Extracellular Matrix and Its Artificial Substitutes: Comparison
View Latest Version
Please note this is a comparison between Version 2 by Amina Yu and Version 1 by Aleksandra Bandzerewicz.

The existence of orderly structures such as tissues and organs is made possible by cell adhesion, i.e. the process by which cells attach to neighbouring cells and a supporting substance in the form of the extracellular matrix. The extracellular matrix is a three-dimensional structure composed of collagens, elastin and various proteoglycans and glycoproteins. It is a storehouse for multiple signalling factors. Tissue disruption often prevents the natural reconstitution of the matrix. The use of appropriate implants is then required. The possibilities of regenerating damaged tissues using an artificial matrix substitute are described, detailing the host response to the implant. An important issue is the surface properties of such an implant and the possibilities of their modification.

  • extracellular matrix
  • cellular receptors
  • cell adhesion
  • cell signalling
  • scaffolds
  • biomaterials

1. The Extracellular Matrix—Composition, Structure, Functions

1.1. Two Types of the Extracellular Matrix

Although the basic organisation of the  extracellular matrix (ECM) structure is the same throughout, two basic types of the matrix are distinguished by their location and composition: the interstitial matrix, which forms a three-dimensional porous network surrounding the cells (especially connective tissues), and the pericellular matrix, which is more compact and forms a layer adjacent to the cells [15,16][1][2] (see Figure 1).
Cells 11 00914 g001 550
Figure 1. Simplified extracellular matrix structure: three-dimensional macromolecular network composed of various proteins and polysaccharides. The pericellular matrix forms a layer adjacent to the cells: integrins bind to polymerised laminin, which, in turn, is connected via nidogen to the type IV collagen. Interstitial matrix forms porous network of fibrillar collagens, elastic fibres, and proteoglycans.
The interstitial matrix can be equated with the “proper” matrix, as it forms the structural scaffolding for the cells. Its basic components are heterotypic fibrils, composed mainly of type I collagen with small amounts of type III and V collagens in variable proportions, both playing an important role in fibrillogenesis [16][2]. The collagens of the interstitial matrix are mostly secreted by fibroblasts [17][3]. Important components of this “amorphous three-dimensional gel” also include fibronectin and elastin, involved in the organisation of the structure [18,19][4][5].
A typical example of the pericellular matrix is the basement membrane, a delicate and flexible nanostructure that separates the epithelium from the deeper layers of connective tissues. It ensheathes smooth, skeletal, and cardiac muscle fibres, Schwann cells, and adipocytes. The basement membrane forms a specific boundary of many organs in mature tissues, often surrounding their functional units [16,20,21,22][2][6][7][8]. It is mainly composed of type IV collagen, laminins, nidogens and heparan sulfate proteoglycans (HSPGs): perlecan and agrin [21][7]. The basement membrane contains so-called matricellular proteins that do not contribute to its physical stability or structural integrity, although they may be connected to building components. Instead, they have regulatory functions and interact with surface receptors, proteases, hormones or other biologically active molecules. They may be tissue-specific in terms of function and structure [23,24,25,26][9][10][11][12]. Matricellular proteins include SPARC (secreted protein acidic and rich in cysteine, or osteonectin; characteristic of mineralising tissues, mainly bone), thrombospondin-1 (which is rich in platelet α-granules; when secreted, it causes, among other things, activation of TGF-β1, i.e., transforming growth factor-beta 1), and tenascin-C (the gene of this protein is expressed during embryonic life, while in adult tissues, tenascin-C is very poorly detectable, being present rather in the course of pathological processes [27,28,29][13][14][15]. The tasks of the basement membrane include regulation of tissue development, function, and regeneration by controlling the cellular response. It is a storehouse of growth factors and modulates their activity and concentration. It serves to maintain the phenotype of the cells it surrounds [30][16]. The interstitial matrix and the basement membrane are closely interconnected, ensuring the integrity of the tissue [16][2].
The functional equivalent of the basement membrane described above is a type of pericellular matrix that surrounds chondrocytes in articular cartilage [31][17]. It acts as a physical barrier that filters molecules entering and leaving the cells. Together with an adjacent thin layer of matrix, each chondrocyte forms a structural unit called a chondron [32][18]. The morphology of chondrons varies. They can take a discoid/ellipsoid/rounded shape and a variable orientation, which depends on the position, i.e., the depth of location in the cartilage. In some cases, a chondron comprises more than one cell (up to four) [33][19]. IAn this case, an eessential component of the pericellular matrix is type VI collagen, although it generally constitutes a negligible percentage of the collagens of cartilage tissue [34][20]. However, because of its specific presence in the chondrocyte environment of articular cartilage, it often serves as a marker of chondrons [34,35][20][21]. A characteristic feature of articular cartilage is the small number of chondrocytes compared to the extensive extracellular (interstitial) matrix for which synthesis, organisation, and maintenance they are responsible [34][20].

1.2. Major Components of the Extracellular Matrix and Their Functions

1.2.1. Collagens

Collagen proteins account for up to 30% of all proteins in vertebrates and are major extracellular matrix components. The basic collagen macromolecules are composed of three same (homotrimers) or different (heterotrimers) polypeptide chains. They are characterised by the repetitive Gly-X-Y sequences, where X usually stands for proline and Y for 4-hydroxyproline. The intertwined chains form a specific triple helix structure [17,36,37][3][22][23].
Due to their supramolecular organisation, fibrillar (types I, II, III, V, XI, XXIV, and XXVII) and non-fibrillar collagens are distinguished. Characteristic for the non-fibrillar collagens is a disrupted continuity of the typical structure. Compared to fibrillar collagens, they contain shorter (although more numerous) helical (collagenous) domains interspersed with so-called telopeptides, i.for example., non-helical domains. As a result, they may occur in various forms, forming, e.g., network systems (types IV, VIII, X), anchor fibres (type VII), beaded filaments (type VI), or belong to the FACIT group (i.e.for example, fibril-associated collagen with interrupted triple helix, types XI, XII, XIV, XVI, XIX-XXII). The terminology and affiliation are not fully systematised. Collectively, collagens form a family of 28 proteins [38,39,40,41,42,43,44][24][25][26][27][28][29][30].
Historically, collagens were thought to have only a supportive function. Although their main function is indeed to form the structural scaffolding of cells (especially for types I, II, and III), it is known that their role is much broader [15,18,37,45][1][4][23][31]. Collagens are involved in regulating the course of cell adhesion (as ligands of cell receptors) [46[32][33][34][35],47,48,49], cell migration (contact guidance) [50,51,52][36][37][38] and tissue reconstruction and remodelling [37,40,45,53,54][23][26][31][39][40]. Not only is the physical deposition or movement of cells itself important, but also the processes conditioned by this,for e.g.xample,, wound healing, immune response, etc and so on. Although collagens are present in most body tissues and affect their mechanical properties, their distribution varies, e.g.for example, type I collagen is characteristic of bone, skin and tendon, and type II collagen of cartilage tissue [55,56][41][42].

1.2.2. Elastin

Elastin is a hydrophobic fibrillar protein, which owes its characteristic elastic properties to extensive covalent cross-linking of the structure [57][43]. The monomer from which the mature insoluble protein is formed is tropoelastin, secreted by fibroblasts, smooth muscle cells, endothelial cells, respiratory epithelial cells, chondrocytes, and keratinocytes [58,59,60,61,62][44][45][46][47][48]. After secretion into the intercellular space, tropoelastin spontaneously associates into larger particles through interactions between hydrophobic domains in a process called coacervation [63][49]. Such precursors undergo oxidative deamination of lysine residues in tropoelastin. The process is catalysed by LOX family enzymes (lysyl oxidases). The result is the formation of allysine from lysine. Cross-linking occurs via the reaction between lysine and allysine residues (Schiff base reaction) or by aldol condensation of two allysine residues [64,65,66,67][50][51][52][53]. The final fibres are not composed of elastin alone. Elastin forms a core (about 90% of the whole structure), covered by an envelope of microfibrils composed mainly of glycoproteins from the fibrillin group (fibrillin-1 and -2) [58,68,69][44][54][55]. In this way, elastic fibres are formed, giving tissues susceptibility to stretching. They are a particularly important component of blood vessel walls, skin, lungs, heart, tendons, ligaments, bladder, elastic cartilage tissue (for e.g.xample, auricle, larynx, epiglottis), etc and so on. [15,68,70][1][54][56].
The gene expression and formation of elastic fibres occur at early development stages —prenatal and early childhood. De novo production of elastin in adult organisms is unlikely to occur, which is quite uncommon among the ECM components [58,71,72,73,74][44][57][58][59][60]. However, elastin has high metabolic stability and a half-life of approximately 70 years, making the limited synthesis time sufficient (by comparison, the half-life of type VII collagen is estimated to be approximately one month [75,76][61][62]. The adult organism cannot reconstitute elastic fibres that become damaged or degrade progressively with age. They are then repaired incorrectly and consequently do not perform their normal functions. The tissues become too stiff, leading to cardiovascular disease, lung disease or typical signs of ageing, such as loss of skin elasticity [15,67,77,78][1][53][63][64].

1.2.3. Proteoglycans

Proteoglycans are macromolecules of a complex three-dimensional structure. They are composed of a protein core covalently linked to one or more chains of glycosaminoglycans (GAGs), a type of linear, unbranched heteropolysaccharides. The glycosaminoglycan chains may belong to one or different types. Based on localisation, four basic groups of proteoglycans can be distinguished: intracellular and those occurring on the cell surface, in the pericellular space (basement membrane) or intercellular space [15,79,80,81][1][65][66][67].
GAG chains are built by repeating disaccharide units, where one residue is an amino sugar (N-acetylated hexosamine), and the other is uronic acid (D-glucuronic or L-iduronic acid). GAGs differ in the type of monosaccharide residues and the geometry of the linkages between the constituent units (α- and β-glycosidic linkages) and the degree of sulfation of the polysaccharide backbone and the position of this substitution. Based on the chemical structure of the chain, four basic groups of glycosaminoglycans are distinguished: heparan/heparan sulfate, keratan sulfate, chondroitin sulfate/dermatan sulfate, and hyaluronic acid [15,82,83,84,85,86][1][68][69][70][71][72]. Hyaluronic acid represents the simplest type of structure. It is the only one that does not contain sulfate groups (hydroxyl groups are not esterified with sulfate groups) and does not undergo complex modifications in the Golgi apparatus [84,87,88][70][73][74]. Unlike other GAGs, it does not form covalent bonds with proteins and, therefore, is not part of typical proteoglycans. Instead, it can exist in the form of non-covalent complexes with other protein components of the ECM [85,86,89,90,91][71][72][75][76][77].
Hyaluronan has excellent water retention ability. It is abundant in the skin, cartilage, brain, vitreous body, umbilical cord, and synovial fluid. Its physical and physiological properties depend on molecular weight and concentration in the tissue. When highly concentrated, hyaluronan molecules form a three-dimensional meshwork structure exhibiting remarkable viscoelasticity. The organised structure acts as a molecular sieve of proteins and other macromolecules. Hyaluronan is reported to modulate cellular behaviours via the reprogramming of cellular metabolism coupled to its production [92][78]. Hyaluronan activates signalling cascade by interacting with CD44 receptor. CD44 was originally identified as a hyaluronan and hyaluronic acid receptor but can bind to various other ligands. It also serves as a marker for stem cells of several types [93][79].
Glycosaminoglycan chains (and, therefore, the proteoglycans) are negatively charged. It is the result of carboxyl and sulfate residues in their structure [88,94][74][80]. Due to the strong negative charge, these molecules tend to elongate in solution under physiological conditions. This allows them to bind large amounts of water and form a gel. Such properties provide tissues with resistance to deformation by high physical forces, as exemplified by aggrecan, the most important cartilage proteoglycan [79,86,95,96][65][72][81][82]. The proteoglycan family also includes compounds, such as syndecans (trans-membrane receptors; they bind numerous ligands present in the ECM, mediate signal transduction, cell adhesion, migration et al.) [97[83][84],98], serglycin (the only known intracellular proteoglycan; found in leukocyte granules, regulates granulopoiesis) [99[85][86][87],100,101], perlecan and agrin (characteristic of the basement membrane, regulators of many cellular processes; agrin is involved in the formation of neuromuscular synapses) [102,103,104,105,106,107][88][89][90][91][92][93] and fibromodulin (involved in the collagen fibrillogenesis) [108,109][94][95].

1.2.4. Glycoproteins

Like proteoglycans, glycoproteins are composed of covalently linked protein and carbohydrate parts. However, the saccharide chains are much shorter, contain no (or few) repeating units, and are usually branched [2,110,111][96][97][98]. Glycoproteins often act as connectors in the ECM, as they have functional groups capable of binding other proteins, growth factors, or receptors [2,112,113][96][99][100]. Their participation is essential for many biological processes: fertilisation, immune and inflammatory response, blood coagulation, wound healing, etcand so on. [112,114,115,116,117,118,119,120][99][101][102][103][104][105][106][107]. The two most important glycoproteins are fibronectin and laminin. The glycoprotein family also includes fibulins [121][108], tenascin [122][109], fibrinogen [123][110], vitronectin [124][111], osteonectin [27][13], bone sialoprotein [125][112], and reelin [126][113].
The basic structural unit of fibronectin is a dimer composed of two nearly identical polypeptide chains linked by a pair of disulfide bonds. Each such chain is built by irregularly repeating amino acid units (types I, II, and III), forming a mosaic structure of the protein. The molecules consist of domains, for i.e.example, differently structured sections with different functions [127,128,129][114][115][116]. Fibronectin contains domains capable of interacting with the ECM proteins (for e.g.xample, collagen), glycosaminoglycans, surface receptors and other fibronectin molecules. Due to these properties, fibronectin can simultaneously bind to cells and components of the surrounding matrix [128,130,131,132,133,134][115][117][118][119][120][121]. In the body, fibronectin exists in two forms: soluble plasma fibronectin (synthesised by hepatocytes and secreted into the blood) and insoluble cellular fibronectin (produced by fibroblasts, endothelial cells, chondrocytes, myocytes, and others). The insoluble form is a fibrillar cross-linked structure on the cell surface and in the ECM. It is responsible for cell adhesion, proliferation, migration, and the ECM protein deposition [128,135,136,137,138,139][115][122][123][124][125][126]. Both forms of fibronectin are encoded by one gene, while structural differences result from alternative mRNA splicing [140,141][127][128].
Laminins are a group of large, multi-domain glycoproteins of a heterotrimeric structure. The three subunits (α, β, and γ chains) connected by a pair of disulfide bonds form a characteristic Latin cross-shaped structure (a Y-shape/rod shape form is also possible [142,143,144][129][130][131]). The three shorter arms (their globular N-terminal domains) are mainly involved in laminin polymerisation and network self-assembly. At the same time, the longer one mediates cell–cell interactions by binding to receptors [145,146,147,148,149][132][133][134][135][136]. Proteins of the laminin family are an integral part of the basement membrane and play an essential role in forming and maintaining its structure. A critical step in developing the basement membrane is the polymerisation of laminin [149,150,151][136][137][138]. This process is initiated by binding laminin molecules to the cell surface. A connection is formed between the long arm of the protein and the receptors—cognate integrin and dystroglycan. As a result, there is a local increase in the concentration of laminin, and after exceeding a critical value, polymerisation occurs. The structure, thus, formed binds to nidogens and HSPGs (perlecan). The entire network is further stabilised by polymerising type IV collagen [151,152,153,154,155,156,157,158][138][139][140][141][142][143][144][145]. The basement membrane layer built up by the complex network of the described components is called lamina densa (the middle layer between the lamina lucida and the lamina fibroreticularis [159][146].

1.3. The Dynamic Structure of the Extracellular Matrix

The structure of the extracellular matrix undergoes continuous remodelling, during which changes in its composition and overall architecture occur. Cells embedded in the ECM are actively involved in its reorganisation. In addition to synthesising and secreting building components, they are also the source of enzymes that degrade these components. Remodelling processes are complex and must be tightly regulated to maintain environmental homeostasis [19,160,161,162][5][147][148][149].
Protein-degrading enzymes belong to the class of hydrolases and are called proteases (proteinases). Depending on the mechanism of catalysis, they can be divided into several families, including serine proteases (serine residue in the enzyme active site), cysteine proteases (cysteine residue) or metalloproteases (they require the presence of a metal cation in the active centre). These enzymes can be secreted by the cell into its external environment or remain anchored in the cell membrane [163,164][150][151].
The main group of enzymes involved in ECM degradation are the zinc-dependent matrix metalloproteinases (MMPs). More than 20 representatives of this group are known, capable of degrading different types of collagen, gelatin, elastin, laminin, fibronectin and many others [165,166,167][152][153][154]. The sources of MMPs are mainly connective tissue cells (fibroblasts, osteoblasts), inflammatory cells (macrophages, neutrophils, mast cells), and endothelial cells [165,168][152][155]. MMPs are secreted in the form of zymogens, inactive precursors that must undergo biochemical modifications to be activated [19,165,168][5][152][155]. Through controlled degradation of ECM proteins, metalloproteinases facilitate cell migration and trigger the release of growth factors [169,170,171][156][157][158]. They participate in tissue remodelling, an interesting example of which is postpartum uterine involution. In addition, they regulate angiogenesis (blood vessel formation), wound healing, embryonic development, and so etcon. [165,172,173][152][159][160]. In pathological states, their abnormal and/or increased activity contributes to the course of cardiovascular, cancer, autoimmune diseases and s, etco on. [165,174,175,176][152][161][162][163].
The proteolysis occurring in tissues relates not only to the extracellular matrix per se but also concerns the so-called ectodomain shedding, i.e., proteolytic cleavage of cell surface proteins. Modification, degradation, and changes in the activity of these proteins are one of the mechanisms of the cell’s response to changes in microenvironment conditions [177,178][164][165]. Enzymes of the ADAM (a disintegrin and metalloproteases) family, also known as adamalysins, are mainly involved in this process. They have various functions, primarily engaged in intercellular interactions and signal transduction [19,179,180][5][166][167]. The release of biologically active extracellular domains of multiple proteins (cytokines, adhesion molecules, growth factors) from the cell membrane can contribute, e.g., to inflammation (physiological and pathological), as occurs as a result of ADAM17 enzyme activity. The pro-inflammatory action of this sheddase consists of a modification of the cell surface and enrichment of its environment with active soluble molecules [181,182,183,184][168][169][170][171]. The structure and function of ADAM group proteins are similar to the metalloproteinases found in snake venom, responsible for the typical effects of snakebites (haemorrhage, tissue necrosis) [185][172].

1.4. The Extracellular Matrix as a Storehouse of Growth Factors

The ECM significantly influences the cell’s most important natural biological processes: growth, proliferation, and programmed death [186][173]. In addition to mediating interactions and activating relevant mechanisms by contact with its building proteins, the ECM serves as a storehouse of growth factors (and proteases and protease inhibitors). These molecules can be released by proteolytic degradation of the matrix, and the degradation itself regulates the rate, site and intensity of such activation. The fact that growth factors are stored in the vicinity of cells favours increased specificity of their action [19,187,188,189][5][174][175][176].
Growth factors are generally not freely dispersed in the extracellular space but bind, for example, to heparan sulphate proteoglycans. HSPGs then participate in the matrix storage function by preventing the movement and proteolysis of growth factors. They allow their controlled release when necessary. However, another role of HSPGs is also to bind to such molecules to activate them. Then, they act as a coreceptor in ligand–receptor interactions [187,190,191,192][174][177][178][179]. The type of interaction of HSPGs with growth factors depends on the localisation of these proteoglycans. They may remain anchored to the cell membrane or form a structural component of the ECM [193][180].
A well-studied group is the fibroblast growth factors (FGF), which include 22 proteins with key functions in cell development, morphogenesis, tissue repair processes, and angiogenesis. They are among the neurotrophic factors, for i.e.example, those that stimulate and regulate neurogenesis. Some are being investigated for involvement in the development of depression [192,194][179][181]. FGF molecules are mainly bound by heparan sulfate and heparin chains [195,196][182][183]. Proteolytic release of FGF allows subsequent binding of FGF ligands to receptors on the cell surface. This stimulates cell signalling [2][96].

2. Artificial Substitutes of the Extracellular Matrix

2.1. Host Response to Implantation

The body’s first biological reaction to an implant is forming a layer of water on its surface. This happens in just a few nanoseconds. Water molecules form a mono- or bilayer, and the way they are ordered is strongly dependent on surface properties at the atomic level. Water molecules can dissociate on a highly reactive substrate, resulting in the hydroxylation of the implant surface, i.for example., it becomes covered with -OH groups. Water molecules can also be strongly bound but not dissociate. Both of these cases occur as a result of contact with a hydrophilic surface. If the surface is hydrophobic, its interactions with water are much weaker. Therefore, the strength of water-binding determines hydrophobicity or hydrophilicity to the surface. It influences the value of the wetting angle formed between the solid and the plane tangent to the droplet deposited on it. Hydrated ions, such as Cl−, Na+, Ca2+, enter the formed water layer [403,404][184][185].
Once the aqueous layer covers the material’s surface, proteins from body fluids (extravasated blood/tissue fluid) reach it. In the first stage, mainly smaller proteins with the highest mobility are adsorbed, resulting from faster diffusion of small than large molecules. It is a transient state. A dynamic adsorption–desorption equilibrium is established at the contact surface, as proteins with larger size and a stronger affinity for the implanted material, arriving late, can force the desorption of smaller, weak-bound molecules. This phenomenon is called the Vroman effect. It should be kept in mind that fluids in contact with the implant, such as plasma, contain hundreds of different proteins competing for access to the surface. Therefore, the adsorption–desorption process is much more complex and depends on factors, such as the protein concentration in the fluid. The higher the concentration, the greater the primary surface dominance [404,405][185][186].
Proteins usually have an asymmetric structure in which domains of different chemical nature can be distinguished. They have a more or less ellipsoidal shape (globular proteins) [406][187]. As a result of adsorption, conformational changes of the molecule can occur if it is sufficiently susceptible. It is the effect of binding to the substrate with a privileged side in a given case. As a result, the molecule adopts a certain orientation where part of it invariably contacts the body fluid [407,408,409][188][189][190]. Structurally stable proteins do not readily undergo conformational changes. Their adsorption may occur along the longest axis (“side-on”). Otherwise, this axis is perpendicular to the implant surface (“end-on”) [407][188]. The issue is not insignificant in the context of establishing a dynamic adsorption–desorption equilibrium, as the ability to structurally reorient increases the possibility of contact with the substrate [407,410][188][191].
A major problem with implantation is the foreign body response (FBR), a complex process involving different cell types. Neutrophils are the first to reach the implant site and adhere (via proteins) to the protein-coated surface of the material. Activated neutrophils attempt to degrade the implant by secreting factors, such as proteolytic enzymes or reactive oxygen species. They release chemokines that attract other immune cells, mainly monocytes [411,412,413][192][193][194]. These, in turn, reaching their target, differentiate into macrophages [414][195]. The number of macrophages at the implantation site increases due to their progressive proliferation. They replace the initial wave of nucleophiles and release further pro-inflammatory factors. It may lead to implant damage and/or the release of toxic substances into the surrounding tissue environment [415,416][196][197]. Macrophages may fuse into foreign body giant cells (FBGCs) due to chronic cytokine activity. FGBCs can adhere to the material’s surface for an extended time, leading to collagen deposition and fibrous encapsulation (approximately 3–4 weeks after implantation). As a result, the implant is isolated from the surrounding tissues. It prevents integration and vascularisation and ultimately leads to implant loss [412,417][193][198]. The fibrous layer is usually thinner on porous than on solid materials [418,419][199][200]. The presence of mast cells, degranulating upon activation, is also characteristic at the implant site. Among other things, histamine is released from the granules. Histamine dilates blood vessels, improves their permeability and facilitates the arrival of other immune cells. Pro- and anti-inflammatory cytokines and angiogenic or profibrotic factors are also secreted [420,421][201][202].
Immunosuppressive drugs are used to weaken the body’s immune response and prevent implant rejection. A more recent solution is to incorporate anti-inflammatory agents into the implanted material. They must be released in a controlled manner and at an appropriate rate. An additional requirement is to promote angiogenesis [422,423][203][204]. For years, biomaterials engineering has been focused on obtaining biologically inert materials, i.for example.,, minimising the interaction with the organism and reducing the immune response. The contemporary trend is the generation of biomimetic materials, i.e.for example,, mimicking the natural solutions of the organism and stimulating the desired responses. These include enhancing or inhibiting the normal functioning of immune cells [424,425,426][205][206][207].

2.2. Influence of Material Properties on Cell Adhesion

Cells do not experience direct contact with the implanted material but are only ‘informed’ of its physicochemical properties via proteins deposited on the surface. One of the more important characteristics of the material is the wettability of its surface, which, in the case of an aqueous environment, can be equated with hydrophilicity. It is assumed that the ability of cells to adhere increases on hydrophilic surfaces and decreases on hydrophobic surfaces, even though it is hydrophobic surfaces that are generally considered to be more protein-adsorbent [427][208].
The surface protein layer that forms shortly after implantation consists mainly of albumin, fibrinogen, immunoglobulin G, fibronectin, vitronectin et al. The first interactions are usually dominated by albumin due to its relatively small size (66 kDa) and high-concentration in plasma [411,428,429][192][209][210]. It binds much more readily to hydrophobic than hydrophilic surfaces but does not promote cell adhesion. The strong adsorption of albumin reduces the likelihood of being replaced by larger adhesion-promoting proteins, such as fibronectin and vitronectin [430,431,432][211][212][213]. The ability of fibronectin to displace surface-bound albumin is limited on hydrophobic surfaces. As a result of the strong binding of albumin molecules, changes in their secondary structure occur and the degree of denaturation increases [433][214]. Proteins tend to denature as the contact time with the material increases, which occurs when albumin adsorbs onto a hydrophobic material. The binding energy of the adsorbed phase then increases, and, as a result, the probability of desorption decreases [406][187].
Adsorption occurs more readily if there is a charge difference between the protein molecules and the material surface [434][215]. Furthermore, the affinity of the protein for the material may show greater specificity than the distinction between hydrophobicity/hydrophilicity and be based on the recognition of specific functional groups [429,433][210][214]. Additionally, the cells themselves, depending on the type, show a different preference for the functionality of the surface groups [435,436,437,438][216][217][218][219].
Adhesion of cells to the implant surface is made possible by integrins recognising and binding to specific amino acid sequences in the polypeptide chain of the adsorbed protein. It mimics the formation of integrin connections with the ECM proteins under natural conditions. The best known among the pro-adhesive sequences is the tripeptide RGD (arginine-glycine-aspartic acid), present, e.g., in the structure of fibronectin [439,440][220][221]. One way to modify the material to increase biocompatibility is the coating of tripeptide RGD on its surface in the form of immobilised proteins or short synthetic polypeptide ligands. In addition to RGD, the collagen peptide GFOGER (glycine-phenylalanine-hydroxyproline-glycine-glutamate-arginine) and the laminin-specific sequences IKVAV (isoleucine-lysine-valine-alanine-valine) and YIGSR (tyrosine-isoleucine-glycine-serine-arginine), among others, have been identified [441,442,443,444,445][222][223][224][225][226].
Functionalisation of the implant surface with peptides containing the RGD sequence has drawbacks. Integrins that recognise RGD may require the presence of other peptides (synergistic effect) to form a bond. The biological activity of short synthetic peptides is less than that of a whole protein. In turn, modification of these peptides (e.g., by chain elongation) can also result in an undesirable change (increase/reduction) in their activity. Another problem is that cells adhere too strongly to the surface, reducing their movement ability [446,447,448][227][228][229].
An interesting conclusion is provided by the study of cell adhesion on materials exhibiting extreme wettability types. Superhydrophobic surfaces are characterised by a water contact angle value higher than 150°, while superhydrophilic surfaces are around 0°. Although the type of cell determines the contact behaviour, only a few show good adhesion to a surface if the material is superhydrophobic. If the surface has highly hydrophilic and hydrophobic regions, cells will usually selectively attach to the superhydrophilic areas [449][230].
A significant feature of an implant is the topography of its surface, which, like the chemical composition, influences the interactions with integrins and ultimately stimulates the cellular response [450][231]. The shape of the natural matrix at the micro- and nanoscale is understood to be the structure formed by the ECM proteins and the neighbouring cells. For synthetic materials, it is the degree of roughness, the type and size of patterns on the surface. Modifications of these features at the nanoscale affect the activity of the adsorbing proteins by forcing specific changes in their conformation. However, the detailed investigation of such relationships is complicated because the cellular response is always a resultant of the influence of different stimuli. In addition, modifications of topography may be accompanied by changes in surface chemistry [451,452,453,454,455][232][233][234][235][236].
Techniques to create micropatterns on substrates can be divided into two main types: (1) coating portions of the material with an agent that promotes selective adhesion or (2) applying a layer that blocks adhesion and subsequently removing it without harming the cells embedded around it [456][237]. The resulting pattern geometry influences the subsequent formation of cells, for e.g.xample, it promotes cell elongation. Furthermore, it supports/inhibits the spreading of cells on the surface. It is related to facilitating/hindering their movement, respectively, depending on the continuity of the pattern [457][238]. The size of the contact area between cells can influence their differentiation, i.for example., result in different types of daughter cells [458][239]. Discontinuities in topography are the cause of local differences in surface free energy. If the cell can detect it, it will modify the contact orientation by reorganising its cytoskeleton. Mechanical signals transmitted to the cell nucleus affect changes at the level of gene transcription and consequently determine cell behaviour. However, the mechanisms underlying the cellular response are still poorly understood [459,460][240][241].
In addition to patterns characterised by uniformity of shape and size, cell adhesion is influenced by the surface roughness, understood as the overall three-dimensional topography of the substrate, regardless of its regularity. The surfaces of the used materials are rarely smooth at the molecular level, while roughness is not uniformly describable in this case. Cells must be able to recognise a rough surface to react in a certain way, which is dependent on the cell type, as the primary determining factor is the size of the cell. It means that a cell will recognise a surface as smooth if the peak-to-peak distance is greater than the size of the cell [461,462,463][242][243][244].

References

  1. Theocharis, A.D.; Skandalis, S.S.; Gialeli, C.; Karamanos, N.K. Extracellular matrix structure. Adv. Drug Deliv. Rev. 2016, 97, 4–27.
  2. Theocharis, A.D.; Manou, D.; Karamanos, N.K. The extracellular matrix as a multitasking player in disease. FEBS J. 2019, 286, 2830–2869.
  3. Frantz, C.; Stewart, K.M.; Weaver, V.M. The extracellular matrix at a glance. J. Cell Sci. 2010, 123, 4195–4200.
  4. Kular, J.K.; Basu, S.; Sharma, R.I. The extracellular matrix: Structure, composition, age-related differences, tools for analysis and applications for tissue engineering. J. Tissue Eng. 2014, 5, 112.
  5. Bonnans, C.; Chou, J.; Werb, Z. Remodelling the extracellular matrix in development and disease. Nat. Rev. Mol. Cell Biol. 2014, 15, 786–801.
  6. Halfter, W.; Candiello, J.; Hu, H.; Zhang, P.; Schreiber, E.; Balasubramani, M. Protein composition and biomechanical properties of in vivo-derived basement membranes. Cell Adhes. Migr. 2013, 7, 64–71.
  7. Pozzi, A.; Yurchenco, P.D.; Iozzo, R.V. The nature and biology of basement membranes. Matrix Biol. 2017, 57–58, 1–11.
  8. Yurchenco, P.; Patton, B. Developmental and Pathogenic Mechanisms of Basement Membrane Assembly. Curr. Pharm. Des. 2009, 15, 1277–1294.
  9. Bornstein, P.; Sage, E.H. Matricellular proteins: Extracellular modulators of cell function. Curr. Opin. Cell Biol. 2002, 14, 608–616.
  10. Murphy-Ullrich, J.E.; Sage, E.H. Revisiting the matricellular concept. Matrix Biol. 2014, 37, 1–14.
  11. Keller, K.E.; Kelley, M.J.; Acott, T.S. Extracellular matrix gene alternative splicing by trabecular meshwork cells in response to mechanical stretching. Investig. Ophthalmol. Vis. Sci. 2007, 48, 1164–1172.
  12. Bornstein, P. Matricellular proteins: An overview. J. Cell Commun. Signal. 2009, 3, 163–165.
  13. Rosset, E.M.; Bradshaw, A.D. SPARC/osteonectin in mineralized tissue. Matrix Biol. 2016, 52–54, 78–87.
  14. Bornstein, P. Thrombospondins as matricellular modulators of cell function. J. Clin. Investig. 2001, 107, 929–934.
  15. Midwood, K.S.; Chiquet, M.; Tucker, R.P.; Orend, G. Tenascin-C at a glance. J. Cell Sci. 2016, 129, 4321–4327.
  16. Wilson, S.E.; Torricelli, A.A.M.; Marino, G.K. Corneal epithelial basement membrane: Structure, function and regeneration. Exp. Eye Res. 2020, 194, 108002.
  17. Kvist, A.J.; Nyström, A.; Hultenby, K.; Sasaki, T.; Talts, J.F.; Aspberg, A. The major basement membrane components localize to the chondrocyte pericellular matrix-A cartilage basement membrane equivalent? Matrix Biol. 2008, 27, 22–33.
  18. Zhang, Z. Chondrons and the Pericellular Matrix of Chondrocytes. Tissue Eng.-Part B Rev. 2015, 21, 267–277.
  19. Youn, I.; Choi, J.B.; Cao, L.; Setton, L.A.; Guilak, F. Zonal variations in the three-dimensional morphology of the chondron measured in situ using confocal microscopy. Osteoarthr. Cartil. 2006, 14, 889–897.
  20. Poole, C.A. Review. Articular cartilage chondrons: Form, function and failure. J. Anat. 1997, 191, 1–13.
  21. Fraser, S.A.; Crawford, A.; Frazer, A.; Dickinson, S.; Hollander, A.P.; Brook, I.M.; Hatton, P.V. Localization of type VI collagen in tissue-engineered cartilage on polymer scaffolds. Tissue Eng. 2006, 12, 569–577.
  22. Hynes, R.O.; Naba, A. Overview of the matrisome-An inventory of extracellular matrix constituents and functions. Cold Spring Harb. Perspect. Biol. 2012, 4, a004903.
  23. Barnes, M. Update on Collagens: What You Need to Know and Consider. Plast. Surg. Nurs. 2019, 39, 112–115.
  24. Exposito, J.Y.; Valcourt, U.; Cluzel, C.; Lethias, C. The fibrillar collagen family. Int. J. Mol. Sci. 2010, 11, 407–426.
  25. Bella, J.; Hulmes, D.J.S. Fibrillar collagens. Subcell. Biochem. 2017, 82, 457–490.
  26. Mienaltowski, M.J.; Birk, D.E. Structure, Physiology, and Biochemistry of Collagens. In Advances in Experimental Medicine and Biology; Springer: Berlin/Heidelberg, Germany, 2014; Volume 802, pp. 5–29.
  27. Shaw, L.M.; Olsen, B.R. FACIT collagens: Diverse molecular bridges in extracellular matrices. Trends Biochem. Sci. 1991, 16, 191–194.
  28. Ricard-Blum, S.; Ruggiero, F. The collagen superfamily: From the extracellular matrix to the cell membrane. Pathol. Biol. 2005, 53, 430–442.
  29. Ivanova, V.P.; Krivchenko, A.I. Current viewpoint on structure and on evolution of collagens. II. Fibril-associated collagens. J. Evol. Biochem. Physiol. 2014, 50, 273–285.
  30. Shoulders, M.D.; Raines, R.T. Collagen structure and stability. Annu. Rev. Biochem. 2009, 78, 929–958.
  31. Harsha, L.; Brundha, M.P. Role of collagen in wound healing. Drug Invent. Today 2020, 13, 55–57.
  32. Heino, J. The collagen family members as cell adhesion proteins. BioEssays 2007, 29, 1001–1010.
  33. Luckman, S.P.; Rees, E.; Kwan, A.P.L. Partial characterization of cell-type X collagen interactions. Biochem. J. 2003, 372, 485–493.
  34. Smethurst, P.A.; Onley, D.J.; Jarvis, G.E.; O’Connor, M.N.; Graham Knight, C.; Herr, A.B.; Ouwehand, W.H.; Farndale, R.W. Structural basis for the platelet-collagen interaction: The smallest motif within collagen that recognizes and activates platelet Glycoprotein VI contains two glycine-proline-hydroxyproline triplets. J. Biol. Chem. 2007, 282, 1296–1304.
  35. Paavola, K.J.; Sidik, H.; Zuchero, J.B.; Eckart, M.; Talbot, W.S. Type IV collagen is an activating ligand for the adhesion G protein-coupled receptor GPR126. Sci. Signal. 2014, 7, ra76.
  36. Wolf, K.; Alexander, S.; Schacht, V.; Coussens, L.M.; von Andrian, U.H.; van Rheenen, J.; Deryugina, E.; Friedl, P. Collagen-based cell migration models in vitro and in vivo. Semin. Cell Dev. Biol. 2009, 20, 931–941.
  37. Guido, S.; Tranquillo, R.T. A methodology for the systematic and quantitative study of cell contact guidance in oriented collagen gels. Correlation of fibroblast orientation and gel birefringence. J. Cell Sci. 1993, 105, 317–331.
  38. Borgne-Rochet, M.L.; Angevin, L.; Bazellières, E.; Ordas, L.; Comunale, F.; Denisov, E.V.; Tashireva, L.A.; Perelmuter, V.M.; Bièche, I.; Vacher, S.; et al. P-cadherin-induced decorin secretion is required for collagen fiber alignment and directional collective cell migration. J. Cell Sci. 2019, 132, 3189.
  39. Canty, E.G.; Kadler, K.E. Collagen fibril biosynthesis in tendon: A review and recent insights. Comp. Biochem. Physiol. Part A Mol. Integr. Physiol. 2002, 133, 979–985.
  40. Zhang, G.; Young, B.B.; Birk, D.E. Differential expression of type XII collagen in developing chicken metatarsal tendons. J. Anat. 2003, 202, 411–420.
  41. Ricard-Blum, S. The Collagen Family. Cold Spring Harb. Perspect. Biol. 2011, 3, 1–19.
  42. Parenteau-Bareil, R.; Gauvin, R.; Berthod, F. Collagen-based biomaterials for tissue engineering applications. Materials 2010, 3, 1863–1887.
  43. Cocciolone, A.J.; Hawes, J.Z.; Staiculescu, M.C.; Johnson, E.O.; Murshed, M.; Wagenseil, J.E. Elastin, arterial mechanics, and cardiovascular disease. Am. J. Physiol.-Hear Circ. Physiol. 2018, 315, H189–H205.
  44. Mariani, T.J.; Dunsmore, S.E.; Li, Q.; Ye, X.; Pierce, R.A. Regulation of lung fibroblast tropoelastin expression by alveolar epithelial cells. Am. J. Physiol.-Lung Cell. Mol. Physiol. 1998, 274, L47–L57.
  45. Mecham, R.P.; Madaras, J.; McDonald, J.A.; Ryan, U. Elastin production by cultured calf pulmonary artery endothelial cells. J. Cell. Physiol. 1983, 116, 282–288.
  46. Kajiya, H.; Tanaka, N.; Inazumi, T.; Seyama, Y.; Tajima, S.; Ishibashi, A. Cultured human keratinocytes express tropoelastin. J. Investig. Dermatol. 1997, 109, 641–644.
  47. Narayanan, A.S.; Sandberg, L.B.; Ross, R.; Layman, D.L. The smooth muscle cell: III. Elastin synthesis in arterial smooth muscle cell culture. J. Cell Biol. 1976, 68, 411–419.
  48. Nishizaki, T. PKCϵ Increases Extracellular Elastin and Fibulin-5/DANCE in Dermal Fibroblasts. Cell. Physiol. Biochem. 2018, 46, 291–302.
  49. Vrhovski, B.; Weiss, A.S. Biochemistry of tropoelastin. Eur. J. Biochem. 1998, 258, 1–18.
  50. Ozsvar, J.; Yang, C.; Cain, S.A.; Baldock, C.; Tarakanova, A.; Weiss, A.S. Tropoelastin and Elastin Assembly. Front. Bioeng. Biotechnol. 2021, 9, 138.
  51. Vindin, H.; Mithieux, S.M.; Weiss, A.S. Elastin architecture. Matrix Biol. 2019, 84, 4–16.
  52. Lucero, H.A.; Kagan, H.M. Lysyl oxidase: An oxidative enzyme and effector of cell function. Cell. Mol. Life Sci. 2006, 63, 2304–2316.
  53. Wagenseil, J.E.; Mecham, R.P. New insights into elastic fiber assembly. Birth Defects Res. Part C-Embryo Today Rev. 2007, 81, 229–240.
  54. Baldwin, A.K.; Simpson, A.; Steer, R.; Cain, S.A.; Kielty, C.M. Elastic fibres in health and disease. Expert Rev. Mol. Med. 2013, 15, 23.
  55. Thomson, J.; Singh, M.; Eckersley, A.; Cain, S.A.; Sherratt, M.J.; Baldock, C. Fibrillin microfibrils and elastic fibre proteins: Functional interactions and extracellular regulation of growth factors. Semin. Cell Dev. Biol. 2019, 89, 109–117.
  56. Ritty, T.M.; Ditsios, K.; Starcher, B.C. Distribution of the elastic fiber and associated proteins in flexor tendon reflects function. Anat. Rec. 2002, 268, 430–440.
  57. Gabriela Espinosa, M.; Catalin Staiculescu, M.; Kim, J.; Marin, E.; Wagenseil, J.E. Elastic Fibers and Large Artery Mechanics in Animal Models of Development and Disease. J. Biomech. Eng. 2018, 140, 0208031.
  58. Dubick, M.A.; Rucker, R.B.; Cross, C.E.; Last, J.A. Elastin metabolism in rodent lung. BBA-Gen. Subj. 1981, 672, 303–306.
  59. Davidson, J.M.; Smith, K.; Shibahara, S.; Tolstoshev, P.; Crystal, R.G. Regulation of elastin synthesis in developing sheep nuchal ligament by elastin mRNA levels. J. Biol. Chem. 1982, 257, 747–754.
  60. Burnett, W.; Finnigan-Bunick, A.; Yoon, K.; Rosenbloom, J. Analysis of elastin gene expression in the developing chick aorta using cloned elastin cDNA. J. Biol. Chem. 1982, 257, 1569–1572.
  61. Kühl, T.; Mezger, M.; Hausser, I.; Guey, L.T.; Handgretinger, R.; Bruckner-Tuderman, L.; Nyström, A. Collagen VII Half-Life at the Dermal-Epidermal Junction Zone: Implications for Mechanisms and Therapy of Genodermatoses. J. Investig. Dermatol. 2016, 136, 1116–1123.
  62. Shapiro, S.D.; Endicott, S.K.; Province, M.A.; Pierce, J.A.; Campbell, E.J. Marked longevity of human lung parenchymal elastic fibers deduced from prevalence of D-aspartate and nuclear weapons-related radiocarbon. J. Clin. Investig. 1991, 87, 1828–1834.
  63. Yanagisawa, H.; Wagenseil, J. Elastic fibers and biomechanics of the aorta: Insights from mouse studies. Matrix Biol. 2020, 85–86, 160–172.
  64. Yanagisawa, H.; Davis, E.C. Unraveling the mechanism of elastic fiber assembly: The roles of short fibulins. Int. J. Biochem. Cell Biol. 2010, 42, 1084–1093.
  65. Iozzo, R.V.; Schaefer, L. Proteoglycan form and function: A comprehensive nomenclature of proteoglycans. Matrix Biol. 2015, 42, 11–55.
  66. Perrimon, N.; Bernfield, M. Cellular functions of proteoglycans—An overview. Semin. Cell Dev. Biol. 2001, 12, 65–67.
  67. Hardingham, T.E.; Fosang, A.J. Proteoglycans: Many forms and many functions. FASEB J. 1992, 6, 861–870.
  68. Rozario, T.; DeSimone, D.W. The Extracellular Matrix In Development and Morphogenesis: A Dynamic View. Dev. Biol. 2010, 341, 126.
  69. Lindahl, U. A personal voyage through the proteoglycan field. Matrix Biol. 2014, 35, 3–7.
  70. Vynios, D.H.; Karamanos, N.K.; Tsiganos, C.P. Advances in analysis of glycosaminoglycans: Its application for the assessment of physiological and pathological states of connective tissues. J. Chromatogr. B Anal. Technol. Biomed. Life Sci. 2002, 781, 21–38.
  71. Kjellén, L.; Lindahl, U. Proteoglycans: Structures and interactions. Annu. Rev. Biochem. 1991, 60, 443–475.
  72. Gandhi, N.S.; Mancera, R.L. The structure of glycosaminoglycans and their interactions with proteins. Chem. Biol. Drug Des. 2008, 72, 455–482.
  73. Ma, J.; Cai, H.; Long, X.; Cheng, K.; Xu, X.; Zhang, D.; Li, J. Hyaluronic acid bioinspired polymers for the regulation of cell chondrogenic and osteogenic differentiation. Int. J. Biol. Macromol. 2020, 161, 1011–1020.
  74. Ghiselli, G. Drug-Mediated Regulation of Glycosaminoglycan Biosynthesis. Med. Res. Rev. 2017, 37, 1051–1094.
  75. Hardingham, T.E.; Ewins, R.J.F.; Muir, H. Cartilage proteoglycans. Structure and heterogeneity of the protein core and the effects of specific protein modifications on the binding to hyaluronate. Biochem. J. 1976, 157, 127–143.
  76. Hascall, V.C. Interaction of cartilage proteoglycans with hyaluronic acid. J. Supramol. Cell. Biochem. 1977, 7, 101–120.
  77. Rosenberg, L.; Hellmann, W.; Kleinschmidt, A.K. Electron microscopic studies of proteoglycan aggregates from bovine articular cartilage. J. Biol. Chem. 1975, 250, 1877–1883.
  78. Kobayashi, T.; Chanmee, T.; Itano, N. Hyaluronan: Metabolism and function. Biomolecules 2020, 10, 1525.
  79. Senbanjo, L.T.; Chellaiah, M.A. CD44: A multifunctional cell surface adhesion receptor is a regulator of progression and metastasis of cancer cells. Front. Cell Dev. Biol. 2017, 5, 18.
  80. Xu, D.; Esko, J.D. Demystifying heparan sulfate-protein interactions. Annu. Rev. Biochem. 2014, 83, 129–157.
  81. Dudhia, J. Aggrecan, aging and assembly in articular cartilage. Cell. Mol. Life Sci. 2005, 62, 2241–2256.
  82. Walimbe, T.; Panitch, A. Proteoglycans in biomedicine: Resurgence of an underexploited class of ECM molecules. Front. Pharmacol. 2020, 10, 1661.
  83. Elfenbein, A.; Simons, M. Syndecan-4 signaling at a glance. J. Cell Sci. 2013, 126, 3799–3804.
  84. Choi, Y.; Chung, H.; Jung, H.; Couchman, J.R.; Oh, E.S. Syndecans as cell surface receptors: Unique structure equates with functional diversity. Matrix Biol. 2011, 30, 93–99.
  85. Kolset, S.O.; Tveit, H. Serglycin-Structure and biology. Cell. Mol. Life Sci. 2008, 65, 1073–1085.
  86. Henningsson, F.; Hergeth, S.; Cortelius, R.; Åbrink, M.; Pejler, G. A role for serglycin proteoglycan in granular retention and processing of mast cell secretory granule components. FEBS J. 2006, 273, 4901–4912.
  87. Åbrink, M.; Grujic, M.; Pejler, G. Serglycin is essential for maturation of mast cell secretory granule. J. Biol. Chem. 2004, 279, 40897–40905.
  88. Whitelock, J.M.; Graham, L.D.; Melrose, J.; Murdoch, A.D.; Iozzo, R.V.; Anne Underwood, P. Human perlecan immunopurified from different endothelial cell sources has different adhesive properties for vascular cells. Matrix Biol. 1999, 18, 163–178.
  89. Sher, I.; Zisman-Rozen, S.; Eliahu, L.; Whitelock, J.M.; Maas-Szabowski, N.; Yamada, Y.; Breitkreutz, D.; Fusenig, N.E.; Arikawa-Hirasawa, E.; Iozzo, R.V.; et al. Targeting perlecan in human keratinocytes reveals novel roles for perlecan in epidermal formation. J. Biol. Chem. 2006, 281, 5178–5187.
  90. Nugent, M.A.; Nugent, H.M.; Iozzo, R.V.; Sanchack, K.; Edelman, E.R. Perlecan is required to inhibit thrombosis after deep vascular injury and contributes to endothelial cell-mediated inhibition of intimal hyperplasia. Proc. Natl. Acad. Sci. USA 2000, 97, 6722–6727.
  91. Iozzo, R.V. Basement membrane proteoglycans: From cellar to ceiling. Nat. Rev. Mol. Cell Biol. 2005, 6, 646–656.
  92. Bezakova, G.; Ruegg, M.A. New insights into the roles of agrin. Nat. Rev. Mol. Cell Biol. 2003, 4, 295–308.
  93. Zhang, P.; Yang, L.; Li, G.; Jin, Y.; Wu, D.; Wang, Q.M.; Huang, P. Agrin Involvement in Synaptogenesis Induced by Exercise in a Rat Model of Experimental Stroke. Neurorehabil. Neural Repair 2020, 34, 1124–1137.
  94. Jan, A.T.; Lee, E.J.; Choi, I. Fibromodulin: A regulatory molecule maintaining cellular architecture for normal cellular function. Int. J. Biochem. Cell Biol. 2016, 80, 66–70.
  95. Ezura, Y.; Chakravarti, S.; Oldberg, A.; Chervoneva, I.; Birk, D.E. Differential expression of lumican and fibromodulin regulate collagen fibrillogenesis in developing mouse tendons. J. Cell Biol. 2000, 151, 779–787.
  96. Muncie, J.M.; Weaver, V.M. The Physical and Biochemical Properties of the Extracellular Matrix Regulate Cell Fate. In Current Topics in Developmental Biology; Academic Press Inc.: Amsterdam, The Netherlands, 2018; Volume 130, pp. 1–37.
  97. Davis, B.G. Synthesis of glycoproteins. Chem. Rev. 2002, 102, 579–601.
  98. Kornfeld, R.; Kornfeld, S. Comparative aspects of glycoprotein structure. Annu. Rev. Biochem. 1976, 45, 217–237.
  99. Mouw, J.K.; Ou, G.; Weaver, V.M. Extracellular matrix assembly: A multiscale deconstruction. Nat. Rev. Mol. Cell Biol. 2014, 15, 771–785.
  100. Mecham, R.P. Overview of extracellular matrix. Curr. Protoc. Cell Biol. 2012, 57, 10.
  101. Dwek, R.A. Glycobiology: Toward understanding the function of sugars. Chem. Rev. 1996, 96, 683–720.
  102. Preissner, K.T.; Reuning, U. Vitronectin in vascular context: Facets of a multitalented matricellular protein. Semin. Thromb. Hemost. 2011, 37, 408–424.
  103. Weeterings, C.; Adelmeijer, J.; Myles, T.; De Groot, P.G.; Lisman, T. Glycoprotein Ibα-mediated platelet adhesion and aggregation to immobilized thrombin under conditions of flow. Arterioscler. Thromb. Vasc. Biol. 2006, 26, 670–675.
  104. Brockhausen, I.; Kuhns, W. Role of Glycoproteins of the Immune and Blood Coagulation Systems; Springer: Berlin/Heidelberg, Germany, 1997; pp. 77–84.
  105. Rudd, P.M.; Elliott, T.; Cresswell, P.; Wilson, I.A.; Dwek, R.A. Glycosylation and the immune system. Science 2001, 291, 2370–2376.
  106. Jamieson, J.C.; Kaplan, H.A.; Woloski, B.M.R.N.J. Glycoprotein biosynthesis during the acute-phase response to inflammation. Can. J. Biochem. Cell Biol. 1983, 61, 1041–1048.
  107. Gupta, S.K. Role of zona pellucida glycoproteins during fertilization in humans. J. Reprod. Immunol. 2015, 108, 90–97.
  108. Timpl, R.; Sasaki, T.; Kostka, G.; Chu, M.L. Fibulins: A versatile family of extracellular matrix proteins. Nat. Rev. Mol. Cell Biol. 2003, 4, 479–489.
  109. Chiquet-Ehrismann, R.; Tucker, R.P. Tenascins and the importance of adhesion modulation. Cold Spring Harb. Perspect. Biol. 2011, 3, a004960.
  110. Weisel, J.W. Fibrinogen and fibrin. Adv. Protein Chem. 2005, 70, 247–299.
  111. Leavesley, D.I.; Kashyap, A.S.; Croll, T.; Sivaramakrishnan, M.; Shokoohmand, A.; Hollier, B.G.; Upton, Z. Vitronectin-Master controller or micromanager? IUBMB Life 2013, 65, 807–818.
  112. Ganss, B.; Kim, R.H.; Sodek, J. Bone sialoprotein. Crit. Rev. Oral Biol. Med. 1999, 10, 79–98.
  113. Fatemi, S.H. Reelin glycoprotein: Structure, biology and roles in health and disease. Mol. Psychiatry 2005, 10, 251–257.
  114. Pankov, R.; Yamada, K.M. Fibronectin at a glance. J. Cell Sci. 2002, 115, 3861–3863.
  115. Singh, P.; Carraher, C.; Schwarzbauer, J.E. Assembly of fibronectin extracellular matrix. Annu. Rev. Cell Dev. Biol. 2010, 26, 397–419.
  116. Baron, M.; Norman, D.; Willis, A.; Campbell, I.D. Structure of the fibronectin type 1 module. Nature 1990, 345, 642–646.
  117. Potts, J.R.; Campbell, I.D. Fibronectin structure and assembly. Curr. Opin. Cell Biol. 1994, 6, 648–655.
  118. McDonald, J.A.; Kelley, D.G.; Broekelmann, T.J. Role of fibronectin in collagen deposition: Fab’ to the gelatin-binding domain of fibronectin inhibits both fibronectin and collagen organization in fibroblast extracellular matrix. J. Cell Biol. 1982, 92, 485–492.
  119. Tamkun, J.W.; DeSimone, D.W.; Fonda, D.; Patel, R.S.; Buck, C.; Horwitz, A.F.; Hynes, R.O. Structure of integrin, a glycoprotein involved in the transmembrane linkage between fibronectin and actin. Cell 1986, 46, 271–282.
  120. Mao, Y.; Schwarzbauer, J.E. Fibronectin fibrillogenesis, a cell-mediated matrix assembly process. Matrix Biol. 2005, 24, 389–399.
  121. Sekiguchi, K.; Hakomori, S. Functional domain structure of fibronectin. Proc. Natl. Acad. Sci. USA 1980, 77, 2661–2665.
  122. Hymes, J.P.; Klaenhammer, T.R. Stuck in the middle: Fibronectin-binding proteins in gram-positive bacteria. Front. Microbiol. 2016, 7, 1504.
  123. To, W.S.; Midwood, K.S. Plasma and cellular fibronectin: Distinct and independent functions during tissue repair. Fibrogenes. Tissue Repair 2011, 4, 21.
  124. Sottile, J.; Hocking, D.C. Fibronectin polymerization regulates the composition and stability of extracellular matrix fibrils and cell-matrix adhesions. Mol. Biol. Cell 2002, 13, 3546–3559.
  125. Sakai, T.; Johnson, K.J.; Murozono, M.; Sakai, K.; Magnuson, M.A.; Wieloch, T.; Cronberg, T.; Isshiki, A.; Erickson, H.P.; Fässler, R. Plasma fibronectin supports neuronal survival and reduces brain injury following transient focal cerebral ischemia but is not essential for skin-wound healing and hemostasis. Nat. Med. 2001, 7, 324–330.
  126. Maurer, L.M.; Ma, W.; Mosher, D.F. Dynamic structure of plasma fibronectin. Crit. Rev. Biochem. Mol. Biol. 2016, 51, 213–227.
  127. Schwarzbauer, J.E. Identification of the fibronectin sequences required for assembly of a fibrillar matrix. J. Cell Biol. 1991, 113, 1463–1473.
  128. Schwarzbauer, J.E.; Tamkun, J.W.; Lemischka, I.R.; Hynes, R.O. Three different fibronectin mRNAs arise by alternative splicing within the coding region. Cell 1983, 35, 421–431.
  129. Engel, J.; Odermatt, E.; Engel, A.; Madri, J.A.; Furthmayr, H.; Rohde, H.; Timpl, R. Shapes, domain organizations and flexibility of laminin and fibronectin, two multifunctional proteins of the extracellular matrix. J. Mol. Biol. 1981, 150, 97–120.
  130. Senyürek, I.; Kempf, W.E.; Klein, G.; Maurer, A.; Kalbacher, H.; Schäfer, L.; Wanke, I.; Christ, C.; Stevanovic, S.; Schaller, M.; et al. Processing of laminin α chains generates peptides involved in wound healing and host defense. J. Innate Immun. 2014, 6, 467–484.
  131. Aumailley, M.; Bruckner-Tuderman, L.; Carter, W.G.; Deutzmann, R.; Edgar, D.; Ekblom, P.; Engel, J.; Engvall, E.; Hohenester, E.; Jones, J.C.R.; et al. A simplified laminin nomenclature. Matrix Biol. 2005, 24, 326–332.
  132. Hohenester, E. Structural biology of laminins. Essays Biochem. 2019, 63, 285–295.
  133. Odenthal, U.; Haehn, S.; Tunggal, P.; Merkl, B.; Schomburg, D.; Frie, C.; Paulsson, M.; Smyth, N. Molecular analysis of laminin N-terminal domains mediating self-interactions. J. Biol. Chem. 2004, 279, 44504–44512.
  134. Schittny, J.C.; Yurchenco, P.D. Terminal short arm domains of basement membrane laminin are critical for its self-assembly. J. Cell Biol. 1990, 110, 825–832.
  135. Hussain, S.A.; Carafoli, F.; Hohenester, E. Determinants of laminin polymerization revealed by the structure of the α5 chain amino-terminal region. EMBO Rep. 2011, 12, 276–282.
  136. Hohenester, E.; Yurchenco, P.D. Laminins in basement membrane assembly. Cell Adhes. Migr. 2013, 7, 56–63.
  137. McKee, K.K.; Hohenester, E.; Aleksandrova, M.; Yurchenco, P.D. Organization of the laminin polymer node. Matrix Biol. 2021, 98, 49–63.
  138. Sasaki, T.; Fässler, R.; Hohenester, E. Laminin: The crux of basement membrane assembly. J. Cell Biol. 2004, 164, 959–963.
  139. Colognato, H.; Yurchenco, P.D. Form and function: The laminin family of heterotrimers. Dev. Dyn. 2000, 218, 213–234.
  140. Colognato, H.; Winkelmann, D.A.; Yurchenco, P.D. Laminin polymerization induces a receptor-cytoskeleton network. J. Cell Biol. 1999, 145, 619–631.
  141. Pöschl, E.; Schlötzer-Schrehardt, U.; Brachvogel, B.; Saito, K.; Ninomiya, Y.; Mayer, U. Collagen IV is essential for basement membrane stability but dispensable for initiation of its assembly during early development. Development 2004, 131, 1619–1628.
  142. Costell, M.; Gustafsson, E.; Aszódi, A.; Mörgelin, M.; Bloch, W.; Hunziker, E.; Addicks, K.; Timpl, R.; Fässler, R. Perlecan maintains the integrity of cartilage and some basement membranes. J. Cell Biol. 1999, 147, 1109–1122.
  143. McKee, K.K.; Aleksandrova, M.; Yurchenco, P.D. Chimeric protein identification of dystrophic, Pierson and other laminin polymerization residues. Matrix Biol. 2018, 67, 32–46.
  144. Böse, K.; Nischt, R.; Page, A.; Bader, B.L.; Paulsson, M.; Smyth, N. Loss of nidogen-1 and -2 results in syndactyly and changes in limb development. J. Biol. Chem. 2006, 281, 39620–39629.
  145. Li, S.; Liquari, P.; McKee, K.K.; Harrison, D.; Patel, R.; Lee, S.; Yurchenco, P.D. Laminin-sulfatide binding initiates basement membrane assembly and enables receptor signaling in Schwann cells and fibroblasts. J. Cell Biol. 2005, 169, 179–189.
  146. Mak, K.M.; Mei, R. Basement Membrane Type IV Collagen and Laminin: An Overview of Their Biology and Value as Fibrosis Biomarkers of Liver Disease. Anat. Rec. 2017, 300, 1371–1390.
  147. Cambi, A.; Chavrier, P. Tissue remodeling by invadosomes. Fac. Rev. 2021, 10, 39.
  148. Arpino, V.; Brock, M.; Gill, S.E. The role of TIMPs in regulation of extracellular matrix proteolysis. Matrix Biol. 2015, 44–46, 247–254.
  149. Cornfine, S.; Himmel, M.; Kopp, P.; el Azzouzi, K.; Wiesner, C.; Krüger, M.; Rudel, T.; Linder, S. The kinesin KIF9 and reggie/flotillin proteins regulate matrix degradation by macrophage podosomes. Mol. Biol. Cell 2011, 22, 202.
  150. Cawston, T.E.; Young, D.A. Proteinases involved in matrix turnover during cartilage and bone breakdown. Cell Tissue Res. 2010, 339, 221–235.
  151. Oda, K. New families of carboxyl peptidases: Serine-carboxyl peptidases and glutamic peptidases. J. Biochem. 2012, 151, 13–25.
  152. Cui, N.; Hu, M.; Khalil, R.A. Biochemical and Biological Attributes of Matrix Metalloproteinases. Prog. Mol. Biol. Transl. Sci. 2017, 147, 1.
  153. Kapoor; Vaidya, S.; Wadhwan, V.; Hitesh; Kaur, G.; Pathak, A. Seesaw of matrix metalloproteinases (MMPs). J. Cancer Res. Ther. 2016, 12, 28.
  154. Page-McCaw, A.; Ewald, A.J.; Werb, Z. Matrix metalloproteinases and the regulation of tissue remodelling. Nat. Rev. Mol. Cell Biol. 2007, 8, 221.
  155. Chen, Q.; Jin, M.; Yang, F.; Zhu, J.; Xiao, Q.; Zhang, L. Matrix metalloproteinases: Inflammatory regulators of cell behaviors in vascular formation and remodeling. Mediators Inflamm. 2013, 2013, 8315.
  156. Sternlicht, M.D.; Werb, Z. How matrix metalloproteinases regulate cell behavior. Annu. Rev. Cell Dev. Biol. 2000, 17, 463–516.
  157. Imai, K.; Hiramatsu, A.; Fukushima, D.; Pierschbacher, M.D.; Okada, Y. Degradation of decorin by matrix metalloproteinases: Identification of the cleavage sites, kinetic analyses and transforming growth factor-β1 release. Biochem. J. 1997, 322, 809–814.
  158. Giannelli, G.; Falk-Marzillier, J.; Schiraldi, O.; Stetler-Stevenson, W.G.; Quaranta, V. Induction of cell migration by matrix metalloprotease-2 cleavage of laminin-5. Science 1997, 277, 225–228.
  159. Stetler-Stevenson, W.G. Matrix metalloproteinases in angiogenesis: A moving target for therapeutic intervention. J. Clin. Investig. 1999, 103, 1237–1241.
  160. Anacker, J.; Segerer, S.E.; Hagemann, C.; Feix, S.; Kapp, M.; Bausch, R.; Kämmerer, U. Human decidua and invasive trophoblasts are rich sources of nearly all human matrix metalloproteinases. Mol. Hum. Reprod. 2011, 17, 637–652.
  161. Yushchenko, M.; Weber, F.; Mäder, M.; Schöll, U.; Maliszewska, M.; Tumani, H.; Felgenhauer, K.; Beuche, W. Matrix metalloproteinase-9 (MMP-9) in human cerebrospinal fluid (CSF): Elevated levels are primarily related to CSF cell count. J. Neuroimmunol. 2000, 110, 244–251.
  162. Hartung, H.-P.; Kieseier, B.C. The role of matrix metalloproteinases in autoimmune damage to the central and peripheral nervous system. J. Neuroimmunol. 2000, 107, 140–147.
  163. Gonzalez-Avila, G.; Sommer, B.; Mendoza-Posada, D.A.; Ramos, C.; Garcia-Hernandez, A.A.; Falfan-Valencia, R. Matrix metalloproteinases participation in the metastatic process and their diagnostic and therapeutic applications in cancer. Crit. Rev. Oncol. Hematol. 2019, 137, 57–83.
  164. Blobel, C.P. Metalloprotease-Disintegrins: Links to Cell Adhesion and Cleavage of TNFα and Notch. Cell 1997, 90, 589–592.
  165. Werb, Z. ECM and Cell Surface Proteolysis: Regulating Cellular Ecology. Cell 1997, 91, 439–442.
  166. Blobel, C.P. ADAMs: Key components in egfr signalling and development. Nat. Rev. Mol. Cell Biol. 2005, 6, 32–43.
  167. Murphy, G. The ADAMs: Signalling scissors in the tumour microenvironment. Nat. Rev. Cancer 2008, 8, 929–941.
  168. Rose-John, S. ADAM17, shedding, TACE as therapeutic targets. Pharmacol. Res. 2013, 71, 19–22.
  169. Marczynska, J.; Ozga, A.; Wlodarczyk, A.; Majchrzak-Gorecka, M.; Kulig, P.; Banas, M.; Michalczyk-Wetula, D.; Majewski, P.; Hutloff, A.; Schwarz, J.; et al. The Role of Metalloproteinase ADAM17 in Regulating ICOS Ligand–Mediated Humoral Immune Responses. J. Immunol. 2014, 193, 2753–2763.
  170. Hsia, H.E.; Tüshaus, J.; Brummer, T.; Zheng, Y.; Scilabra, S.D.; Lichtenthaler, S.F. Functions of ‘A disintegrin and metalloproteases (ADAMs)’ in the mammalian nervous system. Cell. Mol. Life Sci. 2019, 76, 3055–3081.
  171. Zunke, F.; Rose-John, S. The shedding protease ADAM17: Physiology and pathophysiology. Biochim. Biophys. Acta-Mol. Cell Res. 2017, 1864, 2059–2070.
  172. Van Goor, H.; Melenhorst, W.B.W.H.; Turner, A.J.; Holgate, S.T. Adamalysins in biology and disease. J. Pathol. 2009, 219, 277–286.
  173. Boudreau, N.J.; Jones, P.L. Extracellular matrix and integrin signalling: The shape of things to come. Biochem. J. 1999, 339, 481–488.
  174. Fan, D.; Creemers, E.E.; Kassiri, Z. Matrix as an interstitial transport system. Circ. Res. 2014, 114, 889–902.
  175. Schultz, G.S.; Wysocki, A. Interactions between extracellular matrix and growth factors in wound healing. Wound Repair Regen. 2009, 17, 153–162.
  176. Flaumenhaft, R.; Rifkin, D.B. Extracellular matrix regulation of growth factor and protease activity. Curr. Opin. Cell Biol. 1991, 3, 817–823.
  177. O’Callaghan, P.; Zhang, X.; Li, J.P. Heparan Sulfate Proteoglycans as Relays of Neuroinflammation. J. Histochem. Cytochem. 2018, 66, 305–319.
  178. Li, J.P.; Kusche-Gullberg, M. Heparan Sulfate: Biosynthesis, Structure, and Function. In International Review of Cell and Molecular Biology; Elsevier Inc.: Amsterdam, The Netherlands, 2016; Volume 325, pp. 215–273. ISBN 9780128048061.
  179. Billings, P.C.; Pacifici, M. Interactions of signaling proteins, growth factors and other proteins with heparan sulfate: Mechanisms and mysteries. Connect. Tissue Res. 2015, 56, 272–280.
  180. Matsuo, I.; Kimura-Yoshida, C. Extracellular distribution of diffusible growth factors controlled by heparan sulfate proteoglycans during mammalian embryogenesis. Philos. Trans. R. Soc. B Biol. Sci. 2014, 369, 20130545.
  181. Xu, Y.H.; Zhu, Y.; Zhu, Y.Y.; Wei, H.; Zhang, N.N.; Qin, J.S.; Zhu, X.L.; Yu, M.; Li, Y.F. Abnormalities in FGF family members and their roles in modulating depression-related molecules. Eur. J. Neurosci. 2021, 53, 140–150.
  182. Malkowski, A.; Sobolewski, K.; Jaworski, S.; Bankowski, E. FGF binding by extracellular matrix components of Wharton’s jelly. Acta Biochim. Pol. 2007, 54, 357–363.
  183. Loo, B.M.; Kreuger, J.; Jalkanen, M.; Lindahl, U.; Salmivirta, M. Binding of Heparin/Heparan Sulfate to Fibroblast Growth Factor Receptor 4. J. Biol. Chem. 2001, 276, 16868–16876.
  184. Vogler, E.A. Structure and reactivity of water at biomaterial surfaces. Adv. Colloid Interface Sci. 1998, 74, 69–117.
  185. Kasemo, B.; Gold, J. Implant surfaces and interface processes. Adv. Dent. Res. 1999, 13, 8–20.
  186. Rabe, M.; Verdes, D.; Seeger, S. Understanding protein adsorption phenomena at solid surfaces. Adv. Colloid Interface Sci. 2011, 162, 87–106.
  187. Andrade, J.D.; Hlady, V.; Wei, A.P. Adsorption of complex proteins at interfaces. Pure Appl. Chem. 1992, 64, 1777–1781.
  188. Felgueiras, H.P.; Antunes, J.C.; Martins, M.C.L.; Barbosa, M.A. Fundamentals of protein and cell interactions in biomaterials. In Peptides and Proteins as Biomaterials for Tissue Regeneration and Repair; Barbosa, M.A., Martins, M.C., Eds.; Woodhead Publishing: Thurston, UK, 2018; pp. 1–27. ISBN 9780081008522.
  189. Norde, W.; Giacomelli, C.E. Conformational changes in proteins at interfaces: From solution to the interface, and back. Macromol. Symp. 1999, 145, 125–136.
  190. Norde, W.; Giacomelli, C.E. BSA structural changes during homomolecular exchange between the adsorbed and the dissolved states. J. Biotechnol. 2000, 79, 259–268.
  191. Lundstrom, I. Models of Protein Adsorption on Solid Surfaces. Prog. Colloid Polym. Sci. 1985, 70, 76–82.
  192. Mariani, E.; Lisignoli, G.; Borzì, R.M.; Pulsatelli, L. Biomaterials: Foreign bodies or tuners for the immune response? Int. J. Mol. Sci. 2019, 20, 636.
  193. Ode Boni, B.O.; Lamboni, L.; Souho, T.; Gauthier, M.; Yang, G. Immunomodulation and cellular response to biomaterials: The overriding role of neutrophils in healing. Mater. Horizons 2019, 6, 1122–1137.
  194. Jhunjhunwala, S.; Aresta-DaSilva, S.; Tang, K.; Alvarez, D.; Webber, M.J.; Tang, B.C.; Lavin, D.M.; Veiseh, O.; Doloff, J.C.; Bose, S.; et al. Neutrophil responses to sterile implant materials. PLoS ONE 2015, 10, e0137550.
  195. Desalegn, G.; Pabst, O. Inflammation triggers immediate rather than progressive changes in monocyte differentiation in the small intestine. Nat. Commun. 2019, 10, 1–14.
  196. Mesure, L.; de Visscher, G.; Vranken, I.; Lebacq, A.; Flameng, W. Gene expression study of monocytes/macrophages during early foreign body reaction and identification of potential precursors of myofibroblasts. PLoS ONE 2010, 5, e12949.
  197. Christenson, E.M.; Dadsetan, M.; Anderson, J.M.; Hiltner, A. Biostability and macrophage-mediated foreign body reaction of silicone-modified polyurethanes. J. Biomed. Mater. Res. Part A 2005, 74, 141–155.
  198. Sheikh, Z.; Brooks, P.J.; Barzilay, O.; Fine, N.; Glogauer, M. Macrophages, foreign body giant cells and their response to implantable biomaterials. Materials 2015, 8, 5671–5701.
  199. Ward, W.K.; Slobodzian, E.P.; Tiekotter, K.L.; Wood, M.D. The effect of microgeometry, implant thickness and polyurethane chemistry on the foreign body response to subcutaneous implants. Biomaterials 2002, 23, 4185–4192.
  200. Hulbert, S.F.; Morrison, S.J.; Klawitter, J.J. Tissue reaction to three ceramics of porous and non-porous structures. J. Biomed. Mater. Res. 1972, 6, 347–374.
  201. Zdolsek, J.; Eaton, J.W.; Tang, L. Histamine release and fibrinogen adsorption mediate acute inflammatory responses to biomaterial implants in humans. J. Transl. Med. 2007, 5, 31.
  202. Theoharides, T.C.; Tsilioni, I.; Conti, P. Mast cells may regulate the anti-inflammatory activity of IL-37. Int. J. Mol. Sci. 2019, 20, 3701.
  203. Thevenot, P.T.; Nair, A.M.; Shen, J.; Lotfi, P.; Ko, C.Y.; Tang, L. The effect of incorporation of SDF-1α into PLGA scaffolds on stem cell recruitment and the inflammatory response. Biomaterials 2010, 31, 3997–4008.
  204. Zachman, A.L.; Crowder, S.W.; Ortiz, O.; Zienkiewicz, K.J.; Bronikowski, C.M.; Yu, S.S.; Giorgio, T.D.; Guelcher, S.A.; Kohn, J.; Sung, H.J. Pro-angiogenic and anti-inflammatory regulation by functional peptides loaded in polymeric implants for soft tissue regeneration. Tissue Eng.-Part A 2013, 19, 437–447.
  205. Hosoyama, K.; Ahumada, M.; Goel, K.; Ruel, M.; Suuronen, E.J.; Alarcon, E.I. Electroconductive materials as biomimetic platforms for tissue regeneration. Biotechnol. Adv. 2019, 37, 444–458.
  206. Mason, T.O.; Shimanovich, U. Fibrous Protein Self-Assembly in Biomimetic Materials. Adv. Mater. 2018, 30, 1706462.
  207. Navarro, M.; Michiardi, A.; Castaño, O.; Planell, J.A. Biomaterials in orthopaedics. J. R. Soc. Interface 2008, 5, 1137–1158.
  208. Xu, L.C.; Siedlecki, C.A. Effects of surface wettability and contact time on protein adhesion to biomaterial surfaces. Biomaterials 2007, 28, 3273–3283.
  209. Parhi, P.; Golas, A.; Barnthip, N.; Noh, H.; Vogler, E.A. Volumetric interpretation of protein adsorption: Capacity scaling with adsorbate molecular weight and adsorbent surface energy. Biomaterials 2009, 30, 6814–6824.
  210. Thevenot, P.; Hu, W.; Tang, L. Surface chemistry influences implant biocompatibility. Curr. Top. Med. Chem. 2008, 8, 270–280.
  211. Roach, P.; Farrar, D.; Perry, C.C. Interpretation of protein adsorption: Surface-induced conformational changes. J. Am. Chem. Soc. 2005, 127, 8168–8173.
  212. Roach, P.; Parker, T.; Gadegaard, N.; Alexander, M.R. Surface strategies for control of neuronal cell adhesion: A review. Surf. Sci. Rep. 2010, 65, 145–173.
  213. Chen, S.; Guo, Y.; Liu, R.; Wu, S.; Fang, J.; Huang, B.; Li, Z.; Chen, Z.; Chen, Z. Tuning surface properties of bone biomaterials to manipulate osteoblastic cell adhesion and the signaling pathways for the enhancement of early osseointegration. Colloids Surfaces B Biointerfaces 2018, 164, 58–69.
  214. Zelzer, M.; Albutt, D.; Alexander, M.R.; Russell, N.A. The Role of Albumin and Fibronectin in the Adhesion of Fibroblasts to Plasma Polymer Surfaces. Plasma Process. Polym. 2012, 9, 149–156.
  215. Guo, B.; Anzai, J.I.; Osa, T. Adsorption behavior of serum albumin on electrode surfaces and the effects of electrode potential. Chem. Pharm. Bull. 1996, 44, 800–803.
  216. Faucheux, N.; Schweiss, R.; Lützow, K.; Werner, C.; Groth, T. Self-assembled monolayers with different terminating groups as model substrates for cell adhesion studies. Biomaterials 2004, 25, 2721–2730.
  217. Keselowsky, B.G.; Collard, D.M.; García, A.J. Integrin binding specificity regulates biomaterial surface chemistry effects on cell differentiation. Proc. Natl. Acad. Sci. USA 2005, 102, 5953–5957.
  218. Lan, M.A.; Gersbach, C.A.; Michael, K.E.; Keselowsky, B.G.; García, A.J. Myoblast proliferation and differentiation on fibronectin-coated self assembled monolayers presenting different surface chemistries. Biomaterials 2005, 26, 4523–4531.
  219. McClary, K.B.; Ugarova, T.; Grainger, D.W. Modulating fibroblast adhesion, spreading, and proliferation using self- assembled monolayer films of alkylthiolates on gold. J. Biomed. Mater. Res. 2000, 50, 428–439.
  220. Aiyelabegan, H.T.; Sadroddiny, E. Fundamentals of protein and cell interactions in biomaterials. Biomed. Pharmacother. 2017, 88, 956–970.
  221. Ruoslahti, E.; Pierschbacher, M.D. Arg-Gly-Asp: A versatile cell recognition signal. Cell 1986, 44, 517–518.
  222. Karimi, F.; O’Connor, A.J.; Qiao, G.G.; Heath, D.E. Integrin Clustering Matters: A Review of Biomaterials Functionalized with Multivalent Integrin-Binding Ligands to Improve Cell Adhesion, Migration, Differentiation, Angiogenesis, and Biomedical Device Integration. Adv. Healthc. Mater. 2018, 7, 1701324.
  223. Reyes, C.D.; García, A.J. Engineering integrin-specific surfaces with a triple-helical collagen-mimetic peptide. J. Biomed. Mater. Res.-Part A 2003, 65, 511–523.
  224. Nomizu, M.; Weeks, B.S.; Weston, C.A.; Kim, W.H.; Kleinman, H.K.; Yamada, Y. Structure-activity study of a laminin α1 chain active peptide segment Ile-Lys-Val-Ala-Val (IKVAV). FEBS Lett. 1995, 365, 227–231.
  225. Yin, Y.; Wang, W.; Shao, Q.; Li, B.; Yu, D.; Zhou, X.; Parajuli, J.; Xu, H.; Qiu, T.; Yetisen, A.K.; et al. Pentapeptide IKVAV-engineered hydrogels for neural stem cell attachment. Biomater. Sci. 2021, 9, 2887–2892.
  226. Boateng, S.Y.; Lateef, S.S.; Mosley, W.; Hartman, T.J.; Hanley, L.; Russell, B. RGD and YIGSR synthetic peptides facilitate cellular adhesion identical to that of laminin and fibronectin but alter the physiology of neonatal cardiac myocytes. Am. J. Physiol.-Cell Physiol. 2005, 288, 30–38.
  227. Aota, S.I.; Nomizu, M.; Yamada, K.M. The short amino acid sequence Pro-His-Ser-Arg-Asn in human fibronectin enhances cell-adhesive function. J. Biol. Chem. 1994, 269, 24756–24761.
  228. Cutler, S.M.; García, A.J. Engineering cell adhesive surfaces that direct integrin α5β1 binding using a recombinant fragment of fibronectin. Biomaterials 2003, 24, 1759–1770.
  229. Lebaron, R.G.; Athanasiou, K.A. Extracellular matrix cell adhesion peptides: Functional applications in orthopedic materials. Tissue Eng. 2000, 6, 85–103.
  230. Song, W.; Mano, J.F. Interactions between cells or proteins and surfaces exhibiting extreme wettabilities. Soft Matter 2013, 9, 2985–2999.
  231. Siebers, M.C.; Ter Brugge, P.J.; Walboomers, X.F.; Jansen, J.A. Integrins as linker proteins between osteoblasts and bone replacing materials. A critical review. Biomaterials 2005, 26, 137–146.
  232. Ponche, A.; Bigerelle, M.; Anselme, K. Relative influence of surface topography and surface chemistry on cell response to bone implant materials. Part 1: Physico-chemical effects. J. Eng. Med. 2010, 224, 1471–1486.
  233. Ayala, P.; Desai, T.A. Integrin α3 blockade enhances microtopographical down-regulation of α-smooth muscle actin: Role of microtopography in ECM regulation. Integr. Biol. 2011, 3, 733–741.
  234. Jäger, M.; Zilkens, C.; Zanger, K.; Krauspe, R. Significance of nano- and microtopography for cell-surface interactions in orthopaedic implants. J. Biomed. Biotechnol. 2007, 2007, 69036.
  235. Hulander, M.; Lundgren, A.; Berglin, M.; Ohrlander, M.; Lausmaa, J.; Elwing, H. Immune complement activation is attenuated by surface nanotopography. Int. J. Nanomed. 2011, 6, 2653–2666.
  236. Draghi, L.; Cigada, A. Nanostructured surfaces for biomedical applications. Part I: Nanotopography. J. Appl. Biomater. Biomech. 2007, 5, 61–69.
  237. Folch, A.; Toner, M. Microengineering of cellular interactions. Annu. Rev. Biomed. Eng. 2000, 2, 227–256.
  238. Poellmann, M.J.; Harrell, P.A.; King, W.P.; Johnson, A.J.W. Geometric microenvironment directs cell morphology on topographically patterned hydrogel substrates. Acta Biomater. 2010, 6, 3514–3523.
  239. Solanki, A.; Shah, S.; Memoli, K.A.; Park, S.Y.; Hong, S.; Lee, K.B. Controlling differentiation of neural stem cells using extracellular matrix protein patterns. Small 2010, 6, 2509–2513.
  240. Carthew, J.; Abdelmaksoud, H.H.; Hodgson-Garms, M.; Aslanoglou, S.; Ghavamian, S.; Elnathan, R.; Spatz, J.P.; Brugger, J.; Thissen, H.; Voelcker, N.H.; et al. Precision Surface Microtopography Regulates Cell Fate via Changes to Actomyosin Contractility and Nuclear Architecture. Adv. Sci. 2021, 8, 2003186.
  241. Anselme, K.; Ploux, L.; Ponche, A. Cell/material interfaces: Influence of surface chemistry and surface topography on cell adhesion. J. Adhes. Sci. Technol. 2010, 24, 831–852.
  242. Majhy, B.; Priyadarshini, P.; Sen, A.K. Effect of surface energy and roughness on cell adhesion and growth-facile surface modification for enhanced cell culture. RSC Adv. 2021, 11, 15467–15476.
  243. Hallab, N.J.; Bundy, K.J.; O’Connor, K.; Moses, R.L.; Jacobs, J.J. Evaluation of metallic and polymeric biomaterial surface energy and surface roughness characteristics for directed cell adhesion. Tissue Eng. 2001, 7, 55–70.
  244. Hou, Y.; Xie, W.; Yu, L.; Camacho, L.C.; Nie, C.; Zhang, M.; Haag, R.; Wei, Q. Surface Roughness Gradients Reveal Topography-Specific Mechanosensitive Responses in Human Mesenchymal Stem Cells. Small 2020, 16, 1905422.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/)
ScholarVision Creations
Feedback