Calcifying tendinopathy is a tendon disorder with calcium deposits in the mid-substance presented with symptoms including chronic activity-related pain. It is a special case of tendinopathy and the presence of calcified deposits in calcifying tendinopathy worsens its clinical manifestations. Its underlying pathogenesis is poorly understood and treatment is usually symptomatic. Understanding the pathogenesis of calcifying tendinopathy is essential for its effective evidence-based management.
Chondrocytes phenotype/markers were expressed in clinical samples of tendinopathy and calcifying tendinopathy.1, 2 We reported the presence of chondrocyte phenotype and ectopic ossification in a calcifying tendinopathy model3 and the expression of bone morphogenetic protein-2 (BMP-2) protein at those sites,4 suggesting that BMP-2 might be involved in the pathogenesis. This was further supported by the ectopic overexpression of BMPs in the subacromial bursa of patients with chronic degeneration of rotator cuff.5 Tendons harbored stem cells (tendon-derived stem cells, TDSCs) that could differentiate into chondrocytes and osteoblasts.6 As we observed earlier expression of BMP-2 mRNA and protein at week 2 in healing tendon cells, before the time of its appearance in chondrocyte-like cells and calcified deposits in our calcifying tendinopathy model, we hypothesized that calcification is mediated by the erroneous differentiation of TDSCs to chondrocytes/osteoblasts due to the changes in the mechanical and biological microenvironment.4 As overuse is the major etiological factor for overuse-induced tendinopathy and calcifying tendinopathy, we asked if repetitive cyclic loading of TDSCs would increase the expression of BMP-2 and whether BMP-2 could induce the osteogenic differentiation of TDSCs in vitro.
MATERIALS AND METHODS
Isolation and Culture of Rat TDSCs
All experiments were approved by the Animal Research Ethics Committee of the authors' institution. 4- to 6-week-old male Green Fluorescent Protein (GFP) Sprague–Dawley rats, weighting 250–300 g, were used. The procedure was previously established.6 The patellar tendons were excised from healthy rats overdosed with 2.5% sodium phenobarbital. The tissues were minced, digested with type I collagenase (3 mg/ml; Sigma–Aldrich, St Louis, MO) and passed through a 70 µm cell strainer (Becton Dickinson, Franklin Lakes, NJ) to yield single-cell suspension. The released cells were washed and resuspended in Dulbecco's modified Eagle medium (DMEM; Gibco BRL; Life Technologies, Invitrogen, Carlsbad, CA), 10% fetal bovine serum (FBS), 100 U/ml penicillin, 100 µg/ml streptomycin, and 2 mM L-glutamine (complete culture medium; all from Invitrogen Life Technology, Carlsbad, CA) and cultured at 37°C, 5% CO2. At day 2, the cells were washed with phosphate buffered saline (PBS) to remove the nonadherent cells. At day 7, they were trypsinized and mixed together as passage 0 (P0). The clonogenicity and multi-differentiation potential of the cells were confirmed by colony forming assay, osteogenic, adipogenic, and chondrogenic differentiation assays (Supplementary Appendix 1)6 and were maintained up to P6 (the longest passage that we have tested). Cells from P1 to P5 were used for all experiments. Medium was changed every 3 days.
In Vitro Mechanical Stretching
TDSCs were seeded in specially designed silicone culture plate coated with collagen type I (Sigma–Aldrich) at 5 × 103/cm2. After 3 days, the cells were washed with PBS and placed in serum-free medium (DMEM only) just before stretching. This was to avoid the complex interaction between growth factors in serum with mechanical load. Cyclic uniaxial stretching at 0.5 Hz at 4% or 8% (program 3) was used to induce overuse injury to TDSCs using the in vitro mechanical loading system (ST-140-10, B-Bridge International, Inc., Tokyo, Japan) for 4 h. Cells without stretching (0%) served as controls. The stretching regimen was chosen based on the previous study.7 At 4 h after stretch, cells were harvested for BMP-2 mRNA and protein measurement by quantitative real-time RT-PCR (qPCR) and Western blotting, respectively. As the cells were kept in serum-free medium for 8 h only, we did not observe any effect on the viability of the cells.
The cells were lysed, centrifuged, and the supernatant was then collected for measurement of protein concentration by BCA protein assay (Pierce Biotechnology Inc. Rockford, IL). Ten micrograms of protein was denatured, fractionated by electrophoresis on 12% (w/v) SDS–PAGE and electrophoretically transferred to a PVDF membrane (Millipore, Billerica, MA). The blots were blocked with 5% (w/v) nonfat dry milk in TBS/T solution (DAKO, Glostrup, Denamark), incubated with primary antibody against BMP-2 (1:200; Santa Cruz Biotechnology, Santa Cruz, CA), followed by horseradish peroxidase-conjugated secondary antibody (1:3,000; GE Healthcare, Buckinghamshire, UK). Immunoreactive bands were detected by ECL reagents (Pierce Biotechnology Inc., Rockford, IL). The membranes were stripped with Restore Western blot stripping buffer (Pierce Biotechnology Inc., Rockford, IL) and reprobed with β-actin antibody (R&D Systems, Inc., Minneapolis, MN) as a housekeeping control.
Quantitative Real-Time RT-PCR (qPCR)
qPCR was performed as previously described.4 TDSCs were harvested and homogenized for RNA extraction with RNeasy mini kit (Qiagen, Hilden, Germany). The mRNA was reverse transcribed to cDNA by the First-Strand cDNA kit (Promega, Madison, WI). Five microliters of total cDNA of each sample were amplified in a 25 µl reaction mix using the Platinum SYBR Green qPCR SuperMix-UDG with specific primers for BMP-2 and β-actin using the ABI StepOne Plus system (all from Applied Biosystems, CA; Supplementary Appendix 2). Cycling conditions were: denaturation at 95°C for 10 min, 45 cycles at 95°C for 20 s, optimal annealing temperature (Supplementary Appendix 2) for 25 s, 72°C for 30 s, and finally at 60–95°C with a heating rate of 0.1°C/s. The expression of target gene was normalized to that of β-actin gene. Relative gene expression was calculated with the 2−△CT formula.
Effect of BMP-2 on the Osteogenic Differentiation of TDSCs
TDSCs at P3 were plated at 4 × 103 cells/cm2 in a 24-well plate and cultured in complete culture medium until the cells reached confluence. They were then incubated in complete culture medium with or without rhBMP-2 (100 ng/ml; Wyeth, Cambridge, MA) at 37°C, 5% CO2. At day 3, the alkaline phosphatase (ALP) activity of TDSCs was assessed by ALP activity and ALP cytochemical staining assays. At day 10, the calcium nodule formation in TDSCs was assessed by Alizarin red S staining.
ALP Cytochemical Staining Assay
Cells were washed and fixed with 70% EtOH for 30 min at 4°C. After equilibration of cells with ALP buffer (0.1 M NaCl, 0.1 M Tris–HCl pH 9.5, 50 mM MgCl2, and 0.1% Tween-20), ALP substrate solution [0.5 mg nitro-blue tetrazolium chloride (NBT) and 0.25 mg 5-bromo-4-chloro-3′-indolyphosphate p-toluidine salt (BCIP) in 1 ml ALP buffer] was added to the cells for 20 min at 37°C. The color reaction was stopped by washing the cells with distilled water and the positive staining was viewed under the microscope.
ALP Activity Assay
Cells were washed and lysed with lysis buffer with protease inhibitor cocktail (Thermo Fisher Scientific, Bremen, Germany). The supernatant was assayed for ALP activity with an ALP assay kit (BioSystems, Barcelona, Spain) using 4-nitrophenylphosphate as substrate. Production of p-nitrophenol was measured at OD 405 nm for 30 min at 37°C based on the standard curve prepared with different concentrations of p-nitrophenol. The linear portion of the product curve was used for the calculation of enzyme activity by linear regression. The ALP activity was expressed as nmol p-nitrophenol per minute per mg protein.
Alizarin Red S Staining Assay
Cell/matrix layer was washed with PBS, fixed with 70% ethanol, and stained with 0.5% Alizarin red S (pH 4.1, Sigma).6 To quantitate the amount of Alizarin red S bound to the mineralized nodules, cells were rinsed with water, and extracted with 10% (w/v) cetylpyridinium chloride (CPC) in 10 mM sodium phosphate, pH 7.0 for 15 min at room temperature. The dye concentration in the extracts was determined at OD 562 nm.
Data were presented as mean ± SD and shown in boxplots. Comparison of more than two groups was done using Kruskal–Wallis test followed by post hoc comparison with control group using Mann–Whitney U-test. Comparison of two groups was done using Mann–Whitney U-test. All the data analysis was done using SPSS (version 16.0; SPSS Inc, Chicago, IL). p < 0.050 was regarded as statistically significant.
The cells responded to repetitive tensile stretching by aligning along the direction of tensile force (Fig. 1A,B). There was better alignment at 8% compared to 4% stretching. There was increased expression of BMP-2 protein at both 4% and 8% stretching groups compared to the nonstretched control (0%; Fig. 1C). There was also significant increase in the mRNA expression of BMP-2 in the 4% (p = 0.050) but insignificant difference in the 8% stretching group (p = 0.127) compared to the nonstretched control group (Fig. 1D). There was higher ALP activity staining (Fig. 2A–D) and ALP activity (Fig. 2E, p = 0.050) in TDSCs in the BMP-2 treated group compared to the untreated group. More alizarin red-positive calcium nodules (Fig. 3A–D) and significantly higher dye signal intensity (Fig. 3E, p = 0.050) were observed in the BMP-2 treated group compared to the untreated group.
Our results showed that TDSCs were aligned along the direction of mechanical stretching, indicating that they were sensitive to tensile mechanical load, consistent with previous reports on tendon fibroblasts and bone marrow stem cells.8, 9 Not all, but the majority of the cells in the center of the culture plate were aligned along the direction of mechanical stretching. This might be related to the stretching intensity and duration as we observed slightly better cell alignment at 8% compared to 4% and the cells were stretched for 4 h only. Longer stretching time might be required for the better alignment of the cells. The cells in our study were oriented along, rather than away from, the direction of stretching and the results were reproducible. The differences might be due to differences in cell types, cell–cell interaction and cell–matrix interaction. There has been no report about the sensitivity of TDSCs seeded on a smooth culture surface to mechanical loading. The seeding density used in this study was also relatively high, about 80–90% at the time of mechanical loading. Cell–cell interaction might affect the re-orientation of the cells. The silicon culture plates used for mechanical loading were all coated with collagen type I. Cell–substrate interaction might also affect the response of the cells to mechanical load. We further showed that repetitive tensile load increased the protein and gene expression of BMP-2 in TDSCs. This was consistent with the increased expression of BMP-2 mRNA in hMSC in 3D collagen matrices subjected to 10% uniaxial cyclic tensile strain at 1 Hz for 4 h/day for 7 and 14 days.10 Stretching the cells at 12% at the same frequency also increased the expression of BMP-2 mRNA after 14 days but it was marginally insignificant.10 Siddhivarn et al.11 also reported increased mRNA expression of BMP-2, -6, -7, but not BMP-4, under tensile load in an osteoblastic cell line immediately after cell stretching for 1 h with a 1-h resting period and stretched for another hour at 1 Hz with 1% elongation. Their expressions decreased but remained higher than the nonstretched controls up to 6 h.11 There was no obvious difference in the expression of BMP-2 protein at 4% and 8% stretching in this study, probably the response was saturated. This required further proof. Our unpublished results showed that BMP-2 mRNA expression at 4% stretching also returned to control level at 20 h after stretching. Therefore, the insignificant up-regulation of BMP-2 mRNA at 8%, compared to the nonstretched control group, might be due to faster down-regulation of BMP-2 mRNA at higher strain level while the protein half-life was longer than that of mRNA.
BMP-2 promoted the osteogenic differentiation of TDSCs as demonstrated by the increase in ALP activity and matrix mineralization. The effect of BMP-2 on the osteogenic differentiation of MSC was reported in another study.12 Bi et al.13 reported that tendon progenitors/stem cells were sensitive to BMP-2 stimulation though they have not examined the effect of BMP-2 on the differentiation of their isolated stem cells. There was increased ectopic bone formation in tendon with injection of rhBMP-2.14 The osteogenic effect of BMP-2 on TDSCs in this study might explain for the ectopic bone formation.
We postulate that the damage to the extracellular matrix (ECM) as induced by intratendinous collagenase injection might increase the sensitivity of TDSCs to mechanical loading, resulting in increased production of BMP-2 which promoted ectopic ossification in our calcifying tendinopathy model.3 The role of mechanical loading and/or BMP signaling in erroneous stem cell differentiation and heterotropic calcification was also reported in other diseases.15, 16 Zhang and Wang17 reported that 4% stretching promoted tenogenic differentiation of tendon stem cells (TSCs), while 8% stretching induced some of them to differentiate into nontenocytes. The discrepancy with our data might be explained by the different stretching regimen, animal species of TSCs, and the use of culture dish with microgrooves in their study. The same group also reported increased production of PGE2 after a single bout of rigorous treadmill running and PGE2 could induce differentiation of TSCs into nontenocytes.18 There might be interaction between BMP-2 and PGE2 in the induction of osteogenesis by TSCs upon mechanical loading. However, their order of activation was not clear and could be very complicated. In cultured osteoblasts, BMP-2 transcriptionally induced COX-2 expression, an enzyme regulating PGE2 production, via a Cbfa1 binding site which contributed to both the in vitro and in vivo osteogenic effects of BMP-2.19 PGE2 was also reported to down-regulate BMP-2-mediated phosphorylation of Smads 1, 5, and 8 in chondrocytes.20 However, other studies have shown that PGE2 modulated BMP-2 expression in human mesenchymal stem cells21 and osteosacroma cell lines.22 In addition to tensile loading, the expression of BMPs could also be regulated by hydrostatic pressure, shear stress, and compression. BMP-2 mRNA and protein expression was up-regulated by cellular stretch in high pressure-exposed arteries.23 Shear stress down-regulated BMP-4 gene expression in multiple endothelial cells types from different species and BMP-4 was suggested as a proinflammatory, prohypertensive, and proartherogenic mediator in the vessel wall.24 Compressive strain resulted in rapid induction of BMP-2, RUNX2, smad5, and further enhanced the gene and protein expression of ALP, Col1A1, osteocalcin, osteonectin, and osetopontin required for ECM production in pre-osteoblasts.25 We have not studied the expression of BMP-2 in TDSCs under compressive loading which has also been suggested as the factor leading to ectopic chondrogenesis and ossification in tendinopathy as a result of tendon overuse and this would be the direction of further research.26
The mechanism of mechanical up-regulation of BMP-2 was not clear in this study. A share stress–responsive element was found in the promoter region of mouse BMP-227 and mouse BMP-4.28 Mechanical strain was reported to induce BMP-2 expression and bone nodule formation, but not BMP-4 and -6, via ▵12 prostaglandin J2-depedent pathway in an osteoblastic cell line.11 Further studies of the transcriptional regulation of BMP response to mechanical stress will be required to clarify the issue.
We have not studied the effect of mechanical load on the expression of other osteogenic BMPs such as BMP-4, 6, and 7 as well as the effects of these osteogenic BMPs on the chondrogenic and osteogenic differentiation of TDSCs. It was known that the expressions of BMP-4, 6, 7 were also sensitive to mechanical load.11 Therefore, it was likely that BMP-2 is not the sole BMP affecting TDSC differentiation in response to mechanical stimulation. While these require further studies, our result has confirmed the role of BMP-2 in osteogenic differentiation of TDSCs in vitro. The expression of osteogenic markers, other than ALP, by the cells after mechanical stretching and/or BMP-2 stimulation was not studied in the present experiment. Future studies should examine the expression of these osteogenic markers to better understand the mechanisms. Despite this, the increased ALP activity and calcium nodule formation ability of the cells after BMP-2 stimulation were more definite proof of osteogenesis compared to the expression of osteogenic markers. Another limitation of this study was that the stretching system that we used was unable to control cell reorientation during mechanical loading. The mechanical force experienced by the cells therefore depended on the cell orientation and was not a constant. This should be noted in result interpretation of different load intensities on cellular responses. However, the damage of the ECM during tendon injuries would allow the cells to reorient themselves relatively easier compared to cells embedded in a compact normal matrix and our result provided some hints of the cellular responses to mechanical load which allowed cell reorientation as the case during tendon injuries.
Repetitive tensile load increased the expression of BMP-2 and addition of BMP-2 promoted the osteogenic differentiation of TDSCs in vitro. The expression of BMP-2 in TDSCs in response to mechanical overload might provide a possible explanation for ectopic calcification in calcifying tendinopathy.
This work was supported by equipment/resources donated by the Hong Kong Jockey Club Charities Trust, the Restructuring and Collaboration Fund, the CUHK Direct Grant (2009.1.043). We thank Dr. Juan Chen for her assistance in the Western blotting of BMP-2.