TISSUE ENGINEERING: Part A Volume 15, Number 7, 2009 ª Mary Ann Liebert, Inc. DOI: 10.1089=ten.tea.2008.0203 Original Articles Assessment of Polymer=Bioactive Glass-Composite Microporous Spheres for Tissue Regeneration Applications Hussila Keshaw, M.Sc.,1 George Georgiou, Ph.D.,2 Jonny J. Blaker, Ph.D.,1 Alastair Forbes, M.D.,3 Jonathan C. Knowles, Ph.D.,2 and Richard M. Day, Ph.D.1,3 Conformable scaffold materials capable of rapid vascularization and tissue infiltration would be of value in the therapy of inaccessible wounds. Microporous spheres of poly(D,L-lactide-co-glycolide) (PLGA) containing bioactive glass (BG) were prepared using a thermally induced phase separation (TIPS) technique, and the bioactivity, in vitro degradation, and tissue integration of the microporous spheres were assessed. Microporous spheres containing 10% (w=w) BG stimulated a significant increase in vascular endothelial growth factor secretion from myofibroblasts consistently over a 10-day period ( p < 0.01) compared with the neat PLGA microporous spheres. The microporous spheres degraded steadily in vitro over a 16-week period, with the neat PLGA microporous spheres retaining 82% of their original weight and microporous spheres containing 10% (w=w) BG retaining 77%. Both types of microporous spheres followed a similar pattern of size reduction throughout the degradation study, resulting in a 23% and 20% reduction after 16 weeks for the neat PLGA microporous spheres and PLGA microporous spheres containing 10% (w=w) BG, respectively ( p < 0.01). After in vivo implantation into a subcutaneous wound model, the TIPS microporous spheres became rapidly integrated (interspherically and intraspherically) with host tissue, including vascularization of voids inside the microporous sphere. The unique properties of TIPS microporous spheres make them ideally suited for regenerative medicine applications where tissue augmentation is required. Introduction I n regenerative medicine, bioresorbable polymer scaffolds are used to provide a provisional matrix to guide the growth of cells until complete replacement by host tissue is achieved. Ideally, the scaffold structure and its constituent biomaterial should create an optimal environment to integrate and direct tissue regeneration. Conformable scaffolds for guided tissue regeneration are advantageous for applying to inaccessible tissue defects, such as undermining partial-thickness or full-thickness cutaneous wounds and gastrointestinal fistulae, due to their ability to completely fill the space and be in direct contact with host tissue surfaces, thus facilitating cell infiltration from surrounding tissue. Microspheres are ideal structures for filling inaccessible tissue defects because they can be efficiently packed into asymmetrical spaces. Once implanted, microspheres can act as a scaffold, with predictable interstices produced between adjacent spheres guiding tissue infiltration. As with any tissue engineering scaffold, microspheres should have suitable surface properties that are able to direct tissue in-growth, combined with appropriate mechanical and degradation properties. If the scaffold is resorbable it should also be eventually replaced by the host tissue.1 Poly(D,L-lactideco-glycolide) (PLGA) is a bioresorbable copolymer frequently used in tissue engineering applications, with mechanical and degradation properties controlled by adjusting the molecular weight and copolymer ratio.1–3 Neovascularization is an essential component of wound healing and tissue regeneration, replacing damaged capillaries and reestablishing a supply of oxygen and nutrients. The porosity of a scaffold will dictate the extent of vascular infiltration from host tissue. Targeted delivery of angiogenic agents can be desirable, especially when systemic delivery of the agent could cause damage elsewhere in the body. The introduction of angiogenic growth factors directly into chronic wounds has demonstrated a positive effect on accelerating chronic wound healing. Examples include plateletderived growth factor, available as a topical gel (Becaplermin, [Regranex], Janssen-Cilag Ltd, Buckinghamshire, UK) and 1 Biomaterials and Tissue Engineering Group, Burdette Institute of Gastrointestinal Nursing, Kings College London, London, United Kingdom. Division of Biomaterials and Tissue Engineering, Eastman Dental Institute, University College London, London, United Kingdom. Biomaterials and Tissue Engineering Group, Centre for Gastroenterology & Nutrition, University College London, London, United Kingdom. 2 3 1451 1452 licensed as an adjunct treatment for full-thickness diabetic ulcers. Enhanced healing and angiogenesis after the introduction of naked plasmid DNA encoding the gene for vascular endothelial growth factor (VEGF) has also been achieved in selected patients with ulcers due to vascular occlusive disease.4 Stimulation of angiogenesis both in vivo and in vitro using bioactive glass (BG) has also been reported.5–7 Incorporation of BG into polymer composites for use as an angiogenic stimulus is advantageous because it avoids the risk of denaturing angiogenic peptides with solvents during scaffold fabrication processes. A conformable scaffold material capable of rapid vascularization and tissue infiltration to promote healing of chronic deep inaccessible wounds would be of therapeutic value. Novel porous PLGA microporous spheres containing BG were fabricated using a thermally induced phase separation (TIPS) process, resulting in highly porous structures. The biological activity and mechanical properties of the microporous spheres were assessed, along with their ability to integrate with host tissue in a wound model. Materials and Methods Preparation of PLGA TIPS microporous spheres PLGA (75:25) (Purasorb
PDLG 7507 0.63 dL=g IV; Purac Biomaterials, Gorinchem, The Netherlands) was selected for the study due to its well-characterized properties in tissue engineering applications.1–3 The polymer was dissolved in dimethyl carbonate (>99.9% purity; Sigma-Aldrich, Poole, UK) under magnetic stirring to produce a polymer weight– to–solvent volume ratio of 16% (w=v). The neat PLGA TIPS microporous spheres were prepared by manually delivering the PLGA solution drop-wise from a syringe fitted with a stainless steel nozzle (outer diameter, 0.35 mm; inner diameter, 0.17 mm) into liquid nitrogen to induce phase separation between the polymer and the crystallizing solvent as rapidly as possible.8 PLGA TIPS microporous spheres containing BG were produced by mixing 45S5 BG particles (mean particle size of 4 mm and identical in composition to 45S5 Bioglass
[45% SiO2, 24.5% Na2O, 24.5% CaO, 6% P2O5 wt%];9 a kind gift from Schott Glass, Mainz, Germany) into the polymer solution to produce 10% w=w BG:PLGA. The 45S5 BG was selected for the current study due to its reported ability to stimulate secretion of VEGF.5–7 The solution was sonicated for 20 min to disperse glass particle aggregates and mixed at 200 rpm to ensure homogenous distribution of the BG particles in the polymer solution. The 10% w=w solution was further diluted in the neat PLGA solution to produce 0.1% and 1% w=w BG in PLGA. Control microporous spheres consisting of poly(e-caprolactone) (PCL) were also prepared using the TIPS process. PCL was added to dimethyl carbonate at a ratio of 1:6 w=v, briefly heated in a water bath to 608C to assist polymer dissolution, and stirred at 200 rpm until it had completely dissolved. BG:PLGA and PCL solutions were dropped into liquid nitrogen as described for the neat PLGA. The frozen microporous spheres were subsequently transferred in a polyethylene container to a freezedryer (Edwards Modulyo, Edwards, Crawley, UK) and sublimated overnight to yield the TIPS microporous spheres. The microporous spheres were UV sterilized for 30 min before use. KESHAW ET AL. In vitro assessment of PLGA TIPS microporous spheres VEGF secretion from fibroblasts cultured with microporous spheres. Secretion of VEGF and cell viability was assessed using CCD-18Co myofibroblasts derived from human colon (passages 14–20; CRL-1459; American Type Culture Collection, Rockville, MD). This cell line was selected because of the involvement of myofibroblasts in gastrointestinal fistula healing. Cells were seeded into wells of a 48-well plate at a density of 1
104 cells=well in 500 mL complete medium (Eagle’s minimum essential medium [EMEM; Sigma, Poole, UK] supplemented with 10% fetal bovine serum [Gibco, Paisley, UK], 2 mM L-glutamine [Sigma], 1 mM sodium pyruvate [Sigma], 1% nonessential amino acids, 50 U=mL penicillin, and 50 mg=mL streptomycin [Gibco]), and cultured for 4 days. Before coculturing, the microporous spheres were immersed in phosphate buffered saline (PBS; at 0.13 M, pH 7.4), and air within the porous microporous spheres was removed under vacuum. When the vacuum was removed, the microporous spheres became impregnated with PBS, and sank. Thirty-five TIPS microporous spheres (PLGA, PLGA-BG, or PCL) were transferred into wells containing 500 mL of fresh complete medium in replicates of five. The cells were incubated at 378C in 5% CO2–95% humidity. Conditioned medium was collected from each of the wells and replaced with fresh medium at 1-day intervals for a period of 10 days. Collected medium was stored at 708C until further analysis. The amount of VEGF secreted from the cells cultured with the different types of microporous spheres over the 10day study period was determined using quantitative sandwich enzyme immunoassays (Quantikine
human VEGF; R&D Systems, Abingdon, UK) performed according to the manufacturer’s instructions. Viability of cells cultured with PLGA TIPS microporous spheres. The viability of cells cultured with the microporous spheres was assessed after 10 days using the MTT assay.10 After collection of supernatant on day 10, fresh medium containing 0.5 mg=mL MTT was added to each well and incubated for 4 h at 378C. The resulting formazan product was solubilized with 20% sodium dodecyl sulfate:formamide (1:1) overnight. An aliquot (100 mL) was taken from each well and added to a 96-well plate, and the optical density was measured at 570 nm using a microplate reader. In vitro degradation of PLGA TIPS microporous spheres. An equal number of dry microporous spheres (neat or containing 10% (w=w) BG; n ¼ 30) were weighed (W0) using a four-place digital balance (Mettler Toledo Classic, Mettler-Toledo Ltd., Leicester, UK). The microporous spheres were immersed in PBS, and air within the microporous spheres was removed under vacuum to ensure the degradation medium permeated the porous structure of the microporous spheres. The microporous spheres were placed in 15 mL polypropylene conical tubes containing 10 mL of PBS. The samples were degraded in vitro at 378C for up to 16 weeks, in triplicate. The pH of the solution for each degrading sample was measured at weekly intervals, at which point half of the solution was replaced with 5 mL of fresh PBS. After selected degradation times the microporous spheres were removed from the tubes and weighed (W1) TIPS MICROPOROUS SPHERES after surface blotting on filter paper to remove excess PBS. The samples were then washed in deionized water and vacuum-dried overnight at room temperature before being weighed (W2) again. Percentage water absorption (WA) and percentage weight remaining of microspheres after degradation in PBS (WC) of the microporous spheres were calculated at each time point, using the following equations, respectively: WA ¼ WC ¼ W1  W0 · 100 W0 W2 · 100 W0 Changes to the size of the microporous spheres during degradation were measured from photomicrographs using image analysis software (Image-Pro Plus, Media Cybernetics, Inc., Maryland, USA). A total of 30 microporous spheres were measured at each time point, and the data presented as the mean  the standard error of the mean. Mechanical testing of PLGA TIPS microporous spheres. Changes in the compressive mechanical properties of the PLGA and BG composite TIPS microporous spheres were determined after 0, 1, 2, 4, and 6 weeks of degradation in PBS. The compressive mechanical property of vacuum-dried microporous spheres was measured using a Dynamic Mechanical Analyzer 7e (PerkinElmer Instruments, Massachusetts, USA) operated in the static stress scan mode. Tests were performed on individual microporous spheres at 378C using a parallel plate (rectangle) measuring system. Static force was applied from 1 to 8000 mN at a rate of 500 mN=min. The crosshead speed (mN=min) of the machine was configured for the test and remained constant. However, the geometry of each specimen was taken into account each time. Because the specimens were spherical, the geometry was approximated to that of a cube, and the square of the diameter of each sphere was assumed to be the cross-sectional area. The static modulus of the microporous spheres was determined at 30% strain and plotted as a function of time. The modulus was measured at a given percentage strain because the microporous spheres did not exhibit any sign of elastic behavior. The percentage strain value was given as a point of reference to compare the stiffness properties between each specimen tested. Measurements were taken in replicates of four, and the mean value  the standard error of the mean plotted. Determination of the density and porosity of PLGA TIPS microporous spheres. Density measurements on the porous spheres were taken using a Helium Pycnometer (AccuPyc 1330; Micrometrics, Dunstable, UK), as previously described.11 The envelope and foam densities were measured using an envelope density analyzer (GeoPyc 1360; Micrometrics).11 The GeoPyc determines the envelope volume by measuring the travel of a plunger into a cylinder containing a mixture of sample (at least 20 porous spheres) and graphite powder, which is tapped during measurement. The porosity was determined by dividing sample mass by the envelope density (foam density). Structural morphology of PLGA TIPS microporous spheres. Microporous sphere morphology at each degra- 1453 dation time point was assessed by scanning electron microscopy (SEM). To examine the interior, microporous spheres were bisected with a razor blade. Microporous spheres were mounted onto aluminium stubs via adhesive carbon tabs and sputter coated with gold–palladium alloy for 3 min in an argon atmosphere and viewed under a scanning electron microscope ( JEOL JSM 550LV operated at 20 kV). In vivo assessment of PLGA TIPS microporous spheres Implantation of PLGA TIPS microporous spheres. Implantation studies were performed in compliance with the Animals (Scientific Procedures) Act 1986 on male Wistar rats weighing between 200 and 250 g. All animals were fed on a commercial standard pelleted diet. Rats were anaesthetized with Hypnorm 0.4 mL=kg (fentanyl citrate and fluanisone) and diazepam 5 mg=kg. Twenty neat PLGA TIPS microporous spheres or PLGA TIPS-BG microporous spheres, sterilized by ultraviolet light, were then placed into subcutaneous pockets created on the ventral aspect of each rat and closed with 3=0 Mersilk
sutures (Ethicon
, Gargrave, UK). Twelve rats per group were kept under standard laboratory conditions until sacrifice at 1, 2, 4, and 6 weeks, when the tissue containing the microporous spheres was harvested. The resected tissue constructs were placed into 10% buffered formalin and embedded into paraffin wax for light microscopy. Histological assessment of implanted microporous spheres. Five-micrometer tissue sections were cut and stained with hematoxylin and eosin for histological assessment by light microscopy. Neovascularization was assessed in tissue that had infiltrated the voids inside the microporous spheres. Only clearly delineated voids were selected for assessment. Quantification of blood vessel density was conducted as previously described.7,12 Briefly, blood vessels were identified by the inclusion of erythrocytes within the blood vessel lumen. The number of blood vessels was quantified using a 25-point Chalkley point eyepiece graticule (Graticules, Tonbridge Wells, UK) at a magnification of
250. The graticule was rotated so that the maximum number of graticule points overlaid the blood vessels present in the field of view. The mean of nine Chalkley counts was generated for each type of microporous spheres implanted and used for statistical analysis. The counting was conducted in a blinded manner regarding the inclusion of BG in the PLGA microporous spheres. Data analysis Data were expressed as mean  standard error of the indicated number of observations. Statistical comparisons between groups were performed using a two-tailed unpaired t-test or ANOVA followed by Dunnet’s post hoc test. Differences were considered significant when p < 0.05. Results Microporous sphere morphology The neat PLGA microporous spheres and PLGA microporous spheres containing 10% BG were prepared by solid– liquid phase separation and freeze-drying. The mean diameter of microporous spheres (n ¼ 30), measured by light 1454 KESHAW ET AL. Table 1. Density and Porosity of PLGA TIPS Microporous Spheres Envelope density (g=cm3) Porosity (%) Specific pore volume (cm3=g) 1.26 0.235  0.005 81.3 3.45 1.33 0.232  0.004 82.6 3.56 In vitro characterization microscopy and image analysis software, was 1.91  0.02 mm and 1.82  0.01 mm for the neat and 10% BG microporous spheres, respectively. The porosity of the microporous spheres was 81.3% and 82.6% for the neat and 10% BG microporous spheres, respectively (Table 1). The surfaces of both types of microporous spheres were similar, consisting of a skin about 2 mm thick containing pores ranging from approximately 1–5 mm, frequently arranged in a chevron-like pattern. Cross-sectioned neat microporous spheres or microporous spheres containing 10% BG showed similar highly ordered interconnected tubular morphologies, ranging from approximately 10 to 50 mm, with a ladder-like substructure that was orientated in a radial Secretion of VEGF from cells cultured with microporous spheres. The secretion of VEGF from cells cultured with microporous spheres containing different quantities of BG was assessed over a 10-day period (Fig. 2a). Between days 2 and 10, all compositions of PLGA TIPS microporous spheres stimulated a significant increase in VEGF secretion compared with control cells ( p < 0.01). Although all of the PLGA microporous spheres containing BG stimulated a significant increase in VEGF secretion compared with the neat PLGA microporous spheres, only microporous spheres containing 10% BG produced a significant increase throughout the whole study period ( p < 0.01). PCL microporous spheres, Cells only PLGA + 0% BG PLGA + 0.1% BG PLGA + 1% BG PLGA + 10% BG PCL 400 VEGF (pg/ml) Neat PLGA 10% (w=w) BG–filled PLGA Absolute density (g=cm3) pattern (Fig. 1). Voids were present toward the center of the microporous spheres that were connected to the exterior surface via a neck (Fig. 1a, b). Pores that passed through the microporous sphere also opened out into the void. Pore volume in the BG composite microporous spheres was similar to that of the neat microporous spheres, but the walls of pores contained evenly distributed BG particles. 300 200 100 0 0 1 2 3 4 5 (a) 6 7 8 9 10 Day ** 25000 Cell number 20000 ** 15000 ** 10000 5000 L PC B 10 % PL G A + + G A PL + A G PL G B G 1% 1% 0. 0% + G A PL (b) B B G ly on ls el C FIG. 1. SEM showing the typical morphology of bisected TIPS microporous spheres. (a) The microporous sphere surface consists of a skin about 2 mm thick with pores arranged in a chevron-like pattern. The interior of the microporous spheres shows a highly ordered interconnected tubular morphology with a ladder-like substructure orientated in a radial pattern toward a void (v) inside the microporous sphere that is also connected to the exterior surface via a neck. (b) Pores passing through the microporous sphere open out into the void (v). (c) The walls of pores in TIPSBG microporous spheres contain evenly distributed BG particles (*). G 0 FIG. 2. (a) VEGF secretion from myofibroblasts in response to PLGA microporous spheres containing different concentration of BG or the neat PCL microporous spheres. (b) Cell viability in response to microporous spheres containing different concentrations of BG. All types of microporous spheres produced a significant reduction in cell viability compared with unstimulated control cells ( p < 0.01). Significantly more viable cells were associated with PLGA microporous spheres containing 1% and 10% BG than with the neat PLGA microporous spheres. TIPS MICROPOROUS SPHERES included as a negative control, did not stimulate a significant increase in VEGF secretion, yielding values similar to control cells. Cell viability. The effect of different microporous sphere compositions on the number of viable cells was assessed at the end of the 10-day culture period using the MTT assay (Fig. 2b). All of the different microporous spheres tested produced a significant reduction in the number of viable cells compared with control cells ( p < 0.01), but viability improved with increasing concentrations of BG. Cell viability in response to PLGA microporous spheres containing 1% and 10% BG was significantly greater than that to the neat PLGA microporous spheres. PCL microporous spheres led to a significant decrease in cell viability ( p < 0.01). Based on results from the in vitro cell culture studies, PLGA TIPS-BG microporous spheres containing 10% w=w BG were used for the subsequent detailed characterization and in vivo studies. Degradation of PLGA TIPS microporous spheres. The morphology of both types of TIPS microporous spheres was comparable up to 9 weeks, with the surface porosity and highly ordered interconnected tubular morphology being similar to nondegraded microporous spheres. At 9 weeks, the skin of the microporous spheres appeared slightly thicker, and the pore widths reduced. At 12 weeks, the tubular morphology and ladder-like substructure were still evident in bisected microporous spheres, but the wrinkled surface of the microporous spheres was markedly different, and the small pores arranged in chevron-like pattern had been replaced by a more open porous structure (Fig. 3). The neat PLGA TIPS microporous spheres exhibited a mild and gradual weight loss over the 16-week degradation period, retaining 82.24  2.38% of the starting weight after 16 weeks of degradation in PBS (Fig. 4a). The PLGA TIPS microporous spheres containing 10% BG followed a similar weight loss profile to the neat PLGA microporous spheres, with 76.99  2.61% of the starting weight retained at 16 weeks. The reduction of microporous sphere weight correlated with an overall reduction in size of the microporous spheres (Fig. 4b). Both types of microporous spheres followed a similar pattern of size reduction throughout the degradation study. After 1 week, the size of the neat PLGA microporous spheres was reduced by 15.94  1.05%, and the PLGA microporous spheres containing 10% BG by 17.12  0.93% compared with their original size ( p < 0.01 for both). The maximum reduction in size for both types of microporous spheres occurred after 9 weeks, when the size of microporous spheres was reduced by 26.01  0.84% and 27.82  0.91% for the neat PLGA microporous spheres and PLGA microporous spheres containing BG, respectively ( p < 0.01). After 9 weeks, the size of microporous spheres gradually increased until the end of the study at 16 weeks, when the sizes were reduced by 22.84  0.96% and 20.13  0.95% for the neat PLGA microporous spheres and PLGA microporous spheres containing 10% BG, respectively ( p < 0.01). The neat PLGA TIPS microporous spheres showed a greater initial capacity for water absorption (a weight increase of 285.92  7.92% at day 0 after immersion in PBS compared with their dry weight) than the microporous 1455 spheres containing 10% BG (a weight increase of 246.89  7.81%) at the same time point ( p < 0.05) (Fig. 4c). Water absorption by both types of microporous spheres subsequently decreased from the beginning of study until week 9, when absorption was significantly lower for PLGA microporous spheres containing 10% BG (down to 58.19  0.87%) than for the neat PLGA microporous spheres (down to 89.51  1.41%) ( p < 0.0001). After week 9, water absorption steadily increased again for both types of microporous spheres, reaching 210.96  19.93% and 143.99  5.30% at the end of the study for the neat PLGA microporous spheres and PLGA microporous spheres containing 10% BG, respectively ( p < 0.05). Changes to the pH of the degradation medium for both types of microporous spheres are shown in Figure 4d. The pH of the degradation medium was lower than the starting value (7.4) for both types of microporous spheres at all time points except at 4 weeks, when the pH for both types of microporous spheres increased to between 7.4 and 7.5. The pH was generally higher for microporous spheres containing 10% BG than for the neat PLGA microporous spheres. A drop in pH was recorded at 9 weeks for both types of microporous spheres, after which the pH steadily began to rise before dropping again at 16 weeks. Compressive mechanical tests were performed on the microporous spheres after degradation for 0, 1, 2, 4, and 6 weeks in PBS, corresponding with the in vivo implantation time points. The modulus was increased for both types of microporous spheres throughout the degradation study compared with nondegraded microporous spheres (Fig. 5). After 6 weeks of degradation, the modulus value of PLGA TIPS microporous spheres containing 10% BG was significantly higher than that of the neat PLGA microporous spheres at the same time point ( p < 0.001). In vivo studies Histological assessment of implanted microporous spheres. Microporous spheres (neat PLGA microporous spheres or PLGA microporous spheres containing 10% (w=w) BG) were implanted into subcutaneous pockets created on the ventral aspect of each rat to simulate filling of an undulating wound. At predetermined time points (1, 2, 4, and 6 weeks) the implants and surrounding tissue were resected and processed for histological analysis. The microporous spheres were well tolerated at all time points studied, with no macroscopic differences between the neat PLGA microporous spheres and those containing 10% BG. Six weeks after implantation, degradation of the microporous spheres was evident by an obvious reduction in size compared with their size preimplantation. The microporous spheres were initially implanted as a multilayered cluster, but the loose skin of the rodent wound model resulted in movement of the microporous spheres after implantation. This led to the majority of implanted microporous spheres resting as a single layer rather than maintaining their original cluster formation. Within 1 week of implantation tissue had infiltrated the interstices between packed microporous spheres. This consisted mainly of fibrovascular tissue (Fig. 6). There was no apparent difference between the neat PLGA microporous spheres and those containing 10% BG. Higher magnification revealed cells from the surrounding tissue infiltrating the radial tubular macropores originating at the 1456 surface of the microporous spheres (Fig. 6b). Fibrovascular tissue remained close to the surface of the microporous spheres at all time points studied, becoming denser 6 weeks after implantation. At 1 week postimplantation cells were visible in the voids present toward the center of the microporous spheres. These were completely filled by fibrovascular tissue 2 weeks after implantation (Fig. 6c, d). Quantitative assessment of the number of blood vessels infiltrating the voids at 2 weeks postimplantation revealed no significant difference between the neat PLGA microporous spheres and those containing 10% BG (Fig. 6e). KESHAW ET AL. Discussion Healing of inaccessible wounds that also require tissue augmentation could be accelerated using conformable scaffolds capable of promoting rapid tissue infiltration. Microspheres are ideal structures for creating porous scaffolds for guided tissue regeneration by readily conforming to the shape of the void to be filled. Microsphere-based scaffolds have been investigated for a wide variety of tissue engineering applications, including bone,13 cartilage,14 adipose tissue,15 and skin.16 Many of these studies have involved FIG. 3. SEM of the neat PLGA and PLGA-BG TIPS microporous spheres. Surface porosity, with pores arranged in chevronlike patterns, is similar for both types of microporous spheres up to week 9. At week 12 the surface of both types of microporous spheres appears distorted with the ordered porosity being replaced by a more rugose topography. Bisected microporous spheres reveal the highly ordered interconnected tubular morphologies with a ladder-like substructure largely intact after 12 weeks of degradation. TIPS MICROPOROUS SPHERES 1457 Micro-porous sphere diameter (mm) 100 %i Weight remaining 80 60 40 20 PLGA + 0% BG PLGA + 10% BG 0 0 2 (a) 4 6 8 10 12 14 2.0 PLGA + 0% BG 1.8 PLGA + 10% BG 1.6 1.4 1.2 1.0 16 0 2 4 (b) Degradation time (Weeks) 6 8 10 12 14 16 Degradation time (Weeks) 8.0 350 300 7.6 PLGA + 10% BG 250 7.2 pH % Water Absorption PLGA + 0% BG PLGA + 10% BG PLGA + 0% BG 200 6.8 150 100 6.4 50 6.0 0 0 (c) 2 4 6 8 10 12 14 16 Degradation time (Weeks) 0 2 (d) 4 6 8 10 12 14 16 Degradation time (Weeks) FIG. 4. (a) Percentage weight change of the neat PLGA or PLGA-BG microporous spheres after degradation in PBS. (b) Change in size of the neat PLGA or PLGA-BG microporous spheres after degradation in PBS. Both types of microporous spheres follow similar pattern of size reduction. (c) Water absorption by the neat PLGA or PLGA-BG microporous spheres after degradation in PBS. (d) pH of the degradation medium remained above 7.0 throughout the study period. The pH for PLGA microporous spheres containing BG was slightly increased compared with the neat PLGA microporous spheres. 20 PLGA + 0% BG *** PLGA + 10% BG Static modulus (MPa) using solid microspheres fabricated using conventional oil-in-water emulsion and solvent extraction=evaporation techniques. Despite providing conformable scaffolds, solid microspheres fail to promote both rapid interspherical and intraspherical tissue integration. The microporous spheres described in the current study were produced using TIPS, resulting in highly porous structures.8 Compared with solid microspheres, TIPS microporous spheres of an equal size contain less polymer material (up to 90% less). As a consequence much less degradation product is released at the implantation site as the microporous spheres degrade. This is an important feature, especially with materials such as aliphatic polyesters, including PLGA, that degrade by ester hydrolysis releasing acidic compounds capable of stimulating an inflammatory response at the implant site. Moreover, degradation of solid microspheres is accelerated by autocatalysis. Diffusion of aqueous fluids and their subsequent entrapment in the interior of solid microspheres leads to bulk degradation. If the acidic degradation products cannot readily escape from within the system, this leads to more rapid, protoncatalyzed, polymer degradation.17–19 In contrast to solid microspheres, the porous structure of TIPS microporous spheres investigated in the current study makes them more 15 10 5 0 0 1 2 Time (weeks) 4 6 FIG. 5. Compressive mechanical analysis of the neat PLGA and PLGA microporous spheres containing 10% BG after degradation in PBS. The static modulus was similar for both sets of microporous spheres during degradation, except for microporous spheres containing 10% BG after 6 weeks when the modulus was significantly greater compared with the neat PLGA microporous spheres at the same time point ( p < 0.001). 1458 KESHAW ET AL. FIG. 6. Histological analysis of microporous spheres implanted into subcutaneous tissue. (a) Tissue rapidly infiltrates interstices between packed microporous spheres (neat PLGA microporous spheres; 2 weeks postimplantation). (b) Cells (arrows) also rapidly infiltrate the radial tubular macropores originating at the surface of the microporous spheres with their migration being directed by the orientation of pores (direction of arrows) (neat PLGA microporous spheres; 1 week postimplantation). (c, d) Voids inside the microporous spheres became rapidly filled by fibrovascular tissue (PLGA microporous spheres containing 10% (w=w) BG; 2 weeks postimplantation). (e) Number of blood vessels counted in the voids of microporous spheres 2 weeks after subcutaneous implantation. Blood vessels were counted using a 25-point Chalkley point eyepiece graticule at a magnification of
250. prone to degrade through surface erosion, and allows the acidic degradation products to diffuse away, reducing autocatalysis. A comparison of the degradation kinetics of the two types of TIPS microporous spheres was determined in vitro as a function of the hydrolysis time in PBS. A steady decrease in microporous sphere weight and pH was observed during the 16-week degradation period, rather than a sudden drop in weight and pH typical of autocatalysis.20,21 Even though the pH fluctuated during the degradation study, it did not drop below 7.0, indicating that such varia- tions are unlikely to cause any significant physiological effect when implanted in vivo. The slightly greater weight loss (*5%) by the PLGA microporous spheres containing 10% BG than by the neat PLGA microporous spheres was probably caused by the dissolution and loss of glass particles from the microporous spheres. Similar effects on weight loss have been reported in other studies investigating PLGA TIPS foams containing BG particles.22 The exterior of the TIPS microporous spheres consisted of a skin about 2 mm thick with a semi-smooth surface containing TIPS MICROPOROUS SPHERES pores (1–5 mm) that were frequently arranged in chevron-like patterns, suggested to be caused by the initial freeze front of the solvent across the droplet surface.8 As the freeze fronts progress toward the center of the microporous sphere, the pore structure becomes more ordered, interconnected, and ladder-like. These structural properties make the microporous spheres quite rigid, despite their porosity, and help to maintain their integrity and thus that of the scaffold as a whole during degradation. The presence of BG did not significantly affect the static modulus of the microporous spheres, except after 6 weeks of degradation. The significant increase in modulus for the microporous spheres containing 10% BG at 6 weeks could result from densification of the porous spheres, as suggested by SEM of cross-sectioned spheres at 6 weeks and beyond. However, the percentage weight remaining and the change in diameter at 6 weeks do not show any significant difference between the two types of microporous spheres, indicating that the apparent density of the microporous spheres would not be dissimilar. Water absorption by TIPS microporous spheres was assessed according to a method previously described for TIPS foams.22 Although the assessment of water absorption in the current study reflected fluid trapped in the microporous sphere pores rather than fluid absorption by the pore walls, the results obtained correspond with what was happening to the size of the microporous spheres; that is, as the microporous spheres became reduced in size less, fluid was trapped inside. The neat PLGA TIPS microporous spheres showed a greater capacity for water absorption than microporous spheres containing 10% BG. Reduced pore volume in TIPS foams containing BG has previously been reported;22 therefore, the differences in water absorption occurring from the beginning of the degradation study onward probably result from a smaller volume of bulk fluid being trapped in the pores. The overall smaller values for water absorption in the current study compared with previous studies reflect differences in the porosities of the materials assessed. Previous studies have used higher polymer weight–to–solvent volume ratios resulting in high porosities (>90%) and therefore a greater capacity to trap water in the pores.22,23 Further, the microporous spheres assessed retained their skin. This is less porous than TIPS foam specimens assessed elsewhere, which have been trimmed to expose their more porous internal structure.22 Morphological analysis of the degraded microporous spheres at 9 weeks revealed thickened skins and shrinkage of the interconnected porous network. These structural changes are likely to have caused expulsion of fluid from the microporous spheres and could account for the decreased water absorption seen at 9 weeks. Shrinkage of the microporous spheres, observed both in vitro and in vivo, is likely to have resulted from stresses in the aligned pores radiating from the center of the microporous spheres during degradation, similar to that suggested for the shrinkage of cylindrical disks of TIPS PLGA foam scaffolds.23 Subsequent plasticization due to the presence of water between the polymer chains is likely to have further facilitated their shrinkage in the short term. The surface of the microporous spheres at 12 weeks was blistered with larger pores resulting from polymer degradation. The blistering effect may have caused the observed increase in microporous sphere size, which would allow more fluid to 1459 enter the microporous spheres and account for the increase in water absorption seen at 12 weeks and beyond. Previous studies investigating soft tissue integration with monolithic TIPS polymer foam scaffolds have raised questions regarding the suitability of scaffolds produced using this technique.7,23 For example, studies have shown host tissue infiltration into TIPS PLGA foam cubes cut from a monolith to be dependent on the orientation of the pore structure.7 Further, studies have described tissue in-growth into the pores of TIPS scaffolds being limited by a significant foreign body giant cell response that blocked tissue infiltration into pores smaller than 300 mm.23 With TIPS microporous spheres, pores radiate from the center of the microporous sphere toward the surface; therefore, pore orientation does not need to be controlled for when implanting scaffolds composed of TIPS microporous spheres and macroporosity is maintained due to the predictable gaps between spheres. The semi-smooth porous surface of the TIPS microporous spheres combined with the large void opening onto the surface appears to provide an ideal topology and structure for rapid cell attachment and infiltration into the microporous spheres. Rapid tissue infiltration into the pores on the surface and the large void inside the microporous sphere (created by the entrapment of air as the droplet of polymer solution forms during the microporous sphere fabrication process) occurred within 1 week of implantation. This rapid tissue in-growth is likely to integrate the microporous spheres into the host tissue, preventing subsequent movement. The infiltration of tissue into the voids inside the microporous spheres is likely to occur mainly via movement of cells along a neck that extends from the exterior surface of the microporous sphere. In addition to this, cells may also enter via the pores that open out into the void. The cellularized voids are likely to provide delivery of oxygen, nutrients, and chemotactic signals to cells infiltrating the radial pores from the microporous sphere surface, thus helping to accelerate fibrovascular tissue infiltration and maintaining intraspherical tissue viability. Unlike previous studies investigating the in vivo response to TIPS scaffolds, a foreign body giant cell response was not observed with TIPS microporous spheres. During wound healing in healthy tissue, angiogenesis results in new capillaries sprouting from preexisting vessels and organizing into a microvascular network throughout the granulation tissue. VEGF is a specific and critical regulator of angiogenesis, controlling endothelial proliferation, permeability, and survival. It has been proposed that certain chronic wounds, where revascularization of damaged tissue is unregulated or insufficient, might benefit from molecular manipulation of growth factors, such as VEGF, to enhance microcirculation and promote tissue infiltration into the wound area.24 Recent studies have demonstrated the ability of BG to stimulate angiogenesis both in vitro and in vivo.5–7 Both the neat PLGA TIPS microporous spheres and those containing BG demonstrated their potential angiogenic properties by stimulating a significant increase in the secretion of VEGF from myofibroblasts in vitro. Quantitative assessment of angiogenesis in tissue surrounding microporous spheres that had been implanted subcutaneously was not possible due to the dispersal of microporous sphere cluster at the implant site follow implantation. The extent of intraspherical vascularization of the microporous sphere 1460 voids was not affected by this, but the ability to count vessels within the voids was dependent upon whether tissue sectioning bisected the microporous spheres in an appropriate plane. Therefore, it was only possible to quantitatively assess a limited number of microporous spheres in the current study. A sufficient number of voids for quantitative assessment were visible inside both types of microporous spheres at 2 weeks, but no significant difference in the number of blood vessels infiltrating the voids existed between microporous spheres containing 10% BG or control microporous spheres. The presence of well-vascularized voids inside the neat PLGA TIPS microporous spheres suggests either that the inclusion of an angiogenic stimulus is not necessary to promote neovascularization of the scaffold at the implant site, or that the normal wound healing response in current model, which used healthy animals, masked the angiogenic stimulus initiated by BG. The latter issue could be addressed in future studies by assessing the angiogenic response to TIPS microporous spheres, with or without BG, in wound models created with animals that have impaired angiogenesis, such as spontaneously hypertensive rats, a well-established experimental model of essential hypertension that has documented impaired angiogenesis compared with normotensive rats.25,26 TIPS microporous spheres composed of PLGA demonstrated good integration with host tissue. As the spheres degrade, the volume they occupy becomes reduced allowing space for further tissue infiltration into the interstices. Although the inclusion of BG as an angiogenic stimulus was successfully demonstrated in vitro, the inherent structure of the microporous spheres may also facilitate neovascularization of the microporous spheres. Rapid vascular integration of the TIPS microporous spheres with host tissue is likely to ensure improved viability of cells infiltrating the microporous spheres intraspherically and interspherically. The study demonstrates that PLGA TIPS microporous spheres integrate well with host tissues and degrade at predictable rates, suggesting that they could be readily used as a scaffold=filler material for wounds associated with tissue insufficiency. Acknowledgments The authors appreciate the technical support of Nicky Mordan (SEM). The work conducted in this study was supported by grants from the Sir Halley Stewart Trust and UK Medical Research Council. 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