Next Article in Journal
Cell Growth on Different Types of Ultrananocrystalline Diamond Thin Films
Next Article in Special Issue
Biocompatibility of Chitosan Carriers with Application in Drug Delivery
Previous Article in Journal
Platelet-Rich Plasma Favors Proliferation of Canine Adipose-Derived Mesenchymal Stem Cells in Methacrylate-Endcapped Caprolactone Porous Scaffold Niches
Previous Article in Special Issue
Silica as a Matrix for Encapsulating Proteins: Surface Effects on Protein Structure Assessed by Circular Dichroism Spectroscopy
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
JFB
Volume 3
Issue 3
10.3390/jfb3030569
jfb-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

Extracellular Matrix Molecules Facilitating Vascular Biointegration

by
Steven G. Wise
1,
Anna Waterhouse
2,
Praveesuda Michael
1 and
Martin K.C. Ng
1,*
1
The Heart Research Institute, Eliza Street, Newtown, NSW 2042, Australia
2
Wyss Institute for Biologically Inspired Engineering at Harvard, Boston, MA 02115, USA
*
Author to whom correspondence should be addressed.
J. Funct. Biomater. 2012, 3(3), 569-587; https://doi.org/10.3390/jfb3030569
Submission received: 29 June 2012 / Revised: 1 August 2012 / Accepted: 6 August 2012 / Published: 14 August 2012
(This article belongs to the Special Issue Biocompatibility of Biomaterials)
Browse Figures
Versions Notes

Abstract

:
All vascular implants, including stents, heart valves and graft materials exhibit suboptimal biocompatibility that significantly reduces their clinical efficacy. A range of biomolecules in the subendothelial space have been shown to play critical roles in local regulation of thrombosis, endothelial growth and smooth muscle cell proliferation, making these attractive candidates for modulation of vascular device biointegration. However, classically used biomaterial coatings, such as fibronectin and laminin, modulate only one of these components; enhancing endothelial cell attachment, but also activating platelets and triggering thrombosis. This review examines a subset of extracellular matrix molecules that have demonstrated multi-faceted vascular compatibility and accordingly are promising candidates to improve the biointegration of vascular biomaterials.

1. Introduction

With advances in technology, implantable cardiovascular devices such as grafts, stents, pacemakers and heart valves are playing an increasingly important role in modern cardiovascular medicine. Despite this, biocompatible and clinically effective synthetic materials for cardiovascular devices are currently lacking. Current artificial grafts are predominantly made of either Dacron or expanded polytetrafluoroethylene (ePTFE). These artificial materials lack sufficient elasticity, have poor interactions with vascular cells, are highly thrombogenic and induce chronic inflammation [1].Re-endothelialization of graft surfaces is retarded by the highly hydrophobic surfaces and smooth muscle cells proliferate in response to the inflammatory cascade. Accordingly, current synthetic conduit materials fail uniformly in the treatment of peripheral vascular disease within 5 years due to thrombosis and uncontrolled cellular infiltration [2,3]. There remains a strong demand for novel grafts due to these and other shortcomings.
The problem of innate incompatibility extends beyond graft materials to other vascular devices such as heart valves, pacemaker components and stents. These devices are primarily made of metal alloys, which are highly thrombogenic and induce inflammation at the site of implantation. The thrombogenicity of metal alloys are highlighted by the need for lifelong anticoagulation in metal heart valve recipients [4] and by the worrying propensity for coronary stents to thrombose. While the incidence of stent thrombosis for bare metal platforms is relatively low when used with antiplatelet therapy, such an event is catastrophic, associated with mortality rates of 25% [5]. Drug eluting stents (DES) releasing anti-proliferative agents are highly effective in inhibiting neointimal hyperplasia but also profoundly delay healing and re-endothelialization at the stent deployment site [6]. Consequently, the safety of DES has come into question [7]. DES are not only susceptible to early thrombotic events like BMS, but are also prone to both late (30 days–1 year) and very late (>1 year) stent thrombosis [8]. In stable patients with single vessel disease, late stent thrombosis occurs at a constant rate of 0.6% per year with no indication that this abates in the long term [9]. However, in real world studies incorporating acute coronary syndromes and complex interventions, even higher rates have been reported (0.9–3%/year) [10]. Additionally, DES polymer instability has been associated with chronic inflammatory and hypersensitivity reactions [11].
As a consequence of the lack of biocompatible synthetic materials, a key objective of vascular bioengineering is the development of new biomaterials with appropriate biological properties that mimic that of human arteries. The commonality of vascular implants is the requirement to simultaneously modulate multiple biological processes (Figure 1). Injured and diseased vasculature routinely has damaged and compromised endothelium, while the inflammatory response triggers hyper proliferation of smooth muscle cells (SMCs). In this environment, clotting cascade factors are activated and thrombosis is easily initiated. An optimal medical device will encourage rapid regeneration of the endothelium; inhibit SMC proliferation and migration while having high blood compatibility.
Jfb 03 00569 g001 1024
Figure 1. The multiple facets of vascular biointegration: (a) A healthy vessel is characterized by a complete monolayer of endothelial cells (ECs), quiescent smooth muscle cells (SMCs) and hemostasis; (b) In disease, the endothelium is compromised, SMCs hyper-proliferate and the clotting cascade is triggered; (c) An ideal implanted device will simultaneously enhance EC regrowth, block SMC proliferation and inhibit thrombus formation.
Figure 1. The multiple facets of vascular biointegration: (a) A healthy vessel is characterized by a complete monolayer of endothelial cells (ECs), quiescent smooth muscle cells (SMCs) and hemostasis; (b) In disease, the endothelium is compromised, SMCs hyper-proliferate and the clotting cascade is triggered; (c) An ideal implanted device will simultaneously enhance EC regrowth, block SMC proliferation and inhibit thrombus formation.
Jfb 03 00569 g001

1.1. Vascular Biomimicry

Classically, materials exhibiting biocompatibility perform their function without eliciting an undue host response or resulting in adverse clinical outcomes [12]. However, this paradigm has shifted recently beyond inertness, requiring biocompatible materials to facilitate beneficial cell and tissue interactions. In this context, the design of new vascular devices should aim to promote rapid and appropriate healing. A balance needs to be reached between low thrombogenicity, rapid re-endothelialization and arresting the infiltration and proliferation of SMC’s. An ideal vascular implant would be non-thrombogenic even when in contact with the circulation and in the absence of platelet inhibition.
Biomimetics or imitation of natural methods and processes has been identified as a means of significantly improving the biocompatibility of vascular materials. Biomimicry strategies in the context of vascular devices have been diverse including: whole organ tissue scaffolds [13], host derived vascular grafts [14] and biomolecule coatings with cell interactive characteristics [15]. Previous attempts to develop bioactive coatings for vascular implants have been limited by overly simplistic approaches, often focusing on modulating one facet of biocompatibility (e.g., endothelialization alone) at the expense of others. For example, despite having favorable endothelial cell interactions, collagen, fibronectin and laminin are thrombogenic [16,17,18]. Additionally, vitronectin, collagen (III and IV) and fibronectin promote smooth muscle cell migration [19,20]. Peptides mimicking cell-binding motifs in fibronectin (RGD) and laminin (YIGSR) are also widely used, but suffer the same shortcomings as their parent proteins, failing to achieve multi-faceted effect [21,22]. Similarly, phosphorylcholine (PC), based on a naturally occurring cell membrane lipid, was observed in vitro to be non-thrombogenic when used as a stent coating [23]. In vivo, PC failed to encourage endothelialization and ultimately had no effect on the rate of stent thrombosis [24]. Overall, a successful biomimicry approach will extend beyond emulation of a single desirable vascular property and more broadly seek to address a balance between favorable cell interactions and reducing thrombogenicity.

1.2. Candidate Biomolecules from the Vessel Wall

A range of biomolecules involved in the regulation of thrombosis and cell signaling are found in the vessel wall. The extracellular matrix (ECM), a complex collection of diverse proteins that form an organized network to which cells adhere [25], comprises the underlying structural component of vessels. The ECM modulates cell behavior [26] and signaling [27] and is responsible for the structural support of tissues. The ECM of the vasculature is primarily composed of collagens for structure, elastin for recoil, various glycoproteins (including microfibrils) for adhesion and proteoglycans and glycosaminoglycans, which provide many diverse functions.
Elastic fibers are major ECM assemblies that provide elasticity and resilience. These contain primarily elastin, in combination with fibrillin-1 and co-localized with fibulin-5 [28]. Both fibrillin-1 and fibulin-5 enhance endothelial cell stability and inhibit SMC migration [29]. The elastin rich internal elastic lamina is also a notable structural feature of large arteries, critical in preventing the infiltration of SMCs. The basement membrane, recognized as an essential mediator of cell homeostasis, provides structural support to endothelial cells and regulates their behavior. This sheet-like structure contains numerous proteins including type IV collagen, laminins, small glycoproteins such as nidogen/entactin, and heparin sulphate proteoglycans such as perlecan [30]. Here, perlecan is of particular interest as it is implicated in cell binding and proliferation, growth factor regulation [31] and the inhibition of thrombosis [32].
Despite the wide variety of proteins present in the extracellular matrix and basement membrane, many have features that are not desirable for use as a biomaterial coating, as discussed for collagen and fibronectin above. The ideal candidate biomolecule would enhance healing and endothelialization, reduce smooth muscle cell hyperproliferation caused by inflammation and be non-thrombogenic. Overall, there is a subset of biologically relevant candidates with the potential to improve implant biointegration. These include elastin, fibrillin-1, fibulin-5 and perlecan (Figure 2), which have all been shown to play important development or regulatory roles in the vasculature, while simultaneously influencing re-endothelialization, inhibiting smooth muscle cells and deterring thrombosis (Figure 2). We discuss these four promising candidates in further detail below.
Jfb 03 00569 g002 1024
Figure 2. Candidate biomolecules and their relative locations in the vessel wall, highlighting their proximity to ECs, SMCs and the vessel lumen.
Figure 2. Candidate biomolecules and their relative locations in the vessel wall, highlighting their proximity to ECs, SMCs and the vessel lumen.
Jfb 03 00569 g002

2. Candidate Biomolecules

2.1. Fibrillin-1

Microfibrils are an important component of elastic fibers, providing both mechanical and cell interactive properties. Across tissue types it is proposed that microfibrils are present at the beginning of elastogenesis and act as a scaffold for tropoelastin, the soluble precursor of elastin, deposition and assembly [33]. They are adjacent to tropoelastin producing cells and parallel to the developing fiber [34]. While the specifics of their role in assembly are not known, microfibrils have been described as having a bead-on-a-string appearance [35], while there is also evidence of small filaments that span between the adjacent microfibrils [36] suggesting some lateral assembly interaction. Microfibrils are primarily composed of glycoproteins from the fibrillin family and microfibril-associated glycoprotein (MAGP)-1 and -2. Fibrillin-1 and -2 have so far been implicated as key microfibrillar components, both are found extracellularly, with different but coincident patterns of expression. Of the three fibrillins so far identified, fibrillin-1 has particular importance to the cardiovascular system, while fibrillin-2 and -3 are highly expressed in lung tissue and cartilage, though with differential expression patterns [37].
Fibrillin-1 is a 350 kDa cysteine-rich glycoprotein which has been shown to be particularly important in elastic fiber development [38] and wound repair [39]. It contains 47 epidermal growth factor (EGF) domains, of which 43 are calcium binding (cbEGF)-like domains, and 7 eight-cysteine-containing TB motifs. It is currently theorized that tropoelastin deposition on the microfibril is directed toward the inter-bead region and corresponding concentrations of fibrillin-1 [40], implying an important signaling role in elastic fiber assembly. It interacts strongly with cells via a canonical arginine-glycine-aspartic acid (RGD) motif, as well as with other ECM proteins including decorin, versican and tropoelastin [41].
The importance of fibrillin-1 to the vasculature is demonstrated by autosomal dominant mutations in the fibrillin-1 gene known to cause Marfan Syndrome (MFS) [42]. This disorder has significant skeletal and pulmonary complications, but manifests in the vasculature as aortic aneurysm and weakened heart valves, linked to reduced microfibrillar integrity and vascular mechanics. Marfan patients also exhibit EC dysfunction implying a role for fibrillin-1 in anchoring and signaling to this cell type. Fibrillin-1 knockout mice, used to model MFS also demonstrate progressive fragmentation of their aortic elastic lamellae, fragmentation of microfibrils in other tissues and EC detachment, providing further evidence for its importance [43]. This disease state also points to an important biological role for fibrillin-1 in regulating the bioavailability of transforming growth factor beta (TGF-β), a critically import cell regulatory molecule [44].
Studying the biological effects of fibrillin-1 in vitro has benefited from a molecular biology approach, recombinantly producing discreet domain segments. Details of this approach and the fragment nomenclature have been previously published [45]. Fibrillin-1 constructs spanning the full-length protein have assisted in elucidating important cell interactive regions, such as the RGD present in PF8/PF9 [45], as well as the contribution of surrounding stabilizing domains [46].
Fibrillin-1 binds to a range of cell types including endothelial cells, smooth muscle cells and fibroblasts. This interaction is mediated by αvβ3 and α5β1 integrins binding to the single RGD motif found in fibrillin-1 [47]. Polar residues surround the RGD sequence, making it highly likely to be solvent exposed and accessible for binding interactions. Fibrillin-1 fragments containing the RGD sequence, such as PF8 and PF9 (and constructs such as PF14 which combine PF8 and PF9) are suggested to be responsible for promoting endothelial cell proliferation via this mechanism. However, the binding interaction is complex, requiring contributions from upstream cbEGF domains to position the RGD sequence to achieve full adhesion and signaling [46]. This is exemplified in the differential binding behavior seen between PF9 (RGD only) and PF14 constructs (RGD and surrounding domains), with only PF14 actively signaling into human umbilical vein endothelial cells [48]. The enhanced signaling mediated by PF14 was directly shown to enhance endothelial cell proliferation and migration. These findings are consistent with data from fibrillin-1 null mice, which show thinner and disorganized elastic lamellae leading to a detached endothelial lining [49]. Together these data support an important role for fibrillin-1 in the regulation of endothelial cell binding and signaling as one facet of a wider biological role relevant to arterial morphogenesis and physiology [49].
Importantly, fibrillin-1 fragments have also been shown to signal smooth muscle cells and inhibit their migration. Specifically, PF8 and PF9 constructs have demonstrated dramatically lower migration rates in a Boyden chamber assay in comparison to fibronectin [29]. Fibrillin-1 also appears to regulate the production of matrix metalloproteinase (MMP) and directly impact on the health and phenotype of vascular SMCs. In Fibrillin-1 deficient mice, SMCs are less well adhered, adopt a synthetic and proliferative phenotype and have up-regulated MMP9, leading to fragmentation of elastic lamellae and an enhanced inflammatory response [50]. Similar findings are observed in human MFS patients [51], demonstrating the importance of fibrillin-1 in SMC signaling.
Direct assessment of the blood compatibility of fibrillin-1 (or any fibrillin) has not yet been carried out, but studies of the thrombogenicity of vessel wall components infer their relative compatibility. Decellularized elastic fibers, containing microfibrils and an elastin core, have been shown to bind fewer platelets than collagen or isolated basement membrane [52]. While elastin in particular is highly compatible with blood (discussed in more detail below), microfibrillar components were shown to cause much lower platelet activation than collagen [53]. It has also been assumed that proteins containing an RGD sequence would strongly bind platelets and induce thrombus formation, as is the case for collagen and fibronectin. However, recent work functionalizing materials with RGD inspired peptides and demonstrating improved blood compatibility [54], challenge this model and a direct assessment of the effect of fibrillin-1 on coagulation is needed. Initial work showing the potential benefits of modifying the surface of a biomaterial with fibrillin-1 has been carried out using a polyurethane scaffold. Immobilized PF9 was shown to significantly enhance the binding and spreading of fibroblasts to the material surface and was notably superior to an RGD peptide originating from the sequence of fibronectin [55]. This demonstration supports the idea that not all RGD sites are equivalent.

2.2. Fibulin-5

The fibulins are a family of secreted glycoproteins associated with basement membranes, elastic fibers, and other matrices and are expressed in a variety of tissues. There have been seven fibulins identified, divided into class I and class II based on length and domain structures. Class II fibulins, called “short” fibulins, include fibulin-3, -4, -5 and -7. Of these, fibulin-5 is the most prominently expressed in the vasculature, co-localizing with all layers of elastic lamina and present on the surface of elastic fibers [56].
The importance of fibulin-5 (also called EVEC or DANCE) to normal elastogenesis was first demonstrated by fibulin-5−/− mice that show massively disorganized elastic fibers, resulting in aberrant skin, vascular and lung phenotypes [57]. Given that fibulin-5 is known to interact with the elastic fiber molecules, tropoelastin and fibrillin-1 [58] it was proposed to play an important role in fiber assembly, tethering the developing structure to cell surfaces. However, mice expressing mutant fibulin-5 with a disrupted RGD sequence display normal fiber formation, indicating that their role in elastogenesis requires further investigation [56] and is not limited to cellular interactions.
From a vascular perspective, fibulin-5 is of particular interest as it is abundantly expressed in developing arteries. In adult blood vessels, expression is at low levels, however, the expression can be restarted in response to vascular injury and is enhanced in atherosclerotic plaques [59]. Following vascular injury, fibulin-5−/− mice present excessive SMC proliferation and migration and are more susceptible to thrombus formation. Vessels were found to be less elastic and remodeling impaired, while highlighting an unexpected role for fibulin-5 in SMC regulation [60]. Fibulin-5 also cooperates with fibulin-2 in the formation of the internal elastic lamina. Mice lacking both proteins show severely disorganized lamellae, thinner vessel walls and display higher rates of thrombosis following injury [61].
Fibulin-5 is the only fibulin to contain a conserved RGD motif, known to mediate binding to cell surface integrin receptors present on both endothelial cells and vascular smooth muscle cells [62]. For EC’s, binding is mediated by integrins including αvβ3 and αvβ5, while fibulin-5 also interacts strongly with TGF-βανδ vascular endothelial growth factor (VEGF), implicating a role in regulating angiogenesis and in modulation of endothelial cell function [63]. In vitro, over-expression of fibulin-5 in primary endothelial cells enhanced their attachment, but impaired proliferation. Dual expression with VEGF in the same cell type improved cell proliferation while maintaining a high level of attachment [64]. The importance of fibulin-5 in the protection of ECs is highlighted in hypoxic conditions, where fibulin-5 gene expression is increased and the rate of apoptosis in fibulin-5 knockdown cells is greater [65]. Surfaces coated with fibulin-5 exhibit strong integrin-mediated EC attachment in static and flow conditions at a level comparable to fibrillin-1 and the cells form stable, functional monolayers [29].
The interaction of fibulin-5 with smooth muscle cells occurs via a different subset of integrins and has been shown to have an inhibitory effect. Vascular SMCs from fibulin-5−/− mice have enhanced proliferative and migratory behavior compared to wild-type cells, a response inhibited by overexpression of fibulin-5 [60]. This inhibition occurred in the absence of the αvβ3 integrin demonstrating the presence of additional mediators. The attachment of primary human aortic SMCs to recombinant fibulin-5 was subsequently shown to be through α5β1 and α4β1 integrins. However, SMCs in this study were poorly attached and spread and showed inferior migration and proliferation in comparison to fibronectin [58]. Migration of SMCs to a confluent layer of ECs grown on fibulin-5 was also reduced, in further support of the inhibitory effect of fibulin-5 on SMCs [29].
The blood interactions of fibulin-5 have not been studied, though the increased thrombosis observed in fibulin-5/fibulin-2 double knockout mice infers a role in maintaining hemostasis. As with fibrillin-1, the presence of fibulin-5 in association with elastic fibers is also encouraging given the reduced interactions these have with platelets. As discussed above, the presence of an RGD is not necessarily a barrier to blood compatibility.

2.3. Perlecan

Perlecan is a large (467 kDa) multidomain heparan sulfate proteoglycan (HSPG) that is expressed in most ECM and basement membranes and is essential for the assembly and maintenance of a functional basement membrane. It is the major extracellular HPSG present in blood vessels and consists of a core made up of five distinct domains that each interact with molecules involved in cell proliferation, lipoprotein uptake and cell adhesion [66]. In vertebrates, perlecan functions in a diverse range of developmental and biological processes, including the regulation of wound healing and modulation of vascular biology [31]. It has been found to co-localize with fibrillin-1 and elastin in blood vessels [67] and to play an important regulatory role in sequestering a host of growth factors and cell signaling molecules [68]. Other heparan sulfate proteoglycans include syndecans and glypicans, however these are membrane bound [69]. Secreted HSPGs, agrin and collagen XVIII are of interest; however their roles in the vascular healing process have not been fully elucidated [70].
Perlecan−/− mice display severe phenotypic changes with most embryos dying prior to birth at E10-E12 as a result of ruptured basement membranes and consequential bleeding [71]. Cardiac tissue is especially affected. While basement membranes are present, they lack integrity and holes are observed in the myocardium. The few animals to survive until birth die soon after, suffering serious skeletal defects and defective cephalic development [72], emphasizing the importance of perlecan in a range of tissues. Focusing further on the cardiovascular consequences of perlecan deficiency, heterozygous mice show accelerated atherosclerosis in early lesions, implying a role in retention of lipoproteins [73]. Perlecan−/− mice more seriously demonstrate a high incidence of arterial transposition, valve malformation and hyperplasia of SMC-specific α-actin-positive mesenchymal cells [74].
The modulation of vascular specific cells by perlecan has been clearly demonstrated; inhibiting smooth muscle cells while promoting endothelialization and displaying potent pro-angiogenic activity. The mechanism of these divergent effects is not yet fully understood. It was originally proposed that mouse derived perlecan interacted directly with endothelial cells through β1 and β3 integrins in a substantially RGD dependent manner [75]. However, later studies importantly differentiated that while mouse perlecan contains a prominent RGD motif, the human form contains no RGD at all [76]. An emerging explanation is that cell interactive effects of perlecan are mediated via heparin sulfate side chains sequestering growth factors such as fibroblast growth factor-2 (FGF-2), VEGF and hepatocyte growth factor (HGF). FGF-2 in particular readily binds to these side chains and can stimulate EC proliferation and migration [77].
The inhibition of vascular SMCs has been closely associated with perlecan expression levels in the cell [78]. High levels are found in quiescent phenotypes, with decreased amounts correlating to enhanced proliferation and in cases of vascular injury. As the healing response moves toward completion, perlecan accumulation in the cell induces a reduction in SMC proliferation via a PTEN signaling pathway [79]. This effect has been demonstrated in vivo, using a rat model undergoing balloon induced carotid artery injury. Specifically, expression and deposition of perlecan are low in the early stages following injury, while large amounts of perlecan remain deposited during later stages of lesion development, when SMC proliferation decreases. Up to 6 weeks after injury, perlecan remains abundant, suggesting that turnover is slow in the advanced lesion [80]. Mechanistically, the inhibition of SMCs by perlecan has been linked to the presence of functional heparin sulfate side chains. Mutant SMCs with abrogated side chain secretion show increased proliferation, consistent with the greater intimal hyperplasia observed in the arteries of mice also lacking HS side-chains. As with EC signaling, it is proposed that the effect of perlecan on SMCs is also mediated by the sequestering ability of the side-chains and specifically their binding of FGF-2 [81].
Hemostasis in normal vessels is mediated by a functional monolayer of endothelial cells, which express HSPGs that in turn bind and activate anti-thrombin III, inactivating thrombin [82], as well as inhibiting platelet adhesion and activation due to their highly negative charge [83]. Accordingly, perlecan has been shown to exert a direct influence on the clotting cascade, preventing thrombosis and inhibiting platelet adhesion. In vitro, platelets monitored using a quartz crystal microbalance adhered at similar levels to perlecan with and without HS side-chains, though aggregation was blocked only in the presence of HS chains only [84]. Endothelial cells expressing perlecan at normal levels were also shown to completely prevent occlusive thrombosis in a porcine carotid injury model. In contrast, perlecan-deficient cells had a 23% occlusion rate [32]. Notably, knockdown cells had a reduced ability to interact with FGF-2, a molecule frequently associated with perlecan activity.
The positive effects of perlecan shown in vitro have some evidence of translation to in vivo use. ePTFE vascular grafts dip-coated with perlecan were evaluated in a sheep carotid interposition model. After 6 weeks, perlecan-coated grafts showed a trend to less thrombus formation compared to uncoated controls and despite high inter-animal variability, a higher degree of endothelial cell coverage [84]. Additionally, a novel compound (RUS3108) that induces perlecan over-expression in SMCs was eluted from a stent platform in a rabbit iliac artery model. The cross-sectional area of in-stent neointima in the treated stent group was significantly reduced in comparison to bare stent or polymer coated alone controls [85].

2.4. Tropoelastin

Elastin is one of the major structural components of the vasculature and provides critical mechanical and biological properties [86]. Elastin is formed through the cross-linking of its soluble precursor tropoelastin, which is initiated through the action of the enzyme lysyl oxidase [87]. Tropoelastin is a 60 kDa highly extensible yet elastic monomer [88] that is cross-linked in the extracellular space and assembles to become the major component of elastic fibers [89].
The importance of normal elastin expression to the vasculature is evidenced by the severe disruptions observed in elastin knockout mice. Elastin+/− animals present with hypertension, reduced vessel elasticity and display a reorganized arterial wall structure, but survive to normal age [90]. At birth, vascular function appears normal but progressive remodeling leads to reduced performance in adulthood. Approximately 60% of normal elastin levels are required for normal cardiac function, though this is achieved by significant vessel wall remodeling, notably with an increase in the number of elastic lamellae and layers of SMCs [91]. In elastin−/− mice, arterial development appears normal until E17.5 when the internal elastic lamina (IEL) is severely compromised and subendothelial SMC proliferation becomes uncontrolled. Mice die prematurely of arterial obstruction [92].
The internal elastic lamina is particularly significant in providing signaling and support for luminal endothelial cells [93]. Commensurate with this role, various forms of elastin interact favorably with endothelial cells and support their growth. Elastin solubilized with oxalic acid (α-elastin) enhanced proliferation of ECs at low concentrations [94], while elastin peptides containing the GP/GVGAGVP or VGVAPG motifs are chemotactic for endothelial cells [95] and are consequently pro-angiogenic [96]. Fibrous scaffolds of bovine elastin were also shown to support endothelial cell growth up to 7 days in culture [97] however, the clearest enhancement of endothelialization has been demonstrated with recombinant human tropoelastin (rhTE). Attachment and proliferation of ECs was significantly increased in the presence of tropoelastin immobilized on a plasma deposited surface [15] and shown to be equivalent to fibronectin when coated on tissue culture plastic [98]. The formation of an EC monolayer was also encouraged on three-dimensional fibrous scaffolds containing tropoelastin, applied to the development of a vascular conduit [98]. Recently, it was shown that human coronary artery endothelial cell attachment to tropoelastin is in part mediated through the integrin avβ3, however the site of this interaction as well as other binding interactions has not been elucidated [99].
Elastin is also a potent autocrine regulator of SMC activity that inhibits proliferation and regulates migration. This effect is mediated directly by tropoelastin sequences via a repeating VGVAPG hexapeptide motif. Supply of this hexapeptide activates the same signaling cascade as the full-length protein in a specific and dose dependent manner [100]. In vitro, α-elastin has a strong dose dependent inhibitory effect on SMCs causing a greater than 50% reduction even at low concentrations (0.1 mg/mL) [94]. SMC inhibition has subsequently been demonstrated repeatedly for α-elastin scaffolds [101], hydrogels [102] and electrospun sheets [103]. Studies of SMCs derived from elastin−/− mice provide the strongest evidence for the inhibitory effect of elastin. Cells lacking elastin were highly proliferative and increased by more than twofold by 72 hours, returning to wild type levels following the addition of recombinant tropoelastin to the media [104].
In addition, there is mounting evidence for the haemocompatibility of elastin and its derivatives and this subject has been reviewed in detail elsewhere [105]. In brief, elastin has demonstrated minimal platelet adhesion, degranulation and aggregation and was shown to be one of the least thrombogenic components of the vessel wall. In support of this finding, immobilized rhTE was found to have low thromobengicity on a plasma-activated steel surface [106], while porcine elastin-based scaffolds showed decreased thrombosis compared to commercial ePTFE in acute thrombogenicity tests in a carotid interposition model [107].
Translating the vascular compatibility of elastin-based materials in vivo, recombinant peptides based on the sequence of tropoelastin, have been shown to reduce thrombogenicity when used as surface coatings. Coated polyurethane catheters implanted in a rabbit model demonstrated decreased platelet deposition, reduced fibrin aggregation and ultimately improved patency [108]. Similar improvements in blood contacting properties were seen in a baboon arteriovenous shunt model using 4mm expanded polytetrafluoroethylene grafts coated with elastin-mimetic sequences [109]. Stents coated with an elastin sheet deployed in the coronary arteries of pigs have also demonstrated reduced neointimal hyperplasia compared to uncoated stents [110].

3. Conclusions and Future Prospects

Collectively, the outcomes from commercial graft and stent technologies have shown that simultaneously achieving reductions in neointimal hyperplasia, enhanced endothelialization and low thrombogenicity remains an elusive goal. This article sought to highlight the potential benefits of a biomimicry approach in the production of novel medical devices, employing biomolecules that have been shown to modulate multiple facets of vascular biology. A subset of candidates, chosen from the multitude found in the vasculature, standout as top tier candidates, and their favorable properties are summarized in Table 1.
Table
Table 1. Summary of vascular biological effects of biomolecule candidates.
Table 1. Summary of vascular biological effects of biomolecule candidates.
CandidateEffect on ECsEffect on SMCsBlood CompatibilityTranslation to Date
Fibrillin-1Proliferation, enhancedProliferation, inhibitedNot yet testedEnhanced fibroblast attachment to PU scaffold
Fibulin-5Attachment enhanced; apoptosis reducedProliferation, inhibitedNot yet testedNo translation
TropoelastinAttachment, proliferation, enhanced Proliferation, inhibitedHemocompatible; minimal activation of plateletsImproves EC binding, growth on steel; reduces thrombogenicity of catheters, ePTFE grafts
PerlecanProliferation, enhancedProliferation, hyperplasia (rat model) inhibitedDirect inhibition of thrombosisLess thrombus present on ePTFE grafts, greater ECs; Stents show less neointima
With a growing appreciation of the need for biointegration of vascular devices, it is anticipated that these candidates will increasingly be translated to metal and polymeric substrates for in vivo evaluation. This translation will be achieved more quickly in combination with emerging coating technologies that facilitate binding of biomolecules, in a lasting yet bioactive manner. Recent improvements, set apart from conventional plasma polymerisation, eliminating the need for chemical linkers is one such example [111]. Future developments will continue to aim to produce a novel device platform that can achieve controlled integration with the vasculature through multifaceted biomimicry. Ultimately, the goal remains the development of truly biocompatible materials for a wide range of vascular applications, including stents and graft materials, thereby meeting a large unmet need in cardiovascular medicine.

Conflict of Interest

The authors declare no conflict of interest.

Acknowledgments

We acknowledge grant support from the National Health and Medical Research Council and funding from The Baird Institute.

References

  1. Kannan, R.Y.; Salacinski, H.J.; Butler, P.E.; Hamilton, G.; Seifalian, A.M. Current status of prosthetic bypass grafts: A review. J. Biomed. Mater. Res. B 2005, 74, 570–581. [Google Scholar]
  2. Devine, C.; Hons, B.; McCollum, C. Heparin-bonded dacron or polytetrafluoroethylene for femoropopliteal bypass grafting: A multicenter trial. J. Vasc. Surg. 2001, 33, 533–539. [Google Scholar] [CrossRef]
  3. Devine, C.; McCollum, C. Heparin-bonded dacron or polytetrafluorethylene for femoropopliteal bypass: Five-year results of a prospective randomized multicenter clinical trial. J. Vasc. Surg. 2004, 40, 924–931. [Google Scholar] [CrossRef]
  4. Maegdefessel, L.; Linde, T.; Krapiec, F.; Hamilton, K.; Steinseifer, U.; van Ryn, J.; Raaz, U.; Buerke, M.; Werdan, K.; Schlitt, A. In vitro comparison of dabigatran, unfractionated heparin, and low-molecular-weight heparin in preventing thrombus formation on mechanical heart valves. Thromb. Res. 2010, 126, e196–e200. [Google Scholar] [CrossRef]
  5. Cutlip, D.E.; Baim, D.S.; Ho, K.K.; Popma, J.J.; Lansky, A.J.; Cohen, D.J.; Carrozza, J.P., Jr.; Chauhan, M.S.; Rodriguez, O.; Kuntz, R.E. Stent thrombosis in the modern era: A pooled analysis of multicenter coronary stent clinical trials. Circulation 2001, 103, 1967–1971. [Google Scholar]
  6. Joner, M.; Finn, A.V.; Farb, A.; Mont, E.K.; Kolodgie, F.D.; Ladich, E.; Kutys, R.; Skorija, K.; Gold, H.K.; Virmani, R. Pathology of drug-eluting stents in humans: Delayed healing and late thrombotic risk. J. Am. Coll. Cardiol. 2006, 48, 193–202. [Google Scholar] [CrossRef]
  7. Lemesle, G.; Delhaye, C.; Bonello, L.; de Labriolle, A.; Waksman, R.; Pichard, A. Stent thrombosis in 2008: Definition, predictors, prognosis and treatment. Arch. Cardiovasc. Dis. 2008, 101, 769–777. [Google Scholar] [CrossRef]
  8. Cutlip, D.E.; Windecker, S.; Mehran, R.; Boam, A.; Cohen, D.J.; van Es, G.A.; Steg, P.G.; Morel, M.A.; Mauri, L.; Vranckx, P.; et al. Clinical end points in coronary stent trials: A case for standardized definitions. Circulation 2007, 115, 2344–2351. [Google Scholar] [CrossRef]
  9. Daemen, J.; Wenaweser, P.; Tsuchida, K.; Abrecht, L.; Vaina, S.; Morger, C.; Kukreja, N.; Juni, P.; Sianos, G.; Hellige, G.; et al. Early and late coronary stent thrombosis of sirolimus-eluting and paclitaxel-eluting stents in routine clinical practice: Data from a large two-institutional cohort study. Lancet 2007, 369, 667–678. [Google Scholar]
  10. Kedhi, E.; Joesoef, K.S.; McFadden, E.; Wassing, J.; van Mieghem, C.; Goedhart, D.; Smits, P.C. Second-generation everolimus-eluting and paclitaxel-eluting stents in real-life practice (compare): A randomised trial. Lancet 2010, 375, 201–209. [Google Scholar]
  11. Pendyala, L.; Jabara, R.; Robinson, K.; Chronos, N. Passive and active polymer coatings for intracoronary stents: Novel devices to promote arterial healing. J. Int. Cardiol. 2009, 22, 37–48. [Google Scholar] [CrossRef]
  12. Williams, D.F. On the mechanisms of biocompatibility. Biomaterials 2008, 29, 2941–2953. [Google Scholar] [CrossRef]
  13. Ott, H.C.; Matthiesen, T.S.; Goh, S.K.; Black, L.D.; Kren, S.M.; Netoff, T.I.; Taylor, D.A. Perfusion-decellularized matrix: Using nature’s platform to engineer a bioartificial heart. Nat. Med. 2008, 14, 213–221. [Google Scholar]
  14. Campbell, J.H.; Efendy, J.L.; Campbell, G.R. Novel vascular graft grown within recipient’s own peritoneal cavity. Circ. Res. 1999, 85, 1173–1178. [Google Scholar]
  15. Yin, Y.; Wise, S.G.; Nosworthy, N.J.; Waterhouse, A.; Bax, D.V.; Youssef, H.; Byrom, M.J.; Bilek, M.M.; McKenzie, D.R.; Weiss, A.S.; et al. Covalent immobilisation of tropoelastin on a plasma deposited interface for enhancement of endothelialisation on metal surfaces. Biomaterials 2009, 30, 1675–1681. [Google Scholar] [CrossRef]
  16. Bennett, J.S.; Berger, B.W.; Billings, P.C. The structure and function of platelet integrins. J. Thromb. Haem. 2009, 7, 200–205. [Google Scholar] [CrossRef]
  17. Vyas, S.P.; Vaidya, B. Targeted delivery of thrombolytic agents: Role of integrin receptors. Expert Opin. Drug Deliv. 2009, 6, 499–508. [Google Scholar] [CrossRef]
  18. Maurer, L.M.; Tomasini-Johansson, B.R.; Mosher, D.F. Emerging roles of fibronectin in thrombosis. Thromb. Res. 2010, 125, 287–291. [Google Scholar] [CrossRef]
  19. Dufourcq, P.; Couffinhal, T.; Alzieu, P.; Daret, D.; Moreau, C.; Duplaa, C.; Bonnet, J. Vitronectin is upregulated after vascular injury and vitronectin blockade prevents neointima formation. Card. Res. 2002, 53, 952–962. [Google Scholar] [CrossRef]
  20. Nelson, P.R.; Yamamura, S.; Kent, K.C. Extracellular matrix proteins are potent agonists of human smooth muscle cell migration. J. Vasc. Surg. 1996, 24, 25–33. [Google Scholar] [CrossRef]
  21. Kushwaha, M.; Anderson, J.M.; Bosworth, C.A.; Andukuri, A.; Minor, W.P.; Lancaster, J.R., Jr.; Anderson, P.G.; Brott, B.C.; Jun, H.W. A nitric oxide releasing, self assembled peptide amphiphile matrix that mimics native endothelium for coating implantable cardiovascular devices. Biomaterials 2010, 31, 1502–1508. [Google Scholar]
  22. De Mel, A.; Jell, G.; Stevens, M.M.; Seifalian, A.M. Biofunctionalization of biomaterials for accelerated in situ endothelialization: A review. Biomacromolecules 2008, 9, 2969–2979. [Google Scholar] [CrossRef]
  23. Lewis, A.L. Phosphorylcholine-based polymers and their use in the prevention of biofouling. Coll. Surf. B Bioint. 2000, 18, 261–275. [Google Scholar] [CrossRef]
  24. Whelan, D.M.; van der Giessen, W.J.; Krabbendam, S.C.; van Vliet, E.A.; Verdouw, P.D.; Serruys, P.W.; van Beusekom, H.M. Biocompatibility of phosphorylcholine coated stents in normal porcine coronary arteries. Heart 2000, 83, 338–345. [Google Scholar] [CrossRef]
  25. Adams, J.C.; Watt, F.M. Regulation of development and differentiation by the extracellular matrix. Development 1993, 117, 1183–1198. [Google Scholar]
  26. Mecham, R. Overview of extracellular matrix. Curr. Prot. Cell Biol. 1998, 10, 1–14. [Google Scholar] [CrossRef]
  27. Juliano, R.L.; Haskill, S. Signal transduction from the extracellular matrix. J. Cell Biol. 1993, 120, 577–585. [Google Scholar] [CrossRef]
  28. Stephan, S.; Ball, S.G.; Williamson, M.; Bax, D.V.; Lomas, A.; Shuttleworth, C.A.; Kielty, C.M. Cell-matrix biology in vascular tissue engineering. J. Aant. 2006, 209, 495–502. [Google Scholar]
  29. Williamson, M.R.; Shuttleworth, A.; Canfield, A.E.; Black, R.A.; Kielty, C.M. The role of endothelial cell attachment to elastic fibre molecules in the enhancement of monolayer formation and retention, and the inhibition of smooth muscle cell recruitment. Biomaterials 2007, 28, 5307–5318. [Google Scholar] [CrossRef]
  30. Kalluri, R. Basement membranes: Structure, assembly and role in tumour angiogenesis. Nat. Rev. Cancer 2003, 3, 422–433. [Google Scholar] [CrossRef]
  31. Knox, S.M.; Whitelock, J.M. Perlecan: How does one molecule do so many things? Cell. Mol. Life Sci. 2006, 63, 2435–2445. [Google Scholar] [CrossRef]
  32. 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. [Google Scholar]
  33. Robb, B.W.; Wachi, H.; Schaub, T.; Mecham, R.P.; Davis, E.C. Characterization of an in vitro model of elastic fiber assembly. Mol. Biol. Cell 1999, 10, 3595–3605. [Google Scholar]
  34. Mithieux, S.M.; Weiss, A.S. Elastin. Adv. Protein Chem. 2005, 70, 437–461. [Google Scholar]
  35. Kielty, C.M.; Shuttleworth, C.A. Microfibrillar elements of the dermal matrix. Microsc. Res. Tech. 1997, 38, 413–427. [Google Scholar] [CrossRef]
  36. Davis, E.C.; Roth, R.A.; Heuser, J.E.; Mecham, R.P. Ultrastructural properties of ciliary zonule microfibrils. J. Struct. Biol. 2002, 139, 65–75. [Google Scholar] [CrossRef]
  37. Sabatier, L.; Miosge, N.; Hubmacher, D.; Lin, G.; Davis, E.C.; Reinhardt, D.P. Fibrillin-3 expression in human development. Matrix Biol. 2011, 30, 43–52. [Google Scholar] [CrossRef]
  38. Kielty, C.M.; Sherratt, M.J.; Shuttleworth, C.A. Elastic fibres. J. Cell Sci. 2002, 115, 2817–2828. [Google Scholar]
  39. Amadeu, T.P.; Braune, A.S.; Porto, L.C.; Desmouliere, A.; Costa, A.M. Fibrillin-1 and elastin are differentially expressed in hypertrophic scars and keloids. Wound Repair Regen. 2004, 12, 169–174. [Google Scholar] [CrossRef]
  40. Rock, M.J.; Cain, S.A.; Freeman, L.J.; Morgan, A.; Mellody, K.; Marson, A.; Shuttleworth, C.A.; Weiss, A.S.; Kielty, C.M. Molecular basis of elastic fiber formation: Critical interactions and a tropoelastin-fibrillin-1 cross-link. J. Biol. Chem. 2004, 279, 23748–23758. [Google Scholar]
  41. Kielty, C.M.; Sherratt, M.J.; Shuttleworth, C.A. Elastic fibres. J. Cell Sci. 2002, 115, 2817–2828. [Google Scholar]
  42. Dietz, H.C.; Cutting, G.R.; Pyeritz, R.E.; Maslen, C.L.; Sakai, L.Y.; Corson, G.M.; Puffenberger, E.G.; Hamosh, A.; Nanthakumar, E.J.; Curristin, S.M.; et al. Marfan syndrome caused by a recurrent de novo missense mutation in the fibrillin gene. Nature 1991, 352, 337–339. [Google Scholar]
  43. Charbonneau, N.L.; Carlson, E.J.; Tufa, S.; Sengle, G.; Manalo, E.C.; Carlberg, V.M.; Ramirez, F.; Keene, D.R.; Sakai, L.Y. In vivo studies of mutant fibrillin-1 microfibrils. J. Biol. Chem. 2010, 285, 24943–24955. [Google Scholar]
  44. Massam-Wu, T.; Chiu, M.; Choudhury, R.; Chaudhry, S.S.; Baldwin, A.K.; McGovern, A.; Baldock, C.; Shuttleworth, C.A.; Kielty, C.M. Assembly of fibrillin microfibrils governs extracellular deposition of latent tgf beta. J. Cell. Sci. 2010, 123, 3006–3018. [Google Scholar] [CrossRef]
  45. Rock, M.J.; Cain, S.A.; Freeman, L.J.; Morgan, A.; Mellody, K.; Marson, A.; Shuttleworth, C.A.; Weiss, A.S.; Kielty, C.M. Molecular basis of elastic fiber formation: Critical interactions and a tropoelastin-fibrillin-1 cross-link. J. Biol. Chem. 2004, 279, 23748–23758. [Google Scholar]
  46. Bax, D.V.; Mahalingam, Y.; Cain, S.; Mellody, K.; Freeman, L.; Younger, K.; Shuttleworth, C.A.; Humphries, M.J.; Couchman, J.R.; Kielty, C.M. Cell adhesion to fibrillin-1: Identification of an arg-gly-asp-dependent synergy region and a heparin-binding site that regulates focal adhesion formation. J. Cell. Sci. 2007, 120, 1383–1392. [Google Scholar] [CrossRef]
  47. Bax, D.V.; Bernard, S.E.; Lomas, A.; Morgan, A.; Humphries, J.; Shuttleworth, C.A.; Humphries, M.J.; Kielty, C.M. Cell adhesion to fibrillin-1 molecules and microfibrils is mediated by alpha 5 beta 1 and alpha v beta 3 integrins. J. Biol. Chem. 2003, 278, 34605–34616. [Google Scholar]
  48. Mariko, B.; Ghandour, Z.; Raveaud, S.; Quentin, M.; Usson, Y.; Verdetti, J.; Huber, P.; Kielty, C.; Faury, G. Microfibrils and fibrillin-1 induce integrin-mediated signaling, proliferation and migration in human endothelial cells. Am. J. Phys. Cell Phys. 2010, 299, C977–C987. [Google Scholar] [CrossRef]
  49. Carta, L.; Pereira, L.; Arteaga-Solis, E.; Lee-Arteaga, S.Y.; Lenart, B.; Starcher, B.; Merkel, C.A.; Sukoyan, M.; Kerkis, A.; Hazeki, N.; et al. Fibrillins 1 and 2 perform partially overlapping functions during aortic development. J. Biol. Chem. 2006, 281, 8016–8023. [Google Scholar]
  50. Bunton, T.E.; Biery, N.J.; Myers, L.; Gayraud, B.; Ramirez, F.; Dietz, H.C. Phenotypic alteration of vascular smooth muscle cells precedes elastolysis in a mouse model of marfan syndrome. Circ. Res. 2001, 88, 37–43. [Google Scholar]
  51. Nataatmadja, M.; West, M.; West, J.; Summers, K.; Walker, P.; Nagata, M.; Watanabe, T. Abnormal extracellular matrix protein transport associated with increased apoptosis of vascular smooth muscle cells in marfan syndrome and bicuspid aortic valve thoracic aortic aneurysm. Circulation 2003, 108 Suppl. 1, II329–II334. [Google Scholar]
  52. Baumgartner, H.R.; Muggli, R.; Tschopp, T.B.; Turitto, V.T. Platelet adhesion, release and aggregation in flowing blood: Effects of surface properties and platelet function. Thromb. Haem. 1976, 35, 124–138. [Google Scholar]
  53. Legrand, Y.; Karniguian, A.; Fauvel, F.; Gutman, N. The molecular interaction between platelet and vascular wall. Blood Cells 1983, 9, 263–274. [Google Scholar]
  54. Zheng, W.; Wang, Z.; Song, L.; Zhao, Q.; Zhang, J.; Li, D.; Wang, S.; Han, J.; Zheng, X.L.; Yang, Z.; et al. Endothelialization and patency of rgd-functionalized vascular grafts in a rabbit carotid artery model. Biomaterials 2012, 33, 2880–2891. [Google Scholar]
  55. Jozwiak, A.B.; Kielty, C.M.; Black, R.A. Surface functionalization of polyurethane for the immobilization of bioactive moieties on tissue scaffolds. J. Mater. Chem. 2008, 18, 2240–2248. [Google Scholar]
  56. 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. [Google Scholar] [CrossRef]
  57. Yanagisawa, H.; Davis, E.C.; Starcher, B.C.; Ouchi, T.; Yanagisawa, M.; Richardson, J.A.; Olson, E.N. Fibulin-5 is an elastin-binding protein essential for elastic fibre development in vivo. Nature 2002, 415, 168–171. [Google Scholar]
  58. Lomas, A.C.; Mellody, K.T.; Freeman, L.J.; Bax, D.V.; Shuttleworth, C.A.; Kielty, C.M. Fibulin-5 binds human smooth-muscle cells through alpha5beta1 and alpha4beta1 integrins, but does not support receptor activation. Biochem. J. 2007, 405, 417–428. [Google Scholar] [CrossRef]
  59. Freeman, L.J.; Lomas, A.; Hodson, N.; Sherratt, M.J.; Mellody, K.T.; Weiss, A.S.; Shuttleworth, A.; Kielty, C.M. Fibulin-5 interacts with fibrillin-1 molecules and microfibrils. Biochem. J. 2005, 388, 1–5. [Google Scholar]
  60. Spencer, J.A.; Hacker, S.L.; Davis, E.C.; Mecham, R.P.; Knutsen, R.H.; Li, D.Y.; Gerard, R.D.; Richardson, J.A.; Olson, E.N.; Yanagisawa, H. Altered vascular remodeling in fibulin-5-deficient mice reveals a role of fibulin-5 in smooth muscle cell proliferation and migration. Proc. Natl. Acad. Sci. USA 2005, 102, 2946–2951. [Google Scholar]
  61. Chapman, S.L.; Sicot, F.X.; Davis, E.C.; Huang, J.; Sasaki, T.; Chu, M.L.; Yanagisawa, H. Fibulin-2 and fibulin-5 cooperatively function to form the internal elastic lamina and protect from vascular injury. Arterioscler. Thromb. Vasc. Biol. 2010, 30, 68–74. [Google Scholar]
  62. Yanagisawa, H.; Schluterman, M.K.; Brekken, R.A. Fibulin-5, an integrin-binding matricellular protein: Its function in development and disease. J. Cell Commun. Sig. 2009, 3, 337–347. [Google Scholar] [CrossRef]
  63. Nakamura, T.; Ruiz-Lozano, P.; Lindner, V.; Yabe, D.; Taniwaki, M.; Furukawa, Y.; Kobuke, K.; Tashiro, K.; Lu, Z.; Andon, N.L.; et al. Dance, a novel secreted rgd protein expressed in developing, atherosclerotic, and balloon-injured arteries. J. Biol. Chem. 1999, 274, 22476–22483. [Google Scholar]
  64. Preis, M.; Cohen, T.; Sarnatzki, Y.; Ben Yosef, Y.; Schneiderman, J.; Gluzman, Z.; Koren, B.; Lewis, B.S.; Shaul, Y.; Flugelman, M.Y. Effects of fibulin-5 on attachment, adhesion, and proliferation of primary human endothelial cells. Biochem. Biophys. Res. Commun. 2006, 348, 1024–1033. [Google Scholar]
  65. Guadall, A.; Orriols, M.; Rodriguez-Calvo, R.; Calvayrac, O.; Crespo, J.; Aledo, R.; Martinez-Gonzalez, J.; Rodriguez, C. Fibulin-5 is up-regulated by hypoxia in endothelial cells through a hypoxia-inducible factor-1 (hif-1alpha)-dependent mechanism. J. Biol. Chem. 2011, 286, 7093–7103. [Google Scholar]
  66. Whitelock, J.; Melrose, J. Heparan sulfate proteoglycans in healthy and diseased systems. Wiley Inter. Rev. Sys. Biol. Med. 2011, 3, 739–751. [Google Scholar] [CrossRef]
  67. Hayes, A.J.; Lord, M.S.; Smith, S.M.; Smith, M.M.; Whitelock, J.M.; Weiss, A.S.; Melrose, J. Colocalization in vivo and association in vitro of perlecan and elastin. Histo. Cell Biol. 2011, 136, 437–454. [Google Scholar]
  68. Whitelock, J.M.; Melrose, J.; Iozzo, R.V. Diverse cell signaling events modulated by perlecan. Biochemistry 2008, 47, 11174–11183. [Google Scholar]
  69. Alexopoulou, A.N.; Multhaupt, H.A.; Couchman, J.R. Syndecans in wound healing, inflammation and vascular biology. Int. J. Biochem. Cell Biol. 2007, 39, 505–528. [Google Scholar] [CrossRef]
  70. Kirkpatrick, C.A.; Selleck, S.B. Heparan sulfate proteoglycans at a glance. J. Cell. Sci. 2007, 120, 1829–1832. [Google Scholar] [CrossRef]
  71. Olsen, B.R. Life without perlecan has its problems. J. Cell Biol. 1999, 147, 909–912. [Google Scholar] [CrossRef]
  72. Arikawa-Hirasawa, E.; Watanabe, H.; Takami, H.; Hassell, J.R.; Yamada, Y. Perlecan is essential for cartilage and cephalic development. Nat. Genet. 1999, 23, 354–358. [Google Scholar] [CrossRef]
  73. Vikramadithyan, R.K.; Kako, Y.; Chen, G.; Hu, Y.; Arikawa-Hirasawa, E.; Yamada, Y.; Goldberg, I.J. Atherosclerosis in perlecan heterozygous mice. J. Lipid Res. 2004, 45, 1806–1812. [Google Scholar] [CrossRef]
  74. Costell, M.; Carmona, R.; Gustafsson, E.; Gonzalez-Iriarte, M.; Fassler, R.; Munoz-Chapuli, R. Hyperplastic conotruncal endocardial cushions and transposition of great arteries in perlecan-null mice. Circ. Res. 2002, 91, 158–164. [Google Scholar] [CrossRef]
  75. Hayashi, K.; Madri, J.A.; Yurchenco, P.D. Endothelial cells interact with the core protein of basement membrane perlecan through beta 1 and beta 3 integrins: An adhesion modulated by glycosaminoglycan. J. Cell Biol. 1992, 119, 945–959. [Google Scholar] [CrossRef]
  76. Whitelock, J.M.; Graham, L.D.; Melrose, J.; Murdoch, A.D.; Iozzo, R.V.; Underwood, P.A. Human perlecan immunopurified from different endothelial cell sources has different adhesive properties for vascular cells. Matrix Biol. 1999, 18, 163–178. [Google Scholar] [CrossRef]
  77. Segev, A.; Nili, N.; Strauss, B.H. The role of perlecan in arterial injury and angiogenesis. Cardiovasc. Res. 2004, 63, 603–610. [Google Scholar] [CrossRef]
  78. Weiser, M.C.; Belknap, J.K.; Grieshaber, S.S.; Kinsella, M.G.; Majack, R.A. Developmental regulation of perlecan gene expression in aortic smooth muscle cells. Matrix Biol. 1996, 15, 331–340. [Google Scholar] [CrossRef]
  79. Garl, P.J.; Wenzlau, J.M.; Walker, H.A.; Whitelock, J.M.; Costell, M.; Weiser-Evans, M.C. Perlecan-induced suppression of smooth muscle cell proliferation is mediated through increased activity of the tumor suppressor pten. Circ. Res. 2004, 94, 175–183. [Google Scholar] [CrossRef]
  80. Kinsella, M.G.; Tran, P.K.; Weiser-Evans, M.C.; Reidy, M.; Majack, R.A.; Wight, T.N. Changes in perlecan expression during vascular injury: Role in the inhibition of smooth muscle cell proliferation in the late lesion. Arterioscler. Thromb. Vasc. Biol. 2003, 23, 608–614. [Google Scholar] [CrossRef]
  81. Tran, P.K.; Tran-Lundmark, K.; Soininen, R.; Tryggvason, K.; Thyberg, J.; Hedin, U. Increased intimal hyperplasia and smooth muscle cell proliferation in transgenic mice with heparan sulfate-deficient perlecan. Circ. Res. 2004, 94, 550–558. [Google Scholar] [CrossRef]
  82. Guyton, J.R.; Rosenberg, R.D.; Clowes, A.W.; Karnovsky, M.J. Inhibition of rat arterial smooth muscle cell proliferation by heparin. In vivo studies with anticoagulant and nonanticoagulant heparin. Circ. Res. 1980, 46, 625–634. [Google Scholar] [CrossRef]
  83. Mysliwiec, M.; Borawski, J.; Naumnik, B.; Rydzewska-Rosolowska, A. Endothelial dysfunction, atherosclerosis and thrombosis in uremia—possibilities of intervention. Rocz. Akad. Med. Bialymst. 2004, 49, 151–156. [Google Scholar]
  84. Lord, M.S.; Yu, W.; Cheng, B.; Simmons, A.; Poole-Warren, L.; Whitelock, J.M. The modulation of platelet and endothelial cell adhesion to vascular graft materials by perlecan. Biomaterials 2009, 30, 4898–4906. [Google Scholar]
  85. Segev, A.; Nili, N.; Osherov, A.B.; Qiang, B.; Wong, A.J.; Pillarisetti, S.; Strauss, B.H. A perlecan-inducing compound significantly inhibits smooth muscle cell function and in-stent intimal hyperplasia: Novel insights into the diverse biological effects of perlecan. EuroIntervention 2010, 6, 134–140. [Google Scholar] [CrossRef]
  86. Almine, J.F.; Bax, D.V.; Mithieux, S.M.; Nivison-Smith, L.; Rnjak, J.; Waterhouse, A.; Wise, S.G.; Weiss, A.S. Elastin-based materials. Chem. Soc. Rev. 2010, 39, 3371–3379. [Google Scholar]
  87. Kagan, H.M.; Li, W. Lysyl oxidase: Properties, specificity, and biological roles inside and outside of the cell. J. Cell. Biochem. 2003, 88, 660–672. [Google Scholar] [CrossRef]
  88. Baldock, C.; Oberhauser, A.F.; Ma, L.; Lammie, D.; Siegler, V.; Mithieux, S.M.; Tu, Y.; Chow, J.Y.; Suleman, F.; Malfois, M.; et al. Shape of tropoelastin, the highly extensible protein that controls human tissue elasticity. Proc. Natl. Acad. Sci. USA 2011, 108, 4322–4327. [Google Scholar]
  89. Wise, S.G.; Weiss, A.S. Tropoelastin. Int. J. Biochem. Cell Biol. 2009, 41, 494–497. [Google Scholar] [CrossRef]
  90. Wagenseil, J.E.; Nerurkar, N.L.; Knutsen, R.H.; Okamoto, R.J.; Li, D.Y.; Mecham, R.P. Effects of elastin haploinsufficiency on the mechanical behavior of mouse arteries. Am. J. Phys. Heart Circ. Phys. 2005, 289, H1209–H1217. [Google Scholar] [CrossRef]
  91. Wagenseil, J.E.; Ciliberto, C.H.; Knutsen, R.H.; Levy, M.A.; Kovacs, A.; Mecham, R.P. Reduced vessel elasticity alters cardiovascular structure and function in newborn mice. Circ. Res. 2009, 104, 1217–1224. [Google Scholar] [CrossRef]
  92. Li, D.Y.; Brooke, B.; Davis, E.C.; Mecham, R.P.; Sorensen, L.K.; Boak, B.B.; Eichwald, E.; Keating, M.T. Elastin is an essential determinant of arterial morphogenesis. Nature 1998, 393, 276–280. [Google Scholar] [CrossRef]
  93. Sims, F.H.; Gavin, J.B.; Edgar, S.; Koelmeyer, T.D. Comparison of the endothelial surface and subjacent elastic lamina of anterior descending coronary arteries at the location of atheromatous lesions with internal thoracic arteries of the same subjects: A scanning electron microscopic study. Pathology 2002, 34, 433–441. [Google Scholar] [CrossRef]
  94. Ito, S.; Ishimaru, S.; Wilson, S.E. Effect of coacervated alpha-elastin on proliferation of vascular smooth muscle and endothelial cells. Angiology 1998, 49, 289–297. [Google Scholar] [CrossRef]
  95. Long, M.M.; King, V.J.; Prasad, K.U.; Freeman, B.A.; Urry, D.W. Elastin repeat peptides as chemoattractants for bovine aortic endothelial cells. J. Cell. Phys. 1989, 140, 512–518. [Google Scholar] [CrossRef]
  96. Robinet, A.; Fahem, A.; Cauchard, J.-H.; Huet, E.; Vincent, L.; Lorimier, S.; Antonicelli, F.; Soria, C.; Crepin, M.; Hornebeck, W.; et al. Elastin-derived peptides enhance angiogenesis by promoting endothelial cell migration and tubulogenesis through upregulation of mt1-mmp. J. Cell Sci. 2005, 118, 343–356. [Google Scholar] [CrossRef]
  97. Lee, S.J.; Yoo, J.J.; Lim, G.J.; Atala, A.; Stitzel, J. In vitro evaluation of electrospun nanofiber scaffolds for vascular graft application. J. Biomed. Mater. Res. A 2007, 83, 999–1008. [Google Scholar]
  98. Wise, S.G.; Byrom, M.J.; Waterhouse, A.; Bannon, P.G.; Ng, M.K.; Weiss, A.S. A multilayered synthetic human elastin/polycaprolactone hybrid vascular graft with tailored mechanical properties. Acta Biomater. 2011, 7, 295–303. [Google Scholar]
  99. Wilson, B.D.; Gibson, C.C.; Sorensen, L.K.; Guilhermier, M.Y.; Clinger, M.; Kelley, L.L.; Shiu, Y.T.; Li, D.Y. Novel approach for endothelializing vascular devices: Understanding and exploiting elastin-endothelial interactions. Ann. Biomed. Eng. 2010, 39, 337–346. [Google Scholar]
  100. Karnik, S.K.; Wythe, J.D.; Sorensen, L.; Brooke, B.S.; Urness, L.D.; Li, D.Y. Elastin induces myofibrillogenesis via a specific domain, vgvapg. Matrix Biol. 2003, 22, 409–425. [Google Scholar] [CrossRef]
  101. Leach, J.B.; Wolinsky, J.B.; Stone, P.J.; Wong, J.Y. Crosslinked alpha-elastin biomaterials: Towards a processable elastin mimetic scaffold. Acta Biomater. 2005, 1, 155–164. [Google Scholar] [CrossRef]
  102. Ito, S.; Ishimaru, S.; Wilson, S.E. Inhibitory effect of type 1 collagen gel containing alpha-elastin on proliferation and migration of vascular smooth muscle and endothelial cells. Cardiovasc. Surg. 1997, 5, 176–183. [Google Scholar] [CrossRef]
  103. Miyamoto, K.; Atarashi, M.; Kadozono, H.; Shibata, M.; Koyama, Y.; Okai, M.; Inakuma, A.; Kitazono, E.; Kaneko, H.; Takebayashi, T.; et al. Creation of cross-linked electrospun isotypic-elastin fibers controlled cell-differentiation with new cross-linker. Int. J. Biol. Macromol. 2009, 45, 33–41. [Google Scholar] [CrossRef]
  104. Karnik, S.K.; Brooke, B.S.; Bayes-Genis, A.; Sorensen, L.; Wythe, J.D.; Schwartz, R.S.; Keating, M.T.; Li, D.Y. A critical role for elastin signaling in vascular morphogenesis and disease. Development 2003, 130, 411–423. [Google Scholar] [CrossRef]
  105. Waterhouse, A.; Wise, S.G.; Ng, M.K.; Weiss, A.S. Elastin as a nonthrombogenic biomaterial. Tissue Eng. Part B 2011, 17, 93–99. [Google Scholar]
  106. Waterhouse, A.; Yin, Y.B.; Wise, S.G.; Bax, D.V.; McKenzie, D.R.; Bilek, M.M.M.; Weiss, A.S.; Ng, M.K.C. The immobilization of recombinant human tropoelastin on metals using a plasma-activated coating to improve the biocompatibility of coronary stents. Biomaterials 2010, 31, 8332–8340. [Google Scholar]
  107. Hinds, M.T.; Rowe, R.C.; Ren, Z.; Teach, J.; Wu, P.-C.; Kirkpatrick, S.J.; Breneman, K.D.; Gregory, K.W; Courtman, D.W. Development of a reinforced porcine elastin composite vascular scaffold. J. Biomed. Mater. Res. A 2006, 77A, 458–469. [Google Scholar] [CrossRef]
  108. Woodhouse, K.A.; Klement, P.; Chen, V.; Gorbet, M.B.; Keeley, F.W.; Stahl, R.; Fromstein, J.D.; Bellingham, C.M. Investigation of recombinant human elastin polypeptides as non-thrombogenic coatings. Biomaterials 2004, 25, 4543–4553. [Google Scholar]
  109. Jordan, S.W.; Haller, C.A.; Sallach, R.E.; Apkarian, R.P.; Hanson, S.R.; Chaikof, E.L. The effect of a recombinant elastin-mimetic coating of an eptfe prosthesis on acute thrombogenicity in a baboon arteriovenous shunt. Biomaterials 2007, 28, 1191–1197. [Google Scholar] [CrossRef]
  110. Karnik, S.K.; Brooke, B.S.; Bayes-Genis, A.; Sorensen, L.; Wythe, J.D.; Schwartz, R.S.; Keating, M.T.; Li, D.Y. A critical role for elastin signaling in vascular morphogenesis and disease. Development 2003, 130, 411–423. [Google Scholar] [CrossRef]
  111. Bilek, M.M.; Bax, D.V.; Kondyurin, A.; Yin, Y.; Nosworthy, N.J.; Fisher, K.; Waterhouse, A.; Weiss, A.S.; dos Remedios, C.G.; McKenzie, D.R. Free radical functionalization of surfaces to prevent adverse responses to biomedical devices. Proc. Natl. Acad. Sci. USA 2011, 108, 14405–14410. [Google Scholar]

Share and Cite

MDPI and ACS Style

Wise, S.G.; Waterhouse, A.; Michael, P.; Ng, M.K.C. Extracellular Matrix Molecules Facilitating Vascular Biointegration. J. Funct. Biomater. 2012, 3, 569-587. https://doi.org/10.3390/jfb3030569

AMA Style

Wise SG, Waterhouse A, Michael P, Ng MKC. Extracellular Matrix Molecules Facilitating Vascular Biointegration. Journal of Functional Biomaterials. 2012; 3(3):569-587. https://doi.org/10.3390/jfb3030569

Chicago/Turabian Style

Wise, Steven G., Anna Waterhouse, Praveesuda Michael, and Martin K.C. Ng. 2012. "Extracellular Matrix Molecules Facilitating Vascular Biointegration" Journal of Functional Biomaterials 3, no. 3: 569-587. https://doi.org/10.3390/jfb3030569

APA Style

Wise, S. G., Waterhouse, A., Michael, P., & Ng, M. K. C. (2012). Extracellular Matrix Molecules Facilitating Vascular Biointegration. Journal of Functional Biomaterials, 3(3), 569-587. https://doi.org/10.3390/jfb3030569

Article Metrics

Back to TopTop