Enzymatic degradation of heparin-modified hydrogels and its effect on bioactivity
Introduction
Tissue engineered scaffolds are designed to mimic the extracellular matrix (ECM) by their structure and interactions with the surrounding host environment. Heparan sulfate (HS) molecules, which are a major part of the ECM, are continually synthesised and replaced in the body, a process that helps maintain homeostasis and regulate angiogenesis. HS is synthesised as a part of HS proteoglycan (HSPG), which consists of a protein core that carries linear HS chains composed of alternating hexuronic acid and glucosamine residues. These residues are differentially sulfated to create polydisperse chains with variably charged regions [1]. The HSPG sequesters numerous enzymes, growth factors and cytokines on the cell surface and in the ECM, often acts as an inactive reservoir [2]. Cleavage of the HSPG or HS chains releases fragments of these proteins or polysaccharides from the ECM and can affect their bioactivity [3]. In humans, the protein core of HSPG is susceptible to degradation by several proteases while the HS chains are cleaved by a single heparanase enzyme, an endo-β-glucuronidase that is often referred to as heparanase-1.
Heparanase cleaves HS at specific sites to release HS fragments as well as proteins such as growth factors that are trapped within the ECM or on cell surfaces [4], [5], making them available to the cell surface receptors and causing a change in the environmental conditions that may modulate local tissue responses. One of the primary functions of heparanase is to degrade basement membrane HSPG at sites of injury or inflammation [6], [7], [8], allowing the migration of inflammatory cells and releasing HS-bound growth factors that promote the migration and survival of endothelial cells [9]. The release of basic fibroblast growth factor (FGF-2) by the heparanase action on HS has traditionally been correlated with cell invasion during angiogenesis and inflammation [10].
Platelets have been shown to be a good source of heparanase and the heparanase activity has been reported to be optimum at a pH range of 5.1–6.8 [6], [11], [12], [13]. Other sources of heparanase include endothelial cells (EC) and smooth muscle cells (SMC), although the HS-degrading activity from these cells has been reported to be less that 10% that of platelet extract [13]. Nevertheless, cell lysates from both EC and SMC have been shown to contain factors that can further activate platelet heparanase.
Enzymatic degradation studies of biologically derived hydrogels have been performed to examine substrate availability on the crosslinked biopolymers for enzyme digestion. Collagen, for example, is degraded by metalloproteases and serine proteases, allowing degradation to be locally controlled by cells present in the engineered tissue [14]. Most of the studies have often used bacterial enzymes instead of those from mammalian sources, such as in the degradation studies of hydrogels formulated from chondroitin sulfate (CS) [15] and fibrin [16]. Mammalian heparanase differs from other HS-degrading enzymes such as heparinase and heparitinase from Flavobacterium heparinum, which are eliminases that typically cleave HS into di- or tri-saccharide units [17]. Therefore, to simulate the degradation of heparin in vivo and examine the activity of the resulting heparin fragments, human platelet extract (PE) was used in this study as a source of heparanase.
The aim of this study was to expose heparin-functionalised poly(vinyl alcohol) (PVA) hydrogels to enzymatic degradation, in order to simulate a condition that the co-hydrogels were likely to encounter in vivo. Work has been done to gain a better understanding of the HS-degrading action of heparanase on a molecular level [6], [10], but no reports have explored the effect of platelet heparanase on the activity of heparin-based hydrogels. The heparin-degrading activity of heparanase (PE) was examined by analysing the molecular weight of heparin following incubation with PE, as well as the anticoagulation and FGF-2 signalling properties of the heparin fragments. The degradation capacity of PE on the hydrogel-bound heparin was then examined. Following PE-treatment, extracts from the PVA/heparin co-hydrogels were collected and analysed to assess their molecular weight and the corresponding bioactivities.
Section snippets
Materials
Unless otherwise specified, all materials were from Sigma–Aldrich and used without further purification. Human platelet-rich serum (Australian Red Cross Blood Service) was used as received. Three heparin sodium salt samples, from porcine intestinal mucosa, were used: heparin (grade I-A, 17–19 kDa), low molecular weight (LMW) heparin (4–6 kDa), and Deligoparin™ (2–3 kDa, Opocrin, Italy). Coomassie Plus Reagent (Promega), bovine serum albumin (BSA), casein, monoclonal antibodies, biotinylated
Heparanase activity of the platelet extract
Platelets have been shown to be a reliable source from which heparanase could be obtained [7], [13], [25]. The overall protein concentration of the PE used in this study was determined in the Coomasie Plus assay to be 810 ± 28 μg/mL. To determine the presence of heparanase, an ELISA was performed on the PE using HPA-1 as a probe for heparanase. Antibodies against perlecan (7B5) or serglycin (a-ser) served as negative controls while DPBS was the blank control. The PE and DPBS were probed with
Conclusion
The aim of this study was to investigate the activity of heparin that has been functionalised and polymerised as part of a biosynthetic hybrid biomaterial, and incubated with PE in an attempt to model what might happen when the material is exposed to blood or activated plasma, both of which may be present at sites of injury. The heparanase activity of the PE was shown to be pH dependent and consistent with the action of glucuronidase-type enzymes. The PE was also found to be capable of
Acknowledgements
The authors thank Bill Cheng for help with the preparation of the platelet extract, Litania Lie for additional data on the clotting time and cell assays, and the Australian Research Council Discovery Grant (DP0557863) for project funding. AN wishes to acknowledge support from the UNSW Faculty of Engineering Research Scholarship and the Royal Perth Hospital during the preparation of this manuscript.
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