An injectable hydrogel incorporating mesenchymal precursor cells and pentosan polysulphate for intervertebral disc regeneration

Biomaterials 34 (2013) 9430e9440

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An injectable hydrogel incorporating mesenchymal precursor cells and pentosan polysulphate for intervertebral disc regeneration Jessica E. Frith a, Andrew R. Cameron a, Donna J. Menzies a, Peter Ghosh b, Darryl L. Whitehead c, Stan Gronthos d, Andrew C.W. Zannettino e, Justin J. Cooper-White a, f, g, * a

Australian Institute for Bioengineering and Nanotechnology, University of Queensland, St Lucia, Queensland 4072, Australia Mesoblast Ltd, Level 39, 55 Collins Street, Melbourne 3000, Australia School of Biomedical Sciences, University of Queensland, St Lucia, Queensland 4072, Australia d Mesenchymal Stem Cell Laboratory, School of Medical Sciences, Faculty of Health Sciences, University of Adelaide, Adelaide, 5005 South Australia, Australia e Myeloma Research Laboratory, School of Medical Sciences, Faculty of Health Science, University of Adelaide, Adelaide, South Australia 5000, Australia f School of Chemical Engineering, University of Queensland, St. Lucia, Queensland 4072, Australia g CSIRO Materials Science and Engineering Division, Clayton, 3168 Victoria, Australia b c

a r t i c l e i n f o

a b s t r a c t

Article history: Received 13 June 2013 Accepted 22 August 2013 Available online 16 September 2013

Intervertebral disc (IVD) degeneration is one of the leading causes of lower back pain and a major health problem worldwide. Current surgical treatments include excision or immobilisation, with neither approach resulting in the repair of the degenerative disc. As such, a tissue engineering-based approach in which stem cells, coupled with an advanced delivery system, could overcome this deficiency and lead to a therapy that encourages functional fibrocartilage generation in the IVD. In this study, we have developed an injectable hydrogel system based on enzymatically-crosslinked polyethylene glycol and hyaluronic acid. We examined the effects of adding pentosan polysulphate (PPS), a synthetic glycosaminoglycan-like factor that has previously been shown (in vitro and in vivo) to this gel system in order to induce chondrogenesis in mesnchymal precursor cells (MPCs) when added as a soluble factor, even in the absence of additional growth factors such as TGF-b. We show that both the gelation rate and mechanical strength of the resulting hydrogels can be tuned in order to optimise the conditions required to produce gels with the desired combination of properties for an IVD scaffold. Human immunoselected STRO-1þ MPCs were then incorporated into the hydrogels. They were shown to retain good viability after both the initial formation of the gel and for longer-term culture periods in vitro. Furthermore, MPC/ hydrogel composites formed cartilage-like tissue which was significantly enhanced by the incorporation of PPS into the hydrogels, particularly with respect to the deposition of type-II-collagen. Finally, using a wild-type rat subcutaneous implantation model, we examined the extent of any immune reaction and confirmed that this matrix is well tolerated by the host. Together these data provide evidence that such a system has significant potential as both a delivery vehicle for MPCs and as a matrix for fibrocartilage tissue engineering applications. Crown Copyright Ó 2013 Published by Elsevier Ltd. All rights reserved.

Keywords: Mesenchymal progenitor cell Hydrogel Nucleus pulposus Pentosan polysulphate Fibrocartilage

1. Introduction Degeneration of the intervertebral disc (IVD) is a leading cause of lower back pain and a significant societal and economic problem worldwide. During early stages, IVD degeneration can be treated with physiotherapy and painkillers but in more severe cases the

* Corresponding authors. School of Chemical Engineering, University of Queensland, St. Lucia, Queensland 4072, Australia. E-mail address: [email protected] (J.J. Cooper-White).

only currently available options include fusion of the vertebrae or discectomy. These procedures are invasive, reduce mobility and do not fully restore the function of the disc. Therefore there is an urgent need for an effective strategy to promote tissue regeneration and provide a long-term solution to IVD degeneration. When developing strategies to restore disc function, regeneration of the nucleus pulposus (NP) tissue is seen as a key target. The NP is rich in glycosaminoglycans, proteoglycans and type-IIcollagen and is highly hydrated so that the osmotic pressure it generates readily dissipates any mechanical forces transmitted through the spine. Degeneration of the IVD is characterised by a loss

0142-9612/$ e see front matter Crown Copyright Ó 2013 Published by Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biomaterials.2013.08.072

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of hydration and degradation of the extracellular matrix (ECM) of the NP, which in turn leads to a progressive decrease in disc height, thereby reducing the ability of the IVD to function in cushioning forces applied to the spine. For this reason, successful regeneration of the NP has significant potential to restore disc function. In the development of tissue engineering strategies to promote NP regeneration, mesenchymal stem/stromal cells (MSC) are seen as attractive candidates due to their ready availability, multilineage differentiation potential and low immunogenicity [1e3]. It is currently unknown whether mesenchymal stem cells (MSCs) can directly differentiate into an NP cell (primarily because no specific markers for NP cells have been identified), but in any case, it has been demonstrated that MSCs are able to develop a chondrogenic phenotype and express fibrocartilage-like ECM of the appropriate composition to aid in NP repair [4,5]. In this work, we have focused on using mesnchymal precursor cells (MPCs); a subset of the heterogeneous bone marrow-derived mesenchymal stem cell population that are selected based on the expression of the cell-surface antigens STRO-1, STRO-3, STRO-4, CD146 (MUC-18) and CD106 (VCAM-1) [6,7]. These cells have been shown to have beneficial properties compared to unselected MSCs [8] and have additionally shown promising results when used to treat disc degeneration. For example, a recent study using an ovine model of IVD degeneration showed that injecting MPCs into the disc resulted in some repair of the disc tissue and restoration of function [9]. To fully harness the potential of MPCs (and MSCs) for IVD regeneration, an appropriate vehicle is required to facilitate effective delivery of the cells into the disc and additionally provide an optimised environment for tissue regeneration once the cells are in situ. For such applications, hydrogels are an attractive option and hydrogels of different compositions are being increasingly investigated for NP regeneration [10e13]. Like native NP tissue, hydrogels are highly hydrated and have appropriate permeability to both oxygen and nutrients. Using controllable crosslinking chemistries, gels with different mechanical properties may be constructed. Moreover, regulating the crosslinking kinetics allows the material to be injected into the disc space prior to gelation. This not only eliminates the need for more invasive surgery but also allows for the hydrogel to mould to the exact shape of the defect, a feature which cannot be achieved using conventional pre-formed scaffolds. To date several different hydrogels have been synthesised for NP tissue-engineering applications. These use varying combinations of materials including polyethylene glycol (PEG), gellan gum, type-IIcollagen, hyaluronic acid (HA), gelatin and polyurethane with differing chemistries and could support the viability of either MSCs or cells derived from the IVD [10,14e18]. In particular, Calderon et al. [14] showed high viability of rat MSCs within a gel comprised of HA and type-II-collagen, although differentiation to a chondrogenic phenotype was only possible in the presence of medium supplemented with TGFb1. However, a further advantage of hydrogels is that they can be loaded with bioactive molecules (such as drugs and growth factors) to promote cell viability, differentiation and tissue regeneration. The semi-synthetic sulphated polysaccharide, pentosan polysulphate (PPS) has previously been shown to promote both the viability and chondrogenic differentiation of MPCs and enhance the production of matrix components that are relevant to NP regeneration, such as type-II-collagen, glycosaminoglycans (GAGs) and proteoglycans (PGs) [19]. For this reason, PPS is of interest as a molecule to promote NP regeneration by MPCs. Furthermore, when MPCs were combined with PPS and implanted (within Gelfoam scaffolds) into the disc space of an ovine IVD degeneration model, there was significantly more cartilage formation than in control groups treated with Gelfoam alone or Gelfoam combined with MPCs but in the absence of PPS [20].

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In this study we aimed to build upon the existing data that indicates strong potential coupling of MPCs and PPS as a treatment for IVD degeneration by developing an injectable hydrogel system within which both MPCs and PPS could be incorporated. It is anticipated that the development of an effective biomaterial, which is both injectable and able to provide an optimised physical environment for cell viability and differentiation, would significantly enhance the promising results already obtained and have considerable potential in the future development of an effective therapy to regenerate the NP and improve IVD function. 2. Materials and methods 2.1. Materials Hyaluronic acid of 215 kDa molecular weight was purchased from Lifecore. Amine terminated 8-arm poly(ethylene glycol) (PEG, tripentaerythritol backbone, Mw ¼ 40 kDa) was purchased from JenKem, USA. 3-(4-hydroxyphenyl)propionic acid (HPA), N-(3-dimethylaminopropyl)-N0 -ethylcarbodiimide hydrochloride (EDC), sulpho-N-hydroxysuccinimide (NHS), peroxidase, Type I from horseradish (HRP, 113 U/mg solid) were all purchased from Sigma. Sodium PPS (Batch Q18) was supplied by bene-Arzneimittel GmbH (Munich, Germany). All other reagents were purchased from Gibco unless otherwise stated. 2.2. Functionalisation of HA with tyramine to facilitate crosslinking Using methods outlined by Darr and Calabro [21], HA was reacted with tyramine hydrochloride and 1-Ethyl-3-[3-dimethylaminopropyl]carbodiimide hydrochloride/ N-hydroxysuccinimide (EDC/NHS) in a 1:10:0.1 M ratio in 0.1 M MES buffer, pH 4.8. Briefly solutions of HA and tyramine were mixed to produce the desired concentration, the pH readjusted to 4.8 and EDC/NHS added. After 15 min, sodium hydroxide was used to adjust the pH to 5.8 and the reaction left to proceed for 3 h at room temperature with gentle agitation. Unreacted products were removed by dialysis in 3500 MWCO tubing (ThermoFisher) for 48 h each against 150 mM NaCl, 10% ethanol and distilled water and the purified samples freeze-dried. 2.3. Functionalisation of PEG with HPA to facilitate crosslinking PEG was reacted with 3-4-hydroxyphenylpropionic acid (HPA) using EDC/NHS chemistry. Briefly HPA was dissolved in 0.1 M MES buffer to a concentration of 5 mM and the pH adjusted to 4.8. A solution of 500 mM EDC and 50 mM NHS was added and reacted for 15 min at room temperature. A solution of PEG in 0.1 M MES buffer was added to give a final concentration of 62.5 mM and the PH adjusted to 5.8. The reaction was left to proceed for 3 h at room temperature with gentle agitation and repeated up to four times to achieve the desired degree of functionalisation. Dialysis was performed for 24 h after each reaction in 150 mM NaCl solution using 3500 MWCO tubing. After the final reaction, of the product was dialysed for 24 h in 150 mM NaCl, 24 h in 20% ethanol and a further 24 h in distilled water was performed and the samples freeze-dried. 2.4. NMR Functionalisation of both HATYR and PEGHPA was determined by quantitative 1H NMR (Supp Fig. 1) with spectra collected on a Bruker Avance 750 high-resolution NMR spectrometer. The chemical shifts were referenced to the solvent resonance (D2O) at d ¼ 4.77 ppm. The degree of substitution of HPA on the PEG molecules (ds) was calculated from the relative integral of the methylene resonance from PEG (d ¼ 3.7 ppm) compared to that of the aromatic resonances attributed to the HPA (d ¼ 6.8 and 7.1 ppm), normalising to the number of contributing protons. To ensure the peaks observed for HPA were arising from HPA covalently bound to PEG, pulsed diffusion gradient 1H NMR was employed and the variation of the aromatic peak intensities compared to the quantitative scans against the normalised methylene peak of the PEG. Similarly, the degree of substitution of TYR on the HA-TYR conjugates was calculated by comparing the relative peak integrals of the aromatic protons of TYR (d ¼ 6.8 and 7.1 ppm) and the HA methyl protons (d ¼ 1.9 ppm). 2.5. Hydrogel formation HA/PEG hydrogels were formed by mixing 25 mg/ml HA, 100 mg/ml PEG, 10 U/ ml HRP and 100 mM H2O2 (all in solution in sterile PBS), to provide the desired concentrations. Mixtures were prepared combining all components except H2O2 and thoroughly mixing the solution by vortex. To initiate gelation, the H2O2 was added, the solution vortexed briefly and then the gels added to the required culture vessel/ rheometer as required. For the formation of hydrogels including encapsulated MPCs, the required number of MPCs were pelleted and resuspended evenly in the hydrogel mixture prior to the addition of H2O2.

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2.6. Rheology All rheological measurements were obtained using an AR G2 rheometer (TA Instruments, New Castle, DE) in oscillatory mode using a 20 mm diameter stainless steel flat plate and lower Teflon Pelletier surface. The gels were synthesised and, upon addition of the H2O2, immediately vortexed and loaded between the plates of the rheometer with a gap of 1000 mm. Gelation kinetics and final moduli were determined by time sweeps at 37  C, conducted with a controlled strain of 1% and an angular frequency of 6.28 rad/s. Frequency sweeps were performed between 5.0E3e50 rad/s at 1% strain. Stress sweeps confirmed that this was within the linear viscoelastic region (Supp Fig. 2). 2.7. Swelling and degradation To determine the degree of swelling, 50 ml gels were synthesised and the dry weight obtained after freeze-drying for 24 h (Wd). The gels were then incubated in 500 ml PBS for 24 h at 37  C. The PBS was removed and the gels re-weighed (Ws). The degree of swelling was determined by ((Ws  Wd)/Wd) using triplicate gels for each condition. Degradation of 50 ml gels was measured over a period of three months using triplicate samples for each condition. After synthesis, the gels were incubated in PBS/ 0.01% sodium azide at 37  C with weekly changes of buffer. At specific time points, all excess PBS was removed and the gels weighed. The degree of degradation was expressed as a % of the original gel mass.

dilution. Secondary antibodies conjugated to Alexa-Fluor 488 and Alexa Fluor 568 were used for detection and samples counterstained with Hoechst. 2.13. In vivo assessment Hydrogels of 150 ml were pre-formed using a silicon rubber mould and implanted subcutaneously into Wistar rats (Female, 6e12 weeks). Six replicates were performed for each gel and time point. After 7 and 14 days the animals were sacrificed and implants removed for analysis. Samples were fixed in 4% paraformaldehyde and processed for histology as described previously. Sections for each sample were taken at two different intervals of 500 mm to allow for analysis of different sites throughout the implant. Prior to staining, the samples were dewaxed in xylene and rehydrated to water through a graded ethanol series. Haematoxylin and eosin staining was performed using standard procedures. High resolution images were acquired using a slide scanner (Aperio) to allow for analysis of multiple sections [8e12] for each implant. To detect CD68þ macrophage, dewaxed and hydrated samples were treated with IHC revealer (MP Biomedicals) for 10 min at room temperature, blocked in 2%BSA/2% goat serum/0.1% Tween 20 in Tris-buffered saline (TBS) and incubated overnight with anti-CD68 (clone ED1, Serotec) at a 1:50 dilution. A secondary antibody conjugated to Alexa-Fluor 488 was used for detection and samples counterstained with Hoechst. Images were acquired using an Olympus IX81 microscope with the same exposure for images across all conditions.

3. Results 2.8. In vitro injection model A solution of 3% (w/v) agarose in PBS was made and poured into moulds. Just prior to setting, an air bubble was introduced by syringe and the agarose incubated at 4  C for a further 30 min. Samples were equilibrated at 37  C in PBS for 1 h and then a syringe used to inject HA/PEG containing 0.001% red food dye into the cavity. The samples were incubated at 37  C in PBS for a further 2 h and then dissected to examine gel formation. 2.9. MPC culture Stro3-selected human MPCs were prepared by Lonza (Walkersville, USA) for Mesoblast Ltd (Melbourne, Australia) according to the isolation procedure described by Gronthos et al. [8,22]. MPCs were cultured in alphaMEM supplemented with 100 U/ml penicillin, 100 mg/ml streptomycin (Gibco/Invitrogen Carlsbad, CA, USA), 10% batch-tested foetal bovine serum (FBS), 2 mM L-Glutamine, 1 mM Sodium Pyruvate, and 100 mM L-ascorbate-2-Phosphate. Tissue culture flasks were maintained at 37  C in 5% CO2 in an atmosphere with 95% humidity. Upon reaching 70% confluence MPCs were passaged, replating at 2000 cells/cm2. 2.10. MPC encapsulation and culture in HA/PEG hydrogels MPCs (P4-6) were encapsulated in gels containing 15 mg/ml HATYR, 16.5 mg/ml PEGHPA, 0.25 U/ml horseradish peroxidase (HRP) and varying amounts of hydrogen peroxide (H2O2). Soluble PPS was added into the gels prior to crosslinking at a final concentration of 5 mg/ml. MPCs were resuspended to a concentration of 5  106 cells/ ml in solution containing all of the components except H2O2 and thoroughly mixed. Crosslinking was then initiated by the addition of H2O2 and the gels spotted out in 50 ml spots into low-binding 12-well tissue culture plates. After 15 min, growth media was added to the well and the cell/gel composites cultured with media changes every 3 days. 2.11. Cell viability analysis MPC viability was determined by Live/Dead assay (Life Technologies) according to the manufacturer’s instructions. Staining was carried out at time points of 24 h and 7 days after encapsulation. Images were obtained using a Zeiss LSM710 confocal microscope taking z-stacks through a depth of 100 mm. 2.12. Histological analysis After 21 days of culture, hydrogels were washed with PBS and fixed in 4% PFA for 30 min at RT C. Samples were dehydrated through a graded ethanol series, incubated in xylene for 45 min and embedded in paraffin for sectioning (8 mm sections). Prior to staining, the samples were dewaxed in xylene and rehydrated to water through a graded ethanol series. Histological staining was performed using Haematoxylin and Eosin, Alcian blue, Safranin O and Picrosirius red using standard techniques. To detect Collagen-I and -II, sections were rehydrated to water, treated with 0.01% Pepsin (10 min, 37  C) and 0.1% hyaluronidase (60 min, RT C) and blocked in 2%BSA/2% goat serum in Tris-buffered saline (TBS). Sections were incubated overnight with polyclonal rabbit anti-collagen I (Cedarlane Labs, Canada) and monoclonal mouse anti-collagen II (Lab Vision, USA) primary antibodies both at a 1:50

3.1. Development and characterisation of an enzymatically crosslinked HA/PEG hydrogel system Hydrogels were formed via enzyme-mediated crosslinking of tyramine-functionalised HA (HATYR) and HPA-functionalised PEG (PEGHPA) using horseradish peroxidase (HRP) and H2O2, according to the chemistry originally described by Kurisawa and colleagues [23,24] and more recently by Menzies et al. [25]. These studies demonstrated that the concentration of both HRP and H2O2 affect the gelation speed and final mechanical strength of the resulting hydrogels. In view of this, we first optimised the composition of the HA-PEG hydrogels to produce hydrogels of differing moduli, whilst maintaining an ability to handle the solution and crosslinking kinetics that would render them suitable for future use in our targeted (IVD) clinical setting. Our initial analysis (data not shown) suggested that gels containing 0.25 U/ml HRP would gel within an appropriate timeframe and was chosen to systematically examine the effect of H2O2 concentration on the final modulus and crosslinking time of HA-PEG hydrogels at this constant concentration of HRP. Rheological characterisation confirmed that the elastic modulus of the resulting HA/PEG hydrogels could be tuned by changing the concentration of H2O2. With increasing concentrations of H2O2, the final modulus (as determined at a set time point of 5000 s after initiation of gelation) increased to a maximum of 5.5 kPa when crosslinked using 2.5 mM H2O2, subsequently decreasing with higher H2O2 concentrations (Fig. 1A). At low concentrations of H2O2 (1.5 mM H2O2 or less) the time sweep data revealed a sudden plateau in modulus, which is likely to have been caused by exhaustion of the available H2O2 (Fig. 1B). We also determined how gelation speed was affected by differing H2O2 concentrations and found that the speed of crosslinking (as determined by the time taken for the gels to reach a modulus of 1 kPa) increased as the concentration of H2O2 increased (Fig. 1C, R2 ¼ 0.919). However, the time taken for the gels to reach a modulus of 1 kPa was between 3.6 and 9.5 min across all H2O2 concentrations and was therefore deemed to be within a suitable range for future use. 3.2. Characterisation of enzymatically crosslinked HA/PEG gels incorporating pentosan polysulphate (PPS) Mesenchymal stem cell differentiation is influenced by both soluble and mechanical signalling mechanisms and any effective

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Fig. 1. Rheological testing of the effect of H2O2 concentration on crosslinking and final modulus. A. Peak G0 of HA/PEG hydrogels crosslinked using differing concentrations of H2O2. B. Time sweeps of HA/PEG hydrogels crosslinked using differing concentrations of H2O2. C. Time required to reach a G0 of 1 kPa using differing concentrations of H2O2.

tissue engineering-based strategy will incorporate both of these elements. We therefore aimed to optimise both the soluble signalling and mechanical properties of the HA/PEG hydrogel for IVD repair. Due to a lack of consensus in the elastic modulus of NP tissue (previous measurements range from 3.25 to 15.6 kPa with most in the 3e7 kPa range [26e28]) we tested hydrogels of two different moduli within this region and compared HA/PEG hydrogels alone to hydrogels incorporating PPS, a potent inducer of chondrogenic differentiation in MPCs [19]. To ensure that incorporation of PPS into the hydrogels did not change the mechanical properties, rheological testing was performed to compare the characteristics of hydrogels formed from both 4 mM (3.9 kPa) and 2.5 mM (5.5 kPa) H2O2with and without added PPS. The addition of 5 mg/ml PPS to the HA/PEG gels caused a very slight, but consistent, reduction in the resulting G0 , decreasing from 5.55 to 5.46 kPa and from 3.99 to 3.47 kPa for gels crosslinked with 2.5 mM or 4 mM H2O2 respectively (Table 1). As observed in Fig. 1, the HA/PEG gels crosslinked with the higher concentration of H2O2 were slower to reach a modulus of 1 kPa. Addition of PPS to the system also caused a decrease in gelation speed. Frequency sweeps were used to examine the viscoelastic properties of the HAPEG gels. For frequencies between 5 and 50 rad/s, the storage modulus (G0 ) was always larger than the loss modulus (G00 ) indicating that the hydrogels display a predominantly elastic behaviour (Fig. 2). Additional testing via stress sweeps confirmed that this was within the linear viscoelastic range (Supp Fig. 1). From the stress sweeps we also measured the critical strain (defined as the % strain at which the linear viscoelastic range was breached) showing a breakdown of the gels at approximately 4% and 10% for 2.5 mM and 4 mM H2O2 concentrations respectively. We then examined the swelling and degradation properties of the hydrogels. The swelling ratio for all gels was high, ranging from 21 to 27 percent after 24 h. There was a trend of decreased swelling for the gels incorporating PPS, although this was only statistically significant for the hydrogels synthesised using 2.5 mM H2O2 (Fig. 3A). Degradation profiles showed little degradation over a 3 month period, with no significant differences between any of the different gel compositions. However, it should be noted that these tests were done in the absence of cells and degradation is likely to

be enhanced in the presence of cells (as was observed, see Fig. 6) due to their secretion of hyaluronidase and other enzymes. To assess the utility of these hydrogels as an injectable system, we developed an in vitro injection model to confirm hydrogel crosslinking under physiological conditions. Using this model, we demonstrated that the HA/PEG gels can be injected through a needle into a fully-hydrated synthetic tissue (agarose) at 37  C and crosslink under these conditions (Fig. 4). Excision of the hydrogels from the agarose capsule after 2 h showed that the hydrogels were fully crosslinked and had formed a consistent matrix which retained the shape of the filled defect, even once removed from the agarose (Fig. 4C). 3.3. MPC morphology, viability and differentiation in HA/PEG hydrogels MPCs were encapsulated in the hydrogels at a density of 5  106 cells/ml. Gels were spotted onto non-adhesive tissue culture plates prior to crosslinking and subsequently maintained in standard MPC growth media (Fig. 5A). Under these conditions MPCs took on a rounded morphology that was maintained for the duration of the culture period (Fig. 5B). To determine whether the use of H2O2 during the cross-linking process had an adverse effect on MPC viability, live/dead staining was performed 24 h after encapsulation. The majority of MPCs were stained green, showing that there was good viability, equivalent to monolayer MPCs that had not come into contact with H2O2 (Fig. 5C). Additional staining after 7 days of culture showed good viability of the cells under all conditions, with very few dead (red) cells, indicating the suitability of the HA/PEG hydrogel system for long term maintenance of MPCs (Fig. 5D). We next investigated the structure of the tissues formed from the MPC/hydrogel composites, using histological analysis to identify matrix deposition and structure for samples after 21 days of culture. Haematoxylin and eosin (H þ E) staining showed that all the samples contained significant numbers of cells and that these were relatively evenly distributed throughout the matrix (Fig. 6A). Interestingly, the structure of the composites revealed that the MPCs were within pockets where the matrix had been broken

Table 1 Rheological properties of the HA/PEG hydrogels both with and without addition of 5 mg/ml PPS. HATYR (mg/ml)

PEGHPA (mg/ml)

HRP (U/ml)

H2O2 (mM)

PPS (mg/ml)

Time to 1 kPa (s)

G0 at 5000 s (Pa)

G00 at 5000 s (Pa)

Critical strain (%)

15 15 15 15

16.5 16.5 16.5 16.5

0.25 0.25 0.25 0.25

2.5 2.5 4 4

0 5 0 5

475 501 552 616

5554 5463 3992 3470

18.0 12.2 2.4 2.3

4.4 4.0 13.0 9.8

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Fig. 2. Frequency sweep testing of HA/PEG gels. Frequency sweeps conducted at a constant strain of 1% and frequencies from 5 to 50 rad/sec. A. 2 mM alone B. 2 mM þ PPS C. 4 mM alone D. 4 mM þ PPS.

down, analogous to the lacunae that chondrocytes reside within native NP tissue. Using light microscopy we confirmed that the encapsulated MPCs retained a rounded morphology, although some protrusions could be seen extending across and even wrapping around the periphery of the lacunae (Fig. 6A, arrows). All the hydrogel samples were positive for Alcian blue with areas of less intense staining present around the MPCs, which corresponded to the lacunae observed via H þ E staining (Fig. 6B). However, for the gels incorporating PPS, the area immediately surrounding the lacunae was more intensely stained for the more highly sulphated GAGs (Fig. 6B, inset), with rings of staining present in the pericellular region only in those samples containing PPS. No significant differences were observed between the 2.5 and 4 mM gels. Due to the HA content of the hydrogels, all were positive for Safranin O staining but imaging again showed areas where the MPCs had begun to remodel the matrix immediately surrounding

the cell body (Fig. 6C). Picrosirius red staining for collagens was only evident (above background levels) in the 4 mM hydrogels containing PPS. This was evident as an intense halo directly surrounding the MPC (Fig. 6D, arrows). No differences in staining were observed between any of the different gel compositions in the absence of cells and there was no evidence of degradation analogous to the ‘lacunae’ around the cells in these samples (Supp Fig. 2). Immunostaining was also performed to determine the expression of type-I and type-II-collagen. Little difference was seen in the level of type-I-collagen, although expression was somewhat reduced in 2.5 mM hydrogels incorporating PPS compared to the other samples. Collagen-II deposition was moderately increased when PPS was incorporated into the hydrogels although no significant differences were observed between the different moduli (Fig. 7A). Further examination of Collagen-II deposition in whole mount samples confirmed this finding, with z-stacks showing that

Fig. 3. Swelling and degradation properties of HA/PEG hydrogels with and without PPS. A. Degree of swelling of HA-PEG hydrogels. Data is presented as mean  SD, n ¼ 3 (*) p < 0.05. B. Degradation profiles of HA/PEG hydrogels.

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Fig. 4. In vitro injection model. A) Agarose containing a cavity to be filled by the HA/PEG hydrogel within a bath of PBS. B) Injection of hydrogel solution coloured with red food colouring (at 37 C). C) The composite immediately following injection. D) The crosslinked hydrogel inside the agarose tissue (back) and once excised from the matrix (front).

this deposition was predominantly of a pericellular localisation, with some evidence of the matrix extending more broadly throughout gels with higher collagen-II levels (Fig. 7B and inset). 3.4. In vivo response to HA/PEG hydrogels To characterise the local tissue response to these materials, hydrogels (2.5 mM, with and without PPS) in the absence of MPCs were implanted subcutaneously into a wild type rat model and explanted after seven and 14 days for analysis. At both time points

the hydrogels were clearly visible under the skin, indicating that they had not yet degraded significantly and no macroscopic signs of inflammation or toxicity were evident in the tissue surrounding the implants (Fig. 8A). Furthermore, gross examination of the spleen of the rats identified no enlargement of the organ that would be indicative of a systemic immune reaction (data not shown). Haematoxylin and Eosin staining of sections throughout the explanted tissue showed evidence of capsule formation around the implants (Fig. 8A). However, this minimal initial inflammatory response was significantly reduced by day 14 (compared to day 7)

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Fig. 5. Morphology and viability of MPCs encapsulated in HA/PEG gels. A. Examples of HA/PEG hydrogels. Scale bar ¼ 10 mm. B. MPC morphology 7 days after encapsulation. Scale bar ¼ 200 mm. C. Live/dead staining of MPCs 24 h and 7 days after encapsulation. All cells are stained green and dead cells are red. Scale bars represent 50 mm. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)

Fig. 6. Histological analysis of HA/PEG hydrogels with and without PPS. HA/PEG hydrogels were sectioned and stained after 21 days of culture with MPCs. A. Haematoxylin and eosin White arrows show examples of cells extending membrane protrusions. B. Alcian blue pH 2.5 (insets pH 1.0). C. Safranin O. D. Picrosirius red. Arrows denote areas of more intense pericellular staining. Scale bar ¼ 50 mm.

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Fig. 7. Collagen expression in HA/PEG hydrogels with and without PPS. A) Immunostaining of sections through HA/PEG hydrogels showing Collagen-I (red), Collagen-II (green) and Hoechst staining (blue). Scale bar ¼ 50 mm. B) Collagen-II staining of whole-mount HA/PEG gels. Scale bar ¼ 50 mm.

and there was no apparent infiltration of cells into the hydrogels and no evidence of foreign body giant cells. No significant differences were observed between hydrogels with and without PPS, with the exception of one of the replicates of the day seven PPScontaining hydrogels, which contained vasculature running through the implanted material (Fig. 8B, arrow). Macrophages were identified by immunostaining for CD68 (ED1) and were present immediately surrounding the implanted material. Correlating with the decrease in capsule size at 14 days, the thickness of the layer containing macrophages was reduced on day 14 compared to day seven, by which time they were present only as a thin layer directly adjacent to the implanted material. 4. Discussion We have developed and characterised an injectable hydrogel system suitable for the encapsulation of MPCs for applications in IVD regeneration. Extending upon chemistry initially developed by Kurisawa et al. [23,24] and more recently by ourselves [25], we utilised HRP and H2O2 to crosslink tyramine-functionalised HA (HATYR) and 3-4-hydroxyphenylpropionic acid-functionalised PEG (PEGHPA) and demonstrated that we could form hydrogels with the appropriate mechanical properties for IVD regeneration whilst retaining a composition that it could easily be injected through a needle. This was highlighted by our newly developed ‘in vitro injection model’, which also confirmed that the hydrogels were able to crosslink effectively under physiological conditions similar to those that would be present within the IVD. Initial testing using HATYR alone showed that hydrogels of the appropriate modulus could not be produced whilst keeping a solution of workable viscosity. It would be possible to synthesise hydrogels with the desired range of mechanical properties from PEGHPA alone [25]. However, unlike PEG, HA is a naturally occurring polymer that has been shown to be beneficial for cell survival and chondrogenesis and it is present in the NP, as well as other cartilage-based tissues [29]. Cells can directly bind to HA via cell surface receptors such as CD44 and RHAMM [30], unlike PEG which is biologically inert. Combining both biological and synthetic polymers to obtain materials with the desired mechanical and biological properties has also shown some merit in previous studies [31,32]. Therefore, this dual system of HATYR and PEGHPA was ideal in having the capability to support cells through biological

interactions, as well as being injectable and able to crosslink in an appropriate timespan to produce hydrogels with the desired range of moduli. Systematic investigation into the effect of H2O2 concentration showed that the peak modulus of 5.5 kPa was achieved using an intermediate level of H2O2 (2.5 mM). This differs from the results of Kurisawa et al., who suggest that modulus continually increases with rising H2O2 levels, due to the potential for more di-tyramine crosslinks to be formed [17]. However, bearing in mind that one molecule of H2O2 is consumed for each crosslink that forms between either two tyramine or two HPA molecules, the observed peak modulus correlates reasonably well with the with the theoretical concentration of H2O2 required to produce a stoichiometric balance between the number of tyramine and HPA moieties available for participation in the crosslinking reactions and the number of H2O2 molecules required to crosslink these (1.4 mM). Below these levels, H2O2 was limiting (as demonstrated by the abrupt plateau in our time sweep data for hydrogels crosslinked using 1.0 and 1.5 mM H2O2), but above this level the H2O2 became inhibitory, perhaps due to some inhibition of the HRP enzyme at high H2O2 concentrations. Additionally, we found that gelation speed and modulus were not independent of each other, as suggested by Kurisawa et al. However, as demonstrated by previous work in our group, this may be attributed to factors such as the concentration of the polymers, the degree of functionalisation and the presence of any residual Tyramine or HA or any inactive adducts [25]. The moduli of hydrogels incorporating PPS were similar to those without PPS and the speed of gelation was slower. It is probable that this reduction in gelation speed was caused by steric hindrance of the PPS molecule interrupting the formation of crosslinks between HA and/or PEG molecules. This is backed up by work in which we covalently linked PPS into the hydrogel matrix and observed a similar decrease in gelation kinetics (unpublished). Nonetheless, the change in gelation rate was not deemed to be a problem as it could still produce crosslinked gels well within a timeframe suitable for future clinical use. The change in modulus or gelation speed was small. We also observed a slight decrease in swelling when the gels contained PPS. These results are counterintuitive as the negative charge from the sulphate groups on the PPS would be expected to attract positively charged ions which are usually accompanied by a large number of water molecules [33]. However, this same effect was observed in hydrogels containing

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Fig. 8. In vivo implantation of HA/PEG hydrogels with and without PPS. A) Images of implants after 7 and 14 days and after fixation. Arrows show position of implants under the skin. B) Haematoxylin and eosin staining after 7 and 14 days. Scale bars ¼ 2 mm and 400 mm for low and high magnification images respectively. Arrow denotes vascularisation through implant. C) CD68 immunolocalisation after 7 and 14 days. White lines denote implant boundary. Inset images show corresponding nuclear staining. Scale bar ¼ 100 mm.

chondroitin sulphate (another sulphated GAG), which had a lower swelling ratio than gels containing HA. This was attributed an increase in the overall biopolymer concentration, as well as changes to the charge density [34]. The high viability of the encapsulated MPC population confirmed that there were no detrimental effects from the use of H2O2 to crosslink the gels. This is likely to be due to the fact that the

H2O2 is rapidly consumed by the crosslinking reaction and the cells are not exposed to it for any considerable period of time, a hypothesis supported by the fact that we have previously observed cell death in monolayer cultures treated with equivalent concentrations of H2O2. Furthermore, we also demonstrated that this high viability was maintained after 7 days, thereby confirming the ability of the HA/PEG gels to support longer term maintenance.

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Although PPS is known to enhance MPC viability [19], we observed no significant difference in cell viability between the gels with or without PPS. However, the extremely low levels of cell death in the encapsulated MPC population may mean that there would be little potential for PPS to enhance viability outcomes. To investigate the utility of this system for NP regeneration, we also investigated the structure and composition of the tissue formed after 21 days culture, comparing the different hydrogel compositions. Encouragingly, the cell/hydrogel composites showed a structure analogous to NP tissue in which the cells were present within small pockets within the gel matrix, similar to the lacunae in which NP cells are typically resident. Furthermore, in all samples the encapsulated MPCs had a rounded morphology, which again has been shown to be conducive to chondrogenic differentiation [35]. With compressive moduli of 3.9 kPa and 5.5 kPa, both of our hydrogels had mechanical properties within the range of native NP tissue, for which different studies have determined compressive elastic modulus values ranging from 3 to 15 kPa. Previous studies using hydrogels to promote chondrogenesis of MSCs suggested that matrices of lower modulus (5 vs. 10 kPa) were more conducive to differentiation [36,37] and so it was deemed important to examine more than one modulus within this range. However, although substrate stiffness or compressive modulus is a strong determinant of MSC fate [38], over the range of moduli assessed in this study, we did not find any significant difference in tissue formation. Hydrogels incorporating PPS showed enhanced deposition of some NP-associated matrix components over native hydrogels. This was demonstrated by the increase in GAG deposition around the cells, as well as increased collagen-II deposition. This is important, as a high level of collagen-II is associated with fibrocartilage, a tissue similar in structure to NP tissue, the target for this study. The observed deposition of GAGs and collagen-II is required to produce the highly hydrated environment that will restore the function of the NP in dissipating forces transmitted through the spine. In addition, these results correlate well with a previous study that showed that PPS enhances chondrogenesis and inhibits osteogenesis in MPCs [19]. As PPS is a drug that is already used to treat osteoarthritis and has regulatory approval, this makes the incorporation of PPS into the HA/PEG hydrogel system a promising formulation, although further studies will be required to translate this to the clinic. A subcutaneous rat model was used to determine the inflammatory response to the hydrogels. In this study the hydrogels were implanted in the absence of cells as incorporation of the human MPCs would trigger an immune reaction in the rat. These data provide a useful measure of the extent of reaction to the material itself. PEG is expected to be relatively inert due to its low protein adsorption and cell attachment properties and both HA and PEG have previously been shown to be well-tolerated by the body [39e41]. Our data correlated well with these expectations; although some reaction was evident, no foreign body giant cells were detected and the capsule around the cells, together with the macrophage population, decreased between days seven and 14 corresponding with the typical duration of the acute inflammatory response [42]. No significant differences were observed between the native gels and those containing PPS. However, if any differences were to occur it would be likely that the PPS would improve tolerance, as PPS has previously been shown to be chondroprotective through its ability to downregulate NFkb and other inflammatory markers [43e45]. Together these findings suggest that the reaction to the HA/PEG hydrogels is minimal and indicate that the materials should be well tolerated under their intended application when injected into the intervertebral disc. Further studies using hydrogels incorporating MPCs will be required to further examine these effects but given that MPCs are well-known

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have immune-suppressive properties [3], there is clearly significant potential for the future clinical application of this combined cellhydrogel system as a whole. 5. Conclusions We have developed a hydrogel system with tunable mechanical properties within which MPCs can be encapsulated and cultured over prolonged time periods. Within this matrix MPCs were able to synthesise ECM components relevant to NP tissue and this was moderately enhanced by the incorporation of PPS. Importantly, this was possible without the incorporation of exogenous growth factors such as TGFb, which would make translation of such systems to the clinic both more complicated and more costly. Furthermore, in vivo testing confirmed that the materials used in this system are well tolerated within a wild type rat model. Overall, this system holds significant promise in the development of a tissueengineered therapy for IVD degeneration. Acknowledgements We would like to thank Tania Banks and Yu Qian Chau for expert surgical assistance. The authors also acknowledge funding from the Australian Research Council Linkage Grants Scheme (LP 0882371) in collaboration with Mesoblast Ltd. This work was performed in part at the Queensland node of the Australian National Fabrication Facility, a company established under the National Collaborative Research Infrastructure Strategy to provide nano and microfabrication facilities for Australia’s researchers. Appendix A. Supplementary data Supplementary data related to this article can be found at http:// dx.doi.org/10.1016/j.biomaterials.2013.08.072. References [1] Pittenger MF, Mackay AM, Beck SC, Jaiswal RK, Douglas R, Mosca JD, et al. Multilineage potential of adult human mesenchymal stem cells. Science 1999;284(5411):143e7. [2] Prockop DJ. Marrow stromal cells as stem cells for nonhematopoietic tissues. Science 1997;276(5309):71e4. [3] Ren G, Zhang L, Zhao X, Xu G, Zhang Y, Roberts AI, et al. Mesenchymal stem cell-mediated immunosuppression occurs via concerted action of chemokines and nitric oxide. Cell Stem Cell. 2008;2(2):141e50. [4] Purmessur D, Schek RM, Abbott RD, Ballif BA, Godburn KE, Iatridis JC. Notochordal conditioned media from tissue increases proteoglycan accumulation and promotes a healthy nucleus pulposus phenotype in human mesenchymal stem cells. Arthritis Res Ther 2011;13(3):R81. [5] Bertolo A, Mehr M, Aebli N, Baur M, Ferguson SJ, Stoyanov JV. Influence of different commercial scaffolds on the in vitro differentiation of human mesenchymal stem cells to nucleus pulposus-like cells. Eur Spine J 2012;21(Suppl. 6):826e38. [6] Shi S, Gronthos S. Perivascular niche of postnatal mesenchymal stem cells in human bone marrow and dental pulp. J Bone Miner Res 2003;18(4):696e704. [7] Gronthos S, McCarty R, Mrozik K, Fitter S, Paton S, Menicanin D, et al. Heat shock protein-90 beta is expressed at the surface of multipotential mesenchymal precursor cells: generation of a novel monoclonal antibody, STRO-4, with specificity for mesenchymal precursor cells from human and ovine tissues. Stem Cells Dev 2009;18(9):1253e62. [8] Gronthos S, Zannettino ACW, Hay SJ, Shi S, Graves SE, Kortesidis A, et al. Molecular and cellular characterisation of highly purified stromal stem cells derived from human bone marrow. J Cell Sci 2003;116(9):1827e35. [9] Ghosh P, Moore R, Vernon-Roberts B, Goldschlager T, Pascoe D, Zannettino A, et al. Immunoselected STRO-3þ mesenchymal precursor cells and restoration of the extracellular matrix of degenerate intervertebral discs. J Neurosurg Spine 2012;16(5):479e88. [10] Chen YC, Su WY, Yang SH, Gefen A, Lin FH. In situ forming hydrogels composed of oxidized high molecular weight hyaluronic acid and gelatin for nucleus pulposus regeneration. Acta Biomater 2013;9(2):5181e93. [11] Cheng YH, Yang SH, Lin FH. Thermosensitive chitosan-gelatin-glycerol phosphate hydrogel as a controlled release system of ferulic acid for nucleus pulposus regeneration. Biomaterials 2011;32(29):6953e61.

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[12] Su WY, Chen YC, Lin FH. Injectable oxidized hyaluronic acid/adipic acid dihydrazide hydrogel for nucleus pulposus regeneration. Acta Biomater 2010;6(8):3044e55. [13] Vernengo J, Fussell GW, Smith NG, Lowman AM. Evaluation of novel injectable hydrogels for nucleus pulposus replacement. J Biomed Mater Res B Appl Biomater 2008;84(1):64e9. [14] Calderon L, Collin E, Velasco-Bayon D, Murphy M, O’Halloran D, Pandit A. Type II collagen-hyaluronan hydrogelea step towards a scaffold for intervertebral disc tissue engineering. Eur Cell Mater 2010;20:134e48. [15] Benz K, Stippich C, Osswald C, Gaissmaier C, Lembert N, Badke A, et al. Rheological and biological properties of a hydrogel support for cells intended for intervertebral disc repair. BMC Musculoskelet Disord 2012;13:54. [16] Benz K, Stippich C, Fischer L, Mohl K, Weber K, Lang J, et al. Intervertebral disc cell- and hydrogel-supported and spontaneous intervertebral disc repair in nucleotomized sheep. Eur Spine J 2012;21(9):1758e68. [17] Collin EC, Grad S, Zeugolis DI, Vinatier CS, Clouet JR, Guicheux JJ, et al. An injectable vehicle for nucleus pulposus cell-based therapy. Biomaterials 2011;32(11):2862e70. [18] Hu J, Chen B, Guo F, Du J, Gu P, Lin X, et al. Injectable silk fibroin/polyurethane composite hydrogel for nucleus pulposus replacement. J Mater Sci Mater Med 2012;23(3):711e22. [19] Ghosh P, Wu J, Shimmon S, Zannettino AC, Gronthos S, Itescu S. Pentosan polysulfate promotes proliferation and chondrogenic differentiation of adult human bone marrow-derived mesenchymal precursor cells. Arthritis Res Ther 2010;12(1):R28. [20] Goldschlager T, Ghosh P, Zannettino A, Gronthos S, Rosenfeld JV, Itescu S, et al. Cervical motion preservation using mesenchymal progenitor cells and pentosan polysulfate, a novel chondrogenic agent: preliminary study in an ovine model. Neurosurg Focus 2010;28(6):E4. [21] Darr A, Calabro A. Synthesis and characterization of tyramine-based hyaluronan hydrogels. J Mater Sci Mater Med 2009;20(1):33e44. [22] Gronthos S, Fitter S, Diamond P, Simmons PJ, Itescu S, Zannettino AC. A novel monoclonal antibody (STRO-3) identifies an isoform of tissue nonspecific alkaline phosphatase expressed by multipotent bone marrow stromal stem cells. Stem Cells Dev 2007;16(6):953e63. [23] Kurisawa M, Chung JE, Yang YY, Gao SJ, Uyama H. Injectable biodegradable hydrogels composed of hyaluronic acid-tyramine conjugates for drug delivery and tissue engineering. Chem Commun (Camb) 2005;14(34):4312e4. [24] Lee F, Chung JE, Kurisawa M. An injectable hyaluronic acid-tyramine hydrogel system for protein delivery. J Control Release 2009;134(3):186e93. [25] Menzies DJ, Cameron A, Munro T, Wolvetang E, Grondahl L, Cooper-White JJ. Tailorable cell culture platforms from enzymatically cross-linked multifunctional poly(ethylene glycol)-based hydrogels. Biomacromolecules 2013;14(2): 413e23. [26] Cloyd JM, Malhotra NR, Weng L, Chen W, Mauck RL, Elliott DM. Material properties in unconfined compression of human nucleus pulposus, injectable hyaluronic acid-based hydrogels and tissue engineering scaffolds. Eur Spine J 2007;16(11):1892e8. [27] Umehara S, Tadano S, Abumi K, Katagiri K, Kaneda K, Ukai T. Effects of degeneration on the elastic modulus distribution in the lumbar intervertebral disc. Spine (Phila Pa 1976) 1996;21(7):811e9 discussion 20.

[28] Recuerda M, Cote SP, Villemure I, Perie D. Influence of experimental protocols on the mechanical properties of the intervertebral disc in unconfined compression. J Biomech Eng 2011;133(7):071006. [29] Haberstroh K, Enz A, Zenclussen ML, Hegewald AA, Neumann K, Abbushi A, et al. Human intervertebral disc-derived cells are recruited by human serum and form nucleus pulposus-like tissue upon stimulation with TGF-beta3 or hyaluronan in vitro. Tissue Cell 2009;41(6):414e20. [30] Entwistle J, Hall CL, Turley EA. HA receptors: regulators of signalling to the cytoskeleton. J Cell Biochem 1996;61(4):569e77. [31] Lee CT, Kung PH, Lee YD. Preparation of poly(vinyl alcohol)-chondroitin sulfate hydrogel as matrices in tissue engineering. Carbohyd Polym 2005;61(3): 348e54. [32] Chung C, Burdick JA. Influence of three-dimensional hyaluronic acid microenvironments on mesenchymal stem cell chondrogenesis. Tissue Eng Part A 2009;15(2):243e54. [33] Hardingham T, Bayliss M. Proteoglycans of articular cartilage: changes in aging and in joint disease. Semin Arthritis Rheum 1990;20(3 Suppl. 1):12e33. [34] La Gatta A, Schiraldi C, D’Agostino A, Papa A, De Rosa M. Properties of newlysynthesized cationic semi-interpenetrating hydrogels containing either hyaluronan or chondroitin sulfate in a methacrylic matrix. J Funct Biomater 2012;3:225e38. [35] Gao L, McBeath R, Chen CS. Stem cell shape regulates a chondrogenic versus myogenic fate through Rac1 and N-Cadherin. Stem Cells 2010;28(3):564e72. [36] Toh WS, Lim TC, Kurisawa M, Spector M. Modulation of mesenchymal stem cell chondrogenesis in a tunable hyaluronic acid hydrogel microenvironment. Biomaterials 2012;33(15):3835e45. [37] Bryant SJ, Bender RJ, Durand KL, Anseth KS. Encapsulating chondrocytes in degrading PEG hydrogels with high modulus: engineering gel structural changes to facilitate cartilaginous tissue production. Biotechnol Bioeng 2004;86(7):747e55. [38] Engler AJ, Sen S, Sweeney HL, Discher DE. Matrix elasticity directs stem cell lineage specification. Cell. 2006;126(4):677e89. [39] Turner WS, Schmelzer E, McClelland R, Wauthier E, Chen W, Reid LM. Human hepatoblast phenotype maintained by hyaluronan hydrogels. J Biomed Mater Res B Appl Biomater 2007;82(1):156e68. [40] Jia X, Yeo Y, Clifton RJ, Jiao T, Kohane DS, Kobler JB, et al. Hyaluronic acidbased microgels and microgel networks for vocal fold regeneration. Biomacromolecules 2006;7(12):3336e44. [41] Scott MD, Murad KL. Cellular camouflage: fooling the immune system with polymers. Curr Pharm Des 1998;4(6):423e38. [42] Anderson JM, Rodriguez A, Chang DT. Foreign body reaction to biomaterials. Semin Immunol 2008;20(2):86e100. [43] Lupia E, Zheng F, Grosjean F, Tack I, Doublier S, Elliot SJ, et al. Pentosan polysulfate inhibits atherosclerosis in Watanabe heritable hyperlipidemic rabbits: differential modulation of metalloproteinase-2 and -9. Lab Invest 2012;92(2):236e45. [44] Troeberg L, Fushimi K, Khokha R, Emonard H, Ghosh P, Nagase H. Calcium pentosan polysulfate is a multifaceted exosite inhibitor of aggrecanases. FASEB J 2008;22(10):3515e24. [45] Troeberg L, Mulloy B, Ghosh P, Lee MH, Murphy G, Nagase H. Pentosan polysulfate increases affinity between ADAMTS-5 and TIMP-3 through formation of an electrostatically driven trimolecular complex. Biochem J 2012;443(1):307e15.