 3.75% FeCl₃. In contrast, ADAP^−/− mice typically exhibited cyclic flow changes indicative of dispersion and/or embolization of growing thrombi over time, a pattern previously observed with defects in outside-in αIIbβ3 signaling.^32-34  The average time to first occlusion was prolonged in ADAP^−/− mice (P < .01; Figure 1B), and 30 minutes after the lesion, 78% of ADAP^−/− but 0% of ADAP^+/+ mice exhibited patent arteries (n = 8; P < .01). Histologic examination of carotid arteries excised 30 minutes after FeCl₃ injury confirmed the presence of large, occlusive thrombi in ADAP^+/+ arteries as opposed to smaller, nonocclusive thrombi in ADAP^−/− arteries (Figure 1C left and middle). Arteries of ADAP^+/+ mice treated with isotonic saline instead of FeCl₃ were free of thrombus (Figure 1C right). Thus, under these experimental conditions, the absence of ADAP reduces the capacity of mice to develop a stable, occlusive carotid artery thrombus after a less severe vessel injury.

Given the particular hemostatic phenotype observed in ADAP^−/− mice, we examined the function of ADAP^−/− platelets in whole blood under conditions of shear stress, focusing on the known αIIbβ3 contribution to stable platelet attachment and spreading under these conditions.^23,35  Indeed, stable attachment of ADAP^−/− platelets to fibrinogen-coated coverslips was compromised under shear flow. Under the conditions used, an average of 900 and 800 ADAP^+/+ and ADAP^−/− platelets, respectively, contacted the surface in equivalent image fields examined over a 70-second period. ADAP^−/− platelets consistently showed a decrease in the percent of total surface contacting platelets able to sustain attachment to fibrinogen for between 2 and 30 seconds, relative to ADAP^+/+ platelets. Approximately 30% fewer platelets remained attached for 2 to 4 seconds, and longer-term adhesion was increasingly ADAP dependent, because approximately 40% to 50% fewer platelets remained attached for 6 seconds or up to 30 seconds (n = 4; P < .05, P < .01; paired Student t test; Figure 2A). As controls, no stable platelet adhesion was observed during an equivalent time period when ADAP^+/+ blood was perfused over coverslips coated with bovine serum albumin or over fibrinogen in the presence of EDTA (ethylenediaminetetraacetic acid), which blocks αIIbβ3 interaction with fibrinogen. In sharp contrast to these results, ADAP^−/− platelets adhered and spread onto fibrinogen normally under static conditions, whether platelets were unstimulated, activated by a PAR4 agonist to induce inside-out signaling, or treated with MnCl₂ to directly activate αIIbβ3 (Figure 2B-C; supplemental Figure 1C). Thus, ADAP is required for stable, αIIbβ3-dependent adhesion of platelets to fibrinogen under conditions of shear flow.

To evaluate the nature of the adhesion defect in more detail, platelet spreading was examined at shear rates (γ) between 50 s⁻¹ and 600 s⁻¹, corresponding to the range of venous and low arterial shear rates. Platelets were perfusion fixed and stained with an antibody to GP IX to identify the plasma membrane. At all γ, even at relatively early time points (30-45 seconds), it was possible to distinguish between ADAP^−/− and ADAP^+/+ platelets; more detailed analyses were conducted at γ = 500 s⁻¹. As shown in Figure 3A, platelets from the 2 mouse strains differed in morphology and surface area: ADAP^+/+ platelets lost the symmetrical, spherical appearance of unactivated cells; formed lamellipodia; and developed cusps from which filopodia extended (arrowhead, Figure 3A top). In contrast, ADAP^−/− adherent platelets largely retained a spherical morphology with extended filopodia, but few lamellipodia and exhibited a significant decrease in mean surface area when perfusion-fixed (6.66 ± 1.3 μm²), as compared with ADAP^+/+ platelets (8.93 ± 0.4 μm²; n = 5; P < .05; Figure 3B).

To examine the contact areas of unfixed platelets with fibrinogen in real time under shear flow and to avoid possible fixation artifacts, we used RICM. This technique reveals objects that are at a distance of 100 nm or less from a surface and with a progression from black to white on a gray scale as the distance increases. Early ADAP^+/+ platelet attachment points (dark areas) were small (area < 1 μm²) and central, soon followed by close adhesion contacts at filopodial tips. Central dark regions, corresponding to close contact areas, grew rapidly in ADAP^+/+ platelets as bonding surfaces extended during spreading (Figure 3C top) but remained smaller in ADAP^−/− platelets (Figure 3C bottom; supplemental Video 1) at all time points tested. Quantification of the average close contact area/platelet over time (an example of a thresholded and binarized image used for this measurement is shown in Figure 3D), confirmed the smaller size (P < .05) in ADAP^−/− platelets as compared with ADAP^+/+ platelets (Figure 3E). The surface contact area of ADAP^−/− platelets on fibrinogen was similar to that seen on a dmA1A2 VWF domain fragment (Figure 3E) that lacks binding sites for αIIbβ3 and supports only transient interactions. Continued monitoring by RICM indicated that establishment of close contact formation in ADAP^−/− platelets remained reduced even after 3 minutes from flow initiation. Moreover, the defective ability of ADAP^−/− platelets to form close contacts with and spread on fibrinogen could not be rescued by stimulation of platelets with a PAR4 agonist or with MnCl₂ (Figure 3F-G). Taken together, these results indicate that ADAP is required for the normal growth of platelet contact areas and spreading on fibrinogen under shear flow conditions. Furthermore, because MnCl₂ bypasses any need for inside-out signaling to activate αIIbβ3, the results are consistent with a defect in outside-in αIIbβ3 signaling in ADAP^−/− platelets.

Because of the structural role of the actin cytoskeleton and its potential contribution to platelet anchorage under flow conditions, we sought first to establish the normal F-actin distribution in ADAP^+/+ platelets captured onto fibrinogen within 45 seconds, 1.5 minutes, or 3 minutes of shear flow. From the earliest time point analyzed, we observed the formation of small actin-rich structures (Figure 4A) that increased in number and size over time. Concurrently, platelets increased their total surface areas, and the actin structures persisted and accumulated during the course of 3 minutes, with surprisingly few platelets producing the actin stress cables characteristic of platelets spread under static conditions.³⁶  We hypothesized that if ADAP were to play a role in actin polymerization and spreading, then it would potentially be found associated with actin or with actin-regulatory molecules. Indeed, we found that ADAP colocalized with actin in these structures in ADAP^+/+ platelets (Figure 4A top; Figure 4B). However, in ADAP^−/− platelets captured from flow, actin-rich structures were greatly reduced and usually asymmetrically distributed (Figure 4A bottom), even after 3 minutes. Vinculin, a hallmark of focal complexes and adhesions, localized to the actin-rich structures in ADAP^+/+ platelets but was not clustered in ADAP^−/− platelets (not shown). To quantify the frequency and size of these structures in platelet images, we calculated an arbitrary “cluster index” equal to (average cluster area) × (average cluster number/platelet). Single-cell analysis of the actin cluster index for a minimum of 150 ADAP^+/+ and ADAP^−/− platelets confirmed the decreased assembly of actin-rich structures in ADAP^−/− platelets adherent to fibrinogen (Figure 4C), which had a cluster index similar to that for ADAP^+/+ platelets unstably interacting with a dmA1A2 VWF fragment. The vinculin cluster index was also significantly lower in ADAP^−/− platelets (0.32 ± 0.02) than in ADAP^+/+ platelets (0.84 ± 0.07; n = 3; P < .01). In contrast, under static conditions, actin-rich structure formation was normal in ADAP^−/− platelets adherent to fibrinogen (not shown). Thus, under shear flow conditions, ADAP is required for αIIbβ3-dependent actin organization into structures that also contain vinculin.

Platelet adhesion to fibrinogen via αIIbβ3 initiates a signaling cascade (Figure 5A) that includes activation of Src and Syk protein tyrosine kinases and ADAP phosphorylation. Mice deficient in the ADAP binding partner, Fyn, have normal adhesion under flow and spreading on full-length VWF.³⁷  However, several ADAP binding partners have been implicated in the regulation of actin downstream of integrins, both in primary cells and model cell systems. For example, SLP-76 can promote lamellipodia formation when overexpressed in Chinese hamster ovary cells,¹⁷  polarization of T cells,³⁸  and platelet spreading on fibrinogen^39,40 ; similarly, VASP can promote actin polymerization⁴¹  and filopodia formation⁴² ; and SKAP-HOM can function as a phosphoinositol-gated switch for sustaining membrane ruffles.⁴³  However, the functional hierarchy of these molecules relative to ADAP has not been established in platelets. Accordingly, we investigated how ADAP may interface with its binding partners in fibrinogen-adherent platelets under flow.

We found that platelet adhesion to fibrinogen under flow induced the phosphorylation of Src at Tyr-418, an event concomitant with Src activation, and this was observed even in the absence of ADAP. Staining of perfusion-fixed ADAP^+/+ and ADAP^−/− platelets captured onto fibrinogen showed similar overall localization of Src pTyr-418 to filopodial protrusions (Figure 5B), but there was an additional localization of activated Src to actin-rich clusters in ADAP^+/+ platelets. Image quantification revealed no differences between ADAP^+/+ and ADAP^−/− platelets in Src pTyr-418 signal (Figure 5C). In contrast, staining for SLP-76 revealed major abnormalities in ADAP^−/− platelets. In ADAP^+/+ platelets, SLP-76 was dispersed throughout the cytoplasm and occasionally found as punctae after 45 to 90 seconds of flow, but by 3 minutes SLP-76 colocalized to actin-rich structures (Figure 6A left). However in ADAP^−/− platelets, SLP-76 remained primarily dispersed throughout the cytoplasm with occasional punctae (Figure 6A right). Vav1 is a SLP-76 binding partner that functions downstream of αIIbβ3 as an exchange factor for the actin regulator known to promote lamellipodia formation, Rac1.^17,34  Therefore, we assessed the localization of Vav1 with an antibody to Vav1 pTyr-174, a phospho-site associated with increased Vav1 exchange activity.⁴⁴  ADAP^+/+ platelets captured from flow exhibited strong phosphorylation of Vav1 Tyr-174 that was localized in large part to the actin-rich structures (Figure 6B, top). In contrast, almost no phosphorylation of Vav1 Tyr-174 was detectable in ADAP^−/− platelets (Figure 6B, bottom), and quantification of stained images indicated a greater than 60% reduction in this phosphorylation signal (n = 3, P < .05; Figure 6C). Thus, ADAP is required to correctly localize a pool of SLP-76 and phospho-Vav1 in adherent platelets under flow, providing one potential explanation for the cytoskeletal and spreading defects of ADAP^−/− platelets.

Because lamellipodial growth has been correlated with the presence of VASP at the membrane,⁴⁵  and ADAP may regulate VASP by targeting it to membrane regions, we co-stained perfusion-fixed platelets with antibodies against VASP and GP IX, the latter as a membrane marker. When 3-dimensional images were reconstructed from stacks of 0.1-μm platelet sections, VASP was found to be targeted to the membrane and to actin-rich structures in ADAP^+/+ platelets (Figure 7A). VASP targeting was significantly decreased in ADAP^−/− platelets (% inhibition membrane targeting = 53 ± 13%, n = 3; P < .05; Figure 7A-B). However, when VASP^−/− platelets were perfused over fibrinogen and compared with VASP^+/+ platelets, they showed no defects in spreading, close surface contact with the matrix, assembly of actin-rich structures, or colocalization of ADAP to these structures (Figure 7C-D). Similarly, platelets from SKAP-HOM–deficient mice functioned normally under flow (supplemental Figure 2). These results suggest that if any of the functions of VASP or SKAP-HOM are disrupted in platelets lacking ADAP, this cannot account for the phenotype of ADAP^−/− platelets under flow.

Studies primarily in T lymphocytes and model cell systems have implicated ADAP in outside-in signaling because ligand binding to integrins induces ADAP phosphorylation and association with actin regulatory proteins concomitant with cell motility.^9,38  However, no direct functional role for ADAP in outside-in signaling in platelets has been established. We have performed quantitative studies by using mouse models, intracellular protein staining, and live videomicroscopy of platelets in whole blood to uncover an essential role for ADAP in outside-in αIIbβ3 signaling under shear stress conditions. Our studies lead to the following conclusions: (1) ADAP is important for platelet function in vivo because ADAP-deficient mice exhibit increased frequency of rebleeding from tail bleeding wounds¹⁶  and form unstable thrombi in the carotid artery when challenged with FeCl₃ (Figure 1). (2) Hydrodynamic shear stresses act as physical cues to induce αIIbβ3-dependent remodeling of the cytoskeleton in wild-type platelets (mechanotransduction) to produce platelet morphologies that minimize flow impact and maximize platelet-surface contact areas. (3) ADAP is an essential component of platelet mechanotransduction mechanisms under shear flow because ADAP^−/− platelets exhibit poor stable attachment to fibrinogen, reduced surface contact areas in spreading platelets, and loss of actin-rich structures under flow. (4) ADAP transduces outside-in αIIbβ3 signals to organize the localization of proteins, including vinculin and VASP, and under shear stress by enabling signal relay from Src family kinases to the platelet actin cytoskeleton to appropriately localize SLP-76 and to promote Vav1 tyrosine phosphorylation and lamellipodia formation.

Thus, the required role for ADAP in flow-dependent platelet adhesion and spreading onto fibrinogen, and possibly other αIIbβ3 ligands, suggests that ADAP enhances the stability of thrombi formed in the arterial circulation. The selective abnormality in thrombus formation in ADAP^−/− mice at lower ferric chloride concentrations may furthermore reflect differences in the relative role of ADAP in tissue factor/fibrin versus collagen driven pathways.^46,47  The presence of abundant actin-rich structures under shear stress conditions in ADAP^+/+ platelets correlated with stable adhesion and thrombus formation. Similar structures were recently reported by Calaminus et al, and were hypothesized to be an early stage in platelet spreading.⁴⁸  Interestingly, under shear stress these structures persist even when platelets are well spread but largely lacking actin stress fibers, and they colocalize with active Src pTyr-418 (Figure 5). Thus, these actin-rich structures may help to promote stable anchorage of platelets to the substratum under shear stress.

Signaling pathways triggered by shear stress provide cues for differentiation of stem cells,⁴⁹  and they induce changes in cell morphology, stiffness, and gene expression in endothelial cells,^3,50  where stress-induced signals are superposed on ongoing signaling sustained through cell-matrix adhesions. In platelets, signaling to, and force transmission by, the cytoskeleton under the stringent time and force constraints of shear flow is expected to be an important mechanism for stable adhesion and spreading. In theory, shear stresses may trigger αIIbβ3 signaling pathways by direct structural modification of αIIbβ3, by membrane fluidity changes affecting αIIbβ3 clustering, or by altering associations of regulatory proteins with the cytoplasmic tails of αIIb and β3.⁸  How might ADAP function as a mechanotransducer of αIIbβ3 signals under these conditions? Because no direct actin regulation by ADAP is known, mechanical forces transmitted to ADAP may modulate the actin-regulatory function of one or more of its binding partners, including SLP-76, Fyn, VASP, and SKAP-HOM. For example, ADAP may itself undergo shear-dependent phosphorylation and increase in binding affinity for its partners. Alternatively, shear forces on ADAP might enhance intracellular transport of associated binding partners along microtubule networks by its association with dynein, a microtubule motor protein,^51,52  or enhance membrane retention of the ADAP signaling module via ADAP's phospholipid binding domains.¹⁵ 

Of the several binding partners for ADAP, only SLP-76 has been shown to have an unambiguous role in outside-in αIIbβ3 signaling. SLP-76 expression drives lamellipodia formation,¹⁷  and SLP-76–deficient platelets have a spreading defect, even under static conditions.³⁹  Mice have been generated with phenylalanine substitutions of 1 or more of 3 SLP-76 tyrosine residues that when phosphorylated recruit actin-regulatory proteins such as Vav1 and the Itk/Btk/Tec protein tyrosine kinases.^40,53,54  In particular, the loss of 2 phosphotyrosines in SLP-76 that anchor Vav1 (pTyr-112; pTyr-128) resulted in reduced platelet spreading on fibrinogen under static conditions, indicative of a key role for the SLP-76/Vav1 module in outside-in αIIbβ3 signaling.⁴⁰  Indeed, major findings in the present study were a loss of SLP-76 and Vav1 localization to actin structures and a failure of Vav1 to become phosphorylated on Tyr-174 upon adhesion of ADAP^−/− platelets to fibrinogen under fluid shear stress (Figure 5).

These results further highlight key roles for SLP-76 and Vav1, and they establish ADAP as a critical member of this signaling module for cytoskeletal regulation and the stability of platelet adhesion to fibrinogen under shear. The involvement of Vav1 in these events presumably reflects its Rac1 exchange function and the induction of lamellipodia by Rac1.³⁴  Although Vav1 additionally has GEF activity toward Cdc42, filopodia formation appears to be unaffected in ADAP^−/− platelets under flow. This finding suggests that in ADAP^−/− platelets under flow, alternate pathways may be operating that lead to activation of Cdc42 or formins, both of which have been shown to regulate filopodia formation.⁵⁵  In addition to promoting lamellipodia formation,⁵⁶  phospho-Vav1 has been shown to regulate the stability of clusters containing Vav1 and other proteins in T cells⁵⁷  and to regulate decoupling of cortical actin from the membrane leading to increased cell deformation with applied force.⁵⁸  Because the spherical morphology of fibrinogen-adherent ADAP^−/− platelets appears to represent an early stage of platelet spreading under flow,⁵⁹  we speculate that mislocalization of SLP-76 and loss of Vav1 functions may contribute, at least in part, to the ADAP^−/− platelet phenotype we observed. Interestingly, we found no gross morphologic abnormalities in platelets under flow in platelets deficient in either one of ADAP's 2 other binding partners, SKAP-HOM or VASP.

In conclusion, this study provides a new understanding of the responses of platelets to mechanotransduction initiated through αIIbβ3 under hydrodynamic stress conditions by establishing an essential role for ADAP in this process. Further studies will be required to determine precisely how mechanical forces engendered by hemodynamic shear stresses affect ADAP's function downstream of αIIbβ3 to transmit adhesive signals to the platelet cytoskeleton.

We are grateful to Alexander Clowes for providing VASP-deficient mice, Gary Koretzky for providing ADAP-deficient mice, and Ben Neel for the SKAP-HOM–deficient mice. We thank Meri Ripani for technical assistance with image analysis and Per Fogelstrand for providing expertise on surgical procedures in the mouse.

This work was supported by grants HL78784, HL31950, and HL56595 from the National Institutes of Health

National Institutes of Health

Contribution: A.K.-F. performed the experiments and analyzed the data; Z.M.R. supplied vital reagents; and A.K.-F., Z.M.R., and S.J.S. designed the research and wrote the paper.

Conflict-of-interest disclosure: The authors declare no competing financial interests.

Correspondence: Dr Ana Kasirer-Friede, Department of Medicine, University of California, San Diego, 9500 Gilman Dr, Mail Code 0726, La Jolla, CA 92093-0726; e-mail: akasirer@ucsd.edu/sshattil@ucsd.edu.