d
Original Contribution
EFFECTS OF LOW-INTENSITY PULSED ULTRASOUND, DEXAMETHASONE/
TGF-b1 AND/OR BMP-2 ON THE TRANSCRIPTIONAL EXPRESSION OF GENES
IN HUMAN MESENCHYMAL STEM CELLS: CHONDROGENIC VS. OSTEOGENIC
DIFFERENTIATION
C
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AI,
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AIzk{* Department of Biomedical Engineering, Chung Yuan Christian University, Chung Li, Taiwan, ROC;yDepartment of Physical Medicine and Rehabilitation, Taipei Medical University and Hospital, Taipei, Taiwan, ROC;zGraduate Institute of Medical Sciences, College of Medicine, Taipei Medical University, Taipei, Taiwan, ROC;xDepartment of Orthopedics, Taipei County
Hospital, Taipei, Taiwan, ROC;kBrain Imaging Center, McLean Hospital, Belmont, MA, USA; and{Institute of Graduate Clinical Medicine, Taipei Medical University, Taipei, Taiwan, ROC
(Received 1 December 2009; revised 17 March 2010; in final form 17 March 2010)
Abstract—The effects of low-intensity pulsed ultrasound (LIPUS) on the differentiation of human mesenchymal stem cells (hMSCs) were investigated in this study. hMSCs were subjected to LIPUS with or without dexametha-sone/transforming growth factor-b1 (TD) or bone morphogenetic protein-2 (BMP-2) and the effects of this treat-ment were assessed. TD-treated hMSCs exhibited characteristic chondrogenic morphology and increased messenger RNA (mRNA) expression of chondrogenic markers and LIPUS enhanced the chondrogenic differenti-ation of hMSCs treated with TD. The expression of Runx2, an osteogenic transcription factor was not altered in either TD treatment group; however, a significant increase was detected in the LIPUS only group. The osteogenic appearance exhibited 3 days after LIPUS and/or BMP-2 treatment. Increases in the mRNA expression levels of osteogenic markers, Runx2 and ALP were also detected. There was no additive or altered effect with combined LIPUS and BMP-2 treatment. LIPUS alone can increase osteogenic differentiation of hMSCs and LIPUS enhances TD-mediated chondrogenic differentiation of hMSCs. Clinically, LIPUS may differentially influence bone vs.
cartilage repair. (E-mail:[email protected]@[email protected]) Ó 2010
World Federation for Ultrasound in Medicine & Biology.
Key Words: Human mesenchymal stem cells, Low-intensity pulsed ultrasound, Transforming growth factor, Chondrogenic differentiation, Osteogenic differentiation.
INTRODUCTION
Cartilage is an avascular musculoskeletal tissue that possesses little capacity for self-repair after damage (Buckwalter and Mankin 1998). Various treatments for cartilage regeneration including the use of carbon (Brittberg et al. 1994a), periosteum (Hoikka et al. 1990), perichondrium (Homminga et al. 1990), autologous chon-drocyte transplantation (Brittberg et al. 1994b; Peterson
1996) and subchondral drilling have failed to achieve much success (Bouwmeester et al. 2002; Kon et al. 2008; Minas and Nehrer 1997).
Mesenchymal stem cells (MSCs) are thought to have significant potential for facilitating tissue repair (Bruder et al. 1998; Goshima et al. 1991; Maniatopoulos 1988; Nakahara et al. 1990). MSC tissue repair occurs either as a result of MSC migration from bone marrow or by MSC transplantation to the injured area where new cartilage or bony reparative tissue is formed (Cheung et al. 1980; Furukawa et al. 1980). However, because implantation of undifferentiated MSCs for cartilage regeneration may pose a high risk of undesirable effects (such as bone spur formation), this therapeutic approach should be applied cautiously (van der Kraan and van den Berg 2007). In addition, morbidity is frequently asso-ciated with failure of bone regenerative techniques and
Address correspondence to: Walter H. Chang, Ph.D., Department of Biomedical Engineering, Chung Yuan Christian University, 200 Chung-Pei Road, Chung-Li City 32023, Taiwan. E-mail:whchang@ cycu.edu.twor Wen-Fu Thomas Lai, DMD, MScD, DMSc, Institute of Graduate Clinical Medicine, Taipei Medical University, 250 Wuhsing Street, Taipei, Taiwan 110, ROC. E-mail:[email protected] Brain Imaging Center, McLean Hospital, 115 Mill Street, Belmont, MA 02478 USA. E-mail:[email protected]
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complications from bone grafts (Griffin et al. 2008; Kwong and Harris 2008). Therefore, an optimal therapy for promoting cartilage and bone regeneration is still being sought.
Attempts have already been made to enhance chon-drogenic and osteogenic differentiation by treating MSCs with cytokines such as bone morphogenetic proteins (BMPs) (Lecanda et al. 1997; Selvamurugan et al. 2007) and dexamethasone/transforming growth factor-b1 (TD) (Chen et al. 2005; Johnstone et al. 1998). In addition, stimulation with mechanical stresses such as cyclic hydrostatic pressure and noninvasive low-intensity pulsed ultrasound (LIPUS) have shown regenerative potential (Linkhart et al. 1996; Sant’Anna et al. 2005).
LIPUS has been reported to have various biologic effects and triggers different signaling events involved in bone formation (Azuma et al. 2001; Takayama et al. 2007; Warden et al. 2001) and cartilage regeneration (Choi et al. 2006; Cook et al. 2001; Parvizi et al. 1999; Zhang et al. 2003) in vivo and in vitro. Findings from previous studies, for example, have suggested that exposure to LIPUS enhances aggrecan synthesis in rat chondrocytes (Parvizi et al. 1999). Early studies have shown that LIPUS can accelerate bone healing in animal models (Pilla et al. 1990) and in humans (Heckmann et al. 1994). Recent studies continue to confirm the positive effects of LIPUS on healing of bone (Esteki et al. 2010; Rutten et al. 2008), articular cartilage (Jia et al. 2005) and the tendon graft-bone interface (Papatheodorou et al. 2009). Few studies, however, have investigated the effects of LIPUS on the differentiation of human mesenchymal stem cells (hMSCs). It has been recently reported that chon-drogenic differentiation of both animal and human MSCs is enhanced by the application of LIPUS in studies utilizingin vitro and three-dimensional (3-D) models (Ebisawa et al. 2004; Lee et al. 2006, 2007; Schumann et al. 2006). However, there have been no reports describing how LIPUS regulates the signaling of chondrogenic vs. osteogenic differentiation in hMSCs. We hypothesized that LIPUS may differentially modulate the signaling of chondrogenic and osteogenic differentiation in hMSCs in vitro.
The aim of this study was to investigate the regula-tory effects of LIPUS on chondrogenic and osteogenic differentiation of hMSCs induced by cytokines such as transforming growth factor (TGF)-b1, dexamethasone and bone morphogenetic protein-2 (BMP-2). We first examined the effects of these cytokines on differentiation by observing morphologic changes in hMSCs. Second, we used quantitative real-time polymerase chain reaction (PCR).
PCR to measure levels of chondrogenic and osteo-genic markers, such as Sox9, Runx2, aggrecan and type
I and II collagen, expressed by hMSCs after exposure to cytokines. Finally, we examined the effect of combined LIPUS and cytokine treatment on chondrogenic and oste-ogenic differentiation of hMSCs.
MATERIALS AND METHODS
Subjects
Bone marrow was harvested from individuals under-going surgical treatment of femur fractures in the Ortho-pedic Section of Taipei Municipal Chung-Hsin Hospital, Taipei, Taiwan. Exclusion criteria included any history of endocrine disease or hormone replacement therapy. The study was explained to the patients and all patients provided informed consent for harvesting of bone marrow during their surgical procedure. Bone marrow was har-vested from femur fracture sites by proximal femur aspira-tion during surgical treatment procedures. The study was approved by the Institutional Review Board of our hospital.
Isolation and culture of MSCs
MSCs were isolated from human bone marrow. Bone marrow was mixed with sodium-heparin and diluted with five volumes of phosphate buffered saline. The cell suspension was fractionated on a Percoll gradient (60% initial density, Phamacia, Sweden). The MSC-enriched interface fraction was collected and cultured in Dulbec-co’s modified eagle medium with 1 g/mL glucose (DMEM/LG: Sigma, St. Louis, MO, USA), 10% fetal bovine serum (FBS), 1% penicillin streptomycin ampi-cillin (PSA), 0.2% amphotericin B and 1% gentamycin. The culture medium was changed every 3 days. Adherent cell colonies formed during primary culture were passaged when cells exhibited subconfluent proliferation. Second-passage cells were selected to identify the mecha-nisms of differentiation, as reported previously (Chen et al. 2005). The hematopoietic portion of the bone marrow was depleted, as described previously (Chen et al. 2006; Jiang et al. 2002).
Analysis of chondrogenic differentiation in monolayer culture
The hMSCs were seeded onto 48 35-mm plates after primary culture. The medium was changed after 24 h. The 48 plates were divided into four groups: (1) control group: hMSCs cultured in basic medium, without TGF-b1, dexamethasone, or ultrasound (US); (2) LIPUS group: hMSCs cultured in basic medium with LIPUS treatment every 24 h; (3) TD group: hMSCs cultured in chondrogenic medium containing TGF-b1 (10 ng/mL) and dexametha-sone (1027 M) as previously described (Chen et al. 2005); and (4) TD/LIPUS group: hMSCs cultured in
chondrogenic medium containing TGF-b1 (10 ng/mL) and dexamethasone (1027M) with LIPUS treatment every 24 h. There were 12 plates for each group and three plates for each condition. The basic medium contained DMEM/ LG (Sigma D5523), 0.2% FBS, 1% PSA, 0.2% amphoter-icin B and 1% gentamycin. All samples were cultured at 37C with 5% CO2. Culture medium was changed every
3 days until the samples were harvested. Cells exposed to LIPUS were treated with a single 20 min exposure every 24 h over periods of 1, 2, 3 and 4 weeks. Cells not exposed to LIPUS treatment were exposed to sham ultrasound stimulation (the LIPUS generator was not turned on). Samples from the four groups were harvested after 1, 2, 3 and 4 weeks.
Analysis of osteogenic differentiation in monolayer culture
After primary culture, hMSCs were seeded into 12 35-mm plates, which contained basic medium as described above. The 12 plates were divided into four groups: (1) control group: hMSCs cultured in basic medium without BMP-2 (recombinant human BMP-2: Wyeth Research, Cambridge, MA, USA) or LIPUS; (2) LIPUS group: hMSCs cultured in basic medium with LIPUS treatment every 24 h; (3) BMP group: hMSCs cultured in osteogenic medium containing BMP-2 (100 ng/mL) (Lecanda et al. 1997); and (4) BMP/LIPUS group: hMSCs cultured in osteogenic medium with LIPUS treatment every 24 h. There were three plates for each condition. LIPUS treatment involved a single 20 min exposure every 24 h for three days of culture. This treatment was chosen as previous studies have demonstrated that the expression levels of osteogenic genes are maximal on day three (Gleizal et al. 2006; Sant’Anna et al. 2005).
Application of LIPUS
A modified clinical LIPUS device (Sonopuls 490; Delft Instruments, Enraf-Nonius, The Netherlands) was used as previously described (Li et al. 2002, 2003). A diagram of the device is presented inFigure 1. The device consists of a circular surface transducer (T) with a diameter of 2.6 cm, 5.8 cm2circular surface area and an effective radiating area of 5.0 cm2that was placed at the bottom of a glass tank (GT) filled with distilled water (DW) and a culture well with an inner diameter of 33 mm located 30 mm above the transducer (Fig. 1a). A special absorp-tion chamber (SAC) on top of the culture well directly coupled to the culture medium was used to minimize reflections of the LIPUS signal. The SAC was as described in our previous study (Li et al. 2002). Briefly, the chamber consists of a 32 mm diameter covering film (CF) attached to the culture medium (M). This was tightly sealed with an acrylic base covering (AC), which was
sustained by the wall edge of the culture dish (CD). A 40-mm diameter absorption tube film (TF) was in contact with distilled water (DW) and tightly sealed with one end of an acrylic absorption tube (AT). The other end of the AT was sealed with an absorption wall (AW) and the AT contained 5-mm2 absorption particles (APs). Both the AW and APs were made from sound-absorbing rubber foam (CALMFIEX F-6, Pineer Conductor, Taiwan) (Fig. 1b). The acrylic base covering and film were steril-ized with 70% alcohol and then exposed to ultraviolet rays for at least 24 h before the experiments. This
Fig. 1. (a) Diagram of the experimental set-up for exposure of mesenchymal stem cells (MSCs) grown as a monolayer to low-intensity pulse ultrasound (LIPUS). (b) An enlarged diagram of culture dish (CD) covered by specific absorption chamber (SAC) of the LIPUS exposure system. AP 5 absorption particle; AW 5 absorption wall; AT 5 acrylic base absorption tube; SAC 5 special absorption chamber; AR 5 absorption rubber; CD 5 culture dish; D 5 distance from transducer to cell flask, 30 mm; DW 5 distilled water; R 5 x-y rotator; SM 5 stepping motor; T 5 transducer; TC 5 temperature controller; M 5 culture medium; CF 5 covering film; AC 5
absorption chamber did not reflect any ultrasound at the bottom of the film as detected by hydrophone (TNU 001A; NTR Systems, Seattle, WA, USA). The water temperature was modulated with a temperature controller and then degassed using a degassing tank. A stepping motor (SM) and rotator system (R) rotated the US trans-ducer within a 10 mm radius in a horizontal circular plane around the axis of the culture dish at 0.1 Hz to increase the effective exposure area and avoid the production of standing waves (Fig. 1a). The water bath was filled with distilled, deionized and demineralized (3d) water, which was changed before each experiment. The inferior and side walls of the tank were covered with US-absorbing rubber (AR).
The apparatus was set to 1 MHz, pulsed 1:4 (2 ms ‘‘on’’ and 8 ms ‘‘off’’) and the intensity was set to 200 mW/cm2(spatial-average temporal peak intensity) with a repetition rate of 100 Hz. The acoustic output power was measured and calibrated using a precision LIPUS power meter (UPM-DT-10; Ohmic instruments Co., St. Michaels, MD, USA). Findings from our previous study indicated that 1 MHz, pulsed 1:4 (2 ms ‘‘on’’ and 8 ms ‘‘off’’) with a repetition rate of 100 Hz had an optimal effect on osteoblast differentiation (Li et al. 2002, 2003). In contrast to our findings, Ebisawa et al. reported that a 200 ms tone burst repeating at 1.0 kHz, 30mW/cm2 may better stimulate aggrecan synthesis (Ebisawa al. 2004). In addition, other studies found that an intensity of 200 mW/cm2 resulted in more pronounced MSC chondrogenesis in vivo and in vitro (Cui et al. 2006, 2007; Lee et al. 2006). Based on these reports, we chose to use a higher intensity (200 mW/cm2) to promote MSC chondrogenesis and a repetition rate of 100 Hz to optimally stimulate osteoblast differentiation and MSC osteogenesis.
Calibration was considered to be in the acceptable range if the error accuracy of the output readings was #10%. The intensity was previously determined using a calibrated needle hydrophone (TNU 001A, NTR Systems) at the Biomedical Engineering Center, Industrial Technology Research Institute, Taiwan. The output inten-sity varied with axial distance from the transducer head surface as shown inFigure 2. The output intensity varied with axial distance from the transducer head surface as shown inFigure 2. The distance from the face of the trans-ducer to the bottom of the culture dish is 30 mm. At this distance, the power intensity is 164.93 mW/cm2before the culture dish and is 126.40 mW/cm2after the culture dish. The thickness of the bottom of the culture dish is 1 mm, thus, multiple reflections within the bottom of the dish may affect the ultrasound exposure of the cells. The field distribution from the transducer after passing through the culture well bottom was measured as previously described (Li et al. 2002).
Real-time PCR
Total RNA was extracted from each subconfluent monolayer culture (approximately 106MSCs) at the end of the incubation period using Trizol Reagent (Invitrogen Life Technologies, Carlsbad, CA, USA) according to the manufacturer’s instructions. Trizol extracts for the subcon-fluent monolayer cultures for each treatment group were collected separately. Total RNA was reverse-transcribed using the Superscript II System (Invitrogen Life Technolo-gies) to yield complementary DNA (cDNA), which served as the template for real-time PCR to determine the messenger RNA (mRNA) expression levels of chondro-genesis- and osteochondro-genesis-related genes. Primer sequences used in the experiment were as follows: Runx2, 5’-TTA CTT ACA CCC CGC CAGTC-3’and 5’-CAG CGT CAA CAC CAT TC-3’ (Cho et al. 2005); collagen type II, 5’-TGGTCTTGGTGGAAACTTTGC-3’ and 5’-GCC CATTGGTCCTTGGATTA-3’; aggrecan, 5’-TGCATTC CACGAAGCTAACCT-3’ and 5’-CGCCTCGCCTTCTT GAAAT-3’; Sox9, 5’-CAGTACCCGCACTTGCACAA-3’ and 5’-CTCGTTCAGAAGTCTCCAGAGCTT-3’ (Yagi et al. 2005); type I collagen, 5’-TTCCCCCAGCCA-CAAAGAGTC-3’ and 5’-CGTCATCGCACAACACCT-3’ (Miosge et al. 2004); b1 integrin, 5’- AGTGAATGG GAACAACGAGGTC-3’ and 5’-CAATTCCAGCAAC CACACCA-3’ (Chen et al. 2006); and alkaline phosphatase (ALP) 5’-ACCATTCCCACGTCTTCACATTTG-3’ and 5’-AGACATTCTCTCGTTCACCGCC-3’ (Diefenderfer et al. 2003).
Real-time quantification was performed with the LightCycler assay, using a fluorogenic SYBR Green I reaction mixture with the LightCycler instrument (Roche, Mannheim, Germany). Expression levels of all genes were normalized to levels of b-actin mRNA within the same sample. The b-actin primer was designed using PRIMER3 software (http://frodo.wi.mit.edu/cgi-bin/ primer3/primer3_www.cgi) with published sequence data from the NCBI database (GeneBank NID:
Fig. 2. Measurement of axial attenuation of the transducer trans-mission of acoustic waves (1 MHz, 200 mW) through distilled
NM_001101). Primer sequences for b-actin were 5’-GCA TCCCCCAAAGTTCACAA-3’ and 5’-AGGACTGGGC CATTCTCCTT-3’.
Statistical analysis
Continuous variables are presented as mean and stan-dard deviation and were compared between groups by one-way analysis of variance (ANOVA). To identify significant between group differences compared, Bonfer-roni post-hoc tests were performed. Statistical significance was set at 0.05 for one-way ANOVA and for Bonferroni post-hoc tests, a corrected a (a’) 5 0.0083 (i.e., 0.05/6) was used. Statistical analyses were performed using SPSS 15.0 statistical software (SPSS Inc., Chicago, IL, USA).
RESULTS
Morphologic changes associated with chondrogenic differentiation of hMSCs
Control hMSCs exhibited a fibroblast-like morphology at the end of 1 week (Fig. 3a) whereas a less dense cell morphology was noted among the other three groups (Fig. 3b-d). Compared with the other condi-tions, more cuboidal cells were observed in the TD groups, both with and without LIPUS exposure (Fig. 3c and d).
The hMSCs retained fibroblast-like morphology after 2 weeks (Fig. 3e). A lower density of cells with increased intercellular space was evident after LIPUS treatment (Fig. 3f). In comparison with the observations from the first week, cells treated with TD exhibited less of a spindle-shaped morphology and a lower cell density (Fig. 3g). The cells treated with TD and LIPUS attained a mildly rectangular and nodular-like morphology (Fig. 3h).
After 3 weeks of culture, the spindle-shaped morphology of hMSCs was retained and cells were present at a higher density (Fig. 3i). After LIPUS treat-ment, hMSCs appeared loosely arranged, with increased intercellular space, similar to that seen in the second week (Fig. 3j). After TD treatment, hMSCs displayed a mild to moderate rectangular and nodular shape. A more pronounced nodular shape was noted after applying LIPUS to the TD treated cells (Fig. 3k and l).
After 4 weeks of culture, hMSCs retained a spindle-shaped fibroblastic morphology (Fig. 3m). After LIPUS treatment, cells also exhibited a spindle-shaped fibro-blastic morphology and grew at a lower density (Fig. 3n). Cells treated with TD were of a somewhat more rectangular-like shape compared with those at week three (Fig. 3o). Rectangular- and nodular-like morphology was apparent with abundant extracellular matrix in cells exposed to TD with LIPUS (Fig. 3p).
Real-time PCR to detect changes in mRNA levels of genes associated with chondrogenic differentiation of hMSCs
To further support the chondrogenic differentiation of hMSCs induced by applying LIPUS after exposure to cytokines, the expression levels of genes involved in chondrogenic differentiation were analyzed by real-time PCR. The genes analyzed included integrin b1, Sox9, ag-grecan, type I collagen, type II collagen and Runx2.Table 1presents a comparison of the mean expression levels of these genes. Expression levels were measured in hMSCs cultured for 1, 2, 3 and 4 weeks, for each of the four treat-ment groups (control, LIPUS, TD and TD/LIPUS).
A significant difference in integrin b1 mRNA expres-sion levels was noted between the four groups at the second and third weeks (bothp , 0.001). Integrin b1 expression in the TD and TD/LIPUS group were significantly higher compared with in the LIPUS and control groups. Integrin b1 expression levels were optimal in all groups by the second week and were significantly lower at week four (p , 0.001). The integrin b1 expression levels detected at week four were similar to those measured at week one (Fig. 4a).
Expression levels of Sox9 mRNA were significantly different between the four groups at each week analyzed (allp , 0.001), with the exception of week one. In the TD and TD/LIPUS groups, levels of Sox9 mRNA were significantly higher compared with the LIPUS and control groups. Within groups, the mean expression levels during the 4 weeks showed an increasing trend between weeks one and two and then decreased by week four in all groups. Significant differences existed between weeks in the LIPUS group (p 5 0.031), the TD group (p , 0.001) and the TD/LIPUS group (p , 0.001) (Fig. 4b).
Levels of aggrecan mRNA expression differed signifi-cantly between groups at weeks three (p , 0.001) and four (p 5 0.002). However, no there was no between group differ-ences at weeks one and two. Aggrecan mRNA expression levels in the TD and TD/LIPUS groups were significantly higher than levels in the LIPUS and control groups. Signifi-cant differences existed between weeks in all four groups (p , 0.05). There were increasing trends from week one to week four within all four groups, with the highest levels of ag-grecan mRNA detected at week four (Fig. 4c).
Type II collagen mRNA levels were significantly different between groups at weeks three and four (p , 0.001). Levels of type II collagen mRNA were signifi-cantly higher in the TD and TD/LIPUS groups compared with the LIPUS and control groups. Significant differences existed between weeks in all four groups (p , 0.001). The mean expression levels of type II collagen within groups showed increasing trends from weeks one to three, with a slight decrease at week four. In contrast, minimal expres-sion of type I collagen was apparent during weeks one and two. Mean type I collagen mRNA levels in the LIPUS
group were significantly higher than in the TD and TD/LI-PUS groups at each week except for week two. In addition, significant differences were detected between weeks in each group (allp , 0.05). There was a significant trend for the mean levels of type I collagen mRNA to increase from week one to week four in each group (Fig. 4d and e). Interestingly, expression levels of Runx2 mRNA were not altered in the TD groups, with or without LIPUS. Significant between group differences were apparent at every week (p , 0.001). The mean expression level of Runx2 mRNA in the LIPUS group was significantly higher than in the other three groups at each week. There was a trend for Runx2 expression levels to increase from week one to week four, however, the differences were not statistically significant (Fig. 4f).
Morphologic changes associated with osteogenic differentiation of hMSCs
hMSCs exhibited a fibroblast-like morphology after being cultured for 3 days with basic medium while cell density decreased after treatment with LIPUS (Fig. 5a and b). hMSCs treated with BMP and LIPUS exhibited cuboidal morphology (Fig. 5d), with a lower density and increased intercellular space compared with cells in the BMP only group (Fig. 5c).
Real-time PCR results regarding the osteogenic differentiation of hMSCs
To investigate osteogenic differentiation induced by LIPUS and BMP-2 treatment, 12 hMSCs samples were collected and divided into four groups: Control, LIPUS,
Fig. 3. Morphologic changes in the chondrogenic differentiation of human mesenchymal stem cells (hMSCs) in monolayer culture (3100). Cellular morphology and cell density were analyzed by microscopy and representative images of hMSCs grown in monolayer cultures for 1 week (a-d), 2 weeks (e-h), 3 weeks (i-l) and 4 weeks (m-p) are shown. The images repre-sent four conditions analyzed for each week. hMSCs received either no treatment (control, a, e, i, m), low-intensity pulsed ultrasound (LIPUS) only (b, f, j, n), dexamethasone/transforming growth factor b1 (TD) only (c, g, k, o) or combined TD/ LIPUS treatment (d, h, l, p). In general, hMSCs displayed fibroblast-like morphology (a, e, i, m). When hMSCs were treated with only LIPUS, the cells appeared spindle-shaped but had increased intracellular space relative to the control cells. In addi-tion, a lower density of cells was noted (b, f, j, n). Cells treated with TD (c, g, k, o) and TD/LIPUS (d, h, l, p) displayed a more
BMP and BMP/LIPUS. For each treatment group, the mRNA expression levels of four osteogenic markers were measured by real-time PCR. These results are de-picted in Figure 6. The osteogenic markers analyzed were alkaline phosphate (ALP), Runx2, type I collagen and type II collagen. There were significant between group differences in expression levels of ALP (p 5 0.004) and Runx2 (p 5 0.001). The increase in ALP mRNA expres-sion was slightly higher in the BMP/LIPUS group compared with both the LIPUS and BMP groups (Fig. 6a). The mean expression level of Runx2 mRNA in both the LIPUS and BMP groups was significantly increased compared with the control group, however, these levels were slightly lower compared with in the LIPUS/ BMP co-treatment group (Fig. 6b). There were no signifi-cant between group differences in either type I collagen (Fig. 6c) or type II collagen (Fig. 6d) mRNA expression levels. There was low expression of type I collagen and almost no expression of type II collagen in all groups.
DISCUSSION
There is an extensive body of literature reporting on the chondrogenic and osteogenic differentiation of MSCs (Maniatopoulos 1988; Nakahara et al. 1990). Factors
promoting chondrogenic or osteogenic differentiation include the extracellular matrix (ECM) (Chen et al. 2005; Ou et al. 2009), local cytokines (Johnstone et al. 1998) and mechanical stress (Friedl et al. 2007). These factors enable local MSCs to accumulate, proliferate and terminally differentiate into chondrocytes or osteocytes. The use of MSCs in tissue repair has great potential clin-ical significance. Cell culture studies have shown that LIPUS can stimulate expression of Runx2, collagen, alka-line phosphatase, osteocalcin, IGF -1 and TGF-ß1 in fibro-blasts, osteoblasts and bone marrow stromal cells (Azuma et al. 2001; Li et al. 2002; Sant’Anna et al. 2005; Warden et al. 2001).
LIPUS has been reported to have various biologic effects and triggers different signaling events involved in bone formation (Azuma et al. 2001; Takayama et al. 2007; Warden et al. 2001) and cartilage regeneration (Choi et al. 2006; Cook et al. 2001; Parvizi et al. 1999; Zhang et al. 2003) in vivo and in vitro and is currently used clinically for stimulation of bone healing. In addition, there is also considerable interest in the clinical application of BMPs for bone healing. As early as the 1990s LIPUS was shown to accelerate bone healing in rabbit models (Pilla et al. 1990) and in a randomized, double-blind study accelerated healing of tibial shaft
Table 1. Summary of the mRNA expression levels of integrin b1, Sox9, aggrecan, type I collagen, type II collagen and Runx2
Group
Control LIPUS TD TD/LIPUS p value
Integrin b1 Week 1 0.367 (0.058) 0.5 (0.1) 0.5 (0.1) 0.533 (0.058) 0.137 2 0.5 (0.1) 1.1 (0.1)ay 1.067 (0.115)ay 1.133 (0.058)ay ,0.001* 3 0.467 (0.058) 0.8 (0.1)a 0.833 (0.058)a 0.933 (0.058)a ,0.001* 4 0.4 (0.1) 0.5 (0.1)z 0.467 (0.058)z 0.567 (0.058)zx 0.17 Sox9 Week 1 0.012 (0.001) 0.015 (0.001) 0.015 (0.002) 0.017 (0.002) 0.067 2 0.016 (0.003) 0.022 (0.003) 0.034 (0.004)ay 0.049 (0.005)aby ,0.001* 3 0.015 (0.003) 0.021 (0.003) 0.03 (0.003)ay 0.048 (0.004)abcy ,0.001* 4 0.014 (0.001) 0.017 (0.002) 0.024 (0.002)a 0.033 (0.003)abyzx ,0.001* Aggrecan Week 1 0.027 (0.006) 0.03 (0.01) 0.027 (0.006) 0.03 (0.01) 0.916 2 0.053 (0.006) 0.063 (0.021) 0.067 (0.012) 0.077 (0.015) 0.33 3 0.107 (0.015)y 0.143 (0.021)y 0.173 (0.023)yz 0.257 (0.025)abcyz ,0.001* 4 0.113 (0.032)y 0.157 (0.038)y 0.197 (0.032)yz 0.273 (0.032)ayz 0.002* Type I collagen Week 1 0.049 (0.008) 0.066 (0.006) 0.032 (0.003)b 0.03 (0.003)b ,0.001* 2 0.113 (0.015) 0.14 (0.02) 0.087 (0.021) 0.083 (0.025) 0.032* 3 0.233 (0.065) 0.52 (0.053)yz 0.153 (0.025)by 0.16 (0.03)by ,0.001* 4 0.343 (0.09)yz 0.567 (0.09)yz 0.23 (0.026)byz 0.23 (0.036)byz 0.001* Type II collagen Week 1 0.005 (0.001) 0.007 (0.002) 0.006 (0.001) 0.007 (0.003) 0.525
2 0.29 (0.053)y 0.387 (0.095) 0.42 (0.095)y 0.543 (0.186) 0.147 3 0.467 (0.055)y 0.683 (0.146)y 1.087 (0.099)ayz 1.733 (0.134)abcyz ,0.001* 4 0.423 (0.078)y 0.593 (0.122)y 0.967 (0.131)yz 1.667 (0.2)abcyz ,0.001* Runx2 Week 1 0.00065 (0.00011) 0.00156 (0.00012)a 0.00053 (0.00014)b 0.00048 (0.0001)b ,0.001* 2 0.00069 (0.00013) 0.00157 (0.00015)a 0.00052 (0.00008)b 0.00049 (0.00014)b ,0.001* 3 0.00083 (0.00011) 0.00167 (0.00014)a 0.00067 (0.00011)b 0.00073 (0.00014)b ,0.001* 4 0.00097 (0.00017) 0.00163 (0.00019)a 0.00067 (0.00013)b 0.00074 (0.00013)b ,0.001* *p , 0.05 as determined by one-way ANOVA.ySignificant difference between the indicated week and week 1 for a given group as determined by Bonferroni post hoc test.zSignificant difference between the indicated week and week 2 for a given group as determined by Bonferroni post hoc test. xSignificant difference between the indicated week and week 3 for a given group as determined by Bonferroni post hoc test.aSignificant difference
between the indicated group and the control group for a given week as determined by Bonferroni post-hoc test.bSignificant difference between the indi-cated group and the LIPUS group for a given week as determined by Bonferroni post-hoc test.cSignificant difference between the indicated group and the
fractures in humans (Heckmann et al. 1994). More recently, LIPUS was shown to accelerate clinical fracture healing of delayed unions of the fibula (Rutten et al. 2008). The authors found that LIPUS increased osteoid thickness, mineral apposition rate and bone volume at the front of new bony callus formation, which indicated increased osteoblast activity. Other authors have reported accelerated healing of articular cartilage (Jai et al. 2005) and tendon graft-bone interface (Papatheodorou et al. 2009) in rabbit models. Few studies, however, have inves-tigated the effects of LIPUS on the differentiation of human MSCs (hMSCs). We hypothesized that a combina-tion of these two therapies, LIPUS and BMPs, may provide better results than either treatment alone.
This is also true regarding cartilage regeneration research (Jia et al. 2005). If damage to the articular
cartilage extends to within the bone, pluripotential mesen-chymal cells migrate to the site of injury (Buckwalter and Mankin 1998; O’Driscoll et al. 1988). The present study attempted to characterize the effects of LIPUS alone and in combination with cytokines TD and BMP-2 on chon-drogenic and osteogenic differentiation and has potential clinical significance given that LIPUS has already been demonstrated to accelerate bone growth during fracture healing and distraction osteogenesis (Azuma et al. 2001; Reuter et al. 1984; Sant’Anna et al. 2005; Takikawa et al. 2001; Warden et al. 2001).
This study examined the effect of combined TD and LIPUS treatment to induce chondrogenic differentiation of hMSCs. The results demonstrated that TD and TD/LI-PUS induced morphologic changes associated with chon-drogenic differentiation (Fig. 2). In addition, we detected
Fig. 4. Box plots of mRNA expression levels of integrin b1, Sox9, aggrecan, type I collagen, type II collagen and Runx2 under different treatment conditions (graphic presentation ofTable 1data).
increases in the mRNA expression levels of chondrogenic markers such as Sox9, aggrecan and type II collagen in hMSCs treated with TD and TD/LIPUS (Table 1). Our findings are in accordance with what has been reported in the literature. Chondrogenesis of bone marrow progen-itors was reported to be stimulated in the presence of the TD (Chen et al. 2005; Grigoriadis et al. 1996) and in a prior study, we found that levels of type II collagen and Sox9 mRNA increased when hMSCs were treated with TD (Chen et al. 2005).
This study provides evidence of an enhanced effect of LIPUS and TD on chondrogenic differentiation of
hMSCs. Our data indicate that LIPUS enhanced chondro-genic differentiation of TD-treated hMSCs but not of non-TD treated cells. This finding is in agreement with that reported by Ebisawa and colleagues. Combined, our find-ings indicate that the TGF-b1 signal may be mediated by specific membrane receptors and Smad signaling (Attisano and Wrana 2002; van der Kraan et al. 2009). Our data suggests that the effect observed during LIPUS treatment occurs via an integrin-mediated mechanotrans-duction pathway. Specifically, in our three experimental groups, we observed a significant increase in the mRNA expression of integrin b1 after 2 and 3 weeks of hMSC
Fig. 5. Morphologic changes in the osteogenic differentiation of human mesenchymal stem cells (hMSCs) in monolayer culture (3100x). hMSCs exhibited fibroblast-like morphology after three days of culture in basic medium and became less dense after treatment with LIPUS (a, b). When supplemented with bone morphogenetic protein (BMP) and LIPUS, cells exhibited cuboidal morphology (d), with a lower density and more intercellular space compared with cells in BMP only
group (c).
Fig. 6. Effect of low-intensity pulsed ultrasound (LIPUS) and/or bone morphogenetic protein (BMP) treatment on expres-sion of genes related to osteogenic differentiation of human mesenchymal stem cells (hMSC). Quantitative real-time PCR was performed to determine the effects of various treatments on the expression of mRNAs encoding (a) alkaline phosphate (ALP), (b) Runx2, (c) type I collagen and (d) type II collagen. Cells were treated for three days under the following condi-tions: A nontreated control, LIPUS only, BMP only and LIPUS/BMP. The data represent mRNA levels for each gene that
treatment. The increase was followed by a dramatic decrease in integrin b1 mRNA expression at week four to a level similar to that assessed at week one. These results also agree with previous studies reporting that mechanical signaling was mediated by integrins (Lee et al. 2006; Zhou et al. 2004).
Integrins are heterodimeric transmembrane glyco-proteins consisting of one a- and one b-subunit. The b1 subfamily of integrins consists of many ECM receptors, including those for fibronectin, collagens and laminin. For example, integrin a2b1 is the preferential receptor for type II collagen (Loeser et al. 2000). Integrin binding stimulates intracellular signaling which can affect gene expression and alter cellular expression and affinity of in-tegrins (Humphries 1990; Hynes 1992; Loeser et al. 2000). The current hypothesis is that integrins function by inducing a conformational change which results in repositioning of the ligand binding site to a more accessible position away from the cell surface. This conformational change also triggers intracellular signaling. This hypothesis is supported by data demonstrating that, in addition to playing a role promoting adhesion to ECM ligands or counter-receptors on adjacent cells, integrins serve as a transmem-brane mechanical link between extracellular contacts and the cytoskeleton (Hynes 2002). In addition, many integ-rins are not constitutively active; in fact, they are normally expressed on the cellular surface in an inactive or ‘‘OFF’’ state, a state in which they do not bind ligands and do not signal (Hynes 2002). We hypothesize that LIPUS must somehow activate integrins, perhaps by facilitating a mechanical change in the conformation of the integrin molecule. The mRNA expression levels of integrin b1 and the chondrogenic transcription factor, Sox9, were significantly increased at week two. Aggrecan mRNA and type II collagen mRNA levels were increased at weeks three and four. The mRNA levels of integrin b1 (the receptor) and Sox9 (a transcription factor) were upre-gulated more rapidly in this study than mRNA levels of aggrecan and type II collagen.
Our studies revealed that mRNA expression levels of aggrecan and type II collagen were increased by LIPUS treatment. These data confirm findings from previous studies reporting that LIPUS treatment enhanced aggrecan mRNA expression and proteoglycan synthesis of chon-drocytes cultured in a monolayer (Parvizi et al. 1999). Our 2-D data showed that aggrecan and type II collagen did not increase in the presence of LIPUS, with or without TD, until week three. This may be explained by the fact that ECM synthesis is followed by cytokine or mechanical stimulation.
In our study of osteogenic differentiation of hMSCs, mRNA levels of Runx2, an osteogenic transcription factor, did not significantly increase from week one to
four in any group except the LIPUS group. Runx2 mRNA expression levels in the LIPUS treatment group were increased at week one. Expression of type I collagen mRNA did not increase until weeks three and four in the LIPUS group. These results indicate that in hMSCs, LIPUS treatment can only stimulate the expression of genes associated with osteogenesis. Levels of Runx2 mRNA were upregulated at week one while type I collagen mRNA levels did not increase until week three in this study.
It has previously been reported that LIPUS and BMP-2 individually increase Cbfa-1 and ALP mRNA expression, with maximal levels being attained on day three (Sant’Anna et al. 2005). However, combined treat-ment with LIPUS and BMP-2 did not result in a synergistic effect. In another study, it was found that BMP-2 enhanced the osteogenic differentiation of human bone marrow stromal cells in a dose-dependent manner (Lecanda et al. 1997). In this study, we observed a morphologic shift from elongated fibroblast-like cells to shorter and shaped cuboidal cells 3 days after LIPUS and BMP treatment. In addition, we observed increases in the mRNA levels of osteogenic markers such as Runx2 and ALP. This is in keeping with findings from previous reports. We found that there were no significant differences in mRNA expression of Runx2 and ALP among the LIPUS, BMP-2 and LIPUS/BMP-2 groups. BMP-2 receptors act through the Smads signaling proteins, which influence the function of Runx2 (Hanai et al. 1999; Zwijsen et al. 2003).
We acknowledge that our study has several limita-tions. It was difficult to compare week four-induced-chondrogenesis to day three-induced-osteogenesis. In particular, we need to provide additional evidence that cytokines can increase the production of integrins at the cell surface before LIPUS is applied. It appears that LI-PUS treatment can somehow activate integrins by an unknown mechanism.
In conclusion, LIPUS enhances chondrogenic differ-entiation of hMSCs treated with TD. The combined treat-ment of LIPUS and TD induced integrin b1, Sox9, aggrecan and type II collagen mRNA expression. LIPUS alone can increase osteogenic differentiation. LIPUS with BMP induced ALP and Runx2 mRNA expression. Clini-cally, LIPUS may differentially influence bone vs. carti-lage repair.
Acknowledgements—This work was supported by Taipei Medical University, Taiwan, ROC, under grants 99-TMU-TMUH-018.
REFERENCES
Attisano L, Wrana JL. Signal transduction by the TGF-beta super family. Science 2002;296:1646–1647.
Azuma Y, Ito M, Harada Y, Takagi H, Ohta T, Jingushi S. Low-intensity pulsed ultrasound accelerates rat femoral fracture healing by acting
on the various cellular reactions in the fracture callus. J Bone Miner Res 2001;16:671–680.
Bouwmeester PS, Kuijer R, Homminga GN, Bulstra SK, Geesink RG. A retrospective analysis of two independent prospective cartilage repair studies: Autogenous perichondrial grafting versus subchondral dril-ling 10 years post-surgery. J Orthop Res 2002;20:267–273. Brittberg M, Faxe´n E, Peterson L. Carbon fiber scaffolds in the treatment
of early knee osteoarthritis. A prospective 4-year follow-up of 37 patients. Clin Orthop Relat Res 1994a;307:155–164.
Brittberg M, Lindahl A, Nilsson A, Ohlsson C, Isaksson O, Peterson L. Treatment of deep cartilage defects in the knee with autologous chon-drocyte transplantation. N Engl J Med 1994b;331:889–895. Bruder SP, Kurth AA, Shea M, Hayes WC, Jaiswal N, Kadiyala S. Bone
regeneration by implantation of purified, culture-expanded human mesenchymal stem cells. J Orthop Res 1998;16:155–162. Buckwalter JA, Mankin HJ. Articular cartilage: Degeneration and
osteo-arthritis, repair, regeneration, and transplantation. Instr Course Lect 1998;47:487–504.
Chen CW, Boiteau RM, Lai WF, Barger SW, Cataldo AM. sAPPalpha enhances the transdifferentiation of adult bone marrow progenitor cells to neuronal phenotypes. Curr Alzheimer Res 2006;3:63–70. Chen CW, Tsai YH, Deng WP, Shih SN, Fang CL, Burch JG, Chen WH,
Lai WF. Type I and II collagen regulation of chondrogenic differen-tiation by mesenchymal progenitor cells. J Orthop Res 2005;23: 446–453.
Chen Z, Evans WH, Pflugfelder SC, Li DQ. Gap junction protein con-nexin 43 serves as a negative marker for a stem cell-containing pop-ulation of human limbal epithelial cells. Stem Cells 2006;24: 1265–1273.
Cheung HS, Lynch KL, Johnson RP, Brewer BJ.In vitro synthesis of tissue-specific type II collagen by healing cartilage. I. Short-term repair of cartilage by mature rabbits. Arthritis Rheum 1980;23: 211–219.
Cho HH, Park HT, Kim YJ, Bae YC, Suh KT, Jung JS. Induction of oste-ogenic differentiation of human mesenchymal stem cells by histone deacetylase inhibitors. J Cell Biochem 2005;96:533–542. Choi BH, Woo JI, Min BH, Park SR. Low-intensity ultrasound
stimu-lates the viability and matrix gene expression of human articular chondrocytes in alginate bead culture. J Biomed Mater Res A 2006;79:858–864.
Cook SD, Salkeld SL, Popich-Patron LS, Ryaby JP, Jones DG, Barrack RL. Improved cartilage repair after treatment with low-intensity pulsed ultrasound. Clin Orthop Relat Res 2001; 391(Suppl):S231–S243.
Cui JH, Park K, Park SR, Min BH. Effects of low-intensity ultrasound on chondrogenic differentiation of mesenchymal stem cells embedded in polyglycolic acid: Anin vivo study. Tissue Eng 2006;12:75–82. Cui JH, Park SR, Park K, Choi BH, Min BH. Preconditioning of
mesen-chymal stem cells with low-intensity ultrasound for cartilage forma-tionin vivo. Tissue Eng 2007;13:351–360.
Diefenderfer DL, Osyczka AM, Garino JP, Leboy PS. Regulation of BMP-induced transcription in cultured human bone marrow stromal cells. J Bone Joint Surg Am 2003;85(A Suppl. 3):19–28.
Ebisawa K, Hata K, Okada K, Kimata K, Ueda M, Torii S, Watanabe H. Ultrasound enhances transforming growth factor beta-mediated chondrocyte differentiation of human mesenchymal stem cells. Tissue Eng 2004;10:921–929.
Esteki A, Yasrebi B, Shadmehr A. Pulsed low-intensity ultrasound enhances healing rate in the osteoperforated tibia in a rabbit model. J Orthop Trauma 2010;24:170–175.
Friedl G, Schmidt H, Rehak I, Kostner G, Schauenstein K, Windhager R. Undifferentiated human mesenchymal stem cells (hMSCs) are highly sensitive to mechanical strain: Transcriptionally controlled early osteo-chondrogenic response in vitro. Osteoarthr Cartilage 2007; 15:1293–1300.
Furukawa T, Eyre DR, Koide S, Glimcher MJ. Biochemical studies on repair cartilage resurfacing experimental defects in the rabbit knee. J Bone Joint Surg Am 1980;62:79–89.
Gleizal A, Li S, Pialat JB, Beziat JL. Transcriptional expression of calva-rial bone after treatment with low-intensity ultrasound: Anin vitro study. Ultrasound Med Biol 2006;32:1569–1574.
Goshima J, Goldberg VM, Caplan AI. The origin of bone formed in composite grafts of porous calcium phosphate ceramic loaded with marrow cells. Clin Orthop Relat Res 1991;269:274–283.
Griffin XL, Costello I, Costa ML. The role of low-intensity pulsed ultra-sound therapy in the management of acute fractures: A systematic review. J Trauma 2008;65:1446–1452.
Grigoriadis AE, Heersche JN, Aubin JE. Analysis of chondroprogenitor frequency and cartilage differentiation in a novel family of clonal chondrogenic rat cell lines. Differentiation 1996;60:299–307. Hanai J, Chen LF, Kanno T, Ohtani-Fujita N, Kim WY, Guo WH,
Imamura T, Ishidou Y, Fukuchi M, Shi MJ, Stavnezer J, Kawabata M, Miyazono K, Ito Y. Interaction and functional cooperation of PEBP2/ CBF with Smads. Synergistic induction of the immunoglobulin germ line C alpha promoter. J Biol Chem 1999;274:31577–31582. Heckman JD, Ryaby JP, McCabe J, Frey JJ, Kilcoyne RF. Acceleration
of tibial fracture-healing by noninvasive, low-intensity pulsed ultra-sound. J Bone Joint Surg Am 1994;76:26–34.
Hoikka VE, Jaroma HJ, Ritsila¨ VA. Reconstruction of the patellar artic-ulation with periosteal grafts. Four-year follow-up of 13 cases. Acta Orthop Scand 1990;61:36–39.
Homminga GN, Bulstra SK, Bouwmeester PS, van der Linden AJ. Peri-chondral grafting for cartilage lesions of the knee. J Bone Joint Surg Br 1990;72:1003–1007.
Humphries MJ. The molecular basis and specificity of integrin-ligand interactions. J Cell Sci 1990;97:585–592.
Hynes RO. Integrins: Bidirectional, allosteric signaling machines. Cell 2002;110:673–687.
Hynes RO. Integrins: Versatility, modulation, and signaling in cell adhe-sion. Cell 1992;69:11–25.
Jia XL, Chen WZ, Zhou K, Wang ZB. Effects of low-intensity pulsed ultrasound in repairing injured articular cartilage. Chin J Traumatol 2005;8:175–178.
Jiang Y, Jahagirdar BN, Reinhardt RL, Schwartz RE, Keene CD, Ortiz-Gonzalez XR, Reyes M, Lenvik T, Lund T, Blackstad M, Du J, Aldrich S, Lisberg A, Low WC, Largaespada DA, Verfaillie CM. Pluripotency of mesenchymal stem cells derived from adult marrow. Nature 2002;418:41–49.
Johnstone B, Hering TM, Caplan AI, Goldberg VM, Yoo JU.In vitro chondrogenesis of bone marrow-derived mesenchymal progenitor cells. Exp Cell Res 1998;238:265–272.
Kon E, Delcogliano M, Filardo G, Montaperto C, Marcacci M. Second generation issues in cartilage repair. Sports Med Arthrosc 2008;16: 221–229.
van der Kraan PM, van den Berg WB. Osteophytes: Relevance and biology. Osteoarthr Cartil 2007;15:237–244.
van der Kraan PM, Blaney Davidson EN, Blom A, van den Berg WB. TGF-beta signaling in chondrocyte terminal differentiation and osteoarthritis Modulation and integration of signaling pathways through receptor-Smads. Osteoarthr Cartilage 2009;17:1539–1545. Kwong FN, Harris MB. Recent developments in the biology of fracture
repair. J Am Acad Orthop Surg 2008;16:619–625.
Lecanda F, Avioli LV, Cheng SL. Regulation of bone matrix protein expression and induction of differentiation of human osteoblasts and human bone marrow stromal cells by bone morphogenetic protein-2. J Cell Biochem 1997;67:386–396.
Lee HJ, Choi BH, Min BH, Park SR. Low-intensity ultrasound inhibits apoptosis and enhances viability of human mesenchymal stem cells in three-dimensional alginate culture during chondrogenic differenti-ation. Tissue Eng 2007;5:1049–1057.
Lee HJ, Choi BH, Min BH, Son YS, Park SR. Low-intensity ultrasound stimulation enhances chondrogenic differentiation in alginate culture of mesenchymal stem cells. Artif Organs 2006;30:707–715. Li JG, Chang WH, Lin JC, Sun JS. Optimum intensities of ultrasound for
PGE(2) secretion and growth of osteoblasts. Ultrasound Med Biol 2002;28:683–690.
Li JK, Chang WH, Lin JC, Ruaan RC, Liu HC, Sun JS. Cytokine release from osteoblasts in response to ultrasound stimulation. Biomaterials 2003;24:2379–2385.
Linkhart TA, Mohan S, Baylink DJ. Growth factors for bone growth and repair: IGF, TGF beta and BMP. Bone 1996;19(1 Suppl):1S–12S. Loeser RF, Sadiev S, Tan L, Goldring MB. Integrin expression by
a differential role for alpha1beta1 and alpha2beta1 integrins in medi-ating chondrocyte adhesion to types II and VI collagen. Osteoarthr Cartilage 2000;8:96–105.
Maniatopoulos C, Sodek J, Melcher AH. Bone formationin vitro by stromal cells obtained from bone marrow of young adult rats. Cell Tissue Res 1988;254:317–330.
Minas T, Nehrer S. Current concepts in the treatment of articular cartilage defects. Orthopedics 1997;20:525–538.
Miosge N, Hartmann M, Maelicke C, Herken R. Expression of collagen type I and type II in consecutive stages of human osteoarthritis. His-tochem Cell Biol 2004;122:229–236.
Nakahara H, Bruder SP, Haynesworth SE, Holecek JJ, Baber MA, Goldberg VM, Caplan AI. Bone and cartilage formation in diffusion chambers by subcultured cells derived from the periosteum. Bone 1990;11:181–188.
O’Driscoll SW, Keeley FW, Salter RB. Durability of regenerated artic-ular cartilage produced by free autogenous periosteal grafts in major full-thickness defects in joint surfaces under the influence of contin-uous passive motion. A follow-up report at one year. J Bone Joint Surg Am 1988;70:595–606.
Ou KL, Wu J, Lai WF, Yang CB, Lo WC, Chiu LH, Bowley J. Effects of the nanostructure and nanoporosity on bioactive nanohydroxyapa-tite/reconstituted collagen by electrodeposition. J Biomed Mater Res A 2010;92:906–912.
Papatheodorou LK, Malizos KN, Poultsides LA, Hantes ME, Grafanaki K, Giannouli S, Ioannou MG, Koukoulis GK, Protopappas VC, Fotiadis DI, Stathopoulos C. Effect of transosseous application of low-intensity ultrasound at the tendon graft-bone inter-face healing: Gene expression and histological analysis in rabbits. Ultrasound Med Biol 2009;35:576–584.
Parvizi J, Wu CC, Lewallen DG, Greenleaf JF, Bolander ME. Low-intensity ultrasound stimulates proteoglycan synthesis in rat chondrocytes by increasing aggrecan gene expression. J Orthop Res 1999;4:488–494. Peterson L. Articular cartilage injuries treated with autologous
chondro-cyte transplantation in the human knee. Acta Orthop Belg 1996; 62(Suppl 1):196–200.
Pilla AA, Mont MA, Nasser PR, Khan SA, Figueiredo M, Kaufman JJ, Siffert RS. Noninvasive low-intensity pulsed ultrasound accelerates bone healing in the rabbit. J Orthop Trauma 1990;4:246–253. Rutten S, Nolte PA, Korstjens CM, van Duin MA, Klein-Nulend J.
Low-intensity pulsed ultrasound increases bone volume, osteoid thickness
and mineral apposition rate in the area of fracture healing in patients with a delayed union of the osteotomized fibula. Bone 2008;43:348–354. Reuter U, Strempel F, John F, Knoch HG. Modification of bone fracture healing by ultrasound in an animal experiment model. Z Exp Chir Transplant Kunstliche Organe 1984;17:290–297.
Sant’Anna EF, Leven RM, Virdi AS, Sumner DR. Effect of low-intensity pulsed ultrasound and BMP-2 on rat bone marrow stromal cell gene expression. J Orthop Res 2005;23:646–652.
Schumann D, Kujat R, Zellner J, Angele MK, Nerlich M, Mayr E, Angele P. Treatment of human mesenchymal stem cells with pulsed low-intensity ultrasound enhances the chondrogenic phenotype in vitro. Biorheology 2006;43:431–443.
Selvamurugan N, Kwok S, Vasilov A, Jefcoat SC, Partridge NC. Effects of BMP-2 and pulsed electromagnetic field (PEMF) on rat primary osteoblastic cell proliferation and gene expression. J Orthop Res 2007;25:1213–1220.
Takayama T, Suzuki N, Ikeda K, Shimada T, Suzuki A, Maeno M, Otsuka K, Ito K. Low-intensity pulsed ultrasound stimulates osteo-genic differentiation in ROS 17/2.8 cells. Life Sci 2007;80: 965–971.
Takikawa S, Matsui N, Kokubu T, Tsunoda M, Fujioka H, Mizuno K, Azuma Y. Low-intensity pulsed ultrasound initiates bone healing in rat nonunion fracture model. J Ultrasound Med 2001;20: 197–205.
Warden SJ, Favaloro JM, Bennell KL, McMeeken JM, Ng KW, Zajac JD, Wark JD. Low-intensity pulsed ultrasound stimulates a bone-forming response in UMR-106 cells. Biochem Biophys Res Commun 2001;286:443–450.
Yagi R, McBurney D, Laverty D, Weiner S, Horton WE Jr. Intrajoint comparisons of gene expression patterns in human osteoarthritis suggest a change in chondrocyte phenotype. J Orthop Res 2005;23: 1128–1138.
Zhang ZJ, Huckle J, Francomano CA, Spencer RG. The effects of pulsed low-intensity ultrasound on chondrocyte viability, proliferation, gene expression and matrix production. Ultrasound Med Biol 2003;29: 1645–1651.
Zhou S, Schmelz A, Seufferlein T, Li Y, Zhao J, Bachem MG. Molecular mechanisms of low- intensity pulsed ultrasound in human skin fibro-blasts. J Biol Chem 2004;279:54463–54469.
Zwijsen A, Verschueren K, Huylebroeck D. New intracellular compo-nents of bone morphogenetic protein/Smad signaling cascades. FEBS Lett 2003;546:133–139.
