Microenvironment and phenotypic stability specify tissue formation by human articular cartilage-derived cells in vivo

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Experimental Cell Research 287 (2003) 16 –27

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Microenvironment and phenotypic stability specify tissue formation by human articular cartilage-derived cells in vivo Francesco Dell’Accio, Cosimo De Bari, and Frank P. Luyten* Laboratory for Skeletal Development and Joint Disorders, Department of Rheumatology, University Hospitals, Katholieke Universiteit, Leuven, Belgium Received 5 September 2002, revised version received 28 November 2002

Abstract During in vitro expansion, adult human articular cartilage-derived cells (HACDC) lose their phenotypic stability and capacity to form cartilage in vivo after 4 – 6 population doublings (PD). Nevertheless, HACDC can be efficiently expanded for up to 20 PD. Here we show that HACDC can generate cartilage, fibrous tissue, skeletal muscle, bone, and adipocytes depending on the balance between phenotypic stability and environmental cues. When 5 ⫻ 106 cells were injected intramuscularly into nude mice, early-passage (EP)-HACDC formed cartilage; late-passage (LP)-HACDC formed mostly fibrous tissue, but a limited number of cells contributed to muscle formation. When 0.5 ⫻ 106 cells were injected into regenerating mouse muscle, both EP- and LP-HACDC integrated with host myofibers and expressed muscle genes, but a number of EP-HACDC maintained collagen type II expression. HACDC seeded into Collagraft and implanted subcutaneously into nude mice formed scattered bone islands displaying immunoreactivity for human osteocalcin, and expressing human bone-specific genes. Importantly, neither collagen type II transcript nor cartilage tissue was detected at 8 weeks after implantation. Myogenic, osteogenic, and adipogenic differentiation was induced in vitro using specific culture conditions. These findings provide evidence that in vivo tissue formation by HACDC is specified by a balance between environmental cues and the inherent phenotypic stability. © 2003 Elsevier Science (USA). All rights reserved. Keywords: Cartilage, articular; Cell differentiation; Cells, cultured; Osteoarthritis; Chondrocytes; Cartilage repair; Stem cells; Tissue engineering; Myogenesis; Osteogenesis

Introduction Adult articular cartilage is an avascular tissue presumed to be composed by a single type of cells, the articular chondrocytes, which are dispersed in an abundant extracellular matrix, rich in proteoglycans and collagen type II [1]. Articular cartilage is traditionally considered a terminally differentiated tissue with limited capacity for repair. In vitro expanded autologous chondrocytes are currently used to promote the healing of symptomatic joint surface defects in human patients [2]. The capacity of the expanded human articular cartilage-derived cells (HACDC)1 to contribute to

* Corresponding author. Laboratory for Skeletal Development and Joint Disorders, Onderwijs & Navorsing, Herestraat 49, 3000 Leuven, Belgium. Fax: ⫹32-16-346200. E-mail address: [email protected] (F.P. Luyten). 1 Abbreviations used: HACDC, human articular cartilage-derived cells; EP, early-passage; LP, late passage; PD, population doublings; CFA, cartilage formation assay; PBS, phosphate-buffered saline; ISH, in situ

the regeneration of stable cartilage tissue is therefore of paramount importance in cartilage tissue engineering [3,4]. We have previously shown that HACDC progressively lose, throughout in vitro expansion, their capacity to form stable cartilage in vivo in a standardized cartilage formation assay (CFA) [3]. The intramuscular injection of 5 ⫻ 106 primary HACDC into the thighs of nude mice reproducibly forms a stable cartilage implant in this assay within 14 days after injection. This cartilage formation capacity is lost with cell expansion after 4, occasionally 6, population doublings

hybridization; SQ-RT-PCR, semiquantitative reverse transcription polymerase chain reaction; TA, tibialis anterior; DMEM, Dulbecco’s modified Eagle medium; FBS, fetal bovine serum; RT, reverse transcription; COL2A1, collagen type II ␣1 chain; MyHC-IIx/d, myosin heavy chain type IIx/d; MA, myogenesis assay; CK, creatine kinase; FI, freshly isolated; DAPI, 4,6-Diamidino-2-phenylindole; OC, osteocalcin; OP, osteopontin; BSP, bone sialoprotein; COL10A1, collagen type X ␣1 chain; PPAR␥-2, peroxisome proliferator-activated receptor ␥2; aP2, Fatty acidbinding protein aP2.

0014-4827/03/$ – see front matter © 2003 Elsevier Science (USA). All rights reserved. doi:10.1016/S0014-4827(03)00036-3

F. Dell’Accio et al. / Experimental Cell Research 287 (2003) 16 –27

(PD), and is dependent on cell culture conditions [3]. No implant was macroscopically retrieved when late-passage HACDC were injected [3]. Despite the loss of the spontaneous cartilage-forming ability in vivo, dedifferentiated HACDC can be expanded in vitro for up to 20 PD with a roughly linear growth curve, and can be at least partially recovered to the cartilage phenotype when cultured in lowmelting-temperature agarose [3,5] suggesting that some expanded cells are still capable of differentiation. In this study we explored the biological behavior of expanded human articular cartilage cell populations in response to environmental cues.

Materials and methods Cartilage harvest and chondrocyte isolation Normal articular cartilage was obtained from the femoral condyles of human donors within 12 h postmortem. Cartilage was sliced full thickness, excluding the external borders of the condyles, the mineralized cartilage, and the subchondral bone to avoid synovial and bone marrow contamination. Release from articular cartilage and in vitro culture of HACDC was performed as previously described [3]. Routinely, cultures were passaged at a 1:4 dilution (about 2 PD per passage). We call hereafter “early passage” (EP), HACDC that have been expanded for no longer than 4 PD, and “late passage” (LP), HACDC that have been expanded in vitro for more than 10 PD.

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In vivo assay for myogenic differentiation Twenty-five microliters of 10 ␮M cardiotoxin (Latoxan, France) was injected into the tibialis anterior (TA) muscle of NMRI nu⫺/⫺ mice. Twenty-four hours later, 0.5 ⫻ 106 viable HACDC suspended in 25 ␮l PBS were transplanted into the same TA muscle as described with bone marrowderived cells [6]. The mice were sacrificed 4 weeks after HACDC implantation, unless stated otherwise. In vitro myogenesis assay The in vitro myogenesis assay was performed as described elsewhere [7]. Briefly, HACDC were seeded into 35-mm dishes at a density of 5000 cells/well in growth medium: high-glucose Dulbecco’s modified Eagle medium (DMEM, Life Technologies, Merelbeke, Belgium) containing 10% fetal bovine serum (FBS, BioWhittaker, Verviers, Belgium), 200 units/ml penicillin, 200 ␮g/ml of streptomycin, and 0.5 ␮g/ml of amphotericin B (Life Technologies). Beginning 24 h after seeding, cultures were treated for 24 h with 10 ␮M 5-azacytidine (Sigma, Bornem, Belgium). Cultures were then washed twice with HBSS, and the medium was changed either to growth medium or to myogenic medium (growth medium supplemented with 5% horse serum, 50 ␮M hydrocortisone (Sigma), and 4 ng/ml basic fibroblast growth factor (a kind gift of TK Sampath, Creative BioMolecules, Hopkinton, MA, USA)). The medium was changed twice a week for 4 weeks, until the experiment was terminated. The experiment was carried out in quadruplicate. Immunostaining for skeletal muscle-specific myosin heavy chain was performed as described elsewhere [8].

Animal care and maintenance

In vivo osteogenesis assay

All the procedures on animals were approved by the local ethical committee. Eight-week-old, female NMRI nu⫺/⫺ nude mice were maintained in isolator cages in pathogen-free conditions until the end of the experiment.

HACDC were seeded into Collagraft matrix (NeuColl, Campbell, CA, USA) and implanted subcutaneously into nude mice as described with bone marrow stromal cells [9]. Briefly, 3-mm3 Collagraft cubes were wetted in DMEM, blotted onto a gauze compress, and immersed into 50 ␮l of HACDC suspension (105 cells/␮l in growth medium) for 1 h at 37°C to allow attachment of the cells to the Collagraft matrix. The percentage of attaching cells varied routinely between 45 and 60%. The cell–Collagraft constructs were implanted subcutaneously into nude mice. At different time points (between 4 and 12 weeks) the mice were killed and the constructs dissected. The explants were then cut in 2, and one half was destined to RNA extraction, and the other fixed overnight in 4% formaldehyde. Fixed samples were decalcified overnight in Decal (Serva, Heidelberg, Germany) and paraffin-embedded. Hematoxylin– eosin and Masson’s trichrome stainings were performed according to standard protocols. Immunostaining for human osteocalcin was carried out as follows. Eight-micrometer-thick sections were deparaffinized and rehydrated in an ethanol series. After washing in PBS, nonspecific binding was blocked with 1% Blocking

In vivo assay for stable cartilage formation The capacity to form stable cartilage in vivo was assessed as described elsewhere [3]. Briefly, 5 ⫻ 106 cells were resuspended in 50 ␮l phosphate-buffered saline (PBS) and injected intramuscularly into the posterior compartment of the thighs of nude mice. After 2 weeks, unless stated otherwise, the injected muscles were dissected and either fixed in 4% paraphormaldehyde and paraffin-embedded for histology, histochemistry, or in situ hybridization (ISH), or used for gene expression analysis by semiquantitative reverse transcription polymerase chain reaction (SQ-RTPCR). The immunodetection of collagen type I was performed as described below for ostocalcin using a monoclonal antibody antihuman collagen type I (Chemicon).

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Reagent (Roche Molecular Biochemicals, Brussels, Belgium) in PBS supplemented with 0.2% (v/v) Triton X-100. Slides were incubated 1 h at room temperature with a rabbit anti-human osteocalcin antibody (Chemicon, Hofheim, Germany) diluted 1:800 in 1% Blocking Reagent in PBS supplemented with 0.2% Triton X-100. After extensive washing in PBS– 0.2% Triton X-100, slides were incubated 1 h in a goat anti-rabbit antibody conjugated with alkaline phosphatase by a dextran backbone (EnVision Kit, Dako Diagnostics, Heverlee, Belgium), washed extensively in PBS– 0.2% Triton X-100, and stained using the DAKO fuchsin ⫹ substrate system (Dako Diagnostics) in the presence of 0.2 mM levamisole (Sigma) to inhibit endogenous alkaline phosphatase activity. Sections were washed in water, counterstained with hematoxylin, and mounted with Mowiol (Calbiochem Merck Belgolabo, Overijse, Belgium). In vitro osteogenesis assay Osteogenesis was induced in vitro as described previously for synovial membrane-derived mesenchymal stem cells [8]. All experiments were performed in quadruplicate. The cultures were either stained for the presence of calcium deposits by alizarin red S staining or destined to RNA extraction and gene expression analysis. In vitro adipogenesis assay The in vitro adipogenesis assay was performed as previously described for synovial membrane-derived mesenchymal stem cells [8]. Briefly, confluent monolayer cultures were primed to adipogenesis by the addition of adipogenic induction medium, consisting of growth medium supplemented with 1 ␮M dexamethasone (Sigma), 0.5 mM methyl-isobutylxanthine (Sigma), 10 ␮g/ml insulin (Sigma), and 100 mM indomethacin (Sigma). After 72 h, the medium was changed to adipogenic maintenance medium consisting of 10 ␮g/ml insulin in growth medium for 24 h. Cells were treated with adipogenic induction medium four times. The cultures were subsequently kept in adipogenic maintenance medium. Controls for the adipogenic treatment were cells cultured in growth medium supplemented with vehicle solutions of the compounds of the adipogenic media. Experiments were performed in quadruplicate. After 3 weeks, cells were either stained for lipids with oil red O or destined to RNA extraction for gene expression analysis.

were analyzed by gel electrophoresis, stained with ethidium bromide, and visualized by UV transillumination. Gene expression of human cells within mouse tissue was evaluated using primers that specifically recognize human genes. We used the PCR conditions that allowed efficient amplification at the highest stringency. In experiments involving the detection of human cDNAs within mouse tissues injected with HACDC, uninjected mouse tissues were used to control the species specificity of the primers. cDNAs from uninjected mouse tissues were equalized to the sample with the highest ␤-actin expression within the experiment using primers that amplify both human and mouse ␤-actin. In situ hybridization for human-specific Alu genomic repeats ISH for human specific Alu repeats was performed as described elsewhere [9] with a few modifications. Briefly, sections were deparaffinized and rehydrated. Matrix digestion was obtained by enzymatic treatment with 10 ␮g/ml proteinase K in 0.1 M Tris–HCl, 50 mM EDTA, pH 8, for 30 min at 37°C. Sections were immediately postfixed in 3% formaldehyde for 10 min, washed in PBS, dehydrated in an ethanol series, and dried for 20 min at room temperature. After rehydration and equilibration in PBS, sections were acetylated in 0.25% acetic acid containing 0.1 M triethanolamine (pH 8) for 10 min, and prehybridized with 50% deionized formamide containing 4x SSC at 37°C for 15 min. Sections were covered with hybridization buffer (1x Denhardt’s solution, 0.2 mg/ml denatured sheared salmon sperm DNA, 4x SSC, and 50% deionized formamide) containing 1 ng/␮l digoxigenin-labeled double-stranded DNA probe specific for human Alu genomic repeats and covered with a glass coverslip. Denaturation of both the probe and the genomic DNA template was achieved by heating the slides at 95°C for 45 s. Hybridization was performed at 42°C overnight. Sections were washed twice for 30 min at room temperature with 2x SSC and 0.1x SSC and for 30 min at 50°C in 0.1x SSC. Digoxigenin was detected using a commercially available kit (DIG Nucleic Acid Detection Kit, Roche Molecular Biochemicals, Brussels, Belgium) according to the manufacturer’s protocol. DIG-labeled human-specific Alu probe was generated as described elsewhere [9].

Results Total RNA extraction and SQ-RT-PCR analysis Total RNA was extracted using TRIzol reagent (Life Technologies) according to the manufacturer’s instructions. cDNAs were obtained by reverse transcription (RT) of 1 ␮g of total RNA (Thermoscript, Life Technologies) with oligo(dT)20 primer. PCRs were carried out in a Perkin Elmer Thermal Cycler 9600 (Applied Biosystems, Lennik Belgium). The sequences of the primers and the expected sizes of the PCR products are listed in Table 1. PCR products

Persistence and phenotype of LP-HACDC in the cartilage formation assay in vivo We have previously shown that HACDC lose their capacity to spontaneously form cartilage in the CFA when expanded in vitro for more than 4 – 6 PD. No implant was macroscopically retrieved when dissecting LP-HACDC-injected mouse muscles [3]. To localize and subsequently characterize LP-HACDC within the injected muscle, we

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Table 1 Primers used for SQ-RT-PCR analysisa Gene

Primer sequence

Amplicon size (bp)

h-␤ actin

Forward 5⬘-CCGACAGGATGCAGAAGGAG-3⬘ Reverse 5⬘-GGCACGAAGGCTCATCATTC-3⬘ Forward 5⬘-TGACGGGGTCACCCACACTGTGCCCATCTA-3⬘ Reverse 5⬘-CTAGAAGCATTTGCGGTGGACGATGGAGGG-3⬘ Forward 5⬘-CTGCTCGTCGCCGCTGTCCTT-3⬘ Reverse 5⬘-AAGGGTCCCAGGTTCTCCATC-3⬘ Forward 5⬘-CATTGTTGGTCTGCCTGGTC-3⬘ Reverse 5⬘-TTCTCTCTCTGCCCTAAGCC-3⬘ Forward 5⬘-ATAGGAACACCCAAGCCATC-3⬘ Reverse 5⬘-TTTGCGTAGACCCTTGACAG-3⬘ Forward 5⬘-GGCACAATGACAACAAGAGC-3⬘ Reverse 5⬘-GAAAAGAAGAGGACCCTGCC-3⬘ Forward 5⬘-CGATTTCCAGTTCAGGGCAG-3⬘ Reverse 5⬘-TCCATTGTCTCCTCCGCTGC-3⬘ Forward 5⬘-TCACACTCCTCGCCCTATTG-3⬘ Reverse 5⬘-GAAGAGGAAAGAAGGGTGCC-3⬘ Forward 5⬘-GCTAAACCCTGACCCATCTC-3⬘ Reverse 5⬘-ATAACTGTCCTTCCCACGGC-3⬘ Forward 5⬘-AATCCCTGGACCGGCTGGAATTTC-3⬘ Reverse 5⬘-TTGATGCCTGGCTGTCCTGGAACC-3⬘ Forward 5⬘-AGGACTCAGGGTGGTTCAGC-3⬘ Reverse 5⬘-AGGAGCAGAGCAAAGAGGTG-3⬘ Forward 5⬘-TATGAAAGAAGTAGGAGTGGGC-3⬘ Reverse 5⬘-CCACCACCAGTTTATCATCCTC-3⬘ Forward 5⬘-CGTGGTGACAAGGGTGAGAC-3⬘ Reverse 5⬘-TAGGTGATGTTCTGGGAGGC-3⬘

662

mh-␤ actin h-COL2A1 m-Col2A1 h-MyHCIIx/d h-MCK h-BSP h-OC h-OP h-COL10A1 PPAR␥-2 h-aP2 Collagen type I

661 432 collagen type IIA 225 collagen type IIB 750 599 721 426 362 547 267 474 290 827

a The prefix “h-” means that the primer set allows specific amplification of the human cDNA. The prefix “m-” means that the primer set allows specific amplification of the mouse cDNA. The prefix “mh-” means that the primer set does not allow discrimination between mouse and human cDNA.

performed ISH for human-specific Alu genomic repeats at 2 and 8 weeks after implantation of LP-HACDC (16 PD). In both cases, human nuclei were mostly clustered in large cell aggregates (Fig. 1A). These cell aggregates formed fibrous tissue, according to the histological appearance (Fig. 1B) and type I collagen immunoreactivity (Fig. 1C). Interestingly, some sporadic human nuclei were scattered within the muscle tissue, suggesting their possible integration into host myofibers (Figs. 1D, E). Our in vivo assay can be considered a chimeric experiment where human cells have been injected into a mouse host. This offers the opportunity to selectively monitor gene expression of human cells within the mouse tissue by SQRT-PCR, using human-specific primers. In Fig. 1F we show that, under our experimental conditions, the implants obtained by injecting EP-HACDC expressed human collagen type IIA and IIB (COL2A1), confirming a mature cartilage phenotype [10]. COL2A1 was not detected in the LPHACDC-derived implants and in monolayer culture, confirming the loss of their mature chondrocytic phenotype on serial passaging and the inability of LP-HACDC to spontaneously form stable cartilage in vivo. Surprisingly, human myosin heavy chain type IIx/d (MyHC-IIx/d) was detected in the muscles injected with LP-HACDC, indicating that at least some human cells had acquired a muscle phenotype. Under our experimental conditions, MyHC-IIx/d was detected neither in LP-HACDC monolayers nor in the muscle injected with 5 ⫻ 106 EP-HACDC. Taken together, these

data indicate that LP-HACDC persisted within the mouse muscle, generating mostly fibrous tissue. A subset of cells appeared to differentiate into skeletal muscle. Characterization of the myogenic potential of HACDC To study the myogenic potential of HACDC, we have employed a well-established in vivo myogenesis assay (MA) [6]. This assay consists of injecting 0.5 ⫻ 106 cells into the regenerating TA muscle of nude mice. To induce muscle regeneration, the muscle was damaged 24 hours earlier by a local injection of the snake venom cardiotoxin. We assessed the myogenic potential of LP-HACDC in the MA. At the same time, we investigated whether EPHACDC, which form stable cartilage in the CFA, could be induced to differentiate into muscle in this strongly myogenic microenvironment [6]. At 3 weeks after injection, skeletal muscle-specific human genes, such as MyHC-IIx/d and creatine kinase (CK), were detected by SQ-RT-PCR in the regenerating muscles injected with either EP- or LPHACDC (Fig. 2A), indicating muscle differentiation of the injected human cells. In the regenerating muscles injected with EP-HACDC, human COL2A1 expression was maintained for at least 4 weeks in all the samples tested. Despite the maintenance of COL2A1 expression, we did not detect any cartilage tissue when TA muscles were serially sectioned and stained with toluidine blue at 2 and 4 weeks after the implantation of EP-HACDC. The presence of human

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Fig. 1. Persistence and phenotypic characterization of late-passage adult human articular cartilage-derived cells (LP-HACDC) in a cartilage formation assay (CFA) in vivo. Five million LP-HACDC were injected intramuscularly into the thighs of nude mice in a well-standardized CFA. (A) In situ hybridization for human-specific Alu genomic repeats showing the persistence and localization of the human LP-HACDC within the mouse muscle at 4 weeks after implantation. (B) Detail of a parallel, nonconsecutive section corresponding to the squared area in (A), stained with hematoxylin and eosin. Areas occupied by LP-HACDC generated a fibrous-like tissue. (C) Immunohistochemistry for human collagen type I in another parallel, nonconsecutive section, showing intense positivity in the areas occupied by most human cells in (A). (D) Detail of (A). In areas where the human cells were not clustered, the position of some human nuclei (arrow) suggests their possible integration into host myofibers. (E) False-colored image of the brightfield shown in (D) superimposed on the fluorescence image of the same section showing the DAPI nuclear counterstaining. ALU-positive (human) nuclei are displayed in red; Alu-negative, DAPI-stained nuclei are green. (F) EP- and LP-HACDC were injected into the CFA 2 weeks after implantation and the HACDC-injected muscles were dissected and subjected to gene expression analysis by SQ-RT-PCR. Lane 1 (m-SkM): uninjected mouse muscle. Lane 2 (m-joint): cDNA from a mouse joint containing bone and cartilage tissues. Lane 3 (EP implant): mouse muscle injected with primary confluent EP-HACDC. Lane 4 (LP monolayer): cDNA from LP-HACDC—about 16 population doublings (PD)—in monolayer. Lane 5 (LP implant): mouse muscle injected with the LP-HACDC as in lane 4. Lane 6 (W): water control. All the samples containing human cells were normalized for expression of the housekeeping gene ␤-actin using human-specific primers. The presence of template cDNA in the samples that did not contain human cells is documented by the expression of ␤-actin using primers that detect both human and mouse ␤-actin (mh-␤actin). Human collagen types IIA and IIB (COL2A1) were detected only in muscle injected with EP-HACDC, indicating the presence of mature chondrocytes. The size of the amplification product is 432 bp for collagen type IIA and 225 bp for collagen type IIB. Human adult skeletal muscle myosin heavy chain type II x/d (MyHC-IIx/d) was detected in the muscle injected with LP-HACDC, indicating skeletal muscle differentiation of the human cells. Mouse collagen type II was present only in the mouse joint positive control, confirming the species specificity of the COL2A1 detection, and indicating that mouse cells did not contribute significantly to cartilage formation when EP-HACDC were injected. Bars ⫽ 800 ␮m (A, C), 100 ␮m (D, E), and 50 ␮m (B).

cells in these sections was confirmed by ISH for humanspecific Alu repeats (not shown). Skeletal muscle consists of multinucleated myofibers that, on injury, regenerate by recruiting mononucleated satellite cells [11]. To investigate whether HACDC fused with host myofibers in the MA, we stained sections from mouse TA at 2 weeks after HACDC transplantation for human ␤2-microglobulin. In Fig. 2B we show that ␤2-microglobulin was detected on isolated cells, as well as on some muscle fibers, documenting fusion of the human cells with the positive myofibers. Centronucleation is a feature of myofiber regeneration [12]. We observed sporadic muscle fibers with centrally located human nuclei (Fig. 2C, D), confirming the participation of some human cells in the myofiber

regeneration process. Taken together, these results indicate that, under conditions strongly promoting myogenesis, both EP- and LP-HACDC can be recruited to skeletal muscle repair, fuse with host myofibers, and express muscle-specific genes. The capacity of in vitro-expanded HACDC to reprogram their phenotype could result from an at least partial “dedifferentiation” process associated with the monolayer culture. To investigate whether the de-differentiation associated with in vitro culture in monolayer is required for myogenic differentiation of HACDC, we injected in the MA freshly isolated (FI) HACDC from two donors (44 and 72 years old), immediately after enzymatic release from the cartilage tissue. As shown in Fig. 2E, human MyHC-IIx/d

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Fig. 2. In vivo myogenic potential of HACDC. Muscle damage was induced in the tibialis anterior (TA) muscle of nude mice by local injection of cardiotoxin. Twenty-four hours later, 5 ⫻ 105 EP- or LP-HACDC were injected into the regenerating TA muscle. (A) SQ-RT-PCR analysis at 4 weeks after HACDC injection. Lane 1 (m-TA): cardiotoxin-treated, uninjected mouse TA muscle. Lane 2 (h-SkM): human muscle. Lane 3: monolayer (M) culture of primary confluent (2 PD) EP-HACDC. Lane 4: mouse TA muscle treated with cardiotoxin and injected (I) 24 h later with the EP-HACDC as in lane 3. Lane 5,

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was detected by SQ-RT-PCR in the injected muscle at 4 weeks after implantation, indicating muscle differentiation. MyHC-IIx/d was not detectable in FI-HACDC before injection. As with EP-HACDC, human COL2A1 expression was maintained after the injection in the regenerating mouse TA muscles, indicating the persistence of phenotypic characteristics of the original cell population. These data show that growth and expansion in monolayer culture are not required for myogenic differentiation of HACDC. It has been reported that the muscle phenotype is dominant when a nonmuscle cell (including a chondrocyte) is experimentally fused to a myotube or a myoblast [13]. To investigate whether the fusion with preexisting myofibers is required for nuclear reprogramming, we have challenged EP- and LP-HACDC in an in vitro assay for myogenic differentiation [7]. MyHC-IIx/d mRNA was detected by SQ-RT-PCR in the myogenic cultures, but not in the control cultures (Fig. 2F). In contrast to C2C12 myoblast cultures, used as a positive control, no typical myosin heavy chain-positive multinucleated myotubes were documented. Immunoreactivity was observed in scattered mononucleated cells (data not shown). Taken together, these data indicate that the cytoplasm of muscle fibers is not required for muscle gene expression by HACDC. Osteogenic differentiation The possibility of inducing myogenic differentiation prompted us to investigate whether HACDC could differentiate also to other mesenchymal lineages. Osteogenic culture conditions induced calcium deposition in both EP- and LP-HACDC monolayer cultures as evaluated by alizarin red staining (Fig. 3A), and upregulation of the bone markers osteopontin, osteocalcin, and bone sialoprotein as evaluated by SQ-RT-PCR (data not shown). Calcium deposits were detected neither in control HACDC cultures, nor in skin fibroblasts, either treated or not, which were used as cell negative controls (not shown). To investigate whether HACDC can form bone in vivo,

we adopted an assay for bone formation validated with bone marrow cells [9]. HACDC were seeded into Collagraft matrix, and the cell–Collagraft constructs were implanted subcutaneously into nude mice. As shown in Fig. 3, at 8 weeks after implantation, scattered areas of bone were observed after Masson’s trichrome or hematoxylin– eosin staining. Most of the areas of the sections consisted of fibrous and osteoid tissues. Similar results were obtained with EP- and LP-HACDC. To prove the human origin of the bone tissue, we performed an immunostaining using an antibody specific to human osteocalcin. Discrete areas, which morphologically appeared as bone, stained positive for human osteocalcin (Fig. 3D). Mouse bone was used as a control for human specificity of the antibody (Fig. 3F). We also detected human nuclei within the bone areas by ISH for human Alu genomic repeats (not shown), demonstrating that these bone tissue areas were contributed, at least in part, by human cells. To examine the molecular dynamics during bone formation in vivo, we analyzed gene expression by SQ-RT-PCR using human-specific primers (Fig. 4A). The bone markers osteopontin, osteocalcin, and bone sialoprotein, absent in the monolayer cultures, were progressively upregulated in the HACDC–Collagraft implants. Similar dynamics were observed using either primary or late-passage HACDC. The expression of the cartilage marker COL2A1, present in the primary HACDC monolayers, progressively decreased at 4 and 8 weeks. No COL2A1 was detected in LP-HACDC at any time before or after implantation. The expression of collagen type X never exceeded the base levels of EPHACDC in monolayer culture. We tested reproducibility using HACDC from donors of various ages (range: 25–77 years) in this bone formation assay at 4 weeks (when available) and at 8 weeks after implantation. Under our experimental conditions, the upregulation of bone markers was reproducible and consistent, regardless of donor age or cell passage number, with some variability at the 4-week time point as shown in Fig. 4B. To investigate to which extent the environment could

LP-HACDC (12 PD) in monolayer. Lane 6: mouse TA muscle treated with cardiotoxin and injected 24 h later with LP-HACDC. Lane 7: water control. All the samples containing human cells were normalized for human-specific ␤-actin. The uninjected mouse muscle was normalized to the sample with the highest ␤-actin (mouse and human) and was included as a control for species specificity of PCR amplification. Human MyHC-IIx/d and human muscle creatine kinase (MCK) were detected in the TA muscles injected with either EP- or LP-HACDC, indicating skeletal muscle differentiation. Human MyHC-IIx/d and h-MCK were not detected in the monolayer cultures. EP-HACDC maintained detectable levels of collagen type IIA and type IIB also after injection into the MA. (B) Immunostaining for human ␤2-microglobulin on a section of a regenerating mouse TA 4 weeks after EP-HACDC injection. Several myofibers were positive (green) indicating fusion of at least one human cell into each positive myofiber. Bar ⫽ 50 ␮m. (C) Brightfield image of an ISH for human Alu repeats showing a human nucleus centrally located in a myofiber 4 weeks after the injection of LP-HACDC. (D) False-colored image of the brightfield shown in (C) superimposed on the fluorescent image of the same section showing DAPI nuclear counterstaining. The Alu-positive (human) nucleus is displayed in red, and the DAPI-positive/Alu-negative nuclei in green. Bar ⫽ 50 ␮m. (E) To test whether in vitro expansion is required for muscle differentiation, freshly isolated (FI)-HACDC were challenged in the MA. SQ-RT-PCR analysis showed detectable human-specific MyHC-IIx/d in the mouse TA injected with FI-HACDC (lane 3, I-m-TA). Collagen type IIA and IIB expression was maintained after the injection at levels comparable to that of FI-HACDC before injection (lane 2). Lane 1 (m-TA): cardiotoxin-treated, uninjected mouse TA muscle was used as control for primers specificity to human genes. Lane 4 (w): water control. Lane 5 (RT⫺): RT-negative control of lane 3, used to exclude genomic contamination. Lane 6 (h-M): positive control for human genes. (F) SQ-RT-PCR analysis of EP-HACDC cultured under myogenic in vitro conditions. After 4 weeks of culture, MyHC-IIx/d was detected by SQ-RT-PCR in the myogenic culture (lane 1, My) but not in control culture (lane 2, C). Lane 3 (RT⫺): control of lane 1, to control for genomic contamination. Lane 4 (W): water control. cDNA samples were equalized for the expression of ␤-actin. The experiment was conducted in quadruplicate.

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Fig. 3. Bone differentiation. (A) HACDC were cultured for 3 weeks in the presence of osteogenic [14] or growth (control) medium. Calcium deposits are stained with alizarin red. (B–E) To induce bone differentiation in vivo, EP- and LP-HACDC were seeded into Collagraft scaffolds, and the constructs were implanted subcutaneously into nude mice. Eight weeks after implantation the constructs were explanted. (B) Minute ossicles (arrows) were retrieved in Collagraft seeded with either EP-HACDC (Masson’s trichrome) or LP-HACDC (C, hematoxylin– eosin). Fibrous and osteoid tissues were also present. (D) The ossicles stained positive for human osteocalcin as evaluated by immunohistochemistry. (E) Masson’s trichrome staining of a parallel, nonconsecutive section. (F) The antibody used for osteocalcin immunodetection was specific to human osteocalcin, as it did not bind to adult mouse bone. Bars ⫽ 50 ␮m.

determine the type of tissue formed in vivo starting from the same cell pool, we implanted two aliquots of the same EP-HACDC culture either in the CFA or subcutaneously as a cell–Collagraft construct. At 8 weeks, the implants were dissected and analyzed histologically after staining with Masson’s trichrome and safranin O. As shown in Fig. 5, we retrieved fibrous and bone tissues, but no cartilage, in the subcutaneously implanted Collagraft construct. In contrast, abundant safranin O-positive cartilage was retrieved in the thighs injected with the 5 ⫻ 106 HACDC suspension (CFA). Taken together, these data suggest that the interplay of environmental cues and phenotypic stability specifies the type of tissue formed by HACDC in vivo. Adipogenic differentiation in vitro Adipogenic culture conditions [8,14] promoted the accumulation of oil red O-positive lipid vacuoles by EP-and LP-HACDC (Fig. 6). This phenomenon was observed in clusters of cells across the culture dish. SQ-RT-PCR analysis revealed upregulation of the adipogenic markers PPAR␥-2 and aP2 in the treated cultures. Discussion Adult articular cartilage is traditionally considered a terminally differentiated tissue devoid of significant intrinsic

repair mechanisms. Its repair is thought to rely on mesenchymal progenitors incoming from the underlying bone marrow [15] or from the synovial membrane [8]. Some lines of evidence, however, suggest that articular cartilage may contain resident cells that are sensitive to environmental cues and responsive to cartilage damage. First, in human diseases restricted to the articular cartilage, such as chondromalacia patellae, intense cell proliferation has been reported within the lesion as well as within the surrounding cartilage [16]. Second, especially in young animals, small partial-thickness articular cartilage defects display significant repair [17–19]. In this study we have shown that articular cartilage from adult human individuals contains cells that, when released, can enter myogenic, osteogenic, and adipogenic differentiation, maintaining this capacity throughout passaging, within the ranges examined. The HACDC plasticity cannot be explained solely by fusion with cells resident in the host tissues [20,21], since myogenic, adipogenic, and osteogenic differentiation was, at least partially, also induced in vitro, in the absence of mature host cells. In vivo, differentiation appears to be co-determined by phenotypic stability and environmental cues. The importance of the interplay between environmental factors and phenotypic stability in tissue determination is underscored by findings described in chick– quail embryo transplantation experiments [22]. This study showed that

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Fig. 4. Gene expression dynamics during bone formation in Collagraft. (A) Expression of the bone markers osteocalcin (OC), osteopontin (OP), and bone sialoprotein (BSP) and the cartilage marker COL2A1 was monitored by SQ-RT-PCR in the HACDC monolayers, 4 and 8 weeks after implantation of the Collagraft constructs. The primers used were specific to the human genes with the exception of mh-␤-actin. Lane 1 (E): Collagraft implanted without human cells. Lane 2 (mj): mouse joint containing bone and cartilage tissues included as control for primer specificity to human genes. Lane 3 (bm): human bone marrow stromal cells implanted into Collagraft, used as a positive control for osteogenesis. Lane 4: EP-HACDC in monolayer culture. Lanes 5 and 6: EP-HACDC–Collagraft constructs, 4 and 8 weeks after implantation, respectively. Lane 7: LP-HACDC in monolayer. Lanes 8 and 9: LP-HACDC–Collagraft constructs 4 and 8 weeks after implantation, respectively. Lane 10 (w): water control. All the samples containing human cells were normalized for expression of the housekeeping gene ␤-actin using human-specific primers. The samples that did not contain human cells were normalized to the highest expression of ␤-actin using primers for human/mouse ␤-actin. The bone markers, undetectable in the monolayer cultures, were progressively upregulated after 4 and 8 weeks in Collagraft. (B) Reproducibility. Three EP-HACDC (2 PD) and 3 LP-HACDC (16 PD) from different donors were seeded into Collagraft and implanted into nude mice. Expression of human osteocalcin was monitored in the monolayers before implantation and at 4 and 8 weeks after implantation as indicated. The ages of the donors are indicated.

fragments of quail sclerotomes from stage XII or older retained their commitment to cartilage when transplanted dorsally into age-matched (Embryonic Days 4 to 6) chick recipients, but not when grafted into younger recipients. Sclerotome fragments of earlier stages never retained their cartilage commitment, but integrated into the muscle and dermis. In vitro-expanded HACDC retain for 4 to 6 PD the capacity to spontaneously form stable cartilage in vivo [3]. On further passaging, a partial chondrocyte differentiation could still be recovered under specific in vitro conditions, e.g., by culture in low-melting-temperature agarose, but the capacity to spontaneously form stable cartilage in vivo was lost [3]. When injected in a well-standardized CFA, LPHACDC did not form any cartilage tissue, but persisted for at least 8 weeks, mostly forming a collagen type I-rich fibrous tissue. Some cells, however, acquired a skeletal

muscle phenotype. In the same assay, EP-HACDC formed a cartilage nodule, expressed human COL2A1, but human MyHC-IIx/d was not detected. In the MA, however, like LP-HACDC, also EP-HACDC expressed muscle markers and fused with host myofibers. This apparent discrepancy could be explained by a better exposure of the injected individual cells to potent myogenic signals in the MA. First, in the MA the cardiotoxin-induced muscle damage elicits muscle regeneration and, therefore, the production of potent myogenic signals [6]. Second, in the MA the injection of a 10-times-lower number of HACDC into a severely injured muscle reduces cell aggregation and ensures a more even distribution of the individual cells between the regenerating myofibers (data not shown). More limited muscle damage may also occur in the CFA, because of the needle injury and the injection of a relatively large volume of cell suspension. This damage, however, is minimal when compared with that caused by cardiotoxin, and may not reach the “threshold” levels required to recruit EP-HACDC to myogenesis. This threshold may be higher for EP-HACDC in the CFA by virtue of their remarkable phenotypic stability in vivo and the “community effect” [23] generated by their distribution in large cellular aggregates. The multilineage differentiation observed may be explained either by the reprogramming of differentiated chondrocytes or by the existence of a separate undifferentiated, multipotent cell population within articular cartilage. In the first hypothesis, the chondrocytes would cease the expression of the cartilage markers and acquire the myogenic, osteogenic, or adipogenic ones. In the second, chondrocytes would be overgrown by a progenitor cell subpopulation. As a third possibility, chondrocytes would persist throughout passaging as de-differentiated cells, accounting for the loss of their capacity to form cartilage in vivo, while a separate, constant subpopulation of multipotent progenitors would be responsible for the formation of bone, adipocytes, and muscle in the assays employed in this study. The hypothesis of the existence of a separate, undifferentiated progenitor cell population within HACDC finds support in the focal nature of differentiation in all the assays used. Alternative explanations for the focal nature of differentiation are (i) that the assays employed may be suboptimal for the complete differentiation of all the cells, or (ii) that not all the cells in the culture are equally sensitive, at the same time, to differentiation signals in function, for instance, of the cell cycle. In addition, in all the in vivo assays employed, the distribution of the cells was uneven. Consequently, different individual cells were exposed to different environmental cues. For instance, the cells that clustered in the CFA, and though to a much lesser extent also in the MA, were exposed to a microenvironment that was substantially different from that of the isolated human cells dispersed between the host myofibers. The present data do not allow us to discriminate whether the observed phenomena are due to the existence of a distinct subpopulation of progenitor cells within adult articular cartilage or to a phenotypic plasticity of articular chon-

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Fig. 5. Influence of the microenvironment on tissue formation. Two aliquots out of the same primary HACDC culture were implanted either subcutaneously in Collagraft (A–C) or intramuscularly into the thighs (CFA) (B) of nude mice and dissected for histological analysis 8 weeks after implantation. (A), Masson’s trichrome showing the formation of fibrous tissue and minute ossicles. (B), Detail of (A) delimited by the square showing mature ossicles. (C) Safranin O staining of a parallel section of (A) showing that no cartilage tissue is detected within the EP-HACDC–Collagraft construct. (D), Safranin O staining of the implant obtained from the intramuscular EP-HACDC injection (CFA) showing abundant safranin O-positive cartilage. Bars ⫽ 800 ␮m in (A, C) and 50 ␮m (D).

drocytes. The surprisingly broad differentiation potential that we report may be the result of the loss of phenotypic stability associated with in vitro culture [24]. De-differentiated, unstable cells may be more sensitive to environmental cues. We have shown that freshly isolated HACDC, which had never been cultured in vitro, could enter myogenesis in the MA, indicating that the in vitro culture is not required for myogenic differentiation in vivo. We cannot, however, exclude that the enzymatic digestion of the native articular cartilage, required to release HACDC, may be responsible for some de-differentiation. As for bone marrow MSCs [25], it is also possible that the number of cell divisions, including those possibly occurring in vivo after implantation, may directly influence the differentiation of the grafted cells.

The in vivo phenotypic stability of EP-HACDC is retained for a certain number of cell divisions in culture and is influenced by cell culture conditions. This phenotypic stability allows EP-HACDC to override, to a certain extent, the environmental signals. Indeed, the CFA is not per se inducing cartilage formation, since periosteal cells, which under specific conditions have chondrogenic potential [26], fail to form cartilage in this assay [3]. Even in the presence of strongly myogenic signals, such as in the MA, the expression of COL2A1 was retained by EP-HACDC, and the failure to form mature cartilage tissue in this assay may be linked simply to the small number of injected cells. The phenotypic stability may be dependent on either epigenetic factors, such as DNA methylation and histone deacetylation [27], or on an as yet unidentified autocrine–paracrine sig-

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Fig. 6. Adipogenic differentiation. EP- and LP-HACDC were cultured for 3 weeks in either growth medium (A and D, respectively) or in adipogenic medium (B and E). (C) Detail of (B). (F) SQ-RT-PCR analysis demonstrated the upregulation of the adipogenic markers PPAR␥2 and aP2 in the treated cultures. c, control medium; a, adipogenic medium; w, water. Bars ⫽ 100 ␮m.

nal(s). These two hypotheses are not mutually exclusive, and the balance between the determinants of stability and the signals coming from the microenvironment would ultimately determine the type of tissue formed in vivo. We cannot exclude contamination by multipotent cells from other origins, such as the bone marrow [28] incoming from involuntary violation of the subchondral plate, from the synovial membrane [8], and from the vasculature [29]. This possibility seems remote because adult human articular cartilage is avascular and because of the high consistency and reproducibility of the results. When harvesting the articular cartilage for the obtainment of HACDC, extreme caution was taken to avoid the calcified layer and the cartilage at the borders of the articular surface to avoid contamination by cells from the bone marrow and the synovial membrane. In addition, the cartilage tissue was washed extensively before enzymatic digestion. At this point, we cannot identify a subpopulation of cells, within HACDC, that has distinct progenitor properties. The lack of molecular markers that identify unequivocally mesenchymal stem cells makes further characterization of a hypothetical specific subpopulation difficult and eventually enrichment through cell sorting a challenge. In addition, as one can also evince from this set of data, cells are dynamic entities that can respond to environmental cues. The expression of molecular markers is affected by the culture conditions and possibly by the tissue dissection and cell isolation [30]. This is particularly relevant for cell types such as HACDC, which are known to be phenotypically not stable in vitro [3,5]. Under specific experimental conditions, molecular markers represent a pow-

erful tool to monitor the phenotypic changes resulting from in vitro manipulations, but one should be aware that this might have little to do with their origin or the intrinsic nature of the cells [30]. Our experiments do not address whether the multipotency of HACDC is inherent to single cells, or whether there are several different unipotent progenitors. An experiment of cell cloning starting from 50 single cells from a secondary culture yielded five independent clones. None of these clones, however, could be expanded to the extent required to complete the experiments needed to claim multipotency (data not shown). This limited expandability in vitro may indicate that the clonogenic cells from adult articular cartilage may be different from the bone marrow-derived, or synovial membrane-derived mesenchymal stem cells that display a longer life span [8,14]. The existence of distinct progenitor cells within adult articular cartilage has not been proven so far. In addition, it is difficult to explain what multipotency would be needed for in articular cartilage. Multipotency could be an intrinsic property of “reserve cells” that are not committed to any specific lineage. Alternatively, the observed multipotency may be the result of the disruption of a mechanism of control of differentiation caused by in vitro manipulations or in vivo microenvironmental factors. Further understanding these mechanisms may represent an opportunity to restore the phenotypic stability in those cells that have lost it due to in vitro manipulation or pathologic conditions, or to redirect cell differentiation into a specific direction in protocols for cell-based tissue repair. Importantly, a balance between plasticity and phenotypic

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stability, as in phenotypically stable EP-HACDC, may be an important requirement to guarantee stable cartilage formation in autologous chondrocyte implantation protocols, preventing the undesired heterotopic tissue formation described with mesenchymal stem cells [31,32].

Acknowledgments This work was supported by FWO Grant G.0192.99 and IWT Grant N,000259 We thank Przemko Tylzanowski for critically reviewing the manuscript. We acknowledge Johan Vanlauwe and Johan Bellemans (Department of Orthopedics, University Hospitals Katholicke Universiteit, Leuven, Belgium) for providing articular cartilage samples. Special thanks are also due the people working in the mortuary of the University Hospitals Katholicke Universiteit, Leuven. We are grateful to J. Neys and I. Derese for processing tissue blocks for histology.

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