1. Introduction
Vascular intimal injury occurs following cardiovascular disease treatments including venous bypass grafting, arteriovenous dialysis accesses, balloon angioplasty, and stenting [1,2]. Intimal hyperplasia progresses after vascular intimal injury, which gradually narrows the vessel lumen, reduces blood flow, and eventually induces ischemia with life-threatening consequences for patients [1,3]. As the main component of the media vasculature, vascular smooth muscle cells (VSMCs) abnormally migrate, proliferate, and differentiate to induce the thickening of the vascular wall and narrowing of the vascular lumen after vascular intimal injury [3,4]. Nonetheless, molecular mechanisms underlying intimal hyperplasia after injury have remained incompletely understood.
Platelets are anucleated fragments that are formed and released into the bloodstream by precursor cells called megakaryocytes in the bone marrow [5,6]. After the intimal injury, platelets are recruited and attached to the site of injury, and the adhered platelets are activated by exposed collagen to produce massive platelet-derived microvesicles (PMVs) [7]. Then, the fenestrae on the internal elastic lamina (1-3 μm in width) allow PMVs (10-1000 nm in diameter) to diffuse into the middle layer [2,8]. This process has been found to participate in various cardiovascular diseases including intimal hyperplasia [2].
Podosomes, as highly dynamic adhesion structures, are identified early on as structures mediating cell-matrix contact [9]. They have been recognized as multipurpose structures that mediate various functions, such as mechanosensing, matrix degradation, and directional migration, allowing cells to sense and interact with the surroundings [9,10]. Podosomes secrete matrix metalloproteinase-2 (MMP-2), and MMP-9 and deliver membrane type 1-MMP to the tips through microtubules in a variety of cells including VSMCs, which induces ECM degradation [11]. In addition, VSMCs leading edge exhibits numerous podosomes during the wound healing process to assist cell directional migration [12].
Our recent work has established a direct causal role of PMVs in modulating VSMC energy metabolism, migration, and dedifferentiation [2,13,14]. In the present study, we explored the role of PMVs in podosome formation and VSMC migration. Indeed, we identified a novel role of podosome in mediating MMP-9 dependent VSMCs migration after co-culture with platelets. Activated platelets indirectly induced VSMC migration via releasing PMVs. Podosome was formed in VSMCs to induce cell migration. Of note, MMP-9 activity was robustly activated in VSMCs to implement the function of the podosome. Collectively, our data elucidate a new mechanism that PMVs promote VSMC migration via forming podosomes and inducing MMP-9 activity.
2. Materials and methods
2.1. Platelet isolation, purification and activation
As shown in Fig. 1A, Sprague Dawley (SD) Rat blood was collected from the right cardiac ventricle into 3.2% sodium citrate buffer (pH 7.4) and diluted with an equal volume of 37 °C normal saline (apyrase 1 U/ml, A6410 Sigma, PGE1 0.1 μg/ml, P5515 Sigma). Platelet-rich plasma (PRP) was prepared by centrifugation at 1300 g for 10 min, adding 1 U/ml apyrase, 0.1 μg/ml PGE1, and 5 mM EDTA. Platelets were prepared from PRP by centrifugation at 2100 g for 10 min and resuspended at 108 platelets/ml in HEPES/Tyrode's buffer [15,16,17]. The resuspended platelets were activated with 1 μg/ml collagen I (C3867, Sigma) at 37 °C for 1 h as described previously [18].
Fig. 1. Platelet microvesicles (PMVs) drive VSMCs migration. The platelets were collected and activated, VSMCs were isolated, identified, and treated with PMVs. (A) The flow chart showed the processes of platelet isolation, purification, and activation. (B) Representative immunofluorescent image of VSMCs. VSMCs were identified by SMA (red). Nuclei (blue) were stained with DAPI. Scale bar = 50 μm. (C) Schematic diagram of VSMCs co-cultures with platelets. (D) Representative images of VSMC migration. VSMCs were co-cultured with platelets or HEPES/Tyrode's buffer for 24 h and images were captured at 0 h, 3 h, 6 h, 9 h, 12 h, and 24 h respectively. The migration rate was detected by the wound healing assay. Scale bar = 200 μm. (E) The line chart showed the wound closure percentage of VSMCs after co-culture with platelets (n = 4). Data are presented as Mean ± SEM, ∗∗P < 0.01. |
2.2. Cell culture
Primary VSMCs were obtained from the thoracic aortas of male SD rats via an explant method and cultured as previously described [2,19,20,21,22,23]. Briefly, Male SD rats (160-180 g) were euthanized with intraperitoneal pentobarbitone injection, and thoracic aortas was cut off. After the removal of the adventitia and endothelium, the media of thoracic aortas were isolated surgically and minced into small pieces, which were plated onto a 21 cm2 cell culture dish. VSMCs were cultured in DMEM (Gibco) containing 10% calf serum (FCS, Gibco), and 1% penicillin-streptomycin (Gibco) at 37 °C in a 5% CO2 atmosphere. The VSMCs were characterized by antibodies specific to α-smooth muscle actin (1:200) [24]. In all of the experiments, the purity of the cell populations was universally higher than 95%, and the VSMC monolayers were passaged every 2-3 days after trypsinization. Only VSMCs between passages 4 and 7 were used for the following experiments.
2.3. Platelets and VSMCs co-culture system
Coverslips suitable for six-well plates were treated with 0.1% gelatin at 37 °C for 30 min. VSMCs suspension with a cell density of 5 × 105/well was placed on the surface of coverslips in the six-well plate. Transwells with 1.0 μm pores were immersed into the plate carefully with a sterile forceps and 1 ml activated platelets were added in the upper chamber. HEPES/Tyrode's buffer was used as a control. The co-culture system was placed at 37 °C with 5% CO2 for 24 h, and then the cells on the coverslips were used for follow-up experiments (Fig. 1C).
2.4. Wound healing assay
The wound healing assay was performed as described before [2]. After VSMCs reached 95% confluence, a line was scratched across a monolayer of cells using a sterile 200-μL pipette tip. Images of the scratched line were captured at 0 h, 3 h, 6 h, 9 h, 12 h, and 24 h, respectively, after co-culture with activated platelets using a microscope (4X objective, IX-71, Olympus, Japan). The wound area was measured using ImageJ software (NIH, USA). The cell migration activity was calculated as (S0 - St)/S0, where S0 is the wound area at the initial time point, and St is the wound area at the observation time point.
2.5. GO analysis
Human platelet protein composition was obtained from the primary source literature as described previously [25]. The top 300 proteins expressed in platelets were used for GO analysis by DAVID (https://david.ncifcrf.gov/tools.jsp). Nine biological processes associated with VSMC phenotype were elected and 60 proteins included in these processes were categorized by Cellular Component.
2.6. Immunofluorescent staining
Cells were fixed in 4% paraformaldehyde for 15 min and permeabilized with 0.3% Triton-X for 5 min. After washing 3 times in PBS, cells were blocked with 10% goat serum (Gibco) for 1 h and then immunostained with specific antibodies against cortactin (1:200, Cell Signaling Technology) at 4 °C overnight. After washing in PBS for 3 times, the coverslips were incubated with a green fluorochrome-conjugated second antibody and rhodamine phalloidin (1:100, Cytoskeleton) in a blocking buffer for 1 h. For gelatin degradation, cells were incubated with rhodamine phalloidin for 30 min only. Coverslips were sealed by antifade mounting medium with DAPI (Beyotime). Images were captured using laser-scanning confocal microscopy (Olympus IX81). Ten random fields were selected and the podosome number was calculated with Image J software (NIH, USA).
2.7. Gelatin zymography
The protein concentration was measured by a BCA protein assay kit (Beyotime) and diluted to the same concentration. Then the samples were added to 7.5% polyacrylamide gels containing sodium dodecyl sulfate (SDS) and 0.1% gelatin. After electrophoresis, the gels were washed 2 times with eluent solution (2.5% Triton X-100, 50 mM Tris-HCl, 5 mM CaCl2, PH 7.5) for 30 min to remove the SDS. Then the gels were washed with incubation solution (50 mM Tris-HCl, 10 mM CaCl2, 0.9% NaCl, PH 7.5) for 10 min. With fresh incubation solution, the gels were incubated at 37 °C for 24 h to develop zymolytic bands. Finally, the gels were stained with 0.3% Coomassie Brilliant Blue G-250 for 30 min and decolored with boiling water for 5 min. The different protease bands were analyzed with ImageJ software (NIH, USA).
2.8. Gelatin degradation assay
The coverslips were coated with fluorescent gelatin as previously described [26]. Briefly, the Oregon Green 488-conjugated gelatin (G13186, Sigma) was prepared according to the manufacturer's instructions. The coverslips were coated with 50 μg/ml poly-L-lysine and incubated with 0.5% glutaraldehyde. Gelatin was used to coat the coverslips in 24 well plates at 37 °C and incubated at 4 °C in the dark for 10 min. Then, washed the coverslips three times with PBS and added 500 μl of freshly made 5 mg/ml sodium borohydride (NaBH4) for 15 min at room temperature to reduce and inactivate residual glutaraldehyde. Washed each well three times with PBS and incubated coverslips in 70% ethanol for 30 min at room temperature. Rinsed the coverslips three times with sterile PBS and transferred the coverslips to a new 6-well plate. VSMCs were placed on the coated coverslips and cultured in the co-culture system. Gelatin degradation (visualized as darker areas) was captured by confocal microscopy (LV1000; Olympus, Tokyo, Japan). Five random fields were selected and the degradation area was calculated using Image J software (NIH, USA).
2.9. Statistical analysis
All data are expressed as Mean ± SEM. Statistical analyses were performed using Student's t-test for comparison between the two groups. All analyses were performed using GraphPad Prism Software. A p value of <0.05 was considered to be statistically significant and is presented as ∗∗p < 0.01, or ∗∗∗p < 0.001.
3. Results
3.1. Platelets promote VSMC migration in a contactless way
Platelets can be internalized by VSMCs and can induce VSMC differentiation [15]. Besides, we found that PMVs released by platelets can adhere to VSMCs to promote their migration [2,18]. To distinguish the effects of platelets and their secretions on VSMC function, VSMCs were co-cultured with activated platelets. Platelets were isolated and activated following the procedures as shown in Fig. 1A. VSMCs were identified using α-smooth muscle actin by immunofluorescent (Fig. 1B). The contactless crosstalk between platelets and VSMCs was performed by co-culture system (Fig. 1C). Notably, platelets were blocked by a permeable membrane of transwell while PMVs moved freely. Wound healing assay indicated that freely diffused PMVs contact and promote VSMC migration in a time-dependent way (Fig. 1D). Of note, PMVs robustly increased wound closure percent from 34.8% to 63.9% after 24 h treatment (Fig. 1E). These results indicated that PMVs promote VSMC migration.
3.2. PMVs promote VSMC migration via inducing podosomes formation
Podosomes are highly dynamic adhesion structures to mediate cell-matrix contact [9]. They have been found to implicate in mechanosensing, matrix degradation, and directional migration [9,10]. Here, we analyzed the human platelet protein composition by GO analysis using the data from the primary source [25]. The top 300 proteins expressed in platelets were used for GO analysis by DAVID and 9 biological processes associated with VSMC phenotype including Positive Regulation of Podsome Assembly were elected (Fig. 2A). Besides, 60 proteins were found to participate in these processes and the 60 proteins were retrieved to explore the Cellular Component containing these proteins, Extracellular Exosome was the one that contains the most (51/60, 85%) of these proteins (Fig. 2B). Therefore, we explored the presence of podosomes in VSMCs. Podosomes were found in VSMCs by immunofluorescent staining of cortactin (Fig. 2C), which is an Src substrate colocalizing with F-actin in podosomes and was used as a marker of podosome that was found to co-localize with F-actin cores [27,28]. To investigate whether podosome was modulated by PMVs, VSMCs were treated with 1 ml activated platelets or 1 ml HEPES/Tyrode's buffer using a transwell co-culture system. Ten random fields of every group were analyzed to quantify the number of podosomes. The average number of podosomes per VSMC significantly increased from 2.303 ± 0.312 to 4.265 ± 0.830 after co-culture with platelets (Fig. 2D and E). Therefore, podosome as an invasive protrusion was formed in VSMCs to induce cell migration.
Fig. 2. PMVs drive VSMC migration via inducing podosome formation. Human platelet protein composition was obtained from the primary source [25] and used for GO analysis. (A) The top 300 proteins expressed in platelets were used for GO analysis by DAVID (https://david.ncifcrf.gov/tools.jsp). Nine biological processes associated with VSMC phenotype were elected, platelets were found to participate in the positive regulation of podosome assembly and 60 proteins were found to involve in these processes. (B) Sixty proteins included in 9 biological processes were categorized by Cellular Component analysis, 51 of them were found in extracellular exosomes. (C) Representative immunofluorescent image of podosomes in VSMCs. Podosomes were identified based on the co-localization of cortactin (green) with F-actin (red). Nuclei (blue) were stained with DAPI. Scale bar = 50 μm. The platelets were collected and activated, and VSMCs were isolated, identified, and treated with PMVs. (D) Representative immunofluorescent image of podosomes in VSMCs, indicating podosome formation after PMV treatment. Podosomes were identified based on the co-localization of cortactin (green) with F-actin (red). Nuclei (blue) were stained with DAPI. Arrows indicate the podosomes in the PMVs treating group compared to the control group. Scale bar = 50 μm. (E) Quantitative analysis of podosome number. Platelets increased the number of podosomes in VSMC. Data are presented as Mean ± SEM, ∗∗∗P < 0.001. |
3.3. PMVs promote VSMC migration via elevating MMP-9 activity
Podosomes are proven to execute their function via secreting MMPs [11]. Therefore, VSMC MMP-9 activity was detected by gelatin zymography. VSMCs were treated with 1 ml activated platelets or 1 ml HEPES/Tyrode's buffer for 24 h and harvested for gelatin zymography assay. The activity of MMP-9 in VSMCs increased significantly with a 10.71-fold upregulation after co-culture with platelets compared to the control group (Fig. 3A and B). Furthermore, the activity of MMP in the intact cells after co-culture with platelets was detected. Gelatin degradation assay showed that MMP activity was strongly activated after co-culture with platelets (Fig. 3C and D). These results indicated that MMP-9 activity was robustly activated in VSMCs to implement the function of the podosome. Taken together, our data elucidated a novel mechanism that PMVs promote VSMC migration via forming podosomes and inducing MMP-9 activity robustly (Fig. 4).
Fig. 3. PMVs drive VSMC migration via inducing MMP-9 activity in podosomes. The platelets were collected and activated, and VSMCs were isolated, identified, and treated with PMVs. (A) The MMP activity in VSMCs co-cultured with platelets or HEPES/Tyrode's buffer for 24 h was detected by gelatin zymography. (B) Quantitative analysis of MMP-9 activity. Platelets increased the activity of MMP-9 in VSMC. (C) Gelatin degradation was detected using a gelatin degradation assay. VSMCs were stained with DAPI (blue) and rhodamine-phalloidin (red). Oregon Green 488 gelatin was green. Arrows indicate the degradation areas (dark). Scale bar = 30 μm. (D) Quantitative analysis of gelatin degradation. Data are presented as Mean±SEM, ∗∗P<0.01, ∗∗∗P<0.001. |
Fig. 4. PMVs drive VSMC migration by forming podosomes and promoting MMP-9 activity. The number of podosomes was increased, MMP-9 activity was elevated and the VSMC migration rate was promoted after PMV treatment. |
4. Discussion
The important finding of the present study is the demonstration of the function of podosome in mediating MMP-9 dependent VSMC migration after PMVs treatment (Fig. 4). Collagen I activated platelets promote VSMC podosome (an invasive protrusion) formation via releasing PMVs. Besides, MMP-9 activity is strongly activated in VSMCs to implement the function of the podosome. Finally, this leads to VSMC migration. Taken together, our findings in the present study reveal a novel mechanism that PMVs promote VSMC migration via forming podosomes and inducing MMP-9 activity.
Under the physiological condition, platelets flow is close to the blood vessel wall under laminar shear stress and the platelets are inactivated by prostacyclin derived from endothelial cells (ECs) [29]. However, the platelets are activated by collagen after vascular intimal injury to promote VSMC dedifferentiation, proliferation, and migration [30,31,32]. The activated platelets released numerous microvesicles (PMVs) which are rich in proteins, lipids, and genetic information [13]. We had elucidated that PMVs regulate VSMC energy metabolism via α catalytic subunit of AMP-activated protein kinase after intimal injury [13]. Besides, PMVs promoted VSMC dedifferentiation via Src/Lamtor1/mTORC1 axis after intimal injury [14]. Of note, PMVs were also found to increase Col8a1 secretion and vascular stiffness in the intimal injury model [18]. The present research is consistent with our previous findings that platelets induce VSMC migration via PMVs, we further verified that platelets induced VSMC migration via podosomes.
Podosomes are found in lots of cell lines including ECs, VSMCs, osteoclasts, macrophages, and dendritic cells [9]. The size of the podosome ranges from 0.5 to 1 μm in diameter and from 0.5 to 0.8 μm in height. As multipurpose structures, podosomes were found to mediate a variety of functions including matrix degradation, mechanosensing, and directional migration [10,33]. These functions enable cells to sense and interact with their environment and participate in its remodeling [10,33]. However, the precise role of podosomes in VSMC and vascular remodeling after intimal injury has remained unknown. Cortactin was used as a marker for the identification of podosome in this research. The number of yellow pixels, which represent the co-location of F-actin and cortactin, were counted as podosome numbers. Podosomes can hardly be observed in the normal VSMC. According to our results, one normal VSMC contains no more than two podosomes on average. And it seems that podosomes are more inclined to a single arrangement with minimal rings and clusters in the normal VSMC. However, podosome rings and clusters appear more frequently after platelet treatment. In the present study, we observed that podosomes are formed in VSMCs to induce cell migration. Importantly, MMP-9, a gelatinase involved in the breakdown of extracellular matrix during the progress of many vascular diseases [34,35], was robustly activated in VSMCs to implement the function of the podosome. Therefore, our data elucidate a novel mechanism that PMVs induce podosome formation and increase MMP-9 activity to promote VSMC migration.
In summary, PMVs drive VSMC migration via forming podosomes and promoting MMP-9 activity. Notably, the molecular mechanism in depth of podosomes in vascular remodeling after vascular intimal injury needs to be explored in our future research.
Conflict of interest
None declared.
Acknowledgment
This research was supported by grants from the National Natural Science Foundation of China [grant numbers 12032003 and 12102261].
