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Keywords:

  • degranulation;
  • phospholipase D;
  • platelet;
  • signaling;
  • thrombosis

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. Disclosure of Conflict of Interests
  9. References
  10. Supporting Information

Summary.  Background: Platelet activation and aggregation are crucial for primary hemostasis, but can also result in occlusive thrombus formation. Agonist-induced platelet activation involves different signaling pathways leading to the activation of phospholipases, which produce second messengers. The role of phospholipase C (PLC) in platelet activation is well established, but less is known about the relevance of phospholipase D (PLD) .

Objective and methods:  The aim of this study was to determine a potential function of PLD2 in platelet physiology. Thus, we investigated the function of PLD2 in platelet signaling and thrombus formation, by generating mice lacking PLD2 or both PLD1 and PLD2. Adhesion, activation and aggregation of PLD-deficient platelets were analyzed in vitro and in vivo.

Results:  Whereas the absence of PLD2 resulted in reduced PLD activity in platelets, it had no detectable effect on the function of the cells in vitro and in vivo. However, the combined deficiency of both PLD isoforms resulted in defective α-granule release and protection in a model of FeCl3-induced arteriolar thrombosis, effects that were not observed in mice lacking only one PLD isoform.

Conclusion:  These results reveal redundant roles of PLD1 and PLD2 in platelet α-granule secretion, and indicate that this may be relevant for pathologic thrombus formation.


Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. Disclosure of Conflict of Interests
  9. References
  10. Supporting Information

Blood platelets ‘monitor’ the integrity of the vascular system, and, in response to altered vascular surfaces, e.g. at sites of traumatic injury or pathologic alteration of the endothelial layer, they rapidly become activated. This activation causes a morphologic change of the cells from a discoid to a spherical shape, their firm adhesion and spreading on the reactive surface, the exocytosis of α-granules and dense granules, and platelet–platelet interactions. This multistep process, which involves multiple membrane receptors and intracellular signaling pathways [1], is crucial to form a platelet plug that seals the wound site and thereby limits blood loss after tissue trauma. However, in diseased arteries, these events may lead to thrombotic vessel occlusion and ischemic tissue damage [2], such as in myocardial infarction and ischemic stroke [3,4].

At sites of injury, exposure of subendothelial collagens triggers platelet activation via tyrosine phosphorylation cascades downstream of glycoprotein (GP) VI, with its associated immunoreceptor tyrosine activation motif (ITAM) [5]. Activated platelets release the soluble agonists thromboxane A2 (TxA2) and ADP, which – together with locally generated thrombin – stimulate receptors that couple to heterotrimeric G-protein-coupled receptors and induce distinct downstream signaling pathways [6]. Stimulation of Gq or ITAM-coupled receptors induces the activation of phospholipase C (PLC) family proteins, which trigger calcium mobilization and the activation of protein kinase C (PKC) via diacylglycerol (DAG). All of these signaling events converge in the ‘final common pathway’ of platelet activation, the functional upregulation of integrin adhesion receptors, most notably αIIbβ3 integrin. This so-called ‘inside-out’ activation enables integrins to efficiently bind their ligands [7], thus promoting firm adhesion and platelet aggregation by binding to ligands such as von Willebrand factor (VWF), fibronectin, or fibrinogen. Furthermore, platelets secrete their granule contents upon activation, and thereby reinforce the local thrombotic activity. Dense granules contain predominantly small molecules, such as ADP and serotonin, whereas α-granules contain a vast array of proteins, ranging from adhesion molecules and coagulation factors to chemokines and cytokines [8].

Phospholipase D (PLD) isoforms also become activated downstream of major platelet signaling pathways. PLD is a widely distributed enzyme that hydrolyzes phosphatidylcholine to choline and phosphatidic acid (PA) [9]. PA, as well as its metabolites DAG and lysoPA, are important intracellular messengers that have critical roles in several cellular functions [9]. In the presence of a primary alcohol, the alcohol, and not water, is the preferred substrate, leading to the generation of phosphatidyl alcohol instead of PA. This transphosphatidylation is unique to PLD, and is commonly used in assays to measure the activity of the enzyme [10]. Two mammalian PLD isoforms exist, and are ubiquitously expressed: PLD1 [11] and PLD2, which share ∼ 50% sequence homology [12]. PLD1 is believed to be activated by PKC and GTPases of the ADP ribosylation factor and Rho families [13–15]. The regulation of PLD2 is still a matter of debate. Because of the high basal activity of PLD2 in vitro [16], it has been speculated that its activity is not inducible, but others have shown that it increases upon stimulation [17,18]. In platelets, both PLD isoforms are present, and PLD activity has been reported to be upregulated upon platelet activation by collagen, thrombin, and the TxA2 analog U46619 [19,20]. It has been suggested that PLD is required for the secretion of dense granules [21] and lysosomes [20]. However, until recently [22–24], no PLD-deficient mice were reported, and potential downstream targets of PLD were identified mostly by correlation studies (linking PLD activity to simultaneously occurring cellular events) or by inhibiting PA generation with 1-butanol to divert PLD activity. Yet, primary alcohols only partially prevent PA production, even at the maximal applicable concentrations, and have off-target effects on cell behavior that may confound interpretation of the obtained results [25]. We recently reported the analysis of Pld1−/− platelets, and revealed that PLD1 is required for proper integrin activation in response to weak agonist signals. Consequently, mice lacking PLD1 were protected against arterial thrombosis and ischemic stroke [23]. A very recent study, which used a PLD inhibitor, suggested a role of PLD2 as negative regulator of platelet secretion [26]. However, by using Pld2−/− and Pld1−/−/Pld2−/− mice, we show that the isolated loss of PLD2 has no effect on platelet activation, whereas the combined deficiency of both PLD isoforms results in a defect in α-granule release and protection from ferric chloride-induced thrombosis.

Materials and methods

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. Disclosure of Conflict of Interests
  9. References
  10. Supporting Information

Animals

Animal studies were approved by the district government of Lower Franconia (Bezirksregierung Unterfranken). Pld2−/− mice were generated as follows. A VGB6 embryonic stem (ES) cell clone (BD1) in which the Pld2 gene was deleted and replaced with a lacZ-flanked and loxP-flanked neocassette was obtained from KOMP Repository (http://www.komp.org). Male chimeric mice originating from the injection of ES cells into albino C57Bl/6 blastocyts were bred to C57Bl/6 females to generate Pld2−/− mice, which were intercrossed to produce Pld2−/− mice. Mice were genotyped by PCR, with 5′-AAGCAACACCACACATTCCA-3′ and 5′-CTTCCCGACTCACAGCTTTC-3′ for the wild type, and 5′-TCATTCTCAGTATTGTTTTGCC-3′ and 5′-GGAGGAAGAGTGAGATGAAG-3′ for the Pld2-null allele. Pld2−/− mice were intercrossed with Pld1−/− mice [23], which had meanwhile been backcrossed with a C57Bl/6 background, to generate Pld1−/−/Pld2−/− mice.

Chemicals and antibodies

The anesthetic drugs used included medetomidine (Pfizer, Karlsruhe, Germany), midazolam (Roche, Grenzach-Wyhlen, Germany), and fentanyl (Janssen-Cilag, Neuss, Germany). To reverse anesthesia, atipamezol (Pfizer) and flumazenil and naloxon (both from Delta Select, Dreieich, Germany) were used according to local authority regulations. ADP was from Sigma-Aldrich (Schnelldorf, Germany), protease-activated receptor 4 (PAR4)-activating peptide was from Thermo Fisher Scientific (Dreieich, Germany), U46619 was from Alexis Biochemicals (San Diego, CA, USA), thrombin was from Roche, collagen (Kollagenreagent Horm) was from Nycomed (Munich, Germany), apyrase type III was from GE Healthcare (Chalfont St. Giles, UK), human fibrinogen, high molecular weight heparin and rabbit anti-PLD2 (P5993) were from Sigma-Aldrich, rabbit anti-tubulin (MAB1864) was from Chemicon (Hofheim, Germany), and anti-rabbit IgG–horseradish peroxidase (HRP) was from Dako Cytomation (Hamburg, Germany). Indomethacin was purchased from a local pharmacy. All monoclonal anti-platelet GP antibodies conjugated with fluorescein isothiocyanate, phycoerythrin or DyLight 488 were obtained from Emfret Analytics (Eibelstadt, Germany). Collagen-related peptide (CRP) was generated as previously described [27].

RT-PCR analysis

Murine platelet mRNA was isolated with Trizol reagent and detected by RT-PCR, according to the manufacturer’s protocol (Invitrogen, Karlsruhe, Germany). The following primers were used to detect the Pld2 transcript: 5′-GTGCCACTGTGCAGGTCTTGAGG-3′ and 5′-GCAGAATAGCCTGGATGGAG-3′.

Immunoblotting

Proteins of lysed platelets were separated by SDS-PAGE and blotted onto poly(vinylidene difluoride) membranes. After blocking, the membrane was incubated with antibody overnight at 4 °C. HRP-conjugated secondary antibodies were incubated for 1 h at room temperature, and enhanced chemiluminescence was used for visualization.

Platelet preparation and PLD activity measurements

Washed platelets were prepared as previously described [28]. To determine PLD activity, the formation of [3H]phosphatidylethanol ([3H]Ptd-EtOH) was measured with standard protocols [10,20]. Washed platelets were adjusted to a final concentration of × 108 platelets mL−1, and were labeled with 3.7 kBp mL−1 [3H]palmitic acid (Perkin Elmer, Rodgau, Germany) at 37 °C for 1 h. Aliquots of 80 μL were preincubated with 0.5% ethanol for 10 min, and then stimulated with thrombin (0.1 U mL−1) or CRP (10 μg mL−1) in the presence of 2 mm CaCl2. Reactions were stopped by adding 500 μL of ice-cold chloroform/methanol and placing on ice. To extract the lipids, 500 μL of ice-cold chloroform and 350 μL of water were added; the lipids were then collected in the organic phase, and separated by TLC. [3H]Ptd-EtOH bands were identified through comigration with standards, and quantified by scintillation. PLD activity is depicted as percentage of phosphatidylethanol of total H3-labeled phospholipids.

Platelet aggregation/ATP release and flow cytometry

Washed platelets (160 μL with 1.56 × 108 platelets mL−1) were analyzed in the presence of 70 μg mL−1 human fibrinogen. Transmission was recorded on a four-channel aggregometer (Fibrintimer; APACT, Hamburg, Germany) for 10 min, and was expressed in arbitrary units, with buffer representing 100% transmission. ATP secretion was measured with CHRONO-LUME reagent, according to the manufacturer’s protocol, on a Chronolog aggregometer (Chrono-Log Corp., Havertown, PA, USA). Luciferase (25 μL) was added directly to the platelets (0.5 × 106 platelets μL−1) under constant stirring, and the indicated concentrations of various agonists were added to study ATP release. The luminescence intensity was measured at a setting of × 0.01. For flow cytometry, heparinized whole blood was diluted 1 : 20, incubated with the appropriate fluorophore-conjugated mAbs for 15 min at room temperature, and analyzed on a FACSCalibur (Becton Dickinson, Heidelberg, Germany).

Measurement of VWF and platelet factor 4 (PF4) release

Washed platelets were adjusted to a final concentration of 0.5 × 106 platelets μL−1, and activated in the presence of 1.4 μm indomethacin and 2 U mL−1 apyrase with the indicated agonists for 15 min at 37 °C. Platelets were immediately centrifuged. VWF in the supernatant was quantified by ELISA, with unconjugated and HRP-conjugated anti-human VWF (hVWF) antibodies (DakoCytomation). The amount of secreted PF4 was quantified with a PF4 ELISA (RayBiotech, Norcross, GA, USA).

Adhesion under flow conditions

Rectangular coverslips (24 × 60 mm) were coated with 0.2 mg mL−1 fibrillar type I collagen (Nycomed) or rabbit anti-hVWF antibody (1 : 500; AOO82; DakoCytomation) for 1 h at 37 °C, and blocked with 1% bovine serum albumin. For flow adhesion experiments on VWF, blocked coverslips were incubated with murine plasma for 2 h at 37 °C to allow binding of VWF to immobilized anti-VWF antibody. Heparinized whole blood was labeled with a DyLight 488-conjugated anti-GPIX Ig derivative at 0.2 μg mL−1, and perfusion was performed as previously described [29]. Image analysis was performed off-line with metavue software (Visitron, Munich, Germany). Thrombus formation was expressed as the mean percentage of total area covered by thrombi, and as the mean integrated fluorescence intensity per square millimeter. Platelet adhesion on the VWF matrix was analyzed by counting adhesive platelets per visual field.

Intravital microscopy of thrombus formation in FeCl3-injured mesenteric arterioles

Four-week-old mice were anesthetized, and the mesentery was exteriorized through a midline abdominal incision [30]. Arterioles (35–60 μm diameter) were visualized with a Zeiss Axiovert 200 inverted microscope (× 10/0.3 NA objective) (Carl Zeiss, Göttingen, Germany) equipped with a 100-W HBO fluorescent lamp source, and a CoolSNAP-EZ camera (Visitron). Digital images were recorded and analyzed off-line with metavue software. Injury was induced by topical application of a 3-mm2 filter paper saturated with FeCl3 (20%) for 10 s. Adhesion and aggregation of fluorescently labeled platelets (Dylight-488-conjugated anti-GPIX Ig derivative) in arterioles were monitored for 40 min or until complete occlusion occurred (blood flow stopped for > 1 min).

Aorta occlusion model

A longitudinal incision was used to open the abdominal cavity and expose the abdominal aorta of anesthetized mice. An ultrasonic flow probe was placed around the vessel, and thrombosis was induced by a single firm compression with forceps. Blood flow was monitored for 30 min.

Transient middle cerebral artery (MCA) occlusion (tMCAO) model

Experiments were conducted on 8–10-week-old mice according to previously published recommendations for research in mechanism-driven basic stroke studies [31]. tMCAO was induced under inhalation anesthesia with the intraluminal filament (Doccol Company, Sharon, VT, USA) technique [32]. After 60 min, the filament was withdrawn to allow reperfusion. For measurements of ischemic brain volume, mice were killed 24 h after induction of tMCAO, and brain sections were stained with 2% 2,3,5-triphenyltetrazolium chloride (Sigma-Aldrich). Brain infarct volumes were calculated and corrected for edema [32]. Neurologic function and motor function were assessed by two independent and blinded investigators 24 h after tMCAO, as previously described [32].

Bleeding time

Mice were anesthetized, and a 2-mm segment of the tail tip was removed with a scalpel. Tail bleeding was monitored by gently absorbing blood with filter paper at 20 s intervals without making contact with the wound site. When no blood was observed on the paper, bleeding was determined to have ceased. Experiments were stopped after 20 min.

Statistics

Results from at least three experiments per group are presented as means ± standard deviations. Differences between two groups were assessed with the Mann–Whitney U-test. For the stroke model, infarct volumes and functional data were tested for Gaussian distribution with the D’Agostino and Pearson omnibus normality test, and then analyzed with the two-tailed Student’s t-test. For statistical analysis, PrismGraph 4.0 software (GraphPad Software, La Jolla, CA, USA) was used. Differences between more than two groups were analyzed with one-way anova with Dunnett’s T3 as post hoc test, with spss 20. A P-value of < 0.05 was considered to be statistically significant.

Results

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. Disclosure of Conflict of Interests
  9. References
  10. Supporting Information

Generation of Pld2−/− mice

To directly test the function of PLD2 in platelet activation and thrombus formation, we disrupted the Pld2 gene in mice. In line with a previous report [24], mice heterozygous for Pld2-null mutation and Pld2−/− mice were born in the expected Mendelian ratio, developed normally, and were viable and fertile. The mice appeared to be healthy, and did not show any signs of spontaneous bleeding. RT-PCR (Fig. 1A) and western blot analysis (Fig. 1B) confirmed the absence of PLD2 in platelets and other tissues. Pld1 mRNA expression was unaltered in Pld2−/− platelets, indicating that Pld1 mRNA expression is not upregulated upon Pld2 gene deletion. Blood cell counts, mean platelet volume, platelet lifespan and the expression of prominent surface GP receptors were similar between wild-type and Pld2−/− mice (Table S1 and data not shown). Together, these results indicate that megakaryopoiesis and platelet formation occur independently of PLD2.

image

Figure 1.  Genetic ablation of Pld2 results in decreased phospholipase D (PLD) activity in platelets. (A) Analyses of the presence of Pld1 and Pld2 mRNA in platelets with the indicated genotypes by RT-PCR. Actin mRNA served as control. (B) Western blot of organ lysates of mice with the indicated genotypes. Expression of tubulin was used as loading control. (C) Platelets were labeled with [3H]palmitic acid and stimulated with the indicated agonists. PLD activity is shown as percentage of phosphatidylethanol (Ptd-EtOH) of total 3H-labeled phospholipids. Data are mean ± standard deviation of four mice per group. *P < 0.05, **P < 0.01, ***P < 0.001. CRP, collagen-related peptide; WT, wild-type.

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To assess the effect of PLD2 deficiency on platelet PLD activity, a widely accepted in vivo PLD assay was used that detects phosphatidylethanol, a non-degradable product of PLD. Only low basal PLD activity was detected in wild-type platelets (∼ 5% of maximal activity, Fig. 1C), but platelet activation with classic agonists, such as thrombin or CRP, resulted in a pronounced increase in PLD activity. Remarkably, PLD activity detected at early time points was reduced in Pld2−/− platelets (Fig. 1C), demonstrating that PLD activity in platelets is tightly regulated and that PLD2 significantly contributes to the total PLD activity in these cells. However, the high residual inducible PLD activity in Pld2−/− platelets, which was indistinguishable from that observed in wild-type platelets at late time points, confirmed the previous notion that PLD1 is the dominant isoform in these cells [23].

Absence of PLD2 has no effect on platelet activation in vitro

We investigated whether reduced PLD activity in Pld2−/− platelets influences agonist-induced platelet activation. Flow cytometric analysis of αIIbβ3 activation and degranulation, determined as the surface expression of P-selectin (Fig. 2A) and ATP release (not shown), yielded indistinguishable results for Pld2−/− and wild-type platelets in response to all tested agonists at all concentrations. Similarly, no differences in reactivity were noted between Pld2−/− and wild-type platelets in standard aggregometry with different agonists (Fig. 2B). Our recent study of Pld1−/− platelets [23] revealed a signaling role for PLD1 downstream of GPIb, which is difficult to assess under static conditions. Consequently, flow adhesion assays were performed in which whole blood was perfused over a collagen-coated or VWF-coated surface at different shear rates. Wild-type and Pld2−/− platelets formed stable aggregates on collagen to the same extent and with the same kinetics (Fig. 2C). Likewise, rolling and firm adhesion on VWF was indistinguishable between the two genotypes (Fig. 2D).

image

Figure 2.  Absence of phospholipase D2 has no effect on platelet function in vitro. (A) Flow cytometric analyses of αIIbβ3 integrin activation (JON/A–phycoerythrin [PE]) and degranulation-dependent P-selectin exposure in response to the indicated agonists. Results are mean fluorescence intensity (MFI) ± standard deviation (SD) of six mice per group. (B) Washed platelets were stimulated with the indicated agonists, and light transmission was recorded with a Born aggregometer. Representative aggregation curves of three individual experiments are shown. (C) Whole blood was perfused over a collagen-coated (0.2 mg mL−1) surface. Representative pictures are shown (top). Blood was perfused for 4 min, and washing with Tyrode’s buffer was performed for a period equal to the perfusion time. Platelet surface coverage (bottom, left) and relative thrombus volume as measured by the integrated fluorescent intensity (IFI) per square millimeter (bottom, right) ± SD of four mice per group were measured. Bar: 50 μm. (D) Whole blood was perfused over a von Willebrand factor-coated surface, and then washed with Tyrode’s buffer for a period equal to the perfusion time. Tethered platelets were counted after 100 s of blood perfusion (left). The number of firmly adherent platelets was counted at the end of the washing step (right). Bar graphs depict mean values ± SD of four or more mice per group. *< 0.05, **< 0.01, ***< 0.001. CRP, collagen-related peptide; CVX, convulxin; FITC, fluorescein isothiocyanate; Rhod, rhodocytin; U46, U46619; WT, wild-type.

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PLD2 has been proposed to play a role in cytoskeletal rearrangements [15,33]. To test this directly, F-actin levels were measured in thrombin-activated and resting platelets with a flow cytometric approach. No differences were detected between wild-type and mutant platelets (not shown). Furthermore, wild-type and Pld2−/− platelets were allowed to spread on a fibrinogen-coated surface in the presence of thrombin. Pld2−/− platelets spread to a similar extent and with the same kinetics as wild-type platelets (Fig. S1). Collectively, these data demonstrate that the absence of PLD2 has no effect on platelet activation in vitro.

Normal hemostasis and occlusive thrombus formation in Pld2−/− mice

Next, we analysed the effect of PLD2 deficiency on platelet function in hemostasis and thrombosis. Bleeding times after amputation of the tail tip of wild-type and Pld2−/− mice were similar (Fig. 3A). Similarly, the application of 20% FeCl3 on the exteriorized mesenteric arterioles of Pld2−/− mice resulted in fast platelet adhesion and thrombus growth leading to irreversible vessel occlusion after 14.2 ± 4.1 min, which was similar to the kinetics of thrombus formation observed in wild-type mice (13.8 ± 3.4 min; Fig. 3B,C). As Pld1−/− mice have been shown to be protected in another in vivo model, in which the abdominal aorta is mechanically injured and blood flow is monitored with an ultrasonic perivascular Doppler flowmeter [23], Pld2−/− mice were also subjected to this model. However, also in this system, no difference between wild-type and Pld2−/− mice was observed (Fig. 3D).

image

Figure 3.  Lack of phospholipase D2 has no influence on thrombus formation, ischemic stroke or hemostatic function. (A) Tail bleeding times of wild-type and Pld2−/− mice. Each symbol represents one individual. (B, C) Thrombus formation in small mesenteric arterioles was induced by topical application of 20% FeCl3. For monitoring of thrombus formation by intravital microscopy, platelets were labeled fluorescently. Time to stable occlusion (B) and representative pictures (C) are shown. Each symbol represents one individual. (D) Mechanical injury of the abdominal aorta was induced by tight compression with forceps, blood flow was monitored for 30 min, and time to stable occlusion was recorded. Each symbol represents one individual. (E) Brain infarct volumes of wild-type and Pld2−/− mice that were subjected to the transient middle cerebral artery occlusion model (60 min of occlusion, 24 h after reperfusion). Data are mean ± standard deviation of 10 mice per group.

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To test for a potential role of PLD2 in focal cerebral ischemia, the development of neuronal damage in Pld2−/− mice following tMCAO was studied. For this, a thread was advanced through the carotid artery into the MCA, reducing regional cerebral flow by > 90% to induce cerebral ischemia, and was removed after 1 h to allow reperfusion [32,34]. At 24 h after reperfusion, both infarct volumes (Fig. 3E) and neurologic scores (not shown) were indistinguishable between wild-type and Pld2−/− mice.

Together, these results demonstrate that thrombus formation, hemostasis and ischemic brain infarct development occur independently of PLD2.

Partially redundant function of PLD1 and PLD2 in platelet degranulation

PLD1 and PLD2 share a high degree of sequence homology, making it likely that one isoform is able to compensate, at least partially, for the loss of the other. In order to address this question, Pld2−/− mice were intercrossed with Pld1−/− mice [23] to obtain mice lacking both PLD isoforms. The combined deficiency of PLD1 and PLD2 completely abolished agonist-induced PLD activity in platelets (Fig. 4A). Nevertheless, Pld1−/−/Pld2−/− mice developed normally, were viable, were fertile, and appeared to be healthy. Blood cell counts, mean platelet volume and the expression of prominent surface GP receptors were similar between wild-type and Pld1−/−/Pld2−/− mice, apart from slightly elevated white blood cell counts and increased β1 integrin levels on the platelet surface (Table S2). Together, these results indicate that PLD activity is dispensable for megakaryopoiesis and platelet production.

image

Figure 4.  Combined deficiency of phospholipase D1 and phospholipase D2 results in abolished phospholipase D (PLD) activity and decreased integrin activation and degranulation downstream of protease-activated receptors (PARs). (A) Platelets were labeled with [3H]palmitic acid and stimulated with the indicated agonists. PLD activity is shown as percentage of phosphatidylethanol (Ptd-EtOH) of total 3H-labeled phospholipids. Data are mean ± standard deviation (SD) of four mice per group. (B, D) Flow cytometric analyses of degranulation-dependent P-selectin exposure (B) and αIIbβ3 integrin activation (JON/A–phycoerythrin [PE]) (D) in response to the indicated agonists. Results are mean fluorescence intensity (MFI) ± SD of six mice per group. (C) Measurement of secreted von Willebrand factor (VWF) in the supernatant of resting or activated wild-type and Pld1−/−/Pld2−/− platelets. Data are presented as OD450 nm ± SD of four mice per group. (E) Measurement of secreted platelet factor 4 (PF4) in the supernatant of resting or activated wild-type and Pld1−/−/Pld2−/− platelets. Data are presented as OD450 nm ± SD of four mice per group. (F) ATP secretion after platelet stimulation with the indicated thrombin concentration was measured with a Chronolog aggregometer. Results are mean of ATP concentration (nm) ± SD of four mice per group. (G) Washed platelets were stimulated with the indicated agonists, and light transmission was recorded with a Born aggregometer. Representative aggregation curves of three individual experiments are shown. *< 0.05, **< 0.01, ***< 0.001. CRP, collagen-related peptide; CVX, convulxin; FITC, fluorescein isothiocyanate; Rhod, rhodocytin; U46, U46619; WT, wild-type.

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Next, we tested whether the absence of PLD activity influences agonist-induced platelet activation. As the lack of PLD1 alone results in mild defects in platelet integrin activation [23], flow cytometric analyses were performed with wild-type, Pld1−/− and Pld1−/−/Pld2−/− platelets, to enable direct comparison of single-deficient and double-deficient cells. Degranulation was indistinguishable between wild-type and Pld1−/− platelets, but Pld1−/−/Pld2−/− platelets showed reduced α-granule release upon stimulation of PAR4 with either thrombin or PAR4-activating peptide, as determined by the levels of surface-exposed P-selectin (Fig. 4B). Furthermore, the secretion of the α-granular proteins VWF and PF4 was decreased in Pld1−/−/Pld2−/− platelets as compared with wild-type platelets (Fig. 4C,E), whereas platelets lacking only one PLD isoform did not show a defect in secretion of VWF (Fig. S2). In order to exclude the possibility that this defect results from defective α-granule biogenesis in Pld1−/−/Pld2−/− platelets, we performed transmission electron microscopy. Microscopic analyses showed that the amount and localization of α-granules in Pld1−/−/Pld2−/− platelets were similar to those in wild-type platelets (Fig. S3). To test whether the secretion defect was specific for α-granules, we assessed ATP release as a measure of dense granule secretion. Remarkably, agonist-induced ATP release was indistinguishable between wild-type and Pld1−/−/Pld2−/− platelets (Fig. 4F). No ATP secretion was detectable in platelets of either genotype at lower thrombin concentrations (not shown). This indicated that PLD1 and PLD2 have redundant functions in the secretion of α-granules upon weak stimulation of platelet PAR4. In Pld1−/− platelets, integrin activation was slightly reduced upon stimulation with low and intermediate concentrations of thrombin or PAR4-activating peptide, confirming previous results [23]. This defect was even more pronounced in platelets lacking both PLD isoforms (Fig. 4D).

Despite these defects in integrin activation and degranulation, no differences in reactivity were noted between Pld1−/−/Pld2−/− and wild-type platelets in aggregometry (Fig. 4G). This supports previous reports stating that robust aggregation can occur in this assay under conditions of suboptimal integrin activation [35,36].

Absence of both PLD isoforms protects mice from FeCl3-induced arteriolar thrombosis

In vivo, platelet activation and thrombus formation occur in flowing blood, where locally produced soluble mediators are rapidly diluted. Under these conditions, subtle defects in integrin activation may translate into reduced thrombus formation. Thus, we investigated the effect of PLD double deficiency in a model of arteriolar thrombosis. We have previously shown that Pld1−/− mice are protected in arterial thrombosis models induced by chemical injury of the carotid artery or mechanical injury of the abdominal aorta [23]. Therefore, we chose the FeCl3-induced mesenteric arteriole injury model to investigate a potential redundant function of the two PLD isoforms. The application of 20% FeCl3 on the exteriorized mesenteric arterioles of Pld1−/− mice resulted in fast platelet adhesion and thrombus growth leading to irreversible vessel occlusion after 19.4 ± 4.1 min, which was similar to the kinetics of thrombus formation observed in wild-type mice (20.8 ± 3.6 min; Fig. 5A,B). In contrast, although initial platelet adhesion in Pld1−/−/Pld2−/− mice was similar to that in wild-type or Pld1−/− mice, the final outcome was more variable. In 12 of 34 Pld1−/−/Pld2−/− vessels, no full occlusion occurred within the observation period of 40 min, whereas only four of 38 wild-type vessels remained open. In the cases in which full occlusion of the vessels was observed, time to full occlusion was significantly prolonged in Pld1−/−/Pld2−/− mice (15.5 ± 3.9 min vs. 19.8 ± 7.7 min; Fig. 5C,D). This indicates that the two PLD isoforms do have partially redundant functions that are relevant for thrombus formation in vivo.

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Figure 5.  Mice lacking both phospholipase D isoforms showed defective thrombus formation in vivo. (A–D) Thrombus formation in small mesenteric arterioles was induced by topical application of 20% FeCl3. For monitoring of thrombus formation by intravital microscopy, platelets were labeled fluorescently. Representative pictures (A) and time to stable occlusion of wild-type and Pld1−/− mice and representative pictures (C) and time to stable occlusion (D) of wild-type and Pld1−/−/Pld2−/− mice are shown. Each symbol represents one individual. *< 0.05. WT, wild-type.

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Pld1 −/− /Pld2 −/− mice are protected from ischemic stroke while having unaltered hemostasis

Next, we analyzed the effect of the combined deficiency of both PLD isoforms on the outcome after cerebral ischemia. Wild-type and Pld1−/−/Pld2−/− mice were subjected to the tMCAO model. In Pld1−/−/Pld2−/− mice, infarct volumes 24 h after reperfusion were reduced to ∼ 60% as compared with the infarct volumes of wild-type mice (87.0 ± 24.8 mm3 vs. 51.5 ± 31.0 mm3; Fig. 6A). Reduction of infarct size was functionally relevant, as the Bederson score, which assesses global neurologic function, was significantly better in Pld1−/−/Pld2−/− mice (2.9 ± 0.6 vs. 1.9 ± 0.9; Fig. 6B). The grip test, which specifically measures motor function and coordination, showed a similar tendency, although the difference was not significant (3.9 ± 1.6 vs. 4.6 ± 0.5, data not shown). The protection of Pld1−/−/Pld2−/− mice was similar to that observed in Pld1−/− mice [23], suggesting that PLD2 made only a very minor contribution to infarct growth in this model.

image

Figure 6. Pld1−/−/Pld2−/− mice are protected from ischemic stroke and have normal tail bleeding times. (A) Representative pictures of three corresponding coronal sections and brain infarct volumes of wild-type and Pld1−/−/Pld2−/− mice that were subjected to the transient middle cerebral artery occlusion (tMCAO) model. Data are mean ± standard deviation of 10 mice per group. (B) Neurologic Bederson score assessed 24 h after tMCAO. (C) Tail bleeding times of wild-type and Pld1−/−/Pld2−/− mice. Each symbol represents one individual. *< 0.05. WT, wild-type.

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To test whether the reduced thrombus stability in Pld1−/−/Pld2−/− mice impaired hemostasis, tail bleeding experiments were performed. No significant differences in bleeding times were found between wild-type and Pld1−/−/Pld2−/− mice (Fig. 6D), demonstrating that PLD activity is dispensable for normal hemostasis.

Discussion

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. Disclosure of Conflict of Interests
  9. References
  10. Supporting Information

Our results show, for the first time, that PLD2 is dispensable for platelet production and function. However, combined deficiency of PLD1 and PLD2 resulted in a selective defect in α-granule secretion in response to submaximal thrombin stimulation and protection in a model of arterial thrombosis, making PLDs attractive targets for safe antithrombotic treatments.

PLD has been implicated in many elementary cell functions, leading to the assumption that the enzyme might be required for development [9]. In line with this assumption, the first reports on Pld knockout mice led to the hypothesis that the absence of one PLD isoform can be compensated for by the other [23,37]. However, the present study, which is the first report on mice lacking both PLD isoforms, shows that these animals do not display obvious defects, indicating that PLD is not required for normal development. Most previous studies on the potential role of PLD were performed with inhibitors, mostly primary alcohols, which might have off-target effects and thereby lead to an overestimation of the significance of PLD. Thus, previous results obtained with inhibitors should be carefully re-evaluated, making use of knockout mice.

Previously, high constitutive PLD activity has been reported in platelets [20] and attributed to PLD2, owing to its high basal activity in vitro [16]. In contrast to this, an older study showed negligible basal PLD activity [19], which we also observed (Fig. 1A). This discrepancy might be explained by the fact that the authors of the former study used [3H]arachidonic acid to measure PLD activity, which might have caused platelet activation via the thromboxane receptor triggering PLD activity [19]. In our analyses, PLD activity upon platelet stimulation in Pld2−/− platelets was decreased as compared with PLD activity in wild-type platelets, demonstrating that both PLD isoforms are inducible upon platelet stimulation but that, in line with previous reports [20,23], PLD1 appears to be the major PLD isoform in platelets.

Our results show that the decreased PLD activity resulting from the absence of PLD2 alone does not alter platelet function, at least under the experimental conditions used in this study. Thus, we generated mice lacking both PLD isoforms to determine potential redundant functions of the two enzymes. In our initial report on Pld1−/− mice [23], we observed a slight defect in integrin activation upon submaximal platelet activation with thrombin/PAR4-activating peptide or CRP. In the present study, we did not detect reduced CRP responses in Pld1−/− mice (Fig. 4B,D), whereas the defect downstream of thrombin or GPIb (as assessed by flow adhesion experiments with a VWF-coated surface) persisted (Fig. 4B,D and data not shown). It is of note that, in contrast to our initial report, for which we used mice on an Sv129 × C57Bl/6 mixed background, for this study we used mice on a pure C57Bl/6 background. The different genetic background of the mice might affect GPVI responses to CRP, e.g. by slight alterations of GPVI surface expression levels. Previously, it has been demonstrated that, especially, CRP responses depend to a great extent on GPVI expression levels [38], providing a possible explanation for this discrepancy between this study and our previous report [23]. Alternatively, GPVI signaling could be influenced by modifier genes that differ between the two mouse strains. Such a strong impact of modifiers was reported to be the reason for variable in vivo results with Gp6−/− mice [39].

The direct comparison of Pld1−/− with Pld1−/−/Pld2−/− platelets (Fig. 4) revealed a strong defect in integrin activation and α-granule release downstream of submaximal stimulation with thrombin, which was not observed in either single-knockout platelet population. Our observation of reduced secretion in PLD-deficient platelets contradicts a recently proposed model suggesting that PLD2 is a negative regulator of platelet degranulation [26]. However, the authors of that study used an inhibitor (5-fluoro-2-indolyl des-chlorohalopemide [FIPI]) that not only blocks both PLD isoforms but might also have off-target effects. This is especially problematic in that particular case, as the FIPI concentration was titrated until effects on platelet degranulation (not on PLD activity) could be observed. Consequently, the inhibitor concentration used by Elvers et al. was 100-fold higher than that necessary to abrogate PLD activity [25], suggesting that this may not have been the optimal approach to study PLD function. The α-granule release defect in platelets lacking both PLD isoforms might at least partially explain the more pronounced defect in integrin activation, as reduced α-granule secretion results in reduced mobilization of intracellularly stored αIIbβ3 to the platelet surface.

A role for PLD in secretion and secretory vesicle formation has been described for several cell types [9], including platelets [19,20]. However, degranulation of α-granules or dense granules was not affected in platelets lacking only one PLD isoform, whereas the combined deficiency resulted in defective α-granule secretion downstream of submaximal thrombin stimulation. PKC is essential for granule secretion [40], and PLD and PKC regulate each other via a positive feedback loop [41]. However, it is difficult to explain why an effect of PLD on PKC would affect secretion from α-granules but not from dense granules. Nevertheless, the absence of dense granules but not of α-granules in platelets lacking PKCα [42] demonstrates that one PKC isoform does have differential effects on the release of the major granule subtypes. Thus, one could speculate that the selective α-granule defect observed in platelets lacking PLD results from differential regulation of PKC isoforms. Another explanation might be that PLD, or its product PA, regulates α-granule cargo proteins and thereby the final steps of vesicle membrane fusion. It has been demonstrated that the N-ethylmaleimide-sensitive fusion protein attachment protein receptors (SNAREs) syntaxin 4 and VAMP-8 are crucial for α-granule secretion [8,43]. In vitro studies with syntaxin 4 vesicles have demonstrated that addition of PA enhances the rate of fusion [44]. Via this mechanism, PLD might promote the degranulation of platelet α-granules. Stimulation with other agonists or higher thrombin doses results in degranulation of Pld1−/−/Pld2−/− platelets similar to that observed in wild-type platelets (Fig. 4), demonstrating that PLD is not strictly required for platelet secretion. Further studies will be required to reveal the exact mechanism whereby PLD contributes to platelet degranulation.

In conclusion, we have shown that PLD1 and PLD2 have redundant functions in platelet α-granule secretion downstream of protease-activated receptors. Considering that current PLD inhibitors are isoform-selective at best, but definitively not specific, the observation that mice lacking both PLD isoforms display no obvious hemostatic defect is of significant interest. Thus, interfering with PLD activity and thereby dampening GPIb-triggered integrin activation [23] and platelet degranulation might be a promising and safe strategy for antithrombotic therapy.

Disclosure of Conflict of Interests

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. Disclosure of Conflict of Interests
  9. References
  10. Supporting Information

This work was supported by the Deutsche Forschungsgemeinschaft (grant Ni556/8-1 to B. Nieswandt and grant SFB 688 to B. Nieswandt and G. Stoll). The other authors state that they have no conflict of interest.

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  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. Disclosure of Conflict of Interests
  9. References
  10. Supporting Information
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Supporting Information

  1. Top of page
  2. Abstract
  3. Introduction
  4. Materials and methods
  5. Results
  6. Discussion
  7. Acknowledgements
  8. Disclosure of Conflict of Interests
  9. References
  10. Supporting Information

Data S1. Methods.

Figure S1. Spreading is not affected by the lack of PLD2.

Figure S2. vWF secretion of Pld1-/-- or Pld2-/- platelets is unaltered.

Figure S3. α-granule localization and abundancy is unchanged in PLD deficient platelets.

Table S1. Absence of PLD2 does not affect platelet glycoprotein expression.

Table S2. Lack of PLD1 and PLD2 has no effect on platelet glycoprotein expression.

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JTH_4924_sm_FigureS1-S3-TableS1-S2.pdf175KSupporting info item

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