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Keywords:

  • BONE REPAIR;
  • HETEROTOPIC OSSIFICATION;
  • PLASMINOGEN;
  • ANGIOGENESIS;
  • MACROPHAGE

ABSTRACT

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

The further development in research of bone regeneration is necessary to meet the clinical demand for bone reconstruction. Plasminogen is a critical factor of the tissue fibrinolytic system, which mediates tissue repair in the skin and liver. However, the role of the fibrinolytic system in bone regeneration remains unknown. Herein, we investigated bone repair and ectopic bone formation using plasminogen-deficient (Plg–/–) mice. Bone repair of the femur is delayed in Plg–/– mice, unlike that in the wild-type (Plg+/+) mice. The deposition of cartilage matrix and osteoblast formation were both decreased in Plg–/– mice. Vessel formation, macrophage accumulation, and the levels of vascular endothelial growth factor (VEGF) and transforming growth factor-β (TGF-β) were decreased at the site of bone damage in Plg–/– mice. Conversely, heterotopic ossification was not significantly different between Plg+/+ and Plg–/– mice. Moreover, angiogenesis, macrophage accumulation, and the levels of VEGF and TGF-β were comparable between Plg+/+ and Plg–/– mice in heterotopic ossification. Our data provide novel evidence that plasminogen is essential for bone repair. The present study indicates that plasminogen contributes to angiogenesis related to macrophage accumulation, TGF-β, and VEGF, thereby leading to the enhancement of bone repair.


Introduction

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

The regeneration of a bone defect remains a challenge.[1] Although an autogenous bone graft is the gold-standard procedure for the reconstruction of a bone defect, it has several disadvantages, including the limited size of the graft and injury to the donor site.[1] The clarification of the mechanism for bone repair and regeneration is, therefore, necessary to meet the clinical demand for bone reconstruction.

Bone is a highly vascularized tissue that is formed through intramembranous or endochondral ossification.[2] Endochondral ossification is observed in most bones, including the femur and tibia. During endochondral ossification, cartilage is formed from mesenchymal stem cells, and the cartilage is replaced by bone accompanied by the invasion of blood vessels. This vascular invasion is a critical process for endochondral ossification. Vascular endothelial growth factor (VEGF) is one of the important mediators of vascular formation and invasion during endochondral ossification,[3] and a previous study indicated that VEGF stimulates bone repair by promoting angiogenesis.4 VEGF also promotes osteogenesis by stimulating the migration and differentiation of osteoblasts.[5]

Plasminogen, a pivotal component of the fibrinolytic system, is an inactive proenzyme that is converted to the active serine protease plasmin by tissue-type plasminogen activator (t-PA) and urokinase-type PA (u-PA).[6] The fibrinolytic system has many physiological and pathophysiological functions in mammals beyond its proteolytic effect on thrombi. Plasmin activates the tissue proteolytic system.[7] In addition, plasmin modulates the release of growth factors, such as VEGF and transforming growth factor-β (TGF-β), through the degradation of the extracellular matrix (ECM).[8, 9]

The healing of skin wounds is impaired by plasminogen deficiency owing to the failure of ECM degradation.[10] Plasminogen deficiency also impairs liver remodeling after a toxic injury.[11] Recently, we reported that plasminogen is essential for the formation of granulation tissue during liver repair.[12] These findings indicate that the tissue fibrinolytic system is involved in the tissue-repair process of the skin and liver. In a previous study on plasminogen-deficient mice, we showed that bone metabolism is regulated by plasminogen/plasmin through the expression of osteoprotegerin, a decoy receptor for receptor activator of NF-κB (RANK) ligand, in osteoblastic cells and the increase in the number of osteoclasts.[13] Furthermore, the mineral density of trabecular and cortical bone in the tibia is decreased by plasminogen deficiency, as shown in that study.[13] However, whether plasminogen/plasmin plays some role in bone repair remains unknown.

In the present study, we investigated the role of plasminogen in ossification using mice with a plasminogen gene deficiency (Plg−/−) and their wild-type (Plg+/+) counterparts.

Materials and Methods

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Materials

Anti-Smad2, anti-phosphorylated Smad2/3, anti-bone morphogenetic protein (BMP)-2, and anti-hypoxia inducible factor (HIF)-1α antibodies were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Anti-Osterix, anti-CD31, anti-VEGF, and anti-TGF-β antibodies were obtained from Abcam (Cambridge, UK). Anti-F4/80, anti-Smad3, anti-alkaline phosphatase (ALP), and anti-β-actin antibodies were from AbD Serotec (Raleigh, NC, USA), Zymed Laboratories (San Francisco, CA, USA), Abnova (Taipei, Taiwan), and Cell Signaling Technology (Danvers, MA, USA), respectively. Human recombinant BMP-2 was provided by Pfizer Inc. (Groton, CT, USA).

Animals

Male and female mice with plasminogen gene deficiency (Plg−/−) and their wild-type counterparts (Plg+/+), each weighing 18 to 25 g and 7 to 10 weeks old, with a mixed C57BL/6J (75%) and 129/SvJ (25%) background, were used. These mice were kindly provided by Professor D Collen (University of Leuven, Leuven, Belgium). All experiments were performed according to the guidelines of the National Institutes of Health and the institutional rules for the use and care of laboratory animals at Kinki University.

Bone defect model

A bone defect was induced in the mice according to the method described previously[14] with some modifications. Briefly, under anesthesia induced by 2% isoflurane, the anterior skin over the mid-femur of the right leg was incised longitudinally for 5 mm in length. After splitting the muscle, the surface of femoral bone was exposed. Thereafter, a hole was made using a drill with a diameter of 0.9 mm. The hole was irrigated with saline to prevent thermal necrosis of the margins. The incised skin was then sutured in a sterile manner, and the anesthesia was discontinued. The body temperature was maintained at 37°C during surgery by the use of a heating pad.

Heterotopic ossification model

The heterotopic ossification was induced as described previously.[15] Briefly, a 5-µL aliquot containing 5 µg recombinant human BMP-2 was adsorbed onto a 5-mm × 5-mm-thick atelocollagen sponge (Koken Co., Tokyo, Japan). Under anesthesia induced using 2% isoflurane, the sponge was implanted into a pouch created in the quadriceps muscle of the mice. The body temperature was maintained at 37°C during surgery by using a heating pad.

In vivo quantitative computed tomography (qCT) analysis

The mice were anesthetized using 2% isoflurane, and the femur or BMP-2-induced heterotopic bone was scanned using an X-ray CT system (Latheta LCT-200; Hitachi Aloka Medical, Tokyo, Japan). Parameters used for the CT scans were as follows: tube voltage, 50 kVp; tube current, 500 µA; integration time, 3.6 ms; axial field of view, 48 mm, with an isotropic voxel size of 48 µm. Images were generated by integration of two and three signal averages for the femur and heterotopic bone, respectively. Total scan time was approximately 5 minutes for the femur and 9 minutes for the heterotopic bone. Volume-rendered 3-dimensional CT pictures were reconstructed using the VGStudio MAX2.1 software (Nihon Visual Science, Tokyo, Japan). The area of bone defect in the femur was quantified by using an image-processing program (ImageJ, http://rsbweb.nih.gov/ij/download.html). The mineral content of the heterotopic bone was calculated using Latheta software (Hitachi Aloka Medical). A threshold density of 160 mg/cm3 was selected to distinguish mineralized from unmineralized tissue. The density range was calibrated daily with a manufacturer-supplied phantom.

Histological analysis

The mice were anesthetized using pentobarbital sodium (50 mg/kg, intraperitoneally) on days 1, 4, 7, and 14 after surgery. The femur and heterotopic bone were removed, fixed in 4% paraformaldehyde, demineralized in 22.5% formic acid and 340 mM sodium citrate solution for 24 hours, and embedded in paraffin. Thereafter, 4-µm-thick sections were obtained. The sections were processed for hematoxylin and eosin (H&E), Alcian blue, and toluidine blue staining. The metachromatic area in the sections stained with toluidine blue was quantified using image-processing software (Mac SCOPE; Mitani Co., Fukui, Japan) in a blinded evaluation.

The sections were stained with tartrate-resistant acid phosphatase (TRAP) by using a TRAP staining kit (Wako Pure Chemical, Osaka, Japan). The number of TRAP-positive multinucleated cells on the bone surface was measured at the damaged site of the femur and heterotopic bone in a blinded evaluation.

Immunostaining was performed as described previously.[12, 16] Briefly, the sections were incubated with the anti-ALP antibody at a dilution of 1:100, anti-Osterix antibody at a dilution of 1:200, anti-CD31 antibody at a dilution of 1:100, anti-VEGF antibody at a dilution of 1:200, or anti-F4/80 antibody at a dilution of 1:1000 followed by incubation with the appropriate secondary antibody conjugated with horseradish peroxidase. Positive signals were visualized using a tyramide signal amplification system (PerkinElmer, Waltham, MS, USA). These sections were counterstained with 4′,6-diamidino-2-phenylindole (DAPI) and photographed under a fluorescence microscope (E800; Canon, Tokyo, Japan) with a CCD camera.

The number of ALP-, Osterix-, and F4/80-positive cells in the microscopic fields was quantified in a blinded evaluation. The number and the total luminal area of CD31-positive vessels in the microscopic fields were quantified by using ImageJ in a blinded evaluation, as described previously.[12, 17]

Quantitative real-time PCR

Total RNA was isolated from the cells and tissues by using an RNeasy Mini Kit or an RNeasy Micro Kit (Qiagen, Hilden, Germany), according to the manufacturer's instructions. Reverse transcription was performed using a High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Foster City, CA, USA). Quantitative real-time PCR was assessed by the incorporation of SYBR Green into double-stranded DNA and performed on an ABI StepOne Real-Time PCR System (Applied Biosystems). The PCR primers are listed in Supplemental Table S1. The specific mRNA amplification of the target was determined as the Ct value followed by normalization with the GAPDH level.

Western blot analysis

A 5-mm piece of femur containing the damaged site and intact femur of the contralateral side were removed and homogenized in a tissue lysis buffer (Cell Signaling Technology) supplemented with protease inhibitors. The cultured cells were also lysed with the same buffer. Total protein concentration was determined using the BCA assay reagent (Pierce, Rockford, IL, USA). The same amount of protein aliquots were denatured in sodium dodecyl sulfate (SDS) sample buffer and separated on 10% polyacrylamide-SDS gels. The proteins were transferred to a PVDF membrane. The blots were blocked with a buffer containing 20 mM Tris-HCl (pH 7.6), 137 mM NaCl, 0.1% Tween 20, and 3% skim milk. The membrane was incubated with the anti-VEGF antibody at a dilution of 1:4000, anti-TGF-β antibody at a dilution of 1:4000, anti-phosphorylated Smad2/3 at a dilution of 1:1000, anti-Smad2 antibody at a dilution of 1:1000, anti-Smad3 antibody at a dilution of 1:500, anti-BMP-2 antibody at a dilution of 1:1000, anti-HIF-1α antibody at a dilution of 1:1000, or anti β-actin antibody at a dilution of 1:4000. The immune complexes were visualized using the appropriate secondary antibody and the ECL advance detection system (GE Healthcare, Tokyo, Japan), according to the manufacturer's instructions.

Isolation of F4/80 and CD11b-double positive cells from the femur

The mice were anesthetized using pentobarbital sodium (50 mg/kg, intraperitoneally) on day 4 after the femoral bone defect was created, and the damaged femur was removed. The bone marrow cells were flushed out into Dulbecco's modified Eagle medium (DMEM) with 1% penicillin streptomycin. The cells were labeled at 4°C for 30 minutes with the optimal dilution of phycoerythrin-conjugated rat monoclonal anti-mouse F4/80 antibody (AbD Serotec, Raleigh, NC, USA) and peridinin-chlorophyll protein-Cy5.5-conjugated rat monoclonal anti-mouse CD11b (BD Biosciences, Tokyo, Japan), as well as with the respective isotype control antibodies. After lysis of the erythrocytes, F4/80 and CD11b-double positive cells (5.0 × 105) were isolated using the FACSVantage SE DiVa (BD Biosciences) and analyzed by real-time PCR as described above.

Local treatment of VEGF

The biodegradable polylactic acid (PLA) depot solution was prepared from a mixture of 40% (wt/wt) PLA (Resomer R 202 H, Sigma, St. Louis, MO, USA), 5% (wt/wt) benzyl alcohol (Wako Pure Chemical), and 55% (wt/wt) benzyl benzoate (Wako Pure Chemical).[4] Recombinant murine VEGF (ProSpec, East Brunswick, NJ, USA) was homogenized in the PLA depot solution for 3 minutes at 5500 rpm. After a femoral bone defect was made, as described above, 10 µL of PLA depot solution containing either 10 µg VEGF or only solvent was added to the damaged site on the femur before suturing the incised skin.

Fibrinogen depletion

Fibrinogen depletion was induced by treatment with recombinant batroxobin (ProSpec), a defibrinogenating drug, as described previously.[12] Batroxobin was administered intravenously (3 BU/kg) to Plg+/+ and Plg−/− mice at 1 hour before and subcutaneously (1.5 BU/kg) every 12 hours over 4 days from 3 hours after the bone defect. On day 4 after the bone defect, the mice were euthanized, and bone sections were prepared and stained with the anti-Osterix antibody or anti-CD31 antibody as described above. The control groups received saline using the same schedule as for the administration of batroxobin. The concentration of plasma fibrinogen was measured and calculated from standard curves, as described previously.[18] The concentration of plasma fibrinogen decreased to an undetectable level at 1 hour after the first intravenous administration of batroxobin. Higher and larger numbers of doses of batroxobin are lethal in Plg−/− mice.

Preparation of primary osteoblasts and bone marrow stromal cells

Calvarial osteoblastic cells were obtained from Plg+/+ and Plg−/− mice, according to the method described previously.[19] Briefly, the calvaria was removed from 3-day-old mice, cleaned of soft tissue, and digested 4 times with 1 mg/mL type IV collagenase and 0.25% trypsin for 20 minutes at 37°C with gentle agitation. Cells from the second, third, and fourth digestions were plated and grown in α-MEM with 10% fetal bovine serum (FBS). The cells were seeded in a 6-well plate (1.0 × 105 cells/well) and grown until confluent. For osteogenic differentiation, the medium was changed to α-MEM with 10% FBS, 50 µg/mL of ascorbic acid, and 10 mM β-glycerophosphate.

Bone marrow stromal cells were obtained from Plg+/+ and Plg−/− mice as described previously.[20] Briefly, femurs and tibias were removed from the mice and cleaned of soft tissue. The bone marrow cells were flushed out into DMEM. After the cells were grown in DMEM with 10% FBS for 24 hours, the nonadherent cells were removed using phosphate-buffered saline. For osteogenic differentiation of the bone marrow stromal cells, the medium was changed to α-MEM with 10% FBS, 100 ng/mL BMP-2, 50 µg/mL ascorbic acid, and 10 mM β-glycerophosphate after the cells were passaged once.

The medium was supplemented with 1% penicillin-streptomycin and changed twice a week.

Mineralization assay

Mineralization of calvarial osteoblastic cells and bone marrow stromal cells was determined in 6-well plates by Alizarin Red staining, as described previously.[21] The calvarial osteoblastic and bone marrow stromal cells were cultured in the osteogenic differentiation medium for 3 and 2 weeks, respectively. The cells were fixed with ice-cold 70% ethanol and stained with Alizarin Red S solution (Wako Pure Chemical) to detect calcification of the cells. For quantification, the stained cells were destained with 10% cetylpyridinium chloride (Wako Pure Chemical), the extracted stain was transferred to a 96-well plate, and absorbance was measured at 570 nm.

Statistical analysis

Data are expressed as the mean ± the standard error of the mean (SEM). Statistical significance was evaluated using Student's t test for two-group comparisons. The significance level was set at p < 0.05.

Results

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Delayed bone repair after a femoral bone defect in Plg−/− mice

The damaged site on the femur was repaired progressively until day 14 in Plg+/+ mice, as assessed by qCT (Fig. 1A). Conversely, the damaged site still remained on day 14 in Plg−/− mice (Fig. 1B). The bone repair was significantly delayed in Plg−/− mice on days 4, 7, and 14 compared with Plg+/+ mice (Fig. 1C). In H&E-stained sections, bone repair was found to be delayed in Plg−/− mice (Fig. 1H–K) compared with Plg+/+ mice (Fig. 1D–G). Newly generated bone tissue and bone lining cells were observed from day 7 in Plg+/+ mice (Fig. 1L–O). Although the newly generated bone tissue appeared on day 14 in Plg−/− mice (Fig. 1P–S), there was less tissue than in Plg+/+ mice, and bone lining cells were not observed on day 14 in Plg−/− mice (Fig. 1S). No fibrin clots were observed in the blood vessels of Plg+/+ or Plg−/− mice on day 14 after a femoral bone defect (data not shown). These data indicate that plasminogen plays a crucial role in bone repair.

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Figure 1. Delayed bone repair after a femoral bone defect in Plg−/− mice. (A, B) Three-dimensional images of the damaged site after a femoral bone defect in Plg+/+ (A) and Plg−/− (B) mice, as assessed by quantitative computed tomography (qCT). The arrowheads indicate the damaged site. (C) Quantification of area of bone defect on days 1, 4, 7, and 14 in Plg+/+ and Plg−/− mice. Data represent the mean ± standard error of the mean (SEM) of 4 mice. *p < 0.05. **p < 0.01. n.s. = not significant. (D–K) H&E-stained vertical sections collected from the damaged site in Plg+/+ (D–G) and Plg−/− (H–K) mice after a femoral bone defect. The arrows indicate the edge of the damaged site. Ct = cortical bone. (L–S) Enlarged images at the damaged site in Plg+/+ (L–O) and Plg−/− (P–S) mice. Scale bars = 100 µm (D–K) and 50 µm (L–S). The results represent experiments performed on 4 mice in each group.

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Decrease in chondrogenesis during the repair of a femoral bone defect in Plg−/− mice

In the sections stained with Alcian and toluidine blue, there was significantly less cartilage matrix in Plg−/− mice than in Plg+/+ mice (Fig. 2A–H). The levels of type II and X collagens in the damaged femur of Plg−/− mice were decreased compared with Plg+/+ mice (Fig. 2I, J). These data indicate that chondrogenesis is impaired by plasminogen deficiency during bone repair.

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Figure 2. Decrease in cartilage formation at the damaged site after a femoral bone defect in Plg−/− mice. (A–F) Microphotographs of Alcian blue- (A, D) or toluidine blue-stained sections (B, C, E, F) at the damaged site in Plg+/+ (A–C) and Plg−/− (D–F) mice on day 7 after a femoral bone defect. (C) and (F) are enlarged photographs of the squares in (B) and (E), respectively. The arrows indicate the edge of the damaged site. Scale bars = 200 µm (A, B, D, E) and 100 µm (C, F). The results represent experiments performed on 4 mice in each group. (G, H) Quantification of the area and height of the metachromatic region in the sections stained with toluidine blue in Plg+/+ and Plg−/− mice on day 7 after a femoral bone defect. (I, J) Relative mRNA levels of type II collagen (Col II) (I) and type X collagen (Col X) (J) in the damaged femur of Plg+/+ and Plg−/− mice on day 7. Data represent the mean ± SEM of 4 mice (G–J). *p < 0.05. Ct = cortical bone.

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Localization of osteoblasts and osteoclasts at the damaged site after a femoral bone defect

ALP-positive cells were abundant on the bone surface at the damaged site in Plg+/+ mice on day 7 (Fig. 3A, B). In contrast, the number of ALP-positive cells and the levels of ALP mRNA in Plg−/− mice were significantly decreased on day 7 compared with Plg+/+ mice (Fig. 3C–F). The number of Osterix-positive cells and the levels of Osterix mRNA in Plg−/− mice were significantly decreased compared with Plg+/+ mice (Fig. 3G–J). The number of TRAP-positive multinucleated cells was not significantly different between Plg+/+ and Plg−/− mice on days 7 and 14 (Fig. 3K–O). These results suggest that bone formation is impaired by plasminogen deficiency during bone repair.

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Figure 3. Localization of ALP-, Osterix-, and TRAP-positive cells at the damaged site after a femoral bone defect. (A–D) Microphotographs of ALP-positive cells at the damaged site on days 4 (A, C) and 7 (B, D) after a femoral bone defect in Plg+/+ (A, B) and Plg−/− (C, D) mice. (E) The number of ALP-positive cells at the damaged site on day 7 after a femoral bone defect in Plg+/+ and Plg−/− mice. (F) Relative mRNA levels of ALP in the damaged femur of Plg+/+ and Plg−/− mice. (G, H) Microphotographs of Osterix-positive cells at the damaged site on day 4 after a femoral bone defect in Plg+/+ (G) and Plg−/− (H) mice. (I) The number of Osterix-positive cells at the damaged site on day 4. (J) Relative mRNA levels of Osterix in the damaged femur of Plg+/+ and Plg−/− mice. (K–N) Microphotographs of TRAP-positive cells at the damaged site on days 7 (K, M) and 14 (L, N) in Plg+/+ (K, L) and Plg−/− (M, N) mice. The inset shows an enlarged image of the small square in (K). The arrows indicate the edge of the damaged site. (O) The number of TRAP-positive multinucleated cells on the bone surface at the damaged site on day 7 in Plg+/+ and Plg−/− mice. Scale bars = 50 (A–D, G, H) and 100 µm (K–N). The results represent experiments performed on 4 mice in each group (A–D, G, H, K–N). Data represent the mean ± SEM of 4 mice (E, F, I, J, O). *p < 0.05. **p < 0.01. n.s. = not significant.

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Decrease in angiogenesis at the damaged site during bone repair in Plg−/− mice

Because vessel formation is a critical event for bone repair, we examined the effect of plasminogen deficiency on vessel formation. Blood vessels were abundant at the damaged site in Plg+/+ mice on days 4 and 7, although they were scarce in Plg−/− mice (Fig. 4A–D). The number and luminal area of vessels in Plg−/− mice were significantly decreased compared with Plg+/+ mice (Fig. 4E, F). These results indicate that angiogenesis was impaired by plasminogen deficiency during bone repair.

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Figure 4. Impaired angiogenesis at the damaged site after a femoral bone defect in Plg−/− mice. (A–D) Microphotographs of CD31-positive blood vessels at the damaged site on days 4 (A, C) and 7 (B, D) after a femoral bone defect in Plg+/+ (A, B) and Plg−/− (C, D) mice. (E, F) Quantification of blood vessels at the damaged site on days 4 and 7 in Plg+/+ and Plg−/− mice. The number (E) and total luminal area (F) of blood vessels in the damaged site are shown. (G, H) Microphotographs of VEGF immunostaining at the damaged site on day 4 after a femoral bone defect in Plg+/+ (G) and Plg−/− (H) mice. The arrows indicate the edge of the damaged site. Ct = cortical bone. (I–L) Western blot analysis for VEGF (I), TGF-β, Smad2/3 phosphorylation (pSmad2/3), Smad2, Smad3 (J), BMP-2 (K), and HIF-1α (L) in the damaged and contralateral intact femur on day 4 after a bone defect in Plg+/+ and Plg−/− mice. The results represent experiments performed on 4 mice in each group (A–D, G–L). Data represent the mean ± SEM of 4 mice (E, F). Scale bars = 50 (A–D) and 200 µm (G, H). **p < 0.01.

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VEGF is upregulated by several growth factors, eg, TGF-β and BMP-2, and HIF-1α at the damaged site after bone injury;[5, 22-24] therefore, we examined their expression levels during bone repair. Although VEGF expression was clearly observed at the damaged site in Plg+/+ mice, it was greatly decreased in Plg−/− mice (Fig. 4G, H). The upregulation of VEGF protein levels in the damaged femur was suppressed in Plg−/− mice (Fig. 4I). Similarly, the amount of TGF-β was markedly upregulated in the damaged femur of Plg+/+ mice, although its increase was less in Plg−/− mice (Fig. 4J). The phosphorylation of Smad2/3 in the damaged femur of Plg−/− mice was decreased compared with Plg+/+ mice (Fig. 4J). The upregulation of BMP-2 and HIF-1α in the damaged femur was not significantly different between Plg+/+ and Plg−/− mice (Fig. 4K, L). These data suggest that angiogenesis is suppressed by plasminogen deficiency, presumably through a reduction of TGF-β, rather than by decreased levels of BMP-2 and HIF-1α.

Role of plasminogen in the accumulation of macrophages and their expression of TGF-β during bone repair

Because recruited macrophages are thought to be one of the major sources of TGF-β during tissue repair, we examined macrophage accumulation and TGF-β levels in macrophages in the damaged femur. The accumulation of macrophages in Plg−/− mice was significantly reduced at the damaged site of the femur on day 4 compared with Plg+/+ mice (Fig. 5A–C). TGF-β levels in macrophages were decreased in the damaged femur of Plg−/− mice compared with Plg+/+ mice (Fig. 5D). There was no significant difference in macrophage VEGF levels between Plg+/+ and Plg−/− mice (Fig. 5E). These results indicate that the accumulation of macrophages and TGF-β expression in macrophages are suppressed by plasminogen deficiency during bone repair.

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Figure 5. Effect of plasminogen deficiency on macrophage accumulation and TGF-β and VEGF levels in macrophages after a femoral bone defect. (A, B) Microphotographs of F4/80-positive cells at the damaged site on day 4 after a femoral bone defect in Plg+/+ (A) and Plg−/− (B) mice. The arrows indicate the edge of the damaged site. Scale bars = 200 µm. Ct = cortical bone. The results represent experiments performed on 4 mice in each group. (C) The number of F4/80-positive cells at the damaged site on day 4. (D, E) Relative mRNA levels of TGF-β (D) and VEGF (E) in F4/80 and CD11b-double positive cells derived from the damaged femur of Plg+/+ and Plg−/− mice on day 4. Data represent the mean ± SEM of 4 (C) and 7 mice (D, E). *p < 0.05. n.s. = not significant.

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Effect of local treatment of VEGF on the bone repair after a femoral bone defect

We examined the effects of local treatment of VEGF on bone repair after a femoral bone defect in Plg+/+ and Plg−/− mice. The delayed bone repair in Plg−/− mice was significantly reversed by local treatment with VEGF on day 7 after the femoral bone defect (Fig. 6A, B). A decrease in the number of ALP-positive cells by plasminogen deficiency was reversed by local treatment with VEGF in Plg−/− mice (Fig. 6C–G). Moreover, decreases in the number and luminal area of vessels by plasminogen deficiency were significantly reversed by local treatment with VEGF in Plg−/− mice (Fig. 6H–M). These data suggest that local treatment with VEGF partly reverses delayed bone repair by plasminogen deficiency.

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Figure 6. Effect of local treatment of VEGF on bone repair after a femoral bone defect. (A) Three-dimensional images of the damaged site on day 7 after a femoral bone defect in Plg+/+ and Plg−/− mice treated with solvent (control) or VEGF, as assessed by qCT. The arrowheads indicate the damaged site. (B) Quantification of area of bone defect on day 7 in Plg+/+ and Plg−/− mice. (C–F) Microphotographs of ALP-positive cells at the damaged site on day 7 after a femoral bone defect in Plg+/+ (C, D) and Plg−/− (E, F) mice treated with solvent (control) or VEGF. (G) The number of ALP-positive cells at the damaged site on day 7 after a femoral bone defect in Plg+/+ and Plg−/− mice. (H–K) Microphotographs of CD31-positive blood vessels at the damaged site on day 7 after a femoral bone defect in Plg+/+ (H, I) and Plg−/− (J, K) mice treated with solvent (control) or VEGF. (L, M) Quantification of blood vessels at the damaged site on day 7 in Plg+/+ and Plg−/− mice treated with solvent (control) or VEGF. The number (L) and total luminal area (M) of blood vessels in the damaged site are shown. Scale bars = 50 µm (C–F, H–K). The results represent experiments performed on 6 mice in each group (A, C–F, H–K). Data represent the mean ± SEM of 6 mice (B, G, L, M). *p < 0.05.

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Effect of fibrinogen depletion on the recruitment of Osterix-positive cells and angiogenesis

Defibrination did not affect the recruitment of Osterix-positive cells or angiogenesis at the damaged site of the femur on day 4 in Plg+/+ and Plg−/− mice (Supplemental Fig. S1). These results suggest that circulating fibrinogen might not be involved in the recruitment of osteoblastic cells or angiogenesis at the damaged site during bone repair in the presence or absence of plasminogen.

Role of plasminogen in heterotopic ossification

Next, we examined the role of plasminogen in the heterotopic ossification induced by BMP-2 in muscles. In Plg+/+ and Plg−/− mice, heterotopic bone was formed in a similar manner on day 14, as assessed by qCT (Fig. 7A). No significant difference was observed in total bone mineral content of the heterotopic bone between Plg+/+ and Plg−/− mice (Fig. 7B). There was no significant difference in the formation of cartilage tissue between Plg+/+ and Plg−/− mice (Supplemental Fig. S2A–D). In addition, angiogenesis (Fig. 7C–F), VEGF expression (Fig. 7G–I), macrophage accumulation (Fig. 7J–L), and the number of ALP-positive cells (Fig. 7M–P), Osterix-positive cells (Supplemental Fig. S2E–H), and TRAP-positive multinucleated cells (Supplemental Fig. S2I–K) in the heterotopic bone were not significantly different between Plg+/+ and Plg−/− mice. These results indicate that heterotopic ossification is not impaired by plasminogen deficiency.

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Figure 7. BMP-2-induced heterotopic ossification in Plg+/+ and Plg−/− mice. (A) Three-dimensional images of the heterotopic bone on days 7 and 14 in Plg+/+ and Plg−/− mice, as assessed by qCT. The arrows indicate the heterotopic bone. Scale bars = 5 mm. (B) Total mineral content in the heterotopic bone on day 14 in Plg+/+ and Plg−/− mice. (C, D) Microphotographs of CD31-positive blood vessels in the heterotopic bone on day 7 in Plg+/+ (C) and Plg−/− (D) mice. (E, F) Quantification of blood vessels in the heterotopic bone on day 7 in Plg+/+ and Plg−/− mice. The number (E) and total luminal area (F) of blood vessels in the heterotopic bone are shown. (G, H) Microphotographs of VEGF immunostaining in the heterotopic bone on day 7 in Plg+/+ (G) and Plg−/− (H) mice. (I) Relative mRNA levels of VEGF in the heterotopic bone on day 7 in Plg+/+ and Plg−/− mice. (J, K) Microphotographs of F4/80 immunostaining in the heterotopic bone on day 7 in Plg+/+ (J) and Plg−/− (K) mice. (L) The number of F4/80-positive cells in the heterotopic bone on day 7 in Plg+/+ and Plg−/− mice. (M, N) Microphotographs of ALP-positive cells in the heterotopic bone on day 7 in Plg+/+ (M) and Plg−/− (N) mice. (O) The number of ALP-positive cells in the heterotopic bone on day 7 in Plg+/+ and Plg−/− mice. (P) Relative mRNA levels of ALP in the heterotopic bone on day 7 in Plg+/+ and Plg−/− mice. The results represent experiments performed on 4 mice in each group (A, C, D, G, H, J, K, M, N). Scale bars = 50 µm (C, D, G, H, J, K, M, N). Data represent the mean ± SEM of 4 mice in each group (E, F, I, L, O, P). n.s. = not significant.

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Role of osteoblasts in plasminogen deficiency

We examined the role of osteoblasts in the impaired bone repair by plasminogen deficiency using primary calvarial osteoblastic cells from Plg+/+ and Plg−/− mice. The mRNA levels of Osterix, ALP, and osteocalcin in calvarial osteoblastic cells from Plg−/− mice were elevated compared with those from Plg+/+ mice (Supplemental Fig. S3B, C, E). These findings were similar to those observed in cells cultured in osteogenic medium for 7 days. Conversely, the mRNA levels of Runx2 and type I collagen in calvarial osteoblastic cells were not significantly different between Plg+/+ and Plg−/− mice before and after the cells were cultured in osteogenic medium for 7 days (Supplemental Fig. S3A, D). The mineralization of calvarial osteoblastic cells was significantly increased in Plg−/− mice compared with Plg+/+ mice (Supplemental Fig. S3F, G).

The mRNA and protein levels of VEGF and TGF-β in calvarial osteoblastic cells were not significantly different between Plg+/+ and Plg−/− mice before and after the cells were cultured in osteogenic medium for 7 days (Supplemental Fig. 3H–J). Exogenous plasmin had no effect on VEGF levels in calvarial osteoblastic cells from Plg+/+ mice (Supplemental Fig. 3K).

Mineralization and the expression of osteogenic markers, VEGF, and TGF-β in bone marrow stromal cells during osteogenic differentiation

The mRNA levels of osteogenic markers, including Runx2, Osterix, ALP, type I collagen, and osteocalcin, were elevated in bone marrow stromal cells from both genotypes that were cultured in osteogenic medium for 7 days. However, they were not significantly different between Plg+/+ and Plg−/− mice (Supplemental Fig. S4A–E). Similar levels of mineralization were observed in bone marrow stromal cells from Plg+/+ and Plg−/− mice cultured in osteogenic medium for 14 days (Supplemental Fig. S4F). VEGF levels seemed to decrease during osteogenic differentiation in bone marrow stromal cells from Plg−/− mice compared with Plg+/+ mice (Supplemental Fig. S4G, I). Although TGF-β mRNA levels were not significantly different in bone marrow stromal cells from Plg+/+ and Plg−/− mice that were cultured in osteogenic medium for 7 days (Supplemental Fig. S4H), TGF-β protein was slightly elevated in bone marrow stromal cells from Plg+/+ mice, but not in Plg−/− mice, during osteogenic differentiation (Supplemental Fig. 4I). Exogenous plasmin did not affect the levels of VEGF and TGF-β in bone marrow stromal cells from Plg+/+ mice (Supplemental Fig. S4J).

Discussion

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

The bone-repair process can generally be divided into three phases: acute inflammation, repair, and remodeling.[25] In the acute inflammation phase, neutrophils are recruited rapidly to the damaged site in response to various mediators, coinciding with an increase in macrophage infiltration.[25] The recruited leukocytes release cytokines and growth factors, which stimulate ECM production and angiogenesis. In the repair phase, the femur is repaired through the process of endochondral ossification, ie, the formation of cartilage and the replacement of the cartilage by bone accompanied by the invasion of blood vessels.[25] In the remodeling phase, bone formation and resorption are tightly regulated by osteoblasts and osteoclasts.[25] Plasminogen is a pivotal component of the tissue fibrinolytic system,[6] and plasmin plays crucial roles in the activation of the tissue proteolytic system and the release of growth factors, including TGF-β and VEGF, from the ECM.[7-9] Previous studies indicate that plasminogen is important for tissue repair in the skin and liver beyond the degradation of fibrin.[12, 26, 27] Our present study revealed that plasminogen deficiency impaired the formation of cartilage and bone tissues during bone repair, indicating that plasminogen plays a crucial role in bone repair. Moreover, the tissue fibrinolytic system might be essential for the repair and regeneration of bone.

The newly generated blood vessels support cell proliferation and differentiation at the damaged site during bone repair. They enable the recruitment of mesenchymal stem cells, which differentiate into osteoblasts.[25] Furthermore, the invasion of blood vessels into cartilage tissue is essential for bone formation during endochondral ossification.[2] We revealed that plasminogen deficiency decreased the recruitment of osteoblastic cells and angiogenesis at the damaged site. These findings suggest that impaired angiogenesis is responsible for the delayed bone repair caused by plasminogen deficiency. Moreover, we showed that VEGF expression was similar between the damaged and intact femurs of plasminogen-deficient mice, although VEGF levels were elevated in the damaged femurs of wild-type mice. These findings suggest that VEGF is induced in a plasminogen-dependent manner at the damaged site of the femur. VEGF expression is regulated by several growth factors, including TGF-β and BMP-2, as well as HIF-1α.[5, 22-24] In the present study, plasminogen deficiency suppressed the upregulation of TGF-β and phosphorylation of Smad2/3 in the damaged femur; however, it did not affect the levels of BMP-2 or HIF-1α. These findings indicate that BMP-2 and HIF-1α are not involved in the elevation of VEGF at the damaged femur. Previous studies demonstrated that plasmin activates the latent form of TGF-β and releases its active form from the ECM.[8, 28] Taken together, our findings indicate that plasminogen enhances VEGF expression through TGF-β during bone repair.

TGF-β is produced by several types of cells, eg, endothelial and inflammatory cells.[29] Activated macrophages are thought to be one of the major sources of TGF-β during tissue repair.[30] In the present study, we showed that plasminogen deficiency decreased the levels of TGF-β in F4/80 and CD11b-double positive cells, which are most likely macrophages. These findings indicate that plasminogen is crucial for the induction of TGF-β in macrophages at the damaged site of the femur. The phenotype of macrophages is modulated by the tissue fibrinolytic system during liver repair.[17, 31] Li and colleagues reported that plasmin stimulates the release of cytokines from monocytes/macrophages.[32] Moreover, previous studies indicated that activated macrophages are crucial for osteogenesis.[33-35] We revealed that plasminogen deficiency suppresses the accumulation of macrophages at the damaged site in the present study. These findings indicate that plasminogen plays a pivotal role in the accumulation of macrophages at the damaged site during bone repair.

We showed that the local treatment of VEGF reversed delayed bone repair by plasminogen deficiency. These data indicate that the decrease in VEGF levels at the damaged site might be responsible for the delayed bone repair in Plg−/− mice. However, the effects of VEGF on bone repair were partial in Plg−/− mice. We cannot rule out the possibility that the efficacy of local treatment with VEGF was limited in the present study because the decrease in vessel formation by plasminogen deficiency was only partially reversed by local treatment with VEGF in Plg−/− mice. Alternatively, proteolysis of ECM is important for angiogenesis and requires activation of the tissue proteolytic system, including the tissue fibrinolytic and matrix metalloproteinase systems.[36] Our findings, therefore, might be partially explained by the mechanism, such as the plasmin-activated matrix metalloproteinase system.[7]

The major substrate of plasmin is fibrin. Bugge and colleagues reported that the abnormalities observed in plasminogen-deficient mice, eg, delayed wound healing and hepatocyte necrosis, are rescued by fibrinogen deficiency.[37] These data suggest that an essential physiological role of plasmin is the degradation of fibrin. However, Bezerra and colleagues showed that the impaired liver remodeling in plasminogen-deficient mice was not rescued by fibrinogen deficiency;[11] moreover, fibrinogen deficiency did not completely rescue the impaired wound healing of plasminogen-deficient mice.[27] These data suggest that plasminogen is involved in tissue repair in a fibrin degradation-independent manner. The present study showed that pharmacological defibrination does not affect the recruitment of osteoblastic cells and angiogenesis at the damaged site of the femur in both Plg+/+ and Plg−/− mice. These findings suggest that plasminogen is involved in bone repair independently of the degradation of fibrin by plasmin.

A large number of proinflammatory cytokines and growth factors are released in the acute inflammatory phase after a bone injury.[38, 39] Among them, BMP-2 has a potent osteogenic capacity and plays a role in initiating the fracture repair through endochondral ossification.[40] BMP-2 induces heterotopic ossification in soft tissues through endochondral ossification.[41] Because the process of BMP-2-induced heterotopic ossification is analogous to the repair process of the femur,[41] we examined whether plasminogen plays some role in heterotopic ossification in the present study. We showed that plasminogen deficiency does not affect BMP-2-induced heterotopic ossification, in contrast to a marked delayed bone repair in the femur by plasminogen deficiency. Moreover, the accumulation of macrophages was related to the delayed bone repair in plasminogen-deficient mice. Because numerous bone marrow cells exist in the microenvironment of a damaged site after a femoral bone defect, the presence of sufficient bone marrow cells is thought to be advantageous for the accumulation of macrophages at a damaged site after a femoral bone defect. In addition, the accumulation of macrophages in the heterotopic ossification model seems to be less than in the bone repair model (data not shown). Taken together, macrophages might not be important in heterotopic ossification compared with bone repair; therefore, plasminogen deficiency might have no effect on heterotopic ossification.

Daci and colleagues reported that the absence of both t-PA and u-PA enhances osteoblastic differentiation and mineralization in primary osteoblastic cells.[42] The recent study showed that plasminogen deficiency suppressed osteoprotegerin expression in osteoblastic cells, resulting in an increase in the number of osteoclasts.[13] These data suggest that the tissue fibrinolytic system could affect osteoblastic cells. The present study showed that plasminogen deficiency enhanced mineralization and the levels of several osteogenic markers, including Osterix, ALP, and osteocalcin in calvarial osteoblastic cells, suggesting that plasminogen suppresses the differentiation and mineralization of osteoblasts. We revealed that plasminogen deficiency decreases angiogenesis as well as the accumulation of macrophages at the damaged site during bone repair in the present study. Taken together, the effect of the change on the tissue fibrinolytic system in osteoblasts is not responsible for delayed bone repair caused by plasminogen deficiency. Moreover, the effect of plasminogen on bone repair might be owing to a non-cell autonomous effect in osteoblasts in vivo but not its direct effects on osteoblasts. There were no significant differences in osteogenic differentiation and mineralization of Plg+/+ and Plg−/− mice in bone marrow stromal cell cultures in our study. This finding also supports the fact that the effects of plasminogen are a non-cell autonomous effect in osteogenic cells. We showed that plasminogen deficiency decreases cartilage formation, osteoblastogenesis, and angiogenesis at the damaged site of femur after bone defect, suggesting the possibility that plasminogen might be crucial for endochondral ossification during fracture repair. However, the role of plasminogen in intramembranous ossification is unknown in our study. Further studies are in progress in our laboratory to clarify the details of the role of macrophages in delayed bone repair by plasminogen deficiency and the role of the tissue fibrinolytic system in intramembranous ossification.

Plasminogen activator inhibitor-1 (PAI-1) is a negative regulator of the fibrinolytic system, and the ratio of PAI to plasminogen is important for plasminogen function in various tissues.[43] Rundle and colleagues reported that the absence of PAI-1 increases the size of the callus as well as the formation of callus cartilage during fracture healing in their study in PAI-1-deficient mice.[44] Moreover, the remodeling of the fracture callus was found to be accelerated in PAI-1-deficient mice than in wild-type mice. These data suggest an enhancement of the tissue fibrinolytic system by PAI-1 deficiency that stimulates fracture healing through facilitation of ECM remodeling; this was compatible with our present findings that the plasminogen deficiency impairs bone repair. Taken together, the tissue fibrinolytic system might play an important role in the acute inflammatory phase as well as in the repair and remodeling phases during fracture healing.

The use of autogenous bone grafts for the reconstruction of bone defects has several disadvantages.[1] Although tissue-engineering techniques using biomimetic scaffolds and osteoprogenitor cells are expected to be a promising approach for the reconstruction of bone defects, they are insufficient at the present time.[1] Tissue-engineering techniques require three components: cells, scaffold, and environment.[45] Because our findings imply that plasminogen is critical for the formation of a suitable microenvironment for bone repair, the spatiotemporal regulation of the tissue fibrinolytic system is a promising strategy to meet the clinical needs for the regeneration of bone tissue for reconstruction.

In conclusion, our data provide novel evidence that plasminogen is essential for bone repair. The present study indicates that plasminogen contributes to angiogenesis related to macrophage accumulation, TGF-β, and VEGF, thereby leading to the enhancement of bone repair.

Acknowledgments

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

We thank Mr S Kurashimo (Life Science Research Institute, Kinki University) for supporting the flow cytometry analysis. This study was partly supported by a grant from the Osaka Medical Research Foundation for Intractable Diseases and a Grant-in-Aid for Young Scientists (B: 23790260) to NK, a Grant-in-Aid for Scientific Research (C: 24590289) to HK, and a grant from the Global COE Program F11 from the Ministry of Education, Culture, Sports, Science, and Technology of Japan.

Authors' roles: Study design: NK and HK. Study conduct: HK. Data collection: NK, YT, KO, MY, and KO. Data analysis: NK and HK. Data interpretation: NK, OM, and HK. Drafting manuscript: NK and HK. HK takes responsibility for the integrity of the data analysis.

References

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Supporting Information

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Additional Supporting Information may be found in the online version of this article.

FilenameFormatSizeDescription
jbmr1921-sm-0001-SupFig1.tif4599KSupplementary Figure 1
jbmr1921-sm-0002-SupFig2.tif3590KSupplementary Figure 2
jbmr1921-sm-0003-SupFig3.tif981KSupplementary Figure 3
jbmr1921-sm-0004-SupFig4.tif1116KSupplementary Figure 4
jbmr1921-sm-0005-SupTabs1.doc34KSupplementary Table s1
jbmr1921-sm-0006-SupFigLegend.doc41KSupplementary Figure Legend

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