Microvesicles (MVs) represent a subset of extracellular vesicles secreted by blebbing and shedding from plasma membranes. They are thought to play a significant role in transcellular signal transduction (1, 2).
MVs contain diverse signaling factors (e.g., miRNAs, surface molecules, and proteins) and are of major importance in cell-to-cell communication (2). Different stimuli, such as cytokines, bacterial lipopolysaccharides, reactive oxygen species, thrombin, and C-reactive protein, result in significant changes in both the secretion patterns and contents of MVs (3). These alterations can result in a fundamental change in MV-related signaling transduction (4). Moreover, MVs selectively bind to cells via specific surface molecules rather than nonspecifically interacting with various cell types (2, 5).
Due to the diversity of cell types (e.g., platelets, macrophages, endothelial cells, osteoblasts, and osteoclasts) involved in the complex process of fracture healing, synergistic interactions between these cells are essential. MVs are thought to play an important role in intercellular communication of the abovementioned cells (6–8). For example, in vitro studies demonstrated that MVs originating from either endothelial cells or mesenchymal stem cells (MSCs) had the potential to stimulate angiogenesis (9, 10). In addition, studies showed that MVs secreted by osteoblasts contained the receptor activator of the nuclear factor-kappa B (NF-κB) ligand (RANKL), which is known to be involved in the regulation of osteoclast function (11). Furthermore, isolated MVs from bone marrow were shown to have the potential to stimulate the differentiation of osteoblasts (12).
Despite the aforementioned findings, knowledge about the role of MVs during fracture healing remains sparse, especially the effects of systemically derived MVs on fracture healing. Therefore, in this study, we aimed to characterize the pattern of systemic release of MVs after induction of a femoral fracture and throughout the subsequent healing process. Furthermore, whether these MVs influence function of osteoblasts was investigated.
MATERIALS AND METHODS
This study was approved by the North Rhine-Westphalian State Agency for Nature, Environment and Consumer Protection (Reg. no.: 84-02.04.2015.A078). All the experiments were performed in accordance with “Guide for the Care and Use of Laboratory Animals” by the National Research Council (US) Committee for the Update of the Guide for the Care and Use of Laboratory Animals. In total, 24 female Sprague Dawley (SD) rats aged 8 to 10 weeks were used. Due to the protective effects of female hormones in cases of inflammatory stimuli, all the animals were in the same phase of the estrous cycle as determined by vaginal swabs. An overview of the study protocol is provided in Figure 1.
The rats were randomly allocated into a control group and three treatment groups (groups A, B, and C; 6 animals/group). The animals in group A were killed 3 days postfracture, whereas those in groups B and C were killed 1 and 2 weeks postfracture, respectively.
Animal fracture model
The femur fracture model has been previously described (13) and was applied with minor changes. Briefly, the rats were anesthetized with an intraperitoneal (i.p.) injection of ketamine 100 mg/kg i.p. (Pfizer, New York, NY) and xylazine 2% 10 mg/kg i.p. (Xylapan, Vetoquinol, Ravensbrug, Germany) before operation. Analgesia was induced with an additional subcutaneous (s.c.) application of buprenorphine hydrochloride (0.03–0.05 mg/kg) (Reckitt Benckiser Healthcare Ltd., UK). Afterward, isoflurane (2%–2.5% by volume) was also given consistently via a nasal mask during operation. The adequate anesthesia was assured by toe pinch reflex test. After anesthesia, a Kirschner wire (1.4 × 26 mm) was inserted intramedullary in the right hind limb femur in treatment groups A, B, and C. A fracture of the femoral shaft was then induced by a 1 kg plumb-cuboid blunt guillotine, which was dropped from a height of 20 cm. Fracture induction and the correct position of the Kirschner wire were confirmed by X-rays (Fig. 2A). Buprenorphine hydrochloride (0.03 mg/kg s.c. every 6–8 h for 48 h, subsequently every 12 h for 3 weeks) and Meloxicam (Metacam; Ingelheim, Germany, Boehringer Ingelheim GmbH) (1 mL/300 mL into drinking water for 1 week) were given after fracture induction.
At the end of the study period, the animals were killed by cardiac puncture under adequate anesthesia and analgesia. Blood was collected in ethylenediamine tetraacetic acid embedded tubes (SARSTEDT, Nuembrecht, Germany) and centrifuged at 1,200 g for 5 min at 4°C. The plasma was then stored at −80°C.
MV counts and sizes
Analysis of the concentrations and sizes of MVs in plasma was performed by nanoparticle tracking analysis (NanoSight 300; NTA, NanoSight Ltd., Amesbury, UK). For the analysis, 100 μL of plasma was centrifuged at 5,000 g for 20 min to remove platelets and apoptotic bodies. The plasma was then diluted to 200 μL with phosphate-buffered saline (PBS), which was filtered through a 0.2-μm sterile syringe filter (Corning, NY). Samples were then analyzed by NanoSight 300 in accordance with the manufacturer's instructions.
MV isolation and MV-free plasma generation
MV isolation was performed according to a previously reported method (14). Briefly, the plasma obtained from each animal was centrifuged at 5,000 g for 20 min. The supernatant was then centrifuged again at 17,000 g for 90 min. The supernatant after second centrifuge was collected as MV-free plasma. The pellet (including the MVs) was resuspended in PBS. The MVs were washed a second time, centrifuged at 17,000 g for 90 min, and dissolved in PBS.
Transmission electron microscopy of MVs
Unfixed isolated MVs were allowed to adsorb for 10 min on formvar–carbon-coated nickel grids (200 mesh), which was glow discharged for 2 min (Maxtaform; Plano, Wetzlar, Germany). After washing twice with Aqua Dest (Science Services GmbH, Munich, Germany), the samples on grids were stained using a drop of 0.5% uranyl acetate in Aqua Dest. After air-drying, the samples were examined using a LEO 906 (Carl Zeiss, Oberkochen, Germany) transmission electron microscope, operating at an acceleration voltage of 60 kV.
Osteoblast isolation, culture, and identification
Osteoblast isolation (rat cranial bone) and culture were performed from a neonatal SD rat according to a previously described protocol (15). The culture medium (CM) was changed every 3 days until the cells covered around 80% of the plate. After trypsinization (Gibco; Thermo Scientific), the cells were collected and seeded into culture flasks (Culturestar, Greine rbio-one GmbH, Frickenhausen, Germany). Third-passage cells were used for further analyses.
To identify osteoblasts, a bone formation assay was performed (Fig. 2B). Isolated osteoblasts were cultured on a six-well plate with 2 mM β-GP (β-glycerol phosphate; Sigma Aldrich, St. Louis, Mo), 10 nM dexamethasone (Merck Serono GmbH, Darmstadt, Germany), and 50 μg/mL of ascorbate (Sigma Aldrich). The pH was adjusted to 7.4, and 50% of the CM was changed every third day. After 14 days, the cells were washed with PBS 2 more times, fixed in formaldehyde for 15 min, and washed twice with PBS. After air-drying, the cells were stained with Alizarin Red S (Sigma Aldrich) for 5 min and washed with 50% ethanol (AppliChem, Darmstadt, Germany) 3 times.
Imaging of fluorescently labeled MVs
The MVs and the same volume of PBS as a control were incubated with WGA-Alexa Fluor 594 solution (0.5 μg/mL Wheat Germ Agglutinin, Alexa Fluor 594 conjugate; Thermo Fisher, Waltham, Mass) for 10 min, washed twice with DMEM (Dulbecco's Modified Eagle Medium; Thermo Fisher) and after centrifugation at 17,000 g resuspended in DMEM (Dulbecco's Modified Eagle Medium; Thermo Fisher). The osteoblasts were cultured in a 24-well plate on sterilized cover slips. The cells were incubated with the stained MVs for 10 min, 30 min, and 1 h. The control plate was cultured for 1 h. Subsequently, the cover slips were washed with PBS, liquid was removed with blotting paper, and mounting medium (SlowFade Diamond Antifade Mountant with DAPI (4’,6-diamidino-2-phenylindole); Thermo Fisher) was used to mount the cells and stain the nuclei. The samples were analyzed by confocal microscopy (LSM 710; Carl Zeiss Microscopy GmbH, Jena, Germany). A Z-stack was generated, and a 3D image reconstructed using ZEN 2.3 software (Blue edition; Carl Zeiss Microscopy GmbH) to identify intracellular MVs.
Transforming growth factor-β1 and insulin-like growth factor 1 analysis in MVs and MV-free plasma
Commercialized enzyme-linked immunosorbent assay kits were used to determine the concentration of transforming growth factor-β1 (TGF-β1) (Boster Biological Technology, Pleasanton, Calif) and insulin-like growth factor 1 (IGF-1) (Cloud Clone Corp., Wuhan, China). MVs were isolated from 100 μL plasma and lysed with 100 μL peqGOLD TriFast (peqlab; VWR International GmbH, Radnor, Pa). The MV solution and MV-free plasma was then analyzed according to the protocol.
Four kinds of CMs were used, and for the following assays four kinds of CM were prepared:
- Normal CM was prepared with DMEM, 100 U/mL of penicillin, 50 μg/mL of streptomycin sulfate, and 10% fetal bovine serum (FBS; Thermo Fisher).
- Normal CM containing MVs isolated from plasma resuspended in a comparable amount of FBS (MV-contained culture medium, MVCM) was prepared with DMEM, 100 U/mL of penicillin, 50 μg/mL of streptomycin sulfate, and 10% FBS and MV.
- CM containing plasma from groups A, B, C, or the control (plasma-contained culture medium, PCM) was prepared with DMEM, 100 U/mL of penicillin, 50 μg/mL of streptomycin sulfate, 5% plasma from the animals from groups A, B, and C, and 5% FBS.
- MV-free plasma CM was prepared with DMEM, 100 U/mL of penicillin, 50 μg/mL of streptomycin sulfate, 5% FBS, and 5% MV-free plasma from the animals in groups A, B, and C.
Osteoblasts (5,000 per well) were seeded in 24-well plates (TPP, Trasadingen, Switzerland) and cultured with 500 μL of PCM. For each test, triple wells were used as repeats. The medium was changed every third day. After 3, 5, and 8 days, the cells were digested with trypsin, and the total number of cells was counted with Neubauer chamber.
3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay
An MTT assay was performed to evaluate the viability and proliferation of the osteoblasts (16). For the assay, 10,000 osteoblasts per well were seeded in 96-well plates and cultured in 200 μL of four kinds of CMs (1 to 4, see Experimental CM part) for 4 days. Each CM experiment was repeated 3 times. On the fourth day, 20 μL of 5 mg/mL of MTT (Sigma Aldrich) were added into each well and incubated for 4 h. The supernatant was then extracted, and 100 μL of dimethyl sulfoxide (Sigma Aldrich) was added in each well and gently shaken in the dark for 10 min. The optical density value was measured with wavelengths of 490 nm and 690 nm (for correction).
Quantitative real-time polymerase chain reaction
The osteoblasts were cultured with four kinds of CMs (1. to 4.) for 5 days and then dissolved with peqGOLD TriFast. Total RNA was extracted, and cDNA was synthesized with cDNA kits (Thermo Scientific). The quantitative real-time polymerase chain reaction (qRT-PCR) was performed with SYBR Green real-time PCR Master Mix Reagent (Thermo Scientific) on a StepOne Real-Time PCR System (Applied Biosystems, Waltham, Mass). The expression of runt-related transcription factor 2 (RUNX2), osteocalcin, and collagen 1A were analyzed. Peptidylprolyl isomerase A (PPIA) was used as the housekeeping gene. The sequences of the primers are listed in Table 1. The analysis of gene expression was performed using the 2−ΔΔCT method (%PPIA) (17).
Excel 2016 (Microsoft, Redmond, Wash) and Graphpad Prism 7.0 (GraphPad Software, La Jolla, Calif) were used for data collection and statistical analysis. The Kolmogorov–Smirnov test was applied to test for a normal distribution. An analysis of variance, Student t test, and Kruskal–Wallis test were performed to test significance, and P < 0.05 was accepted as denoting statistical significance.
To obtain plasma from the trauma model, three groups of animals underwent femur fracture and fixation of right hind limbs under anesthesia. All the animals survived until the end of the group-specific study period. Correct fixation by intramedullary pins was verified by an X-ray. Osteosynthesis resulted in appropriate stabilization of the femoral fracture (Fig. 2A). No sign of infections or other complications were observed. Animals were killed after 3 days, 1 week, and 2 weeks, and blood plasma was collected through heart puncture.
Identification of MVs
To identify MVs from the animal model, TEM was used. Figure 3, A and B shows representative TEM images of MVs. Nanoparticle tracking analysis was used to assess size and amount of MVs. Figure 3C shows a typical presentation of MVs obtained by nanoparticle tracking analysis (from 2 weeks group). The average size of the MVs was 185.1±4.7 nm. A typical size distribution is shown in Figure 3D. There was no significant difference in the MV concentrations of the control and study groups (Fig. 3E).
MV uptake of osteoblasts
To verify that MVs from the trauma model could be taken up by osteoblasts, fluorescently labelled MVs were cocultured with primary cranial osteoblasts. Condensed red fluorescence was observed inside the osteoblasts indicating incorporation of MVs (Fig. 4A). The amount of incorporated MVs increased with the time of incubation. No intracellular red fluorescence was observed in the control group without labeled MVs. The 3D reconstruction of confocal images (Fig. 4B) showed that the MVs were located around the nuclei. These results indicated that the cultured osteoblasts were able to incorporate MVs generated under trauma conditions located in the plasma.
Concentrations of TGF-β1 and IGF-1 in MVs and MV-free plasma
To locate and quantify osteogenic cytokines in the plasma, concentrations of TGF-β1 and IGF-1 were measured with ELISA kits. Compared with the control group, TGF-β1 and IGF-1 concentrations in MV-free plasma were significantly lower in group A (TGF-β1, P = 0.0320; IGF-1, P = 0.0211). The levels of TGF-β1 and IGF-1 within MVs were not significantly different between the control and study groups. The levels of TGF-β1 and IGF-1 were significantly higher in MV-free plasma than in MVs (Fig. 5), which proved that TGF-β1 and IGF-1 were mostly located within the MV-free plasma rather than in MVs.
Growth and differentiation of osteoblasts cultured in PCM
To assess the influence of full trauma plasma on proliferation and differentiation of primary osteoblasts, osteoblasts were cultured in PCM, made by plasma from study animals. A growth assay was performed, and mRNA expression of osteogenic genes was analyzed. As shown in Figure 6, plasma from groups B and C at day 5 and 8 after incubation, respectively, significantly stimulated the growth of osteoblasts comparing with the control group (day 5, P = 0.0318; day 8, P = 0.009). PCM was associated with an increase in osteoblast viability over time, with significantly higher viability in group C as compared with that in the control group (P = 0.0393; Fig. 7A). As previously reported, RUNX 2 and collagen 1A synthesis served as markers of early-phase osteogenesis, and osteocalcin synthesis as late-phase osteoblastic differentiation markers (18). Osteocalcin mRNA expression of osteoblasts was reduced in the early posttrauma phase as compared with that in the controls (group B: P
= 0.0172; Fig. 7C), with an increase in group C (B vs. C, P = 0.0021). Comparisons of collagen 1A and RUNX2 revealed no significant differences in any of the groups (Figs. 7, B and D). These results indicated that plasma from traumatized animals influences growth and differentiation of the primary osteoblasts.
Viability/proliferation and differentiation of osteoblasts cultured in MVCM
The functions of MVs from trauma plasma on primarily cultured osteoblasts were assessed. MVs from trauma and control plasma were isolated, and cocultured with the primary osteoblasts in CM made from FBS (MVCM). As depicted in Figure 8A, stimulation with MVs resulted in a significant increase in osteoblast viability in all the study groups (A, P = 0.0276; B, P = 0.0295; C, P = 0.0407). Osteocalcin was lower in group C as compared with that in the control group (P = 0.0395; Fig. 8C). There were also decreased osteocalcin expression levels in groups A and B (P (A) = 0.0686, P (B) = 0.0708); however, they did not show statistical significance. Analysis of RUNX 2 and collagen 1A revealed no significant differences between the control and study groups (Fig. 8, B and D). These results showed the ability of MVs from trauma plasma to influence viability and differentiation of osteoblasts.
Viability/proliferation and differentiation of osteoblasts cultured in MV-free plasma CM
To further distinguish the function of MVs and soluble components of the trauma plasma, an MTT assay and mRNA expression of osteogenic genes were also analyzed on primary osteoblasts cultured with MV-free plasma from study animals. MV-free plasma from group A significantly stimulated viability of primary osteoblasts (P = 0.0171; Fig. 9A), as compared with the control group. However, no significant differences in viability of primary osteoblasts were found between all the other groups. Osteocalcin expression was significantly higher in group C as compared with that in the control (P = 0.0454). No differences were detected in RUNX2 and collagen 1A expression (Figs. 9, B–D). Comparing MVs in the trauma plasma, MV-free plasma stimulated viability and differentiation of the primary osteoblasts at certain time points.
Successful fracture healing involves a complex interaction between a variety of cellular components and signaling molecules (1). Dysfunction of any of these cellular components or disturbances of signaling mechanisms affects fracture healing. In this study, we aimed to investigate the interaction of systemic MVs from a rat fracture model with osteoblasts. Our main results can be summarized as follows:
- MVs isolated after a femoral fracture were time-dependently incorporated in osteoblasts and concentrated around the nucleus.
- The stimulatory effect of trauma plasma on osteoblast proliferation depends on the postfracture time.
- MVs from trauma plasma increased the viability of osteoblasts, particularly in the late phase (i.e., 2 weeks postfracture) after a femoral fracture.
- Late-phase differentiation of osteoblasts was not stimulated by the MVs in a high extent. In contrast, MV-free plasma seemed to have a stimulatory effect on differentiation in the late phase (2 weeks after fracture).
Previous studies described increased osteoblast growth 2 weeks after a fracture (19, 20). In accordance with this finding, in a rat model, we found enhanced growth of osteoblasts stimulated with plasma from 1 and 2 weeks after fracture. The stimulatory effect increased with the time elapsed since trauma. Therefore, plasma, specifically plasma mediators, has the potential to modulate fracture healing. Among these mediators, studies reported that TGF-β and IGF-I represented important growth factors for osteoblasts (21). TGF-β and IGF-1 were secreted after different inflammatory stimuli and seemed to be good indicators of the fracture-healing process (21). In our study, both TGF-β1 and IGF-1 in MV-free plasma decreased in the first week postfracture. These findings are consistent with those of previous publications (21, 22) and therefore indicate the reliability of our model. To the best of our knowledge, no other studies have described the characteristics and kinetics of TGF-β1 and IGF-1 concentrations in MVs after fractures. We found significantly lower levels of TGF-β1 and IGF-1 in MVs compared with MV-free plasma, with no relevant changes in intravesicular TGF-β1 and IGF-1 levels over the entire observation period.
MVs originate from a variety of cells and indicate the functional status of an organism at a specific time point. In previous studies, MVs were detected in the systemic circulation after diverse insults (e.g., severe trauma, stress) (23–25). Previous studies provided evidence that MVs seemed to be of major importance for intercellular communication to regulate bone metabolism (11, 12, 26, 27). In this context, osteoblastic MVs were found to be important for cell–cell communication of osteoblasts with both osteoclasts (11) and MSCs (27); these MVs were able to incorporate into cells (e.g., integration of MV-derived osteoclasts into osteoblasts) (28–30). These findings are in line with those obtained in our study, which revealed the integration of plasma-derived MVs into osteoblasts and their aggregation around the nuclei. Based on the observed accumulation, it can be postulated that MV integration is faster than intracellular degradation of MVs. These results could explain both the potential impact of MVs on cellular function and the importance of MVs as mediators of intracellular communication.
As indicated by the results of the MTT assay in our study, plasma-derived MVs seemed to have the potential to stimulate the viability and proliferation of osteoblasts, particularly in the late phase (i.e., 2 weeks) after fractures, whereas MV-independent mechanisms seemed to be more relevant in the early phase after trauma. Accordingly, others also found stimulatory effects of both bone marrow-derived MVs on osteoblast function (12) and effects of pluripotent stem cell-derived MVs on angiogenesis and osteogenesis (26).
Previous studies have provided evidence that incorporated extracellular vesicles (e.g., originating from bone marrow stem cells or prostate cancer cell lines) affect osteoblast differentiation (12). In this context, several studies demonstrated that osteoblast-derived MVs had the potential to stimulate the differentiation of stem cells into osteoblasts (31, 32). In contrast, we found that late-phase differentiation of osteoblasts, represented by osteocalcin synthesis (18), was almost exclusively stimulated by mediators located in plasma but not by plasma-derived MVs alone. These diverse findings might be explained by different factors. First, the low osteocalcin expression after plasma-derived MVs stimulation in our study might be associated with the stimulatory effects of these MVs on the viability and proliferation of osteoblasts. In this context, Owen et al. found low expression of osteocalcin mRNA during the proliferation phase of osteoblasts (33). Furthermore, studies observed that different factors (e.g., mediators and dynamic cell stretching) that promote the proliferation of osteoblasts simultaneously inhibited osteocalcin expression (34, 35). Second, it is possible that the different origins of the investigated MVs (plasma vs. bone marrow stem cells) result in different effects on osteocalcin expression. Therefore, additional studies are needed to investigate the characteristics (e.g., content and receptor status) of MVs originating from diverse cell types.
As determined by the analysis of RUNX 2 and collagen 1A synthesis (early-phase osteoblastic differentiation markers) (18), we observed no significant differences in the early-phase differentiation of osteoblasts among the control and experimental groups following MVs and MV-free plasma stimulation. First, these findings may point to a comparable effect of both MVs and plasma on early osteoblastic differentiation. Second, as the synthesis of RUNX 2 and collagen 1A in the trauma groups were not significantly different to that in the controls, the findings may indicate that the primary osteoblasts from this protocol are in an early differentiation phase, with a stable expression of RUNX2 and collagen 1A.
First, as our findings were derived from an animal model, they are not directly transferable to humans. However, the results from this study showed a potential role of osteoblast modulation by MVs, which may help in modulating osteogenesis after fracture or orthopaedic operations. Further in vivo and translational studies will be conducted in our laboratory in the future. Second, the amount of plasma harvested from this rat trauma model was limited, whereas the consumption of the cell culture and tests was high. This limited the possibility of further analyses, such as MV phenotype, miRNA analysis, and in vivo studies. Thus, additional studies are needed to further elucidate the role of MVs in osteoblastic function postfracture. In the following studies, larger animals may be needed in studies to increase both the availability of material for analysis, as well as the translational relevance. Finally, the centrifuge protocol for MV isolation does not exclude the presence of small amounts of other components (e.g., exosomes) that might have an impact on osteoblast functions (14).
The fracture healing process involves a complex network of signal transduction between a variety of cells. Our study showed a potential effect of MVs on regulating fracture healing, by modulating the viability and proliferation of osteoblasts. Therefore, MVs may possibly have potential therapeutic uses in cases of fracture healing disturbances. This should be clarified in further translational studies.
This work was supported by the Confocal Microscopy Facility of the Interdisciplinary Center for Clinical Research (IZKF) Aachen.
1. Qiao Z, Greven J, Horst K, Pfeifer R, Kobbe P, Pape HC, Hildebrand F. Fracture healing and the underexposed role of extracellular vesicle-based cross talk. Shock
49 (5):486–496, 2018.
2. Cocucci E, Racchetti G, Meldolesi J. Shedding microvesicles
: artefacts no more. Trends Cell Biol
19 (2):43–51, 2009.
3. Dignat-George F, Boulanger CM. The many faces of endothelial microparticles. Arterioscler Thromb Vasc Biol
31 (1):27–33, 2011.
4. Lawson C, Vicencio JM, Yellon DM, Davidson SM. Microvesicles
and exosomes: new players in metabolic and cardiovascular disease. J Endocrinol
228 (2):R57–R71, 2016.
5. Losche W, Scholz T, Temmler U, Oberle V, Claus RA. Platelet-derived microvesicles
transfer tissue factor to monocytes but not to neutrophils. Platelets
15 (2):109–115, 2004.
6. Laffont B, Corduan A, Rousseau M, Duchez AC, Lee CH, Boilard E, Provost P. Platelet microparticles reprogram macrophage gene expression and function. Thromb Haemost
115 (2):311–323, 2016.
7. Laffont B, Corduan A, Ple H, Duchez AC, Cloutier N, Boilard E, Provost P. Activated platelets can deliver mRNA regulatory Ago2*microRNA complexes to endothelial cells via microparticles. Blood
122 (2):253–261, 2013.
8. Dang PN, Dwivedi N, Phillips LM, Yu X, Herberg S, Bowerman C, Solorio LD, Murphy WL, Alsberg E. Controlled dual growth factor delivery from microparticles incorporated within human bone marrow-derived mesenchymal stem cell aggregates for enhanced bone tissue engineering via endochondral ossification. Stem Cells Transl Med
5 (2):206–217, 2016.
9. Zhang J, Ren J, Chen H, Geng Q. Inflammation induced-endothelial cells release angiogenesis associated-microRNAs into circulation by microparticles. Chin Med J (Engl)
127 (12):2212–2217, 2014.
10. Deregibus MC, Cantaluppi V, Calogero R, Lo Iacono M, Tetta C, Biancone L, Bruno S, Bussolati B, Camussi G. Endothelial progenitor cell derived microvesicles
activate an angiogenic program in endothelial cells by a horizontal transfer of mRNA. Blood
110 (7):2440–2448, 2007.
11. Deng L, Wang Y, Peng Y, Wu Y, Ding Y, Jiang Y, Shen Z, Fu Q. Osteoblast
: a novel mechanism for communication between osteoblasts and osteoclasts. Bone
12. Qin Y, Wang L, Gao Z, Chen G, Zhang C. Bone marrow stromal/stem cell-derived extracellular vesicles regulate osteoblast
activity and differentiation in vitro and promote bone regeneration in vivo. Sci Rep
13. Wang SJ, Lewallen DG, Bolander ME, Chao EY, Ilstrup DM, Greenleaf JF. Low intensity ultrasound treatment increases strength in a rat femoral fracture model. J Orthop Res
12 (1):40–47, 1994.
14. Dey-Hazra E, Hertel B, Kirsch T, Woywodt A, Lovric S, Haller H, Haubitz M, Erdbruegger U. Detection of circulating microparticles by flow cytometry: influence of centrifugation, filtration of buffer, and freezing. Vasc Health Risk Manag
15. Orriss IR, Taylor SE, Arnett TR. Rat osteoblast
cultures. Methods Mol Biol
16. Choi H, Srikanth S, Atti E, Pirih FQ, Nervina JM, Gwack Y, Tetradis S. Deletion of Orai1 leads to bone loss aggravated with aging and impairs function of osteoblast
lineage cells. Bone Rep
17. Livak KJ, Schmittgen TD. Analysis of relative gene expression data using real-time quantitative PCR and the 2−ΔΔCT method. methods
25 (4):402–408, 2001.
18. Tang Q, Tong M, Zheng G, Shen L, Shang P, Liu H. Masquelet's induced membrane promotes the osteogenic differentiation of bone marrow mesenchymal stem cells by activating the Smad and MAPK pathways. Am J Transl Res
10 (4):1211–1219, 2018.
19. Iwaki A, Jingushi S, Oda Y, Izumi T, Shida JI, Tsuneyoshi M, Sugioka Y. Localization and quantification of proliferating cells during rat fracture repair: detection of proliferating cell nuclear antigen by immunohistochemistry. J Bone Miner Res
12 (1):96–102, 1997.
20. Wildemann B, Schmidmaier G, Ordel S, Stange R, Haas NP, Raschke M. Cell proliferation and differentiation during fracture healing are influenced by locally applied IGF-I and TGF-beta1: comparison of two proliferation markers, PCNA and BrdU. J Biomed Mater Res B Appl Biomater
65 (1):150–156, 2003.
21. Fischer C, Doll J, Tanner M, Bruckner T, Zimmermann G, Helbig L, Biglari B, Schmidmaier G, Moghaddam A. Quantification of TGF-ss1, PDGF and IGF-1 cytokine expression after fracture treatment vs. non-union therapy via masquelet. Injury
47 (2):342–349, 2016.
22. Chen QQ, Wang WM. Expression of FGF-2 and IGF-1 in diabetic rats with fracture. Asian Pac J Trop Med
7 (1):71–75, 2014.
23. Matijevic N, Wang YW, Holcomb JB, Kozar R, Cardenas JC, Wade CE. Microvesicle phenotypes are associated with transfusion requirements and mortality in subjects with severe injuries. J Extracell Vesicles
24. Curry N, Raja A, Beavis J, Stanworth S, Harrison P. Levels of procoagulant microvesicles
are elevated after traumatic injury and platelet microvesicles
are negatively correlated with mortality. J Extracell Vesicles
25. Fleshner M, Crane CR. Exosomes, DAMPs and miRNA: features of stress physiology and immune homeostasis. Trends Immunol
38 (10):768–776, 2017.
26. Qi X, Zhang J, Yuan H, Xu Z, Li Q, Niu X, Hu B, Wang Y, Li X. Exosomes secreted by human-induced pluripotent stem cell-derived mesenchymal stem cells repair critical-sized bone defects through enhanced angiogenesis and osteogenesis in osteoporotic rats. Int J Biol Sci
12 (7):836–849, 2016.
27. Xu JF, Yang GH, Pan XH, Zhang SJ, Zhao C, Qiu BS, Gu HF, Hong JF, Cao L, Chen Y, et al. Altered microRNA expression profile in exosomes during osteogenic differentiation of human bone marrow-derived mesenchymal stem cells. PLoS One
9 (12):e114627, 2014.
28. Sun W, Zhao C, Li Y, Wang L, Nie G, Peng J, Wang A, Zhang P, Tian W, Li Q, et al. Osteoclast-derived microRNA-containing exosomes selectively inhibit osteoblast
activity. Cell Discov
29. Li D, Liu J, Guo B, Liang C, Dang L, Lu C, He X, Cheung HY, Xu L, Lu C, et al. Osteoclast-derived exosomal miR-214-3p inhibits osteoblastic bone formation. Nat Commun
30. Ye Y, Li SL, Ma YY, Diao YJ, Yang L, Su MQ, Li Z, Ji Y, Wang J, Lei L, et al. Exosomal miR-141-3p regulates osteoblast
activity to promote the osteoblastic metastasis of prostate cancer. Oncotarget
8 (55):94834–94849, 2017.
31. Nair R, Santos L, Awasthi S, von Erlach T, Chow LW, Bertazzo S, Stevens MM. Extracellular vesicles derived from preosteoblasts influence embryonic stem cell differentiation. Stem Cells Dev
23 (14):1625–1635, 2014.
32. Cui Y, Luan J, Li H, Zhou X, Han J. Exosomes derived from mineralizing osteoblasts promote ST2 cell osteogenic differentiation by alteration of microRNA expression. FEBS Lett
590 (1):185–192, 2016.
33. Owen TA, Aronow M, Shalhoub V, Barone LM, Wilming L, Tassinari MS, Kennedy MB, Pockwinse S, Lian JB, Stein GS. Progressive development of the rat osteoblast
phenotype in vitro: reciprocal relationships in expression of genes associated with osteoblast
proliferation and differentiation during formation of the bone extracellular matrix. J Cell Physiol
143 (3):420–430, 1990.
34. Viereck V, Siggelkow H, Tauber S, Raddatz D, Schutze N, Hufner M. Differential regulation of Cbfa1/Runx2 and osteocalcin gene expression by vitamin-D3, dexamethasone, and local growth factors in primary human osteoblasts. J Cell Biochem
86 (2):348–356, 2002.
35. Kaspar D, Seidl W, Neidlinger-Wilke C, Ignatius A, Claes L. Dynamic cell stretching increases human osteoblast
proliferation and CICP synthesis but decreases osteocalcin synthesis and alkaline phosphatase activity. J Biomech
33 (1):45–51, 2000.