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Overview: Biomaterials for Regenerative Medicine Vol. 64, No.4 pp. 74-82
Engineering Cell Attachments to Scaffolds in
Cartilage Tissue Engineering

Andrew J. Steward, Yongxing Liu, and Diane R. Wagner
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Integrin binding to a tissue engineering scaffold promotes focal adhesion formation and association with the actin cytoskeleton. (a) An integrin with a and b subunits attaches to a tissue engineering scaffold and to the actin cytoskeleton through linking proteins talin (tal) and vinculin (vin). (b) Focal adhesions, containing multiple integrins, form in response to integrin binding and are a concentrated site of actin filaments and proteins such as talin, vinculin, paxillin (pax), focal adhesion kinase (FAK), and Src.



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Cell attachment varies depending on the scaffold type and surface modification. (a) Cells in natural polymer scaffolds such as collagen and fibrin show strong integrin attachment. (b) An adhesive protein layer may adsorb on the surface of a porous polymer scaffold to promote cell binding. (c) Cells do not bind to hydrogels such as agarose, alginate, or PEG. (d) To modify cell attachment, hydrogels may be conjugated with peptides such as the RGD sequence.



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(a) Chondrocyte and (b) collagen fiber organization in articular cartilage.13



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Horizontal view of circumferential collagen organization in the deep layer showing chondrocyte (C), pericellular matrix (*), pericellular capsule (arrowheads), territorial matrix (TM) and interterritorial matrix (IM). Figure adapted from Poole et al.15



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Cell adhesion density and type have not been optimized for some cell types, and in fact results from the literature are in conflict on the question of whether integrin binding is beneficial for or inhibitory to cartilage tissue engineering.







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One of the challenges of tissue engineering, a promising cell-based treatment for damaged or diseased cartilage, is designing the scaffold that provides structure while the tissue regenerates. In addition to the scaffold material's biocompatibility, mechanical properties, and ease of manufacturing, scaffold interactions with the cells must also be considered. In cartilage tissue engineering, a range of scaffolds with various degrees of cell attachment have been proposed, but the attachment density and type have yet to be optimized. Several techniques have been developed to modulate cell adhesion to the scaffold. These studies suggest that the need for cell attachment in cartilage tissue engineering may vary with cell type, stage of differentiation, culture condition, and scaffold material. Further studies will elucidate the role of cell attachment in cartilage regeneration and enhance efforts to engineer cell-based cartilage therapies.


…describe the overall significance of this paper?
Tissue engineering is a promising treatment for damaged or diseased cartilage and usually requires a scaffold to provide structure while cells produce new cartilage matrix. The scaffold supplies a substrate for cell attachment to support cell survival, differentiation, and cartilage matrix deposition. In cartilage tissue engineering, cell adhesion to the scaffold via integrin binding may vary, but the results of these attachments are not fully understood. Efforts to understand the role of integrin binding and focal adhesion formation in cartilage tissue engineering could impact cell-based therapies for cartilage repair.

…describe this work to a materials science and engineering professional with no experience in your technical specialty?
Cell adhesion to a tissue engineering scaffold is an important regulator of cell behavior and tissue regeneration. Cell adhesion to a material may be modulated by treatments that improve wettability or by grafting molecules that provide sites for cell anchorage to the surface. These techniques have allowed for investigations into the consequences of cell attachment on cartilage tissue engineering, and are expected to enhance cartilage repair.

…describe this work to a layperson?
In biological applications where materials interact with cells, the properties of the material that control cell attachment are critically important to cell function. Here, we review the role of scaffolds as a substrate for cell attachment in the context of cartilage tissue engineering. The effect of cell attachment in cartilage regeneration may vary with cell type, stage of differentiation, culture condition and scaffold material.


Cartilage disease and injury are substantial health issues; osteoarthritis (OA) alone afflicts an estimated 15% of the U.S. population (nearly 40 million persons),1 and costs the U.S. economy more than $60 billion per year.2 The prevalence of OA increases rapidly with age and with an aging U.S. population it is expected that the incidence and associated costs will increase dramatically in the future.3 In spite of the magnitude of this problem, there exist few adequate treatment options, in part due to the fact that articular cartilage exhibits limited capacity for self-repair. Essentially no consistent repair occurs in cartilage defects that do not penetrate the subchondral bone.4 While full-thickness defects that do penetrate the subchondral bone show partial repair by bone marrow-derived mesenchymal stromal cells (MSCs), the repair tissue typically consists of a less functional fibrocartilage, rather than articular cartilage.5 Current therapies for cartilage damage include abrasion arthroplasty, microfracture, autologous osteochondral transplantation, autologous chondrocyte transplantation, and prosthetic joint replacement;6 however, each of these treatments has disappointing limitations such as deficient long-term repair, inadequate donor tissue/cell availability, donor site morbidity, and limited durability.

A promising treatment for damaged or diseased cartilage is tissue engineering, a technique that leverages the principles of engineering and the life sciences to develop substitutes that restore, maintain, or improve the function of tissues such as cartilage.7 A successful tissue engineering solution for cartilage repair will require a combination of several components, including the appropriate cell type; biochemical and biomechanical signals to encourage and maintain cell metabolism and direct cell phenotype; and a temporary artificial and/or macromolecular scaffold to provide structure for the regenerating tissue.8 The scaffold plays an important role in maintaining cell function and guiding tissue growth and has four basic performance requirements: three-dimensional and porous to allow nutrient and waste transport; biocompatible and bioresorbable; mechanical properties similar to the native tissue; and appropriate cell attachment that supports cell survival, differentiation and matrix production.9

The focus of this article is on the role of scaffolds as a substrate for cell attachment in the context of engineering cartilage tissue. Although many of the challenges in the design and manufacture of a scaffold for tissue engineering are the same as those for bio-inert medical implants, tissue engineering scaffolds must also be developed with cell-scaffold interactions in mind, as they directly regulate the biological function of the cells. To engineer most tissues, cell attachment sites are assumed to be necessary features of the scaffold, but in cartilage tissue engineering a wide range of scaffolds with varying levels of direct cell attachment are regularly reported. See the sidebar for a description of cartilage structure and composition.


Cartilage Structure and Composition

Articular cartilage is found on the ends of all diarthrodial joints and is critical for joint motion.10 The ability to resist compression and distribute loads allows cartilage to decrease the peak stresses in subchondral bone.11 Cartilage is an aneural, avascular, connective tissue composed mostly of extracellular matrix (ECM) molecules and water. Approximately 70–80% of cartilage consists of water, which is vital for nutrient transfer and load distribution. Chondrocytes, the cellular component of cartilage, make up approximately 1% by volume of articular cartilage.11 The main function of chondrocytes is to produce and organize the ECM, comprising collagen (60% of dry weight), proteoglycans (25–35% of dry weight), and non-collagenous proteins (15–20% of dry weight).11 Collagen is a fibrous molecule that increases the tensile strength and organization of the tissue. Proteoglycans have a protein core and numerous sulphated glycosaminoglycans (sGAG) branches. The branches are made up of disaccharide units; the type and number of these units determine the specific properties of the proteoglycan.

The major types of proteoglycans are chondroitin sulfate, keratan sulfate, dermatan sulfate, heparan sulfate, and hyaluronan. Hyaluronan is not sulphated or attached to a protein core, but it is the most abundant proteoglycan in cartilage and plays a role in binding other proteoglycans in order to form larger complexes.12 Proteoglycans have a negative charge that attracts ions, causing an osmotic imbalance. This imbalance leads to absorption of water which helps hydrate the tissue and induces a swelling pressure that increases the tissue's resistance to compression.12 Cartilage also has a unique zonal organization (Figure A). Chondrocytes and collagen fibers at the surface of the cartilage are aligned parallel to the surface to resist shear stresses in the superficial zone. The chondrocytes and collagen fibers are more randomly aligned in the middle zone in order to distribute the load throughout the tissue. Finally, the cells and fibers are aligned perpendicular to the surface to resist compressive forces in the deep zone.14 Clearly, although cartilage appears to be a simple aneural, avascular, connective tissue, there are many levels of complexity in its composition and structure.

While cartilage has vertical organizational variation, differences in matrix composition that radiate from the chondrocytes are also important. Three regions of the ECM are distinguished by different proteins and functions: the interterritorial, territorial, and pericellular matrices (Figure B). The interterritorial matrix occupies the spaces furthest from the cells and consists of keratin sulfate-rich proteoglycans and thick collagen bundles. The territorial matrix contains chondroitin sulfate-rich proteoglycans and smaller more radially organized collagen bundles. Finally, the pericellular matrix (PCM) is made up of small diameter collagen fibers that form a tightly woven capsule immediately adjacent to the cell.15 Type II collagen is the most abundant collagen in articular cartilage, but it is accompanied by types XI, IX, VI, III, XII, XIV, and X collagen as well.16 Although type VI collagen is a small percentage of the total collagen content of articular cartilage, it has been shown to be highly concentrated in the PCM. Type VI collagen is believed to provide pericellular architecture and also to improve signaling between the cell and its microenvironment.10 The PCM also consists of sulphated proteoglycans, an assortment of glycoproteins, hyaluronan, biglycan, fibronectin, and laminin.10 In native cartilage tissue, cells interact with the distinct extracellular matrix in the pericellular microenvironment. Providing an environment in which the cells reproduce this surrounding structure may be critical to the success of cartilage tissue engineering.

The primary linkage between the tissue engineering scaffold and the interior of any cell occurs through integrin molecules on the cell surface (Figure 1a). The integrin family comprises cell adhesion receptors that span the cell membrane and bind to a wide range of extracellular matrix (ECM) components and scaffold materials.17 Internal to the cell, integrins attach to the actin cytoskeleton through intermediate proteins.18,19 Integrins consist of two non-covalently associated subunits, denoted a and b. To date, 18 a and 8 b subunits have been identified, which occur in 24 different combinations.20 The particular combination of the a and b subunits determines the ligand binding specificity of the integrin. Some integrin receptors bind to a limited number of protein sequences, but most bind to many sequences which could be part of the same or different macromolecules of the ECM or scaffold.21 Each cell type expresses a specific subset of the known integrins, which controls the cells' interactions with their microenvironment.21,22 Additionally, different matrices or scaffolds, or different arrangements of the same matrix or scaffold component, can transmit distinct signals to a cell through the same integrin.21 Cellular attachment to the ECM or scaffold via integrin binding is essential for migration, proliferation, differentiation, and survival in a variety of cell types,21,23–28 and as a result integrins play a role in embryonic development; tissue growth, remodeling and maintenance; and in tissue engineering. Integrin attachments modulate cellular processes mainly through focal adhesions (Figure 1b). Focal adhesions are large clusters of integrins, often located at the periphery of the cell, that form in response to integrin binding. In addition to assuring cell-scaffold attachment, these integrin-based focal adhesion complexes also provide an intracellular concentration of more than 50 proteins29 including scaffolding proteins, GTPases and enzymes which provide critical signaling between the cell exterior and interior.30 Focal adhesions are also a targeted location for the anchorage of actin filaments, a key component of the cytoskeletal structure of most cells, which are linked to the b integrin subunits through various proteins.31,32 The interactions between focal adhesions and the actin cytoskeleton are bidirectional: actin stress fibers regulate the assembly and growth of the focal adhesions, while the focal adhesions regulate the assembly of the actin cytoskeleton.33 In some cell types, focal adhesions have been shown to be the site of a concentration of growth factor receptors such as those for basic fibroblast growth factor, platelet-derived growth factor and epidermal growth factor; focal adhesion formation subsequently leads to an enhanced response to growth factor stimulation.34,35 Focal adhesions also play a key role in sensing the stiffness and surface topography of the scaffold to which they are attached. Therefore, scaffold stiffness and surface topography contribute to the lineage commitment of undifferentiated stem cells and to the maintenance of a differentiated morphology via focal adhesions.33,36–38

A key regulator of focal adhesion assembly and signaling is focal adhesion kinase (FAK). FAK is a tyrosine kinase that localizes only to focal adhesions39 and has been implicated as a mechanosensor.40 Following attachment, integrin conformation is shifted, allowing FAK interaction with b1 integrin, resulting in autophosphorylation of FAK at Y397 and kinase activation.41 Once activated, FAK phosphorylates other scaffolding proteins and triggers a signaling cascade.42,43 FAK contains protein binding domains and also serves as a scaffolding protein within the focal adhesions, independent of kinase activity.44 Following FAK recruitment to the focal adhesion and activation, the resulting phosphorylation of target proteins activates cell proliferation, survival and migration pathways.45–48 This is a fraction of FAK's potential role in focal adhesion signaling, as FAK activation and downstream pathways are extremely complex.


Integrin signaling has been studied extensively in articular chondrocytes;it has been implicated in cartilage tissue maintenance and arthritis,49,50 and genetic mutations affecting integrin expression result in cartilage abnormalities and disease.51 Cell-matrix interactions are necessary for survival, proliferation, and differentiation to the chondrocyte phenotype.52 Chondrocytes have been shown to attach to PCM proteins through integrin binding in experiments that use specific inhibitors of integrin-mediated attachment. The addition of either synthetic peptide arginine-glycine-aspartic acid (RGD) or antibodies that block the function of b1 or b3-integrins inhibits cell attachment to ECM proteins such as fibronectin, vitronectin, and collagen II.53

In cartilage development, cell attachment to fibronectin is particularly important. Investigations with chick embryonic limb bud cells have determined that cell adhesion to fibronectin via integrin binding is necessary for condensation, which is the first step in developmental chondrogenesis.54,55 However, as the cells differentiated into chondrocytes, fibronectin expression decreased, which implies that integrin binding may be important in early differentiation, but not as necessary to maintain the differentiated state.54 Fibronectin expression in chick limb bud cells is modulated by members of the TGF-b family; when TGF-b was added to these cells prior to condensation, fibronectin expression and chondrogenic differentiation increased, but when it was added after condensation the increased fibronectin expression had no effect on differentiation. These results further confirm that cell-fibronectin interactions, and therefore integrin binding, are necessary for initial condensation and early differentiation but not for maintenance of the chondrocytic phenotype.56 These results suggest that providing the optimal cell attachment to a scaffold may be particularly critical when engineering tissues from undifferentiated cells such as mesenchymal stem cells (MSCs).


A large variety of scaffolds have been used in cartilage tissue engineering, each with strengths and weaknesses with respect to the design criteria stated earlier. One classification scheme for the different scaffolds is based on whether they promote cell attachment. For example, many of the natural hydrogels that are common in cartilage tissue engineering research, such as alginate and agarose, do not promote cell attachment. Other materials such as collagen and fibrin are known to promote integrin binding and have shown promise in cartilage repair strategies.

Non-Adherent Materials
Natural Hydrogels
Alginate and agarose are linear polysaccharides derived from seaweed and algae. Agarose as a scaffold has been shown to both promote chondrogenic induction, and maintain the induced cartilage phenotype.57 Agarose requires higher than physiologic temperatures to melt, but due to the development of low-melting point agarose, the cell viability concerns with the high temperatures have been overcome.58 Alginate gels, on the other hand, form by the addition of divalent cations such as Sr2+, Ba2+, and Ca2+, with calcium ions being the most common due to their physiologic nature and availability.59 Like many hydrogels, there is no integrin binding between cells and either alginate or agarose, and encapsulated cells retain their rounded shape. The retention of the round cell shape and lack of cell attachments enhances chondrogenesis relative to cells seeded in monolayer. Dedifferentiated cells cultured in monolayer but covered in agarose showed no tendencies towards a chondrogenic phenotype, whereas cells encapsulated in an agarose gel differentiated into chondrocytes. This evidence supports the idea that agarose does not directly affect the chondrogenic phenotype, but rather promotes chondrogenesis based on secondary effects such as cell shape and attachment.60 The main weakness of alginate and agarose hydrogels as scaffolds for cartilage tissue engineering is their poor mechanical properties; they have a compressive moduli approximately 15% of native tissue.61 A further concern is that a four-fold increase in the amount of collagen type VI was observed around chondrocytes embedded in agarose as compared to native tissue; the collagen fibers also ran perpendicular to the cell surface rather than tangentially as in native cartilage. The differences in composition and organization could have significant impacts on the tissue engineering of cartilage.62

Synthetic Hydrogels
Natural hydrogels, such as alginate and agarose, have advantages in their inherent biocompatibility, but synthetic and photocrosslinkable hydrogels offer greater control of the final macroscopic properties of the gel. Poly(ethylene glycol) and poly(vinyl alcohol) can be modified for photopolymerization, which enables spatial and temporal control of the gelation process and also provides a mechanism to polymerize the gel in situ (i.e., in a tissue defect).63 Photopolymerized PEG hydrogels have been shown to support MSC survival, differentiation, and accumulation of chondrogenic ECM in the presence of TGF-b1.64

Cell Adherent Materials
Collagen is the most abundant protein in mammals and is composed of fibrils made up of tropocollagen triple helices. These triple helices allow collagen to withstand high tensile forces.12 Cells adhere to collagen through integrin binding, which could lead to a spread and flattened morphology and to chondrocyte dedifferentiation over time. Collagen types I and II may be fabricated into fibrous sponges through lyophilization or hydrogels by neutralizing an acidic collagen solution. Collagen sponges seeded with bovine articular chondrocytes showed superior chondrogenesis relative to cells seeded in monolayer based on sGAG accumulation and chondrogenic gene expression. The porous structure of the collagen sponges allows cells to aggregate and therefore they may retain their native spherical morphology.65 Cell-seeded collagen gels contract a large amount which can be problematic for tissue engineering applications, but methods have been developed to reduce this contraction.66,67 Human synovium-derived MSCs seeded in a collagen hydrogel have been shown to increase both chondrogenic gene expression and protein synthesis in the presence of BMP-2 and TGF-b3.68

Fibrin is a naturally occurring protein involved in wound healing. Fibrinogen, when combined with thrombin, polymerizes into a fibrin hydrogel. Cells attach to fibrin, and MSCs seeded in fibrin gels in the presence of chondrogenic growth factors differentiate to the chondrogenic lineage.69 Fibrin gels may also enhance collagen synthesis relative to type I collagen gels in some cell types.70 One of the main disadvantages of fibrin is that it contracts when seeded with stem cells, leading to poor control over the size of the implanted scaffold. However, this problem is diminished when fibrin is seeded with chondrocytes.71 Fibrin also tends to degrade quickly when implanted, but addition of aprotinin or e-amino-n-caproic acid slows fibrin's rate of degradation.72,73


Modulating cell-scaffold interactions through alterations of the scaffold material allows for investigations into the influence of integrin attachment on cartilage matrix production. The choice of scaffold material(s) directly determines cell adhesion, which has significant effects on other cell functions, such as proliferation and differentiation. Scaffolds composed of natural polymers such as fibrin74,75 and collagen76–79 are often used to achieve direct cell anchorage (Figure 2a). These macromolecules provide good cell adhesion, making them an obvious scaffold material when cell attachment is desired. Biodegradable synthetic polymers such as poly(alpha-hydroxy esters), poly(lactic acid, PLA), poly(glycolic acid, PGA), and their copolymer poly(lactic-co-glycolic acid) (PLGA) are attractive candidates for cartilage tissue engineering scaffolds that promote cell attachment in an indirect manner. These synthetic polymers are usually hydrophobic, allowing a protein layer to quickly adsorb on the scaffold surface upon contact with body fluid in vivo or culture medium in vitro to establish cell adhesion sites (Figure 2b).80 However, their high hydrophobicity may impair cell adhesion due to denaturing of the adsorbed proteins and inaccessibility of the adhesion motifs to cells.81 In addition, the hydrophobicity directly inhibits cell seeding in porous scaffolds, because the penetration of the aqueous cell suspension is retarded, resulting in uneven cell distribution.82,83 Surface modifications for improved wettability can not only effectively promote cell seeding efficiency but also sustained cell adhesion. Typical surface modification methods include ethanol treatment,84 alkaline hydrolysis,85 and plasma oxidization.86 In plasma oxidization of polystyrene, for example, the outermost surface of the polymer is disrupted and hydrophilic functional groups are produced which can interact strongly with proteins such as fibronectin87,88 and collagen89 to improve cell anchorage. Ultimately, a balance between hydrophobicity and hydrophilicity is required for cell attachment through protein adsorption; Groth et al.81 identified the optimal water contact angle as 55–75Ί based on a fibroblast study.

Grafting ECM proteins such as collagen to a hydrophobic surface is another attractive technique to improve cell seeding efficiency and promote cell adhesion as well. A collagen coating simultaneously produces two beneficial effects: it increases hydrophilicity to facilitate cell infiltration and homogenous cell distribution, and it directly provides cell anchorage sites on the scaffold surface.82,89,90 For instance, PLGA sponges were coated with a thin collagen layer by introducing a collagen solution under vacuum followed by centrifugation and freeze drying.82 Collagen may also be incorporated into PLGA scaffolds in the form of a microsponge that fills the scaffold pores.83 The resulting hybrid scaffold significantly promoted cartilage matrix production and mechanical properties of the engineered tissue.83 Hydrogels are advantageous for tissue engineering scaffolds because they encapsulate cells homogenously and provide intrinsic porosity, but many hydrogels such as alginate, hyaluronan, agarose and chitosan lack cell adhesion motifs to support direct cell anchorage (Figure 2c). Additionally, unlike rigid polymers, the highly hydrated state of the hydrogel molecules inhibits adsorption of adhesive proteins from the environment.91 The encapsulated cells generally maintain spherical shapes in the gels and are unable to spread or migrate. The lack of anchorage to the gel matrix may adversely affect the behaviors of stem cells such as MSCs which proliferate only to a limited extent in the non-cell adhesive gels.91,92 Some effort has been made to blend non-adhesive hydrogels with cell adhesive macromolecules such as collagens and their denatured form gelatin to achieve cell attachment in hydrogels.93,94

A recent study showed that the mere manipulation of surface topography may change the nonadherent hydrogel poly(ethylene glycol) to be cell adhesive; significant cell adhesion was induced with microscale surface patterns, while there was no cell adhesion to smooth substrates of the same material.95 One explanation for these results is that the microtopography may have enhanced the adsorption of adhesive proteins to the surface of the scaffolds, although the mechanism for this is not yet known. Clearly, material-protein interactions and the consequent effects on cell adhesion are complex, and more study is needed to fully understand the optimal design parameters for tissue engineering scaffolds.

Synthetic peptides, particularly those containing the arginine-glycineaspartic acid (RGD) sequence, have been incorporated into a variety of biomaterials to provide natural cellmatrix interactions that are effective at enhancing cell adhesion and biological activity23 (Figure 2d). The RGD motif was identified as the minimum essential cell adhesion peptide sequence in fibronectin.96 Since then, the RGD peptide sequence has been found in ECM components such as vitronectin, fibrinogen, von Willebrand factor, thrombospondin, laminin, entactin, tenascin, osteopontin, bone sialoprotein, and under some conditions, collagens.97 The RGD sequence is the most effective and most commonly used synthetic peptide sequence for studies of cell adhesion due to its widespread distribution throughout the organism, its biological impact on cell behavior and survival, and the fact that it is a ligand for multiple integrins.23 Cells seeded on a non-adhesive hydrogel surface that has been conjugated with RGD peptides will attach and spread through integrin binding.23,98–100 Other peptide sequences, such as the GFOGER sequence from collagen, also promote beneficial cell-matrix interactions.79,101 Although synthetic peptides lack the specificity and function of native ECM proteins, they have important advantages when conjugated to nonadhesive hydrogels, as they allow direct control over ligand type and density. Because the synthetic peptides are typically designed to represent only a single motif, in some cases they can be chosen such that they selectively activate particular adhesion receptors, allowing for a focused investigation of signaling pathways.23 The RGD peptide sequence, however, is bound by nearly half of the 24 integrins, including α5β1, α8β1, αIIbβ3, αvβ1, αvβ3, αvβ5, αvβ6, αvβ8, and others that show weaker affinity.97


Although cartilage is one of the few tissues for which cell attachment to the scaffold is not absolutely necessary for tissue engineering, cell-scaffold interactions can both enhance and diminish the effectiveness of matrix production (Table I). The role of cell attachment in cartilage tissue engineering is complicated, and depends on the scaffold, culture conditions, and stage of differentiation of the cells. For example, cell attachment via integrin binding may be necessary for MSCs to undergo chondrogenesis via condensation.54,55 Additionally, scaffolds that promote cell attachment, such as collagen gels, enhance chondrocyte proliferation, which could be beneficial in promoting tissue growth.92 On the other hand, not all effects of cell attachment increase cartilage matrix deposition. For example, long-term integrin binding can lead to chondrocyte dedifferentiation and formation of calcified cartilage or fibrocartilage,103 while mature chondrocytes implanted in non-adherent alginate or agarose hydrogels maintain their phenotype for long periods without undergoing dedifferentiation.60,103 Furthermore, MSCs and adipose-derived stromal cells (ASCs), which are generally anchorage-dependent, undergo significant chondrogenic differentiation in alginate or agarose hydrogels both in vitro and in vivo.104–106 Cells seeded in agarose were more metabolically active than those seeded in collagen gels, which implies that a lack of cellular attachment enhances biosynthesis.107 However, cells seeded in fibrin hydrogels increased collagen deposition relative to those in collagen gels as well.108 Clearly, distinguishing the effects of cell attachment in different materials can be difficult due to many inherent material differences. Therefore, the ability to modify a material to either enhance or inhibit cell attachment can better elucidate the role of cell attachment as opposed to comparing two distinct materials.

Conjugation of the integrin-attachment motif RGD to a scaffold that otherwise does not promote cell attachment provides a direct method of altering integrin binding and has been employed in several studies investigating chondrogenesis of MSCs for cartilage tissue engineering. MSCs seeded in RGD modified alginate gels expressed significantly lower levels of the chondrogenic genes aggrecan, collagen II, and Sox 9, and accumulated less sGAG than in the control samples of alginate conjugated to non-binding RGE. These results suggest that RGD and integrin attachment inhibited chondrogenesis of MSCs.109 In a follow-up study, MSC attachment to RGD-modified agarose gels inhibited sGAG production in a density-dependent manner through the RhoA/Rock pathway and actin cytoskeleton interactions.110 In a separate investigation, however, it was shown that cell attachment in early differentiation stages was beneficial to MSC chondrogenesis. Using PEG hydrogels conjugated to enzymatically cleavable RGD peptides that are released by the MSCs in later stages of differentiation, this study demonstrated enhanced chondrogenesis when the MSCs bind initially but later release the RGD attachments relative to persistent cell attachments to uncleavable RGD.111 Another research group demonstrated beneficial effects of cell attachment with lyophilized RGDalginate sponge scaffolds for cartilage tissue engineering. Human MSCs seeded on the RGD sponges attained a flattened morphology and formed focal adhesions; the RGD group also showed higher expression of the chondrogenic genes Sox 9 and collagen II than the control. They concluded that the cells seeded in unmodified alginate clustered together, but the cells in the RGD-alginate attached and stayed separated, which could have enhanced nutrient flow to the cells and led to a more chondrogenic phenotype.112 Cumulatively, these results using RGD to modulate cell attachment are somewhat confusing, as some studies report an inhibition in chondrogenesis with integrin binding, and others report an enhancement. However, the different studies used different cell types, scaffold architectures, and peptide concentrations, which may explain the difference in the results. The scaffold architecture in particular could affect the number of cell attachment sites, nutrient availability, cell shape, and cell aggregation which could have a significant effect on cellular behavior and chondrogenic potential. Additionally, these RGD studies demonstrate that cell-scaffold attachment is important in cartilage tissue engineering, but has yet to be optimized.

Alterations to a scaffold's physiochemical structure can also allow insight into the role of cell attachment on chondrogenesis. PEGT/PBT copolymer scaffolds were constructed and their wettability modified, while the overall porosity and composition of the scaffolds remained equal. The more hydrophilic scaffolds adsorb less protein (specifically fibronectin which strongly promotes integrin attachments), and the cells maintained a more spheroidal morphology, produced more proteoglycan, had a higher collagen type II/I ratio, and had reduced staining for actin relative to the scaffold with a less hydrophilic surface.113 Similarly, hybrid scaffolds composed of polycaprolactone and biopolymers such as hyaluronan, chitosan, fibronectin, and collagen were fabricated in order to explore the effect of cell attachment with specific ECM molecules.114,115 These scaffolds demonstrated that chondrocyte interaction with hyaluronan and chitosan promotes neocartilage formation and may be promising components of a cartilage tissue engineering scaffold.114 Therefore, by modifying the protein adsorption or bioactivity of a scaffold, insights into the role of cell attachment in chondrogenesis may be better elucidated.

Although type I and II collagen both support cell attachment, differences in the chemical structures may lead to cell binding through different integrins or to different spatial arrangements of integrin attachments when cells are seeded in fibrous collagen sponges and hydrogels. Although type II collagen is more abundant in native cartilage than type I, no significant differences were found between type I and type II collagen sponges seeded with chondrocytes after 20 days of in vitro culture.116 GAG-collagen copolymer sponges were formed using either collagen type I or II, and the GAG-collagen type II sponge maintained a chondrogenic phenotype better than the GAG-collagen type I copolymer.117 Both collagen type I and II sponges were implanted into full thickness defects in rabbit knees. Collagen type I sponges were found to recruit more subchondral progenitor cells than collagen II sponges, but promoted calcified cartilage and fibrocartilage formation. Collagen type II sponges, however, supported a more chondrogenic phenotype.102 Therefore a matrix composed of both cell types with a deep layer of type I collagen and a superficial layer of type II collagen was proposed.

Inhibition of the signaling molecules that are activated by cell attachment, such as focal adhesion kinase (FAK), can also add insight into the effects of cell attachment on chondrogenesis. While the addition of exogenous type II collagen to chondrocytes increased FAK signaling, knockdown of FAK by siRNA inhibited expression of type II collagen and cell proliferation, indicating that FAK is required for communication with collagen II and is involved in the regulation of type II collagen expression and cell proliferation.118 However, suppression of FAK also enhances the early stages of chondrogenesis; FAK inhibition in micromass cultures increased expression of chondrogenic genes, suggesting that cell-cell interactions rather than cell-ECM interactions drive condensation and early chondrogenesis.119


Although current cartilage tissue engineering strategies use a wide range of scaffold materials, the above data demonstrate that cell adhesion is involved in the regulation of cartilage matrix production in engineered tissues. Providing the correct microenvironmental attachment signals to the cell via the scaffold may be necessary to replicate the complex structure and organization of cartilage tissue. The optimal scaffold type and surface treatment have yet to be determined, but may vary with cell type, stage of differentiation, culture condition and scaffold material. Cell type may have an especially large influence on the effect of cell attachment, as chondrocytes form a PCM and quickly bind to the proteins found there; the question of cell attachment may be less crucial to these cells as long as dedifferentiation is not induced. Formation of the PCM and its interaction with the chondrocytes may also explain the relative success in engineering cartilage with so many different scaffold materials.

Although chondrocytes are an obvious choice as a source of cells for cartilage tissue engineering, their limited number, slow proliferation, as well as donor site morbidity have led to intense interest in alternate cell types that can differentiate to the chondrogenic lineage, such as MSCs and ASCs. With these undifferentiated cells, cell-scaffold attachments may be especially important as they may be involved in driving the cell to the chondrogenic differentiation pathway. However, cell adhesion density and type have not been optimized for these cells, and in fact results from the literature are in conflict on the question of whether integrin binding is beneficial for or inhibitory to cartilage tissue engineering (Table I). We expect that further, more detailed studies will elucidate the role of cell attachment in chondrogenesis and will enhance efforts to engineer cell-based cartilage therapies.


The authors acknowledge support from the University of Notre Dame Adult Stem Cell Initiative, the U.S. Army Medical Research and Materiel Command W81XWH-07-0662 and W81XWH-09-1-0741, and the Naughton Graduate Fellowship.


1. R.C. Lawrence et al., Arthritis Rheum,, 41 (5) (1998), pp. 778–799.
2. J.A. Buckwalter, C. Saltzman, and T. Brown, Clin. Orthop. Relat. Res. (427 Suppl) (2004), pp. S6–S15.
3. Arthritis Foundation, Association of State and Territorial Health Officers (1999).
4. H.J. Mankin, N. Engl. J. Med., 291 (24) (1974), pp. 1285–1292.
5. F. Shapiro, S. Koide, and M.J. Glimcher, J. Bone Joint Surg. Am., 75 (4) (1993), pp. 532–553.
6. S.N. Redman, S.F. Oldfield. and C.W. Archer, Eur. Cell. Mater., 9 (2005), pp. 23–32.
7. R. Langer and J.P. Vacanti, Science, 260 (5110) (1993), pp. 920–926.
8. D.L. Butler, S.A. Goldstein, and F. Guilak, J. Biomech. Eng., 122 (2000), pp. 570–575.
9. D.W. Hutmacher, Biomaterials, 21 (24) (2000), pp. 2529–2543.
10. C.A. Poole, J. Anatomy, 191 (1) (1997), pp. 1–13.
11. J.A. Buckwalter and H.J. Mankin, J. Bone and Joint Surg., 79 (4) (1997), pp. 600–611.
12. E.M. Culav, C.H. Clark, and M.J. Merrilees, Phys. Ther., 79 (3) (1999), pp. 308–319.
13. J.A. Buckwalter, V.C. Mow, and A. Ratcliffe, J. Am. Acad. Orthop. Surg., 2 (4) (1994), pp. 192–201.
14. T.J. Klein, J. Malda, R.L. Sah, and D.W. Hutmacher, Tissue Eng. Part B: Rev., 15 (2) (2009), pp. 143–157. 15. C. Poole, M.H. Flint, and B.W. Beaumont, J. Anat., 138 (Pt 1) (1984), pp. 113–138.
16. D.R. Eyre, Arthritis Research, 4 (1) (2002), pp. 30–35.
17. R.O. Hynes, Cell, 69 (1) (1992), pp. 11–25. 18. C. Brakebusch and R. Fδssler, EMBO J., 22 (10) (2003), pp. 2324–2333.
19. I. Delon and N.H. Brown, Curr. Opin. Cell Biol., 19 (1) (2007), pp. 43–50.
20. A. van der Flier and A. Sonnenberg, Cell Tissue Res., 305 (3) (2001), pp. 285–298.
21. D.G. Stupack and D.A. Cheresh, J. Cell. Sci., 115 (19) (2002), pp. 3729–3738.
22. B.H. Luo, C.V. Carman, and T.A. Springer, Annu. Rev. Immunol., 25 (2007), pp. 619–647.
23. U. Hersel, C. Dahmen, and H. Kessler, Biomaterials, 24 (24) (2003), pp. 4385–4415.
24. J.A. Eble and J. Haier, Current Cancer Drug Targets, 6 (2) (2006), pp. 89–105.
25. D. Docheva, C. Popov, W. Mutschler, and M. Schieker, J. Cell. Mol. Med., 11 (1) (2007), pp. 21–38.
26. K. Raymond, M.A. Deugnier, M.M. Faraldo, and M.A. Glukhova, Curr. Opin. Cell Biol., 21 (5) (2009), pp. 623–629.
27. E.A. Clark and J.S. Brugge, Science, 268 (5208) (1995), pp. 233–239.
28. A. Howe, A.E. Aplin, S.K. Alahari, and R.L. Juliano, Curr. Opin. Cell Biol., 10 (2) (1998), pp. 220–231.
29. E. Zamir and B. Geiger, J. Cell. Sci., 114 (20) (2001), pp. 3583–3590.
30. R.O. Hynes, Cell, 110 (6) (2002), pp. 673–687. 31. R. Zaidel-Bar, S. Itzkovitz, A. Ma'ayan, R. Iyengar, and B. Geiger, Nat. Cell Biol., 9 (8) (2007), pp. 858– 867.
32. B. Geiger, A. Bershadsky, R. Pankov, and K.M. Yamada, Nat. Rev. Mol. Cell Biol., 2 (11) (2001), pp. 793–805.
33. B. Geiger, J.P. Spatz, and A.D. Bershadsky, Nat. Rev. Mol. Cell Bio., 10 (1) (2009), pp. 21–33.
34. G.E. Plopper, H.P. McNamee, L.E. Dike, K. Bojanowski, and D.E. Ingber, Mol. Biol. Cell, 6 (10) (1995), pp. 1349–1365.
35. S. Miyamoto, H. Teramoto, J.S. Gutkind, and K.M. Yamada, J. Cell Biol., 135 (6) (1996), pp. 1633–1642.
36. A.J. Engler, S. Sen, H.L. Sweeney, and D.E. Discher, Cell, 126 (4) (2006), pp. 677–689.
37. D.E. Discher, P. Janmey, and Y. Wang, Science, 310 (5751) (2005), pp. 1139–1143.
38. V. Vogel and M. Sheetz, Nat. Rev. Mol. Cell Bio., 7 (4) (2006), pp. 265–275.
39. M. Schaller, C.A. Borgman, B.S. Cobb, R.R. Vines, A.B. Reynolds, and J.T. Parsons, Proc. Nat. Acad. of Sci., 89 (11) (1992), pp. 5192–5196.
40. H.B. Wang, M. Dembo, S.K. Hanks, and Y. Wang, Proc. Nat. Acad. of Sci., 98 (20) (2001), pp. 11295–11300.
41. J. Zhao and J.L. Guan, Cancer Metastasis Rev., 28 (1) (2009), pp. 35–49.
42. J.T. Parsons, J. Cell. Sci., 116 (8) (2003), pp. 1409–1416.
43. S. Roy, P.J. Ruest, and S.K. Hanks, J. Cell. Biochem., 84 (2) (2002), pp. 377–388.
44. M.C. Beckerle, M.D. Schaller, J.D. Hildebrand, and J.T. Parsons, Mol. Biol. Cell, 10 (10) (1999), pp. 3489–3505.
45. A.P. Gilmore, T.W. Owens, F.M. Foster, and J. Lindsay, Curr. Opin. Cell Biol., 21 (5) (2009), pp. 654–661.
46. A. Gilmore and L.H. Romer, Mol. Biol. Cell, 7 (8) (1996), pp. 1209–1224.
47. D. Ilic et al., Nature (London), 377 (6549) (1995), pp. 539–544.
48. L.A. Cary, J.F. Chang, and J.L. Guan, J. Cell. Sci., 109 ( Pt 7) (1996), pp. 1787–1794.
49. R.F. Loeser, Biorheology, 37 (1) (2000), pp. 109– 116.
50. S. Millward-Sadler and D.M. Salter, Ann. Biomed. Eng., 32 (3) (2004), pp. 435–446.
51. A. Woods, G. Wang, and F. Beier, J. Biol. Chem., 280 (12) (2005), pp. 11626–11634.
52. M.S. Hirsch, L.E. Lunsford, V. Trinkaus-Randall, and K.K.H. Svoboda, Developmental Dynamics, 210 (3) (1997), pp. 249–263.
53. R.F. Loeser, Arthritis & Rheumatism, 36 (8) (1993), pp. 1103–1110.
54. S. Tavella et al., J. Cell. Sci., 110 ( Pt 18) (1997), pp. 2261–2270.
55. A.L. Gehris, E. Stringa, J. Spina, M.E. Desmond, R.S. Tuan, and V.D. Bennett, Dev. Biol., 190 (2) (1997), pp. 191–205.
56. E.F. Roark and K. Greer, Am .J. Anat., 200 (2) (1994), pp. 103–116.
57. A.Y. Thompson, K.A. Piez, and S.M. Seyedin, Exp. Cell Res., 157 (2) (1985), pp. 483–494.
58. J. Raghunath, J. Rollo, K.M. Sales, P.E. Butler, and A.M. Seifalian, Biotechnol. Appl. Biochem., 46 (Pt 2) (2007), pp. 73–84.
59. O. Smidsrψd, Faraday Discuss. Chem. Soc., 57 (1974), pp. 263–274.
60. P.D. Benya and J. D. Shaffer, Cell, 30 (1) (1982), pp. 215–224.
61. J.L. Drury and D.J. Mooney, Biomaterials, 24 (24) (2003), pp. 4337–4351.
62. M. Dimicco, J.D. Kisiday, H. Gong, and A.J. Grodzinsky, Osteoarthritis and Cartilage/OARS, Osteoarthritis Research Society, 15 (10) (2007), pp. 1207–1216.
63. S.J. Bryant and K.S. Anseth, J. Biomed. Mater. Res., 59 (1) (2002), pp. 63–72.
64. C.G. Williams, T.K. Kim, A. Taboas, A. Malik, P. Manson, and J. Elisseeff, Tissue Eng., 9 (4) (2003), pp. 679–688.
65. K.E. Yates, F. Allemann, and J. Glowacki, Cell Tissue Banking, 6 (1) (2005), pp. 45–54.
66. A.M. Freyria et al., Tissue Eng. Part A, 15 (6) (2008), pp. 1233–1245.
67. E. Gentleman, E.A. Nauman, K.C. Dee, and G.A. Livesay, Tissue Eng., 10 (3-4) (2004), pp. 421–427.
68. A. Yokoyama, I. Sekiya, K. Miyazaki, S. Ichinose, Y. Hata, and T. Muneta, Cell Tissue Res., 322 (2) (2005), pp. 289–298.
69. G.I. Im, Current Applied Physics, 5 (5) (2005), pp. 438–443.
70. E. Grassl, T.R. Oegema, and R.T. Tranquillo, J. Biomed. Mater. Res. Part A, 60 (4) (2002), pp. 607– 612.
71. G.M. Peretti, M.A. Randolph, M.T. Villa, M.S. Buragas, and M.J. Yaremchuk, Tissue Eng., 6 (5) (2000), pp. 567–576.
72. Q. Ye et al., European J. Cardio-Thoracic Surg., 17 (5) (2000), pp. 587–591.
73. A. Mol et al., Biomaterials, 26 (16) (2005), pp. 3113–3121.
74. G.M. Peretti et al., Ann. Plast. Surg., 46 (5) (2001), pp. 533–537.
75. D.A. Hendrickson et al., J. Orthop. Res., 12 (4) (1994), pp. 485–497.
76. S.R. Frenkel, B. Toolan, D. Menche, M.I. Pitman, and J.M. Pachence, J. Bone Joint Surg. Br., 79 (5) (1997), pp. 831–836.
77. M.C. Ronziere, S. Roche, J. Gouttenoire, O. Demarteau, D. Herbage, and A.M. Freyria, Biomaterials, 24 (5) (2003), pp. 851–861.
78. A.E. Sams, R.R. Minor, J.A. Wootton, H. Mohammed, and A.J. Nixon, Osteoarthritis Cartilage, 3 (1) (1995), pp. 61–70.
79. S.Q. Liu, Q. Tian, J.L. Hedrick, J.H. Po Hui, P.L. Ee, and Y.Y. Yang, Biomaterials, 31 (28) (2010), pp. 7298–7307.
80. R.A. Latour, Encyc. Biomater. and Biomed. Eng., (2005), pp. 1–15.
81. T. Groth, G. Altankov, A. Kostadinova, N. Krasteva, W. Albrecht, and D. Paul, J. Biomed. Mater. Res., 44 (3) (1999), pp. 341–351.
82. G. Chen et al., J. Biomed. Mater. Res. B—Appl. Biomater., 90 (2) (2009), pp. 864–872.
83. W. Dai, N. Kawazoe, X. Lin, J. Dong, and G. Chen, Biomaterials, 31 (8) (2010), pp. 2141–2152.
84. A.G. Mikos, M.D. Lyman, L.E. Freed, and R. Langer, Biomaterials, 15 (1) (1994), pp. 55–58.
85. J. Gao, L. Niklason, and R. Langer, J. Biomed. Mater. Res., 42 (3) (1998), pp. 417–424.
86. J. Yang, Y. Wan, J. Yang, J. Bei, and S. Wang, J. Biomed. Mater. Res. A., 67 (4) (2003), pp. 1139–1147.
87. T.G. van Kooten, H.T. Spijker, and H.J. Busscher, Biomaterials, 25 (10) (2004), pp. 1735–1747.
88. A.P. Kowalczyk and P.J. McKeown-Longo, J. Cell. Physiol., 152 (1) (1992), pp. 126–134.
89. J. Yang, J. Bei, and S. Wang, Biomaterials, 23 (12) (2002), pp. 2607–2614.
90. Z. Ma, C. Gao, Y. Gong, and J. Shen, Biomaterials, 26 (11) (2005), pp. 1253–1259.
91. J.T. Oliveira and R.L. Reis, J. Tissue Eng. Regen. Med., (2010), in press.
92. H. Yamaoka et al., J. Biomed. Mater. Res. A, 78 (1) (2006), pp. 1–11.
93. D. Bosnakovski, M. Mizuno, G. Kim, S. Takagi, M. Okumura, and T. Fujinaga, Biotechnol. Bioeng., 93 (6) (2006), pp. 1152–1163.
94. C. Yang, H. Frei, F.M. Rossi, and H.M. Burt, J. Tissue Eng. Regen. Med., 3 (8) (2009), pp. 601–614.
95. V.A. Schulte, M. Diez, M. Moller, and M.C. Lensen, Biomacromolecules, 10 (10) (2009), pp. 2795–2801.
96. M.D. Pierschbacher and E. Ruoslahti, Nature, 309 (5963) (1984), pp. 30–33.
97. E. Ruoslahti, Ann. Rev. Cell and Devel. Bio., 12 (1) (1996), pp. 697–715.
98. B. Jeschke et al., Biomaterials, 23 (16) (2002), pp. 3455–3463.
99. J.A. Rowley and D.J. Mooney, J. Biomed. Mater. Res., 60 (2) (2002), pp. 217–223.
100. J.A. Rowley, G. Madlambayan, and D.J. Mooney, Biomaterials, 20 (1) (1999), pp. 45–53.
101. C.D. Reyes and A.J. Garcνa, J. Biomed. Mater. Res. Part A, 65 (4) (2003), pp. 511–523.
102. P. Buma et al., Biomaterials, 24 (19) (2003), pp. 3255–3263.
103. J.L. van Susante et al., Acta Orthop. Scand., 66 (6) (1995), pp. 549–556.
104. C.T. Buckley et al., J. Biomech., 43 (5) (2010), pp. 920–926.
105. H.L. Ma, S.C. Hung, S.Y. Lin, Y.L. Chen, and W.H. Lo, J. Biomed. Mater. Res. A, 64 (2) (2003), pp. 273–281.
106. K.H. Bouhadir, K.Y. Lee, E. Alsberg, K.L. Damm, K.W. Anderson, and D.J. Mooney, Biotechnol. Prog., 17 (5) (2001), pp. 945–950.
107. A. Guaccio, C. Borselli, O. Oliviero, and P.A. Netti, Biomaterials, 29 (10) (2008), pp. 1484–1493.
108. R.A. Clark, L.D. Nielsen, M.P. Welch, and J.M. McPherson, J. Cell. Sci., 108 ( Pt 3) (Pt 3) (1995), pp. 1251–1261.
109. J.T. Connelly, A.J. Garcνa, and M.E. Levenston, Biomaterials, 28 (6) (2007), pp. 1071–1083.
110. J.T. Connelly, A.J. Garcia, and M.E. Levenston, J. Cell. Physiol., 217 (1) (2008), pp. 145–154.
111. C.N. Salinas and K.S. Anseth, Biomaterials, 29 (15) (2008), pp. 2370–2377.
112. T. Re'em, O. Tsur-Gang, and S. Cohen, Biomaterials, 31 (26) (2010), pp. 6746–6755.
113. S. Miot et al., Biomaterials, 26 (15) (2005), pp. 2479–2489.
114. J. Schagemann et al., J. Biomed. Mater. Res. Part A, 93 (2) (2010), pp. 454–463.
115. J.C. Schagemann et al., Biomaterials, 31 (10) (2010), pp. 2798–2805.
116. T. Ohno, K. Tanisaka, Y. Hiraoka, T. Ushida, T. Tamaki, and T. Tateishi, Mater. Sci. and Eng.: C, 24 (3) (2004), pp. 407–411.
117. S. Nehrer et al., J. Biomed. Mater. Res. Part B: Appl. Biomater., 38 (2) (1997), pp. 95–104.
118. Y.H. Kim and J.W. Lee, J. Cell. Physiol., 218 (3) (2009), pp. 623–630.
119. D. Pala et al., J. Biol. Chem., 283 (14) (2008), pp. 9239–9247.

Andrew J. Steward, graduate student, Yongxing Liu, visiting research assistant professor, and Diane R. Wagner, assistant professor, are with the Department of Mechanical Engineering and Bioengineering Graduate Program, University of Notre Dame, Notre Dame, IN 46556 USA. Steward is also with the Department of Mechanical and Manufacturing Engineering and Trinity Centre for Bioengineering, Trinity College Dublin, Dublin, Ireland. Dr. Wagner can be reached at the Department of Aerospace and Mechanical Engineering, 145 Multidisciplinary Research Building, Notre Dame, IN 46556 USA; (574) 631- 5735; e-mail: