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The response of primary articular chondrocytes to micrometric surface topography and sulphated hyaluronic acid-based matrices
Cell Biology International 29 (2005) 605e615 www.elsevier.com/locate/cellbi
The response of primary articular chondrocytes to micrometric surface topography and sulphated hyaluronic acid-based matrices D.W. Hamilton a, M.O. Riehle a,*, R. Rappuoli b, W. Monaghan c, R. Barbucci b, A.S.G. Curtis a b
a Centre for Cell Engineering, University of Glasgow, Glasgow, UK CRISMA and Department of Chemical and Biosystems Sciences and Technologies, University of Siena, Siena, Italy c Department of Electronic and Electrical Engineering, University of Glasgow, Glasgow, UK
Received 22 August 2004; revised 10 January 2005; accepted 3 March 2005
Abstract Understanding the response of chondrocytes to topographical cues and chemical patterns could provide invaluable information to advance the repair of chondral lesions. We studied the response of primary chondrocytes to nano- and micro-grooved surfaces, and sulphated hyaluronic acid (HyalS). Cells were grown on grooves ranging from 80 nm to 9 mm in depth, and from 2 mm to 20 mm in width. Observations showed that the cells did not spread appreciably on any groove size, or alter morphology or F-actin organization, although cells showed accelerated movement on 750 nm deep grooves in comparison to flat surfaces. On chemical patterns, the cells migrated onto, and preferentially attached to, HyalS and showed a greater degree of spreading and F-actin rearrangement. This study shows that 750 nm deep grooves and sulphated hyaluronic acid elicit responses from primary chondrocytes, and this could have implications for the future direction of cartilage reconstruction and orthopaedic treatments in general. Ó 2005 International Federation for Cell Biology. Published by Elsevier Ltd. All rights reserved. Keywords: Topography; Sulphated hyaluronic acid; Ovine chondrocytes; Cell adhesion; Cell migration; F-actin
1. Introduction Cartilage has only a small capacity for self-repair and has become a focus for cell and tissue engineering. Most research involves the combination of in vitro expanded chondrocytes with constructs comprised of materials such as polyglycolic acid, or polylactic acid (Vacanti et al., 1993; Freed et al., 1998). Initial seeding and subsequent cartilage development is enhanced using specially designed bioreactors. When used in such applications, chondrocytes seem unable to reproduce
* Corresponding author. Fax: C44 (0)141 330 3730. E-mail address: [email protected] (M.O. Riehle).
cartilage tissue as found in vivo; the cartilage rarely has the same morphology or biocomposition as healthy cartilage and usually breaks down (Freed et al., 1994; Temenoff and Mikos, 2000). However, in many of these studies, little attention was paid to the response of the chondrocytes to the material properties of the construct. There is a requirement to understand more about the chondrocyteematerial interaction, and more specifically, what part surface chemistry and surface topography play (Boyan et al., 1996). In attempts to advance cartilage engineering, most attention has been paid to surface chemistry, and materials such as titanium (Boyan et al., 1999), poly (L-lactic acid) (PLLA) (Cui et al., 2003), as well as natural materials such as collagen (Freyria et al., 2004), and
1065-6995/$ - see front matter Ó 2005 International Federation for Cell Biology. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.cellbi.2005.03.013
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hyaluronic acid (Barbucci et al., 2000) have been used. Although polymers such as PGA and PLLA support chondrocyte adhesion and formation of cartilaginouslike tissue (Vunjak-Novakovic et al., 1999; Cui et al., 2003), naturally occurring extracellular matrix proteins such as fibronectin have been investigated (Lawlor et al., 1996) in attempts to more accurately mimic in vivo situations. In particular, hyaluronic acid or hyaluronan (Hyal) is abundant in cartilage (Wiebkin et al., 1974), and there are many hyaluronic acid scaffolds being researched for cartilage reconstruction, which have had varying success in repairing lesions (Bloebaum et al., 1992; Aigner et al., 1998). Unfortunately Hyal alone is very unstable in physiological conditions, and many of the hyaluronic based matrices in use have been modified by adding reactive groups (Glass et al., 1996), which have been shown to alter the adhesion of some cell types (Chen et al., 1997). It has been shown in vitro that the introduction of sulphate groups in the chain of Hyal increases the resistance to degradation by hyaluronidases (Abatangelo et al., 1997). However, there is still a gap in the understanding of how sulphated hyaluronic acid (HyalS) influences chondrocyte adhesion, cytoskeletal organization and motility. In addition, when considering material characteristics, the topography of the material must also be taken into account. There have been no studies investigating chondrocyte response to topographical cues, but microfabricated grooves have proven to be an excellent model for investigating the influence of topography alone on cell behaviour (Curtis and Wilkinson, 1998). The reaction of many cell types to such substrata has been previously reported (Brunette, 1986; Wojciak-Stothard et al., 1996; Walboomers et al., 1998), and grooves have been shown to affect cell orientation, adhesion, cytoskeletal organization and motility, as well as changes in gene expression. To date, one investigation of chondrocyte reaction to topography has been reported. On titanium, it has been observed that chondrocytes react mainly to the roughness, which was the most important variable in governing adhesion, proliferation and matrix synthesis (Boyan et al., 1999). Additionally, Boyan et al. (1999) highlighted the observation that the maturation state of the chondrocytes affected how they responded to the material roughness. In the studies investigating topography, many of these studies have employed established cell lines, and there are only a few reports on how primary cells respond to topographical and chemical cues (Webb et al., 1995; Rajnicek and McCaig, 1997). Clearly, there is a gap in the understanding of primary chondrocyte response to material properties, especially given the dramatic change in the cells when grown in vitro. We report here on the response of primary ovine chondrocytes to repeating grooved substrata and to photoimmobilised patterns of sulphated hyaluronic based matrices.
2. Materials and methods 2.1. Fabrication of topography Multiple grooved substrata were prepared in fused silica slides and coverslips using photolithography methods as described in Clark et al. (1990). This method produces substrata with a uniform surface chemistry. 2.2. Fabrication of HyalS structures Poly(ethylene terephthalate) (PET) film was purchased from Akita Sumitomo Bake Co. (Akita, Japan), hyaluronic acid (Hyal, MW 200,000) was provided by F.A.B. (Fidia Advanced Biopolymers, AbanoTerme, Padova, Italy). HyalS was prepared as previously reported (Barbucci et al., 1998). Photoimmobilization was carried out by photolithography techniques. The disks were then washed with distilled water and the pattern of the immobilized polymer was confirmed to be that of the original photomask. 2.3. Cell isolation Articular chondrocytes were isolated from the metacarpalephalangeal joints of 8-month-old lambs as previously described (Hamilton et al., 2005). The cells were plated immediately on test and control surfaces at a density of 5!103 cells per cm2. 2.4. Interference reflection microscopy Interference reflection microscopy (IRM) was used to estimate cell-substrate separation as described previously (Curtis, 1964). Taking into account that the more complex predictions of Gingell and Todd (1979) would be more accurate, but since the control over refractive index of the cell will always be limited and we compared like with like, we assumed the refractive index of the cells to be similar and uniform. Primary chondrocytes were seeded on either grooved or flat control surfaces, and cellular densities were still low enough after 4 days to exclude the possibility of contact inhibition. The cells were then mounted on a cavity slide filled with a CO2 independent DMEM formulation (160 ml 25 mM HEPES water, 18 ml DMEM 10!, 20 ml foetal calf serum). The IRM was set up using a Vickers M15 microscope equipped for epifluorescence with a mercury vapour lamp and a neutral semitransparent reflector, using an interference filter (546 nm) to create a monochromatic beam of collimated light whose intensity could be adjusted using neutral density filters (to prevent damage to the cells) to illuminate the cell/substrate interface. The intensity of the reflected light in a cell-free area and the
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intensity of the light in an area with a cell was used to calculate the cellesubstrate separation according to Curtis (1964). A calibration curve was obtained using a series of ND filters to show that the system light sourceemicroscopeecameraecomputer responded linearly within the range of intensities observed within one image. The reflectivity, p, of the background is at a given wavelength determined by refractive indices of the medium (n1Z1.34) and the refractive index of the substrate (n0Z1.515) by: pZ((n1ÿn0)/(n1Cn0))1.34). To calculate the cellesubstrate separation, we used the reflectivity of the background obtained by an intensity measurement at a cell-free part of the image to map the crossover point of the reflectivity vs distance curve, and then used an intensity measurement under a cell, which was mapped onto the reflectivity vs distance curve to finally obtain the cellesurface separation at that point in nm. A spreadsheet (Excel) set up to work with this model can be obtained from the authors. Light areas of the cells correspond to the regions of the cell furthest from the substrata, and dark areas, those closest. Twenty cells were measured in four different experiments to obtain the values in Fig. 3C.
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not move continuously the average time spent, distance travelled and number of ‘‘runs’’ (uninterrupted continuous movement), as well as the number of stops, was also determined. Statistical analysis of all cell movement parameters was performed using ANOVA and post hoc testing using t-tests assuming unequal variance with a Bonferroni adjustment. The level of significance was selected at p!0.05 ( p>0.0125 for a four group comparison). 2.6. Atomic force microscopy Atomic force microscopy (AFM) was carried out by Dr Mathis Riehle using a digital Nanoscope III AFM (digital, Leatherhead, Surrey) in the contact mode using air-dried samples.
2.5. Time-lapse observations and analysis of cell movement In order to measure the dynamics of cell movement, time lapse video recordings were taken of chondrocytes on the various grooved substrates using a 10! objective on either a Leitz Diavert or a Leitz Labovert microscope and a COHU 4095 high performance CCD camera and stored on S-VHS tape at 1 frame per minute. The video was digitized (1 frame/3 h real time) and analysis performed on a Macintosh computer using the public domain NIH image program (developed at the US National Institutes of Health and available on the internet at http://rsb.info.nih.gov/nihimage). The x,y coordinates of the cells were determined interactively using NIH image. The resulting x,y coordinates of 20 cells, from three independent experiments, meant that over 60 time-points each were combined and used for the calculations described below. The time-lapse videos were analyzed using a designated spreadsheet created by Dr Mathis Riehle (freely available for download at http://www.gla.ac.uk/centres/ cellengineering/mathis) and by visual observations. The analysis yielded the following data: the number of cells moving, the time the cells spent moving, the distance covered, the integrated path length, the average velocity (inclusive and exclusive of periods where the cells had stopped), the time the cells spent not moving (stopping time), the number of stops and the overall, as well as the integrated direction of the movement, and the average angle of change in direction. Since cells do
Fig. 1. Orientation of cells on plain and grooved silica. (A) Effect of groove width and (B) groove depth; 90 would be aligned to the long axis of the groove. Box plot shows the distribution of angle of cell axis with respect to the groove long axis. Central bar represents the mean, the box contains 50% of the data, the barsZ75% data and 90% of the data lies within the outer points.
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2.7. Scanning electron microscopy Samples were prepared for electron microscopy as previously described (Bell, 1981). The samples were then critical point dried, mounted on aluminium stubs and sputter coated with gold. Following this, the samples were viewed using a Philips 500 scanning electron microscope at 3 kV accelerating voltage. 2.8. Indirect immunofluorescence and F-actin staining Chondrocytes were plated on flat and grooved structures at a concentration of 5!103cells/ml and fixed after 96 h for F-actin staining. For CD44 staining, cells were plated on HyalS structures at the same density and fixed after 96 h. The medium was removed and the attached cells and structures washed three times for 5 min using warm (37 C) PBS. Cells were then fixed in 4% formalin (Sigma, Poole, UK) in PBS for 5 min, and permeabilised with 0.5% Triton X-100 in PBS for 15 min. F-actin staining and immunostaining were performed as previously described (Wojciak-Stothard et al., 1996). Following staining, all samples were washed three times in PBS and mounted in mowiol. Fluorescent preparations were examined using a confocal laser microscope
or a Hamamatsu CCD camera on a Vickers M15 fluorescent microscope.
3. Results 3.1. Primary chondrocyte adhesion, morphology and spreading Primary chondrocytes showed no significant alignment to grooves of any width or depth in the ranges tested in these experiments (Fig. 1A,B). There was a large spread of angles of the cell axis with respect to the groove long axis. Scanning electron microscope examination and time lapse video observations further demonstrated that chondrocytes did not alter their spherical morphology (Fig. 2A). However, at 96 h, some cells on the flat control surface demonstrated a slight degree of spreading (Fig. 2B,C). Furthermore, any cell in contact with another also remained spherical in shape (Fig. 2D). Interference reflection microscopy demonstrated that, on flat and grooved substrates, there was a predominance of light areas as opposed to dark areas (Fig. 3A,B); areas of close contact were confined to small areas. The values calculated for the furthest (light) and closest (dark) areas of the cells are shown in Fig. 3C.
Fig. 2. Scanning electron microscope images of primary chondrocytes grown on flat control surfaces. (A) Chondrocytes after 24 h in culture. In (B) and (C), the cells have been cultured for 96 h and are beginning to spread on the surface. In (D), chondrocytes maintain their spherical morphology if in contact with another cell (shown by arrow), rather than the substrate. In all cases, the bar is 5 mm (original magnification !400).
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3.2. Time-lapse observations and analysis of cell movement Observations of the time-lapse video confirmed no change in the spherical shape of the cells during movement. Guidance of the cells movement along the groove long axis was evident on nanometric topography. Fig. 4AeF shows the movement values measured for the cells on each surface tested. In summary, primary chondrocytes show the largest increased response in movement to grooves of 750 nm depth. There was a higher percentage of cells moving at any one time, and the average run length was longer and the stop time shorter than on any other surface tested. The velocity of the cells was not significantly different and the average distance travelled per run was similar to that seen on the other surfaces. On grooves 3 mm deep, the cell response was decreased below the level of movement evident on control surfaces, cells on 8 mm gratings were not significantly different from controls. 3.3. F-actin distribution and chondrocyte phenotype Chondrocytes remained round on experimental and control surfaces and F-actin was on all surfaces, distributed around the cell margins (Fig. 5A), and there was no extension of lamellapodia or microspikes. There was some evidence of cell spreading on flat surfaces by 96 h, with F-actin distributed round the edge of the cell, although no stress-fibre formation or F-actin condensation along the groove/ridge boundaries was evident (Fig. 5B). In addition, all cells maintained type II collagen expression over the duration of the experiments (data not shown).
Fig. 3. Interference reflection images of primary chondrocytes on (A) flat silica coverslips, (B) 80 nm deep grooves, and (C) the distance between the ventral surface of primary chondrocytes and the substrate below. Values are the mean of 20 cells from four experimentsG1 standard deviation.
4. Discussion 3.4. Chondrocyte response to PET and sulphated hyaluronic acid AFM analysis demonstrated that the average height of the patterned HyalS was 1 mm (Fig. 6A). Time-lapse video recording demonstrated a migration from PET surface to bands of HyalS 48 h after plating out. Significantly more cells were present on HyalS after 48 h and there was an overall increase in cell number after 72 h (Fig. 6B,C). Analysis of cell movement demonstrated that cells on HyalS migrated significantly shorter distances and for shorter run times than cells on PET (Fig. 7A,B). F-actin staining of cells on HyalS showed that they had leading lamellapodia (Fig. 7DeG). Staining for CD44 adhesion molecule (Fig. 8AeC) revealed that receptors were more numerous in cells on HyalS and were located around the cell edge. Cells on PET exhibited fewer receptors and those present were located in the centre of the cells. There was also evidence of matrix production by the chondrocytes on HyalS (Fig. 8D).
Synonymous with growth in tissue culture, chondrocytes undergo a phenotypic change from a near spherical form to an elongated fibroblastic type with a loss of markers, including type II collagen and aggrecan (Kuroda, 1964; Von der Mark, 1986; Temenoff and Mikos, 2000). Previous studies have shown that cells such as macrophages, osteoblasts, neurons, epithelial cells and fibroblasts (Curtis and Wilkinson, 1998) can sense and respond to artificial micrometric topography and exhibit changes in cell morphology, orientation, cytoskeletal organization, proliferation and gene expression. We hypothesized that topographical cues, such as repeating grooved substrata as well as patterned HyalS, may provide an alternative way to investigate primary chondrocyte response to material characteristics. In this paper, we report on the adhesion, morphology, migration and cytoskeletal organization of primary chondrocytes on surfaces with three different types of chemistry and three types of topography.
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Fig. 4. Analysis of primary chondrocyte movement over a period of 20 h after initial seeding. (A) Percentage of cells moving, (B) average time on the run, (C) average velocity, (D) average distance travelled, (E) number of stops, and (F) average length of stops. *Comparison with flat surface, P!0.0125; t-test assuming unequal variance with a Bonferroni adjustment.
We show here that primary chondrocytes maintain the spherical shape associated with their phenotype over at least 4 days of culture, and show no evidence of F-actin reorganization, which would be expected if phenotype change had taken place (Lawlor et al., 1996). Analysis of cell movement covered the percentage of cells moving, average velocity, average run and stop time, persistence of movement and number of stops over a 24 h culture period. It was noted that cells moved on all
groove depths, but responded most significantly to grooves 750 nm deep in terms of distance moved and average time spent moving. Moreover, grooves with a depth of 3 mm or more reduced cell movement in all aspects investigated in comparison to control surface. The average transition time of the chondrocytes on all surfaces was once per hour, but on 3 mm deep grooves this dropped to once every 2 h, indicating a strong inhibition of movement on this surface (data not shown).
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Fig. 5. (A) F-actin arrangement in cells on plain silica (column A) and grooved silica (column B), 5 mm wide, 3 mm deep after 24 and 96 h. There is no evident change in the F-actin arrangement in cells cultured on flat or grooved substrates for 4 days. BarZ3 mm. (B) F-actin arrangement in a spread primary chondrocyte 96 h after plating on a flat surface. F-actin is dispersed and is not as prominent in the periphery (compare with A). Membrane ruffles are evident around the edge of the cell.
It is interesting that the cells migrated most on the 750 nm deep grooves. Although the cells did not change to an elongated morphology, they migrated relatively rapidly, especially on 750 nm deep silica grooves with relatively good guidance in the direction of the groove long axis. Curtis et al. (1995) observed that on topographical cues, such as repeating grooves, cells must align before they begin to move, and that the initiation of movement appears to be related to cytoskeletal reorganization. On grooves, chondrocytes did not alter their spherical shape or F-actin arrangement, and very few microspikes or subsequent lamellapodia were observed. It is possible that, when in the spherical morphology, chondrocytes are not able to form adhesions, other than those directly under the cell (as shown by IRM); the cells are not sufficiently flexible. The movement exhibited by primary chondrocytes is similar to that which has been reported for neutrophils (Mandeville et al., 1997). The focal adhesions of the cells are confined to a very small percentage of their apparent contact area. It seems that the cells may teeter along the grooves or ridges. Clearly morphological and
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cytoskeletal organizations are not a pre-requisite for migration on repeating grooved substrata. This type of motion exhibited by primary chondrocytes and neutrophils has received very little attention to date. This type of movement is very different from that which has been reported for other cell types, such as endothelia, dorsal root ganglion neurons, and epitenon fibroblasts (Curtis et al., 1995). There are only a few studies that have investigated chondrocyte movement, mostly in response to cytokines, such as hepatocyte scatter factor, which are known to promote migration in other cell types (Takebayashi et al., 1995). Only one study assessed the in vitro movement of primary chondrocytes, and it demonstrated that approximately 30% of the total cell number was capable of migration (Chang et al., 2003). In that study, the chondrocytes were given 3 days to recover from enzymatic digestion. However, in this current investigation, we seeded the cells directly on the substrate after isolation from cartilage and 70% of the cells exhibited migration on 750 nm deep grooves after 4 h. It is clear from this investigation that certain chondrocyte populations can migrate if grown on suitable substrata in vitro, and further investigations should be aimed at identifying the phenotype of these particular cells, which could potentially be of great benefit in orthopaedic applications in the future. In addition to assessing the chondrocyte base response to materials, maintenance of the chondrocyte phenotype is also of great importance; F-actin has been associated with its loss (Von der Mark, 1986). Previous studies have shown that chondrocytes do not produce type I collagen until they have stress-fibre networks (Loeser and Lee, 1997), a cytoskeletal arrangement not seen in situ (Langelier et al., 2000). When primary chondrocytes were labelled for F-actin, the arrangement was the same on both control and grooved surfaces 96 h after plating. As it was the same in situ, it is possible that the cells require a period of time to alter the cytoskeleton to be able to respond. F-actin is known to be central in cellular reactions to topographies such as repeating grooves (Curtis et al., 1995), thus the lack of elongation and alignment of chondrocytes to grooved surfaces in comparison with other cell types (Curtis and Wilkinson, 1998) is not unexpected. In response to chemical patterns, primary chondrocytes preferentially migrated from the PET areas to the HyalS, where the cells spread and adopted a more elongated morphology. This migration is unlikely to have been due to chemotaxis, because the material was thoroughly washed free of soluble products in its preparation. Two alternative, but related, explanations seem more likely. The first is that the HyalS is so adhesive that any cell moving randomly onto it cannot leave (a fly paper effect) and the second is that the edge topography of the HyalS (1 mm in height) is not conducive to
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Fig. 6. (A) AFM image showing height profile of HyalS. (B) Coomassie blue staining of primary chondrocytes on PET and HyalS (darker stripes) after 96 h. The cells cluster on the HyalS but not on PET. BarZ100 mm. (B) The same preparation at a higher magnification. The cells are more spread on HyalS compared to their appearance on PET. Arrow indicates cells that have aligned to the boundary between substrates. BarZ50 mm. (C) Number of primary chondrocytes adhering to PET and HyalS surfaces over a 96 h period. BarsZstandard deviation, *P!0.01, Student’s paired t-test.
re-emigration to the PET surface. With regard to the morphological changes of primary chondrocytes on HyalS, this is further evidence of the inability of chemical patterns to prevent the spreading associated with loss of the chondrogenic phenotype (Loeser and Lee, 1997). Given that cells remain round on PET, it would appear to be a better substrate on which to maintain the chondrocyte phenotype. However, the alteration in CD44 receptor organization on HyalS and the increased matrix production (see Fig. 8C) suggests that the material can still be recognized by the cells even although it is modified. It is known that chondrocytes bind to hyaluronan through CD44 (Ishida et al., 1997), so the
addition of sulphate groups appears not to alter the recognition of the hyaluronan unit by the chondrocytes. When considering the response of cells to substrata used in this study, a common criticism is whether or not the cells are responding to the topography or the chemistry of the material. The HyalS bands are 1 mm in height, as observed using AFM. Cells are known to align to repeating grooved structures with depths significantly lower than this (Curtis and Wilkinson, 1998). It is relevant that there are a number of cells concentrated at the junction of the PET and the HyalS, and this is likely to be a topographic reaction. This is another example of a general finding commented on by
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Fig. 7. Cell movement of cells on PET and HyalS. (A) Average distance travelled and (B) average time on the run. *P!0.05, Student’s paired t-test. (C) Number of cells crossing the boundaries between HyalS and PET over a 96 h time frame. Data shows a directed migration from PET to HyalS. Error barsZ5% standard error. F-actin arrangement in primary chondrocytes grown on (E,F) photoimmobilised HyalS and (G,H) PET. Cells on HyalS showed a higher degree of spreading than those on PET. Bars in (E) and (F) are 10 mm, in (G) and (H) 5 mm.
Curtis and Wilkinson (1998) and Riehle et al. (2002). The effect of the topography of chemical patterns on cell behaviour has been argued in general terms (Curtis and Wilkinson, 1998), but, although there are strong arguments for topographic effects, we cannot yet exclude the possibility of chemical ones. These types of effects could
again be most valuable in locating chondrocytes to specific positions in constructs. With regard to the silica grooves, the cell response is likely to be topographical in nature because of the uniform surface chemistry that such surfaces probably have, due to blanket etching (Clark et al., 1990).
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Fig. 8. CD44 receptor arrangement on primary chondrocytes cultured on (A) PET and (B) HyalS. For explanation, see text. (C) SEM micrograph of primary chondrocytes on HyalS. Arrow indicates matrix production by the cells, which are encased in the matrix. BarZ5 mm.
It is clear that chondrocyte response to materials needs further investigation. Chemical and topographical cues are clearly important, but this investigation demonstrates that neither is more dominant than the other, both capable of altering cell behaviour. In selecting biomaterials for cartilage repair, surface structure and chemistry must be investigated to optimize the response of the cells, which in turn could aid ex vivo cartilage tissue development.
Acknowledgements The authors would like to thank Mr E. Robertson and Mrs M. Mullin for their help with the SEM images. The authors would also like to thank Sandyford Abattoir, Paisley, for kindly supplying all the lambs legs which provided the cell material used in this work. This work was supported by the EPSRC and Smith and Nephew plc, York.
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Glass JR, Dickerson KT, Stecker K, Polarek JW. Characterization of a hyaluronic acid-Arg-Gly-Asp peptide cell attachment matrix. Biomaterials 1996;17(11):1101e8. Hamilton DW, Riehle MO, Monaghan W, Curtis AS. Articular chondrocyte passage number: influence on adhesion, migration, cytoskeletal organization and phenotype in response to micro- and nanometric topography. Cell Biol Int 2005; in press. Ishida O, Tanaka Y, Morimoto I, Takigawa M, Eto S. Chondrocytes are regulated by cellular adhesion through CD44 and hyaluronic acid pathway. J Bone Miner Res 1997;12(10):1657e63. Kuroda Y. Studies on cartilage cells in vitro. 1. Morphology and growth of cartilage cells in monolayer culture. Exp Cell Res 1964;35:326e36. Langelier E, Suetterlin R, Hoemann CD, Aebi U, Buschmann MD. The chondrocyte cytoskeleton in mature articular cartilage: structure and distribution of actin, tubulin and vimentin filaments. J Histochem Cytochem 2000;48(10):1307e20. Lawlor P, Bebbington A, Ireland GW, Archer CW. Rate of chondrocyte modulation on serum adhesive proteins is mediated by cell shape. Cell Eng 1996;1(4):159e67. Loeser RF, Lee GM. Adhesion mediated signaling in articular chondrocytes. Mol Biol Cell 1997;8S:400a. Mandeville JTH, Lawson MA, Maxfield FR. Dynamic imaging of neutrophil migration in three dimensions: mechanical interactions between cells and matrix. J Leukoc Biol 1997;61(2):188e200. Rajnicek AM, McCaig CD. Guidance of CNS growth cones by substratum grooves and ridges: effects of inhibitors of the cytoskeleton, calcium channels and signal transduction pathways. J Cell Sci 1997;110(23):2915e24. Riehle M, Dalby MJ, Johnstone H, Gallagher JO, Wood MA, Casey B, et al. Nanometric surface patterns for tissue engineering: fabrication and biocompatibility in vitro. In: Merhari KEGL, Dobisz EA, Angelopoulos M, Herr D, editors. Nanopatterning: from ultralarge-scale integration to biotechnology, MRS Proceedings, 705. Takebayashi T, Iwamoto M, Jikko A, Matsumura T, EnomotoIwamoto M, Myoukai F, et al. Hepatocyte growth factor/scatter factor modulates cell motility, proliferation, and proteoglycan synthesis of chondrocytes. J Cell Biol 1995;129(5):1411e9. Temenoff JS, Mikos AG. Review: Tissue engineering for regeneration of articular cartilage. Biomaterials 2000;21:431e40. Vacanti CA, Kim W, Upton J, Vacanti MP, Mooney D, Schloo B, et al. Tissue-engineered growth of bone and cartilage. Transplant Proc 1993;25:1019e21. Von der Mark K. Differentiation, modulation and dedifferentiation of chondrocytes. Rheumatology 1986;10:272e315. Vunjak-Novakovic G, Martin I, Obradovic B, Treppo S, Grodzinsky AJ, Langer R, et al. Bioreactor cultivation conditions modulate the composition and mechanical properties of tissue-engineered cartilage. J Orthop Res 1999;17(1):130e8. Walboomers XF, Croes HJE, Ginsel LA, Jansen JA. Growth behavior of fibroblasts on microgrooved polystyrene. Biomaterials 1998; 19(20):1861e8. Webb A, Clark P, Skepper J, Compston A, Wood A. Guidance of oligodendrocytes and their progenitors by substratum topography. J Cell Sci 1995;108(8):2747e60. Wiebkin OW, Hardingham TE, Muir H. The interaction of proteoglycans and hyaluronic acid and the effect of hyaluronic acid on proteoglycan synthesis by chondrocytes of adult cartilage. In: Burleigh PM, Poole AR, editors. Dynamics of connective tissue macromolecules. Amsterdam: North-Holland; 1975. p. 82e104. (QU 532.C7 S989d 1974). Wojciak-Stothard B, Curtis A, Monaghan W, MacDonald K, Wilkinson C. Guidance and activation of murine macrophages by nanometric scale topography. Exp Cell Res 1996;223(2):426e35.
The response of primary articular chondrocytes to micrometric surface topography and sulphated hyaluronic acid-based matrices D.W. Hamilton a, M.O. Riehle a,*, R. Rappuoli b, W. Monaghan c, R. Barbucci b, A.S.G. Curtis a b
a Centre for Cell Engineering, University of Glasgow, Glasgow, UK CRISMA and Department of Chemical and Biosystems Sciences and Technologies, University of Siena, Siena, Italy c Department of Electronic and Electrical Engineering, University of Glasgow, Glasgow, UK
Received 22 August 2004; revised 10 January 2005; accepted 3 March 2005
Abstract Understanding the response of chondrocytes to topographical cues and chemical patterns could provide invaluable information to advance the repair of chondral lesions. We studied the response of primary chondrocytes to nano- and micro-grooved surfaces, and sulphated hyaluronic acid (HyalS). Cells were grown on grooves ranging from 80 nm to 9 mm in depth, and from 2 mm to 20 mm in width. Observations showed that the cells did not spread appreciably on any groove size, or alter morphology or F-actin organization, although cells showed accelerated movement on 750 nm deep grooves in comparison to flat surfaces. On chemical patterns, the cells migrated onto, and preferentially attached to, HyalS and showed a greater degree of spreading and F-actin rearrangement. This study shows that 750 nm deep grooves and sulphated hyaluronic acid elicit responses from primary chondrocytes, and this could have implications for the future direction of cartilage reconstruction and orthopaedic treatments in general. Ó 2005 International Federation for Cell Biology. Published by Elsevier Ltd. All rights reserved. Keywords: Topography; Sulphated hyaluronic acid; Ovine chondrocytes; Cell adhesion; Cell migration; F-actin
1. Introduction Cartilage has only a small capacity for self-repair and has become a focus for cell and tissue engineering. Most research involves the combination of in vitro expanded chondrocytes with constructs comprised of materials such as polyglycolic acid, or polylactic acid (Vacanti et al., 1993; Freed et al., 1998). Initial seeding and subsequent cartilage development is enhanced using specially designed bioreactors. When used in such applications, chondrocytes seem unable to reproduce
* Corresponding author. Fax: C44 (0)141 330 3730. E-mail address: [email protected] (M.O. Riehle).
cartilage tissue as found in vivo; the cartilage rarely has the same morphology or biocomposition as healthy cartilage and usually breaks down (Freed et al., 1994; Temenoff and Mikos, 2000). However, in many of these studies, little attention was paid to the response of the chondrocytes to the material properties of the construct. There is a requirement to understand more about the chondrocyteematerial interaction, and more specifically, what part surface chemistry and surface topography play (Boyan et al., 1996). In attempts to advance cartilage engineering, most attention has been paid to surface chemistry, and materials such as titanium (Boyan et al., 1999), poly (L-lactic acid) (PLLA) (Cui et al., 2003), as well as natural materials such as collagen (Freyria et al., 2004), and
1065-6995/$ - see front matter Ó 2005 International Federation for Cell Biology. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.cellbi.2005.03.013
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hyaluronic acid (Barbucci et al., 2000) have been used. Although polymers such as PGA and PLLA support chondrocyte adhesion and formation of cartilaginouslike tissue (Vunjak-Novakovic et al., 1999; Cui et al., 2003), naturally occurring extracellular matrix proteins such as fibronectin have been investigated (Lawlor et al., 1996) in attempts to more accurately mimic in vivo situations. In particular, hyaluronic acid or hyaluronan (Hyal) is abundant in cartilage (Wiebkin et al., 1974), and there are many hyaluronic acid scaffolds being researched for cartilage reconstruction, which have had varying success in repairing lesions (Bloebaum et al., 1992; Aigner et al., 1998). Unfortunately Hyal alone is very unstable in physiological conditions, and many of the hyaluronic based matrices in use have been modified by adding reactive groups (Glass et al., 1996), which have been shown to alter the adhesion of some cell types (Chen et al., 1997). It has been shown in vitro that the introduction of sulphate groups in the chain of Hyal increases the resistance to degradation by hyaluronidases (Abatangelo et al., 1997). However, there is still a gap in the understanding of how sulphated hyaluronic acid (HyalS) influences chondrocyte adhesion, cytoskeletal organization and motility. In addition, when considering material characteristics, the topography of the material must also be taken into account. There have been no studies investigating chondrocyte response to topographical cues, but microfabricated grooves have proven to be an excellent model for investigating the influence of topography alone on cell behaviour (Curtis and Wilkinson, 1998). The reaction of many cell types to such substrata has been previously reported (Brunette, 1986; Wojciak-Stothard et al., 1996; Walboomers et al., 1998), and grooves have been shown to affect cell orientation, adhesion, cytoskeletal organization and motility, as well as changes in gene expression. To date, one investigation of chondrocyte reaction to topography has been reported. On titanium, it has been observed that chondrocytes react mainly to the roughness, which was the most important variable in governing adhesion, proliferation and matrix synthesis (Boyan et al., 1999). Additionally, Boyan et al. (1999) highlighted the observation that the maturation state of the chondrocytes affected how they responded to the material roughness. In the studies investigating topography, many of these studies have employed established cell lines, and there are only a few reports on how primary cells respond to topographical and chemical cues (Webb et al., 1995; Rajnicek and McCaig, 1997). Clearly, there is a gap in the understanding of primary chondrocyte response to material properties, especially given the dramatic change in the cells when grown in vitro. We report here on the response of primary ovine chondrocytes to repeating grooved substrata and to photoimmobilised patterns of sulphated hyaluronic based matrices.
2. Materials and methods 2.1. Fabrication of topography Multiple grooved substrata were prepared in fused silica slides and coverslips using photolithography methods as described in Clark et al. (1990). This method produces substrata with a uniform surface chemistry. 2.2. Fabrication of HyalS structures Poly(ethylene terephthalate) (PET) film was purchased from Akita Sumitomo Bake Co. (Akita, Japan), hyaluronic acid (Hyal, MW 200,000) was provided by F.A.B. (Fidia Advanced Biopolymers, AbanoTerme, Padova, Italy). HyalS was prepared as previously reported (Barbucci et al., 1998). Photoimmobilization was carried out by photolithography techniques. The disks were then washed with distilled water and the pattern of the immobilized polymer was confirmed to be that of the original photomask. 2.3. Cell isolation Articular chondrocytes were isolated from the metacarpalephalangeal joints of 8-month-old lambs as previously described (Hamilton et al., 2005). The cells were plated immediately on test and control surfaces at a density of 5!103 cells per cm2. 2.4. Interference reflection microscopy Interference reflection microscopy (IRM) was used to estimate cell-substrate separation as described previously (Curtis, 1964). Taking into account that the more complex predictions of Gingell and Todd (1979) would be more accurate, but since the control over refractive index of the cell will always be limited and we compared like with like, we assumed the refractive index of the cells to be similar and uniform. Primary chondrocytes were seeded on either grooved or flat control surfaces, and cellular densities were still low enough after 4 days to exclude the possibility of contact inhibition. The cells were then mounted on a cavity slide filled with a CO2 independent DMEM formulation (160 ml 25 mM HEPES water, 18 ml DMEM 10!, 20 ml foetal calf serum). The IRM was set up using a Vickers M15 microscope equipped for epifluorescence with a mercury vapour lamp and a neutral semitransparent reflector, using an interference filter (546 nm) to create a monochromatic beam of collimated light whose intensity could be adjusted using neutral density filters (to prevent damage to the cells) to illuminate the cell/substrate interface. The intensity of the reflected light in a cell-free area and the
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intensity of the light in an area with a cell was used to calculate the cellesubstrate separation according to Curtis (1964). A calibration curve was obtained using a series of ND filters to show that the system light sourceemicroscopeecameraecomputer responded linearly within the range of intensities observed within one image. The reflectivity, p, of the background is at a given wavelength determined by refractive indices of the medium (n1Z1.34) and the refractive index of the substrate (n0Z1.515) by: pZ((n1ÿn0)/(n1Cn0))1.34). To calculate the cellesubstrate separation, we used the reflectivity of the background obtained by an intensity measurement at a cell-free part of the image to map the crossover point of the reflectivity vs distance curve, and then used an intensity measurement under a cell, which was mapped onto the reflectivity vs distance curve to finally obtain the cellesurface separation at that point in nm. A spreadsheet (Excel) set up to work with this model can be obtained from the authors. Light areas of the cells correspond to the regions of the cell furthest from the substrata, and dark areas, those closest. Twenty cells were measured in four different experiments to obtain the values in Fig. 3C.
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not move continuously the average time spent, distance travelled and number of ‘‘runs’’ (uninterrupted continuous movement), as well as the number of stops, was also determined. Statistical analysis of all cell movement parameters was performed using ANOVA and post hoc testing using t-tests assuming unequal variance with a Bonferroni adjustment. The level of significance was selected at p!0.05 ( p>0.0125 for a four group comparison). 2.6. Atomic force microscopy Atomic force microscopy (AFM) was carried out by Dr Mathis Riehle using a digital Nanoscope III AFM (digital, Leatherhead, Surrey) in the contact mode using air-dried samples.
2.5. Time-lapse observations and analysis of cell movement In order to measure the dynamics of cell movement, time lapse video recordings were taken of chondrocytes on the various grooved substrates using a 10! objective on either a Leitz Diavert or a Leitz Labovert microscope and a COHU 4095 high performance CCD camera and stored on S-VHS tape at 1 frame per minute. The video was digitized (1 frame/3 h real time) and analysis performed on a Macintosh computer using the public domain NIH image program (developed at the US National Institutes of Health and available on the internet at http://rsb.info.nih.gov/nihimage). The x,y coordinates of the cells were determined interactively using NIH image. The resulting x,y coordinates of 20 cells, from three independent experiments, meant that over 60 time-points each were combined and used for the calculations described below. The time-lapse videos were analyzed using a designated spreadsheet created by Dr Mathis Riehle (freely available for download at http://www.gla.ac.uk/centres/ cellengineering/mathis) and by visual observations. The analysis yielded the following data: the number of cells moving, the time the cells spent moving, the distance covered, the integrated path length, the average velocity (inclusive and exclusive of periods where the cells had stopped), the time the cells spent not moving (stopping time), the number of stops and the overall, as well as the integrated direction of the movement, and the average angle of change in direction. Since cells do
Fig. 1. Orientation of cells on plain and grooved silica. (A) Effect of groove width and (B) groove depth; 90 would be aligned to the long axis of the groove. Box plot shows the distribution of angle of cell axis with respect to the groove long axis. Central bar represents the mean, the box contains 50% of the data, the barsZ75% data and 90% of the data lies within the outer points.
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2.7. Scanning electron microscopy Samples were prepared for electron microscopy as previously described (Bell, 1981). The samples were then critical point dried, mounted on aluminium stubs and sputter coated with gold. Following this, the samples were viewed using a Philips 500 scanning electron microscope at 3 kV accelerating voltage. 2.8. Indirect immunofluorescence and F-actin staining Chondrocytes were plated on flat and grooved structures at a concentration of 5!103cells/ml and fixed after 96 h for F-actin staining. For CD44 staining, cells were plated on HyalS structures at the same density and fixed after 96 h. The medium was removed and the attached cells and structures washed three times for 5 min using warm (37 C) PBS. Cells were then fixed in 4% formalin (Sigma, Poole, UK) in PBS for 5 min, and permeabilised with 0.5% Triton X-100 in PBS for 15 min. F-actin staining and immunostaining were performed as previously described (Wojciak-Stothard et al., 1996). Following staining, all samples were washed three times in PBS and mounted in mowiol. Fluorescent preparations were examined using a confocal laser microscope
or a Hamamatsu CCD camera on a Vickers M15 fluorescent microscope.
3. Results 3.1. Primary chondrocyte adhesion, morphology and spreading Primary chondrocytes showed no significant alignment to grooves of any width or depth in the ranges tested in these experiments (Fig. 1A,B). There was a large spread of angles of the cell axis with respect to the groove long axis. Scanning electron microscope examination and time lapse video observations further demonstrated that chondrocytes did not alter their spherical morphology (Fig. 2A). However, at 96 h, some cells on the flat control surface demonstrated a slight degree of spreading (Fig. 2B,C). Furthermore, any cell in contact with another also remained spherical in shape (Fig. 2D). Interference reflection microscopy demonstrated that, on flat and grooved substrates, there was a predominance of light areas as opposed to dark areas (Fig. 3A,B); areas of close contact were confined to small areas. The values calculated for the furthest (light) and closest (dark) areas of the cells are shown in Fig. 3C.
Fig. 2. Scanning electron microscope images of primary chondrocytes grown on flat control surfaces. (A) Chondrocytes after 24 h in culture. In (B) and (C), the cells have been cultured for 96 h and are beginning to spread on the surface. In (D), chondrocytes maintain their spherical morphology if in contact with another cell (shown by arrow), rather than the substrate. In all cases, the bar is 5 mm (original magnification !400).
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3.2. Time-lapse observations and analysis of cell movement Observations of the time-lapse video confirmed no change in the spherical shape of the cells during movement. Guidance of the cells movement along the groove long axis was evident on nanometric topography. Fig. 4AeF shows the movement values measured for the cells on each surface tested. In summary, primary chondrocytes show the largest increased response in movement to grooves of 750 nm depth. There was a higher percentage of cells moving at any one time, and the average run length was longer and the stop time shorter than on any other surface tested. The velocity of the cells was not significantly different and the average distance travelled per run was similar to that seen on the other surfaces. On grooves 3 mm deep, the cell response was decreased below the level of movement evident on control surfaces, cells on 8 mm gratings were not significantly different from controls. 3.3. F-actin distribution and chondrocyte phenotype Chondrocytes remained round on experimental and control surfaces and F-actin was on all surfaces, distributed around the cell margins (Fig. 5A), and there was no extension of lamellapodia or microspikes. There was some evidence of cell spreading on flat surfaces by 96 h, with F-actin distributed round the edge of the cell, although no stress-fibre formation or F-actin condensation along the groove/ridge boundaries was evident (Fig. 5B). In addition, all cells maintained type II collagen expression over the duration of the experiments (data not shown).
Fig. 3. Interference reflection images of primary chondrocytes on (A) flat silica coverslips, (B) 80 nm deep grooves, and (C) the distance between the ventral surface of primary chondrocytes and the substrate below. Values are the mean of 20 cells from four experimentsG1 standard deviation.
4. Discussion 3.4. Chondrocyte response to PET and sulphated hyaluronic acid AFM analysis demonstrated that the average height of the patterned HyalS was 1 mm (Fig. 6A). Time-lapse video recording demonstrated a migration from PET surface to bands of HyalS 48 h after plating out. Significantly more cells were present on HyalS after 48 h and there was an overall increase in cell number after 72 h (Fig. 6B,C). Analysis of cell movement demonstrated that cells on HyalS migrated significantly shorter distances and for shorter run times than cells on PET (Fig. 7A,B). F-actin staining of cells on HyalS showed that they had leading lamellapodia (Fig. 7DeG). Staining for CD44 adhesion molecule (Fig. 8AeC) revealed that receptors were more numerous in cells on HyalS and were located around the cell edge. Cells on PET exhibited fewer receptors and those present were located in the centre of the cells. There was also evidence of matrix production by the chondrocytes on HyalS (Fig. 8D).
Synonymous with growth in tissue culture, chondrocytes undergo a phenotypic change from a near spherical form to an elongated fibroblastic type with a loss of markers, including type II collagen and aggrecan (Kuroda, 1964; Von der Mark, 1986; Temenoff and Mikos, 2000). Previous studies have shown that cells such as macrophages, osteoblasts, neurons, epithelial cells and fibroblasts (Curtis and Wilkinson, 1998) can sense and respond to artificial micrometric topography and exhibit changes in cell morphology, orientation, cytoskeletal organization, proliferation and gene expression. We hypothesized that topographical cues, such as repeating grooved substrata as well as patterned HyalS, may provide an alternative way to investigate primary chondrocyte response to material characteristics. In this paper, we report on the adhesion, morphology, migration and cytoskeletal organization of primary chondrocytes on surfaces with three different types of chemistry and three types of topography.
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Fig. 4. Analysis of primary chondrocyte movement over a period of 20 h after initial seeding. (A) Percentage of cells moving, (B) average time on the run, (C) average velocity, (D) average distance travelled, (E) number of stops, and (F) average length of stops. *Comparison with flat surface, P!0.0125; t-test assuming unequal variance with a Bonferroni adjustment.
We show here that primary chondrocytes maintain the spherical shape associated with their phenotype over at least 4 days of culture, and show no evidence of F-actin reorganization, which would be expected if phenotype change had taken place (Lawlor et al., 1996). Analysis of cell movement covered the percentage of cells moving, average velocity, average run and stop time, persistence of movement and number of stops over a 24 h culture period. It was noted that cells moved on all
groove depths, but responded most significantly to grooves 750 nm deep in terms of distance moved and average time spent moving. Moreover, grooves with a depth of 3 mm or more reduced cell movement in all aspects investigated in comparison to control surface. The average transition time of the chondrocytes on all surfaces was once per hour, but on 3 mm deep grooves this dropped to once every 2 h, indicating a strong inhibition of movement on this surface (data not shown).
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Fig. 5. (A) F-actin arrangement in cells on plain silica (column A) and grooved silica (column B), 5 mm wide, 3 mm deep after 24 and 96 h. There is no evident change in the F-actin arrangement in cells cultured on flat or grooved substrates for 4 days. BarZ3 mm. (B) F-actin arrangement in a spread primary chondrocyte 96 h after plating on a flat surface. F-actin is dispersed and is not as prominent in the periphery (compare with A). Membrane ruffles are evident around the edge of the cell.
It is interesting that the cells migrated most on the 750 nm deep grooves. Although the cells did not change to an elongated morphology, they migrated relatively rapidly, especially on 750 nm deep silica grooves with relatively good guidance in the direction of the groove long axis. Curtis et al. (1995) observed that on topographical cues, such as repeating grooves, cells must align before they begin to move, and that the initiation of movement appears to be related to cytoskeletal reorganization. On grooves, chondrocytes did not alter their spherical shape or F-actin arrangement, and very few microspikes or subsequent lamellapodia were observed. It is possible that, when in the spherical morphology, chondrocytes are not able to form adhesions, other than those directly under the cell (as shown by IRM); the cells are not sufficiently flexible. The movement exhibited by primary chondrocytes is similar to that which has been reported for neutrophils (Mandeville et al., 1997). The focal adhesions of the cells are confined to a very small percentage of their apparent contact area. It seems that the cells may teeter along the grooves or ridges. Clearly morphological and
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cytoskeletal organizations are not a pre-requisite for migration on repeating grooved substrata. This type of motion exhibited by primary chondrocytes and neutrophils has received very little attention to date. This type of movement is very different from that which has been reported for other cell types, such as endothelia, dorsal root ganglion neurons, and epitenon fibroblasts (Curtis et al., 1995). There are only a few studies that have investigated chondrocyte movement, mostly in response to cytokines, such as hepatocyte scatter factor, which are known to promote migration in other cell types (Takebayashi et al., 1995). Only one study assessed the in vitro movement of primary chondrocytes, and it demonstrated that approximately 30% of the total cell number was capable of migration (Chang et al., 2003). In that study, the chondrocytes were given 3 days to recover from enzymatic digestion. However, in this current investigation, we seeded the cells directly on the substrate after isolation from cartilage and 70% of the cells exhibited migration on 750 nm deep grooves after 4 h. It is clear from this investigation that certain chondrocyte populations can migrate if grown on suitable substrata in vitro, and further investigations should be aimed at identifying the phenotype of these particular cells, which could potentially be of great benefit in orthopaedic applications in the future. In addition to assessing the chondrocyte base response to materials, maintenance of the chondrocyte phenotype is also of great importance; F-actin has been associated with its loss (Von der Mark, 1986). Previous studies have shown that chondrocytes do not produce type I collagen until they have stress-fibre networks (Loeser and Lee, 1997), a cytoskeletal arrangement not seen in situ (Langelier et al., 2000). When primary chondrocytes were labelled for F-actin, the arrangement was the same on both control and grooved surfaces 96 h after plating. As it was the same in situ, it is possible that the cells require a period of time to alter the cytoskeleton to be able to respond. F-actin is known to be central in cellular reactions to topographies such as repeating grooves (Curtis et al., 1995), thus the lack of elongation and alignment of chondrocytes to grooved surfaces in comparison with other cell types (Curtis and Wilkinson, 1998) is not unexpected. In response to chemical patterns, primary chondrocytes preferentially migrated from the PET areas to the HyalS, where the cells spread and adopted a more elongated morphology. This migration is unlikely to have been due to chemotaxis, because the material was thoroughly washed free of soluble products in its preparation. Two alternative, but related, explanations seem more likely. The first is that the HyalS is so adhesive that any cell moving randomly onto it cannot leave (a fly paper effect) and the second is that the edge topography of the HyalS (1 mm in height) is not conducive to
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Fig. 6. (A) AFM image showing height profile of HyalS. (B) Coomassie blue staining of primary chondrocytes on PET and HyalS (darker stripes) after 96 h. The cells cluster on the HyalS but not on PET. BarZ100 mm. (B) The same preparation at a higher magnification. The cells are more spread on HyalS compared to their appearance on PET. Arrow indicates cells that have aligned to the boundary between substrates. BarZ50 mm. (C) Number of primary chondrocytes adhering to PET and HyalS surfaces over a 96 h period. BarsZstandard deviation, *P!0.01, Student’s paired t-test.
re-emigration to the PET surface. With regard to the morphological changes of primary chondrocytes on HyalS, this is further evidence of the inability of chemical patterns to prevent the spreading associated with loss of the chondrogenic phenotype (Loeser and Lee, 1997). Given that cells remain round on PET, it would appear to be a better substrate on which to maintain the chondrocyte phenotype. However, the alteration in CD44 receptor organization on HyalS and the increased matrix production (see Fig. 8C) suggests that the material can still be recognized by the cells even although it is modified. It is known that chondrocytes bind to hyaluronan through CD44 (Ishida et al., 1997), so the
addition of sulphate groups appears not to alter the recognition of the hyaluronan unit by the chondrocytes. When considering the response of cells to substrata used in this study, a common criticism is whether or not the cells are responding to the topography or the chemistry of the material. The HyalS bands are 1 mm in height, as observed using AFM. Cells are known to align to repeating grooved structures with depths significantly lower than this (Curtis and Wilkinson, 1998). It is relevant that there are a number of cells concentrated at the junction of the PET and the HyalS, and this is likely to be a topographic reaction. This is another example of a general finding commented on by
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Fig. 7. Cell movement of cells on PET and HyalS. (A) Average distance travelled and (B) average time on the run. *P!0.05, Student’s paired t-test. (C) Number of cells crossing the boundaries between HyalS and PET over a 96 h time frame. Data shows a directed migration from PET to HyalS. Error barsZ5% standard error. F-actin arrangement in primary chondrocytes grown on (E,F) photoimmobilised HyalS and (G,H) PET. Cells on HyalS showed a higher degree of spreading than those on PET. Bars in (E) and (F) are 10 mm, in (G) and (H) 5 mm.
Curtis and Wilkinson (1998) and Riehle et al. (2002). The effect of the topography of chemical patterns on cell behaviour has been argued in general terms (Curtis and Wilkinson, 1998), but, although there are strong arguments for topographic effects, we cannot yet exclude the possibility of chemical ones. These types of effects could
again be most valuable in locating chondrocytes to specific positions in constructs. With regard to the silica grooves, the cell response is likely to be topographical in nature because of the uniform surface chemistry that such surfaces probably have, due to blanket etching (Clark et al., 1990).
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Fig. 8. CD44 receptor arrangement on primary chondrocytes cultured on (A) PET and (B) HyalS. For explanation, see text. (C) SEM micrograph of primary chondrocytes on HyalS. Arrow indicates matrix production by the cells, which are encased in the matrix. BarZ5 mm.
It is clear that chondrocyte response to materials needs further investigation. Chemical and topographical cues are clearly important, but this investigation demonstrates that neither is more dominant than the other, both capable of altering cell behaviour. In selecting biomaterials for cartilage repair, surface structure and chemistry must be investigated to optimize the response of the cells, which in turn could aid ex vivo cartilage tissue development.
Acknowledgements The authors would like to thank Mr E. Robertson and Mrs M. Mullin for their help with the SEM images. The authors would also like to thank Sandyford Abattoir, Paisley, for kindly supplying all the lambs legs which provided the cell material used in this work. This work was supported by the EPSRC and Smith and Nephew plc, York.
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