Establishment of tendon-derived cell lines exhibiting pluripotent mesenchymal stem cell-like property

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Experimental Cell Research 287 (2003) 289 –300

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Establishment of tendon-derived cell lines exhibiting pluripotent mesenchymal stem cell-like property R. Salingcarnboriboon,a H. Yoshitake,a K. Tsuji,a M. Obinata,b T. Amagasa,c A. Nifuji,a and M. Nodaa,* a

Department of Molecular Pharmacology, Medical Research Institute, Tokyo Medical and Dental University, Tokyo, Japan b Department of Cell Biology, Institute of Development, Aging and Cancer, Tohoku University, Sendai, Japan c First Department of Oral and Maxillofacial Surgery, Tokyo Medical and Dental University, Tokyo, Japan Received 4 October 2002, revised version received 14 January 2003

Abstract Development of the musculoskeletal system requires coordinated formation of distinct types of tissues, including bone, cartilage, muscle, and tendon. Compared to muscle, cartilage, and bone, cellular and molecular bases of tendon development have not been well understood due to the lack of tendon cell lines. The purpose of this study was to establish and characterize tendon cell lines. Three clonal tendon cell lines (TT-E4, TT-G11, and TT-D6) were established using transgenic mice harboring a temperature-sensitive mutant of SV40 large T antigen. Proliferation of these cells was significantly enhanced by treatment with bFGF and TGF-␤ but not BMP2. Tendon phenotyperelated genes such as those encoding scleraxis, Six1, EphA4, COMP, and type I collagen were expressed in these tendon cell clones. In addition to tendon phenotype-related genes, expression of osteopontin and Cbfal was observed. These clonal cell lines formed hard fibrous connective tissue when implanted onto chorioallantoic membrane in ovo. Furthermore, these cells also formed tendon-like tissues when they were implanted into defects made in patella tendon in mice. As these tendon cell lines also produced fibrocartilaginous tissues in tendon defect implantation experiments, mesenchymal stem cell properties were examined. Interestingly, these cells expressed genes related to osteogenic, chondrogenic, and adipogenic lineages at low levels when examined by RT-PCR. TT-G11 and TT-E4 cells differentiated into either osteoblasts or adipocytes, respectively, when they were cultured in cognate differentiation medium. These observations indicated that the established tendon cell line possesses mesenchymal stem cell-like properties, suggesting the existence of mesenchymal stem cell in tendon tissue. © 2003 Elsevier Science (USA). All rights reserved. Keywords: Tendon; Cell line; Adipogenic; Osteogenic; Stem cell-like

Introduction Musculoskeletal tissue formation requires coordinated development of distinct types of tissues during embryogenesis. To date, developmental biology on bone, cartilage, and muscle has been the major field of the study. In contrast, regulatory mechanisms involved in tendon and ligament formation have not been well understood. Tendon is a spe-

* Corresponding author. Department of Molecular Pharmacology, Medical Research Institute, Tokyo Medical and Dental University, 3-10 Kanda-Surugadai, 2-chome, Chiyoda-ku, Tokyo 101, Japan. Fax: ⫹81-35280-8066. E-mail address: [email protected] (M. Noda).

cific connective tissue that links skeletal elements and muscles. Like cartilage, tendon originates from lateral plate mesoderm. Drosophila studies suggest that mature tendon cells are singled out from a cluster of competent cells by the influences of approaching muscle cells [1]; however, whether this is also the case in vertebrae is still to be elucidated. In avians, development of proximal limb tendons is closely in association with muscle development [2]. In contrast, distal limb tendons develop from muscle and their morphogenesis is more correlated with that of skeletal elements [2]. The initial autonomous differentiation of tendon primordial cells precedes muscle binding, and it takes place even in limbs that are devoid of muscles. However, when muscles are experimentally removed in limbs, the

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Table 1 Primers used in RT-PCR Primer

Forward primer

Reverse primer

Product size

Cycle

Scleraxis Six-1 EphA4 COMP Col-1 Sox9 Col-X Col-ll

5⬘-AAC ACG GCC TTC ACT GC-3⬘ 5⬘-ACC AGT TCT CGC CTC ACA ATC-3⬘ 5⬘-ACC TTA AGA GTC TTC CGG GCG TCA TG-3⬘ 5⬘-CGC AGC TGC AAG ACG TGA GAG AGC TGT-3⬘ 5⬘-TTT GTG GAC CTC CGG CTC-3⬘ 5⬘-GGC ATG AGT GAG GTG CAC TC-3⬘ 5⬘-AGG CAA GCC AGG CTA TGG AA-3⬘ 5⬘-CTC GCG GTG AGC CAT GAT CCG-3⬘

5⬘-CTT CGA ATC GCC GTC TT-3⬘ 5⬘-TTT TCG GTG TTC TCC CTT TCC-3⬘ 5⬘-GCG TCG ACC ATT CTG CTC CTC GTG CCC A-3⬘ 5⬘-CCG AAT TCC GCT GGT CTG GGT TTC GA-3⬘ 5⬘-AAG CAG AGC ACT CGC CCT-3⬘ 5⬘-CGG AGT TCT GAT GGT CAG CG-3⬘ 5⬘-GCT GTC CTG GAA AGC CGT TT-3⬘ 5⬘-GAG GGC CAG GAG GTC CTC TGG-3⬘

27 27 28 28 23 32 30 32

OPN OC ALP Osterix aP2 PPAR-␥ GAPDH

5⬘-CGA CGA TGA TGA CGA TGA TGA T-3⬘ 5⬘-CTC TGT CTC TCT GAC CTC ACA G-3⬘ 5⬘-ATT GCC CTG AAA CTC CAA AAC C-3⬘ 5⬘-CTG GGG AAA GGA GGC ACA AAG AAG-3⬘ 5⬘-GAA GCT TGT CTC CAG TCA AAA C-3⬘ 5⬘-AAG CTC CAA TAC CAA AGT G-3⬘ 5⬘-ACC ACA GTC CAT GCC ATC AC-3⬘

5⬘-CTG GCT TTG GAA CTT GCT TGA C-3⬘ 5⬘-CAG GTC CTA AAT AGT GAT ACC G-3⬘ 5⬘-CCT CTG GTG GCA TCT CGT TAT C-3⬘ 5⬘-GGG TTA AGG GGA GCA AAG TCA GAT-3⬘ 5⬘-AGT CAC GCC TTT CAT AAC ACA T-3⬘ 5⬘-GGT GAT TTG TCC GTT GTC TTT C-3⬘ 5⬘-TCC ACC ACC CTG TTG CTG TA-3⬘

321 337 490 878 224 690 452 180 380 460 230 460 473 420 520 452

tendon tissues cease their development and degenerate [2– 4]. Signaling molecules in this process have not been fully understood. Tendon and ligament cells have been reported to respond to a variety of growth factors [5,6]. Signaling by TGF-␤ superfamily members, especially BMPs, has been thought to be a key regulator in the development of tendon during chick embryogenesis [7]. TGF-␤ and bFGF mRNAs have been reported to be expressed in both intrinsic tenocytes and extrinsic inflammatory cells in tendon after injury and repair, suggesting the significance of these molecules during tendon development and repair [8 –10]. One of the intriguing features of tendon is its possible plasticity. Conversion of tendon into cartilage has been observed to occur as a consequence of surgical or nonsurgical trauma in human patients as well as animals [11,12]. Thus, in addition to the molecular mechanisms underlying tendon-specific cell differentiation, certain signaling pathways may exist to alter the fate of tendon cells. Though tendons and ligaments are derived from mesenchymal cells, which also give rise to bone, cartilage, fat, and muscle, the presence of precursors or mesenchymal stem cells in tendon has not been studied. If there are such cells, development and repair of tendon would be under the control of certain types of extrinsic signals such as cytokines and intrinsic signals including transcription factors. Scleraxis, a twist-related bHLH transcription factor, is specifically expressed in mesenchymal precursors of connective tissues in early development, while in later developmental stages in mice, scleraxis transcripts are selectively expressed in tendons and their progenitors [13–15]. On the other hand, scleraxis expression is downregulated in developing bones at the onset of ossification [16]. Other transcription factors such as two murine homeobox-containing genes, Six1 and Six2, are expressed in a complementary fashion during the development of limb tendons [17].

bp bp bp bp bp bp bp and bp bp bp bp bp bp bp bp

23 30 28 30 27 30 23

In addition to the transcription factors, several other molecules are expressed in tendon tissues. EphA4 is an Eph family member receptor tyrosine kinase and is expressed in response to regulatory signals during limb patterning. Later in limb development, EphA4 is expressed in cell condensations that form tendons and their attachments to cartilage rudiments [18,19]. Another type of tendon-related molecule is a pentameric noncollagenous glycoprotein, COMP, which is a member of the thrombospondin gene family of extracellular calcium-binding proteins. COMP is also expressed in cartilage and ligament in addition to tendon tissues [20 – 22]. The restricted tissue distribution and expression of COMP in developing as well as adult tendon tissues suggest the involvement of this protein in the regulation of tendon formation [23]. The structural similarities between COMP and thrombospondin could imply that they may perform similar functions implicated in interactions with extracellular matrix components and cells [24]. Although fragmented information has been accumulated, the studies on tendon biology are still in an early stage. Tendon cell research has been hampered by the lack of appropriate cell lines. The purpose of this study is to establish and characterize tendon cell lines. We established three cell lines derived from the Achilles tendon of transgenic

Table 2 cDNA probe list cDNA probes

Fragment

Probe size

Scleraxis (mouse) Type I collagen (mouse) Osteopontin (mouse) Cbfa1 (mouse) Osteocalcin (mouse) Alkaline phosphatase (mouse)

EcoRI, EcoRV EcoRI EcoRI EcoRI EcoRI EcoRI

1.1 kb 1.5 kb 1.4 kb 1.8 kb 520 bp 2.4 kb

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Fig. 1. Morphology of tendon-derived cell lines (A–C). Photomicrograph pictures of the cells were taken using a phase-contrast microscope. Figures are indicating the morphology of TT-E4 (A), TT-G11 (B), and TT-D6 (C).

mice harboring a temperature-sensitive mutant of SV40 large T antigen [25–27].

Materials and methods Isolation and cloning of tendon-derived cells The tendon cells were isolated from Achilles tendon of 8-week-old transgenic mice harboring the SV40 large T antigen gene [25–28]. The clones of tendon-derived cells were established by a limiting dilution technique [27–29]. Briefly, Achilles tendons of transgenic mice were excised and rinsed three times in ␣-MEM containing 10% FBS and antibiotic–antimicotic solution, and then digested in ␣-MEM containing collagenase at 37°C for 20 min. The dissociated cells were resuspended in 2 ml of medium containing 10% FBS and plated into 35-mm cell culture dishes (Corning) at 37°C, 5% CO2. After 24 h, the cells were trypsinized and single cells were diluted and were transferred to 96-well plates (Corning) to yield one cell per well. Survived single cells were recovered from the 96-well plates and were used to generate clones. Three cell clones were selected based on their growth at 33°C, a permissive temperature for the temperature-sensitive mutant of large T antigen, under the 0.5% FBS condition. To confirm immortality, the cells were maintained in continuous culture for at least 1 year in ␣-MEM supplemented with 0.5% FBS at

33°C, 5% CO2 before initiation of characterization. MC3T3E1 and NIH3T3 cells were cultured in ␣-MEM and DMEM supplemented with 10% FBS, respectively, at 37°C, in the atmosphere of 5% CO2. The medium was replaced every 3 days. Total RNA used for reverse transcription polymerase chain reaction (RT-PCR) experiments was prepared from the cultures of these cells at their early confluence. Growth of the cells in response to serum The cells were plated in 4-well tissue culture plates at 1 ⫻ 104 cells/cm2 in ␣-MEM culture medium supplemented with 0.5, 1, and 10% FBS. The numbers of cells were counted at four time points (Days 0, 2, 4, and 6), after harvesting the cells with 1 ml of 0.125% trypsin, and resuspended in 9 ml of Isoton III (Coulter Electronics Ltd. Bedfordshire, England) using a Coultor Counter Model ZM (Coulter Electronics Ltd.). Alkaline phosphatase assay The cells were seeded in 96-well plates at 1 ⫻ 104 cells/cm2 in ␣-MEM culture medium supplemented with 0.5% FBS and cultured until they became confluent. The confluent cells were treated with 500 ng/ml rhBMP2 for 3 days. Cell lysates were prepared by rinsing the cells twice with 0.9% saline solution, and scraping the cell layer into 50

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␮l of a buffer containing 10 mM Tris–HCl, pH 7.5, 0.5 mM MgCl2, and 0.1% Triton X-100. After the cell lysates were subjected to repeated freeze-and-thaw, alkaline phosphatase activities were measured by incubation in an aliquot of 50 ␮l substrate solution [2 mM PNPP (Na2-p-nitrophenyl phosphate), 20 mM MgCl2, 1 M AMP (2-amino-2-methyl1-propanol), H2O; mixed at 1:1:1:6 ratio composition] for 1 h. Samples were subjected to spectrophotometry at 415 nm using a microplate reader (Bio-Rad Model 450). Specific alkaline phosphatase activities were calculated by standardizing absorbance values per mg protein/h. MTT assay

Fig. 2. (A) Effect of serum on the growth of the tendon cell lines. The tendon cell line were cultured in 4-well plates in ␣-MEM medium supplemented with 0.1, 0.5, and 10% of fetal bovine serum (FBS). The cells were cultured up to 6 days and at each time point cells were subjected to count by using a Coulter counter as described under Materials and methods. For each bar, 3 wells were used for obtaining the values. The figure indicates the percentage of the data obtained from one of the two independent experiments with similar results. (B) Effect of the cytokines on the growth of the tendon cell lines. The cells were cultured in 96-well plates in the presence or the absence of 5 ng/ml bFGF, 500 ng/ml BMP2, and 4 ng/ml TGF-␤. After 3 days cultured in the absence or presence of the cytokines, an MTT assay was conducted. The results are normalized against the value on Day 1. For each bar, 5 wells of the cells were used and the data represented one of the two experiments with similar results.

Cells were seeded onto two 96-well plates (normalized and test plates) at 5 ⫻ 103 cells/cm2 in 50 ␮l ␣-MEM supplemented with 0.5% FBS. After 24 h, 50 ␮l of fresh medium containing 500 ng/ml BMP2, 4 ng/ml TGF-␤, or 5 ng/ml bFGF was added to the wells and the cells were cultured for 3 days. The plates were subjected to MTT assay. Briefly, the cells were incubated in a solution containing 3-(4, 5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide for 3 h at 37°C. After the solution was removed, 50 ␮l of isopropanol was added and the plate was shaken vigorously for 30 s. The plate was then subjected to absorbance measurement using a microplate reader (BioRad Model 450) at 570-nm test wavelength and 630-nm reference wavelength [30]. RT-PCR analysis RT was carried out using 1 ␮g of total RNA, 0.2 ␮g of oligo(dT) primer, dNTP mix (10 mM/l), and MMLV-RT (100 units) in a final volume of 20 ␮l. RNA and primer were incubated together at 65°C for 10 min and then cooled rapidly on ice before addition of other reagents. RT was carried out at 37°C for 1 h. Complementary DNA was

Fig. 3. (A and B) Analysis of expression of the gene related to tendon phenotype. The tendon cell lines were cultured and RNAs were extracted as described under Materials and methods. For RT-PCR, RNAs were used to reverse transcribe to cDNA and primers described in Table 1 were used to generated the data (A). Northern blot of the expression of scleraxis and type I collagen in the three tendon-derived cell lines. RNA was extracted as described previously and the Northern blot was conducted using the cDNA probes for each of the genes (B).

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Fig. 4. (A–B) In ovo chorioallantoic membrane culture of the tendon cell lines. Tendon cell lines were seeded into the collagen sponge and then implanted into chorioallantoic membrane as described under Materials and methods (A–D). For control (E–H) sponge without cells was implanted into the chorioallantoic membranes. After the termination of in ovo culture the sections were made and they were stained for H&E (B and F), Van Gieson (C and G), and immunostained for large T (D and H).

amplified by PCR according to the “hot-start” method in a 25-␮l reaction volume containing 2.5 mM dNTP mix, 10 ␮M specific primers, and rTaq DNA polymerase (1 unit). After an initial denaturation at 94°C for 2 min in a Gene Amp PCR System 9700 (PE Biosystems), amplifications were performed at 94°C for 40 s, 60°C for 1 min, and 72°C for 1 min. Reaction products were analyzed by electrophoresis in 1.5% agarose gels containing ethidium bromide. The primers used for our RT-PCR experiments are listed in Table 1. RNA preparation and Northern blot analysis Total RNA was extracted from the confluent cells grown in a 56-cm2 dish, according to the acid guanidium thiocyanate–phenol– chloroform method [31]. Total cellular RNA

was subjected to fractionation in 1.0% agarose gels containing 0.66 M formaldehyde. The RNAs were then transferred to nylon filters (Genescreen, Life Science product) by electroblotting for 18 h [32]. The cDNA probes specific for osteopontin, type I collagen, alkaline phosphatase, osteocalcin, scleraxis, and Cbfal were labeled with [␣-32P]dCTP using a BcaBEST labeling kit (TAKARA). The profiles of cDNA probes are listed in Table 2. Filters were prehybridized overnight in a hybridization buffer containing 50% formaldehyde, 5x SSC, 5x Denhardt’s solution, 0.1% SDS, and 50 ␮g/ml sheared and denatured herring sperm DNA. Hybridization was performed in fresh hybridization solution containing labeled DNA probes at 42°C overnight and then the filters were washed three times in 1.0x SSC, 0.1% SDS at room temperature for 5 min and once in 0.2x SSC, 0.1% SDS at 60°C for 30 min. The filters were exposed to X-ray

Fig. 5. At the periphery of the implanted collagen sponge some fibrous tissues were also observed (A and B). These tissues were stained for H&E (A) and for the immunodetection using anti-large T antigen antibody (B).

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films (RX-U, Fuji medical X-ray film) using intensifying screens at ⫺80°C. In vitro differentiation assay The cells were cultured in either osteogenic differentiation medium (␣-MEM containing 10% FBS, 500 ng/ml rhBMP2, L-ascorbic acid, ␤-glycerophosphate) or adipogenic differentiation medium (␣-MEM containing 0.5% FBS, 10⫺7 M dexamethazone, 10 ␮g/ml insulin, IBMX for 2 days followed by ␣-MEM containing 0.5% FBS, 10 ␮g/ml insulin for an additional 3 weeks.) The chick chorioallantoic membrane (CAM) assay [33] was performed using 9-day-old chicken embryos incubated at 37°C. One small hole was made directly over the air sac in the shell with a needle to create a window. A 3-mm3 collagen sponge was preincubated with TT-D6 cells (107 cells in 100 ␮l of ␣-MEM) and was placed on the chorioallantoic membrane near large blood vessels. Holes were sealed with a piece of clear adhesive tape and incubation was carried out at 37°C for 10 days. The implants were harvested and fixed in 4% paraformaldehyde solution in PBS overnight before being subjected to histological examination. In vivo tendon defect model Adult GFP transgenic mice (kindly provided by Prof. M. Okabe at Osaka University) were anesthetized by intraperitoneal injection with pentobarbital. Patellar tendon defects were created by a transverse cut distal to the patella. Cell sheets were formed in culture dishes by prolonged cultures (14 days) of the over confluent cells; they were then scraped out and were put into the tendon defects. The same defect was produced on the contralateral side in the same mouse as a control, where no cell sheets were implanted. The specimens were harvested after 12 weeks. All animal experiments were approved by the Animal Welfare Committee of our institute. Histological and immunohistochemical studies Tissues were fixed overnight in 4% paraformaldehyde in PBS, decalcified, and embedded in paraffin according to standard procedure. Then, 8-␮m sections were made, dewaxed in xylene, and rehydrated through decreasing grades of alcohol in PBS. The sections were then subjected to stain with hematoxylin & eosin (H&E), toluidine blue, and/or Van Gieson staining. Immunohistological detection was performed by incubating the sections in a 0.3% H2O2 in PBS solution for 10 min and in a 5% normal goat serum in PBS for 30 min to quench endogenous peroxidases and to block nonspecific sites, respectively. Sections were incubated overnight at 4°C in a 1:100 dilution of the anti-large T antigen mouse monoclonal IgG (sc-147, Santa Cruz Biotechnology, Inc.) in 5% normal goat serum/PBS. As a neg-

ative control, PBS was used instead of first antibody. The sections were washed twice with PBS and incubated with anti-mouse biotin conjugated (Sigma) as a secondary antibody at dilution 1:200 in 5% normal goat serum/PBS at room temperature for 1 h. Excess antibody was then removed by three rinses with PBS before further detection with the ABC staining kit and DAB staining substrate. The sections were overlaid with Immuno-Mount. Statistical analysis Statistical significance of the data was evaluated by Fisher’s PLSD.

Results Three tendon cell lines (TT-E4, TT-G11, and TT-D6) were obtained using the SV40 large T antigen transgenic mice. These cells exhibited fibroblastic morphology (Figs. 1A–C). As mentioned under Materials and methods, the three tendon cell lines were established based on survival and proliferation under the 0.5% serum condition at 33°C for over a year. This condition was set to let the temperature mutant of large T antigen be in an active conformation at 33°C and to select the cells which can overcome the low serum concentration condition based on the activity of the large T antigen. Therefore, these cells may have lost their growth response to serum. To test this point, the effects of serum on the growth rate of these cell lines were examined. The number of cells cultured in medium containing different concentrations of serum (0.5, 1, and 10%) revealed dose-dependent serum enhancement on the growth of the three cell lines. The levels of growth responses to serum in three tendon cell lines were similar (Fig. 2A). The presence of receptors for cytokines was examined further. As a previous study reported that bFGF injected locally can increase the number of proliferating cells in the repairing tendon [10], we examined FGF response in these cells. Treatment with bFGF for 3 days significantly enhanced proliferation in all three tendon cell lines (Fig. 2B). This FGF effect was specific as BMP2 did not alter proliferation in these cells (Fig. 2B). Interestingly, TGF-␤ modestly but consistently enhanced proliferation in TT-E4 and TT-G11 but not in TT-D6 cells, revealing the individual difference among these cells (Fig. 2B). To characterize the tendon cell property, gene expression profile was examined in the three tendon cell lines by RT-PCR (Fig. 3A). These cell lines expressed tendon phenotype-related genes encoding scleraxis, six-1, COMP, EphA4, and type I collagen. In contrast, fibroblasts such as NIH3T3 cells and osteoblast-like MC3T3E1 cells did not express scleraxis and COMP genes although they expressed six-1 and EphA4 genes (Fig. 3A). Northern blot analysis also indicated the expression of scleraxis and type I collagen in the three tendon cell lines (Fig. 3B). Among the three cell

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Fig. 6. Formation of tendon-like fibrous tissues in the cell sheets cultured on chorioallantoic membrane (A–C). Cells were cultured to over confluence and then the cell sheets were detached, subjected to implantation, and cultured on the chorioallantoic membrane. After 10 days in culture on the chorioallantoic membrane the tissue was removed and subjected to histological examination. (A) H&E staining; (B) anti-large T antigen antibody staining; and (C) negative control without primary antibody.

lines, TT-D6 cells expressed highest levels of tendon phenotype-related genes and therefore was used for the subsequent characterization. To examine how these established tendon cells behave in vivo, the cells were implanted in ovo on CAM. For these

experiments, TT-D6 cells were seeded in collagen sponges and were placed on CAM. After 10 days of in ovo implantation, formation of dense connective tissue mass was observed in the implanted collagen sponges where the tendon cells were seeded (Figs. 4A–D), while no such tissue was

Fig. 7. Fibrous tissue formation within the implanted cell sheets in the patella tendon defect in mice (A–D). Cell sheets were formed in culture and implanted into the tendon defect produced in the knees of mice as described under Materials and methods. After 3 months, the sections were prepared and histology was made for each of the defected areas implanted with cell sheets (B and D) or without cells (A and C). The bracket in B indicates the site of implanted cell sheets. The asterisk in D indicates the area of ectopic bone formation in the patella tendon defect. (A and B) Van Gieson staining; (C and D) Toluidine blue staining.

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Fig. 8. Expression of genes related to the differentiated cells derived from mesenchymal tissues. RNA was extracted according to Materials and methods (A–C). Exceptional genes related to chondrocyte phenotype markers (A) and osteoblastic cell markers (B) were examined by RT-PCR. Northern blot analysis was conducting using Cbfa1 and OPN cDNA probes (C).

formed in the absence of cells (Figs. 4E–H). H&E staining indicated the presence of numerous nuclei in the interspace in the collagen sponge implanted with TT-D6 cells (Fig. 4B). These cells secreted extracellular matrix containing collagen which was stained red (Fig. 4C, asterisks) according to the Van Gieson technique, while the collagen sponge was stained a lighter red (Fig. 4C). The origin of these cells was examined by using anti-SV40 large T antigen antibody. In the sponge with TT-D6 cells, the sections were positive for large T antigen (Fig. 4D), indicating that they are derived from the implanted TT-D6 cells. No such signal was detected in the sponge without TT-D6 cells (Fig. 4H). In some cases, at the periphery of the implants, TT-D6 cells were found to form a fibrous connective tissue band wrapping around the collagen sponge, as indicated in Fig. 5A, and these cells were positive for large T antigen (Fig. 5B). These observations indicated that the established tendon cells could form a connective tissue mass in ovo. In ovo culture experiments using collagen sponges as scaffolds revealed that tendon cells can form a connective tissue mass; however, histological sections of such a mass did not indicate oriented fiber formation (Figs. 4B and C). This observation suggests that collagen sponges may not be an appropriate scaffold for tendon fiber formation. Instead, tendon cells may require scaffold of extracellular matrix protein produced by themselves. To address whether established tendon cells could form organized fibers if they were allowed to use their own extracellular matrix as scaffold, the tendon cells (TT-D6) were grown to over confluence in culture to form a cell sheet, which was then detached from the dish and implanted onto chorioallantoic membrane in ovo. After 10 days in ovo cultures of TT-D6 cell sheets on CAM, formation of fibers similar to those in tendon connective tissue was observed (Fig. 6A). The cells forming such fibers were positive for large T antigen (Fig. 6B). No staining was observed when only the secondary antibody was used without anti-large T antibody as shown in Fig. 6C. These observations indicate that TT-D6 cells exhibited a capability to form a tendon-like fiber when they were placed under an appropriate scaffold such as those with extracellular matrix produced by themselves. Although an in ovo culture system could provide an

environment closer to in vivo than in vitro in terms of the ECM, other environmental signals such as those from mature tendon cells or from the cells in adjacent tissues may be missing. Furthermore, tendon cells may need an environment under appropriate tensile stress. Therefore, we implanted TT-D6 cell sheets into the defects produced in the patella tendons in mice and examined the tissues 3 months after operation. Tendon defect implantation experiments demonstrated that TT-D6 cell sheets that were implanted into patellar tendon defects in adult mice form tendon tissues (Fig. 7B, bracket). Toluidine blue staining revealed that TT-D6 cell sheets formed tendon tissues; however, fibrocartilaginous tissues were also formed in part (Fig. 7D, asterisk). Interestingly, a large area of ectopic cartilage and bone formation was observed in the contralateral control patellar tendon defects where no cells were implanted (Figs. 7A and C). The formation of fibrocartilaginous tissues suggests that the established tendon cell lines may possess certain plasticity. As the presence of many tissue stem cells has been suggested, we examined whether the clonal tendon cells express, even at low levels, the genes related to tissues other than tendon. RT-PCR analyses revealed that the tendon cell lines expressed, though at low levels, genes related to chondrocyte phenotype including type X collagen (E4 and D6) and Sox9 (E4, G11, and D6) as shown in Fig. 8A. Type II collagen mRNA levels were very low but still detectable by RT-PCR analysis in E4, G11, and D6 (Fig. 8A). These observations were in accordance with the findings on the fibrocartilaginous tissue formation in part in the tendon cell sheet implantation experiments. As bone is closely related to cartilage and is also derived from similar mesenchymal cells, osteoblast phenotype-related gene expression was examined in the tendon cells by RT-PCR. The analyses revealed that these cells also expressed the genes related to osteoblastic phenotypes such as osteocalcin, osteopontin, and alkaline phosphatase (Fig. 8B). Northern blot analysis only detected the expression of Cbfal and osteopontin but not osteocalcin and alkaline phosphatase (Fig. 8C and unpublished data). Although the levels were low, expression of osteoblast-related genes could be

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Fig. 9. BMP2 effect on alkaline phosphatase activity in the tendon cell lines. (A) Alkaline phosphatase activity in the tendon cell lines was measured after being cultured in the presence or absence of 500 ng/ml BMP2 for 3 days. RNAs of the three cell lines were examined for the expression of alkaline phosphatase mRNA (B) and Osterix mRNA (C) after being cultured in the presence or absence of BMP2 for 3 days. The cell lines were subjected to osteogenic differentiation in the differentiation medium, the cultures were subjected to staining for calcified nodule formation using alizarin red, and the staining was compared to the culture in the absence of differentiation medium (D).

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Fig. 10. (A and B) Adipogenic differentiation potential in the tendon cell lines. RNAs of the tendon cell lines were subjected to RT-PCR analysis for adipocyte-related gene expression (A). After the cells were cultured in the presence of adipocyte differentiation medium and subjected to stain for lipid accumulation, some of the cultures were stained for lipid droplets by Oil red O (B).

the reflection of the capability of these cells to differentiate into osteoblasts. Since BMP2 is a potent inducer of osteoblastic differentiation [34 –37], we examined the effects of BMP2 on the tendon cell lines. TT-D6, TT-E4, and TT-G11 cells expressed alkaline phosphatase activity at very low levels (less than 0.01 ␮mol/min/mg protein). BMP2 treatment enhanced alkaline phosphatase activity in the three tendon cell lines (Fig. 9A). The enhancement of alkaline phosphatase activity was observed heterogeneously in the cells in culture when observed by alkaline phosphatase staining (data not shown). In contrast to its effect on alkaline phosphatase, BMP2 treatment did not significantly alter proliferation of these cells (Fig. 2B and data not shown). Gene expression analysis indicated that BMP treatment enhanced ALP and Osterix mRNA expression (Figs. 9B and C). Furthermore, as a consequence of these changes in gene expression, BMP treatment enhanced formation of calcified nodules stained positive for alizarin red in TT-G11 cells when they were cultured in osteogenic medium (Fig. 9D). As tendons are derived from mesenchymal origins, which also gives rise to cartilage, bone, and fat, we exam-

ined the expression of genes related to adipocyte phenotypes. In addition to the osteoblastic and chondrocytic features, these tendon-derived cell lines also expressed aP2 and PPAR-␥ genes which are expressed in adipocytes (Fig. 10A). When TT-E4 cells were cultured in adipogenic medium, these cells dramatically changed their morphology from fibroblastic-like spindle shape to round shape and some of them were observed to contain small lipid droplets in their cytoplasm, which stained positive for Oil Red O (Fig. 10B). However, neither TT-G11 nor TT-D6 showed adipogenic differentiation capability.

Discussion Novel tendon-derived cell lines were established and these cells express genes, such as scleraxis, six1, EphA4, and COMP, which are expressed in developing tendon during embryogenesis. TT-D6 cells demonstrated high expression levels of tendon lineage-related genes and could differentiate into tendon-like connective tissue when they were

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implanted both in ovo on CAM and in vivo in tendon defects. TT-D6 cells can differentiate into a dense connective tissue-like mass, which stained positive to reveal mature collagen when cultured on collagen sponge in ovo. The appearance of disorganized collagen fibril in cultured collagen sponge and the evidence of chondrogenesis in the early stage of the tendon repair model may be due to the lack of appropriate mechanical stimuli. In the early stage of tendon formation, extracellular tendon matrix and/or mechanical force might provide the regulatory signal directing the fate of tendon cells to form a mature collagen bundle in tendon. Lack of appropriate extracellular matrix and/or mechanical stress due to the condition of in ovo culture could be the reason for the lack of appearance of tendon-like fibers. In fact, in vivo tendon defect experiments revealed more organized fiber formation. TT-D6 cells showed a capability to differentiate into fibrocartilaginous tissue when implanted into patella tendon defects, demonstrating multipotential characteristics. However, these cells failed to form an aggregate mass in in vitro cartilaginous differentiation models (unpublished data). Even though some of them could form an aggregate pellet, they did not seem to produce any proteoglycan-rich extracellular matrix or exhibit any characteristics resembling that of cartilage-like tissue such as lacunae and proteoglycan synthesis. Moreover, these cells also expressed genes which are specific for the other tissues of mesenchymal lineage such as osteogenic, chondrogenic, and adipogenic cells when examined by RT-PCR. However, some of these genes cannot be detected by Northern blot analysis because of the low level of expression. Interestingly, adipogenic and osteogenic potentials were also observed in TT-E4 and TT-G11 cells, respectively, suggesting the capability to differentiate into multiple mesenchymal lineages of these cells. Stem cells can be broadly grouped into two categories based on their origin from either the embryo or the adult. Only the early embryos possess truly pluripotent cells that can give rise to all the cell types of embryos and adult tissues and, therefore, have the potential as a source of stem cells. In the adult tissues, on the other hand, specialized, tissue- or organ-specific stem cell types have been reported to exist to give rise to differentiated cell types. Our observation raises the possibility that tendon might contain stem cells that may generate diverse cell types during regeneration or repair of connective tissues. Although bone and cartilage could be observed in tendon under pathological conditions, it was not known whether the tendon cells contain pluripotent stem cells or whether stem cells are derived from surrounding connective tissue and might have migrated into tendon. Although tendon is a distinct tissue with specific features, molecular and cellular mechanisms underlying tendon cell development have not been fully understood. Our newly established cell lines provide a clue to elucidate the

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molecular mechanisms to explain the differentiation of this tissue.

Acknowledgments This research was supported by the Grants-in-Aid received from the Japanese Ministry of Education (15659352, 14207056, 14034214, 14028022), grants from Japan Space Forum, NASDA, Japan Society for Promotion of Science (JSPS, Research for the Future Program, Genome Science), and a JSPS research grant for foreign post-doctoral fellow.

References [1] T. Volk, Singling out Drosophila tendon cells: a dialogue between two distinct cell types, Trends Genet. 15 (1999) 448 – 453. [2] M. Kieny, A. Chevallier, Autonomy of tendon development in the embryonic chick wing, J. Embryol. Exp. Morphol. 49 (1979) 153– 165. [3] G. Kardon, Muscle and tendon morphogenesis in the avian hind limb, Development 125 (1998) 4019 – 4032. [4] H.J. Jacob, B. Christ, B. Brand, On the development of trunk and limb muscles in avian embryos, Bibl. Anat. (1986) 1–23. [5] C.C. Schmidt, H.I. Georgescu, C.K. Kwoh, G.L. Blomstrom, C.P. Engle, L.A. Larkin, C.H Evans, S.L. Woo, Effect of growth factors on the proliferation of fibroblasts from the medial collateral and anterior cruciate ligaments, J. Orthop. Res. 13 (1995) 184 –190. [6] A.K. Letson, L.E. Dahners, The effect of combinations of growth factors on ligament healing, Clin. Orthop. (1994) 207–212. [7] D. D’Souza, K. Patel, Involvement of long- and short-range signalling during early tendon development, Anat. Embryol. (Berl.) 200 (1999) 367–375. [8] J. Chang, D. Most, R. Thunder, B. Mehrara, M.T. Longaker, W.C. Lineaweaver, Molecular studies in flexor tendon wound healing: the role of basic fibroblast growth factor gene expression, J. Hand Surg. [Am.] 23 (1998) 1052–1058. [9] J. Chang, D. Most, E. Stelnicki, J.W. Siebert, M.T. Longaker, Hui, K. Lineaweaver, Gene expression of transforming growth factor beta-1 in rabbit zone II flexor tendon wound healing: evidence for dual mechanisms of repair, Plast. Reconstr. Surg. 100 (1997) 937–944. [10] F.J. Duffy Jr., R.H. Seiler, J.G. Gelberman, C.A. Hergrueter, Growth factors and canine flexor tendon healing: initial studies in uninjured and repair models, J. Hand Surg. [Am.] 20 (1995) 645– 649. [11] P. Rooney, D. Walker, M.E. Grant, J. McClure, Cartilage and bone formation in repairing Achilles tendons within diffusion chambers: evidence for tendon– cartilage and cartilage– bone conversion in vivo, J. Pathol. 169 (1993) 375–381. [12] J. McClure, The effect of diphosphonates on heterotopic ossification in regenerating Achilles tendon of the mouse, J. Pathol. 139 (1983) 419 – 430. [13] W.R. Atchley, W.M. Fitch, A natural classification of the basic helix–loop– helix class of transcription factors, Proc. Natl. Acad. Sci. USA 94 (1997) 5172–5176. [14] D. Brown, D. Wagner, X. Li, J.A. Richardson, E.N. Olson, Dual role of the basic helix–loop– helix transcription factor scleraxis in mesoderm formation and chondrogenesis during mouse embryogenesis, Development 126 (1999) 4317– 4329. [15] R. Schweitzer, J.H. Chyung, L.C. Murtaugh, A.E. Brent, V. Rosen, E.N. Olson, A. Lassar, C.J. Tabin, Analysis of the tendon cell fate using Scleraxis, a specific marker for tendons and ligaments, Development 128 (2001) 3855–3866.

300

R. Salingcarnboriboon et al. / Experimental Cell Research 287 (2003) 289 –300

[16] P. Cserjesi, D. Brown, K.L. Ligon, G.E. Lyons, N.G. Copeland, D.J. Gilbert, N.A. Jenkins, E.N. Olson, Scleraxis: a basic helix–loop– helix protein that prefigures skeletal formation during mouse embryogenesis, Development 121 (1995) 1099 –1110. [17] G. Oliver, R. Wehr, N.A. Jenkins, N.G. Copeland, B.N. Cheyette, V. Hartenstein, S.L. Zipursky, P. Gruss, Homeobox genes and connective tissue patterning, Development 121 (1995) 693–705. [18] K. Patel, R. Nittenberg, D. D’Souza, C. Irving, D. Burt, D.G. Wilkinson, C. Tickle, Expression and regulation of Cek-8, a cell to cell signalling receptor in developing chick limb buds, Development 122 (1996) 1147–1155. [19] K. Patel, H. Makarenkova, H.S. Jung, The role of long range, local and direct signalling molecules during chick feather bud development involving the BMPs, follistatin and the Eph receptor tyrosine kinase Eph-A4, Mech. Dev. 86 (1999) 51– 62. [20] P. DiCesare, N. Hauser, D. Lehman, S. Pasumarti, M. Paulsson, Cartilage oligomeric matrix protein (COMP) is an abundant component of tendon, FEBS Lett. 354 (1994) 237–240. [21] J.T. Hecht, M. Deere, E. Putnam, W. Cole, B. Vertel, H. Chen, J. Lawler, Characterization of cartilage oligomeric matrix protein (COMP) in human normal and pseudoachondroplasia musculoskeletal tissues, Matrix Biol. 17 (1998) 269 –278. [22] C. Fang, C.S. Carlson, M.P. Leslie, H. Tulli, E. Stolerman, R. Perris, L. Ni, P.E. Di Cesare, Molecular cloning, sequencing, and tissue and developmental expression of mouse cartilage oligomeric matrix protein (COMP), J. Orthop. Res. 18 (2000) 593– 603. [23] R.K. Smith, L. Zunino, P.M. Webbon, D. Heinegard, The distribution of cartilage oligomeric matrix protein (COMP) in tendon and its variation with tendon site, age and load, Matrix Biol. 16 (1997) 255–271. [24] A. Oldberg, P. Antonsson, K. Lindblom, D. Heinegard, COMP (cartilage oligomeric matrix protein) is structurally related to the thrombospondins, J. Biol. Chem. 267 (1992) 22346 –22350. [25] M. Obinata, Possible applications of conditionally immortalized tissue cell lines with differentiation functions, Biochem. Biophys. Res. Commun. 286 (2001) 667– 672. [26] H.N. Matsumoto, M. Tamura, D.T. Denhardt, M. Obinata, M. Noda, Establishment and characterization of bone marrow stromal cell lines that support osteoclastogenesis, Endocrinology 136 (1995) 4084 – 4091. [27] N. Mataga, M. Tamura, N. Yanai, T. Shinomura, K. Kimata, M. Obinata, M. Noda, Establishment of a novel chondrocyte-like cell line

[28]

[29]

[30]

[31]

[32]

[33]

[34]

[35]

[36]

[37]

derived from transgenic mice harboring the temperature-sensitive simian virus 40 large T-antigen gene, J. Bone Miner. Res. 11 (1996) 1646 –1654. D. Feuerbach, E. Loetscher, K. Buerki, T.K. Sampath, J.H. Feyen, Establishment and characterization of conditionally immortalized stromal cell lines from a temperature-sensitive T-Ag transgenic mouse, J. Bone Miner. Res. 12 (1997) 179 –190. K. Wewetzer, B. Seilheimer, Establishment of a single-step hybridoma cloning protocol using an automated cell transfer system: comparison with limiting dilution, J. Immunol. Methods 179 (1995) 71–76. F. Denizot, R. Lang, Rapid colorimetric assay for cell growth and survival. Modifications to the tetrazolium dye procedure giving improved sensitivity and reliability, J. Immunol. Methods 89 (1986) 271–277. P. Chomczynski, N. Sacchi, Single-step method of RNA isolation by acid guanidinium thiocyanate–phenol– chloroform extraction, Anal. Biochem. 162 (1987) 156 –159. P.S. Thomas, Hybridization of denatured RNA and small DNA fragments transferred to nitrocellulose, Proc. Natl. Acad. Sci. USA 77 (1980) 5201–5205. C. Scher, C. Haudenschild, M. Klagsbrun, The chick chorioallantoic membrane as a model system for the study of tissue invasion by viral transformed cells, Cell 8 (1976) 373–382. T. Katagiri, A. Yamaguchi, M. Komaki, E. Abe, N. Takahashi, T. Ikeda, V. Rosen, J.M. Wozney, Fujisawa-Sehara, A. Suda, T. Bone morphogenetic protein-2 converts the differentiation pathway of C2C12 myoblasts into the osteoblast lineage, J. Cell Biol. 127 (1994) 1755–1766. R.S. Thies, M. Bauduy, B.A. Ashton, L. Kurtzberg, J.M. Wozney, V. Rosen, Recombinant human bone morphogenetic protein-2 induces osteoblastic differentiation in W-20-17 stromal cells, Endocrinology 130 (1992) 1318 –1324. J.M. Wozney, V. Rosen, Bone morphogenetic protein and bone morphogenetic protein gene family in bone formation and repair, Clin. Orthop. 346 (1998) 26 –37. A. Yamaguchi, T. Katagiri, T. Ikeda, J.M. Wozney, V. Rosen, E.A. Wang, A.J. Kahn, T. Suda, S. Yoshiki, Recombinant human bone morphogenetic protein-2 stimulates osteoblastic maturation and inhibits myogenic differentiation in vitro, J. Cell Biol. 113 (1991) 681– 687.