RESEARCH ARTICLE Cytoskeleton, May 2015 72:246–255 (doi: 10.1002/cm.21221) C V
2015 Wiley Periodicals, Inc.
Quantitative Single-Cell Motility Analysis of Platelet-Rich Plasma-Treated Endothelial Cells In Vitro Tomoyuki Kawase,1,2* Takaaki Tanaka,3 Kazuhiro Okuda,4 Makoto Tsuchimochi,2,5 Masafumi Oda,6 and Toshiaki Hara7 1
Division of Oral Bioengineering, Institute of Medicine and Dentistry, Niigata University, Niigata, Japan Advanced Research Center, the Nippon Dental University School of Life Dentistry at Niigata, Niigata, Japan 3 Department of Materials Science and Technology, Niigata University, Niigata, Japan 4 Division of Periodontology, Institute of Medicine and Dentistry, Niigata University, Niigata, Japan 5 Department of Oral and Maxillofacial Radiology, the Nippon Dental University School of Life Dentistry at Niigata, Niigata, Japan 6 Institute of Research Collaboration and Promotion, Niigata University, Niigata, Japan 7 Department of Mechanical and Control Engineering, Niigata Institute of Technology, Kashiwazaki, Japan 2
Received 9 December 2014; Revised 22 February 2015; Accepted 25 March 2015 Monitoring Editor: George Bloom
Platelet-rich plasma (PRP) has been widely applied in regenerative therapy due to its high concentration of growth factors. Previous in vitro and in vivo studies have provided evidence supporting the angiogenic activity of PRP. To more directly demonstrate how PRP acts on endothelial cells, we examined the PRP-induced changes in the motility of human umbilical vein endothelial cells by examining the involvement of VEGF. Time-lapse quantitative imaging demonstrated that in the initial phase (2 h) of treatment, PRP substantially stimulated cell migration in a wound-healing assay. However, this effect of PRP was not sustained at significant levels beyond the initial phase. The average net distance of cell migration at 10 h was 0.45 6 0.16 mm and 0.82 6 0.23 mm in control and PRP-stimulated cells, respectively. This effect was also demonstrated with recombinant human VEGF and was significantly attenuated by a neutralizing anti-VEGF antibody. Immunofluorescent examination of paxillin and actin fibers demonstrated that PRP concomitantly upregulated focal adhesion and cytoskeletal formation. Western blotting analysis of phosphorylated VEGFR2 demonstrated that PRP mainly stimulated the phosphorylation of immature VEGFR2 in a dose- and timedependent manner, an action that was completely blocked by the neutralizing antibody. Taken together, these data suggest that PRP acts directly on endothelial Additional Supporting Information may be found in the online version of this article. *Address correspondence to: Tomoyuki Kawase, Division of Oral Bioengineering, Institute of Medicine and Dentistry, Niigata University, Niigata 951-8514, Japan. E-mail: [email protected]. ac.jp Published online 6 April 2015 in Wiley Online Library (wileyonlinelibrary.com).
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cells via the activation of VEGFR2 to transiently upregulate their motility. Thus, the possibility that PRP desensitizes target endothelial cells for a relatively long period of time after short-term activation should be considered when the controlled release system of PRP components is designed. V 2015 Wiley Periodicals, Inc. C
Key Words:
platelet-rich plasma; human umbilical vein endothelial cells (HUVECs); wound healing assay; time-lapse imaging; cell motility; vascular endothelial growth factor
Introduction
P
latelet-rich plasma (PRP) is the plasma fraction that contains high levels of concentrated platelets and their growth factors. Since Marx demonstrated the clinical applicability and effectiveness of PRP in tissue regeneration [Marx, 2004], PRP has been widely applied to regenerative therapies in various fields, including oral surgery, plastic surgery, and orthopedic surgery [Dhillon et al., 2012; Albanese et al., 2013]. In clinical studies [Okuda et al., 2005, 2009, 2013; Yamamiya et al., 2008; Nagata et al., 2012], we have demonstrated that PRP, alone or in combination with cultured periosteal sheets or crushed bone, facilitated periodontal tissue regeneration and alveolar bone formation. Although the effect of PRP on skeletal tissue regeneration remains controversial, it is clear that PRP reproducibly and significantly facilitates wound healing and soft tissue regeneration [Akhundov et al., 2012]. Underlying this biological activity, PRP stimulates the proliferation of connective tissue fibroblasts and the formation of granulation tissue [Nakajima et al., 2012]. However, it is unclear whether concomitant blood capillary formation is due to the direct
or indirect activity of PRP via the formation of granulation tissue. Recently, several groups have demonstrated by both in vitro and in vivo experimental systems that PRP is capable of stimulating endothelial cell proliferation and migration [Kakudo et al., 2014; Roubelakis et al., 2014]. In addition, our group demonstrated that freeze-dried PRP and platelet-rich fibrin (PRF), a modified preparation of PRP [Dohan et al. 2006], have direct angiogenic activities when coated over biodegradable mesh in an in vitro wound-healing assay using human umbilical vein endothelial cells (HUVEC) or the chick chorioallantoic membrane (CAM) [Kobayashi et al., 2012; Nakajima et al., 2012; Horimizu et al., 2013]. The molecular mechanism underlying the effects of PRP has not been vigorously investigated. A recently published article demonstrated that a MAPK-dependent pathway is involved in PRP-induced endothelial cell proliferation [Kakudo et al., 2014]. Nevertheless, the effects of PRP on endothelial cell motility remain poorly understood. To confirm the direct action of PRP on endothelial cells and to demonstrate their motility in response to PRP at the singlecell level, we examined the effects of PRP on HUVECs using an in vitro wound healing assay and a time-lapse imaging system. The wound healing assay is a simple technique that has been widely applied in the study of angiogenesis. It is based on the concept that endothelial cell division represents a key step in the angiogenic process, which is critical in tissue healing [Roussy et al., 2007]. As a result of imaging analysis using cell-tracking software, we found that PRP significantly up-regulated endothelial cell motility in the initial phase via the activation of the VEGF receptor 2 (VEGFR2) and that this activity was not sustained in the following phase.
Results Stimulatory Effect of PRP on Endothelial Cell Migration
Platelet counts and the concentration of VEGF are indicated in Table I. The platelets were concentrated in PRP preparations by 5.8-fold. The VEGF concentration was determined to be 246.0 6 35.2 pg mL21 in frozen PRP preparations. Time-lapse imaging of PRP-treated HUVEC cultures was performed in accordance with the time-schedule shown Table I. Platelet Counts and VEGF Concentrations Whole blood samples
Platelets VEGF
4
25.8 6 6.5 x 10 /lL N.D.
n 5 6. N.D. represents “not determined.”
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PRP preparations
194.9 6 24.1 x 104/lL 236.5 6 37.2 pg mL21
Fig. 1. Time schedules of PRP treatment for time-lapse recording (A) and for immunofluorescence imaging and western blotting analysis (B). In the time-lapse recording, early confluent cell cultures were used. Control and VEGF-treated cells were incubated for 30 min prior to the recording. In the other experiments, subconfluent cell cultures were used.
in Fig. 1A. The tracking data collected for individual cells are shown in Fig. 2. The width of the cell-free gap zone is 500 lm. This cell-free-gap was completely closed in PRPtreated cultures but not in control cultures at 10 h of recording. The cell-tracking data clearly demonstrate the time-dependent nature of the cell migration required to close the gap in the PRP-treated cultures. Changes in the distance of cell migration between each interval (10 min) over the time course of the experiment are shown in Fig. 3 (Upper). Compared with the control cultures, cells in the PRP-treated cultures migrated more actively. Cells of the PRP-treated cultures sustained cell motility at relatively high levels, but the difference between PRP-treated and control cells was not statistically significant. The total distances of cell migration at 1 and 10 h are shown in Fig. 3 (Middle and Lower). Similar data were obtained at both time points. PRP significantly increased the total length of the migration, and this effect was significantly attenuated by the addition of an anti-VEGF neutralizing antibody (3 lg mL21). VEGF (200 ng mL21) mimicked the actions of PRP. Stimulatory Effects of PRP on the Formation of Focal Adhesions and the Actin Cytoskeleton
Cells were treated with PRP according to the time course shown in Fig. 1B. Immunofluorescence photomicrographs of paxillin, a component of focal adhesions, and polymerized actin, a major cytoskeletal fiber, are shown in Fig. 4. In control cultures, actin fibers were poorly developed and paxillin was detected in a few areas, such as at the ends of actin fibers at 1 h of treatment. PRP (0.3%) dramatically upregulated paxillin expression and actin fiber formation. This effect was mimicked by VEGF and was clearly Quantitative Analysis of PRP-induced Endothelial Cell Migration
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Fig. 2. Time-lapse images of HUVEC migration in control (Left) and PRP (0.3%)-treated cultures (Right) at the starting time point (0 h) (Upper) and 10 h (Middle). The cells included in the regions shown in broken lines were tracked. Cell tracking data along the Y axis are shown in the Lower Panels. These data are representative of the data obtained from three to five independent experiments.
attenuated by the addition of anti-VEGF antibody, as observed in the wound healing assay. Furthermore, when the neutralizing antibody was substituted with a corresponding isotype control, attenuation of neither paxillin expression nor actin fiber formation was observed (Supporting Information Fig. S1). Dose- and Time-dependent Effects of PRP on Vegfr2 Phosphorylation
The change in the effect of PRP on VEGFR2 phosphorylation over the time course of the experiment is shown in
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Fig. 5. In the non-treated basal state (control), phosphorylated VEGFR2 was widely distributed, but the levels of receptor phosphorylation were low. Treatment with PRP (0.3%) rapidly stimulated the phosphorylation of VEGFR2. However, the phosphorylated state was not maintained for a longer period of time, and it decreased to basal levels within 60 min. The corresponding isotype control did not show specific staining (Supporting Information Fig. S2). The change in the effect of PRP on the localization of VEGFR2 and VE-cadherin over the experimental time CYTOSKELETON
localization gradually reduced over time. The corresponding isotype control did not show specific staining (Supporting Information Fig. S2). Western blotting analyses of the effects of PRP on the phosphorylation of VEGFR2 are shown in Fig. 7. Consistent with the immunofluorescence findings shown in Fig. 5, PRP (1%) stimulated the phosphorylation of immature VEGFR2 (150 kDa) in a time-dependent manner. At 10 min of treatment, PRP (0.125–2%) stimulated VEGFR2 phosphorylation in a dose-dependent manner. Furthermore, the addition of anti-VEGF antibody substantially attenuated PRPinduced VEGFR2 phosphorylation at 10 min.
Discussion
Fig. 3. Quantitative analysis of HUVEC migration by timelapse recording. (Upper) The distance of cell migration per 10min interval are plotted in a time-course (control vs. PRPtreated cells). The total distance of HUVEC migration at 1 h and 10 h are shown in the Middle and Lower panels, respectively. The data were obtained from a single representative experiment, and similar data were obtained from other independent experiments. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
course is shown in Fig. 6. In the control state, both VEGFR2 and VE-cadherin were widely distributed on the plasma membrane. As observed for VEGFR2 phosphorylation, PRP (0.3%) rapidly increased the expression of the surface proteins, both of which were co-localized on the plasma membrane. The increased expression and coCYTOSKELETON
In this study, we found that PRP directly acted on HUVECs to immediately up-regulate their motility. This cellular response lasted for at least 2 h; after which, their motility declined to levels that were only slightly higher than control levels. In addition, this mechanical response involved the activation of VEGFR2 and the formation of focal adhesions. According to a review article by Lamalice et al. [2007], endothelial cell migration involves three major mechanisms—namely, chemotaxis, haptotaxis, and mechanotaxis. Chemotaxis is the directional migration toward a gradient of soluble chemoattractant such as VEGF or bFGF, whereas haptotaxis is associated with increased endothelial cell migration activated in response to an integrin binding to an ECM component. On the other hand, mechanotaxis is cell migration induced by mechanical forces, such as stiffness gradients of extracellular matrixes, substrate deformation, intercellular tension or intercellular force gradients [Gray et al., 2003]. Recently, more attention has been focused on fluid shear stress—the tangential component of hemodynamic stresses caused by blood flow—to which endothelial cells are constantly subjected in vivo. [Li et al., 2002; Lamalice et al., 2007; Lin and Helmke, 2008]. Because PRP contains VEGF and activates VEGFR2, it has been proposed that PRP is capable of stimulating chemotaxis. Furthermore, the slightly increased migration in the phase after the initial phase could be identified as haptotaxis because the addition of PRP to HUVEC cultures and fibroblasts [Kawase et al., 2003] helps to slowly convert fibrinogen into insoluble fibrin, which contains a RGD motif [Yang et al., 2004] binding site for HUVECs via av integrin [Kronenwett et al., 2002; Humphries et al. 2006]. Furthermore, it is unlikely that mechanotaxis is involved in PRP-induced endothelial cell migration because the cells were placed on a nearly uniform polystyrene culture dish under static conditions. The conventional wound healing assay demonstrates cell migration at experimental endpoints, whereas time-lapse imaging studies can track individual cells and enable the analysis of cell motility at the single-cell level. Using this
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Fig. 4. The effects of stimulation with PRP and VEGF on focal adhesion and actin fiber formation. Subconfluent HUVEC cultures on coverslips were treated with PRP or VEGF for 60 min and fixed with formalin for immunofluorescent detection of paxillin and polymerized actin. Bar 5 20 lm. Similar data were obtained from three additional independent experiments.
technique, we found that the initial acceleration phase lasted for only 2 h. This phenomenon could be explained by the depletion of VEGF ligand or by desensitization of the VEGF receptor. However, it is unlikely that VEGF could be depleted within 2 h because some VEGF is thought to be bound to fibrin and to remain functional [Chen et al., 2010]. In contrast, it has been demonstrated
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that VEGF induces VEGFR2 desensitization or downregulation rapidly and actively by two distinct mechanisms: trafficking to the endosome and interaction with VEcadherin [Bruns et al., 2010]. The activated and phosphorylated VEGFR2 is internalized within the initial 30–45 min and ubiquitinated for lysosomal degradation [Bruns et al., 2010; Nakayama et al., 2013]. This receptor CYTOSKELETON
Fig. 5. Immunofluorescence data indicating the time-course effects of PRP on VEGFR2 phosphorylation. Subconfluent HUVEC cultures were treated with PRP (0.3%) at the indicated concentrations for the indicated periods of time. Fixed cells were permeabilized for the staining. Bar 5 20 lm. Similar data were obtained from three additional independent experiments. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
internalization is balanced by a high rate of recycling [Scott and Mellor, 2009] and is required for endothelial recovery during wound healing [Santos et al., 2007]. Alternatively, the activated VEGFR2 is preserved (i.e., not internalized) on the plasma membrane and desensitized by interaction CYTOSKELETON
with VE-cadherin [Bruns et al., 2010]. Our immunofluorescent examination demonstrated that PRP reduced internalization of VEGFR2 to preserve VEGFR2 on the plasma membrane. However, this receptor preservation did not last longer than 60 min. Therefore, it seems likely that
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Fig. 6. Immunofluorescence data indicating the time-course effects of PRP on the expression and localization of VEGFR2 and VE-cadherin. Subconfluent HUVEC cultures were treated as described in the legend of Figure 5; however, fixed cells were not permeabilized for detection of the surface proteins. Bar 5 20 lm. Similar data were obtained from three additional independent experiments. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
VEGFR2 is desensitized primarily by interaction with VEcadherin and subsequently by internalization of this complex in PRP-treated HUVECs. Further investigations are required to reveal the mechanism underlying this phenomenon.
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In conclusion, a time-lapse imaging system and biochemical analysis have demonstrated that PRP transiently upregulated the motility of individual endothelial cells. Because fibrin has been demonstrated to directly act on endothelial cells to change their morphology and stimulate CYTOSKELETON
is shown in Supporting Information Fig. S3. VEGF concentrations in PRP preparations appeared to be highly correlated with platelet count in PRP preparations (R 5 0.734). The study design and consent forms for all procedures performed on the study subjects were approved by the ethical committee for human subject use at Niigata University Medical and Dental Hospital according to the Helsinki Declaration of 1975, as revised in 2008. Neutralization and Mimicking Experiments
Fig. 7. Western blotting data indicating the time- and dosedependent effects of PRP on the phosphorylation of VGFR2. Subconfluent HUVEC cultures were treated with PRP at the indicated concentrations for the indicated periods of time. At the end of the treatments, the cells were lysed for Western blotting analysis.
tube formation [Chalupowicz et al., 1995; Bach et al., 1998], endothelial cell motility could be up-regulated synergistically by VEGF and fibrin in PRP-treated HUVECs. Of further interest is that the decrement of up-regulated cell motility is preceded by the peak interaction of VEGFR2 and VE-cadherin, a receptor of fibrin. Therefore, we believe that, in addition to the activation process, it is necessary to take PRP-induced desensitization of target endothelial cells into account when the controlled release system of PRP components is designed.
Materials and Methods PRP Preparation
As previously described [Okuda et al. 2003], blood was collected from three healthy and non-smoking volunteers aged 28, 30, and 54 years (two females; one male), and PRP was prepared using the two-step centrifugation protocol. The number of platelets in freshly prepared PRP samples was determined using an automated hematology analyzer (PocH 100iV diff: Sysmex, Kobe, Japan). Then, PRP preparations were frozen and stored at 220 C until further use (usually within 2 weeks). The concentrations of VEGF in frozen PRP samples were determined using a Human VEGF Quantikine ELISA Kit (R&D Systems, Minneapolis, MN). The correlation between platelet count and VEGF concentrations CYTOSKELETON
Human recombinant VEGF and mouse monoclonal antihuman VEGF antibodies (16F1) were obtained from Life Technologies (Carlsbad, CA, USA) and Immuno-biological Laboratories (Fujioka, Japan), respectively. VEGF was diluted to a final concentration of 200 ng mL21 in medium or Hank’s balanced salt solution (HBSS) (Life Technologies). For VEGF neutralization, 3 lg mL21 anti-VEGF antibody, which is expected to block the action of VEGF (50 ng mL21) [Suzuki et al., 1999], was added to the medium or HBSS containing the indicated concentrations of PRP and incubated for 30 min at room temperature prior to experiments. As an isotype control, mouse IgG2a (Medical & Biological Laboratories, Nagoya, Japan) was used at a final concentration of 3 mg mL21. Endothelial Cell Cultures and In Vitro Wound Healing Assay
HUVECs were obtained from AllCells, LLC (Emeryville, CA) and maintained in endothelial cell medium supplemented with growth factors (HuMedia-EB2; KURABO, Osaka, Japan) [Kobayashi et al., 2012]. For the wound healing assay, HUVECs were seeded into double-well culture inserts (Model 80209; ibidi GmbH, Munich, Germany) on a 35-mm dish (Falcon 353001, Corning Incorporated, Corning, NY) at a density of 2.4–3.0 3 102 cells mm22. The time course of this experiment is shown in Fig. 1A. Briefly, after 6–8 h of incubation in a CO2 incubator (5% CO2), the culture inserts were removed and the cells were labeled with 1 lM CellTrackerTM Orange CMTMR dye (Life Technologies) for 15 min. The cells were then washed with medium and incubated in medium containing PRP or VEGF for 30 min in a CO2 incubator (5% CO2). This time period is important for the conversion of fibrinogen to fibrin. Because fibrin is insoluble and often interferes with optical examination of the sample, the culture dishes were gently shaken two or three times during this period. The culture dishes were then set on the stage incubation chamber, which is connected to a CO2, humidity and temperature controller (5% CO2) (UNO; Okolab S.r.l., Naples, Italy), and the time-lapse recording was started. The time-lapse recording system consisted of a monochrome CCD camera (QIClick; Nippon Roper, Tokyo, Japan), LED lamp (TLED Plus, Sutter Instrument, Novato, CA) and personal desktop computer (Endeavour AT992E;
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EPSON, Suwa, Japan) with imaging software (VisiView; Visitron Systems GmbH, Puchheim Germany). The fluorescent images were obtained at an interval of 10 min for a 10 h period. The data were saved as an avi file and analyzed using the particle-tracking function of ImagePro Plus software (Nippon Roper). Immunofluorescence Examination
HUVECs were seeded onto glass coverslips and precultured for 2 days to form sub-confluent cultures. The cells were washed once with warmed HBSS and treated with PRP or VEGF. At the indicated time points, the cells were fixed with 10% formalin for 15 min and washed three times with PBS containing 0.5% Tween 20 (T-PBS) as previously described [Kawase et al., 2014]. After blocking with diluted Block Ace (1:2 in washing buffer; DS Pharma, Osaka, Japan) for 60 min at room temperature, the cells were probed with a rabbit polyclonal anti-paxillin antibody (1:200) (Transduction Laboratories, Lexington, KY) or a rabbit polyclonal phospho-VEGF-R2 (Tyr996) antibody (1:100) (Cell Signaling Technology Japan, Tokyo, Japan) overnight at 4 C. When incubating with rabbit monoclonal anti-VEGF-R2 antibody (D5B1) (1:100) (Cell Signaling Technology) and mouse monoclonal anti-CD144 (VEcadherin) (1:10) (BioLegend, San Diego, CA) overnight at 4 C, fixed cells were neither permeabilized with T-PBS nor blocked with the diluted Block Ace. Instead, cells were washed with PBS and blocked with 2.5% normal horse serum (Vector Labs, Burlingame, CA). The cells were then washed three times with T-PBS or PBS and probed with AlexaFlour 488-conjugated anti-rabbit IgG, AlexaFlour 555-conjugated anti-mouse IgG or AlexaFlour 555-conjugated anti-rabbit IgG, (Abcam, Cambridge, MA) diluted in Immunoshot (Mild) (Cosmo Bio, Tokyo, Japan) for 30 min at room temperature. Simultaneously, the polymerized F-actin fibers were stained with 0.1 lM Acti-StainTM 555 fluorescent phalloidin (Alexa Fluor 555-conjugated phalloidin) (Cytoskeleton, Denver, CO) in the same solution. After three washes with T-PBS, the cells were examined under a fluorescence microscope (Nikon, Tokyo, Japan). Isotype controls for rabbit primary antibody and mouse primary antibody (Life Technologies) were used as negative controls. Western Blotting Analysis
HUVECs were seeded onto 6-well plates at a density of 1 3 104 cells/well and pre-cultured for 2 days to form subconfluent cultures. After washing as shown in Fig. 1B, the cells were treated with PRP or VEGF in HBSS in a CO2 incubator (5% CO2) for the indicated periods of time. After washing twice with ice cold PBS, the cells were lysed with Laemmli sample buffer as previously described [Uematsu et al., 2013]. Protein samples were fractionated using 10%
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SDS-PAGE (ATTO, Tokyo, Japan) and electro-blotted onto PVDF membranes using the Trans-BlotV TurboTM Transfer System (Bio-Rad Laboratories, Hercules, CA). After blocking with diluted Block A (1:2) or 5% BSA (Fraction V, Sigma, St. Louis, MO) in T-TBS for 4–5 h at 4 C, the membranes were probed with the following primary antibodies: rabbit polyclonal anti-phospho-VEGFR2 (Y996; 1:2000 in dilution; Cell Signaling Technology, Danvers, MA), rabbit polyclonal anti-VEGFR2 (D5B1; 1:2000 in dilution; Cell Signaling Technology) or anti-actin antibody (1:1000 dilution; Santa Cruz Biotechnology, Santa Cruz, CA) overnight at 4 C. After washing three times with T-TBS, the membranes were probed with horseradish peroxidase-conjugated goat polyclonal anti-rabbit IgG H&L (1:5000 in dilution; Abcam, Cambridge, MA) or horseradish peroxidase-conjugated donkey anti-goat IgG (Santa Cruz Biotechnology) for 45 min at 4 C. After washing, images were visualized using ClarityTM Western ECL Substrate (Bio-Rad) and imaged using a cooled CCD camera system (Image Capture; ATTO, Tokyo, Japan). R
Statistical Analysis
The data were reported as the mean value 6 standard deviation (S.D.). For multi-group comparisons, statistical analyses were performed to compare the mean values using oneway analysis of variance (ANOVA) followed by Dunn’s multiple comparison test (SigmaPlot 12.5; Systat Software, San Jose, CA). P values < 0.05 were considered significant.
Acknowledgments
The authors thank Dr. Mito Kobayashi (Niigata University) for her technical assistance in determining the number of platelets and the concentrations of VEGF. This project was financially supported by a Grant-in-Aid for New Market Development by the Industrial Creation for 2012 from the Niigata Industrial Creation Organization and JSPS KAKENHI (Grant #24390443 and #24390465). References Akhundov K, Pietramaggiori G, Waselle L, Darwiche S, Guerid S, Scaletta C, Hirt-Burri N, Applegate LA, Raffoul WV. 2012. Development of a cost-effective method for platelet-rich plasma (PRP) preparation for topical wound healing. Ann Burns Fire Disasters 25:207–213. Albanese A, Licata ME, Polizzi B, Campisi G. 2013. Platelet-rich plasma (PRP) in dental and oral surgery: From the wound healing to bone regeneration. Immun Ageing 10:23. Bach TL, Barsigian C, Yaen CH, Martinez J. 1998. Endothelial cell VE-cadherin functions as a receptor for the beta15-42 sequence of fibrin. J Biol Chem 273:30719–30728. Bruns AF, Herbert SP, Odell AF, Jopling HM, Hooper NM, Zachary IC, Walker JH, Ponnambalam S. 2010. Ligand-stimulated vegfr2 signaling is regulated by co-ordinated trafficking and proteolysis. Traffic 11:161–174.
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Quantitative Single-Cell Motility Analysis of Platelet-Rich Plasma-Treated Endothelial Cells In Vitro Tomoyuki Kawase,1,2* Takaaki Tanaka,3 Kazuhiro Okuda,4 Makoto Tsuchimochi,2,5 Masafumi Oda,6 and Toshiaki Hara7 1
Division of Oral Bioengineering, Institute of Medicine and Dentistry, Niigata University, Niigata, Japan Advanced Research Center, the Nippon Dental University School of Life Dentistry at Niigata, Niigata, Japan 3 Department of Materials Science and Technology, Niigata University, Niigata, Japan 4 Division of Periodontology, Institute of Medicine and Dentistry, Niigata University, Niigata, Japan 5 Department of Oral and Maxillofacial Radiology, the Nippon Dental University School of Life Dentistry at Niigata, Niigata, Japan 6 Institute of Research Collaboration and Promotion, Niigata University, Niigata, Japan 7 Department of Mechanical and Control Engineering, Niigata Institute of Technology, Kashiwazaki, Japan 2
Received 9 December 2014; Revised 22 February 2015; Accepted 25 March 2015 Monitoring Editor: George Bloom
Platelet-rich plasma (PRP) has been widely applied in regenerative therapy due to its high concentration of growth factors. Previous in vitro and in vivo studies have provided evidence supporting the angiogenic activity of PRP. To more directly demonstrate how PRP acts on endothelial cells, we examined the PRP-induced changes in the motility of human umbilical vein endothelial cells by examining the involvement of VEGF. Time-lapse quantitative imaging demonstrated that in the initial phase (2 h) of treatment, PRP substantially stimulated cell migration in a wound-healing assay. However, this effect of PRP was not sustained at significant levels beyond the initial phase. The average net distance of cell migration at 10 h was 0.45 6 0.16 mm and 0.82 6 0.23 mm in control and PRP-stimulated cells, respectively. This effect was also demonstrated with recombinant human VEGF and was significantly attenuated by a neutralizing anti-VEGF antibody. Immunofluorescent examination of paxillin and actin fibers demonstrated that PRP concomitantly upregulated focal adhesion and cytoskeletal formation. Western blotting analysis of phosphorylated VEGFR2 demonstrated that PRP mainly stimulated the phosphorylation of immature VEGFR2 in a dose- and timedependent manner, an action that was completely blocked by the neutralizing antibody. Taken together, these data suggest that PRP acts directly on endothelial Additional Supporting Information may be found in the online version of this article. *Address correspondence to: Tomoyuki Kawase, Division of Oral Bioengineering, Institute of Medicine and Dentistry, Niigata University, Niigata 951-8514, Japan. E-mail: [email protected]. ac.jp Published online 6 April 2015 in Wiley Online Library (wileyonlinelibrary.com).
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cells via the activation of VEGFR2 to transiently upregulate their motility. Thus, the possibility that PRP desensitizes target endothelial cells for a relatively long period of time after short-term activation should be considered when the controlled release system of PRP components is designed. V 2015 Wiley Periodicals, Inc. C
Key Words:
platelet-rich plasma; human umbilical vein endothelial cells (HUVECs); wound healing assay; time-lapse imaging; cell motility; vascular endothelial growth factor
Introduction
P
latelet-rich plasma (PRP) is the plasma fraction that contains high levels of concentrated platelets and their growth factors. Since Marx demonstrated the clinical applicability and effectiveness of PRP in tissue regeneration [Marx, 2004], PRP has been widely applied to regenerative therapies in various fields, including oral surgery, plastic surgery, and orthopedic surgery [Dhillon et al., 2012; Albanese et al., 2013]. In clinical studies [Okuda et al., 2005, 2009, 2013; Yamamiya et al., 2008; Nagata et al., 2012], we have demonstrated that PRP, alone or in combination with cultured periosteal sheets or crushed bone, facilitated periodontal tissue regeneration and alveolar bone formation. Although the effect of PRP on skeletal tissue regeneration remains controversial, it is clear that PRP reproducibly and significantly facilitates wound healing and soft tissue regeneration [Akhundov et al., 2012]. Underlying this biological activity, PRP stimulates the proliferation of connective tissue fibroblasts and the formation of granulation tissue [Nakajima et al., 2012]. However, it is unclear whether concomitant blood capillary formation is due to the direct
or indirect activity of PRP via the formation of granulation tissue. Recently, several groups have demonstrated by both in vitro and in vivo experimental systems that PRP is capable of stimulating endothelial cell proliferation and migration [Kakudo et al., 2014; Roubelakis et al., 2014]. In addition, our group demonstrated that freeze-dried PRP and platelet-rich fibrin (PRF), a modified preparation of PRP [Dohan et al. 2006], have direct angiogenic activities when coated over biodegradable mesh in an in vitro wound-healing assay using human umbilical vein endothelial cells (HUVEC) or the chick chorioallantoic membrane (CAM) [Kobayashi et al., 2012; Nakajima et al., 2012; Horimizu et al., 2013]. The molecular mechanism underlying the effects of PRP has not been vigorously investigated. A recently published article demonstrated that a MAPK-dependent pathway is involved in PRP-induced endothelial cell proliferation [Kakudo et al., 2014]. Nevertheless, the effects of PRP on endothelial cell motility remain poorly understood. To confirm the direct action of PRP on endothelial cells and to demonstrate their motility in response to PRP at the singlecell level, we examined the effects of PRP on HUVECs using an in vitro wound healing assay and a time-lapse imaging system. The wound healing assay is a simple technique that has been widely applied in the study of angiogenesis. It is based on the concept that endothelial cell division represents a key step in the angiogenic process, which is critical in tissue healing [Roussy et al., 2007]. As a result of imaging analysis using cell-tracking software, we found that PRP significantly up-regulated endothelial cell motility in the initial phase via the activation of the VEGF receptor 2 (VEGFR2) and that this activity was not sustained in the following phase.
Results Stimulatory Effect of PRP on Endothelial Cell Migration
Platelet counts and the concentration of VEGF are indicated in Table I. The platelets were concentrated in PRP preparations by 5.8-fold. The VEGF concentration was determined to be 246.0 6 35.2 pg mL21 in frozen PRP preparations. Time-lapse imaging of PRP-treated HUVEC cultures was performed in accordance with the time-schedule shown Table I. Platelet Counts and VEGF Concentrations Whole blood samples
Platelets VEGF
4
25.8 6 6.5 x 10 /lL N.D.
n 5 6. N.D. represents “not determined.”
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PRP preparations
194.9 6 24.1 x 104/lL 236.5 6 37.2 pg mL21
Fig. 1. Time schedules of PRP treatment for time-lapse recording (A) and for immunofluorescence imaging and western blotting analysis (B). In the time-lapse recording, early confluent cell cultures were used. Control and VEGF-treated cells were incubated for 30 min prior to the recording. In the other experiments, subconfluent cell cultures were used.
in Fig. 1A. The tracking data collected for individual cells are shown in Fig. 2. The width of the cell-free gap zone is 500 lm. This cell-free-gap was completely closed in PRPtreated cultures but not in control cultures at 10 h of recording. The cell-tracking data clearly demonstrate the time-dependent nature of the cell migration required to close the gap in the PRP-treated cultures. Changes in the distance of cell migration between each interval (10 min) over the time course of the experiment are shown in Fig. 3 (Upper). Compared with the control cultures, cells in the PRP-treated cultures migrated more actively. Cells of the PRP-treated cultures sustained cell motility at relatively high levels, but the difference between PRP-treated and control cells was not statistically significant. The total distances of cell migration at 1 and 10 h are shown in Fig. 3 (Middle and Lower). Similar data were obtained at both time points. PRP significantly increased the total length of the migration, and this effect was significantly attenuated by the addition of an anti-VEGF neutralizing antibody (3 lg mL21). VEGF (200 ng mL21) mimicked the actions of PRP. Stimulatory Effects of PRP on the Formation of Focal Adhesions and the Actin Cytoskeleton
Cells were treated with PRP according to the time course shown in Fig. 1B. Immunofluorescence photomicrographs of paxillin, a component of focal adhesions, and polymerized actin, a major cytoskeletal fiber, are shown in Fig. 4. In control cultures, actin fibers were poorly developed and paxillin was detected in a few areas, such as at the ends of actin fibers at 1 h of treatment. PRP (0.3%) dramatically upregulated paxillin expression and actin fiber formation. This effect was mimicked by VEGF and was clearly Quantitative Analysis of PRP-induced Endothelial Cell Migration
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Fig. 2. Time-lapse images of HUVEC migration in control (Left) and PRP (0.3%)-treated cultures (Right) at the starting time point (0 h) (Upper) and 10 h (Middle). The cells included in the regions shown in broken lines were tracked. Cell tracking data along the Y axis are shown in the Lower Panels. These data are representative of the data obtained from three to five independent experiments.
attenuated by the addition of anti-VEGF antibody, as observed in the wound healing assay. Furthermore, when the neutralizing antibody was substituted with a corresponding isotype control, attenuation of neither paxillin expression nor actin fiber formation was observed (Supporting Information Fig. S1). Dose- and Time-dependent Effects of PRP on Vegfr2 Phosphorylation
The change in the effect of PRP on VEGFR2 phosphorylation over the time course of the experiment is shown in
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Fig. 5. In the non-treated basal state (control), phosphorylated VEGFR2 was widely distributed, but the levels of receptor phosphorylation were low. Treatment with PRP (0.3%) rapidly stimulated the phosphorylation of VEGFR2. However, the phosphorylated state was not maintained for a longer period of time, and it decreased to basal levels within 60 min. The corresponding isotype control did not show specific staining (Supporting Information Fig. S2). The change in the effect of PRP on the localization of VEGFR2 and VE-cadherin over the experimental time CYTOSKELETON
localization gradually reduced over time. The corresponding isotype control did not show specific staining (Supporting Information Fig. S2). Western blotting analyses of the effects of PRP on the phosphorylation of VEGFR2 are shown in Fig. 7. Consistent with the immunofluorescence findings shown in Fig. 5, PRP (1%) stimulated the phosphorylation of immature VEGFR2 (150 kDa) in a time-dependent manner. At 10 min of treatment, PRP (0.125–2%) stimulated VEGFR2 phosphorylation in a dose-dependent manner. Furthermore, the addition of anti-VEGF antibody substantially attenuated PRPinduced VEGFR2 phosphorylation at 10 min.
Discussion
Fig. 3. Quantitative analysis of HUVEC migration by timelapse recording. (Upper) The distance of cell migration per 10min interval are plotted in a time-course (control vs. PRPtreated cells). The total distance of HUVEC migration at 1 h and 10 h are shown in the Middle and Lower panels, respectively. The data were obtained from a single representative experiment, and similar data were obtained from other independent experiments. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
course is shown in Fig. 6. In the control state, both VEGFR2 and VE-cadherin were widely distributed on the plasma membrane. As observed for VEGFR2 phosphorylation, PRP (0.3%) rapidly increased the expression of the surface proteins, both of which were co-localized on the plasma membrane. The increased expression and coCYTOSKELETON
In this study, we found that PRP directly acted on HUVECs to immediately up-regulate their motility. This cellular response lasted for at least 2 h; after which, their motility declined to levels that were only slightly higher than control levels. In addition, this mechanical response involved the activation of VEGFR2 and the formation of focal adhesions. According to a review article by Lamalice et al. [2007], endothelial cell migration involves three major mechanisms—namely, chemotaxis, haptotaxis, and mechanotaxis. Chemotaxis is the directional migration toward a gradient of soluble chemoattractant such as VEGF or bFGF, whereas haptotaxis is associated with increased endothelial cell migration activated in response to an integrin binding to an ECM component. On the other hand, mechanotaxis is cell migration induced by mechanical forces, such as stiffness gradients of extracellular matrixes, substrate deformation, intercellular tension or intercellular force gradients [Gray et al., 2003]. Recently, more attention has been focused on fluid shear stress—the tangential component of hemodynamic stresses caused by blood flow—to which endothelial cells are constantly subjected in vivo. [Li et al., 2002; Lamalice et al., 2007; Lin and Helmke, 2008]. Because PRP contains VEGF and activates VEGFR2, it has been proposed that PRP is capable of stimulating chemotaxis. Furthermore, the slightly increased migration in the phase after the initial phase could be identified as haptotaxis because the addition of PRP to HUVEC cultures and fibroblasts [Kawase et al., 2003] helps to slowly convert fibrinogen into insoluble fibrin, which contains a RGD motif [Yang et al., 2004] binding site for HUVECs via av integrin [Kronenwett et al., 2002; Humphries et al. 2006]. Furthermore, it is unlikely that mechanotaxis is involved in PRP-induced endothelial cell migration because the cells were placed on a nearly uniform polystyrene culture dish under static conditions. The conventional wound healing assay demonstrates cell migration at experimental endpoints, whereas time-lapse imaging studies can track individual cells and enable the analysis of cell motility at the single-cell level. Using this
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Fig. 4. The effects of stimulation with PRP and VEGF on focal adhesion and actin fiber formation. Subconfluent HUVEC cultures on coverslips were treated with PRP or VEGF for 60 min and fixed with formalin for immunofluorescent detection of paxillin and polymerized actin. Bar 5 20 lm. Similar data were obtained from three additional independent experiments.
technique, we found that the initial acceleration phase lasted for only 2 h. This phenomenon could be explained by the depletion of VEGF ligand or by desensitization of the VEGF receptor. However, it is unlikely that VEGF could be depleted within 2 h because some VEGF is thought to be bound to fibrin and to remain functional [Chen et al., 2010]. In contrast, it has been demonstrated
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that VEGF induces VEGFR2 desensitization or downregulation rapidly and actively by two distinct mechanisms: trafficking to the endosome and interaction with VEcadherin [Bruns et al., 2010]. The activated and phosphorylated VEGFR2 is internalized within the initial 30–45 min and ubiquitinated for lysosomal degradation [Bruns et al., 2010; Nakayama et al., 2013]. This receptor CYTOSKELETON
Fig. 5. Immunofluorescence data indicating the time-course effects of PRP on VEGFR2 phosphorylation. Subconfluent HUVEC cultures were treated with PRP (0.3%) at the indicated concentrations for the indicated periods of time. Fixed cells were permeabilized for the staining. Bar 5 20 lm. Similar data were obtained from three additional independent experiments. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
internalization is balanced by a high rate of recycling [Scott and Mellor, 2009] and is required for endothelial recovery during wound healing [Santos et al., 2007]. Alternatively, the activated VEGFR2 is preserved (i.e., not internalized) on the plasma membrane and desensitized by interaction CYTOSKELETON
with VE-cadherin [Bruns et al., 2010]. Our immunofluorescent examination demonstrated that PRP reduced internalization of VEGFR2 to preserve VEGFR2 on the plasma membrane. However, this receptor preservation did not last longer than 60 min. Therefore, it seems likely that
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Fig. 6. Immunofluorescence data indicating the time-course effects of PRP on the expression and localization of VEGFR2 and VE-cadherin. Subconfluent HUVEC cultures were treated as described in the legend of Figure 5; however, fixed cells were not permeabilized for detection of the surface proteins. Bar 5 20 lm. Similar data were obtained from three additional independent experiments. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]
VEGFR2 is desensitized primarily by interaction with VEcadherin and subsequently by internalization of this complex in PRP-treated HUVECs. Further investigations are required to reveal the mechanism underlying this phenomenon.
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In conclusion, a time-lapse imaging system and biochemical analysis have demonstrated that PRP transiently upregulated the motility of individual endothelial cells. Because fibrin has been demonstrated to directly act on endothelial cells to change their morphology and stimulate CYTOSKELETON
is shown in Supporting Information Fig. S3. VEGF concentrations in PRP preparations appeared to be highly correlated with platelet count in PRP preparations (R 5 0.734). The study design and consent forms for all procedures performed on the study subjects were approved by the ethical committee for human subject use at Niigata University Medical and Dental Hospital according to the Helsinki Declaration of 1975, as revised in 2008. Neutralization and Mimicking Experiments
Fig. 7. Western blotting data indicating the time- and dosedependent effects of PRP on the phosphorylation of VGFR2. Subconfluent HUVEC cultures were treated with PRP at the indicated concentrations for the indicated periods of time. At the end of the treatments, the cells were lysed for Western blotting analysis.
tube formation [Chalupowicz et al., 1995; Bach et al., 1998], endothelial cell motility could be up-regulated synergistically by VEGF and fibrin in PRP-treated HUVECs. Of further interest is that the decrement of up-regulated cell motility is preceded by the peak interaction of VEGFR2 and VE-cadherin, a receptor of fibrin. Therefore, we believe that, in addition to the activation process, it is necessary to take PRP-induced desensitization of target endothelial cells into account when the controlled release system of PRP components is designed.
Materials and Methods PRP Preparation
As previously described [Okuda et al. 2003], blood was collected from three healthy and non-smoking volunteers aged 28, 30, and 54 years (two females; one male), and PRP was prepared using the two-step centrifugation protocol. The number of platelets in freshly prepared PRP samples was determined using an automated hematology analyzer (PocH 100iV diff: Sysmex, Kobe, Japan). Then, PRP preparations were frozen and stored at 220 C until further use (usually within 2 weeks). The concentrations of VEGF in frozen PRP samples were determined using a Human VEGF Quantikine ELISA Kit (R&D Systems, Minneapolis, MN). The correlation between platelet count and VEGF concentrations CYTOSKELETON
Human recombinant VEGF and mouse monoclonal antihuman VEGF antibodies (16F1) were obtained from Life Technologies (Carlsbad, CA, USA) and Immuno-biological Laboratories (Fujioka, Japan), respectively. VEGF was diluted to a final concentration of 200 ng mL21 in medium or Hank’s balanced salt solution (HBSS) (Life Technologies). For VEGF neutralization, 3 lg mL21 anti-VEGF antibody, which is expected to block the action of VEGF (50 ng mL21) [Suzuki et al., 1999], was added to the medium or HBSS containing the indicated concentrations of PRP and incubated for 30 min at room temperature prior to experiments. As an isotype control, mouse IgG2a (Medical & Biological Laboratories, Nagoya, Japan) was used at a final concentration of 3 mg mL21. Endothelial Cell Cultures and In Vitro Wound Healing Assay
HUVECs were obtained from AllCells, LLC (Emeryville, CA) and maintained in endothelial cell medium supplemented with growth factors (HuMedia-EB2; KURABO, Osaka, Japan) [Kobayashi et al., 2012]. For the wound healing assay, HUVECs were seeded into double-well culture inserts (Model 80209; ibidi GmbH, Munich, Germany) on a 35-mm dish (Falcon 353001, Corning Incorporated, Corning, NY) at a density of 2.4–3.0 3 102 cells mm22. The time course of this experiment is shown in Fig. 1A. Briefly, after 6–8 h of incubation in a CO2 incubator (5% CO2), the culture inserts were removed and the cells were labeled with 1 lM CellTrackerTM Orange CMTMR dye (Life Technologies) for 15 min. The cells were then washed with medium and incubated in medium containing PRP or VEGF for 30 min in a CO2 incubator (5% CO2). This time period is important for the conversion of fibrinogen to fibrin. Because fibrin is insoluble and often interferes with optical examination of the sample, the culture dishes were gently shaken two or three times during this period. The culture dishes were then set on the stage incubation chamber, which is connected to a CO2, humidity and temperature controller (5% CO2) (UNO; Okolab S.r.l., Naples, Italy), and the time-lapse recording was started. The time-lapse recording system consisted of a monochrome CCD camera (QIClick; Nippon Roper, Tokyo, Japan), LED lamp (TLED Plus, Sutter Instrument, Novato, CA) and personal desktop computer (Endeavour AT992E;
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EPSON, Suwa, Japan) with imaging software (VisiView; Visitron Systems GmbH, Puchheim Germany). The fluorescent images were obtained at an interval of 10 min for a 10 h period. The data were saved as an avi file and analyzed using the particle-tracking function of ImagePro Plus software (Nippon Roper). Immunofluorescence Examination
HUVECs were seeded onto glass coverslips and precultured for 2 days to form sub-confluent cultures. The cells were washed once with warmed HBSS and treated with PRP or VEGF. At the indicated time points, the cells were fixed with 10% formalin for 15 min and washed three times with PBS containing 0.5% Tween 20 (T-PBS) as previously described [Kawase et al., 2014]. After blocking with diluted Block Ace (1:2 in washing buffer; DS Pharma, Osaka, Japan) for 60 min at room temperature, the cells were probed with a rabbit polyclonal anti-paxillin antibody (1:200) (Transduction Laboratories, Lexington, KY) or a rabbit polyclonal phospho-VEGF-R2 (Tyr996) antibody (1:100) (Cell Signaling Technology Japan, Tokyo, Japan) overnight at 4 C. When incubating with rabbit monoclonal anti-VEGF-R2 antibody (D5B1) (1:100) (Cell Signaling Technology) and mouse monoclonal anti-CD144 (VEcadherin) (1:10) (BioLegend, San Diego, CA) overnight at 4 C, fixed cells were neither permeabilized with T-PBS nor blocked with the diluted Block Ace. Instead, cells were washed with PBS and blocked with 2.5% normal horse serum (Vector Labs, Burlingame, CA). The cells were then washed three times with T-PBS or PBS and probed with AlexaFlour 488-conjugated anti-rabbit IgG, AlexaFlour 555-conjugated anti-mouse IgG or AlexaFlour 555-conjugated anti-rabbit IgG, (Abcam, Cambridge, MA) diluted in Immunoshot (Mild) (Cosmo Bio, Tokyo, Japan) for 30 min at room temperature. Simultaneously, the polymerized F-actin fibers were stained with 0.1 lM Acti-StainTM 555 fluorescent phalloidin (Alexa Fluor 555-conjugated phalloidin) (Cytoskeleton, Denver, CO) in the same solution. After three washes with T-PBS, the cells were examined under a fluorescence microscope (Nikon, Tokyo, Japan). Isotype controls for rabbit primary antibody and mouse primary antibody (Life Technologies) were used as negative controls. Western Blotting Analysis
HUVECs were seeded onto 6-well plates at a density of 1 3 104 cells/well and pre-cultured for 2 days to form subconfluent cultures. After washing as shown in Fig. 1B, the cells were treated with PRP or VEGF in HBSS in a CO2 incubator (5% CO2) for the indicated periods of time. After washing twice with ice cold PBS, the cells were lysed with Laemmli sample buffer as previously described [Uematsu et al., 2013]. Protein samples were fractionated using 10%
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SDS-PAGE (ATTO, Tokyo, Japan) and electro-blotted onto PVDF membranes using the Trans-BlotV TurboTM Transfer System (Bio-Rad Laboratories, Hercules, CA). After blocking with diluted Block A (1:2) or 5% BSA (Fraction V, Sigma, St. Louis, MO) in T-TBS for 4–5 h at 4 C, the membranes were probed with the following primary antibodies: rabbit polyclonal anti-phospho-VEGFR2 (Y996; 1:2000 in dilution; Cell Signaling Technology, Danvers, MA), rabbit polyclonal anti-VEGFR2 (D5B1; 1:2000 in dilution; Cell Signaling Technology) or anti-actin antibody (1:1000 dilution; Santa Cruz Biotechnology, Santa Cruz, CA) overnight at 4 C. After washing three times with T-TBS, the membranes were probed with horseradish peroxidase-conjugated goat polyclonal anti-rabbit IgG H&L (1:5000 in dilution; Abcam, Cambridge, MA) or horseradish peroxidase-conjugated donkey anti-goat IgG (Santa Cruz Biotechnology) for 45 min at 4 C. After washing, images were visualized using ClarityTM Western ECL Substrate (Bio-Rad) and imaged using a cooled CCD camera system (Image Capture; ATTO, Tokyo, Japan). R
Statistical Analysis
The data were reported as the mean value 6 standard deviation (S.D.). For multi-group comparisons, statistical analyses were performed to compare the mean values using oneway analysis of variance (ANOVA) followed by Dunn’s multiple comparison test (SigmaPlot 12.5; Systat Software, San Jose, CA). P values < 0.05 were considered significant.
Acknowledgments
The authors thank Dr. Mito Kobayashi (Niigata University) for her technical assistance in determining the number of platelets and the concentrations of VEGF. This project was financially supported by a Grant-in-Aid for New Market Development by the Industrial Creation for 2012 from the Niigata Industrial Creation Organization and JSPS KAKENHI (Grant #24390443 and #24390465). References Akhundov K, Pietramaggiori G, Waselle L, Darwiche S, Guerid S, Scaletta C, Hirt-Burri N, Applegate LA, Raffoul WV. 2012. Development of a cost-effective method for platelet-rich plasma (PRP) preparation for topical wound healing. Ann Burns Fire Disasters 25:207–213. Albanese A, Licata ME, Polizzi B, Campisi G. 2013. Platelet-rich plasma (PRP) in dental and oral surgery: From the wound healing to bone regeneration. Immun Ageing 10:23. Bach TL, Barsigian C, Yaen CH, Martinez J. 1998. Endothelial cell VE-cadherin functions as a receptor for the beta15-42 sequence of fibrin. J Biol Chem 273:30719–30728. Bruns AF, Herbert SP, Odell AF, Jopling HM, Hooper NM, Zachary IC, Walker JH, Ponnambalam S. 2010. Ligand-stimulated vegfr2 signaling is regulated by co-ordinated trafficking and proteolysis. Traffic 11:161–174.
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Quantitative Analysis of PRP-induced Endothelial Cell Migration
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