Cell Tissue Res DOI 10.1007/s00441-014-2018-2
REGULAR ARTICLE
Uniaxial repetitive mechanical overloading induces influx of extracellular calcium and cytoskeleton disruption in human tenocytes Wan Chen & Yinshuan Deng & Jiqiang Zhang & Kanglai Tang
Received: 11 July 2014 / Accepted: 25 September 2014 # Springer-Verlag Berlin Heidelberg 2014
Abstract Tendon calcification is common in the Achilles tendon, and injuries affect not only athletes, but also the general population. However, the underlying cellular mechanisms are not yet fully understood. In this study, we isolated healthy human tenocytes and subjected them to uniaxial mechanical stretching (at 1.0 Hz) for various stretch times (4 h, 8 h, 12 h) or magnitudes (0 %, 4 %, 8 %, 12 %). The extracellular calcium chelator EGTA, calcium channel inhibitor MnCl2, nifedipine, or various doses of exogenous calcium were administered to these cells with or without mechanical overloading. The intracellular calcium concentration was determined by using a Fluo-3/AM fluorescence probe, and the cytoskeleton was revealed by F-actin Phalloidin staining. The intracellular calcium concentration increased in a magnitudeWan Chen and Yinshuan Deng contributed equally to this study. This work was supported by the Natural Science Foundation of China (81230040, 30872620, and 81071464) and Chongqing Science and Technology Committee (CSTC2011BA5010).
and time-dependent manner following stretching. These increases were suppressed by EGTA, MnCl2, or nifedipine. Additionally, cytoskeleton F-actin was disrupted significantly by stretching in a time-dependent manner. When extracellular calcium was applied, the intracellular calcium concentration increased, and F-actin was disrupted dramatically under mechanical stretching compared with non-stretched cells. Thus, repetitive mechanical overloading induces the accumulation of abnormally high concentrations of intracellular calcium resulting from extracellular calcium influx mediated, at least in part, by membrane calcium channels and finally causes cytoskeleton disorganization and tenocyte dysfunction. These findings provide novel experimental evidence for the pathology of tendon calcification and indicate that the blockade of calcium influx is a potential target for the prevention and treatment of calcific tendinopathy. Keywords Tenocyte . Mechanical overloading . Calcium . Cytoskeleton . Tendinopathy
The authors declare no conflicts of interest. Electronic supplementary material The online version of this article (doi:10.1007/s00441-014-2018-2) contains supplementary material, which is available to authorized users. W. Chen : Y. Deng : K. Tang (*) Department of Orthopedic Surgery, Southwest Hospital, Third Military Medical University, Chongqing 400038, People’s Republic of China e-mail: [email protected] Y. Deng Department of Orthopaedics, Lanzhou General Hospital, Lanzhou 730050, People’s Republic of China J. Zhang (*) Department of Neurobiology, Chongqing Key Laboratory of Neurobiology, Third Military Medical University, Chongqing 400038, People’s Republic of China e-mail: [email protected]
Introduction Tendons are load-bearing tissues that are responsible for the transmission of muscular forces to bone and respond to mechanical loading by changing their metabolism and their structural and mechanical properties (Kjaer 2004). Disorders of the tendon, particularly tendinopathy, affect not only athletes, but also the general population, with a cumulative incidence of 52 % in middle- or long-distance runners (Achilles tendinopathy; Koo et al. 2011) and 32–44 % in jumping athletes (patellar tendinopathy; Oh et al. 2010). Studies have shown that, in Caucasian populations, the prevalence of calcific tendinopathy, which is caused by the pathologic deposition of calcium hydroxyapatite crystals in the tendons and is different from insertional calcific tendinopathy
Cell Tissue Res
(Oliva et al. 2012; Rui et al. 2011; Siegal et al. 2009), is 2.7– 22 % (Oliva et al. 2011). The biological changes of tendinopathy are believed to be associated with resident tenocytes. These cells are fibroblastlike and are characterized by their unique elongated cell shape, which is essential for maintaining their phenotype and function. Whereas mechanical loading is known to be essential for the maintenance of the normal structure and function of tenocytes, repetitive mechanical loading has been regarded as a major causative factor of tendinopathy (Crockett et al. 2010; Wang 2006; Wang et al. 2006) because mechanical stretching induces tenocyte apoptosis (Chao et al. 2008; Endlich et al. 2002) and the production of inflammatory factors such as prostaglandin E2 (Wang et al. 2003b), leukotriene B4 (Thampatty et al. 2006), interleukin-1β (Thampatty et al. 2007; Yang et al. 2005), and glucocorticoids (SabetiAschraf et al. 2010). One of the most important early events in the cellular response to mechanical loading is calcium signaling (Wall and Banes 2005). A mechanical stretch-induced increase of intracellular Ca2+ concentration has been found to occur in vascular smooth muscle cells (VSMCs; Mohanty and Li 2002), cardiac muscles (Miura et al. 2001), and urinary bladder myocytes (Wei et al. 2008). In human tendon, although some previous studies have revealed calcium influx in response to fluid flow stimuli, as mentioned by Wall and Banes (2005), whether cyclic mechanical stretching increases the intracellular calcium concentration and the way in which this is achieved remain to be uncovered. Additionally, several clues indicate that Ca2+ influx causes cell dysfunction, particularly in cytoskeletal actin reorganization or damage. For example, in skeletal muscles, cytoskeletal damage and membrane disruption are mediated primarily by an increased Ca2+ influx (Zhang et al. 2008). In neurons, this influx is responsible for cytoskeletal damage, axonal degeneration, and cell death (Kilinc et al. 2009). Studies have also shown that, under cyclic mechanical stretch, the structure and function of the cytoskeleton are profoundly affected, leading to reorganization or breakage and finally to cell dysfunction or even death (Lozupone et al. 1992; Vico et al. 1998). Furthermore, mechanical stretching has been shown to induce the degradation of alpha-actin filaments in VSMCs (Goldman et al. 2003). However, the effect of mechanical overloading on the integration of F-actin, one of the most important components of the cytoskeleton (Stricker et al. 2010) and the preliminary structure in response to mechanical stress (Lu et al. 2008) in human tenocytes are not yet elucidated. To understand the cellular and molecular mechanisms underlying the development of tendinopathy caused by repetitive mechanical overloading, we collected healthy human tenocytes from surgical waste and investigated the manner in which repetitive mechanical stretch affected intracellular calcium levels and cytoskeleton re-organization in tenocytes by
using the well-established unique microgrooved silicon membrane system (Wang et al. 2003a, 2003b, 2004, 2005).
Materials and methods Isolation and identification of human tenocytes Surgical waste materials from the healthy tendons of 34 patients (from March 2009 to June 2011; 11 males and 23 females; from 20–38 years; average age: 27.5 years) were collected for this study. The patients underwent tendon transplantation surgery, the donor tendon being from the patient’s own peroneus brevis tendon. The protocol for obtaining the human tendon samples from fresh surgical waste was approved by the Institutional Review Board of Southwest Hospital, Third Military Medical University. The isolation of tenocytes was conducted according to the procedure of Cao et al. (2006) with slight modifications. Briefly, after being washed in phosphate-buffered saline (PBS), tissue was minced aseptically into pieces 1 mm × 1 mm × 1 mm in size and transferred to a 25-ml flask. Enzyme solution (0.1 % trypsin and 0.02 % EDTA-Na2 in DMEM/F12; 10 ml) was added to the tissue, which was then incubated for 2 h at 37 °C at 200 rpm on a rocking bed. The product of enzyme digestion was centrifuged, the trypsin was removed, and 20 ml 0.1 % type I collagenase prepared in DMEM/F12 and 5 % fetal bovine serum (FBS) was added to the tissue, which was further incubated for 12 h at 37 °C on the same rocking bed (200 rpm). The cell suspension was collected and centrifuged for 5 min at 1500 rpm. The supernatant was removed, and the cells were washed with PBS and seeded into a 75-ml flask containing 10 ml DMEM/F12, 10 % FBS (with 50 U/ml penicillin and 50 U/ml streptomycin). The primary culture was maintained in a humidified atmosphere of 5 % CO 2 at 37 °C, and the medium was changed every 2–3 days. Once the primary cultured tenocytes had reached approximately 80 % confluence, the medium was removed, and cells were washed with PBS. Subsequently, 3 ml enzyme solution (0.25 % trypsin and 0.02 % EDTA in PBS) was added and incubated with the cells for 3 min. Enzyme digestion was terminated by the addition of 3 ml culture medium (10 % FBS in DMEM/F12). The cells were collected by low centrifugation at 1500 rpm for 5 min, re-suspended in the same culture medium, and cultured at an initial density of 5×105 in a 200-ml culture flask. The cells were subcultured every 3–4 days, and passages 3 and 4 were used in the following experiments unless otherwise indicated. Tenocyte identification was carried out based on the immunoreactivity of the cells for vimentin (Bjur et al. 2008) and
Cell Tissue Res
collagen I (Schulze-Tanzil et al. 2004) but their low or negative immunoreactivity for collagen III (Pauly et al. 2010; Xu et al. 2004). The primary cultured cells were subjected to digestion by using the method mentioned above and reseeded on poly-lysine pre-treated coverslips in a 12-well plate at a density of 1–2×105 cells/ml and cultured for 24 h. The medium was removed, the coverslips were washed with PBS, and the cells were fixed in 4 % formaldehyde (freshly prepared from paraformaldehyde) in PBS for 30 min at room temperature. Fluorescence immunocytochemistry was carried out as follows. The fixative was removed, and the coverslips were washed 3 times with PBS for 5 min each. Normal goat serum (5 %) was added to the coverslips and incubated for 30 min at room temperature to block non-specific binding sites. The cells were then incubated with primary antibody overnight at 4 °C, washed with PBS, incubated with the secondary antibody (fluorescein-isothiocyanate-labeled goat anti-mouse, sc-2082; Santa Cruz, Dallas, USA) for 1 h, counterstained with 4,6-diamidino-2-phenylindole (DAPI; H1200; Molecular Probes; Grand Island, USA), and examined first under an Olympus microscope (BX60; Olympus, Tokyo, Japan) and finally by confocal laser scanning microscopy (TCS2NT; Leica, Wetzlar, Germany). The primary antibodies used in the identification were as follows: monoclonal mouse anti-human collagen I (C2456; Sigma-Aldrich, Munich, Germany), mouse anti-human collagen III (C7805; Sigma-Aldrich), and mouse anti-human vimentin (V5255; Sigma-Aldrich).
Cyclic mechanical stretching on the microgroove silicon membrane system Cell stretching was carried out by using the unique uniaxial microgroove silicon membrane system as previously described (Li et al. 2004; Wang et al. 2005). Before transference of the cells to the microgroove silicon membrane, the membrane was pre-treated with 2 ml ProNectin (10 μg/ml; Z378666; Sigma-Aldrich) for 5 min at room temperature to promote cell attachment, and then the cells were plated at a density of 2×104/cm2 together with 3 ml DMEM/F12 containing 10 % FBS, 100 U/ml penicillin, and 100 U/ml streptomycin in silicon dishes. The cells were cyclically stretched 24 h after plating at three different magnitudes (4 %, 8 %, 12 %) and 1.0 Hz for various times (4, 8, 12 h) as described previously (Wang et al. 2005). A control sample represented cells with no stretching (0 % magnitude) unless indicated otherwise. F-actin Phalloidin staining (77418; Sigma-Aldrich) analysis (see below) was used to verify the growth status and mechanical effects of stretching cells on the microgrooves; cells were not stretched or were stretched for 12 h under 12 % magnitude at 1.0 Hz.
Extracellular calcium blocking by EGTA Tenocytes were cultured on the microgroove silicon membrane for 24 h, and then the medium was replaced with 3 ml medium containing various concentrations (0.5 mmol/l or 1.0 mmol/l) of ethylene glycol tetraacetic acid (EGTA; prepared in serum-free DMEM/F12). Next, the cells were stretched at 8 % magnitude and 1.0 Hz for 4 h. After this stretching step, the medium was removed by washes with Hank’s solution. Calcium-fluorescence examination was carried out (see below), and serum-free DMEM/F12 was used as the blank control. Calcium channel blocking by MnCl2 or nifedipine Cell stretching was carried out at 1.0 Hz and 8 % magnitude. Various concentrations of MnCl2 (0, 0.5 mol/l, or 1.0 mmol/l in DMEM/F12 containing 10 % FBS) or nifedipine (0, 20, 40, or 80 μM in DMEM/F12 with 10 % FBS) were added to the cells, which were stretched for 4 h. Calcium-fluorescence examination was carried out (see below). DMEM/F12 with 10 % FBS medium was used as the blank control. Extracellular calcium administration Tenocytes were cultured on the microgroove silicon membrane for 24 h, the medium was removed, and the cells were washed with PBS. Then, the cells were stretched at 8 % magnitude and 1.0 Hz for 4 h and incubated for 4 h with various concentrations of calcium solution (3.0 mmol/l, 5.0 mmol/l, or 10.0 mmol/l in DMEM/F12 containing 10 % FBS). After the stretching step, the medium was removed by washes with Hank’s solution, and calcium-fluorescence detection and F-actin visualization (Phalloidin staining) were carried out as described below. Intracellular calcium concentration analysis The changes of calcium concentration were examined at 1.0 Hz under various conditions. Cells were stretched at 4 %, 8 %, and 12 % for 4 h, 8 h, and 12 h; non-stretched cells were used as the blank control unless indicated otherwise. After the stretching step, the culture medium was removed, and the cells were washed with Hank’s solution 3 times (5 min each). Fluo-3/AM stock solution was prepared as follows: predissolved Fluo-3/AM (46396; Sigma-Aldrich; 1 mmol/l in dimethylsulfoxide [DMSO]) was diluted with Hank’s solution to a final concentration of 5 μmol/l. PluronicF-127 (P2443; Sigma-Aldrich) stock solution was diluted to 18 % in DMSO (w/v). The working solution was prepared by using a ratio of 5 μl Fluo-3/AM stock solution to 1 ml PluronicF127 (P2443; Sigma-Aldrich) stock solution and mixing well. The working solution was added to the microgrooved silicon
Cell Tissue Res
dishes and incubated at 37 °C for 40 min. After being washed with Hank’s solution, the cells were examined and recorded under a 100× objective lens on a Leica confocal laser scanning microscope (TCS2NT; Leica; excitation: 488 nm; detection: 525 nm). For each experimental group, ten viewing fields were selected, and 10 cells/viewing field were collected. Thus, 100 cells were selected for each group, and the fluorescence intensity for each cell was assessed by using Ion Domain Quantify software (Leica, Germany); the mean density of the 100 cells was used as the Ca2+ concentration in each group.
F-actin visualization: Rhodamine-Phalloidin staining Because the cells on the silicon dishes could not be clearly analyzed under high magnification on the confocal laser scanning microscope directly, because of the thickness of the dishes, we transferred the cells after the stretching step to the ordinary culture system by strictly employing the following procedure. The medium in the silicon dishes was removed, and then cells were digested with trypsin. Next, the cells were re-suspended and plated onto 24-well plates and cultured for 8 h to ensure maximal adherence. F-actin visualization was carried out as described previously (Geiger et al. 2006; Ren et al. 2009) with slight modifications. Briefly, cells were fixed in 4 % formaldehyde (freshly prepared from paraformaldehyde) in PBS for 30 min at room temperature, washed with PBS (3 times, 5 min each), and treated with 0.5 % Triton X-100 (in PBS) for 10 min at room temperature. After being washed with PBS and being blocked with 3 % milk-powder in PBS at 4 °C for 1 h, the cells were incubated with RhodaminePhalloidin (0.2 μg/ml; Invitrogen, Carlsbad, USA) prepared with 1 % BSA for 30 min at room temperature, followed by PBS washes and DAPI counterstaining (5 min at room temperature). Finally, cells were examined by using a Leica Fig. 1 Identification of human tenocytes. a Cell orientation was parallel to the grooves on the silicon membrane dishes; cells showed uniaxial stretching. ×100. b F-actin Rhodamine-Phalloidin staining showing unstretched tenocytes. c Cell orientation at a 12 % magnitude stretch for 12 h at 1.0 Hz; no cell shedding is apparent. d–f Primary tenocytes were immunopositive for vimentin (d) and collagen I (e) but were negative for collagen III (f). DAPI staining (blue) was used to identify cell nuclei (b–f). Bars 20 μm
confocal laser scanning microscope (TCS2NT; Leica). However, to verify the growth status of the cells on the silicon membrane, Rhodamine-Phalloidin staining was carried out on the membrane only without transfer to the coverslips.
Statistical analysis The concentration of intracellular calcium was measured as mentioned above; the optical density of F-actin was measured by using Image Pro Plus software (version 4.5; Media Cybernetics, Rockville, USA) under 100× magnification, unless indicated otherwise. The results are shown as the means± S.E. Data analysis was carried out by one- or two-way analysis of variance (ANOVA), and significance analysis was conducted by using the t-test, the Scheffe test, or Tamhane’s test with SPSS version 13.0 (IBM; Chicago, USA). A P-value of less than 0.05 was considered to be statistically significant.
Results Identification of tenocytes The multi-station microgrooved system was set to run at 1.0 Hz in this study, and six microgrooves were set at 4 %, 8 %, and 12 % loading magnitude as shown in Electronic Supplementary Material Fig. S1. Tenocytes were cultured on the silicon membranes with parallel microgrooves on their surface (Fig. 1a). F-Actin visualization revealed the intact cytoskeleton (Fig. 1b), and no obvious cell shedding occurred, even at 12 % overloading and 1.0 Hz for 12 h (Fig. 1c). These primary tenocytes were immunopositive for vimentin and collagen I but negative for collagen III (Fig. 1d-f).
Cell Tissue Res
Intracellular calcium concentration was upregulated by mechanical stretching When cells were stretched at 0 % or 4 % magnitude, calcium concentration increased slightly from 4 h to 8 h (P0.05; Fig. 2a-a’’, b-b’’). Additionally, at 8 % magnitude (Fig. 2c-c’’), the intracellular calcium concentration increased significantly in a timedependent manner from 4 h to 8 h to 12 h (P < 0.05). However, at 12 % magnitude (Fig. 2d-d’’), no significant difference was detected at each time point examined (P>0.05). When the stretch time was set at 4 h, with the increase in magnitude from 0 % (unstretched) to 12 %, a significant Fig. 2 Increase in concentration of intracellular calcium induced by mechanical overloading (green calcium fluorescence). Both the stretch time and magnitude affected the intracellular calcium levels profoundly (n=100). Cells were cultured on microgroove silicon membranes for 4 h, 8 h, or 12 h with or without stretch (see also Table 1). Bars20 μm
magnitude-dependent increase of intracellular calcium fluorescence was detected (Fig. 2a–d); the highest level was detected at 12 % magnitude (P
REGULAR ARTICLE
Uniaxial repetitive mechanical overloading induces influx of extracellular calcium and cytoskeleton disruption in human tenocytes Wan Chen & Yinshuan Deng & Jiqiang Zhang & Kanglai Tang
Received: 11 July 2014 / Accepted: 25 September 2014 # Springer-Verlag Berlin Heidelberg 2014
Abstract Tendon calcification is common in the Achilles tendon, and injuries affect not only athletes, but also the general population. However, the underlying cellular mechanisms are not yet fully understood. In this study, we isolated healthy human tenocytes and subjected them to uniaxial mechanical stretching (at 1.0 Hz) for various stretch times (4 h, 8 h, 12 h) or magnitudes (0 %, 4 %, 8 %, 12 %). The extracellular calcium chelator EGTA, calcium channel inhibitor MnCl2, nifedipine, or various doses of exogenous calcium were administered to these cells with or without mechanical overloading. The intracellular calcium concentration was determined by using a Fluo-3/AM fluorescence probe, and the cytoskeleton was revealed by F-actin Phalloidin staining. The intracellular calcium concentration increased in a magnitudeWan Chen and Yinshuan Deng contributed equally to this study. This work was supported by the Natural Science Foundation of China (81230040, 30872620, and 81071464) and Chongqing Science and Technology Committee (CSTC2011BA5010).
and time-dependent manner following stretching. These increases were suppressed by EGTA, MnCl2, or nifedipine. Additionally, cytoskeleton F-actin was disrupted significantly by stretching in a time-dependent manner. When extracellular calcium was applied, the intracellular calcium concentration increased, and F-actin was disrupted dramatically under mechanical stretching compared with non-stretched cells. Thus, repetitive mechanical overloading induces the accumulation of abnormally high concentrations of intracellular calcium resulting from extracellular calcium influx mediated, at least in part, by membrane calcium channels and finally causes cytoskeleton disorganization and tenocyte dysfunction. These findings provide novel experimental evidence for the pathology of tendon calcification and indicate that the blockade of calcium influx is a potential target for the prevention and treatment of calcific tendinopathy. Keywords Tenocyte . Mechanical overloading . Calcium . Cytoskeleton . Tendinopathy
The authors declare no conflicts of interest. Electronic supplementary material The online version of this article (doi:10.1007/s00441-014-2018-2) contains supplementary material, which is available to authorized users. W. Chen : Y. Deng : K. Tang (*) Department of Orthopedic Surgery, Southwest Hospital, Third Military Medical University, Chongqing 400038, People’s Republic of China e-mail: [email protected] Y. Deng Department of Orthopaedics, Lanzhou General Hospital, Lanzhou 730050, People’s Republic of China J. Zhang (*) Department of Neurobiology, Chongqing Key Laboratory of Neurobiology, Third Military Medical University, Chongqing 400038, People’s Republic of China e-mail: [email protected]
Introduction Tendons are load-bearing tissues that are responsible for the transmission of muscular forces to bone and respond to mechanical loading by changing their metabolism and their structural and mechanical properties (Kjaer 2004). Disorders of the tendon, particularly tendinopathy, affect not only athletes, but also the general population, with a cumulative incidence of 52 % in middle- or long-distance runners (Achilles tendinopathy; Koo et al. 2011) and 32–44 % in jumping athletes (patellar tendinopathy; Oh et al. 2010). Studies have shown that, in Caucasian populations, the prevalence of calcific tendinopathy, which is caused by the pathologic deposition of calcium hydroxyapatite crystals in the tendons and is different from insertional calcific tendinopathy
Cell Tissue Res
(Oliva et al. 2012; Rui et al. 2011; Siegal et al. 2009), is 2.7– 22 % (Oliva et al. 2011). The biological changes of tendinopathy are believed to be associated with resident tenocytes. These cells are fibroblastlike and are characterized by their unique elongated cell shape, which is essential for maintaining their phenotype and function. Whereas mechanical loading is known to be essential for the maintenance of the normal structure and function of tenocytes, repetitive mechanical loading has been regarded as a major causative factor of tendinopathy (Crockett et al. 2010; Wang 2006; Wang et al. 2006) because mechanical stretching induces tenocyte apoptosis (Chao et al. 2008; Endlich et al. 2002) and the production of inflammatory factors such as prostaglandin E2 (Wang et al. 2003b), leukotriene B4 (Thampatty et al. 2006), interleukin-1β (Thampatty et al. 2007; Yang et al. 2005), and glucocorticoids (SabetiAschraf et al. 2010). One of the most important early events in the cellular response to mechanical loading is calcium signaling (Wall and Banes 2005). A mechanical stretch-induced increase of intracellular Ca2+ concentration has been found to occur in vascular smooth muscle cells (VSMCs; Mohanty and Li 2002), cardiac muscles (Miura et al. 2001), and urinary bladder myocytes (Wei et al. 2008). In human tendon, although some previous studies have revealed calcium influx in response to fluid flow stimuli, as mentioned by Wall and Banes (2005), whether cyclic mechanical stretching increases the intracellular calcium concentration and the way in which this is achieved remain to be uncovered. Additionally, several clues indicate that Ca2+ influx causes cell dysfunction, particularly in cytoskeletal actin reorganization or damage. For example, in skeletal muscles, cytoskeletal damage and membrane disruption are mediated primarily by an increased Ca2+ influx (Zhang et al. 2008). In neurons, this influx is responsible for cytoskeletal damage, axonal degeneration, and cell death (Kilinc et al. 2009). Studies have also shown that, under cyclic mechanical stretch, the structure and function of the cytoskeleton are profoundly affected, leading to reorganization or breakage and finally to cell dysfunction or even death (Lozupone et al. 1992; Vico et al. 1998). Furthermore, mechanical stretching has been shown to induce the degradation of alpha-actin filaments in VSMCs (Goldman et al. 2003). However, the effect of mechanical overloading on the integration of F-actin, one of the most important components of the cytoskeleton (Stricker et al. 2010) and the preliminary structure in response to mechanical stress (Lu et al. 2008) in human tenocytes are not yet elucidated. To understand the cellular and molecular mechanisms underlying the development of tendinopathy caused by repetitive mechanical overloading, we collected healthy human tenocytes from surgical waste and investigated the manner in which repetitive mechanical stretch affected intracellular calcium levels and cytoskeleton re-organization in tenocytes by
using the well-established unique microgrooved silicon membrane system (Wang et al. 2003a, 2003b, 2004, 2005).
Materials and methods Isolation and identification of human tenocytes Surgical waste materials from the healthy tendons of 34 patients (from March 2009 to June 2011; 11 males and 23 females; from 20–38 years; average age: 27.5 years) were collected for this study. The patients underwent tendon transplantation surgery, the donor tendon being from the patient’s own peroneus brevis tendon. The protocol for obtaining the human tendon samples from fresh surgical waste was approved by the Institutional Review Board of Southwest Hospital, Third Military Medical University. The isolation of tenocytes was conducted according to the procedure of Cao et al. (2006) with slight modifications. Briefly, after being washed in phosphate-buffered saline (PBS), tissue was minced aseptically into pieces 1 mm × 1 mm × 1 mm in size and transferred to a 25-ml flask. Enzyme solution (0.1 % trypsin and 0.02 % EDTA-Na2 in DMEM/F12; 10 ml) was added to the tissue, which was then incubated for 2 h at 37 °C at 200 rpm on a rocking bed. The product of enzyme digestion was centrifuged, the trypsin was removed, and 20 ml 0.1 % type I collagenase prepared in DMEM/F12 and 5 % fetal bovine serum (FBS) was added to the tissue, which was further incubated for 12 h at 37 °C on the same rocking bed (200 rpm). The cell suspension was collected and centrifuged for 5 min at 1500 rpm. The supernatant was removed, and the cells were washed with PBS and seeded into a 75-ml flask containing 10 ml DMEM/F12, 10 % FBS (with 50 U/ml penicillin and 50 U/ml streptomycin). The primary culture was maintained in a humidified atmosphere of 5 % CO 2 at 37 °C, and the medium was changed every 2–3 days. Once the primary cultured tenocytes had reached approximately 80 % confluence, the medium was removed, and cells were washed with PBS. Subsequently, 3 ml enzyme solution (0.25 % trypsin and 0.02 % EDTA in PBS) was added and incubated with the cells for 3 min. Enzyme digestion was terminated by the addition of 3 ml culture medium (10 % FBS in DMEM/F12). The cells were collected by low centrifugation at 1500 rpm for 5 min, re-suspended in the same culture medium, and cultured at an initial density of 5×105 in a 200-ml culture flask. The cells were subcultured every 3–4 days, and passages 3 and 4 were used in the following experiments unless otherwise indicated. Tenocyte identification was carried out based on the immunoreactivity of the cells for vimentin (Bjur et al. 2008) and
Cell Tissue Res
collagen I (Schulze-Tanzil et al. 2004) but their low or negative immunoreactivity for collagen III (Pauly et al. 2010; Xu et al. 2004). The primary cultured cells were subjected to digestion by using the method mentioned above and reseeded on poly-lysine pre-treated coverslips in a 12-well plate at a density of 1–2×105 cells/ml and cultured for 24 h. The medium was removed, the coverslips were washed with PBS, and the cells were fixed in 4 % formaldehyde (freshly prepared from paraformaldehyde) in PBS for 30 min at room temperature. Fluorescence immunocytochemistry was carried out as follows. The fixative was removed, and the coverslips were washed 3 times with PBS for 5 min each. Normal goat serum (5 %) was added to the coverslips and incubated for 30 min at room temperature to block non-specific binding sites. The cells were then incubated with primary antibody overnight at 4 °C, washed with PBS, incubated with the secondary antibody (fluorescein-isothiocyanate-labeled goat anti-mouse, sc-2082; Santa Cruz, Dallas, USA) for 1 h, counterstained with 4,6-diamidino-2-phenylindole (DAPI; H1200; Molecular Probes; Grand Island, USA), and examined first under an Olympus microscope (BX60; Olympus, Tokyo, Japan) and finally by confocal laser scanning microscopy (TCS2NT; Leica, Wetzlar, Germany). The primary antibodies used in the identification were as follows: monoclonal mouse anti-human collagen I (C2456; Sigma-Aldrich, Munich, Germany), mouse anti-human collagen III (C7805; Sigma-Aldrich), and mouse anti-human vimentin (V5255; Sigma-Aldrich).
Cyclic mechanical stretching on the microgroove silicon membrane system Cell stretching was carried out by using the unique uniaxial microgroove silicon membrane system as previously described (Li et al. 2004; Wang et al. 2005). Before transference of the cells to the microgroove silicon membrane, the membrane was pre-treated with 2 ml ProNectin (10 μg/ml; Z378666; Sigma-Aldrich) for 5 min at room temperature to promote cell attachment, and then the cells were plated at a density of 2×104/cm2 together with 3 ml DMEM/F12 containing 10 % FBS, 100 U/ml penicillin, and 100 U/ml streptomycin in silicon dishes. The cells were cyclically stretched 24 h after plating at three different magnitudes (4 %, 8 %, 12 %) and 1.0 Hz for various times (4, 8, 12 h) as described previously (Wang et al. 2005). A control sample represented cells with no stretching (0 % magnitude) unless indicated otherwise. F-actin Phalloidin staining (77418; Sigma-Aldrich) analysis (see below) was used to verify the growth status and mechanical effects of stretching cells on the microgrooves; cells were not stretched or were stretched for 12 h under 12 % magnitude at 1.0 Hz.
Extracellular calcium blocking by EGTA Tenocytes were cultured on the microgroove silicon membrane for 24 h, and then the medium was replaced with 3 ml medium containing various concentrations (0.5 mmol/l or 1.0 mmol/l) of ethylene glycol tetraacetic acid (EGTA; prepared in serum-free DMEM/F12). Next, the cells were stretched at 8 % magnitude and 1.0 Hz for 4 h. After this stretching step, the medium was removed by washes with Hank’s solution. Calcium-fluorescence examination was carried out (see below), and serum-free DMEM/F12 was used as the blank control. Calcium channel blocking by MnCl2 or nifedipine Cell stretching was carried out at 1.0 Hz and 8 % magnitude. Various concentrations of MnCl2 (0, 0.5 mol/l, or 1.0 mmol/l in DMEM/F12 containing 10 % FBS) or nifedipine (0, 20, 40, or 80 μM in DMEM/F12 with 10 % FBS) were added to the cells, which were stretched for 4 h. Calcium-fluorescence examination was carried out (see below). DMEM/F12 with 10 % FBS medium was used as the blank control. Extracellular calcium administration Tenocytes were cultured on the microgroove silicon membrane for 24 h, the medium was removed, and the cells were washed with PBS. Then, the cells were stretched at 8 % magnitude and 1.0 Hz for 4 h and incubated for 4 h with various concentrations of calcium solution (3.0 mmol/l, 5.0 mmol/l, or 10.0 mmol/l in DMEM/F12 containing 10 % FBS). After the stretching step, the medium was removed by washes with Hank’s solution, and calcium-fluorescence detection and F-actin visualization (Phalloidin staining) were carried out as described below. Intracellular calcium concentration analysis The changes of calcium concentration were examined at 1.0 Hz under various conditions. Cells were stretched at 4 %, 8 %, and 12 % for 4 h, 8 h, and 12 h; non-stretched cells were used as the blank control unless indicated otherwise. After the stretching step, the culture medium was removed, and the cells were washed with Hank’s solution 3 times (5 min each). Fluo-3/AM stock solution was prepared as follows: predissolved Fluo-3/AM (46396; Sigma-Aldrich; 1 mmol/l in dimethylsulfoxide [DMSO]) was diluted with Hank’s solution to a final concentration of 5 μmol/l. PluronicF-127 (P2443; Sigma-Aldrich) stock solution was diluted to 18 % in DMSO (w/v). The working solution was prepared by using a ratio of 5 μl Fluo-3/AM stock solution to 1 ml PluronicF127 (P2443; Sigma-Aldrich) stock solution and mixing well. The working solution was added to the microgrooved silicon
Cell Tissue Res
dishes and incubated at 37 °C for 40 min. After being washed with Hank’s solution, the cells were examined and recorded under a 100× objective lens on a Leica confocal laser scanning microscope (TCS2NT; Leica; excitation: 488 nm; detection: 525 nm). For each experimental group, ten viewing fields were selected, and 10 cells/viewing field were collected. Thus, 100 cells were selected for each group, and the fluorescence intensity for each cell was assessed by using Ion Domain Quantify software (Leica, Germany); the mean density of the 100 cells was used as the Ca2+ concentration in each group.
F-actin visualization: Rhodamine-Phalloidin staining Because the cells on the silicon dishes could not be clearly analyzed under high magnification on the confocal laser scanning microscope directly, because of the thickness of the dishes, we transferred the cells after the stretching step to the ordinary culture system by strictly employing the following procedure. The medium in the silicon dishes was removed, and then cells were digested with trypsin. Next, the cells were re-suspended and plated onto 24-well plates and cultured for 8 h to ensure maximal adherence. F-actin visualization was carried out as described previously (Geiger et al. 2006; Ren et al. 2009) with slight modifications. Briefly, cells were fixed in 4 % formaldehyde (freshly prepared from paraformaldehyde) in PBS for 30 min at room temperature, washed with PBS (3 times, 5 min each), and treated with 0.5 % Triton X-100 (in PBS) for 10 min at room temperature. After being washed with PBS and being blocked with 3 % milk-powder in PBS at 4 °C for 1 h, the cells were incubated with RhodaminePhalloidin (0.2 μg/ml; Invitrogen, Carlsbad, USA) prepared with 1 % BSA for 30 min at room temperature, followed by PBS washes and DAPI counterstaining (5 min at room temperature). Finally, cells were examined by using a Leica Fig. 1 Identification of human tenocytes. a Cell orientation was parallel to the grooves on the silicon membrane dishes; cells showed uniaxial stretching. ×100. b F-actin Rhodamine-Phalloidin staining showing unstretched tenocytes. c Cell orientation at a 12 % magnitude stretch for 12 h at 1.0 Hz; no cell shedding is apparent. d–f Primary tenocytes were immunopositive for vimentin (d) and collagen I (e) but were negative for collagen III (f). DAPI staining (blue) was used to identify cell nuclei (b–f). Bars 20 μm
confocal laser scanning microscope (TCS2NT; Leica). However, to verify the growth status of the cells on the silicon membrane, Rhodamine-Phalloidin staining was carried out on the membrane only without transfer to the coverslips.
Statistical analysis The concentration of intracellular calcium was measured as mentioned above; the optical density of F-actin was measured by using Image Pro Plus software (version 4.5; Media Cybernetics, Rockville, USA) under 100× magnification, unless indicated otherwise. The results are shown as the means± S.E. Data analysis was carried out by one- or two-way analysis of variance (ANOVA), and significance analysis was conducted by using the t-test, the Scheffe test, or Tamhane’s test with SPSS version 13.0 (IBM; Chicago, USA). A P-value of less than 0.05 was considered to be statistically significant.
Results Identification of tenocytes The multi-station microgrooved system was set to run at 1.0 Hz in this study, and six microgrooves were set at 4 %, 8 %, and 12 % loading magnitude as shown in Electronic Supplementary Material Fig. S1. Tenocytes were cultured on the silicon membranes with parallel microgrooves on their surface (Fig. 1a). F-Actin visualization revealed the intact cytoskeleton (Fig. 1b), and no obvious cell shedding occurred, even at 12 % overloading and 1.0 Hz for 12 h (Fig. 1c). These primary tenocytes were immunopositive for vimentin and collagen I but negative for collagen III (Fig. 1d-f).
Cell Tissue Res
Intracellular calcium concentration was upregulated by mechanical stretching When cells were stretched at 0 % or 4 % magnitude, calcium concentration increased slightly from 4 h to 8 h (P0.05; Fig. 2a-a’’, b-b’’). Additionally, at 8 % magnitude (Fig. 2c-c’’), the intracellular calcium concentration increased significantly in a timedependent manner from 4 h to 8 h to 12 h (P < 0.05). However, at 12 % magnitude (Fig. 2d-d’’), no significant difference was detected at each time point examined (P>0.05). When the stretch time was set at 4 h, with the increase in magnitude from 0 % (unstretched) to 12 %, a significant Fig. 2 Increase in concentration of intracellular calcium induced by mechanical overloading (green calcium fluorescence). Both the stretch time and magnitude affected the intracellular calcium levels profoundly (n=100). Cells were cultured on microgroove silicon membranes for 4 h, 8 h, or 12 h with or without stretch (see also Table 1). Bars20 μm
magnitude-dependent increase of intracellular calcium fluorescence was detected (Fig. 2a–d); the highest level was detected at 12 % magnitude (P
