Tendons are dynamic structures; their extracellular matrices are continuously being synthesised and broken down over the course of an individual’s lifetime. The macromolecules, namely collagen, proteoglycans, hyaluronan and the non-collagenous proteins form the extracellular matrix of tendons. In normal tendon exists a fine balance between the synthesis and degradation of these macromolecules resulting in a strong healthy tendon. It is evident that damage to tendons, such as in overuse tendinopathy results in changes to the levels and types of macromolecules present in tendon with decreased levels of collagen and increased levels of proteoglycans, hyaluronan and non-collagenous proteins, causing a weakened tendon that is prone to rupture.
These degenerative features have thus far been partially characterised. By identifying the levels and various types of macromolecules present in normal tendons and tendons exhibiting overuse tendinopathy an understanding of the basis of the condition can be determined and possible ways of preventing or ameliorating tendon degeneration can be considered. The terms overuse tendinopathy and pathological tendon will be used interchangeably throughout this study.
This literature review will attempt to define and characterise the structural and functional properties of tendon and will discuss the current literature regarding the levels, types, synthesis and catabolism of macromolecules present in the extracellular matrix of tendons and also attempt to define and characterise the pathological aspects of overuse tendinopathies. Chapter Two of this thesis will dictate the materials and methodology used in these studies. Chapters Three, Four and Five will present the results of this thesis. Finally, chapter Six will include the discussion and discuss any limitations and future considerations.
1.1 Synovial Joint
Joints are articulations found between adjacent parts of bone that allow controlled frictionless movement (for review see; Mankin & Radin, 1997). In the human body there are three different types of joints and these are grouped according to the type of movement they make. They include the freely movable joints (synovial joints; i.e., most joints of the extremities such as the knee joint), slightly movable (cartilaginous joints; i.e., the vertebrae and ribs) and those that are immovable (fibrous joints; i.e., the skull). The majority of the joints found in the human body are synovial joints (for review see; Mankin & Radin, 1997).
There are six different types of synovial joints including the ball-and-socket joints, hinge joints, saddle joint, pivot joint, gliding joints and condyloid joints. A synovial joint contains a joint cavity that is enclosed by a fibrous capsule linking the adjoining bones. This joint capsule is lined by a synovial membrane that secretes a lubricating and nutritious fluid called synovial fluid that is rich in albumin and hyaluronan. The surface of each bone is typically covered with articular hyaline cartilage or in some circumstances fibrocartilage. In addition, the joint capsule is supported by accessory structures such as tendons and ligaments, which provide stability to the synovial joint (Sledge et al., 2001).
1.1.1 Articular Cartilage
Articular cartilage covers the adjoining ends of bones in joints and has a white colour (for review see; Mankin & Radin, 1997). It is a tissue that is devoid of blood and nerves and provides a wear resistant surface with low frictional properties for the joint and attains its nutrients via diffusion from the synovium into the synovial fluid (for review see; Mankin & Radin, 1997). Furthermore, articular cartilage is resilient and flexible. This allows articular cartilage to withstand large compressive and tensile forces as well as allowing it to distribute load on subchondral bone during joint loading (Kempson, 1980) even though it is only a few millimetres thick (Hardingham, 1998).
Its biomechanical properties are dependent on the structural composition of the extracellular matrix, which is comprised of water (70-80%), collagens (predominantly Type II collagen), proteoglycans (predominantly aggrecan) and non-collagenous proteins (Kuettner et al., 1991; Poole, 1997). The predominant cell type present in articular cartilage is called the chondrocyte. These cells are responsible for the maintenance, synthesis and degradation of all the extracellular matrix components (Kuettner et al., 1991; Buckwalter & Mankin, 1998).
Mature articular cartilage can be divided up into four zones including the superficial (tangential) zone, the middle (transitional) zone, the deep (radial) zone and the zone of calcified cartilage (Huber et al., 2000). The organisation and composition as well as mechanical properties of the extracellular matrix varies within these zones. The deeper zones have high proteoglycan levels and low cellularity whereas the more superficial zones contain low proteoglycan levels and increased cellularity (Aydelotte et al., 1988; Buckwalter & Mankin, 1998).
1.1.2 Joint Capsule and Ligament
The joint capsule is a fibrous connective tissue that is attached to the skeletal parts of a joint beyond their articular surfaces. The principal function of the joint capsule is to seal the joint space and to supply stability by limiting movement (for review see; Mankin & Radin, 1997). Most joint capsules are strengthened by ligaments. Ligaments act together with the joint capsule and the peri-articular muscles to provide stability to the joint preventing excessive movements. They permit free movements when lax, but can stop unwanted movements when tight by virtue of their high tensile strength.
Occasionally joint capsules are strengthened by tendons, such as the extensor tendon in the finger joint. The joint capsule and ligaments proceed to hold the bones together and to guide and limit joint movements. Ligaments attach one bone with another bone and have a limited vascular and neural supply which enable them to repair relatively well after damage (Bray et al., 1990). The knee joint is a good example of different types of ligaments. The medial collateral ligament fuses with the joint capsule, and the cruciate ligaments and the lateral collateral ligament, which are both completely independent of the joint capsule.
1.1.3 Synovial Membrane
The synovial membrane (synovium) lines the non-articular surfaces of a joint such as the joint capsule and ligaments, and is responsible for secreting and absorbing synovial fluid, which contains hyaluronan (Mason et al., 1999). Synovial fluid lubricates the joint and provides at least partly for the nutrition of articular cartilage, invertebral discs and menisci. The synovial extracellular matrix acts as a scaffolding to support synoviocytes and plays an important role in cell migration and differentiation. It is mostly composed of collagen particularly Type III collagen, with smaller amounts of proteoglycans such as decorin and biglycan (Mason et al., 1999), non-collagenous proteins such as fibronectin, elastin and lamina, hyaluronic acid as well as lipids, serum proteins and electrolytes (Hirohata & Kobayashi, 1964).
The synovial membrane has only been detected in vertebrate animals (Henderson & Edwards, 1987). Furthermore, synovial tissue is not arranged into discrete layers, but rather represents a continuum from surface to deep zones. The extracellular matrix of the synovial membrane varies in composition from its surface to its deep zones (Hirohata & Kobayashi, 1964).
Tendons are dense fibrous connective tissues found between muscles and bones (for review see; Benjamin & Ralphs, 1997). The primary role of tendon is to absorb and transmit force generated by muscle to the bone to provide movement at a joint. In addition tendons operate as a buffer by absorbing forces to limit muscle damage. Each individual muscle has two tendons, one that is proximal and the other distal. The attachment of the proximal tendon of a muscle to bone is called a muscle origin and that of the distal tendon an insertion.
A normal tendon has a bright white colour and a fibroelastic texture and enables resistance to mechanical forces. Tendons come in many shapes and this is most likely due to their function, they can be round or oval in cross section or they can come in the form of flattened sheets, fan shaped, ribbon shaped or cylindrical in shape (for review see; Benjamin & Ralphs, 1997). In a muscle like the quadriceps which creates strong forces the tendons are short and broad, while those that are involved in more delicate movements like the finger flexors, long and thin tendons are present (Kannus, 2000).
Tendons are arranged in a hierarchical fashion (see Figure 1.1). A group of collagen fibres form a primary fibre bundle or subfascicle; this is the basic unit of tendon. A group of subfascicles form secondary bundles or fascicles, which form tertiary bundles constituting the tendon as a whole. The primary, secondary and tertiary bundles are encased in a thin connective tissue reticulum called the endotenon (Elliott, 1965; Kastelic et al., 1978; Rowe, 1985). The endotenon carries blood vessels, nerves and lymphatics to deeper areas of the tendon (Elliott, 1965; Hess et al., 1989). The whole tendon is surrounded by an epitenon, which is a dense fibrillar network of collagen (Jozsa et al., 1991).
The epitenon is contiguous with the endotenon and like the endotenon is rich in blood vessels, nerves and lymphatics (Hess et al., 1989). Many tendons are surrounded by a connective tissue called the paratenon. Paratenon allows free movement of the tendon against the surrounding tissues (Schatzker & Branemark, 1969; Hess et al., 1989). The myotendinous junction is the site of union with a muscle, and the osteotendinous junction is the site of union with a bone (Kannus, 2000).
In tendon, blood vessels represent between 1-2% of the entire extracellular matrix (Lang, 1960; Lang, 1963). Some blood vessels may originate from the perimysium at the musculotendinous junction and blood vessels from the osteotendinous junction (Schatzker & Branemark, 1969; Carr & Norris, 1989; Clark et al., 2000). At rest, rabbit tendons have been shown to have blood flow of around one-third that of muscle, and it is known that blood flow in tendon increases with exercise and during healing in animals (Backman et al., 1991). The oxygen consumption of tendons is 7.5 times lower than that of skeletal muscles (Vailas et al., 1978).
1.1.5 Tendon Extracellular Matrix
The major cell type present in tendon is the fibroblast (also known as tenocytes; Ross et al., 1989; Schweitzer et al., 2001; Salingcarnboriboon et al., 2003), which are embedded within an extracellular matrix (see Figure 1.2). These cells are sparsely distributed, comprising only 5% of the dry weight of adult tendon (Ross et al., 1989; Schweitzer et al., 2001; Salingcarnboriboon et al., 2003). These cells lie in longitudinal rows and have many cell extensions that extend into the extracellular matrix (McNeilly et al., 1996). Fibroblasts are responsible for the synthesis and degradation of all the macromolecular components that make up the extracellular matrix of tendon, including the most abundant macromolecule present in tendon, collagen, as well as proteoglycans, hyaluronan and non-collagenous proteins (Vogel & Heinegard, 1985; Curwin, 1997; O’Brien, 1997).
The extracellular matrix is made up of parallel bundles of collagen aligned longitudinally (60-85% of tendon dry weight) associated with elastin fibres which constitutes approximately 1-2% of the dry weight of tendon (Tipton et al., 1975; Hess et al., 1989; Jozsa et al., 1989; Curwin, 1997; Kirkendall & Garrett, 1997; O’Brien, 1997). Tendon consists of 55-70% water, most of which is associated with proteoglycans in the extracellular matrix (Elliott, 1965; Vogel, 1977; Merrilees & Flint, 1980; Riley et al., 1994b; Vogel & Meyers, 1999). The proteoglycan content of tendons is approximately 1% of dry weight of tendons (O’Brien, 1997).Water and proteoglycans have important lubricating and spacing roles in tendons that allow collagen fibres to glide over one another (Amiel et al., 1984).
The structure, composition and the organisation of the tendon matrix is crucial for the physical properties that tendons posses (Riley, 2004). The collagen component gives tendon its great tensile strength (Scott, 2003) whereas it is the proteoglycan component of the tendon matrix that enables tendons to withstand compressive load (Schonherr et al., 1995), while elastin fibres increase tendon extensibility (Scott, 2003).
1.1.6 Tendon cells
The cell population of tendon has so far been poorly characterised (for review see; Riley, 2000), the majority of tendon cells have the appearance of fibroblasts (also known as tenocytes) and constitute about 90-95% of the cells present in tendon (Ross et al., 1989; Schweitzer et al., 2001; Salingcarnboriboon et al., 2003). The remaining 5% to 10% of cells present in tendon are chondrocyte-like cells (fibrochondrocytes), which are mostly present in the fibrocartilaginous regions of tendon where tendon attaches to bone. Also present in tendon are some mast cells, capillary endothelial cells, smooth muscle cells and nerve cells (Hess et al., 1989; Jozsa & Kannus, 1997).
Fibrocartilage cells are large and have an oval shape and they are often packed with intermediate filaments (Merrilees & Flint, 1980; Ralphs et al., 1991). Tendon cells are linked to one another via gap junctions (McNeilly et al., 1996; Ralphs et al., 1998), allowing cell-to-cell interactions (McNeilly et al., 1996). Fibroblasts have a branched cytoplasm surrounding an elliptical, speckled nucleus. The rough endoplasmic reticulum and the Golgi apparatus are well developed with few mitochondria in the cytoplasm (Ippolito et al., 1980; Moore & De Beaux, 1987). Like other connective tissue cells, fibroblasts are derived from mesenchyme.
It is believed that in tendon there are a small number of mesenchymal stem cells that have the ability to differentiate into chondrogenic, osteogenic and adipogenic cells if the conditions allow (Salingcarnboriboon et al., 2003). Tendons have been shown to respond to mechanical load by modifying their extracellular matrix (Banes et al., 1988; Ehlers & Vogel, 1998; Buchanan & Marsh, 2002; Lavagnino & Arnoczky, 2005). Tendon cells receive their vascular supply from the surrounding paratenon.
Tendons were once considered almost static and unable to participate in repair. However, the activity of tendon cells has been shown to be active throughout an individual’s life as they express various matrix components (Chard et al., 1987; Ireland et al., 2001; Riley et al., 2002). Regional differences in cell morphology and activity exists in tendons, synovial-like cells that are found in the endotenon and epitenon surround the main fibre bundles (Banes et al., 1988). A greater proliferative capacity and a different matrix synthetic activity is characteristic of these synovial-like cells compared to the fibroblasts within the fibres, and are the first cells to respond following acute tendon injury (Gelberman et al., 1986; Banes et al., 1988; Garner et al., 1989; Gelberman et al., 1991; Khan et al., 1996b).
Tendon Extracellular Matrix Macromolecules
The following section will discuss the major extracellular matrix proteins and their roles in tendon. This will include the major constituent of tendon, collagen, the small and large proteoglycans and the non-collagenous proteins as well as hyaluronan. This section will also discuss the synthesis of collagens, proteoglycans and hyaluronan.
Collagen is the most copious protein present in the extracellular matrix of connective tissues and accounts for approximately 90% of the total protein of tendons, or 65% to 75% of the dry weight of tendons (von der Mark, 1981; O’Brien, 1992). There are currently 28 different collagen types (numbered I-XXVIII) present in vertebrates with at least 42 different alpha chains (Veit et al., 2006) with this number continuing to mount (Brown & Timpl, 1995; Aumailley & Gayraud, 1998). Collagen molecules can be defined as an extracellular protein that contains at least one triple helical domain (van der Rest & Bruckner, 1993). Collagen provides the tendon with its structural integrity as well as assisting in various physiological functions.
Collagen consists of three polypeptide alpha chains, which combine to form a homotrimer (three identical alpha chains) or a heterotrimer (two or three different alpha chains). Covalent bonds known as collagen cross-links develop between individual collagen molecules in a collagen fibre (Eyre et al., 1984; Bailey et al., 1998; Bailey, 2001; Brady & Robins, 2001). The collagen arrangement gives tendon its great tensile strength. Cross-links are formed from a pathway of different chemical reactions that result in divalent cross-links that join two polypeptide chains, to multivalent, i.e. tri- or even tetravalent, cross-links (Bailey & Lapiere, 1973; Eyre et al., 1984). These cross-links come about from enzymatic modification of lysine or hydroxylysine residues by the copper-dependent enzyme lysine oxidase (Robins, 1988).
Collagens are divided into two subgroups, the fibrillar and non-fibrillar collagens. Non-fibrillar collagens can be further divided into seven subfamilies including microfibril collagens, fibril-associated collagens with interrupted helices (FACIT) collagens, network collagens, MULTIPLEXIN collagens (proteins with multiple triple helix domains and interruptions), basement membrane-associated collagens, transmembrane-associated collagens and epithelium-associated collagens (von der Mark, 1999). The non-fibrillar collagens present in tendon include Types IV, VI, IX, X, XII and XIV (von der Mark, 1999).
The fibrillar collagens present in tendon include, Types I, II, III, V and XI (Kielty et al., 1993; Kadler et al., 1996; Fukuta et al., 1998; von der Mark, 1999). The fibrillar collagens contain a continuous triple helix domain, 300 nm in length, capable of undergoing the staggered, lateral associations required to form fibrils (Mayne, 1997). The resulting fibrils provide the structural support for tissues. All the fibril-forming collagens have a similar structure and size, being composed of a large, continuous central triple-helical domain (COL1) of approximately 1000 amino-acid residues
Occurs in most tissues, tendon, bone, skin etc
Main component of tendon, skin, bone, dentin, cartilage, ligament etc
Hyaline cartilage, invertebral disc
Restricted to fibrocartilage; forms less-organised meshwork
Vessels, kidney, liver, skin, tendon
Normally restricted to endotenon; forms smaller less organised fibrils
Basement membranes, tendon
Basement membrane of tendon blood vessels
Core of Type I collagen fibril - forms template for fibrillogenesis
Vessels, skin, intervertebral disc
Cell associated - found in seams between fibrils
Forms anchoring fibrils in the skin
Descements membrane in the cornea
Forms a lattice
Hyaline cartilage, vitreous humour, tendon
Cell and matrix interactions with Type II collagen fibril surface
Growth plate, tendon
Restricted to insertion fibrocartilage
Core of Type II collagen fibril - forms template for fibrillogenesis
Embryonic tendon and skin, periodontal ligament
Mediates cell/matrix interactions with Type I collagen fibril surface
Adhesion of cells to basement membranes
Foetal skin, tendon
Mediates cell/matrix interactions with Type I collagen fibril surface
Stabilizes skeletal muscle cells and microvessels
Skin, cornea, lung
Connects epithelial cells to the matrix
Endothelial cells, liver, eye
Needed for normal development of the eye
Forms radially distributed aggregates
Corneal epithelium, skin, cartilage and tendon
Binds to collagen fibrils
Matrix assembly of vascular networks in blood vessel formation
Interacts with components of microfibrils
Metastatic tumour cells, heart retina
Cell adhesion, Binds to heparin
Expressed in tissues containing Type I collagen Developing bone and cornea
Regulating Type I collagen fibrillogenesis
May play a role in adherens junctions between neurons
Testis and ovary of adult tissues
Development of the reproductive tissues
Cartilage, ear, eye and lung
Basement membranes around Schwann cells in the peripheral nervous system.
flanked by a variable amino-terminal domain of about 50-520 amino acid residues and a highly conserved non-triple-helical carboxyl-terminal domain of about 250 amino acid residues (for reviews see; Kielty et al., 1993; Fichard et al., 1995; Pihlajaniemi & Rehn, 1995; Prockop & Kivirikko, 1995; Bateman et al., 1996). The amino- and carboxyl-terminal extensions are commonly referred to as amino- and carboxyl- propeptides, respectively. The C-propeptide is called the NC1 domain, whereas the amino-propeptide is divided into sub-domains. The first is a short sequence (NC2) that links the major triple helix to the minor one (COL2) and a globular amino-terminal end (NC3) that shows structural and splicing variations.
Collagen Types II, IX, X and XI (Fukuta et al., 1998) are present at specific sites within the fibrocartilage region of tendon, found at the bone insertion and where the tendon is subjected to shear forces or compression (Fukuta et al., 1998; Waggett et al., 1998). Collagen Types II, IX, X and XI were once thought to occur only in cartilage (Visconti et al., 1996; Fukuta et al., 1998; Riley, 2000). It has now been shown that these collagens are found in the fibrocartilaginous regions of tendon, which wraps under bone. Their presumed function is to help resist compression and shear forces at these sites (Visconti et al., 1996; Fukuta et al., 1998; Waggett et al., 1998).
Collagen also plays an important role in attaching tendons to bone. Where the tendon attaches to bone, tendons commonly widen and give way to fibrocartilage, a transformation where the aligned fibres originating from the tendon are separated by other collagen fibres arranged in a three dimensional network surrounding rounded cells (Liu et al., 1995). This arrangement helps to transmit tensile forces onto a broad area and reduces the chance of failure under excessive loading. The following review will focus on the collagens that are known to exist in tendon; this includes collagen Types I-VI, IX-XII and XIV.
126.96.36.199 Type I Collagen
Type I collagen is the predominant and most studied collagen type present in the extracellular matrix of tendon, ligament and bone representing approximately 95% of the total collagen content or 60% of the tendon dry weight (Evans & Barbenel, 1975; von der Mark, 1981; Riley et al., 1994b; Rufai et al., 1995). It is synthesized by a number of cell types such as fibroblasts, osteocytes and odontoblasts. Type I collagen consists of two α1(I) chains and a shorter α2(I) chain (Kielty et al., 1993), these two chains are products of separate genes and are not a posttranslational modification of a single molecule (for review see; Kivirikko & Prockop, 1995).
The two α1(I) and one α2(I) chains of a monomer of Type I collagen are primarily comprised of approximately 338 repeating tripeptide sequences of Gly-X-Y in which X is frequently proline and Y is frequently hydroxyproline (OHPr). The ends of the α1(I) and one α2(I) chains consist of short telopeptides of between 11-26 amino acids per chain.
In longitudinal sections, the monomers are arranged in fibrils in a head-to-head-to-tail orientation. Each Type I collagen molecule consists of a long central helical region with a short non-helical domain on both the amino- and carboxyl-terminal ends. In tendon, the Type I collagen-containing fibril, organized into fibres (fibril bundles), is the major element responsible for structure stabilization and the mechanical attributes of this tissue. The fibril contains collagen molecules assembled into a quarter-staggered array, and this striated fibril has a 67 nm periodicity (for review see; Kadler et al., 1996; Orgel et al., 2006).
Each alpha chain consists of a repeating triplet of glycine and two other amino acids marked as (Gly-X-Y)n. It is the glycine residues located in every third position that makes it possible for the three alpha chains to coil around the other. It has a molecular weight of 290 kDa. When viewing collagen fibrils under the light microscope they have a crimped appearance, during tendon loading the crimp stretches and the fibrils become aligned, and after loading the crimp will reappear, this is an important elastic component that tendon possesses (O’Brien, 1992).
The Type I collagen α chains contain approximately 290 residues of OHPr per molecule. Proline and OHPr constitute 20% to 25% of all amino acid residues of Type I collagen. The parallel arranged bundles formed by the Type I collagen fibrils gives tissues a high tensile strength with limited elasticity, and therefore is suitable for force transmission. The Type I collagen molecule has the ability to form microfibrils (filaments) as well as larger units of the fibrils or fibres (for review see; Kivirikko & Prockop, 1995). The diameter of the collagen fibril is usually between 20 nm and 150 nm but can range up to 300 nm, this depends on the stage of development (Dyer & Enna, 1976; Jozsa et al., 1984; Fleischmajer et al., 1988).
188.8.131.52 Type II Collagen
The homotrimeric Type II collagen molecule was first discovered in cartilage by Miller and Matukas in 1969 who extracted collagen from cartilage in an experiment that involved pepsin digestion. Type II collagen, although most commonly found in articular and hyaline cartilage is also expressed in tendon particularly around the fibrocartilaginous region and consists of three identical α1(II) chains (Eyre et al., 1992) which forms a meshwork structure that gives Type II collagen the ability to entrap the negatively charged proteoglycan molecules, thereby resisting the swelling pressure of proteoglycans. Each Type II collagen chain has a molecular weight of approximately 95 kDa.
The entire collagen Type II molecule is shaped like a thin rod and is 300 nm long and 1.5 nm wide and has a total combined molecular weight of 295 kDa. This molecule is essential in connective tissues that are subjected to compression such as tendon and articular cartilage. Type II collagen molecules consists of a long central helical region flanked at its amino- and carboxyl-terminus by short non-helical regions termed amino and carboxyl telopeptides (Eyre et al., 1992). As with all fibrillar collagens, Type II collagen molecules are arranged in a quarter-staggered array to form collagen fibrils. Lateral associations of these collagen fibrils forms collagen fibres (Mayne, 1997). In tendon, collagen Types IX and XI as well as the proteoglycans decorin, fibromodulin and lumican inhibit collagen Type II fibril formation reducing fibril thickness (Vogel et al., 1984; Hedbom & Heinegard, 1989; Hedbom & Heinegard, 1993).
184.108.40.206 Type III Collagen
Type III collagen is the second most abundant collagen present in tendon, representing up to 10% of the total collagen content in various tendons (Hanson & Bentley, 1983; Riley et al., 1994b). Type III collagen is a thin collagen fibre consisting of three α1(III) chains with a molecular weight of 290 kDa. In tendon most Type III collagen is found in the endotenon and epitenon (Duance et al., 1977), and is also found in between Type I collagen fibril bundles in aging tendons and at the insertion (Kumagai et al., 1994). It can also be found in skin, blood vessels, ligament and internal organs such as the gastro-intestinal tract but is not found in bone (Epstein & Munderloh, 1978; McCullagh et al., 1980; Amiel et al., 1984). It strengthens the walls of hollow structures like the intestines and uterus.
The fibrils of Type III collagen have a generally thinner diameter compared with Type I collagen fibrils (Lapiere et al., 1977; for review see; Kadler et al., 1996), however the triple helical domain is longer in length being composed of 340 amino acid repeats compared to 338 amino acid repeats in Type I collagen. In the early repair of the injured tendon, Type III collagen fibrils are quickly synthesized to restore strength and elasticity (Williams et al., 1984; Dahlgren et al., 2005). However, the fibrils do not have the same tensile strength quality as Type I collagen and so lack the functional properties needed in a tendon experiencing maximal load. The repair processes continues with Type III fibrils slowly being replaced by Type I collagen fibrils in an attempt to normalize the properties of the tendon (Duance et al., 1977; Williams et al., 1984; Dahlgren et al., 2005).
Type III collagen contains high levels of OHPr and glycine. It has been reported that these high levels of glycine may cause localised helix instability resulting in increased susceptibility to proteolytic cleavage and rapid turnover of the extracellular matrices containing this collagen (Linsenmayer, 1991). The frequency of Type III collagen is considered to be an indicator of tissue age, and is common in the early stages of healing and scar tissue formation where it provides mechanical strength to the matrix (Burgeson & Nimni, 1992).
220.127.116.11 Type IV Collagen
The non-fibrillar collagen, Type IV (Bailey et al., 1979), is a basement membrane-associated collagen (Light & Champion, 1984) composed of triple helical isoforms consisting of six genetically distinct chains [α1(IV) to α6(IV)]. Each chain is characterised by a long collagenous domain of approximately 1400 amino acid residues of Gly-X-Y repeats, that are interrupted at several sites by a short non-collagenous sequence and approximately 15 amino acid residue non-collagenous amino-terminus, and an approximately 230 amino acid residue non-collagenous domain at the carboxyl-terminus (Mayne, 1997). Type IV collagen has been reported to represent approximately 2% of the total collagen content of tendon (Ahtikoski et al., 2003). Unlike the fibrillar collagens discussed so far this collagen does not form fibrillar aggregates but are directly incorporated into the basement membrane without any prior excision of the pro-peptide extensions.
Type IV collagen is found uniquely in the basement membrane of tendon blood vessels (von der Mark, 1981) where it forms a key structural component; it is the only collagen that forms into a meshwork structure where four of these molecules are covalently joined together with their 7-S domains (Risteli et al., 1980; von der Mark, 1981). This lattice links to other basement membrane components such as laminin and entactin, which prevents the movement of cells and the migration of high molecular weight macromolecules. The carboxyl-terminus domain is not removed in post-translational processing and the fibers link head-to-head rather than in parallel. Also, Type IV collagen lacks the regular glycine in every third residue necessary for the tight collagen helix. These two features cause the collagen to form in a sheet.
18.104.22.168 Type V Collagen
Type V collagen is a member of the fibril forming subclass of collagens and has a molecular weight of 300 kDa with a triple helical domain that is 300 nm long. Type V collagen is intercalated into the core of the Type I collagen fibril, where it forms a template for fibrillogenesis and modulates fibril growth (Keene et al., 1987; Linsenmayer, 1991; Waggett et al., 1998). This function may be mediated by retention of the non-collagenous amino-terminal propeptide after Type V collagen molecules are incorporated into fibrils (Birk et al., 1990; Linsenmayer et al., 1993; Niyibizi & Eyre, 1993; Moradi-Ameli et al., 1994). The non-collagenous domain projects outward through the gap between adjacent Type I collagen molecules leaving major portions present on the fibril surface (Birk et al., 1988; Marchant et al., 1996) where they may limit lateral growth of the fibril by steric hindrance and charge interactions (Fichard et al., 1995).
This collagen contains three distinct alpha chains [α1(V), α2(V) and α3(V)] and is a quantitatively minor fibrillar collagen present in tissues where Type I collagen is expressed such as tendon, bone, placenta and skin. This collagen type can form very small diameter fibrils, where Type V collagen triple helical epitopes are exposed, adjacent to cells or basement membranes (Modesti et al., 1984; Gordon et al., 1994). There are several Type V collagen isoforms that differ in chain composition (Fichard et al., 1995). Type V collagen alpha chains also form heterotypic molecules with Type XI collagen alpha chains (Niyibizi & Eyre, 1989; Kleman et al., 1992; Mayne et al., 1993; Mayne et al., 1996) and is found in small amounts inside the fascicles and in the endotenon.
22.214.171.124 Type VI Collagen
Type VI collagen (Furthmayr et al., 1983) is a microfibril collagen consisting of α1(VI), α2(VI) and α3(VI) chains and has a molecular weight of greater than 420 kDa (Jander et al., 1983; Chu et al., 1987; Mayne & Burgeson, 1987) and forms multi molecular filamentous beaded structures after secretion from the cell (Timpl & Chu, 1994). It contains a short triple helical domain of 335–336 amino acids with repeating Gly-Xaa-Yaa sequences flanked by two large globular domains located at the carboxyl- and amino-termini (Chu et al., 1987). These are composed primarily of approximately 200 amino acid subdomains with homology to von Willebrand factor type A domains (Chu et al., 1990).
The amino-terminal globular domain is larger than the carboxyl-terminal domain and consists almost exclusively of the α3(VI) chain, which has nearly twice the mass of the α1(VI) and α2(VI) chains (Kielty et al., 1993; Timpl & Chu, 1994). This collagen forms a sheet-like structure and is usually found co-distributed with Type I collagen fibres in normal tendons (von der Mark, 1981; Waggett et al., 1998). This collagen is present in small amounts in tendon and is known to play a role in binding cells to matrix molecules, including fibrillar collagens, hyaluronan and decorin (Pfaff et al., 1993) as its chains appear to be recognised by cellular receptors (Bonaldo et al., 1990).
Collagen Type VI shows a high affinity, specific interaction with biglycan and decorin by binding to its core protein (Wiberg et al., 2001). The interaction is with the amino-terminal globular domain of the collagen Type VI (Specks et al., 1992; Burg et al., 1996). It may have a role in the development of the matrix supramolecular structure as well as in tissue homeostasis by mediating interactions of cells with the extracellular matrix. More specifically, interactions of collagen Type IV with collagen Type XIV, collagen Type IV and the fibrillar collagens Type I and II, decorin, microfibril-associated glycoprotein-1 and hyaluronan as well as the α1β1 and α2β1 integrins and the cell surface proteoglycan NG2 have been demonstrated (Bonaldo et al., 1990; McDevitt et al., 1991; Bidanset et al., 1992; Brown et al., 1993; Kuo et al., 1997; Pfaff et al., 1993; Burg et al., 1996; Finnis & Gibson, 1997) .
Collagen Type VI also interacts via its triple helical domain with perlecan and fibronectin (Tillet et al., 1994). It is also involved in cell migration and differentiation, and may play a role in bridging cells with the extracellular matrix. Collagen VI assembles intracellularly into antiparallel, overlapping dimers that then align and form tetramers (Engvall et al., 1986).
126.96.36.199 Type IX Collagen
Type IX collagen forms thin-beaded filaments that may interact with fibrils and cells and has a molecular weight of 250 kDa. Type IX collagen is a heterotrimer comprised of three chains [α1(IX), α2(IX) and α3(IX)] and contains three collagenous domains (COL-1, -2 and -3) separated by four non-collagenous domains (NC-1, -2, -3 and -4; Shimokomaki et al., 1980; Ninomiya & Olsen, 1984; Ninomiya et al., 1985; van der Rest et al., 1985; Har-El et al., 1992), these are numbered from the carboxyl-terminus. Present on the α2(IX) chain (NC-3 domain) is a single chondroitin sulphate chain, therefore this collagen can be considered a proteoglycan (Bruckner et al., 1985; Huber et al., 1986; Konomi et al., 1986; McCormick et al., 1987; van der Rest & Mayne, 1987; Olsen, 1989; Ayad et al., 1991; Yada et al., 1992).
The amino-terminal NC-4 domain of the α1(IX) chain is a large globular domain of 266 amino acid residues (Vasios et al., 1988). Type IX collagen is a minor constituent of articular cartilage and tendon (Fukuta et al., 1998). This collagen is a FACIT collagen as its resides on the exterior surface of collagen fibrils and cannot self-associate (Vaughan et al., 1988) and is found covalently cross-linked to collagen Type II (Wu et al., 1992). FACIT's are relatively short collagens, have interruptions in the triple helical domain and can be found at the surfaces of collagen fibrils. COL-1 and COL-2 appear to be involved in interactions with fibrils and COL-3 serves as the arm sticking out of the fibril. Two forms of Type IX collagen exist, with major differences in the NC4 domain (Svoboda et al., 1988).
188.8.131.52 Type X Collagen
Type X collagen was first described in 1982 as a collagenous molecule of 59 kDa per chain in cultures of chondrocytes from developing bone (Schmid & Conrad, 1982). This molecule is a homotrimer [α(X)]3. It forms a meshwork structure and is found in the fibrocartilaginous region of tendon. The amino acid sequence, gene structure and molecular organization of Type X collagen is extremely similar to that of Type VIII collagen.
The triple helical domain is 460 residues long and is also marked by the presence of eight imperfections at locations similar to that of Type VIII collagen. The carboxyl-terminal domain is 162 residues long and the amino-terminal domain is only 52 residues long. The Type X collagen molecule consists of a putative signal-peptide sequence (18 amino acids), an amino-terminal non-collagenous domain (38 amino acids), a triple helix (463 amino acids) and a carboxyl-terminal non-collagenous domain (161 amino acids; Thomas et al., 1991).
184.108.40.206 Type XI Collagen
Type XI collagen is a heterotrimer (Morris & Bachinger, 1987) composed of (α)1, (α)2 and (α)3(XI) chains and is 300 kDa in size with a helical domain length of 320 nm. Type XI collagen is found in small amounts in tendon and is associated with the more abundant Type II collagen fibrils (Burgeson et al., 1982; Mendler et al., 1989; Fichard et al., 1995). Type XI collagen is found in cartilage and vitreous humor of the eye. The cDNA and gene sequences have been identified for all the chains, as well as the chromosomal localizations of the genes: the gene for the α1(XI) chain (COL11A1) is located on chromosome 1p21, the gene for the a2(XI) chain (COL11A2) in chromosome 6p21.2, and the α3(XI) chain on chromosome 12q13-q14 (Prockop & Kivirikko, 1995).
220.127.116.11 Type XII Collagen
Type XII collagen forms a large molecule of 660 kDa. Type XII collagen is a FACIT collagen that is associated with the surface of Type I collagen fibrils (Keene et al., 1991), in particular at the insertion and it also interacts with decorin and fibromodulin (Font et al., 1996). It is highly expressed in fibrous connective tissues such as ligament and tendon and is also found in the dermis, cornea, blood vessel walls, skin, meninges and developing membranous bones containing Type I collagen (Sugrue et al., 1989; Oh et al., 1993), but also in connective tissue of cartilage containing Type II collagen (Lunstrum et al., 1991; Watt et al., 1992).
Type XII collagen is a homotrimer (Dublet et al., 1989) with two triple-helical domains (COL1-2) and three non-triple-helical domains (NC1-3; Gordon et al., 1989; Yamagata et al., 1991). The globular amino-terminal domain (NC3) contains several distinct subdomains homologous to domains found in other molecules, i.e. fibronectin type III repeats, von Willebrand factor A domains and the amino-terminal globular domain found in α1(IX) collagen (the NC4 domain of the long form of the α1(IX) chain). These non-collagenous subdomains make up most of the total length of the Type XII collagen molecule, while the triple-helical domains contribute only a small amount of the entire molecule.
18.104.22.168Type XIV Collagen
Type XIV collagen is a homotrimeric molecule composed of two collagenous domains (COL1-COL2) and three non-collagenous domains (NC1-NC3) and is closely related to Type XII collagen (Dublet & van der Rest, 1991; Gordon et al., 1991; Gerecke et al., 1993; Walchli et al., 1993). This multi-domain collagen can interact with more than one extracellular component simultaneously allowing integration of the developing matrices. Type XIV collagen has a developmental expression pattern in tendon consistent with a role in linear fibril growth.
Type XIV collagen can interact with GAG chains of other proteoglycans, namely with dermatan sulphate of the small proteoglycan decorin (Font et al., 1993), with the heparin sulphate chains of the basement membrane proteoglycan perlecan and with heparin (Brown et al., 1993). This collagen forms a large molecule of 660 kDa.
Type XIV collagen is also able to bind to the small proteoglycan fibromodulin (Font et al., 1996) and to the triple-helical domain of Type VI collagen (Brown et al., 1993). Type XIV has been shown to be commonly expressed in the dermis, tendon, perichondrium, perimysium, the stroma of the lungs and liver and in blood vessels, and also in virtually every tissue containing collagen Type I (Castagnola et al., 1992; Walchli et al., 1994), but also, as with Type XII collagen, in cartilage tissue containing Type II collagen (Lunstrum et al., 1991; Watt et al., 1992). The GAG chain of decorin has been shown to be crucial for binding to this collagen (Ehnis et al., 1996).
Shatton and Schubert first discovered the proteoglycans in 1954. Proteoglycans are defined as containing a protein core with one or more glycosaminoglycan/s (GAG) covalently attached (for review see; Iozzo, 1998; Watanabe et al., 1998; Culav et al., 1999). The function of the different proteoglycans is dictated by the structure of their protein core and GAG side chains. The GAG side chains of the proteoglycans are linear polysaccharides that can attract water and thus contribute to tissue hydration (Iozzo, 1998). They play an essential role in the biochemical, biomechanical and structural properties of the tendon matrix (Roughley & Lee, 1994; Iozzo, 1998; Watanabe et al., 1998; Wight & Merrilees, 2004). They have important roles as cell surface receptors and coreceptors, mediating cell-cell signalling, recognition and binding (Rauch et al., 2001; Selva & Perrimon, 2001; Nakato & Kimata, 2002; Stallcup, 2002; Couchman, 2003; Yoneda & Couchman, 2003).
Proteoglycans make up less than 1% of the dry weight of most tensile regions of tendons. In tendons, they are most commonly found in the extracellular matrix where they are associated with collagen fibrils and matrix proteins, stabilising the extracellular matrix of the tendon (Iozzo & Murdoch, 1996). The proteoglycans found in tendon include decorin, biglycan, fibromodulin, lumican, aggrecan and versican.
Proteoglycans have been classified into two subfamilies according to their molecular weight: 1) the small leucine-rich repeat proteoglycans (SLRPs) and 2) the large aggregating proteoglycans (for review see; Iozzo & Murdoch, 1996).
22.214.171.124 Small Leucine-Rich Repeat Proteoglycans (SLRPs)
All SLRPs contain a small core protein (36-42 kDa) with an amino-terminal domain where GAGs attach, a carboxyl-terminal domain and a central region which contains leucine-rich repeats (LRRs; Iozzo & Murdoch, 1996; Iozzo, 1997; Iozzo, 1999). These small proteoglycans form a horseshoe shaped core protein that is thought to be involved in protein-protein interactions (Scott, 1996). Indeed, these proteoglycans have a role in the mechanical behaviour of tendon through their binding with macromolecules of the tendon matrix. They have been shown to bind to collagen fibrils, growth factors and inhibit collagen fibrillogenesis. The majority of the SLRPs that are present in tendon are substituted with between 1 and 4 chains of GAG which may be chondroitin sulphate, keratan sulphate or dermatan sulphate (for review see; Yoon & Halper, 2005). Occasionally the SLRPs exist in non-glycosylated forms. All connective tissues contain at least one member of the SLRPs (Iozzo, 1997; Iozzo, 1999).
The SLRP gene family contains at least 13 members with more being discovered as time passes (Ameye & Young, 2002). SLRPs interact with various parts of the extracellular matrix having a role in the modulation of matrix formation and integrity, regulation of cell growth, migration and adhesion (Iozzo & Murdoch, 1996; Iozzo, 1998). The core proteins of the SLRPs are characterised by leucine-rich repeats (LRRs; Henry et al., 2001), these structures are involved in the binding of the SLRPs to collagen (Vogel et al., 1984; Hedbom & Heinegard, 1989; Rada et al., 1993; Henry et al., 2001). The central domain of the small proteoglycans can represent up to approximately 80% of the protein moiety and contains approximately 10 fold repeats (with the exception of class III SLRPs) of between 20 to 29 amino acid residue LRR with asparagine and leucine residues (Iozzo, 1999).
Proteoglycans such as fibromodulin also contain 10 LRRs and in addition to this, with the exception of proline arginine-rich end leucine-rich repeat protein (PRELP) have keratan sulphate chains attached to the LRRs (Iozzo, 1999). Proteoglycans in class III such as epiphycan are much smaller and have just six LLRs and contain sulphated tyrosine residues in the amino-terminal extension. The remaining two proteoglycans do not belong to any of the classes mentioned, they have 11 LLRs and based on their amino acid sequence they are more related to one another than any other SLRPs (Iozzo, 1999).
The most abundant proteoglycan present in tendon is decorin (see Figure 1.3; Vogel & Heinegard, 1985; Krusius & Ruoslahti, 1986; Samiric et al., 2004b) representing approximately 80% of the total proteoglycan content (Samiric et al., 2004b). Other members of the SLRPs present in tendon include biglycan (see Figure 1.3; Fisher et al., 1989; Samiric, 2003), lumican (Blochberger et al., 1992; Funderburgh et al., 1993) and fibromodulin (Oldberg et al., 1989). This review will focus on the small proteoglycans that are known to be present in tendon including decorin, biglycan, fibromodulin and lumican.
Decorin is composed of a 36-40 kDa core protein (Lorenzo et al., 2001) that was deduced by cDNA analysis from human fibroblasts (Krusius & Ruoslahti, 1986). This small proteoglycan regulates numerous functions in the extracellular matrix including collagen fibrillogenesis (Reed & Iozzo, 2002), collagen degradation (Bhide et al., 2005), cell growth (Kinsella et al., 2004) and extracellular matrix signalling (Seidler et al., 2006). It is believed that decorin allows collagen fibrils to increase in diameter up to a certain point and then prevents further enlargement (Canty & Kadler, 2002). The core protein contains a 30 amino acid residue propeptide of which the first 16 residues are likely to represent a signal peptide, a central domain containing ten LRRs flanked by disulfide-bonded terminal sequences (Day et al., 1986; Krusius & Ruoslahti, 1986).
A feature of decorin is that it contains one GAG chain of either chondroitin or dermatan sulphate (depending on the tissue) attached to the serine residue at position 4 of the amino-terminal amino acid sequence (Chopra et al., 1985) and the central domain has three attachment sites for amino-linked oligosaccharides. Removal of the GAG chain or the amino-terminal 17 amino acid residues of the decorin protein does not affect the ability of decorin to inhibit fibrillogenesis (Vogel et al., 1987) suggesting this interaction is core protein orientated (Vogel et al., 1987).
Decorin was named due to its electron microscopic appearance on the collagen network, decorating the Type I collagen fibres at the ‘d’ and ‘e’ bands (Scott, 1980; Scott et al., 1981; Scott & Orford, 1981). Decorin can be found in many connective tissues where it is found in great abundance such as tendon, bone, sclera, skin, aorta and cornea, although originally it was isolated from the cartilage and bone. It has been shown that decorin represents up to 1% of tendon dry weight (Vogel & Meyers, 1999; Derwin et al., 2001).
In tendon, decorin constitutes approximately 80% of the total proteoglycan content in the proximal region or the tensile region where tendon attaches to muscle (Samiric, 2003). As a consequence of the binding of decorin to the surface of collagen fibrils, the lateral assembly of triple helical collagen molecules is delayed (Vogel et al., 1984) and the final diameter of the collagen fibrils becomes thinner (Vogel & Trotter, 1987). However, it may be important in the organogenesis and regulation of cell division and differentiation, since its protein core has been shown to bind transforming growth factor-β (TGF-β; Iozzo, 1997). The core protein contains 10 LRRs flanked by disulphide bond stabilised loops on both sides. The core protein contains additional sites for glycosylation (amino-linked glycosylation sites) within the LRRs (Krusius & Ruoslahti, 1986).
Decorin was shown to interact with collagen via its core protein and influence collagen fibrillogenesis (Vogel et al., 1984). Decorin interacts with collagen Types I, II, III, V, VI, XII and XIV to modulate collagen fibrillogenesis (Vogel et al., 1984; Bidanset et al., 1992; Font et al., 1993; Hedbom & Heingard, 1993; Whinna et al., 1993; Thieszen & Rosenquist, 1994; Vogel et al., 1994; Font et al., 1996) as well as fibronectin (Schmidt et al., 1987), thrombospondin (Winnemoller et al., 1992), the complement component C1q (Krumdieck et al., 1992), low-density lipoprotein, the receptor required for its endocytosis (Hausser et al., 1989), epidermal growth factor receptor (EGFR) and TGF-β (Yamaguchi et al., 1990; Hilderbrand et al., 1994; Kovanen & Pentikainen, 1999). Decorin neutralises the growth stimulating effect of TGF-β, thereby regulating various cell processes including cell proliferation, differentiation, adhesion and deposition of the extracellular matrix and has been shown to suppress tumour cell growth by binding to an EGFR (Moscatello et al., 1998). The LRR motif of decorin enables the proteoglycan to take the shape of a horseshoe has been shown to bind to a single triple helix of collagen (Weber et al., 1996).
Decorin binds to collagen primarily via LRRs 4-5, which is composed of 40 amino acid residues (Svensson et al., 1995). The protein core of decorin is compact and horseshoe-shaped, a shape suitable for protein-protein interactions (Scott, 1996). The collagen-binding site is located in the cysteine-free central domain of the decorin core protein (Svensson et al., 1995). Decorin is found primarily at a distance from cells (Bianco et al., 1990). Recent studies have shown that decorin may be capable of forming a dimeric structure (Scott et al., 2003). At specific sites every 64-68 nm, decorin is found connected to collagen fibres where it modulates collagen fibril formation (Scott et al., 1981; Hedbom & Heinegard, 1993). A recent study has identified the sixth leucine motif of decorin as the major collagen I-binding site in vitro (Kresse et al., 1997).
Biglycan is a small proteoglycan and its primary structure has been deduced by cDNA analysis in humans (Fisher et al., 1989). The biglycan core protein is substituted with two GAG chains consisting of either chondroitin sulphate or dermatan sulphate. These GAGs are attached to the amino-terminus of biglycan. Dermatan sulphate is more abundant in most tissues and is attached to serine residues at position 5 and 10 of the amino-terminal amino acid sequence on its 38-42 kDa core protein (Kresse et al., 2001).
Biglycan was originally found like decorin, in cartilage and bone and their amino acid sequences have a high homology (Fisher et al., 1983 Rosenberg et al., 1985), it is also found in tendon, capillary endothelium, skeletal muscle, dermis and skin (Fisher et al., 1989; Bianco et al., 1990; Schonherr et al., 1993; Ameye et al., 2002). In humans the core protein consists of 331 amino acids and is encoded by an exon gene on chromosome X (Danielson et al., 1993).
Interactions of biglycan with collagen Types I, V, VI and XIV have been shown both via its core protein or its GAG chains, however with lower affinity than decorin (Schonherr et al., 1995). The binding affinity for Type I collagen is low, but much higher with Type VI collagen, with which it forms hexagonal networks (Wiberg et al., 2002). Like decorin the core protein contains 10 LRRs flanked by disulphide bond stabilised loops on both sides. Biglycan interacts with collagen Type VI and the complement component C1q (Krumdieck et al., 1992). Biglycan is a Zn2+-binding protein (Yang et al., 1999).
Like decorin, biglycan binds to TGF-β, and therefore has the ability to participate in modulation of cell proliferation (Ruoslahti & Yamaguchi, 1991; Border et al., 1992). Biglycan has the ability to bind to Type I collagen, this interaction is dependent on the presence of amino-linked oligosaccharides of the biglycan core protein. Biglycan has been shown to most commonly occur on the cell surface or in the pericellular space of connective tissues (Bianco et al., 1990; Fedarko et al., 1992).
Another member of the small proteoglycan family is fibromodulin. Fibromodulin was originally identified in cartilage as a 59 kDa protein and contains 10 internal repeats of 24 amino acid residues rich in leucine and is a member of class II proteoglycans (Lorenzo et al., 2001). These repeats are located on the central part of the core protein and make up approximately 80% of all amino acid residues (Ruoslahti & Yamaguchi, 1991; Iozzo & Murdoch, 1996; Lorenzo et al., 2001). It is also found in other connective tissues including tendon and ligament. It contains five asparagine residues in the central region of the LRRs (Asn-X-Thr/Ser; Oldberg et al., 1989) four of which may simultaneously have keratan sulphate (type I keratan sulphate) or amino-linked oligosaccharide bound covalently (Plaas et al., 1990). At the amino-terminal end of the fibromodulin core protein is a tyrosine rich region that is not present in decorin or biglycan (Antonsson et al., 1991).
Fibromodulin is involved in regulating the orientation of collagen fibres and is known to bind to collagen Types I, II, VI, XI and XII as well as TGF-β and inhibits fibrillogenesis of both Type I and Type II collagens. There is some evidence to suggest that fibromodulin binds to these collagens in a different position to decorin (Hedbom & Heinegard, 1993; Hildebrand et al., 1994; Font et al., 1996). This suggests that fibromodulin may have a similar role to decorin within the tendon extracellular matrix. This proteoglycan contains four keratan sulphate chains attached to the amino-terminus end attached via an N-glycosidic linkage to asparagines.
Its molecular weight varies with age, tissue and animal (Ezura et al., 2000). It plays a key role in the formation of mature large collagen fibrils (Jepson et al., 2002). Fibromodulin is expressed at high levels in tendon (Oldberg et al., 1989) and binds to the exact same region on Type I collagen as lumican does (Svensson et al., 1999). This site is however, separate from the decorin-binding site (Hedbom & Heinegard, 1993). Like decorin the protein core of fibromodulin is compact and horseshoe-shaped, a conformation suitable for protein-protein interactions (Scott, 1996). As with decorin, fibromodulin has the ability to decorate the surface of collagen fibres and therefore may have a role in regulating collagen fibril diameter.
Lumican is a highly sulphated keratan sulphate containing proteoglycan that was initially found in the cornea containing two to three keratan sulphate chains (Chakravarti et al., 1995). It is also present in tendon (Funderburgh et al., 1987), articular cartilage (Knudson & Knudson, 2001), intestine (Blochberger et al., 1992), skin, lung (Dolhnikoff et al., 1998) as well as the kidney (Schaefer et al., 2000). There is a lot of information on corneal lumican, much less is known about this molecule in other connective tissues including tendon. Lumican most closely resembles fibromodulin (Blochberger et al., 1992; Grover et al., 2000) and like fibromodulin belongs to the class II proteoglycans (Iozzo, 1997; Iozzo, 1999; Jepsen et al., 2002).
Like fibromodulin it also contains 10 LRRs (Iozzo & Murdoch, 1996; Henry et al., 2001; Lorenzo et al., 2001) with which it shares the same collagen binding site (Jepson et al., 2002), but is slightly smaller with a molecular weight of 38 kDa and contains fewer keratan sulphate chains (2-3; Iozzo & Murdoch, 1996).
Lumican binds to the fibrillar collagens in connective tissues (Blochberger et al., 1992; Chakravarti et al., 1995). It binds to Type I collagen, inhibits the size of collagen fibrils and is involved in the modulation of tendon strength (Rada et al., 1993). The lumican core protein, without the keratan sulphate side chains, is equally efficient in inhibiting collagen fibrillogenesis in vitro, suggesting this function to be entirely core protein mediated (Rada et al., 1993). The protein core of lumican is compact and horseshoe-shaped and is involved in protein-protein interactions (Scott, 1996). Lumican contains additional clusters of tyrosine-sulphate residues in its core protein (Sandy et al., 1997).
1.2.3 Large Aggregating Proteoglycans
A characteristic feature of the large aggregating proteoglycans is the presence of globular domains separated by a GAG attachment region (Abbaszade et al., 1999). The large proteoglycans can be divided into two sub families, the first is the hyalectins which have the ability to bind to hyaluronan (Iozzo & Murdoch, 1996; Aumailley & Gayraud, 1998; Iozzo, 1999). Members of this group of proteoglycans include aggrecan, versican, neurocan and brevican (Iozzo & Murdoch, 1996; Iozzo, 1998). All of the hyalectins have a three-domain structure which includes an amino- terminal domain for hyaluronan binding, a central domain that contains the GAG chains and a carboxyl-terminal lectin-like domain (Iozzo & Murdoch, 1996; Iozzo, 1998).
The second family of large proteoglycans include the non-hyaluronan binding proteoglycans, which include perlecan, agrin and testican which have been less studied than the hyalectins. The large proteoglycans are sometimes called the modular proteoglycans or leticans and are rich in chondroitin and keratan sulphate chains (Iozzo & Murdoch, 1996). They have a large core protein (100-370 kDa) and are mostly entrapped within and between collagen fibrils and fibres (Iozzo, 1998). Due to the large proteoglycans high fixed charge density they are stiffly expanded to provide the collagen fibres with a high capacity to resist high compressive and tensile forces
(Iozzo, 1998). During domain stress these molecules are compressed by approximately 20% (Iozzo, 1998).
The most studied large proteoglycans are aggrecan (see Figure 1.4; Hardingham & Fosang, 1995) and versican (see Figure 1.4; Margolis & Margolis, 1994). Although aggrecan and versican have primarily been studied in articular cartilage and blood vessels respectively, they have also been identified in tendon (Vogel & Heinegard, 1985; Vogel & Thonar, 1988; Vogel et al., 1994; Rees et al., 2000; Samiric, 2003). This review will focus on the large proteoglycans that are present in tendon.
Aggrecan was first described by Shatton and Schubert in 1954 as a mucoprotein. As many as one hundred individual aggrecan monomers interact with hyaluronan to form an aggregate of up to several hundred million in molecular weight (Heinegard & Hascall, 1974; Buckwalter & Rosenberg, 1982). The aggrecan monomer consists of a large core protein (MW 220-250 kDa) containing three globular domains (G1, G2 and G3; Iozzo & Murdoch, 1996). At the amino-terminus end of the core protein is the G1 domain (Heinegard & Hascall, 1974), which non-covalently and specifically binds to hyaluronan (Fosang & Hardingham, 1989). The G1 domain of aggrecan is homologous to link protein (Hering et al., 1997) and is separated from the second homologous domain (G2) by the linear interglobular domain (IGD; Fosang & Hardingham, 1989).
The IGD has been shown to be susceptible to proteolytic degradation by many different classes of proteinases (Hardingham & Fosang, 1995). The G2 domain of aggrecan has no known function. This domain is unique to the aggrecan protein and consists of tandem repeats just like the G1 domain but does not interact with hyaluronan or link protein (Fosang & Hardingham, 1989). At the G3 carboxyl-terminus end are two alternatively spliced epidermal growth factor (EGF-1) like regions (Baldwin et al., 1989; Fulop et al., 1993) a lectin like G3 domain (Doege et al., 1986) and an alternatively spliced domain which has sequence similarity to complement-regulatory proteins.
These motifs suggest that the G3 globular domain has a role in cell adhesion (Siegelman et al., 1990) and is involved in GAG synthesis and secretion of the proteoglycan (Kiani et al., 2002). The C-type lectin motif has been shown to bind to fructose and galactose (Halberg et al., 1988; Saleque et al., 1993; Hardingham et al., 1994) and various proteins (Watanabe et al., 1998), indicating binding to cellular or matrix ligands. Some aggrecan monomers lack the G3 domain possibly due to proteolytic cleavage (Weidemann et al., 1984; Dennis et al., 1990).
Attached to the core protein are many GAG chains. Aggrecan is a highly glycosylated proteoglycan with numerous chondroitin and keratan sulphate chains attached (Iozzo & Murdoch, 1996). A typical aggrecan molecule may contain up to 100 chondroitin sulphate chains and the chains are either 4-sulphated or 6-sulphated, or usually both, which are typically approximately 20 kDa each (Hardingham & Muir, 1973) and as many as 60 keratan sulphate chains and they are usually of shorter length, which are usually 5-15 kDa (Hascall & Riolo, 1972; Doege et al., 1991; Roughley & Lee, 1994) and other oligosaccharides (Hascall & Kimura, 1982).
The chondroitin sulphate attachment domains are the major GAG bearing regions of aggrecan that contain approximately 100 Ser-Gly dipeptide repeats, that are attachment sites for chondroitin sulphate chains (Roughley & Lee, 1994). The chondroitin sulphate domains include chondroitin sulphate subdomain-1 (CS-1) and subdomain-2 (CS-2; Hardingham et al., 1994). The keratan sulphate attachment domain is located in the first portion of the extended region of the core protein between the G2 and G3 domains (Hardingham et al., 1994). The majority of keratin sulphate is found in the keratan sulphate domain.
Approximately 50 keratan sulphate chains are attached to aggrecan in this region (Hascall, 1977). Aggrecan also contains approximately 50 O-linked oligosaccharides and approximately 5-10 N- linked oligosaccharides (Lohmander et al., 1980; Iozzo & Murdoch, 1996) that are covalently attached to the core protein. The O-linked oligosaccharides have a linkage to protein similar to that for keratan sulphate (Iozzo & Murdoch, 1996). The large number of GAG side chain substitution and the resulting fixed charge density attracts counter-ions and water through osmotic processes; this causes a swelling pressure that is crucial for the biomechanical properties of this tissue (Heinegard & Oldberg, 1989), resulting in a stabilized extracellular matrix containing many negatively charged GAG chains which enable aggrecan to attract water (Maroudas et al., 1969). This negatively charged environment leads to increased osmotic pressure, and thus to enhanced tissue hydration.
The aggrecan monomer binds to hyaluronan via the proteoglycan tandem repeat associated with the aggrecan G1 domain and this interaction is stabilised by link protein (Perin et al., 1987; Valhmu et al., 1998). Most of the hyaluronan-binding molecules, including link protein and aggrecan contain proteoglycan tandem repeats. In humans, the protein core structure of aggrecan is the product of a single gene copy residing on chromosome 15q26 (Korenberg et al., 1993) and 15q25→q26.2 (Just et al., 1993).
Parts of tendon that are commonly exposed to compressive forces indicate the presence of fibrocartilage which is rich in aggrecan as well as Type II collagen, these are commonly found in cartilage (Benjamin & Ralphs, 1998). The interactions between aggrecan and collagen reduce the stiffness and viscosity of the collagen network. Aggrecan generates a hydrating osmotic swelling pressure in equilibrium that is opposed by tensile stresses in the surrounding elastic collagen network (Maroudas, 1976).
This protects the bone ends from wear during joint movements. Aggrecan is essential to the viscoelasticity and acts as a lubricant, allowing collagen fibrils to easily move over one another (Benjamin & Ralphs, 1998). The keratan sulphate chains of aggrecan undergoes structural changes and increases in molecular weight with age, due to an increase in fucosylation, sialyation and galactose sulphation (Brown et al., 1998).
Versican is a member of the hyaluronan-binding proteoglycans and is the largest member of this group with a core protein of ~ 400 kDa (Zimmermann & Ruoslahti, 1989; Iozzo & Murdoch, 1996). This proteoglycan is expressed by a large variety of tissues including tendon as well as loose connective tissues, the central and peripheral nervous tissues, embryonic tissue, epidermis and in all three wall layers of veins and elastic arteries (Kimata et al., 1986; Zimmermann & Ruoslahti, 1989; Shinomura et al., 1990; Yamagata et al., 1993; Bode-Lesniewska et al., 1996; Wight, 2002).
The versican gene and protein follow a domain template and contains an amino-terminal G1 domain, homologous to the G1 domain of aggrecan, which binds to both hyaluronan (LeBaron et al., 1992; Watanabe et al., 1997) and link protein, and a carboxyl-terminal G3 domain which consists of set of lectin- like domain (also called carbohydrate recognition domain), two EGF like sequences and complement-binding protein-like subdomains, however it lacks the G2 domain (Zimmermann & Ruoslahti, 1989; Margolis & Margolis, 1994). The carboxyl-terminal G3 domain binds fibulin-1 and -2 (Aspberg et al., 1999; Olin et al., 2001), tenascins (Aspberg et al., 1997) and heparan sulphate proteoglycans (Ujita et al., 1994).
The central region of the versican core protein is encoded by two large exons that specify the chondroitin sulphate attachment regions of versican, of which there are 15 potential sites (Zimmermann & Ruoslahti, 1989; Margolis & Margolis, 1994). It is because of these domains that this ‘versatile’ proteoglycan received its name (Zimmermann & Ruoslahti, 1989).
The structural and functional diversity of versican is increased by variations in the GAG sulphation patterns and the type of GAG chains bound to the core protein. There is a single versican gene; however alternative splicing of its mRNA produces four different versican isoforms that differ in their potential number of GAG chains (Ito et al., 1995; Zako et al., 1995). All isoforms have homologous amino-terminal and carboxyl-terminal domains. The central domain of versican V0 contains both the GAG-α and GAG-β domains (Dours-Zimmermann & Zimmermann, 1994; Naso et al., 1994). The V1 isoform has GAG-β domain, V2 has the GAG-α domain and V3 is void of any GAG attachment domain and only consists of the amino- and carboxyl-terminal globular domains.
The number of amino acid residues for the different splice variants of the mature versican core proteins are 3361 (versican V0), 2380 (versican V1), 1623 (versican V2) and 642 (versican V3). Little information exists about the tissue distributions of the four-versican variants due to the lack of immunological reagents that would distinguish the various versican isoforms (Cattruzza et al., 2002). Versican forms aggregates by binding to hyaluronan through its G1 domain (Margolis & Margolis, 1994). The chondroitin sulphate chains that are attached to the core protein vary in size from 25 kDa to 80 kDa and they contain differing ratios of chondroitin-4-sulphate and chondroitin-6-sulphate (Schonherr et al., 1991).
Versican was originally isolated from fibroblasts and contains 12-15 chains of chondroitin sulphate covalently attached to the central sequence (Krusius et al., 1987; Zimmermann & Ruoslahti, 1989). The entire primary structure of versican has been generated from human (Zimmermann & Ruoslahti, 1989; Dours-Zimmermann & Zimmerman, 1994), murine (Ito et al., 1995), bovine (Schmalfeldt et al., 1998), and chick (Shinomura et al., 1993) cDNA clones. The amino-terminus of versican has an important role in maintaining the integrity of the extracellular matrix by interacting with hyaluronan.
Versican is a member of the hyaluronan binding proteoglycans and plays an important role in cell adhesion (Yamagata et al., 1993), cell growth, migration (Landolt et al., 1995) proliferation and differentiation (Kishimoto et al., 1999). Expansion of the extracellular matrix and increased viscoelasticity of the pericellular matrix that supports cell shape changes that is a necessity for cell proliferation and migration is a common occurrence when versican levels increase (Oldberg et al., 1990).
Versican is found in low levels in the tensional regions of tendon but is found in higher levels in the compressed regions of tendon. Versican is synthesized by fibroblasts and keratinocytes (Zimmermann et al., 1994), arterial smooth muscle cells (Schonherr et al., 1991; Yao et al., 1994), the central nervous system (Bignami et al., 1993) and also mesangial cells of the kidney (Thomas et al., 1994).
1.2.4 Glycosaminoglycans (GAGs)
GAGs are complex unbranched carbohydrate polymers, consisting of repeating disaccharide units that contain one hexosamine and one hexuronic acid that may be sulphated (Poole, 1986). They have a great assortment of biological functions, some of which are known to depend on the presence of specific sulphated sequences that give them the ability to bind to protein ligands and influence their activities (Spillman & Lindahl, 1994). They are O-linked to serine through a linkage sequence at its reducing end (Poole, 1986).
GAGs present in the extracellular matrix play a central role in the modulation of cell signals. All GAGs are negatively charged and form a proteoglycan-GAG complex. The proteoglycan-GAG interaction maintains the hydration of the extracellular matrix; the level of hydration is dependent on the number of GAG chains and on the restriction placed on proteoglycan swelling by the surrounding collagen fibres (Hardingham & Fosang, 1992). GAGs include chondroitin sulphate and dermatan sulphate, which arise from the same precursor, keratan sulphate, heparan sulphate, heparin and hyaluronan.
Decorin, Aggrecan, Versican, Biglycan, Epyphycan, Perlecan, Syndecans-1 and -4, Type IX and XII Collagen
(β1-4)-D-Glucuronic acid (β1-3)N-acetyl-D-galacosamine
Decorin, Biglycan, Epyphycan, Syndecans -1 and -4, Type IX Collagen
(β1-4)-L-Iduronic acid (β1-3) N-acetyl-D-galacosamine
Aggrecan, Fibromodulin, Lumican
(β1-3)-D-Galactose (β1-4)N- acetyl-D-glucosamine
Heparin and Heparan Sulphate
Aggrecan, Perlecan, Syndecans, Glypicans
Hepain: Iduronic acid (β1-4)N-acetyl-D-glucosamine Heparin Sulphate: (β1-4)-D-Glucuronic acid (β1-4)N-acetyl-D-glucosamine or (β1-4)-L-
Exists as a macromolecule within the extracellular matrix
(β1-4)-D-Glucuronic acid (β1-3)N- acetyl-D-glucosamine
Hyaluronan (also called hyaluronic acid or hyaluronate) was first discovered by Meyer and Palmer (1934) when they isolated it from the vitreous body of the eye. It plays a vital role in the formation and stability of the extracellular matrix of tendon via its association with the hyaluronan binding protein called link protein which will be discussed later (Day & Prestwich, 2002; Tammi et al., 2002). In addition, hyaluronan can bind to versican (LeBaron et al., 1992), aggrecan (Hardingham & Muir, 1973), neurocan (Rauch et al., 1992) and the cell surface receptor CD44 (Jalkanen et al., 1987). Indeed, in cartilage where hyaluronan provides load bearing capabilities, removal of link protein (Watanabe & Yamada, 1999) or aggrecan (Watanabe et al., 1997) results in defects in the tissue.
It is a large 100-1000 kDa non-sulphated negatively charged GAG that consists of repeating disaccharide units of glucuronic acid and N-acetylglucosamine (Weissmann & Meyer, 1954). Hyaluronan occurs in most connective tissues including tendon, articular cartilage and ligament (Brandt et al., 1976; Fraser et al., 1997; Vogel & Peters, 2005). An important feature of hyaluronan is its ability to bind large amounts of water (1000-fold of its own weight). Hyaluronan is the only GAG that is not attached to a core protein and is unsulphated.
Hyaluronan plays an important role in tissue remodelling during development, normal tissue homeostasis and disease (Knudson & Knudson, 1993). The interaction of hyaluronan with matrix hyaluronan-binding proteins and cell-surface hyaluronan receptors regulates many aspects of cell activities including cell migration, cell differentiation and cell-cell adhesion (Knudson & Knudson, 1993). Its structure is the simplest of all GAGs. Aggrecan is retained in the extracellular matrix by forming aggregates with hyaluronan via link protein (Hascall & Sajdera, 1969; Hardingham, 1979).
126.96.36.199 Chondroitin Sulphate
Chondroitin sulphate is composed of repeating disaccharide units of differently sulphated and/or unsulphated alternating residues of glucuronic acid and N-acetylgalactosamine (Kelly, 1998). These disaccharides are generally sulphated in the 4 or 6 position of the galactosamine residues (Roughly & Lee, 1994). The sulphate group in the chondroitin-6-sulphate is spatially more freely oriented, which is important for its association various extracellular matrix components particularly with collagen. The attachment of chondroitin sulphate to proteoglycans occurs via xylose-galactose-galactose trisaccharide. The sugar xylose of the trisaccharide is linked by an O-glycoside bond to the serine residue of the successive serine-glycine residues in the proteoglycan core protein.
188.8.131.52 Dermatan Sulphate
Dermatan sulphate is a linear sulphated GAG made up of repeating disaccharide units containing iduronic acid and N-acetylgalactosamine (Roughly & Lee, 1994). It has a molecular weight of 15-40 kDa (Comper & Laurent, 1978). The most significant difference between dermatan sulphate and chondroitin sulphate is by the presence of iduronic acid residues instead of glucuronic acid residues (Roughly & Lee, 1994). The iduronic acid residues may be sulphated at the C2 position and the galactosamine residue may be sulphated at the C4 or C6 position, though disulphated residues sometimes occur (Roughly & Lee, 1994).
It has been suggested that iduronic acid may have a role in the binding site specificity for GAG binding proteins. Dermatan sulphate is attached covalently via an O-xylose linkage to serine residues of core proteins to form dermatan sulphate proteoglycans.
184.108.40.206 Keratan Sulphate
Keratan sulphate was first identified in 1939 by Suzuki from extracts of the cornea (Suzuki, 1939). The keratan sulphate molecule is composed of repeating disaccharide units of D-Galactose and N-acetyl-D-glucosamine. The N-acetyl glucosamine residue is commonly sulphated at its 6-position, but sulphation may also occur at the 6-position of the galactose residues (Plaas et al., 1992; Funderburgh, 2000). In mammalian tissue there are two forms of keratan sulphate including keratan sulphate I (10-26 kDa) and keratan sulphate II (5 kDa).
Keratan sulphate II can be further divided into keratan sulphate II-A and keratan sulphate II-B. Keratan sulphate is the only GAG that does not contain uronic acid residues (Roughly & Lee, 1994). Unlike the other GAGs, keratan sulphate contains oligosaccharide branches. It is however still considered a GAG because it contains a repeating disaccharide of a 6-sulphated N-acetylglucosamine linked to galactose. This portion is unbranched and variable in size.
220.127.116.11 Heparin and Heparan Sulphate
Heparin (heparin monosulphate) and heparan sulphate share the same basic structure of repeating disaccharides of D-glucuronic acid and N-acetylglucosamine (Sasisekharan & Venkataraman, 2000). The size of an individual chain can reach 100 kDa, but normally they are below 50 kDa. Heparan sulphate differs from the above GAGs not only in its structure but also in its restricted presence on cell associated and basement membrane proteoglycans (Gallagher et al., 1986). Heparin is considered a modified form of heparan sulphate (Park et al., 2000).
It utilizes the same biosynthetic machinery, but the polymer modifications occur to a greater extent and a unique core protein is used (Lindahl, 1990). Heparin chains show variability with molecular weights but range between 4 to 40 kDa (MacHarg & Becht, 1983). Heparan sulphate is less sulphated than heparin (Sasisekharan & Venkataraman, 2000). Heparan sulphate and heparin can be found on the cell surface of all animal cells (Park et al., 2000).
1.2.5 Non-Collagenous Proteins
In tendon there are a number of non-collagenous proteins present however little is known about them (Kannus et al., 1998; Riley, 2000). The non-collagenous proteins are predominately involved in cell-matrix organisation or cell-matrix signalling. This review will attempt to analyse the major non-collagenous proteins that are found in the tendon extracellular matrix including elastin, link protein, cartilage oligomeric matrix protein (COMP) and tenascin-C.
Elastin provides elasticity and resilience to tendon as well as elastic cartilage, large arteries, lungs, ligament and skin. It is an insoluble polymer consisting of several or more tropoelastin molecules covalently bound to each other by cross-links. Elastin is a highly hydrated molecule in spite of it being a hydrophobic molecule. Elastin is an
Function in Tendon
Plays a role in cell interactions
Mediates cell-matrix interactions; forms structures with versican
Binds to various extracellular matrix components
Binds hyaluronan to aggrecan, versican etc
Gives tendon elasticity and resilience
Cartilage Oligomeric Matrix Protein
Structural and interactive role with the cell population
Forms fibrillar networks
Forms elastic fibres; provides elastic properties of tissue
Extremely stable molecule; its turnover is extremely slow, so slow in fact that it may last an entire lifetime. Representing less than 2% of the tendon dry weight (Eyre et al., 1984; Hess et al., 1989; Eyden & Tzaphlidou, 2001), elastin is thought to give the extracellular matrix its elastic properties (Elliott, 1965; Butler et al., 1978). Electron microscopy reveals that elastic fibres are approximately 0.3-2.0 mm in diameter and consist of two distinct morphological components: an amorphous central bulk (core) and electro-dense filaments (Jozsa et al., 1979). Elastin does not form helices and is hydrophobic. Very little elastin is found in healing wounds. Elastin contains the two amino acids called desmosine and isodesmosine that form cross-linkages between adjacent tropoelastin chains and are important in imparting the elastic properties to elastin (Rosenbloom, 1993).
18.104.22.168 Link Protein
Link protein is a small non-collagenous protein of approximately 45 kDa and was first discovered by Hascall and Sajdera in 1969. Link protein is a non-collagenous protein that stabilises the interaction of aggrecan with hyaluronan by binding to hyaluronan to the G1 domain of aggrecan (Le Gledic et al., 1982; Neame et al., 1986; Nguyen et al., 1991). When link protein is lacking the aggregates they are a part of are smaller and less stable than they are in the presence of link protein as estimated by rotary shadowing of electron micrographs (Morgelin et al., 1988). Structurally link protein is very similar to the G1 domain of aggrecan as it consists of three disulphide-bonded loops (A, B and B’).
Link protein interacts with aggrecan with 1:1 stoichiometry (Faltz et al., 1979) and exists in three forms -1, -2 and -3. Link proteins -1 and -2 share similar protein structures but contains one or two N-linked oligosaccharides respectively in its amino-terminal region (Roughley et al., 1982). Link protein -3 is a product of the removal of the first 16, 18 or 23 amino acid residues at its amino-terminus (Le Gledic et al., 1983; Nguyen et al., 1991; Neame & Barry, 1993). Link protein is encoded for by a single gene and in humans, the gene is found on chromosome 5 in the q13→q14.1 region (Osbourne-Lawrence et al., 1990).
The amino acid sequence of link protein can be divided into three domains, an amino terminal domain that falls into the immunoglobulin super-family and two carboxyl-terminal domains that are similar to each other. The two carboxyl-terminal domains are responsible for the association with hyaluronan. Link protein stabilizes interactions between aggrecan monomers and hyaluronan (Oldberg et al., 1990; Jarvinen et al., 1997). This stabilisation is achieved via its non-covalent interaction with both hyaluronan and the G1 domain of aggrecan (Nguyen et al., 1991). Link protein has been shown to be important in maintaining aggregates in knockout mice which show signs of cartilage defects causing dwarfism and craniofacial abnormalities (Watanabe & Yamada, 1999). In addition, the absence of link protein causes aggregates to be smaller and less stable than they are in the presence of link protein (Morgelin et al., 1988).
Tenascin-C is a disulphide-linked hexametric protein with subunits of 200-300 kDa in humans; this is created by alternative splicing of a single gene transcript (Gulcher et al., 1991; Chiquet, 1992). In the fibrocartilaginous region of tendon, tenascin-C is predominately cell-associated (similar to Type VI collagen) and may play a part in the development of the chondrocyte cell phenotype (which includes the expression of Type II collagen and aggrecan) in response to compressive load (Riley et al., 1996a).
Each subunit consists of the following structural domains: an amino-terminal cysteine-rich region and heptad motifs, followed by EGF-like repeats, fibronectin Type III domains and a carboxyl-terminal globular domain homologous to fibrinogens. Depending on the species, the number of EGF-like repeats as well as the fibronectin Type III repeats differs (Vrucinic-Filipi & Chiquet-Ehrismann, 1993). Tenascin-C may play a role in collagen fibre alignment and orientation (Mackie & Ramsey, 1996), and is found in abundance in the tendon body and at the osteotendinous and myotendinous junctions (Riley et al., 1996a).
Tenascin-C is generally called an anti-adhesive protein, since many cells do not adhere to tenascin or if they adhere they do not spread. It is predominantly found in the distal regions of tendon, which helps establish and maintain the fibrocartilagenous region by decreasing cell-matrix adhesion (Kannus et al., 1998; Mehr et al., 2000).
22.214.171.124 Cartilage Oligomeric Matrix Protein (COMP)
COMP was first observed in cartilage and has been found in some tendons (Smith et al., 1997; Vogel & Meyers, 1999) where it is distributed around and within tendon bundles (DiCesare et al., 1994). COMP is considered to have a structural and interactive role with cell populations (DiCesare et al., 1994) as well as Type I and II collagen (Rosenberg et al., 1998). COMP as the name implies is commonly found in cartilage but is also present in tendon representing up to 3% of tendon dry weight (Hedbom et al., 1992; DiCesare et al., 1994). COMP is a large 524-kDa-pentameric molecule that is comprised of five disulphide-bonded subunits and is a member of the thrombospondin gene family (thrombospondin 5; Oldberg et al., 1992; Muller et al., 1998).
COMP forms pentamers composed of 5 identical subunits (Hedbom et al., 1992; Morgelin et al., 1992) each of approximately 86 kDa (Zaia et al., 1997) held together at the amino-terminal by a coiled-coil domain (Efimov et al., 1994; Sodersten et al., 2006) and is further stabilized by disulfide bridges within this region. This domain is followed by a flexible region of four EGF like-domains, a region encompassing eight thrombospondin type 3 (calcium-binding) domain, and contains collagen-binding sites at the carboxyl-terminal globular domain (Oldberg et al., 1992), which allows COMP to bind to more than one collagen molecule at the same time (Sodersten et al., 2006).
COMP is a major component of tendon represents up to 3% of the total non-collagenous protein present in tendon (DiCesare et al., 1994; Smith-Mungo & Kagan, 1997; Muller et al., 1998; Smith et al., 1999). COMP binds via each carboxyl-terminal globule equally to one of four sites on collagens I and II, which depends on the presence of zinc ions (Rosenberg et al., 1998) and also has high affinity interactions with Collagen Type IX (Holden et al., 2001; Thur et al., 2001).
126.96.36.199 Other Non-Collagenous Proteins
Fibrillins are extracellular matrix macromolecules whose primary functions are architectural: fibrillins assemble into ultrastructurally distinct microfibrils that are common in the connective tissue space. There are two types of fibrilin in tendon; they include fibrilin-1 (Sakai et al., 1986; Maslen et al., 1991; Corson et al., 1993; Pereira et al., 1993) and fibrilin-2 (Lee et al., 1991; Zhang et al., 1994). It has been shown that fibrilin-1 is a major structural component of connective tissue microfibrils and they play a major role in the maintenance of microfibrils and elastic fibres in humans (Hollister et al., 1990; Dietz et al., 1991) and mice (Pereira et al., 1997; Pereira et al., 1999).
Laminins are a family of macromolecules with numerous functions and are common in basement membranes of connective tissues such as tendon where it is found in the vascular walls. Laminin is found in the vascular walls of tendon and the myotendinous junction, and is involved in cell adhesion (Kannus et al., 1998; Pakkala et al., 2002). In mice lacking laminin, growth retardation and muscle dystrophy occurs.
Matrillin-2 is a large oligomeric matrix protein that forms fibrillar networks via its von Willebrand factor type A - like module in most connective tissues such as tendon (Piecha et al., 1999).
Fibronectin is a high molecular weight non-collagenous protein that binds to receptor proteins that span the cells membrane called integrins. Additionally, they bind to extracellular matrix components such as collagen, fibrin and heparin. Fibronectin can bind directly to several collagens (Johansson & Hook, 1980; Cidadao, 1989) and a collagen-binding region in fibronectin has been characterized (Owens & Baralle, 1986; Obara & Yoshizato, 1997). It helps stabilise the attachment of the extracellular matrix to cells by acting as binding sites for cell surface receptors and helps create a cross-linked network within the extracellular by having binding sites for other extracellular matrix components. Fibronectin participates in cell interactions with the extracellular matrix, and affects a range of cell functions including cell adhesion, cell migration, differentiation, haemostasis, phagocytosis and chemotaxis (Labat-Robert et al., 1990; Aumailly & Gayraud, 1998; Xie et al., 1998; Hopf et al., 1999).
There are five members in the thrombospondin (TSP) family (TSP-1 to -5) of secreted, modular glycoproteins whose functions in the extracellular matrix are diverse and poorly understood (Frazier, 1991; Adams & Lawler, 1993; Bornstein & Sage, 1994). Of the five members of the TSP family (TSP-1 to TSP-5), TSP-1, TSP-2 and TSP-4 are found in the extracellular matrix of tendons (Kannus et al., 1998).
1.2.6 Collagen Synthesis
The fibril forming collagens are the most comprehensively studied of the collagen types particularly Type I collagen. Collagen synthesis is a complex multi step process that starts with the transcription and translation of the individual collagen gene (for reviews see; Kielty et al., 1993; Prockop & Kivirikko, 1995; Bateman et al., 1996). The synthesis of collagen involves many co- and post-translational modifications of their polypeptide chains (Kivirikko & Pihlajaniemi, 1998; Myllyharju, 2003)
The pro-alpha chains are firstly synthesized on the ribosomes of the rough endoplasmic reticulum of fibroblasts (McAnulty & Laurent, 1987). The left-handed helix of each alpha chain is formed in the rough endoplasmic reticulum also. Procollagen is formed following hydroxylation of proline and lysine residues as well as the addition of carbohydrates, this occurs inside the endoplasmic reticulum (Zernicke & Loitz, 1992).
Procollagen molecules are processed through the Golgi apparatus and then collected into secretory vesicles and taken to the cell membrane where they are secreted via exocytosis (Harwood et al., 1976). When the procollagen enters the extracellular matrix the amino- and carboxyl-terminals undergo cleavage by procollagen peptidases (Colige et al., 1997). This allows the formation of the collagen molecule. Stabilisation occurs via lysine derived intramolecular and intermolecular cross-links (Zernicke & Loitz, 1992).
1.2.7 Proteoglycan Synthesis
Proteoglycan synthesis has been initially studied using aggrecan. Proteoglycan core protein synthesis follows the general pattern of all secreted proteins. The proteoglycan core protein is first of all synthesised on ribosomes that are associated with the cytoplasmic surface of the rough endoplasmic reticulum, this process is controlled by a signal peptide (Revel & Hay, 1963; Gallop et al., 1972). The core protein of the proteoglycan is translocated into the lumen of the rough endoplasmic reticulum where the polypeptide core is modified by the addition of N-asparagine-linked mannose-rich oligosaccharides from dolichol phosphate intermediates.
The rough endoplasmic reticulum also accommodates the complex folding of the three globular domains (G1, G2 and G3) which in turn form disulphide bonds and also the trimming of the nascent N-linked oligosaccharides which begins before the product exits the endoplasmic reticulum (Vertel, 1995; Luo et al., 2000). Subsequently the proteoglycan core enters the Golgi apparatus of the cell where the GAG chains and O-linked oligosaccharides are assembled onto the xylosylated core protein.
At the end of GAG polymerisation and sulphation, the entire proteoglycan molecule is taken to a secretatory vacuole at which point the molecule is secreted from the cell by exocytosis (Kimura et al., 1979) and transported to the cell membrane or transported to intracellular compartments. After secretion of each proteoglycan some extracellular processing occasionally takes place.
1.2.8 Hyaluronan Synthesis
The hyaluronan synthases were first characterized in the bacterium Streptococcus pyogenes (DeAngelis et al., 1993). Vertebrates have three hyaluronan synthase (HAS) genes, namely, HAS-1, HAS-2 and HAS-3. All three HAS enzymes have similar sequence and structural features (for review see; Bastow et al., 2008). They contain two amino-terminal and five carboxyl-terminal membrane spanning domains, as well as a central cytoplasmic region. Synthesis of hyaluronan occurs on the inside of the plasma membrane (Prehm, 1983; Heldermon et al., 2001) and is either retained within the cell or taken to the extracellular matrix (Schultz et al., 2007).
The enzyme hyaluronan synthase catalyses its synthesis using substrates uridine diphosphate -N-acetylglucosamine and uridine diphosphate-glucuronic acid with loss of the uridine diphosphate moiety after each addition to the hyaluronan chain, these are sequentially added to the reducing end of the hyaluronan chain (Prehm, 1983).
1.3 Variation in the Structural Composition of the Tendon Extracellular Matrix
The composition of tendons varies greatly along their length allowing tendon to meet specific mechanical requirements (Mehr et al., 2000). Differences in composition are attributed to the synthesis of specific extracellular matrix components to meet the compressive or tensional forces exerted. The composition of the distal region also called the enthesis of tendon where the tendon connects to bone has a structure similar to that of fibrocartilage (Waggett et al., 1998). This region is made up of morphologically distinct fibrocartilaginous tissue due to the compressive forces generated at this site (Vogel & Koob, 1989).
The distal region of tendon contains increased levels of aggrecan and high GAG content of approximately 5% GAG dry weight of tendon of which 65% is chondroitin sulphate in comparison to the tensile region which contains approximately 0.2% GAG dry weight of tendon of which 60% is dermatan sulphate (Merrilees & Flint, 1980; Koob & Vogel, 1987). Consequently, the presence of aggrecan in tendon greatly increases its capacity to absorb water and to withstand compression.
The tensional region of tendon consists of elongated cells surrounded by an arrangement of longitudinal bundles of mainly Type I collagen fibres (Evans & Barbenel, 1975). In total the proteoglycans make up less than 1% of this region with decorin being the predominant proteoglycan present with smaller amounts of biglycan and aggrecan. The large proteoglycans aggrecan and versican represent 10% of the total proteoglycan content of this region (Vogel & Heinegard, 1985; Vogel & Evanko, 1987).
The percentage of GAG varies with mechanical load (Culav et al., 1999). Furthermore, the proportions of different proteoglycans vary with mechanical load with the chondroitin sulphate:dermatan sulphate ratio being higher in tissues that undergo compression and lower in tissues that resist tension (Flint et al., 1980). Chondroitin sulphate is present in higher amounts in the tensional region of tendon whilst keratan sulphate is present in higher quantities in the compressed region of tendon. It has been demonstrated that tendons that are compressed in vitro or that are surgically transferred to a bony pulley, have developed into fibrocartilage with cartilage cells appearing in the tissue.
On the other hand tendons that were transferred from a bony pulley tended to lose their fibrocartilagenous characteristics as the cartilage cells that were present disappeared (Ploetz, 1938; Gillard, 1979; Malaviya et al., 1996). In other work it was observed that the total GAG content of the compressed region of tendon that had been surgically placed in the tensional region of tendon was rapidly reduced to the low levels commonly seen in this tendon region (Gillard et al., 1979). GAG levels were gradually replenished when the tendon was put back to its original position, but the recovery decreased the longer the tendon was left at its translocated site (Gillard et al., 1979).
1.4 Proteinases of the Tendon Extracellular Matrix
The degradation of the tendon extracellular matrix is a feature of normal tendon remodelling as well as in many pathological situations including overuse tendinopathy (for review see; Clark & Murphy, 1999). The endopeptidases are thought to be the key enzymes in extracellular matrix degradation and all four major classes (metallo-, serine, cysteine and apartate) can degrade individual components of the tendon extracellular matrix (for review see; Clark & Murphy, 1999).
The degradation of collagen is primarily mediated by matrix metalloproteinases (MMPs) activity and the degradation of aggrecan in tendon is mostly mediated by the aggrecanases. The following section will discuss the major proteinases of the tendon extracellular matrix including the various matrix metalloproteinases (MMP) and a group of proteinases that are termed A Disintergrin and Metalloproteinase with Thrombospondin Motifs (ADAMTS) as well as their inhibitors the tissue inhibitors of metalloproteinases (TIMPs). This section will also provide information on the catabolism of the major extracellular matrix macromolecules collagen, proteoglycans and hyaluronan.
1.4.1 Metalloproteinases (Metzincins)
Metzincins can be divided into four separate families including astacin, serratia, reprolysin and the matrixins (Hooper, 1994). The metzincins are calcium and zinc dependent endopeptidases that function at neutral pH. These families can be classified according to the residues following the third histidine and by the amino acid sequence surrounding the methionine in the Met-turn. The matrixins contain a serine residue following the third histidine and the reprolysins contain an aspartic acid residue following the third histidine (Hooper, 1994). This review, however will only discuss the matrixins or matrix metalloproteinases and the reprolysins.
1.4.2 Matrix Metalloproteinases (Matrixins)
In mammalian tissue there are currently 28 MMPs that have been identified, with 23 of these occurring in human tissue. MMPs are important in the normal metabolic functions of tendon but also have activity against cell-surface receptors and growth factor precursors (McCawely & Matrisian, 2001). MMPs are implicated in the normal and pathological turnover of the extracellular matrix of tendon. MMPs are referred to as matrixins (Birkedal-Hansen et al., 1993) as they contain zinc at the active site and a conserved methionine eight residues downstream.
They degrade different components of the extracellular matrix at neutral pH (Birkedal-Hansen et al., 1993). Evidence suggests that both the MMPs and tissue inhibitors of matrix metalloproteinases (TIMPs) are not involved to a large degree in intracellular lysosomal phagocytosis, but function extracellularly (Erlichman et al., 2001).
The MMPs belong to a larger family of proteins known as the metzincin superfamily. The gene expression of most matrixins is transcriptionally regulated by growth factors, hormones, cytokines and cellular transformation (Nagase, 1996; Fini et al., 1998). The proteolytic activities of MMPs are controlled during activation from their precursors and inhibition by endogenous inhibitors, α-macroglobulins, and TIMPs.
These enzymes also have an important role in the regulation of numerous cellular activities including cell proliferation, cell death (apoptosis), cell migration and chemotaxis (McCawley & Matrisian, 2001). MMPs have a common multi domain structure; a pre, pro- and catalytic domain followed by a variable hinge region and a hemopexin-like domain (Woessner, 1994) and the pre-domain is removed by secretion (Woessner, 1994).
These MMPs play important roles in embryonic development, morphogenesis, angiogenesis and tissue involution and in diseases associated with degradation of the extracellular matrix such as atherosclerosis, tissue ulceration, periodontal disease, fibrotic lung disease, osteoarthritis and in inflammatory disorders (Nagase, 1994; Woessner, 1994). Initially, the MMPs were described by Gross and Lapiere in 1962 who observed enzymatic activity (collagen triple helix degradation) during tadpole tail metamorphosis. This enzyme was then called interstitial collagenase MMP-1.
The MMPs are subdivided into four groups based on structure and substrate specifity: the collagenases, gelatinases, stromelysins and membrane-type matrix metalloproteinases (MT-MMP; Nagase, 1994; Cawston, 1995; Clark & Parker, 2003). The MMPs share a common domain structure; the three domains are the pro-peptide, the catalytic domain and the haemopexin like carboxyl-terminal domain which is linked to the catalytic domain by a flexible hinge region.
There are currently five MT-MMPs that have been identified (MT-MMP to MT5-MMP; MMP-14 to 17 and MMP-24) to date (Sato et al., 1994; Takino et al., 1995; Will & Hinzmann, 1995; Puente et al., 1996; Nagase, 1997; Bode et al., 1999; Llano et al., 1999). The MT-MMPs are membrane-bound metalloproteinases that contain a transmembrane domain and cytoplasmic tail at its carboxyl-terminal end, which anchors them to the cell surface (Woessner & Nagase, 2000).
The collagenases are a subgroup of matrixins that are capable of degrading fibrillar and non-fibrillar collagens. The collagenases include MMP-1 (Interstitial collagenase-1) MMP-8 (neutrophil collagenase) MMP-13 (collagenase-3). Collagenases specifically cleave the intact Type I collagen molecule in the extracellular environment and occurs at a specific locus in the triple helix between residues 775 and 776 (Cawston, 1995). They are also capable of degrading other extracellular matrix components including aggrecan and link protein (Clark & Murphy, 1999).
The main substrates of the gelatinases are non-fibrillar collagens and gelatine (Clark & Murphy, 1999) these enzymes are distinguished by the presence of an additional domain inserted in the catalytic domain. The gelatinases include gelatinase A (MMP-2) and gelatinase B (MMP-9); both of these are capable of actively degrading denatured collagens (gelatins). The gelatinases, like the collagenases, can also degrade Type I, IV, IX, X, XI and XIV collagen, aggrecan, link protein and fibronectin (Clark & Murphy, 1999).
The stromelysins display a broad ability to cleave extracellular matrix proteins but are unable to cleave the triple helical fibrillar collagens. The stromelysins that are present in connective tissues include stromelysin-1 (MMP-3), stromelysin-2 (MMP-10) and stromelysin-3 (MMP-11). Stromelysin-1 and -2 have the ability to degrade aggrecan (Bonasser et al., 1995), versican, perlecan, link protein, elastin, link protein and collagen Types III, IV, IX and X (Bejarano et al., 1988; Birkedal-Hansen et al., 1993; Clark & Murphy, 1999).
Several MMPs do not belong in any of the above mentioned groups as they have distinctive structural and/or functional properties. They include matrilysin (MMP-7), stromelysin-3 (MMP-11), metalloelastase (MMP-12), enamelysin (MMP-20), MMP-19 and MMP-23 (Velasco et al., 1999).
1.4.3 MMP Inhibition
The tissue inhibitors of metalloproteinases (TIMPs; 21-30 kDa) are a major inhibitor of MMPs in the tissue (Murphy et al., 1994a; Cawston, 1995; Murphy & Willenbrock, 1995; Gomez et al., 1997; Fassina et al., 2000; Cawston & Wilson, 2006). There are currently four types of TIMPs (1-4; Murphy et al., 1994a; Cawston, 1995; Murphy & Willenbrock, 1995; Nagase & Woessner, 1999; Bramono et al., 2004). They have six disulphide bonds and comprise a three-loop amino-terminal domain and an interacting three-loop C-subdomain. TIMPs inhibit active MMPs by forming tight-binding, non-covalent 1:1 enzyme-inhibitor complexes (Brew et al., 2000; Bramono et al., 2004).
Differences in the binding affinity of each type of TIMP for the various MMPs exists (Woessner, 1994; Brew et al., 2000). The balance between the activities of MMPs and TIMPs regulates tendon remodelling and an imbalance produces collagen disturbances in tendons (Dalton et al., 1995). TIMP-1 and TIMP-2 have the capacity to inhibit the activities of all MMPs in particular MMP-2 and MMP-9, respectively.
TIMPs and MMPs are often activated at the same time in response to physical activity (Koskinen et al., 2002), this could mean the simultaneous stimulation and inhibition of degradation. Another MMP inhibitor is α2-macroglobulin, a 772 kDa protein comprising four nearly identical, disulphide-bonded domains which is mainly present in the serum. Inhibition is effected by the presentation of a cleavable region that, once proteolytically cleaved, causes a conformational change that entraps the proteinase, which becomes covalently anchored by transacylation.
The reprolysin or adamlysin family consists of a large number of snake venom metalloproteinases and the structurally related ADAMs (A Disintergrin and Metalloproteinases). This family is made up of 34 members and all share the following common structural features, a pro-, metalloproteinase-like, disintergrin-like, cysteine-rich, EGF-like, transmembrane and cytoplasmic tail (Tang & Hong, 1999).
The ADAM family members are typically cell-surface molecules with additional transmembrane and cytoplasmic domains which span the plasma membrane (Tang & Hong, 1999). These proteins are also referred to as MDC (metalloproteinase/disintergrin/cysteine-rich), disintergrin-metalloproteinases or metalloproteinase-disintergrins (Tang & Hong, 1999).
1.4.5 A Disintergrin and Metalloproteinases with Thrombospondin Motifs (ADAMTS)
A disintergrin and metalloproteinases (ADAM) with thrombospondin (TS) motifs represents a zinc-dependent family possessing both metalloproteinase and disintergrin domains (Kuno et al., 1997; Tang & Hong, 1999; Kaushal & Shah, 2000). Kuno and colleagues first described the ADAMTS proteinases in mice in 1997 (Kuno et al., 1997).
These enzymes show a complex domain organization including signal sequence, propeptide, metalloproteinase domain, disintegrin-like domain, central TS-1 motif, cysteine-rich region, and a variable number of TS-like repeats at the carboxyl- terminal region (Kuno et al., 1997) Aggrecanases were first identified on the basis of their ability to cleave aggrecan at specific Glu-Xaa bonds, with a major site in the IGD of the aggrecan core protein (Glu373-Ala374; Sandy et al., 1991; Ilic et al., 1998; Ilic et al., 2000). This cleavage results in the loss of the GAG-rich portion of the molecule (Sandy et al., 1991; Ilic et al., 1998; Ilic et al., 2000).
There are currently 19 mammalian ADAMTS enzymes that have been identified, however many of these are yet to be fully characterised (Kaushal & Shah, 2000). ADAMTS-4 (aggrecanase-1) and ADAMTS-5 (aggrecanase-2) were the first aggrecanases to be identified (Abbaszade et al., 1999; Tortorella et al., 1999; Kaushal & Shah, 2000). ADAMTS-1 and ADAMTS-5 are best known for their activity against aggrecan but they are able to cleave other extracellular matrix proteoglycans such as versican and brevican (Sandy et al., 2001) and the non-collagenous protein COMP (Dickinson et al., 2003). ADAMTS-1 was initially identified as a novel murine cDNA expressed in a cachexigenic adenocarcinoma cell line that could be up regulated by IL-1 (Kuno et al., 1997).
ADAMTS-2, ADAMTS-3 and ADAMTS-14 are pro-collagen peptidases, and have the ability to function as key regulators of collagen fibril assembly (Colige et al., 1999; Fernandes et al., 2001; Colige et al., 2002). The first aggrecanases to be identified were ADAMTS-4 and ADAMTS-5 (Abbaszade et al., 1999; Tortorella et al., 1999; Kaushal & Shah, 2000). In a recent study involving tendon cells it was reported that there was a small and variable effects IL-1 on ADAMTS-4 gene expression (Tsuzaki et al., 2003). ADAMTS-2 cleaves the amino peptides of Types I, II and III procollagens (Colige et al., 1997; Wang et al., 2003).
ADAMTS-4 has recently been shown to cleave COMP as well as fibromodulin and decorin suggesting that this group of ADAMTSs may have a wider role than just proteoglycan cleavage (Kashiwagi et al., 2004). ADAMTS-13 cleaves the large multimeric von Willebrand factor precursor to generate von Willebrand factor of optimal size for proper coagulation (Fujikawa et al., 2001; Soejima et al., 2001; Zheng et al., 2001). A function for the following ADAMTS proteins is yet to be assigned, ADAMTS-6, -7, -10, -12, -16, -17, -18 and -19.
1.4.6 Collagen Catabolism
The predominant molecule in tendon, Type I collagen is mostly resistant to proteinase degradation due to its strong triple helical structure (Verzijl et al., 2000). However, the collagenases have been shown to have an ability to cleave Type I collagen molecules (Matrisian, 1992; Cawston, 1995; Nagase & Woessner, 1999). Collagenase cleavage occurs at specific sites in the collagen triple helix at residues (Cawston, 1995). The proteinases that are involved in collagen catabolism include MMP-1, MMP-2, MMP-8, MMP-13 and MMP-14 (Billinghurst et al., 1997; Poole et al., 2003). These have been shown to cleave the α1(II) chain of Type II collagen within the triple helical region between the Glu775-Leu776 (Billinghurst et al., 1997; Ohuchi et al., 1997). Cleavage in this region causes two helical fragments representing three-quarter and quarter length fragments (Gadher et al., 1988).
Previous studies have indicated that when using explant cultures of articular and nasal cartilage (Kozaci et al., 1997; Price et al., 1999; van Meurs et al., 1999; Billinghurst et al., 2000) as well as animal models of arthritis (van Meurs et al., 1999) the degradation of the collagen network occurs after the majority of aggrecan molecules are lost from the tissue, increasing the susceptibility to proteolytic attack by various proteinases. Some collagen in tendon is most likely degraded intracellularly after phagocytosis, with fibroblasts and macrophages engulfing the collagen, which is finally broken down by lysosomal enzymes (Everts et al., 1996; Creemers et al., 1998).
There is little knowledge about the catabolism of the non-fibrillar collagens. MMP-3 has the ability to cleave Type IX collagen within the NC-2 domain through its α1(IX), α2(IX) and α3(IX) chains (Wu et al., 1991) and Type X collagen is cleaved by several collagenases as well as gelatanase A (Gadher et al., 1988). Human neutrophil elastase can degrade Type IX, X and XI collagen (Gadher et al., 1988).
1.4.7 Proteoglycan Catabolism
It has been shown then proteoglycans are turned over much more rapidly than the fibrillar collagens. In terms of the catabolism of the small proteoglycans, in particular decorin and biglycan catabolism in happen in two ways; 1) degradation in the extracellular matrix, and 2) intracellular degradation. Studies in ligament and tendon explant cultures showed that 50% of the newly synthesised decorin is lost from the tissue unchanged, while the remainder is taken up and internalised by fibroblasts (Campbell et al., 1996; Winter et al., 2000). ADAMTS-4 has been shown to have activity against brevican, a proteoglycan that is present in nervous tissue (Nakamura et al., 2000).
The degradation of biglycan is less extensive when compared to decorin in fibrous connective tissues (Samiric, 2003; Ilic et al., 2005). Degradation of the core protein of biglycan has been shown in medium from the compressive regions of tendon (Rees et al., 2000). The MMPs and ADAMTSs have been shown to degrade decorin in vitro (Imai et al., 1997; Engle et al., 2004). MMP-2, -3 and -7 cleave decorin in the LRR region. In one study decorin underwent cleavage in the LRR region between Ser210-Leu211 by MMP-2 and -3 and also at Glu243-Leu244 by MMP-7. Furthermore, decorin is also cleaved at Glu154-Leu155 by the MMPs –2, -3, -9 and -14 as well as ADAMTS-4 and -5 (Engle et al., 2004).
Aggrecan catabolism has been extensively studied, particularly in cartilage but also tendon. The aggrecan core protein is particularly vulnerable to cleavage by the proteinases (Arner, 2002). The G1 domain does show some resilience to proteinases. Cleavage of the aggrecan core protein occurs between GAG chains (Campbell et al., 1989; Sandy et al., 1991; Ilic et al., 1992; Samiric, 2003) resulting in aggrecan fragments that do not contain the hyaluronan-binding domain (Sandy et al., 1992). The IGD of aggrecan is vulnerable to cleavage and two sites have been identified between the residues Asn341-Phe342 and Glu373-Ala374 (Hering et al., 1997).
The catabolism of versican is less studied than that of aggrecan, but studies have shown most of the versican fragments present in the tendon matrix do not contain the G1 domain (Samiric, 2003). Versican has been shown to undergo in vivo cleavage at the Glu416-Ala417 site by ADAMTS-1 and -4 (Sandy et al., 2001). Furthermore, cleavage sites between residues Asp1329-Ser1330, Glu2187-Ser2188 and Glu2628-Ser2629 have been found in the GAG-β region of versican. Aggrecanses also cleave versican V1 between Glu441-Ala442 (Sandy et al., 2001) and versican V2 between Glu405-Gln406 (Westling et al., 2004).
1.4.8 Hyaluronan Catabolism
It has been suggested that one third (~ 5 g) of the hyaluronan in the human body turned over every 24 hrs (Laurent & Reed, 1991). The catabolism of hyaluronan varies greatly between different tissues. In the blood its half-life is 2-5 mins (Fraser et al., 1981), in the skin turnover is 1-2 days in the epidermal compartment and in cartilage hyaluronan turnover is 1-3 weeks. The hyaluronan cell surface receptor CD44 has been implicated in the internalisation and catabolism of hyaluronan (Hascall et al., 1999; Knudson et al., 1999). Studies have shown that hyaluronan is catabolised by oxygen free radicals and hyaluronidase in vivo (Schiller et al., 2003; Csoka et al., 2001; Lepperdinger et al., 2001; Adams et al., 2006; Atmuri et al., 2008).
The majority of the hyaluronan produced by the body is removed from the circulatory system via endocytosis in the liver and the lymph nodes, this occurs via the hyaluronan receptor for endocytosis (Weigel & Weigel, 2003). The hyaluronan receptors include CD44, lymphatic vessel endothelial hyaluronan receptor, receptor for hyaluronan-mediated motility, layilin and hyaluronan receptor for endocytosis. In endocytosis these receptors play a local tissue specific role. The family of human hyaluronidase genes include hyaluronidase-1, hyaluronidase-2, hyaluronidase-3, hyaluronidase-4, hyaluronidase P1 and PH-20 with little known about each of these. The hyaluronidases have the ability to catalyse the hydrolysis of hyaluronan (Bastow et al., 2008).
1.5 Overuse Tendinopathy
A healthy tendon is essential for the movements made by all individuals in day to day life from the most active to the sedentary population. When a healthy tendon becomes degenerative people’s lives may be drastically changed. Overuse tendinopathy is a chronic and painful condition that affects the entire population from the most active of individuals to those who are less active (Riley, 2000). This pathological condition produces a significant level of morbidity and the problems that prevail may last for long periods of time in spite of what is considered the appropriate management (Almekinders & Almekinders, 1994).
Very little information about the underlying causes of overuse tendinopathy is available due to a lack of research and therefore the problems are difficult to manage (Almekinders & Temple, 1998). Metabolic change of the tendon is believed to result in degeneration of the tendon matrix most likely caused by proteinases (Riley, 2000). There are several tendons that are more commonly affected than others as these tendons are exposed to high mechanical loads; these include tendons of the shoulder (supraspinatus), elbow (extensor carpi radialis brevis), foot (posterior tibialis), knee (patellar) and ankle (Achilles; Kvist et al., 1985; Kvist et al., 1987; Kvist et al., 1988; Leadbetter, 1992; Kvist, 1994; Myerson & McGarvey, 1998; Kraushaar & Nirschl, 1999; Jonstone, 2000; King et al., 2000). The area that is most commonly affected in overuse tendinopathy is at or proximal to the insertion in the fibrocartilaginous region of the tendon, with the exception of the Achilles tendon (Cook & Khan, 2007).
These tendons/areas have several common features including they are more highly stressed than other tendons, often exposed to repeated strains, including shear or compressive forces with little blood supply relative to other tendon/areas (Riley, 2004). Overuse tendinopathy affecting the enthesis of the patellar tendon have been well documented in a variety of sports. Both ends of the tendon are susceptible, but the most common site of pathology is near the posteromedial portion of the patellar (Yu et al., 1995; Khan et al., 1999). Excessive and repetitive loading of tendons during high intensity physical activity is possibly the main cause for tendon degeneration (Selvanetti et al., 1997).
From a traditional sense the term tendinitis has been used to describe a pathological tendon and this assumes the presence of inflammation (Riley, 2000). However, the pathogenesis of tendinopathy does not appear to involve inflammatory processes (Alfredson & Lorentzon, 2000) but is more likely to involve a failed/slow healing response to overuse injury where the healing process is impeded by the recurring microtrauma and thus results in a degenerate tendon with compromised functional properties (Cook et al., 2002; Riley, 2004). It is therefore of very little surprise that clinical trials using anti-inflammatory drugs have shown very little benefit in overuse tendinopathy (Almekinders & Temple, 1998).
There is little evidence that any conventional treatments (anti-inflammatory drugs) work in overuse tendinopathy (McLauchlan & Handoll, 2001). However, most of the histopathological evidence has come from tendon samples that are in the final stages of overuse tendinopathy rather than at different stages of the disease and this gives the possibility that inflammation may be involved at the beginning or other stages of the disease (Rees et al., 2006). The term tendinosis can be used to describe a degenerative condition with no inflammation (Puddu et al., 1976).
However, it has been shown that in pathological tendon some inflammatory mediators are increased around the tendon (Fu et al., 2002b), indicating the involvement of inflammatory mediators, therefore inflammation cannot be entirely ruled out at some stage of the disease. It has been shown that pathological tendons of the patellar and Achilles exhibit increased gene expression levels of glutamate (Alfredson et al., 1999; Alfredson et al., 2001) which is known to be involved in pain. Furthermore, immunohistochemical analysis of tendon biopsies revealed the presence of ionotrophic glutamate receptor N-methyl-D-aspartate in relation to nerves therefore glutamate may be the cause of pain in overuse tendinopathy (Alfredson et al., 2001). The term tendinopathy is more commonly used today to describe all forms of tendon pathology, as this term does not assume any knowledge of the underlying pathology (Riley, 2000).
1.5.1 Predisposing Factors
Two hypotheses have been put forward to account for overuse tendinopathy, they include; 1) a vascular theory, and 2) a mechanical theory (for review see; Kjaer, 2001). More recently a neural theory has come into circulation, however, at the present time more evidence is required to develop a neural theory for overuse tendinopathy (for review see; Rees et al., 2006). Tendons are metabolically active and need a blood supply to function (Rees et al., 2006), it is thought that a loss or decrease of this blood supply may result in tendon degeneration that is observed in overuse tendinopathy (Rees et al., 2006). It has been proposed that certain tendons are more prone to losing this blood supply than others (Fenwick et al., 2002) including the supraspinatus (Ling et al., 1990), the Achilles (Ahmed et al., 1998) and the tibialis posterior (Frey et al., 1990).
Lesions of the Achilles and supraspinatus tendons generally have an overall reduction in blood supply (Brooks et al., 1992; Leadbetter, 1992; Kannus, 1997b; Ahmed et al., 1998; Fenwick et al., 2002). Several factors may contribute to the decline in blood supply such as aging, vascular disease, trauma, physical disuse or trauma and eventual tissue hypoxia and reduced viability of the tendon cells (Riley, 2004). The major issue with the vascular theory is that in overuse tendinopathy studies have shown an increase in vascularity and cellularity (Astrom & Rausing, 1995) as well as an increase in the blood supply of the tendon (Astrom & Westlin, 1994).
The second theory, the mechanical theory of overuse tendinopathy argues that repetitive loading of the tendon may lead to tendon rupture. Mechanically, tendon lesions are considered to be caused by repetitive microtrauma often called overuse pathologies (Herring & Nilson, 1987; Kannus, 1997a,b; Selvanetti et al., 1997; Riley, 2004). This is due to the biomechanical properties of the tendon and can be explained via the stress-strain curve (for review see; Kjaer, 2001). Under normal circumstances tendon length is increased by no more than 4% of its length (Kvist, 1991). When the tendons length is increased more than this, damage to the tendon may eventuate and when tendon length increases beyond 8-12% the tendon may rupture (Kvist, 1991).
Under normal conditions described above and in particular towards the 4% range degeneration within the tendon may start to occur, especially with repeated and/or prolonged use. This may then lead to a tendon that undergoes changes in its mechanical properties as a result of these repeated microtraumas (Curwin, 1998; Mosler et al., 1985; Wren et al., 2003). This theory goes a long way to explain how repetitive microtraumas to the tendon could accumulate over time and why tendinopathy would be a degenerative condition rather than inflammatory (Rees et al., 2006). Increased overuse tendinopathy with age and in the active individuals is consistent with this theory (Rees et al., 2006).
However, the mechanical theory does not fully explain why certain areas of certain tendons are at increased risk of degenerative change and it does not explain the pain sometimes linked to oversuse tendinopathy (Rees et al., 2006). It is more than likely that the cause of overuse tendinopathy is the result of more than one cause and may be the result of a combination of the two theories described above (Rees et al., 2006) and there is some evidence to suggest that the degenerative process varies among different tendon sites (Kannus & Jozsa, 1991).
Overuse tendinopathy can be acute or chronic and may be associated with a variety of contributing factors (for review see, Riley, 2000). Some of the factors (for reviews see; Riley, 2000; Kjaer, 2004) that may be associated with overuse tendinopathy include age, gender, height, vascularity, genetics (Bauer, 1980; Singer & Jones, 1986), blood type (Jozsa et al., 1989; Kujala et al., 1992), accompanying presence of chronic disease either inherited such as Marfans or Ehlers–Danlos syndromes or acquired such as rheumatoid arthritis or diabetes mellitus (Kannus & Jozsa, 1991) and drug use (Ribard et al., 1992; Huston, 1994).
In addition to the above, high body weight, leg length inequality, foot abnormalities (such as pes cavus or planus), and low joint, tendon, or muscle flexibility are possible contributors to developing overuse tendinopathy (Kannus, 1997a,b). Other factors involved in overuse tendinopathy may include occupation, sport, physical load (excessive force, repetitive loading, high intensity, changes in training pattern, previous injuries, fatigue, poor technique and abnormal or unusual movements), shoes and equipment and environmental conditions such as temperature and the nature of the running surface (Kvist, 1991; Selvanetti et al., 1997; Riley, 2004).
Indeed, in a study by Ferretti et al. (1984) it was shown that there was a strong relationship between the amount of training and the frequency of overuse tendinopathy among volleyball players, and that the harder the floor type on which the volleyball players trained the higher the frequency of overuse tendinopathy.
As tendons age they become more prone to overuse tendinopathy due to a decrease in their biochemical properties. With ageing the collagen, proteoglycan, non-collagenous protein and water content all appear to decrease (Ippolito et al., 1975; Birch et al., 1999). In addition, collagen fibre area and strength decrease making tendons smaller and more prone to rupture. As connective tissues such as tendon age the non-specific cross linking that occurs from the condensation of a reducing sugar with an amino acid group causes an accumulation in advanced glycation end products (Sell & Monnier, 1990; James et al., 1991; Reddy et al., 2002), and this causes the tendon to become more rigid (Verzijl et al., 2000).
In has been shown in animals that the average diameter of fibrils decreases with age (Nakagawa et al., 1994) and disuse (Nakagawa et al., 1989). Overuse tendinopathy is prevalent in all age groups but is most common in those of later middle age (Kannus, 1997a). Tendon rupture not only occurs during high intensity physical activities such as basketball but also is also common in simple activities such as walking or climbing a flight of stairs (Fahlstrom et al., 2002).
The process of tendon healing is both long and even after the fact they do not always return to their original state (Sharma & Maffulli, 2006). Because of the lack of knowledge into tendon biology an effective treatment protocol has not been developed and appears to be a distant solution (Sharma & Maffulli, 2006).
1.5.2 Changes in Collagen Content in Overuse Tendinopathy and Other Pathological Conditions
The evidence of microscopic changes in fibrillar collagens in pathological tendons (Khan et al., 1996a; Riley, 2004) has been corroborated by the biochemical analysis of pathological tendons which showed changes in collagen composition that involved the increase in the ratio of Type III collagen to Type I collagen, increase in the proportion of degraded collagen and increased number of crosslinks in pathological tendon (Riley et al., 1994b; Bank et al., 1999). The limited number of reports on the changes of total collagen content varies between pathologies of different tendons (Riley et al., 1994b; Bank et al., 1999; de Mos et al., 2007).
In a study of the Achilles tendon, cultures from ruptured and tendinopathic tendons revealed an increased production of Type III collagen. It has been shown that ruptured Achilles tendons contain a significantly greater proportion of Type III collagen, which predisposes them to rupture (Maffulli et al., 2000b). The excessive stress that is placed on particular tendons could potentially lead to areas of microtrauma within the tendon (Maffulli et al., 2000b). These areas may heal by the production of Type III collagen, which is an abnormal healing response (Maffulli et al., 2000b).
Many events of microtrauma could result in a critical point where the resistance of the tissue to tensile forces is compromised and tendon rupture results (Maffulli et al., 2000b). It has been speculated by Kannus in 1997a that pathological tendon may begin with horizontal collagen tearing due to mechanical loading, whereas others have suggested that collagen initially separates longitudinally before it tears transversely in chronic tendinosis, this however is still yet to be determined as both situations are present in chronic tendinosis (Cook et al., 2004). This may also indicate that a tear in the collagen network be it transverse or longitudinal is not the beginning of the overuse tendinopathy (Cook et al., 2004).
In a study by Chipman and colleagues in 1993 animals with mutations of the Type I collagen genes developed severe osteogenesis imperfecta a condition that results in weak bones particularly when exposed to mechanical loading. Type III collagen was accumulated at the rupture site probably due to microtraumas and the subsequent healing process (Eriksen et al., 2002). The increased content of Type III collagen can cause thinner collagen fibres, decrease the tensile strength and may finally result in the tendon rupture (Eriksen et al., 2002).
The age-related change in the nature of the cross-link in the carboxyl-terminal telopeptide may contribute to this weakening (Eriksen et al., 2002). Deficiencies in Type V collagen results in symptoms of joint laxity and poor wound healing (Schwarze et al., 2000). In tendons from rabbit, Type V collagen has been linked to high amounts of thin fibrils, suggesting inhibition of fibril assembly (Dressler et al., 2002).
Mutations collagen Type VI have been shown to lead to myopathies such as Bethlem myopathy a dominantly inherited disorder characterized by progressive muscle weakness and wasting suggesting collagen Type VI has a role in tissue integrity (Jobsis et al., 1996). In a recent study, a temporal pattern of changes, from cellular abnormalities, to increased ground substance, longitudinal collagen separation and finally neovascularisation was suggested to proceed to tendinopathy (Cook et al., 2004).
1.5.3 Changes in Proteoglycan and GAG Content in Overuse Tendinopathy and Other Pathological Conditions
Histological studies have promoted our understanding of proteoglycan metabolism in normal and pathological tendons. These studies showed that there is an increase in the levels of GAG in pathological tendons (Chard et al., 1994; Fu et al., 2007). This has yet to be confirmed by the biochemical analysis of GAG levels, proteoglycan gene expression and indeed proteoglycan protein levels (Chard et al., 1994; Riley et al., 1994a; Corps et al., 2004; Corps et al., 2006; Fu et al., 2007; Scott et al., 2007). So far only one study has shown an increase in levels of a proteoglycan, versican, in human pathological tendons (Scott et al., 2007).
In an attempt to fully understand the function of the SLRPs, knockout mice were developed that lack decorin, biglycan, fibromodulin or lumican. In biglycan deficient mice collagen fibrils had smaller diameter and abnormal morphology resulting in weaker tendons with decreased stiffness in the patellar tendon and tails (Ameye et al., 2002a,b). These mice also developed ectopic tendon and joint ossification and at three months of age attained osteoarthritis which could develop from joint instability and mechanically compromised tendons (Ameye et al., 2002a,b).
In a study by Corsi and colleagues 2002, biglycan deficiency led to structural abnormalities in collagen fibrils in bone, dermis and tendon. Indeed, another study of biglycan deficient mice found mice with growth failure, reduced bone formation, and age-related severe osteopenia (Xu et al., 1998). Deficiency of biglycan results in morphological changes in collagen fibrils from many tissues with tendon no exception (Ameye et al., 2002a; Corsi et al., 2002; Goldberg et al., 2003).
Fibromodulin knockout mice have abnormal collagen fibrils, tissue organization and reduced tendon stiffness. In the absence of fibromodulin there is an increase in lumican deposition (Svensson et al., 1999; Ezura et al., 2000; Jepson et al., 2002). It has been shown that in the absence of fibromodulin, that lumican binds to the same sites that fibromodulin does demonstrating that these two small proteoglycans compete for the same binding sites on collagen fibrils (Svensson et al., 2000).
Likewise to mice deficient in biglycan, fibromodulin deficient mice developed ectopic tendon and joint ossification and at three months of age attained osteoarthritis (Ameye et al., 2002). Indeed, from an ultrastructural viewpoint mice have irregular collagen fibril in cross section (Svensson et al., 1999; Ameye et al., 2002) and the average fibril diameter is decreased in patellar and Achilles tendons which could be due to the higher number of small collagen fibrils in the fibril population (Svensson et al., 1999; Ameye et al., 2002).
The importance of the SLRPs were first observed in decorin knockout mice where their role in collagen fibrillogenesis was demonstrated (Danielson et al., 1997). In decorin-knockout mice it was observed that mice exhibited reduced tensile strength of the skin and tendons and had irregularities in the collagen fibril diameter with fibrils being coarse, irregular and randomly arranged as was observed in biglycan knockout mice (Danielson et al., 1997). It was also observed that fewer proteoglycans were collagen bound in the extracellular matrix of skin and tendon (Danielson et al., 1997). In a study of the periodontal ligament collagen fibrils were randomly orientated rather than their normal parallel conformation (Hakkinen et al., 2000).
As mentioned earlier lumican is a major constituent of the cornea. Studies on knockout mice have indicated that lumican is able to substitute functionally for fibromodulin in fibromodulin deficient mice (Jepsen et al., 2002). Whereas, in lumican knockout mice large diameter collagen fibrils forming disorganised matrices were observed in the cornea and skin (Chakravarti et al., 2000). In the cornea of lumican deficient mice thicker, irregular and loosely packed collagen fibrils associated with a dramatic disruption in the organisation of the collagen fibrils occur (Chakravarti et al., 1998; Quantock et al., 2001).
1.5.4 Changes in the Non-Collagenous Proteins in Overuse Tendinopathy and Other Pathological Conditions
Very few studies have investigated the non-collagenous proteins in normal and pathological tendons. Studies using cDNA arrays have shown that some of the non-collagenous proteins are up regulated in Achilles overuse tendinopathy (Alfredson et al., 2003). Tenascin-C is up regulated after injury and is thought to play a role in modulating cell migration and activity (Mackie et al., 1988; Lehto et al., 1990; Amiel et al., 1991). Tenascin-C was increased in ruptured supraspinatus tendons with differences in the expression of isoforms and small peptide fragments, this was attributed to enzyme cleavage (Riley et al., 1996a).
Fibronectin is present in normal tendons in relatively small quantities but following injury it is increased (Jozsa et al., 1989; Lehto et al., 1990; Amiel et al., 1991). In a study using immunostaining, fibronectin was found to be significantly increased in ruptured tendons consistent with wound a healing process (Tillander et al., 2002).
When COMP is lacking as in such genetic diseases as pseudoachondroplasia, individuals have a short stature, lax joints and early onset osteoarthritis (Briggs et al., 1995). In another study by Hecht and colleagues 1998 they found that in mice lacking COMP there were no obvious musculotendinous abnormalities, however in humans lacking COMP, multiple epiphyseal dysphasias are seen which is characterised by short statue and cartilage abnormalities. COMP-deficient mice show no abnormalities and no change in skeletal development (Svensson et al., 2002), this could mean that COMP has a role in the pathological condition because protein gene expression and turnover are increased in such diseases has osteoarthritis (Lorenzo et al., 2004).
1.5.5 Changes in MMPs and TIMPs in Overuse Tendinopathy and Other Pathological Conditions
Several MMPs that have been implicated in overuse tendinopathy with changes in the activities and gene expression of various MMPs and TIMPs (Ireland et al., 2001). In a study of the synovial fluid of individuals with rotator cuff tears the gene expression of MMP-1 and MMP-3 were both significantly increased, with no change in the gene expression of TIMP-1 (Yoshihara et al., 2001). It was also observed that the amount of enzyme activity increased with the size of the tear (Yoshihara et al., 2001).
In a study of ruptured supraspinatus tendons there was increased activity of MMP-1, as well as reduced activity of MMP-2 and MMP-3 with evidence of increased collagen turnover (Riley et al., 2002). Tears of the rotator cuff showed no significant increases in MMP-1 mRNA gene expression (Lo et al., 2004), although the actual activity of MMP-1 may be up regulated, with down regulation of MMP-2 and MMP-3 activity (Riley et al., 2002). The gene expression of MMP-3, TIMP-2, TIMP-3 and TIMP-4 mRNA is decreased in torn rotator cuff tendons (Lo et al., 2004), MMP-3 may therefore play a role in the normal maintenance and remodelling of the rotator cuff tendon, and a decrease in normal MMP-3 activity may represent a failure of normal matrix remodelling and maintenance (Riley et al., 2002). Also, MMP-13 is up regulated at the mRNA and protein level in patients with complete tears of rotator cuff tendons (Lo et al., 2004).
The gene expression of MMP-3 and TIMP-1, TIMP-2, TIMP-3 and TIMP-4 were shown to be down regulated in tendinopathic tendon (Ireland et al., 2001; Alfredson et al., 2003). TIMP-1 was not present in normal tendons, but, after acute tears of the supraspinatus tendon, it was expressed in the tendon edges for a period of two weeks (Choi et al., 2002). Six weeks after the tear, there was no gene expression of TIMP-1, indicating that TIMP-1 may inhibit excessive degradation of extracellular matrix by MMP-2 (Choi et al., 2002).
In a recent study of normal Achilles tendons by Jones and colleagues in 2006 the gene expression of almost all the 23 MMPs that are found in human genes were detected in normal and pathological tendons at variable degrees (Jones et al., 2006). Furthermore, in painful tendinopathy the mRNA of MMP-11, -16 and -24 were up regulated. In ruptured tendons of the same study MMP-1, -9, -11, -14, -17, -19 and -25 were all up regulated. In contrast, the mRNA of MMP-3, -10, -12 and -27 was down regulated in painful tendons and MMP-3, -7, -24 and -28 were down regulated in ruptured tendons. MMP-3 has been proposed as a central regulator of MMP activation (van Meurs et al., 1999) and its down regulation may serve to limit MMP activation within the tissue.
The level of MMP-10 mRNA, which is similar to MMP-3, was also lower in the painful tendon samples by a similar magnitude. The greatest of the observed differences was the 1,000-fold higher level of MMP-1 mRNA in the ruptured tendons, which suggests that there is a high level of collagen degradation occurring in tendons that have ruptured. TIMP-3 is believed to be the primary endogenous inhibitor of the aggrecanase ADAMTS proteinases (Kashiwagi et al., 2001; Hashimoto et al., 2001) and the ADAM-12 and -17 proteinases (Amour et al., 1998; Amour et al., 2000) and a decrease in TIMP-3 might therefore be predicted to influence the activity of these proteinases.
1.5.6 Changes in ADAM and ADAMTS in Overuse Tendinopathy
Few studies have investigated the gene expression of the ADAM and ADAMTSs in normal and pathological tendons. In one study of Achilles tendons by Jones and colleagues in 2006 the gene expression of almost all the 19 ADAMTS and several ADAM genes were detected in normal tendon and pathological tendons at variable degrees (Jones et al., 2006). Furthermore, in painful tendinopathy ADAMTS-2 and -3 were up regulated as was ADAM-12. In ruptured tendon ADAMTS-4 was up regulated as was ADAM-8 and -12. On the other hand ADAMTS-5 was down regulated in painful tendinopathy and ADAM-7 and -13 were also down regulated in ruptured tendons. None of the ADAM family members were down regulated in painful or ruptured tendons (Jones et al., 2006).
The levels of ADAM-12 mRNA were an increased in painful tendon samples than in the normal tendons (Jones et al., 2006). ADAM-12 has been demonstrated to cleave insulin-like growth factor binding proteins 3 and 5, pro-heparin-binding EGF and the extracellular matrix components gelatin, Type IV collagen, and fibronectin and may therefore be significant in painful tendinopathy, both as a regulator of cytokine activity and as a mediator of extracellular matrix degradation (Jones et al., 2006). In a recent study ADAMTS-4 was shown to be significantly up regulated in ruptured Achilles tendons compared to normal and chronic painful tendons, and in tendon extracts that were probed by Western blotting mature ADAMTS-4 was only seen in ruptured tendons while processed ADAMTS-4 was seen in chronic tendinopathy and normal tendons (Corps et al., 2008).
1.5.7 Other Changes in Overuse Tendinopathy
A change in the histological appearance of the extracellular matrix is characteristic of overuse tendinopathy (Khan et al., 1996a). This work suggests that the original tendon matrix undergoes destruction and there is generation of a different and unique tendon matrix. Relatively few biochemical studies of chronic tendinopathy have been undertaken; the few that have been performed have used material collected after the tendon has ruptured (Riley, 2000). Previous studies of ruptured Achilles tendons have shown changes in cellularity and have shown that the tendon cells have a more rounded appearance when compared to normal tendon (Khan et al., 1996b). In another study, histopathological analysis of painful tendons showed a change in cellularity, matrix organisation and increased infiltration of blood vessels (Astrom & Rausing, 1995) and it was proposed that the changes in vascularity may be associated with pain of the tendon (Ohberg et al., 2001).
A change in the turnover of the extracellular matrix has been implicated in degenerative conditions such as osteoarthritis and overuse tendinopathy (Clark & Parker, 2003), suggesting that changes in matrix metabolism may be targets for future drug therapies (Riley, 2000). Various types of degeneration have been observed in tendon with mucoid or lipoid degeneration usually found in the Achilles tendon (for review see; Jozsa & Kannus, 1997). Light microscopy studies of the tendon shows mucoid degeneration with large mucoid patches and vacuoles between fibres. In lipoid degeneration, abnormal intratendinous accumulation of lipid occurs with disorder to the structure of collagen fibres (for review see; Jozsa & Kannus, 1997).
In patellar tendinopathy, mucoid degeneration is commonly seen, although hyaline degeneration rarely occurs (Cook et al., 1997). In rotator cuff tendinopathy, mucoid degeneration is present, with fibrocartilaginous metaplasia, often accompanied by calcium deposition also common (Fukuda et al., 1990). Amyloid deposition in supraspinatus tendons with degenerative tears has also been reported (Cole et al., 2001). Macroscopically, the affected areas of tendon lose their normal brilliant white appearance and become gray-brown and shapeless. It has been proposed that the first signs of morphological change in tendinosis occur in the tenocytes and not the collagen fibres (Cook et al., 2004; Yuan et al., 2002). Previous studies have shown that tendon cells undergo apoptosis in response to hypoxia, oxidative stress or high tensile load (Scott & Duronio, 2003; Scott et al., 2005).
In a study of tendon pathology of the Achilles tendon, biopsies from 40 patients were taken and analysed (14 specimens obtained at autopsy served as controls), it was found that in pathological tendons there was abnormal collagen fibre structure and arrangement, focal variations in cellularity, rounded nuclei, decreased collagen stainability and increased non-collagenous proteins in the extracellular matrix were seen in all pathological biopsy specimens (Movin et al., 1997).
There were some histopathological changes in half of the controls. Increased vascularity was present in two thirds of the biopsy specimens and in one third of the control specimens (Movin et al., 1997). The volume density of GAG-rich areas was higher in the patients 0.47 (0-0.86) than in the controls 0 (0-0.07). In Achilles tendons with achillodynia and ultrasonographic widening there were differences in fibre structure and arrangement as well as increased amounts of GAG (Movin et al., 1997). It was suggested that achillodynia is due to local changes in connective tissue metabolism or circulation or a combination of the two (Movin et al., 1997).
In a study of patients undergoing treatment for subcutaneous rupture of their Achilles tendons as well as normal Achilles tendon biopsies were taken and analysed for fibre structure and arrangement, rounding of the nuclei, regional variations in cellularity, increased vascularity, decreased collagen stainability, hyalinization and GAG. This study showed that ruptured tendons had a significantly higher degree of degeneration than that of the normal tendons (Maffulli et al., 2000a,b).
Histological studies of overuse tendinopathy shows a variety of changes including disordered haphazard healing with absence of inflammatory cells, poor healing response, non-inflammatory intratendinous collagen degeneration, fibre disorientation and thinning, increased cellularity and scattered vascular in growth and increased GAGs (Leadbetter, 1992; Khan & Maffulli, 1998). In a study by Lian et al. in 2007 the role of apoptosis in the pathophysiology of patellar tendinopathy was investigated. They found that the number of apoptotic cells per unit area (4.5 mm2) was 0.91 ± 0.81 (SD) in tendinopathic samples and 0.21 ± 0.21 in controls (P = 0.026), therefore implicating increased apoptotic cell death as a feature of overuse tendinopathy.
Tendons are relatively inactive and have low cellularity and therefore healing is a slow and arduous process. Tendon healing occurs in three overlapping phases. The first phase of tendon healing is called the inflammatory phase where inflammatory cells such as neutrophils enter the injured area. Vasoactive and chemoatractants are released with an increase in the vascular permeability, angiogenesis begins, stimulation of fibroblast proliferation and the release of more inflammatory cells occurs (Murphy et al., 1994b).
Fibroblasts navigate towards the injured area and Type III collagen synthesis begins (Oakes, 2003). The second phase of tendon healing is the remodelling phase which begins just a few days after the initial tendon injury. The synthesis of Type III collagen peaks and lasts for several weeks with water and GAG levels increasing at this time (Oakes, 2003).
Approximately six weeks after the initial tendon injury the final phase of tendon healing begins, this is the modelling phase. During this phase, there is a decrease in cellularity, collagen and GAG synthesis. The modelling phase can be further divided into two phases the consolidation phase and the maturation phase (Tillman & Chasan, 1996). The consolidation phase commences at about six weeks and continues for up to ten weeks. During this time the repaired tissue changes from cellular to fibrous.
Fibroblast metabolism remains high during this period and fibroblasts and collagen fibres become aligned in the direction of stress (Hooley & Cohen, 1979). A higher proportion of Type I collagen is synthesized during this time (Abrahamsson, 1991). After ten weeks the maturation stage occurs with a gradual change of fibrous tissue to scar like tendon tissue over the course of one year (Hooley & Cohen, 1979). During the latter half of this stage fibroblast metabolism and tendon vascularity decline (Amiel et al., 1983). Despite the tendon healing process tendons rarely return to their natural state.
1.7 Aims of this Study
The overview of overuse tendinopathy indicates that a major outcome is a change in the extracellular matrix of tendon as discussed above. This is likely to result from a change in the metabolism of extracellular matrix macromolecules. This could occur at the level of gene expression or synthesis and/or catabolism of individual macromolecules. The aims of the present study were to determine in normal and pathological human patellar tendons the levels and types of collagens and proteoglycans. The gene expression of the proteoglycans, collagens, non-collagenous proteins and various proteinases and their inhibitors were also evaluated. Changes in the metabolism especially rates of synthesis and catabolism of proteoglycans were also determined.