J O U R N A L O F T H E M E C H A N I C A L B E H AV I O R O F B I O M E D I C A L M AT E R I A L S
(
)
–
available at www.sciencedirect.com
journal homepage: www.elsevier.com/locate/jmbbm
Research paper
Dynamic compressive strain influences chondrogenic gene expression in human periosteal cells: A case study I.C. Bonzani a,b , J.J. Campbell d , M.M. Knight d , A. Williams e , D.A. Lee d , D.L. Bader d , M.M. Stevens a,b,c,∗ a Department of Materials, Royal School of Mines, Imperial College London, London SW7 2AZ, UK b Institute for Biomedical Engineering, Imperial College London, London SW7 2AZ, UK c Department of Bioengineering, Imperial College London, London SW7 2AZ, UK d Medical Engineering Division and IRC in Biomedical Materials, School of Engineering and Materials Science, Queen Mary University of
London, London E1 4NS, UK e Chelsea and Westminster Healthcare NHS Trust, Chelsea, London SW10 9NH, UK
A R T I C L E
I N F O
A B S T R A C T
Physical stimuli play a crucial role in skeletogenesis and osteochondral repair and Keywords:
regeneration. Although the periosteum and periosteum-derived cells offer considerable
Periosteal-derived cells Chondrogenic differentiation Dynamic compression Mechanotransduction
therapeutic potential, the molecular mechanisms that control their differentiation are still not fully understood. As an initial case study, this work explores the hypothesis that dynamic compression might selectively enhance chondrogenic and/or osteogenic differentiation in human periosteal cells from two donors. Donor derived human periosteal cells were expanded in monolayer culture before seeding in 3% (w/v) agarose constructs. The ability of this in vitro culture model to support cell viability, chondrogenesis, and mechanotransduction was optimised. The time course of early chondrogenic differentiation was assessed by real time RT-PCR of mRNA expression levels for bone and cartilage specific gene markers. Intermittent dynamic compression (1 Hz, 15% strain) was applied to constructs, in the presence or absence of 10 ng/ml TGF-β3, for up to 4 days. The combined effect of TGF-β3 and compressive loading on the expression levels of the Sox9, Runx-2, ALP, Collagen X, and collagen type I genes was donor dependent. A synergistic effect was noted only in donor two, with peak mRNA expression levels at 24 h, particularly Sox-9 which increased 59.0-fold. These findings suggest that the interactions between mechanical stimuli and TGF-β signalling may be an important mechanotransduction pathway for human periosteal cells and that, importantly, this cellular mechanosensitivity varies between donors. c 2011 Elsevier Ltd. All rights reserved. ⃝
∗ Corresponding author at: Department of Materials, Royal School of Mines, Imperial College London, London SW7 2AZ, UK. Tel.: +44 0 20 7594 6804. E-mail address:
[email protected] (M.M. Stevens). c 2011 Elsevier Ltd. All rights reserved. 1751-6161/$ - see front matter ⃝ doi:10.1016/j.jmbbm.2011.06.015
2 1.
J O U R N A L O F T H E M E C H A N I C A L B E H AV I O R O F B I O M E D I C A L M AT E R I A L S
Introduction
Various tissue engineering and regenerative medicine strategies have exploited the therapeutic potential of periosteal cells through the use of tissue grafts, (Brittberg et al., 1994; Delaney et al., 1989; Stevens et al., 2005) explants (O’Driscoll et al., 1994; Stevens et al., 2004a,b) or isolated cells with or without an associated three-dimensional scaffold (De et al., 2001; Emans et al., 2006; Hutmacher and Sittinger, 2003; Nakahara et al., 1990). Periosteal cells hold promise in osteochondral repair applications due to their ease of isolation, expansion potential in vitro and ability to differentiate into multiple mesenchymal lineages (Choi et al., 2008; De et al., 2006; Stevens et al., 2005). Nevertheless, exploring and optimising the governing factors controlling periosteal cell osteogenesis and chondrogenesis would be of considerable benefit. Periosteal cells play a role in the development and maintenance of the musculoskeletal system, (Malizos and Papatheodorou, 2005; Orwoll, 2003; Rauch, 2005) including long-bone growth, bone fracture repair and bone remodelling. In embryonic and postnatal bone development, radial growth of long bones involves the direct conversion of mesenchymal precursor cells in the periosteum to osteoblasts (Gardner, 1971; Pechak et al., 1986; Taylor, 1992). In fracture healing, the periosteum is responsible for bridging the callus formation and participating in endochondral and intramembranous ossification (Bolander, 1992; Simmons, 1985). The migration, proliferation and differentiation of periosteal precursor cells during these processes are coordinated by a variety of systemic, local and mechanical factors. Indeed, the effects of systemic hormones (Fleisch et al., 2003; Parfitt, 2002; Vanderschueren et al., 2006) (e.g. parathyroid hormone, bisphosphonates, and sex steroids), local growth factors and cytokines (Hanada et al., 2001; Jung et al., 2005; Malizos and Papatheodorou, 2005; Yakar et al., 2002) (e.g. fibroblast growth factors, transforming growth factors, bone morphogenic proteins, and insulin-like growth factors) on periosteal cell proliferation and differentiation have been studied extensively and characterised both in vivo and in vitro. However, the mechanical control mechanisms that influence periosteum function have received relatively little attention. It has been suggested that the in vivo mechanical loading environment is a primary modulator of bone apposition rate (Parfitt et al., 2000; Ruff, 2003; van der Meulen et al., 2003) and secondary chondrogenesis (Archer et al., 2006) on periosteal surfaces during joint formation and post-natal growth. Attempts to characterise the biological responses of the periosteum to mechanical loading have utilised externally applied loading devices in conjunction with small animal models (Gross et al., 1997; Harada et al., 2002; LaMothe et al., 2005; Matsumoto et al., 1998; Meyer et al., 2001). By applying mechanical forces in vivo, expression of a variety of osteogenic and chondrogenic genes in the periosteum can be induced, (Matsumoto et al., 1998; Raab-Cullen et al., 1994) which results in the rapid transformation of quiescent periosteal surfaces to those on which bone formation occurs (Pead et al., 1998). We therefore hypothesised here that the application of mechanical forces in vitro could be used to induce and control periosteal cell differentiation (Carter et al., 2004, 1988; van der Meulen et al., 2003). Previous studies
(
)
–
have shown that compressive loading regimes can stimulate the chondrogenic differentiation of mesenchymal stem cells (Campbell et al., 2006; Miyanishi et al., 2006b) and enhance cartilage-specific matrix formation (Buschmann et al., 1995; Guilak et al., 1994; Lee and Bader, 1997). To date, however, there are no comparable studies published on the effects of mechanical stimulation on human periosteal cells in vitro. This study aims to determine whether the application of compressive strains provides early chondro-inductive effects in isolated human periosteal cells. This aim has been achieved by developing a 3D culture system and determining appropriate controlled external loading regimes, based on a cell/agarose system that was used previously for studying the effects of dynamic compressive strain on chondrocytes. In the present study, the expression profiles of early response genes have been examined in human periosteal cells cultured in agarose constructs under dynamic compression strain in the presence or absence of transforming growth factor-beta3 (TGF-β3).
2.
Materials and methods
2.1.
Human periosteal cell isolation and culture
This study was approved by The Chelsea and Westminster Trust NHS ethical committee (project RREC2700). Periosteal explants were harvested from the proximal tibia of two donors (donor 1–a 32 yr old male, donor 2–a 43 yr old female) using a periosteal elevator. The explants were finely minced, and enzymatically digested in a collagenase solution consisting of 3 mg/ml collagenase D (Roche Applied Science, West Sussex, UK), 3 mg/ml collagenase type II (Sigma-Aldrich, Poole, UK) and 5 mM CaCl2 in growth medium (high glucose Dulbecco’s Modified Eagle Medium containing 10% v/v foetal bovine serum, 1% v/v L-glutamine, 100 units/ml penicillin, 100 µg/ml streptomycin [all from Invitrogen, Paisley, UK]). After incubation at 37 ◦ C for 4.5 h, the cell suspension was washed, passed through a 70 µm cell strainer (Falcon, Becton Dickinson, Oxford UK), and expanded in monolayer at a seeding density of 5 × 104 cells/cm2 per passage for up to 10 passages in the growth medium. To examine the presence of progenitor phenotypes during expansion, periosteal cells were stained with monoclonal antibodies against Stro-1 and ALP (R&D systems, UK) and analysed using flow cytometry as described previously (Stewart et al., 1999).
2.2. Determination of cell viability on 3D cell seeded constructs The effect of nutrient diffusivity on cellular viability was examined to mitigate the challenges associated with the preparation and culture of 3-dimensional cell-seeded constructs. This was achieved by investigating the effects of cell-seeding density on the construct viability. Cellular constructs were produced from passage 3 isolated periosteal cells from a separate donor (65 yr old male) and had final cell densities of 0.5, 2.0, and 8.0 × 106 cells/ml in 3% (w/v) low gelling temperature agarose (type VII; Sigma). The constructs were cultured in individual wells of a 24-well plate (BD
J O U R N A L O F T H E M E C H A N I C A L B E H AV I O R O F B I O M E D I C A L M AT E R I A L S
(
)
3
–
Table 1 – TaqmanTM gene probes used in this study, with details of their sources and specificities. TaqmanTM gene probes (applied biosystems) Gene name Alkaline phosphatase Collagen Type 1, alpha 1 Runt-related transcription factor 2 SRY (sex determining region-y)-box 9 Collagen Type II, alpha 1 Collagen Type X Eukaryotic 18s rRNA
Symbol ALP COL1A1 Runx-2 Sox-9 COL2A1 COL10A1 18s
Catalogue number Hs00758162 Hs00164004 Hs00298328 Hs00165814 Hs00156568 Hs00166657 Hs99999901
Falcon, UK) and supplied with 1 ml of growth medium for up to 10 days. At days 2, 4, and 10, three constructs per cell density were removed from culture, sectioned vertically, and incubated for 1 h at 37 ◦ C with 5 µM calcein-AM and 5 µM ethidium homodimer-1 (Molecular Probes; Paisley, UK) in 2 ml of complete media. A systematic sampling procedure, as described previously (Heywood et al., 2004) was adapted to fluorescently visualise green (live) and red (dead) cells and determine percentage cell viability (mean ± SD) throughout the whole construct and for both peripheral and core regions. A minimum of 4 replicate sampling series were performed per specimen, with 3 specimens examined per cell density per time-point.
2.3. Chondrogenic culture in free-swelling periosteal cellagarose constructs Periosteal cell-agarose constructs, from donor 2 cells at passage 2, were produced at a final density of 2.0×106 cells/ml in 3% (w/v) low gelling temperature agarose (type VII; Sigma) moulds (6 mm diameter and 4 mm depth). The constructs were distributed into a 24-well plate and bathed overnight in 1 ml of chondrogenic media, consisting of high glucose DMEM, 1 µM dexamethasone, 1 mM sodium pyruvate, 0.17 mM ascorbic acid-2-phosphate, 0.35 mM proline and ITS + premix (defined chondrogenic media without TGF-β3, Lonza, Slough, UK). In the following day, designated time 0, the constructs were bathed in 1 ml of chondrogenic media with or without 10 ng/ml TGF-β3 and incubated for up to 16 days in static conditions. Media replacement and TGF-β3 supplementation were performed every other day. Samples were taken at 0.5, 1, 2, 4, 8, and 16 days for the analysis of gene expression using real-time RT-PCR (n = 3 per time point per condition).
2.4. Intermittent dynamic mechanical stimulation of periosteal cell-agarose constructs Mechanical stimulation experiments were adapted from methods described previously (Campbell et al., 2006; Lee et al., 2000). Cell-seeded agarose constructs (3% w/v) with a cell density of 2.0 × 106 cells/ml were prepared using periosteal cells derived from both donors at passage 3. Constructs were incubated overnight in defined chondrogenic media in the absence of TGF-β3. Individual constructs were arranged in the centre of each 24-well plate, and positioned within the custom made cell-straining apparatus (Zwick-Roel,
Accession number NM000478 NM000088 NM001015051 NM000346 NM001844 NM000493 X03205
Chromosome location Chr. 1 21,769,385–21,772,879 Chr. 17 45,632,113–45,633,974 Chr. 6 45,622,542–45,626,797 Chr. 17 67,630,455–67,634,156 Chr. 12 46,679,969–46,684,528 Chr. 6 116,546,814–116,553,363 N/A
Redditch, UK) (Lee and Bader, 1997). This apparatus was positioned into a custom-made cell-straining rig and the individual flat-ended indenters at the base of stainless steel pins were lowered onto the top surface of the cell-seeded constructs to provide a tare load of 0.028 N, associated with the mass of the pin (Campbell et al., 2006). Each mechanical experiment incorporated 12 constructs (6 unloaded + 6 dynamically loaded) which were bathed in 1 ml chondrogenic media without TGF-β3 supplementation and a further 12 constructs bathed in 1 ml chondrogenic media with 10 ng/ml TGF-β3 supplementation. This set-up provided 4 separate conditions per plate. The experimental time course conditions were selected as 0.5, 1, 2, and 4 days, with or without dynamic compressive strain and with medium changed every 2 days. The assembled straining apparatus with cellseeded constructs was transferred to an adapted incubator at 37 ◦ C and 5% CO2 , where the loading frame was secured to the loading actuator. The unconfined dynamic compressive strain with an amplitude of 15% strain and frequency of 1 Hz was applied to the experimental constructs with repeated cycles of loading and a recovery of 1.5 and 4.5 h, respectively (Campbell et al., 2006). In this way, 4 complete loading/unloading cycles of 6 h per 24 h period were performed.
2.5.
Real time RT-PCR
For gene expression analysis, cellular RNA was extracted from cell-agarose constructs using an RNeasy mini kit (QIAGEN, West Sussex, UK) and reverse transcribed into cDNA. Single-plex real-time RT-PCR reactions were carried R out in 20 µl volumes containing 10 µl Taqman⃝ universal mastermix (Applied Biosystems, CA, US), 7 µl 0.1% v/v diethylpyrocarbonate (DEPC) water (Invitrogen Ltd, Paisley, R UK), 2 µl extracted cDNA and 1 µl Taqman⃝ probe (Applied R Biosystems, CA, USA). TaqMan⃝ probes used to target the six selected genes and the housekeeping gene, 18s ribosomal rRNA, are described in Table 1. Each reaction was carried out in triplicate. The PCR reaction was initiated by a 2 min 50 ◦ C and a 10 min 95 ◦ C step to optimise thermal cycling conditions for the ABI Prism 7700 sequence detection system (Applied Biosystems, CA, USA) used to detect relative quantification of gene expression. This was followed by PCR amplifications performed for 40 cycles in a Corbett Rotor-Gene 6000 (Corbett Life Science, Sydney, Australia) at 95 ◦ C for 15 s and 60 ◦ C for 1 min. A measurement of quantitative expression of target sequences, normalised to the abundance of the endogenous housekeeping gene (18s) and expressed relative to day 0 control constructs, was
4
J O U R N A L O F T H E M E C H A N I C A L B E H AV I O R O F B I O M E D I C A L M AT E R I A L S
(
)
–
Table 2 – Percentage cell viability and cell density for the core and periphery of periosteal cell-constructs seeded at different densities and cultured over a period of 10 days in growth medium. The peripheral region represents both the outer third of the construct and the core region represents the central third of the construct. Each value represents the mean ± standard deviation of 20 replicate samplings in each region for 3 constructs per seeding density (∗ p < 0.05—for differences between the peripheral and core regions). The description of the results is given in the Supplementary Information. Seeding density Days in culture Cell viability (%) Cell density (cells/mm2 )
1 4 10 1 4 10
Core
0.5 × 106 cells/ml Periphery P < 0.05
90.2 ± 3.1 85.3 ± 5.0 78.4 ± 6.0 58.6 ± 7.9 48.1 ± 8.9 48.8 ± 10.7
95.6 ± 4.0 89.3 ± 5.4 85.4 ± 10.1 46.7 ± 9.9 46.0 ± 10.1 45.1 ± 9.5
∗ ∗ ∗
Core
2.0 × 106 cells/ml Periphery P < 0.05
84.7 ± 6.2 92.2 ± 7.4 ∗ 80.3 ± 5.6 84.2 ± 5.6 86.0 ± 3.9 88.5 ± 6.6 112.5 ± 17.7 110.6 ± 24.4 105.5 ± 14.7 108.5 ± 13.4 105.3 ± 19.5 97.5 ± 17.5
Core
8.0 × 106 cells/ml Periphery P < 0.05
89.2 ± 6.2 75.4 ± 8.4 73.2 ± 8.3 219.0 ± 38.8 229.9 ± 52.5 225.4 ± 38.3
89.4 ± 6.3 77.3 ± 10.6 77.3 ± 10.2 235.6 ± 51.7 229.6 ± 55.0 191.5 ± 38.1
carried out using the −11CT method. Fold changes in gene expression are presented as mean ± SD change relative to day 0 control constructs. For dynamic gene expression analysis, 3 constructs for each donor per culture condition (unloaded + TGF-β3, loaded + TGF-β3, unloaded no TGF-β3, loaded no TGF-β3) per time point (0.5, 1, and 2 days) were harvested for RT-PCR.
2.6.
Statistics
Mann–Whitney U tests were used to examine the differences in cell viability, density, and RT-PCR results for all experimental conditions. A level of 5% was considered statistically significant (∗ p ≤ 0.05).
3.
Results
3.1. Encapsulation and culture of periosteal cells in agarose The encapsulation of periosteal cells in 3% (w/v) agarose from both donors resulted in a significant change in cell morphology. Periosteal cells in monolayer exhibited spindleshaped, and elongated fibroblastic phenotypes with long membrane extensions, whereas corresponding encapsulated cells were smaller and more rounded (Supplementary Fig. 1). The cell encapsulation method used in this study resulted in a homogeneous distribution of cells throughout the 3D construct, independent of initial cell-seeding density. During culture in growth medium, no significant differences in absolute cell number or density were found between the core and the periphery of the constructs at any seeding density (Table 2). Additionally, there were no significant changes in the overall construct cell density over 10 days in culture, which suggests that in free-swelling conditions agaroseembedded periosteal cells did not proliferate significantly in standard growth medium. At 24 h post-cell encapsulation in the growth medium, periosteal cells from both donors in agarose exhibited significant increases in Runx-2, Sox-9, collagen I, and collagen X mRNA (Fig. 1). In particular, increases in the hypertrophic chondrocyte marker collagen X were by 19.6fold, the osteogenic transcription factor Runx-2 by 9.5-fold,
Fig. 1 – Real time RT-PCR quantification of Sox-9, Runx-2, ALP, Collagen X, and Collagen I gene expression for passage 3 periosteal cells (from Donor 2) encapsulated in 3% (w/v) agarose and incubated for 24 h in standard growth medium. Fold changes in mRNA expression are expressed relative to patient matched monolayer cultured cells (∗ p < 0.05).
and the chondrogenic transcription factor Sox-9 by 4.6-fold. By contrast, the fold increase in mRNA expression for ALP for cells in agarose was not significantly different from cells cultured in monolayer (p > 0.05).
3.2. Temporal profiles of gene expression in free-swelling agarose constructs The temporal profiles for osteogenic (Runx-2, ALP, and Col Ia1) and chondrogenic (Sox-9, Col IIa1, Col X) gene expression by periosteal cells in 3D agarose free-swelling culture are illustrated in Fig. 2. In general, the addition of 10 ng/ml of TGF-β3 during culture increased mRNA levels of each gene examined, over the untreated constructs (−TGF-β3). Peak expression levels of Sox-9, Col X, and Runx-2 occurred at 16 days in the presence of TGF-β3. In the absence of TGFβ3, Runx-2, Col I, and Col X mRNA levels were up-regulated after 0.5 days in culture and subsequently decreased to levels similar to or below baseline day 0 expression levels by 2 days. The expression of the osteoblast marker ALP progressively declined during culture in both treatments (±TGF-β3) (Fig. 2(D)). Conversely, periosteal cells in agarose exhibited a progressive increase in the expression of the hypertrophic
J O U R N A L O F T H E M E C H A N I C A L B E H AV I O R O F B I O M E D I C A L M AT E R I A L S
(A) Sox-9.
(B) Collagen X.
(C) Runx-2.
(D) ALP.
(
)
–
5
(E) Collagen I. Fig. 2 – The relative gene expression of Sox-9 (A), Collagen Type X (B), Runx-2 (C), ALP (D), and Collagen Type I (E) in passage 2 periosteal cell-agarose constructs (Donor 2) cultured for 16 days in chondrogenic media with or without 10 ng/ml TGF-β3. The expression levels are normalised to internal housekeeping gene 18s and the data are presented relative to day 0 levels (>1 = increase and 75% during culture for all densities examined. However, for our model parameters 2 × 106 cells/ml represented the optimal seeding density for sustained periosteal cell viability during in vitro culture. The previous in vitro loading studies using cell and explant culture models have demonstrated that the method of load application, its magnitude, frequency, and duty cycle of loading can influence genetic and biosynthetic response
(Bao et al., 2000; Chowdhury et al., 2003; Miyanishi et al., 2006a; Mukherjee et al., 2001). The level of gross strain used (15%) resulted in local strain values of approximately 15%, which lies within the 0%–20% physiological range of cell strains estimated in intact cartilage (Broom and Myers, 1980; Guilak, 1994). This gross strain was also shown to be equivalent to the cell strains within the construct (Freeman et al., 1994; Knight et al., 1998). In addition, the selected loading frequency of 1 Hz is physiologically viable and has stimulatory effects on chondrocytes in explants and 3D constructs (Elder et al., 2001; Lee and Bader, 1997). In the second phase of this work, chondrogenic differentiation was initiated in free-swelling agarose cultures. Prior to chondrogenic induction, the encapsulation of periosteal cells in agarose gels caused a morphology change, which coincided with significant up-regulation in Sox-9, Runx-2, and Coll I, and X (Fig. 2). The previous studies have reported phenotypic modulation caused by an enforced spherical morphology within hydrogels. For example, Coll II and aggrecan gene expression was evident in marrow stromal cells cultured in alginate and agarose gels, as opposed to collagen gels where cells adopted a more stellate morphology (Diduch et al., 2000). Chondrogenic differentiation was induced through TGF-β and dexamethasone supplementation, as described in previous bone marrow stromal cell cultures (Campbell et al., 2006; Pittenger et al., 1999). The expression of transcription factors Runx-2 and Sox-9, and Col I and X all increased over the 16 day culture period (Fig. 2). The increased expression of Coll I is a feature common with the early stages of chondrogenesis, involving mesenchymal precursor cells (Kosher et al., 1986). Although the accumulation of Coll X mRNA over the 16 day period could indicate differentiation towards the hypertrophic phenotype, the corresponding levels of Runx-2 and ALP would suggest that the cells exhibit a pre-hypertrophic phenotype at day 16 (Goldring et al., 2006). This pre-hypertrophic expression pattern correlates with the
8
J O U R N A L O F T H E M E C H A N I C A L B E H AV I O R O F B I O M E D I C A L M AT E R I A L S
previous human mesenchymal stem cell studies that report the up-regulation of collagens type I and X during the early stages of chondrogenesis (Hardingham et al., 2006; Yoo and Johnstone, 1998). The presence of pre-hypertrophic phenotypes in periosteal cell-constructs during free-swelling culture was further substantiated by the heightened expression levels of both Runx-2 and Sox-9 at day 16. Both transcription factors are highly expressed in osteochondroprogenitor cells and pre-hypertrophic chondrocytes. In this context, the 3-fold increase in Sox-9 expression compared to Runx-2 levels at day 16 suggests that cultures are progressing down the chondrogenic lineage (Hardingham et al., 2006; Zhou et al., 2006). Longer culture studies, however, are needed to examine the complete temporal expression profile of periosteal cells during chondrogenesis and to compare this marker pattern to what is seen in vivo (Bahrami et al., 2000; O’Driscoll, 1999). The expression levels of two important early transcription factors, Runx-2 and Sox-9, were examined in periosteal cells subjected to different loading cycles and culture conditions for up to 4 days. The application of dynamic compressive strains at 15% at a frequency of 1.0 Hz resulted in a donordependent response in human periosteal cells. In cells from donor 1, loading inhibited Runx-2 and Sox-9 expression, as demonstrated by decreased mRNA levels compared to unloaded constructs at nearly all time points. By contrast, loading of cells from donor 2 cells significantly increased gene expression levels, particularly in the presence of TGFβ3. Utilising the intermittent dynamic compression profile, of 1.5 h loading/4.5 h unloading, a short-term temporal response profile was established over 4 days. In agreement with the previous studies, downstream cellular response was greatly influenced by the duration of loading and the number of loading cycles (Chowdhury et al., 2003; Huang et al., 2005; Miyanishi et al., 2006a; Mukherjee et al., 2001). In responsive constructs (donor 2), expression levels peaked in the presence of TGF-β3 at 24 h, constituting 4 separate loading blocks of 1.5 h (5400 cycles). This expression peak was much more evident in the profile of Sox-9 mRNA, which demonstrated a 59-fold increase at 24 h. The profile of Runx-2 expression under each condition, however, remained relatively consistent over the 4 day study. Notably, in periosteal cells from donor 2, a significant increase in Sox9 mRNA levels occurred even in the absence of TGF-β3. However, extended loading studies (≥2 weeks (41)) examining the production of Coll II and aggrecan in more donor populations are needed to confirm whether mechanical signals alone are enough to induce chondrogenesis. The TGF-β3 treatment alone was found to increase the expression levels of Runx-2 and Sox-9 mRNA in periosteal cells from both donors. In donor 2 cells, however, the combination of mechanical stimulation and TGF-β3 was found to significantly up-regulate mRNA levels at all times compared to either stimulation alone; this synergy is consistent with recent studies. For example, in the presence of TGF-β and cyclic compressive strains (10%), increased chondrogenic differentiation and cartilage-related matrix production was reported for human MSCs after 14 days (Terraciano et al., 2007). The interactions between TGF-β signalling and mechanical stimulation could be due to a wide range of potential
(
)
–
mechanisms. The stimulated TGF-β signalling pathways could modulate mechanotransduction, directly or indirectly, by increasing the sensitivity of periosteal cells to loading through the activation of mechanosensitive proteins, such as focal adhesion kinase and paxillin (Luettich and Schmidt, 2003; Wang et al., 2005). Alternatively, downstream targets of TGF-β signalling may be the necessary component for mechanotransduction in periosteal cells. TGF-β has been shown to up-regulate Sox-9 gene expression in differentiating periosteal cells (Figs. 3 and 4) and is known to modulate periosteal chondrogenesis (Hanada et al., 2001; Miura et al., 2002; Mizuta et al., 2002). Thus by increasing the transcription of downstream mechanical stimulation targets, TGF-β signalling may indirectly amplify the effects of mechanotransduction. Conversely, mechanical stimulation may modulate TGF-β signalling. One mechanism may involve the production of TGF-β or its receptors through up-regulation of mRNA expression by mechanical loading. Huang and colleagues found that dynamic compressive loading promoted gene expression and protein production of both TGF-β receptors (TGF-βR-I and II) in rabbit-MSCs in agarose (Huang et al., 2005). This result is important because TGF-βR-1, phosphorylated by TGF-β, initiates intracellular signal transduction, which mediates chondrogenic differentiation of chondroprogenitor cells and mesenchymal stem cells (Hatakeyama et al., 2003). It is also intriguing to consider the possibility that the receptor availability on the cell surface may be regulated by mechanical forces. For instance, in an ideal chondrogenic environment TGF-β receptor could be mainly localised in non-raft membrane domains, where TGF-β signalling is promoted (Di Guglielmo et al., 2003). Although both TGF-β treatment and compressive strain represent strong transcriptional stimuli for early chondrogenic differentiation in periosteal cells cultured in agarose, this response is markedly donor dependent. Similar donor-todonor variability was also seen in the molecular response of human mesenchymal stem cells subjected to dynamic strains (Friedl et al., 2007). In general, the intrinsic heterogeneity of periosteal cells, in terms of lineage hierarchy and differentiation potential, represents a major challenge towards characterising their cellular responses. As shown previously, the differentiation processes in human periosteal cultures are not synchronised throughout the cell populations (Choi et al., 2008; De et al., 2006). In fact, phenotypic and genotypic analysis of periosteal cells and other adult stem cell cultures suggest that early differentiation may be regulated, at least in vitro, by stochastic mechanisms (Candeliere et al., 1999; Madras et al., 2002). This is thought to represent the developmental flexibility found in undifferentiated stem cells, which is lost during the commitment to more mature phenotypic state. In conclusion, a periosteal cell-agarose culture model has been successfully developed and optimised, to allow the study of cell differentiation in a biologically relevant mechanical environment. Ultimately, this study will act as a platform to further develop our understanding of human periosteal cell differentiation and enhance the in vitro development of tissue engineered constructs for cartilage regeneration. Author disclosure statement. No competing financial interests exist.
J O U R N A L O F T H E M E C H A N I C A L B E H AV I O R O F B I O M E D I C A L M AT E R I A L S
Acknowledgements We acknowledge funding from EPSRC (Grant no. EP/C520742/1). ICB thanks the Marshall Commission for the funding of his Ph.D. scholarship.
Appendix. Supplementary data Supplementary material related to this article can be found online at doi:10.1016/j.jmbbm.2011.06.015. REFERENCES
Archer, C., Buxton, P., Hall, B.K., Francis-West, P., 2006. Mechanical regulation of secondary chondrogenesis. Biorheology 43, 355–370. Bahrami, S., Stratmann, U., Wiesmann, H.P., Mokrys, K., Bruckner, P., Szuwart, T., 2000. Periosteally derived osteoblast-like cells differentiate into chondrocytes in suspension culture in agarose. Anat. Rec. 259, 124–130. Bao, X., Clark, C.B., Frangos, J.A., 2000. Temporal gradient in shearinduced signaling pathway: involvement of MAP kinase, cfos, and connexin43. Am. J. Physiol. Heart Circ. Physiol. 278, H1598–H1605. Bolander, M.E., 1992. Regulation of fracture repair by growth factors. Proc. Soc. Exp. Biol. Med. 200, 165–170. Brittberg, M., Lindahl, A., Nilsson, A., Ohlsson, C., Isaksson, O., Peterson, L., 1994. Treatment of deep cartilage defects in the knee with autologous chondrocyte transplantation. N. Engl. J. Med. 331, 889–895. Broom, N.D., Myers, D.B., 1980. A study of the structural response of wet hyaline cartilage to various loading situations. Connect. Tissue Res. 7, 227–237. Buschmann, M.D., Gluzband, Y.A., Grodzinsky, A.J., Hunziker, E.B., 1995. Mechanical compression modulates matrix biosynthesis in chondrocyte/agarose culture. J. Cell Sci. 108, 1497–1508. Campbell, J.J., Lee, D.A., Bader, D.L., 2006. Dynamic compressive strain influences chondrogenic gene expression in human mesenchymal stem cells. Biorheology 43, 455–470. Candeliere, G.A., Rao, Y., Floh, A., Sandler, S.D., Aubin, J.E., 1999. cDNA fingerprinting of osteoprogenitor cells to isolate differentiation stage-specific genes. Nucleic Acids Res. 27, 1079–1083. Carter, D.R., Beaupre, G.S., Wong, M., Smith, R.L., Andriacchi, T.P., Schurman, D.J., 2004. The mechanobiology of articular cartilage development and degeneration. Clin. Orthop. Relate. Res. S69–S77. Carter, D.R., Blenman, P.R., Beaupre, G.S., 1988. Correlations between mechanical stress history and tissue differentiation in initial fracture healing. J. Orthop. Res. 6, 736–748. Choi, Y.S., Noh, S.E., Lim, S.M., Lee, C.W., Kim, C.S., Im, M.W., et al., 2008. Multipotency and growth characteristic of periosteumderived progenitor cells for chondrogenic, osteogenic, and adipogenic differentiation. Biotechnol. Lett. 30, 593–601. Chowdhury, T.T., Bader, D.L., Shelton, J.C., Lee, D.A., 2003. Temporal regulation of chondrocyte metabolism in agarose constructs subjected to dynamic compression. Arch. Biochem. Biophys. 417, 105–111. De, B.C., Dell’Accio, F., Luyten, F.P., 2001. Human periosteumderived cells maintain phenotypic stability and chondrogenic potential throughout expansion regardless of donor age. Arthritis Rheum. 44, 85–95. De, B.C., Dell’Accio, F., Vanlauwe, J., Eyckmans, J., Khan, I.M., Archer, C.W., et al., 2006. Mesenchymal multipotency of adult human periosteal cells demonstrated by single-cell lineage analysis. Arthritis Rheum. 54, 1209–1221.
(
)
–
9
Delaney, J.P., O’Driscoll, S.W., Salter, R.B., 1989. Neochondrogenesis in free intraarticular periosteal autografts in an immobilized and paralyzed limb. An experimental investigation in the rabbit. Clin. Orthop. Relate. Res. 278–282. Di Guglielmo, G.M., Le Roy, C., Goodfellow, A.F., Wrana, J.L., 2003. Distinct endocytic pathways regulate TGF-f receptor signalling and turnover. Nat. Cell Biol. 5, 410–421. Diduch, D.R., Jordan, L.C., Mierisch, C.M., Balian, G., 2000. Marrow stromal cells embedded in alginate for repair of osteochondral defects. Arthroscopy 16, 571–577. Elder, S.H., Goldstein, S.A., Kimura, J.H., Soslowsky, L.J., Spengler, D.M., 2001. Chondrocyte differentiation is modulated by frequency and duration of cyclic compressive loading. Ann. Biomed. Eng. 29, 476–482. Emans, P.J., Pieper, J., Hulsbosch, M.M., Koenders, M., Kreijveld, E., Surtel, D.A., et al., 2006. Differential cell viability of chondrocytes and progenitor cells in tissue-engineered constructs following implantation into osteochondral defects. Tissue Eng. 12, 1699–1709. Fleisch, H., Reszka, A., Rodan, G.A., Rogers, M., 2003. Biophosphanates: mechanisms of action. In: Bilezikian, J.P., Raisz, L.G., Rodan, G.A. (Eds.), Principles of Bone Biology. Academic Press, San Diego, pp. 1361–1386. Freeman, P.M., Natarajan, R.N., Kimura, J.H., Andriacchi, T.P., 1994. Chondrocyte cells respond mechanically to compressive loads. J. Orthop. Res. 12, 311–320. Friedl, G., Schmidt, H., Rehak, I., Kostner, G., Schauenstein, K., Windhager, R., 2007. Undifferentiated human mesenchymal stem cells (hMSCs) are highly sensitive to mechanical strain: transcriptionally controlled early osteo-chondrogenic response in vitro. Osteoarthr. Cartilage 15, 1293–1300. Gardner, E., 1971. Osteogenesis in the human embryo and fetus. In: Broune, G.H. (Ed.), The Biochemistry and Physiology of Bone. Academic Press, New York, pp. 77–118. Goldring, M.B., Tsuchimochi, K., Ijiri, K., 2006. The control of chondrogenesis. J. Cell Biochem. 97, 33–44. Gross, T.S., Edwards, J.L., McLeod, K.J., Rubin, C.T., 1997. Strain gradients correlate with sites of periosteal bone formation. J. Bone Miner. Res. 12, 982–988. Guilak, F., 1994. Volume and surface area measurement of viable chondrocytes in situ using geometric modelling of serial confocal sections. J. Microsci. 173, 245–256. Guilak, F., Meyer, B.C., Ratcliffe, A., Mow, V.C., 1994. The effects of matrix compression on proteoglycan metabolism in articular cartilage explants. Osteoarthr. Cartilage 2, 91–101. Hanada, K., Solchaga, L.A., Caplan, A.I., Hering, T.M., Goldberg, V.M., Yoo, J.U., et al., 2001. BMP-2 induction and TGF-beta 1 modulation of rat periosteal cell chondrogenesis. J. Cell Biochem. 81, 284–294. Harada, Y., Tomita, N., Wakitani, S., Mii, Y., Oka, M., Tsutsumi, S., 2002. Use of controlled mechanical stimulation in vivo to induce cartilage layer formation on the surface of osteotomized bone. Tissue Eng. 8, 969–978. Hardingham, T.E., Oldershaw, R.A., Tew, S.R., 2006. Cartilage, SOX9 and Notch signals in chondrogenesis. J. Anat. 209, 469–480. Hatakeyama, Y., Nguyen, J., Wang, X., Nuckolls, G.H., Shum, L., 2003. Smad signaling in mesenchymal and chondroprogenitor cells. J. Bone Joint Surg. Am. 85-A, 13–18. Heywood, H.K., Sembi, P.K., Lee, D.A., Bader, D.L., 2004. Cellular utilization determines viability and matrix distribution profiles in chondrocyte-seeded alginate constructs. Tissue Eng. 10, 1467–1479. Huang, C.Y., Reuben, P.M., Cheung, H.S., 2005. Temporal expression patterns and corresponding protein inductions of early responsive genes in rabbit bone marrow-derived mesenchymal stem cells under cyclic compressive loading. Stem Cells 23, 1113–1121.
10
J O U R N A L O F T H E M E C H A N I C A L B E H AV I O R O F B I O M E D I C A L M AT E R I A L S
Hutmacher, D.W., Sittinger, M., 2003. Periosteal cells in bone tissue engineering. Tissue Eng. 9 (Suppl. 1), S45–S64. Jung, M., Gotterbarm, T., Gruettgen, A., Vilei, S.B., Breusch, S., Richter, W., 2005. Molecular characterization of spontaneous and growth-factor-augmented chondrogenesis in periosteum bone tissue transferred into a joint. Histochem. Cell Biol. 123, 447–456. Knight, M.M., Ghori, S.A., Lee, D.A., Bader, D.L., 1998. Measurement of the deformation of isolated chondrocytes in agarose subjected to cyclic compression. Med. Eng. Phys. 20, 684–688. Kosher, R.A., Kulyk, W.M., Gay, S.W., 1986. Collagen gene expression during limb cartilage differentiation. J. Cell Biol. 102, 1151–1156. LaMothe, J.M., Hamilton, N.H., Zernicke, R.F., 2005. Strain rate influences periosteal adaptation in mature bone. Med. Eng. Phys. 27, 277–284. Lee, D.A., Bader, D.L., 1997. Compressive strains at physiological frequencies influence the metabolism of chondrocytes seeded in agarose. J. Orthop. Res. 15, 181–188. Lee, D.A., Noguchi, T., Frean, S.P., Lees, P., Bader, D.L., 2000. The influence of mechanical loading on isolated chondrocytes seeded in agarose constructs. Biorheology 37, 149–161. Luettich, K., Schmidt, C., 2003. TGFbeta1 activates c-Jun and Erk1 via alphaVbeta6 integrin. Mol. Cancer 2, 33. Madras, N., Gibbs, A.L., Zhou, Y., Zandstra, P.W., Aubin, J.E., 2002. Modeling stem cell development by retrospective analysis of gene expression profiles in single progenitor-derived colonies. Stem Cells 20, 230–240. Malizos, K.N., Papatheodorou, L.K., 2005. The healing potential of the periosteum: molecular aspects. Injury 36, S13–S19. Matsumoto, T., Nakayama, K., Kodama, Y., Fuse, H., Nakamura, T., Fukumoto, S., 1998. Effect of mechanical unloading and reloading on periosteal bone formation and gene expression in tail-suspended rapidly growing rats. Bone 2, 89S–93S. Meyer, U., Meyer, T., Wiesmann, H.P., Kruse-Losler, B., Vollmer, D., Stratmann, U., et al., 2001. Mechanical tension in distraction osteogenesis regulates chondrocytic differentiation. Int. J. Oral Maxillofac. Surg. 30, 522–530. Miura, Y., Parvizi, J., Fitzsimmons, J.S., O’Driscoll, S.W., 2002. Brief exposure to high-dose transforming growth factor-beta1 enhances periosteal chondrogenesis in vitro: a preliminary report. J. Bone Joint Surg. Am. 84-A, 793–799. Miyanishi, K., Trindade, M.C., Lindsey, D.P., Beaupre, G.S., Carter, D.R., Goodman, S.B., et al., 2006a. Dose- and time-dependent effects of cyclic hydrostatic pressure on transforming growth factor-beta3-induced chondrogenesis by adult human mesenchymal stem cells in vitro. Tissue Eng. 12, 2253–2262. Miyanishi, K., Trindade, M.C., Lindsey, D.P., Beaupre, G.S., Carter, D.R., Goodman, S.B., et al., 2006b. Effects of hydrostatic pressure and transforming growth factor-beta 3 on adult human mesenchymal stem cell chondrogenesis in vitro. Tissue Eng. 12, 1419–1428. Mizuta, H., Sanyal, A., Fukumoto, T., Fitzsimmons, J.S., Matsui, N., Bolander, M.E., et al., 2002. The spatiotemporal expression of TGF-beta1 and its receptors during periosteal chondrogenesis in vitro. J. Orthop. Res. 20, 562–574. Mukherjee, N., Saris, D.B., Schultz, F.M., Berglund, L.J., An, K.N., O’Driscoll, S.W., 2001. The enhancement of periosteal chondrogenesis in organ culture by dynamic fluid pressure. J. Orthop. Res. 19, 524–530. Nakahara, H., Bruder, S.P., Goldberg, V.M., Caplan, A.I., 1990. In vivo osteochondrogenic potential of cultured cells derived from the periosteum. Clin. Orthop. Relate. Res. 223–232. O’Driscoll, S.W., 1999. Articular cartilage regeneration using periosteum. Clin. Orthop. Relate. Res. S186–S203. O’Driscoll, S.W., Recklies, A.D., Poole, A.R., 1994. Chondrogenesis in periosteal explants. An organ culture model for in vitro study. J. Bone Joint Surg. Am. 76, 1042–1051.
(
)
–
Orwoll, E.S., 2003. Toward an expanded understanding of the role of the periosteum in skeletal health. J. Bone Miner. Res. 18, 949–954. Parfitt, A.M., 2002. Parathyroid hormone and periosteal bone expansion. J. Bone Miner. Res. 17, 1741–1743. Parfitt, A.M., Travers, R., Rauch, F., Glorieux, F.H., 2000. Structural and cellular changes during bone growth in healthy children. Bone 27, 487–494. Pead, M.J., Skerry, T.M., Lanyon, L.E., 1998. Direct transformation from quiescence to bone formation in the adult periosteum following a single brief period of bone loading. J. Bone Miner. Res. 3, 647–656. Pechak, D.G., Kujawa, M.J., Caplan, A.I., 1986. Morphology of bone development and bone remodeling in embryonic chick limbs. Bone 7, 459–472. Pittenger, M.F., Mackay, A.M., Beck, S.C., Jaiswal, R.K., Douglas, R., Mosca, J.D., et al., 1999. Multilineage potential of adult human mesenchymal stem cells. Science 284, 143–147. Raab-Cullen, D.M., Thiede, M.A., Petersen, D.N., Kimmel, D.B., Recker, R.R., 1994. Mechanical loading stimulates rapid changes in periosteal gene expression. Calcif. Tissue Int. 55, 473–478. Rauch, F., 2005. Bone growth in length and width: the Yin and Yang of bone stability. J. Muscul. Neur. Interact. 5, 194–201. Ruff, C., 2003. Growth in bone strength, body size, and muscle size in a juvenile longitudinal sample. Bone 33, 317–329. Simmons, D.J., 1985. Fracture healing perspectives. Clin. Orthop. Relate. Res. 100–113. Stevens, M.M., Marini, R.P., Martin, I., Langer, R., Prasad, S.V., 2004a. FGF-2 enhances TGF-beta1-induced periosteal chondrogenesis. J. Orthop. Res. 22, 1114–1119. Stevens, M.M., Marini, R.P., Schaefer, D., Aronson, J., Langer, R., Shastri, V.P., 2005. In vivo engineering of organs: the bone bioreactor. Proc. Natl. Acad. Sci. USA 102, 11450–11455. Stevens, M.M., Qanadilo, H.F., Langer, R., Shastri, V.P., 2004b. A rapid-curing alginate gel system: utility in periosteum-derived cartilage tissue engineering. Biomaterials 25, 887–894. Stewart, K., Walsh, S., Screen, J., Jefferiss, C.M., Chainey, J., Jordan, G.R., et al., 1999. Further characterization of cells expressing STRO-1 in cultures of adult human bone marrow stromal cells. J. Bone Miner. Res. 14, 1345–1356. Taylor, J.F., 1992. The periosteum and bone growth. In: Hall, B.K. (Ed.), Bone. CRC Press, Boca Raton, pp. 21–52. Terraciano, V., Hwang, N., Moroni, L., Park, H.B., Zhang, Z., Mizrahi, J., et al., 2007. Differential response of adult and embryonic mesenchymal progenitor cells to mechanical compression in hydrogels. Stem Cells 25, 2730–2738. van der Meulen, M.C., Beaupre, G.S., Carter, D.R., 2003. Mechanobiologic influences in long bone cross-sectional growth. Bone 14, 635–642. Vanderschueren, D., Venken, K., Ophoff, J., Bouillon, R., Boonen, S., 2006. Clinical review: sex steroids and the periosteum–reconsidering the roles of androgens and estrogens in periosteal expansion. J. Clin. Endocrinol. Metab. 91, 378–382. Wang, S.E., Wu, F.Y., Shin, I., Qu, S., Arteaga, C.L., 2005. Transforming growth factor {beta} (TGF-{beta})-Smad target gene protein tyrosine phosphatase receptor type kappa is required for TGF-{beta} function. Mol. Cell Biol. 25, 4703–4715. Yakar, S., Rosen, C.J., Beamer, W.G., Ackert-Bicknell, C.L., Wu, Y.P., Liu, J.L., Ooi, G.T., Setser, J., Frystyk, J., Boisclair, Y.R., LeRoith, D., 2002. Circulating levels of IGF-1 directly regulate bone growth and density. J. Clin. Investig. 110, 771–781. Yoo, J.U., Johnstone, B., 1998. The role of osteochondral progenitor cells in fracture repair. Clin. Orthop. Relate. Res. S73–S78. Zhou, G., Zheng, Q., Engin, F., Munivez, E., Chen, Y., Sebald, E., et al., 2006. Dominance of SOX9 function over RUNX2 during skeletogenesis. Proc. Natl. Acad. Sci. USA 103, 19004–19009.
