hyaluronic acid/plga core/shell fiber matrices loaded...

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Hyaluronic Acid/PLGA Core/Shell Fiber Matrices Loaded with EGCG Beneficial to Diabetic Wound HealingYong Cheol Shin, Dong-Myeong Shin, Eun Ji Lee, Jong Ho Lee, Ji Eun Kim, Sung Hwa Song, Dae-Youn Hwang, Jun Jae Lee, Bongju Kim, Dohyung Lim, Suong-Hyu Hyon, Young-Jun Lim,* and Dong-Wook Han*

DOI: 10.1002/adhm.201600658

During the last few decades, considerable research on diabetic wound healing strategies has been performed, but complete diabetic wound healing remains an unsolved problem, which constitutes an enormous biomedical burden. Herein, hyaluronic acid (HA)/poly(lactic-co-glycolic acid, PLGA) core/shell fiber matrices loaded with epigallocatechin-3-O-gallate (EGCG) (HA/PLGA-E) are fabricated by coaxial electrospinning. HA/PLGA-E core/shell fiber matrices are composed of randomly-oriented sub-micrometer fibers and have a 3D porous network struc-ture. EGCG is uniformly dispersed in the shell and sustainedly released from the matrices in a stepwise manner by controlled diffusion and PLGA degradation over four weeks. EGCG does not adversely affect the thermomechanical proper-ties of HA/PLGA-E matrices. The number of human dermal fibroblasts attached on HA/PLGA-E matrices is appreciably higher than that on HA/PLGA counter-parts, while their proliferation is steadily retained on HA/PLGA-E matrices. The wound healing activity of HA/PLGA-E matrices is evaluated in streptozotocin-induced diabetic rats. After two weeks of surgical treatment, the wound areas are significantly reduced by the coverage with HA/PLGA-E matrices resulting from enhanced re-epithelialization/neovascularization and increased collagen deposition, compared with no treatment or HA/PLGA. In conclusion, the HA/PLGA-E matrices can be potentially exploited to craft strategies for the accel-eration of diabetic wound healing and skin regeneration.

1. Introduction

Skin is often damaged by scratching, wounding, stabbing, or physical trauma. A slight wound (i.e., partial-thickness wound) naturally heals through a wound healing process without any specific treat-ment.[1] On the other hand, more severe injures (i.e., full-thickness wounds) are quite difficult to naturally heal. A full-thickness wound is defined as extensive damage to the subcutaneous layer of the dermis or further tissues.[2] The healing of full-thickness wounds not only takes a long time, but also requires further treat-ment. In particular, the full-thickness wound healing is significantly impaired and delayed in diabetes owing to the hyperglycemia, which can lead to vascular dysfunction, abnormal collagen metabo-lism, oxidative stress, infection, glycolipid dysbolism, and neuropathy.[3] Therefore, considerable efforts have been made to effectively promote the diabetic wound healing and skin regeneration.[4] As part of

Y. C. Shin, E. J. Lee, J. H. Lee, Prof. D.-W. HanDepartment of Cogno-Mechatronics EngineeringCollege of Nanoscience & NanotechnologyPusan National UniversityBusan 46241, KoreaE-mail: [email protected]. D.-M. ShinResearch Center for Energy Convergence TechnologyPusan National UniversityBusan 46241, KoreaJ. E. Kim, S. H. Song, Prof. D.-Y. HwangDepartment of Biomaterials ScienceCollege of Natural Resources and Life ScienceLife and Industry Convergence Research InstitutePusan National UniversityMiryang 50463, KoreaDr. J. J. Lee, Prof. Y.-J. LimDepartment of ProsthodonticsDental Research InstituteSchool of DentistrySeoul National UniversitySeoul 03080, KoreaE-mail: [email protected]

Dr. B. KimDental Life Science Research InstituteSeoul National University Dental HospitalSeoul 03080, KoreaProf. D. LimDepartment of Mechanical EngineeringSejong UniversitySeoul 05006, KoreaProf. S.-H. HyonCenter for Fiber and Textile ScienceKyoto Institute of TechnologyKyoto 606-8585, Japan

Adv. Healthcare Mater. 2016, DOI: 10.1002/adhm.201600658

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the many efforts to accelerate the wound healing process, many studies have focused on the development of artificial scaffolds to enhance the cellular behavior through a tissue engineering approach.[5] In particular, polymer-based scaffolds incorporated with various biochemical factors have been recently reported one after another. For example, a previous study has shown that the astragalus polysaccharide-loaded poly(lactic-co-glycolic acid, PLGA) fibrous mats accelerate diabetic wound healing by enhancing the restoration of skin microcirculation.[5c] In addi-tion, specific proteins can be also incorporated in polymer scaf-folds to encourage wound healing. It has been documented that the collagen scaffold containing the gap junction connexin 43 (Cx43) antisense oligodeoxynucleotides (Cx43asODN) enhances wound healing process through a sustained release of Cx43asODN, which leads to a downregulation of Cx43 and reduced inflammation.[5e] However, the development of ideal scaffolds for full-thickness wound healing is still a challenge because the wound healing process involves a highly intricate series of cellular responses and interactions among the extra-cellular matrix (ECM), various types of cells, cytokines, and growth factors.[6] An ideal scaffold should not only have excel-lent biocompatibility and bioactivity, but should also promote the complicated repair process. In addition, they should be structurally and dimensionally similar to the natural ECM to support cell growth. To meet these demands, a variety of bioma-terials and techniques have been suggested and examined for the development of wound healing scaffolds. Although a range of techniques, such as emulsion templating, salt leaching, phase separation, and self-assembly, can be used to fabricate a wound healing scaffold, electrospinning is still one of the most effective and applicable techniques. This is because electrospin-ning can easily fabricate a nanoscale fibrous network structure similar to the natural ECM by simply supplying an electric field to polymer solutions.[5b,7] In particular, coaxial electrospinning is quite suitable for fabricating core/shell fibers using a dual or triple concentric nozzle.[7b,8] In addition, electrospun matrices have a large specific surface area-to-volume ratio, high porosity, and ease of control over the diameter, composition, and mor-phology of the constituent fibers.[9]

In the present study, we fabricated hyaluronic acid (HA)/PLGA core/shell fiber matrices loaded with epigallocate-chin-3-O-gallate (EGCG) (HA/PLGA-E), and explored their healing effects on full-thickness wounds in diabetic rats. HA, the main component of the ECM, has been extensively used in the field of biomedical engineering owing to its biological activities, such as wound healing effect and matrix organi-zation.[10] A previous study has revealed that a HA nanofiber wound dressing can promote wound healing by facilitating cell migration and proliferation.[10e] On the other hand, EGCG has attracted considerable attention as a promising biomaterial because of its diverse pharmacological properties. EGCG, the major polyphenolic compound found in green tea, has many pharmacological activities, including antioxidative, cancer pre-ventive, bactericidal, and anti-inflammatory effects.[11] It has been shown that EGCG-incorporated collagen sponge can accel-erate wound healing in diabetic mice due to its potent angio-genic activity.[11c] In addition, EGCG containing membranes can enhance the healing of full-thickness wounds by promoting the formation of new capillaries and the proliferation of epithelial

cells.[11g] Therefore, it was hypothesized that HA/PLGA-E core/shell fiber matrices would effectively promote the healing of the diabetic full-thickness wounds through the synergistic effects of HA and EGCG. The physicochemical and thermomechanical properties of HA/PLGA-E core/shell fiber matrices were deter-mined and the EGCG release behavior was examined. Further-more, the in vivo wound healing activity was investigated using a full-thickness wound model in normal and streptozotocin (STZ)-diabetic Sprague-Dawley (SD) rats to explore their poten-tial as novel scaffolds for tissue regeneration.

2. Results and Discussion

2.1. Physicochemical and Thermomechanical Characterizations of HA/PLGA-E Matrices

HA/PLGA-E core/shell fiber matrices were prepared by coaxial electrospinning of HA and PLGA with 2 and 4 wt% EGCG [HA/PLGA-E(2) and HA/PLGA-E(4) matrices, respectively]. The physicochemical and thermal properties of the HA/PLGA-E core/shell fiber matrices were examined by field emission scan-ning electron microscopy (FESEM), Fourier transform infrared (FTIR) spectroscopy, contact angle measurements, thermo-gravimetric analysis (TGA), and differential scanning calorim-etry (DSC) (Figure 1). The digital photographs showed that the HA/PLGA-E matrices were slightly yellowish red in color due to the EGCG loaded in the PLGA shell (Figure 1A). FESEM images revealed the average diameters of HA/PLGA, HA/PLGA-E(2), and HA/PLGA-E(4) fibers to be 519 ± 88, 365 ± 152, and 292 ± 147 nm, respectively. The average diameter of the fibers gradually decreased when EGCG was loaded in the shell. This decrease in diameter of the HA/PLGA-E core/shell fibers can increase the surface area-to-volume ratio, which leads to an effective interaction between the matrices and cells.[7c,9d,12] On the other hand, the core/shell structure of the HA/PLGA-E fibers was confirmed by FESEM and fluorescence micro scopy (Figure S1, Supporting Information). The cross-sectional FESEM image of the HA/PLGA-E fibers clearly showed their core/shell structure (Figure S1A, Supporting Information). To further prove the core/shell structure, we conducted fluo-rescence microscopy, where the fluorescein-4-isothiocyanate (FITC) was added into the HA core solution. As shown in Figure S1B in the Supporting Information, the green fluores-cence of the FITC, which exists in the core of fibers, was obvi-ously detected along the HA/PLGA-E fibers. Therefore, it is revealed that the core/shell structure of the HA/PLGA-E fibers was finely fabricated by coaxial electrospinning. Figure 1B shows the FTIR spectra of the HA/PLGA and HA/PLGA-E fiber matrices. In the spectrum of the HA/PLGA-E(4) matrix, a noticeable band observed near 1750 cm−1 was assigned to the CO stretching mode from the ester group of PLGA. The char-acteristic band for HA was also found near 1560 cm−1, which can be assigned to the amide II vibration originating from the NH bending coupled with CN stretching vibrations.[9c] On the other hand, the specific bands of EGCG were observed near 850, 1500, and 3600–3400 cm−1, which can be attributed to the CH alkene, CC alkene, and phenylOH stretching mode, respectively.[13] Therefore, the HA/PLGA-E(4) matrices



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were successfully prepared, and the HA and EGCG were uni-formly dispersed in the core/shell fibers. Previous studies have reported that surface hydrophilicity is one of the major factors that can affect the cellular behavior.[14] Therefore, the water con-tact angles and surface energies of the matrices were measured. Figure 1C shows the surface hydrophilicity of the HA/PLGA and HA/PLGA-E fiber matrices. The water contact angles of the HA/PLGA, HA/PLGA-E(2), and HA/PLGA-E(4) matrices were 124.7 ± 2.5, 120.2 ± 1.1, and 119.3° ± 0.7°, respectively. In addition, the surface energy of the HA/PLGA-E(4) matrices was 12.11 mN m−1, which is higher than that of the HA/PLGA matrices (9.58 mN m−1). These results indicated that the surface hydrophilicity of the matrices was significantly increased by the EGCG loading in the matrices due to the hydroxyl groups in EGCG. A series of studies have found that the improvement in the surface hydrophilicity of matrices is beneficial for pro-moting the interactions between the cells and matrices as well as the cellular behaviors including initial attachment and proliferation.[7c,9c,15] Hence, the HA/PLGA-E(4) matrices can provide a favorable microenvironment for cell growth because they have a more hydrophilic surface than other matrices [HA/PLGA and HA/PLGA-E(2) matrices]. The thermal proper-ties of the HA/PLGA-E(4) matrices were also examined by TGA and DSC to investigate the thermal stability of the matrices (Figure 1D). The TGA plots showed that minor weight losses occurred in all matrices over the temperature range, 50–80 °C,

which was followed by major weight losses at temperatures higher than 250 °C. These weight losses were attributed to the glass transition temperature of PLGA (45–55 °C) and thermal decomposition of the matrices.[16] On the other hand, the DSC thermogram revealed two broad endotherm peaks for each matrix near 50 and 350 °C, which were attributed to the glass transition temperature and endothermic melting peak of PLGA, respectively. The endothermic melting peak of the HA/PLGA-E(4) matrices was slightly shifted to a lower tempera-ture region compared to the HA/PLGA matrices, which could be associated with a melting point of EGCG (≈220 °C).[17] These results suggest that the HA/PLGA-E core/shell fiber matrices were sufficiently thermally stable to support cell growth under normal physiological conditions, and the loaded EGCG did not adversely affect the thermal behavior of the matrices.

In vitro EGCG release studies were performed to determine if the required amount of EGCG can be released from the core/shell fibers to have a pharmaceutical effect (Figure 2). EGCG was shown to be sustainedly released from the matrices in a stepwise manner by controlled diffusion and PLGA degrada-tion for 28 d. During the first day, an initial burst release was observed, which was a rapid release of EGCG with ≈10% and 20% of the total EGCG released from the HA/PLGA-E(2) and HA/PLGA-E(4) matrices, respectively. In the following 7 d, the rate of EGCG release increased logarithmically, and approached a plateau. After 28 d, the accumulated release reached about



Figure 1. Physicochemical and thermal characteristics of the HA/PLGA-E core/shell fiber matrices. A) Digital photographs and FESEM images of HA/PLGA, HA/PLGA-E(2), and HA/PLGA-E(4) fiber matrices (magnification: x1000 and x7000 for insertion). All images shown in this figure are repre-sentative of six independent experiments with similar results. B) FTIR spectra of HA/PLGA and HA/PLGA-E fiber matrices. All spectra were recorded in absorption mode over the wavelength range of 500–4000 cm−1 with a resolution of 4.0 cm−1 and 16-times scanning. C) Water contact angles and surface energies of HA/PLGA, HA/PLGA-E(2), and HA/PLGA-E(4) fiber matrices. The different letters denote the significant differences between the control and experimental groups (p < 0.05). If a group is marked with a dual letter (e.g., bc), it has significant difference from the control and another group marked with “a”, but does not have significant difference from the other groups marked with “b” or “c”. D) TGA curves and DSC thermograms of HA/PLGA, HA/PLGA-E(2), and HA/PLGA-E(4) fiber matrices. For TGA and DSC, the samples were heated from 25 to 500 °C at a heating rate of 10 °C min−1.

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65% and 70% of its loaded amounts for the HA/PLGA-E(2) and HA/PLGA-E(4) matrices, respectively. The total amounts of EGCG released during 28 d were 122 × 10–6 m for the HA/PLGA-E(2) matrices and 227 × 10–6 m for HA/PLGA-E(4) matrices. It is noted that considering the period between 1 and 7 d, the daily release of EGCG was around 11 and 19 × 10–6 m d−1 for the HA/PLGA-E(2) and HA/PLGA-E(4) matrices, respectively. Previous studies have reported that the effective concentra-tion of EGCG for pharmacological activities is in the range of 20–400 × 10–6 m.[18] In addition, the reactive oxygen species (ROS) scavenging rate is increased from 52% to 72% with increasing EGCG concentration from 10 to 20 × 10–6 m.[19] This indicates that the HA/PLGA-E(4) matrices are more suitable for diabetic wound healing than HA/PLGA-E(2) matrices. Thus the HA/PLGA-E(2) matrices were excluded from the in vivo studies. Meanwhile, HA release profile was also investigated to deter-mine if the core material (HA) is released from the core/shell fibers (Figure S2, Supporting Information). The HA release pat-tern was similar to that of EGCG. The initial burst release with ≈4% was observed during the first day, followed by a gradual decrease in the release rate of HA. As expected, the released HA was smaller than EGCG because the HA was placed in the

core region of the HA/PLGA-E(4) fiber. The accumulated HA released during 28 d was 18% of its initial loaded amounts. The in vitro degradation behavior of the matrices was exam-ined by measuring the cumulative weight loss of each matrix (Figure S3, Supporting Information). All matrices began to mainly degrade during the first week, and the weight loss rate gradually decreased with time for up to four weeks, as shown in Figure S3A in the Supporting Information. During the testing period, the weight loss of the HA/PLGA-E(4) matrices was higher than that of the HA/PLGA and HA/PLGA-E(2) matrices. After four weeks, the cumulative weight losses for the HA/PLGA, HA/PLGA-E(2), and HA/PLGA-E(4) matrices were 17.81%, 20.69%, and 25.00%, respectively. This greater degrada-tion of the HA/PLGA-E(4) matrices could be due to the release of EGCG. In addition, the improved surface hydrophilicity of the matrices can facilitate water diffusion, and allow the accel-erated hydrolysis of PLGA. Taking the release and degradation profiles into consideration, the main release mechanism may be the controlled diffusion of HA and EGCG with PLGA deg-radation.[20] The morphological changes of the matrices were shown in Figure S3B–D in the Supporting Information. With time, the constituent fibers gradually degraded, and the fibrous structure of the matrices was no longer visible after four weeks of incubation. These morphological changes were in concord-ance with the degradation profiles of the matrices.

Figure 3 presents the mechanical properties of the matrices. The ultimate tensile strength, elastic modulus, and elongation at break were determined from the stress–strain curves. Although the ultimate tensile strength, elastic modulus, and elongation at break were slightly decreased by the EGCG loading, they are not critically dependent on the EGCG loading considering the



Figure 2. In vitro EGCG release profiles from the HA/PLGA-E(2) and HA/PLGA-E(4) core/shell fiber matrices. A) The released EGCG is plotted in units of the percentage of its loaded amounts. EGCG was sustainedly released from the HA/PLGA-E core/shell fibers by controlled diffusion and PLGA degradation. B) Cumulatively released EGCG concentration is plotted in units of μm. After 28 d, the total amounts of released EGCG were about 65% (122 × 10–6 m) and 70% (227 × 10–6 m) of its loaded amounts for the HA/PLGA-E(2) and HA/PLGA-E(4) matrices, respectively.

Figure 3. Mechanical characteristics of the HA/PLGA-E core/shell fiber matrices. Stress–strain curves of the HA/PLGA, HA/PLGA-E(2), and HA/PLGA-E(4) fiber matrices were obtained under a cross-head speed of 10 mm min−1. The ultimate tensile strength, elastic modulus, and elon-gation were determined from the stress–strain curves. Prior to testing, the three types of matrices were cut into a rectangular shape, 40 mm in length and 10 mm in width. The ultimate tensile stress, elastic modulus, and elongation of HA/PLGA-E matrices are not severely dependent on the EGCG loading, suggesting that the EGCG loading does not adversely affect the mechanical properties of the matrices.

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difference between the HA/PLGA and HA/PLGA-E(4) matrices. Therefore, it is suggested that the EGCG loading does not adversely affect the mechanical properties of the matrices.

2.2. Cellular Behaviors of Normal Human Dermal Fibroblasts (nHDFs) on HA/PLGA-E Matrices

Prior to examining the in vivo wound healing activity of the HA/PLGA-E matrices, the in vitro cellular behaviors on the matrices were assessed by investigating the initial attachment and proliferation of nHDFs on the matrices (Figure S4, Sup-porting Information). As shown in Figure S4A in the Supporting Information, the initial attachment of nHDFs was significantly (p < 0.05) enhanced on the HA/PLGA-E(4) matrices compared to that on the tissue culture plastics (TCPs), HA/PLGA, and HA/PLGA-E(2) matrices. This can be explained by the specific binding property of EGCG with various biomolecules facili-tating the interaction with cells. It has been documented that the EGCG can specifically bind with biomolecules, such as amino acid and protein, through the hydrophobic interaction between the functional moieties of EGCG and the hydrophobic residues of biomolecules.[21] Therefore, the HA/PLGA-E(4) matrices can facilitate the interaction between matrices and cells via the binding between EGCG and cell adhesion mol-ecules, which allows an increase in the initial attachment of nHDFs. In addition, the improved surface hydrophilicity of HA/PLGA-E(4) matrices can also participate in the increased initial cell adhesion.[15] Therefore, the specific binding property of EGCG, together with the improved surface hydrophilicity, makes HA/PLGA-E(4) matrices preferable for cell attachment.

On the other hand, the proliferation of nHDFs on the HA/PLGA-E(4) matrices was similar to that on the other matrices, regardless of EGCG loading (Figure S4B, Supporting Informa-tion). It has been described that although the effects of EGCG on the cell growth are found to be dose-dependent, the prolif-eration of nHDFs is not significantly affected at concentrations lower than 200 × 10–6 m.[22] Considering that the total amount of released EGCG from HA/PLGA-E(4) matrices for 28 d was 227 × 10–6 m, the HA/PLGA-E(4) matrices can effectively sup-port the growth of nHDFs without hindering their proliferation. Moreover, EGCG has been shown to exhibit antioxidative and anti-inflammatory effects, which can stem from its molecular structure with parahydroxyl and galloyl groups.[23] Therefore, these antioxidative and anti-inflammatory effects can be of great help for diabetic wound healing, even if the proliferation was somewhat decreased only when compared with the TCP control group. Hence, HA/PLGA-E(4) matrices are expected to be beneficial to diabetic wound healing by exerting both antioxi-dative and anti-inflammatory effects.

2.3. Effects of HA/PLGA-E Matrices on Tissue Regeneration of Wound Skin

To evaluate the diabetic wound healing effects of the HA/PLGA-E core/shell fiber matrices, full thickness incisional models in normal and STZ-diabetic SD rats were designed. The body weight and blood glucose concentration were measured,

as shown in Figure 4. As previously reported, the body weights of the STZ-diabetic SD rats decreased over time (Figure 4A) and the blood glucose concentration significantly increased in all groups (Figure 4B).[24] Therefore, the diabetic conditions were successfully induced in the SD rats. On the other hand, Figure S5 in the Supporting Information shows the body weight and blood glucose concentration of the normal rats during the wound healing process. The body weights were somewhat decreased in normal rats at 14 d. The reduction in body weight, however, could be attributed to the full-thickness wounding and the body dressing. It has been documented that wounding and body dressing can cause occasional loss of body weight in rats.[25] Accordingly, it is considered that the full-thickness wounds and body dressing caused a little discomfort to the rats, which results in a decrease in food consumption and finally in body weight. Nevertheless, the blood glucose concentration was maintained at less than 200 μg mL−1, although the body weight decreased at 14 d. Hence, it is indicated that the normal rats maintained normal health.

The wound healing effects of the HA/PLGA-E core/shell fiber matrices were first investigated by a macroscopic evalu-ation (Figure 5). The healing process of an incisional wound and the remaining wound area of each group were tracked over a 14-d period. As shown in Figure 5A, the HA/PLGA-E(4) group showed a significantly (p < 0.05) higher wound healing rate than the control, PLGA, and HA/PLGA groups, which was even superior to the Rapiderm group, as the positive control. After 7 d, the wound of the HA/PLGA-E(4) group was mark-edly reduced, and the wound was almost completely healed after 14 d. In addition, there was no evidence of inflammation or infection, and the wound was covered with new epidermis and sparse hair. The remaining wound area was quantified as



Figure 4. Changes in A) body weight and B) concentration of blood glucose of STZ-diabetic rats during wound healing processes (14 d). With time, the body weights gradually decreased, and the blood glucose concentrations reached about 600 μg mL−1 in all groups.

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a percentage of the initial wound area (Figure 5B). The wound area in the HA/PLGA-E(4) group was significantly (p < 0.05) decreased after 14 d compared to the other groups. After 14 d, the remaining wound area for the control, PLGA, HA/PLGA, HA/PLGA-E(4), and Rapiderm groups was 49.96%, 48.43%, 40.18%, 10.84%, and 24.87%, respectively. This result is well consistent with the macroscopic evaluation results, as presented in Figure 5A. Interestingly, the wound healing rate was significantly greater in the HA/PLGA-E(4) group than in the other groups. This enhanced wound healing rate could be due to the synergistic effects of HA and EGCG. The HA and EGCG released from the HA/PLGA-E matrices can accelerate the wound healing process by scavenging ROS, mitigating inflammation, stimulating re-epithelialization, and promoting angiogenesis and ECM re-organization.[10,11] Wound healing in diabetes is severely impaired because of several complica-tions, such as vascular dysfunction, glycolipid dysbolism, and neuropathy, caused by hyperglycemia.[5c,6c,26] In addition, the ROS generated by oxidative stress, inflammation, and infec-tion is a major obstacle to wound healing.[27] Therefore, the

synergistic effects of HA and EGCG can significantly enhance full-thickness wound repair by reducing those undesired effects as well as accelerating the wound healing process. Our results were substantially in accordance with previous studies, which demonstrated that ECGC- or HA-incorporated scaffolds can promote wound healing by accelerating re-epithelialization, wound contraction, and angiogenesis.[11c,g] On the other hand, these accelerated wound healing effects were also demon-strated in normal rats (Figure S6, Supporting Information). The remaining wound area of the HA/PLGA-E(4) group was signif-icantly decreased at 7 d compared to the control group, even though a significant difference in the macroscopic changes between control and HA/PLGA-E(4) groups was not apparent. Considering the results of the normal and STZ-diabetic rats, the wound healing effect of the HA/PLGA-E(4) matrices in the STZ-diabetic rats was comparable to that in the normal rats. Therefore, our findings indicated that the HA/PLGA-E(4) matrices can effectively promote diabetic wound healing.

2.4. Effects of HA/PLGA-E Matrices on Formation of Connective Tissue in Wound Skin

Immunohistochemical (IHC) and molecular analysis were conducted to investigate the diabetic wound healing activity of HA/PLGA-E matrices more specifically. Figure 6 shows a his-topathological image, re-epithelialization and CD31 expression in the wounds of each group at 14 d. The histopathological images revealed a continuous and thick epithelial layer in the HA/PLGA-E(4) matrices, while the control and PLGA groups showed an incomplete and thin epithelial layer (Figure 6A). Re-epithelialization, which reflects successful wound healing, was significantly (p < 0.05) increased in the HA/PLGA-E(4) group (53.88 μm) when compared to that of the other groups (40.07, 39.82, 8.21, and 5.85 μm for Rapiderm, HA/PLGA, PLGA, and control group, respectively) (Figure 6B). Interest-ingly, the EGCG showed differential responses depending on the type of cell.[22] In particular, it has been demonstrated that EGCG can stimulate the proliferation and differentiation of keratinocytes, a major population of the epidermis, which leads to an increase in the epidermal thickness.[28] Therefore, the HA/PLGA-E(4) matrices could enhance the proliferation of keratinocytes, resulting in effective re-epithelialization. On the other hand, revascularization of the wound area was estimated by IHC analysis of CD31 expression in wounds (Figure 6C,D). CD31, platelet-endothelial cell adhesion molecule-1, is con-sidered a marker for endothelial cells, reflecting new capillary formation.[29] The expression of CD31 was detected by IHC staining (brown). As shown in Figure 6C, the CD31-positive cells were largely observed in the HA/PLGA-E(4) and Rapiderm groups. In the control, PLGA and HA/PLGA groups, however, CD31-positive cells were rarely observed in the wound area. The number of CD31-positive cells of the HA/PLGA-E(4) group was comparable to that of Rapiderm group, and significantly (p < 0.05) higher than that of the control and HA/PLGA groups (Figure 6D). These results indicated that new capillary forma-tion was greatly improved in the wounds by the HA/PLGA-E(4) matrices, which could be also due to the synergistic effects of HA and EGCG. HA can promote not only revascularization, but



Figure 5. Macroscopic evaluation of the wound healing effects of the HA/PLGA-E core/shell fiber matrices in STZ-diabetic rats. A) Representa-tive macroscopic appearances of the wound area in the control, PLGA, HA/PLGA, HA/PLGA-E(4), and Rapiderm groups on 0, 7, and 14 d (n = 6). The wound healing rate of HA/PLGA-E(4) group was significantly higher than other groups without macroscopic evidence for inflammation or infection. B) Remaining wound area of the control, PLGA, HA/PLGA, HA/PLGA-E(4), and Rapiderm groups. The remaining wound area was quantified as a percentage of the initial wound area at 0 d. After 14 d, the wound area in the HA/PLGA-E(4) group (10.84%) was significantly reduced when compared to that in the other groups (49.96%, 48.43%, 40.18%, and 24.87% for control, PLGA, HA/PLGA, and Rapiderm group, respectively). An asterisk (*) denotes a significant difference compared to the control and PLGA groups (p < 0.05) and a number sign (#) denotes a significant difference between the HA/PLGA-E(4) group and other groups (p < 0.05).

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also ECM re-organization in an animal model, and EGCG can also have potent in vivo angiogenetic effects.[11c,30] These angio-genetic effects of HA and EGCG can allow a seamless provi-sion oxygen and nutrient supplements to allow the wounds to heal, and can effectively accelerate, especially, diabetic wound healing. In addition, the enhanced wound healing effects of the HA/PLGA-E(4) matrices were confirmed in the normal rats (Figure S7, Supporting Information). These results were different from those of STZ-diabetic rats, but the re-epithe-lialization and revascularization obviously increased in the HA/PLGA-E(4) group. These findings provide a fundamental understanding of the mechanism for how the HA/PLGA-E(4) matrices promote diabetic wound healing by showing that the HA/PLGA-E(4) matrices are able to encourage the re-epitheliali-zation and revascularization of the wound area.

Molecular analysis was performed by quantitatively evalu-ating the expression of collagen, matrix metalloproteinases (MMP-1) and CD31. We measured collagen and MMP-1 expres-sion to examine re-epithelialization, and CD31 expression to assess revascularization, respectively (Figure 7). Collagen, a major component of the ECM, plays a significant role in ECM re-organization and tissue re-modeling in wounds. MMP-1 is a calcium-dependent zinc-containing enzyme, and has many

specific functions in the wound healing process, such as regu-lating cell migration, interactions between the ECM and cells, signaling pathway, and tissue re-modeling.[31] The levels of collagen, MMP-1, and CD31 expression were substantially up-regulated in the HA/PLGA-E(4) group: 2.06-fold, 2.35-fold, and 1.95-fold for collagen, MMP-1, and CD31 compared to those in the control group, respectively. These findings are in accordance with the IHC analysis results, and the up-regulated expression of collagen, MMP-1, and CD31 resulted in expedited re-epithelialization and revascularization. It has been shown that MMP-1 plays a crucial role in matrix assembly and the migra-tion of keratinocytes.[31b,c] Thus, the increased collagen and MMP-1 expressions effectively facilitate ECM re-organization, which in turn leads to improved re-epithelialization. On the other hand, as noted in IHC analysis, the significantly elevated CD31 expression reflects the facilitated revascularization by the HA/PLGA-E(4) matrices. These increased expression of col-lagen, MMP-1, and CD31 could be also found in normal rats (Figure S8, Supporting Information). Therefore, considering the IHC and molecular analysis results, the HA/PLGA-E(4) matrices are feasible to remarkably facilitate the diabetic wound healing by promoting both re-epithelialization and revasculari-zation in full-thickness wounds. In summary, it is suggested



Figure 6. IHC analysis of the effect of the HA/PLGA-E core/shell fiber matrices on diabetic wound healing. A) Histopathological observations (H&E staining) of the wounds in the control, PLGA, HA/PLGA, HA/PLGA-E(4), and Rapiderm groups on 14 d (n = 6). A continuous and thick epithelial layer were observed in HA/PLGA-E(4) group. B) Quantitative analysis of re-epithelialization of the wounds in each group at 14 d. The thickness of epidermis for the control, PLGA, HA/PLGA, HA/PLGA-E(4), and Rapiderm groups was 5.85, 8.21, 39.82, 53.88, and 40.07 μm, respectively. C) IHC staining of CD31-positive cells in the wounds on 14 d (n = 6). The CD31-positive cells were IHC stained with the Polink-2 plus polymer HRP detection system and a diaminobenzidine (DAB) reagent kit for anti-CD31 antibody (brown). The arrows indicate the CD31-positive cells. D) Quantification of CD31-positive cells in the wounds. The number of CD31-positive cells for the control, PLGA, HA/PLGA, HA/PLGA-E(4), and Rapiderm groups were ≈68, 74, 124, 284, and 254 cells per mm2, respectively. An asterisk (*) denotes a significant difference compared to the control and PLGA groups (p < 0.05) and a number sign (#) denotes a significant difference between the HA/PLGA-E(4) group and other groups (p < 0.05).

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that the HA/PLGA-E(4) core/shell fiber matrices are clearly beneficial for diabetic wound healing.

3. Conclusion

The aim of the present study was to develop a novel scaffold for full-thickness wound healing, and to explore their in vivo wound healing activity in STZ-diabetic rats. We demonstrated that the HA/PLGA-E core/shell fiber matrices were successfully fabricated by coaxial electrospinning, and the HA and EGCG were released from the matrices in a sustainable manner by controlled diffusion with PLGA degradation. In addition, the in vivo wound healing effects of the HA/PLGA-E matrices were corroborated by an animal study showing that the in vivo full-thickness wound healing rate was significantly accelerated in both normal and STZ-diabetic rats. The possible underlying mechanism can be obtained from IHC and molecular anal-ysis. The results showed that the wound healing activity of the HA/PLGA-E matrices can be attributed to the synergistic effects of HA and EGCG, which prominently promote the re-epithelialization, ECM re-organization, and revascularization in wounds. Therefore, in conclusion, our findings suggest that the HA/PLGA-E core/shell fiber matrices are quite beneficial to diabetic full-thickness wound repair, and can be quite prom-ising candidates for novel scaffolds, particularly for diabetic wound healing.

4. Experimental SectionPreparation of HA/PLGA-E(4) Matrices: The PLGA [75:25

(mol mol−1), molecular weight (MW) = 70–110 kDa] resin used in this study was kindly provided by BMG Inc. (Kyoto, Japan). The EGCG

powder (MW = 458.4) was obtained from DSM Nutritional Products Ltd. (Basel, Switzerland). The solvent used for the shell solution was 1, 1, 1, 3, 3, 3-hexafluoroisopropanol (HFIP) form Sigma-Aldrich Co (St Louis, MO). HA (MW = 0.8–1.8 × 106 Da) was purchased from J&D chem (Cheongju-si, Korea). PLGA 20% (w v−1) and EGCG 2 or 4 wt% of the total PLGA weight were dissolved in HFIP to prepare the shell solution [PLGA-E(2) and PLGA-E(4), respectively]. HA 0.6% (w v−1) was dissolved in distilled water to prepare the core solution. The coaxial spinneret was composed of two-fluid concentric nozzles. The inner needle had a diameter of 0.25 mm, whereas the outer needle had a diameter of 1.07 mm. The HA solution was fed to the inner needle at a flow rate of 0.1 mL h−1 through a syringe pump, whereas the PLGA, PLGA-E(2) or PLGA-E(4) solutions were fed to the outer needle at a flow rate of 0.3 mL h−1 with an another syringe pump. A positive voltage of 15 kV was applied and the working distance between the needle tip and the collector was 17 cm. The HA/PLGA, HA/PLGA-E(2) or HA/PLGA-E(4) core/shell fibers were collected on a grounded steel rotating wheel wrapped with aluminum foil. The rotational speed of the wheel was 20 rpm. The collected core/shell fiber matrices were then dried overnight in a vacuum to remove any residual solvent. For the in vitro cell assays, the as-prepared matrices were cut into a disk shape, 12 mm diameter and 195 ± 21 μm in thickness. For the in vivo animal study, the matrices were cut into a square shape with dimensions of 15 × 15 mm2. All the disks and squares made from the HA/PLGA-E(4) matrices were sterilized by γ-irradiation.

Physicochemical and Thermomechanical Characterizations of HA/PLGA-E Matrices: The surface morphology of the HA/PLGA, HA/PLGA-E(2), and HA/PLGA-E(4) core/shell fiber matrices were observed by FESEM (Hitachi S-4700, Tokyo, Japan) at an accelerating voltage of 5 kV. Prior to analysis, the matrices were coated with an ultrathin layer of gold using an ion coater (E1010, Hitachi). Compositional analysis of the HA/PLGA-E matrices was performed by FTIR spectroscopy. The FTIR spectra were collected by an attenuated total reflectance FTIR spectrophotometer (Spectrum One FTIR Spectrometer, Perkin-Elmer, Boston, MA). All spectra were recorded in absorption mode over the wavelength range of 500–4000 cm−1 with a resolution of 4.0 cm−1 and 16-times scanning. The water contact angles of the matrices were examined at room temperature (RT) by the sessile drop method using a contact angle goniometer (Model CA-X,



Figure 7. Molecular analysis of the effect of the HA/PLGA-E core/shell fiber matrices on diabetic wound healing. A) Immunoblotting for collagen, MMP-1, and CD31 expression in control, PLGA, HA/PLGA, HA/PLGA-E(4), and Rapiderm groups on 14 d. The expression of β-actin was regarded as an internal control. B) Quantification of collagen, MMP-1, and CD31 expression in each group on 14 d. The expression levels were quantified by den-sitometry using an Image Analyzer System, and expressed as the fold-increase over the values of the control group. The levels of collagen (2.06-fold), MMP-1 (2.35-fold) and CD31 expression (1.95-fold) in the HA/PLGA-E(4) group were significantly upregulated compared to those in the control group. An asterisk (*) denotes a significant difference compared to the control and PLGA groups (p < 0.05).

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Kyowa Interface Science Co., Saitama, Japan). The thermal stability of the matrices was investigated by TGA (TGA n-1000, Scinco Co., Seoul, Korea) and DSC (MAC science, Tokyo, Japan). For TGA, the samples were weighed (≈5.8 mg) in open aluminum pans and heated from 25 to 500 °C at a heating rate of 10 °C min−1. For DSC, the matrices were heated from 25 to 500 °C at a rate of 10 °C min−1. The weights of the matrices were 5.78, 5.82, and 5.86 mg for the HA/PLGA, HA/PLGAE-(2), and HA/PLGA-E(4) matrices, respectively. Alpha-Al2O3 was used as a reference. In vitro EGCG release analysis was performed by immersing the matrices in Dulbecco’s phosphate-buffered saline (DPBS, pH 7.4, Gibco BRL, Rockville, MD) at 37 °C for 28 d. The absorbance was measured at 275 nm using a UV spectrophotometer (U-2800A, Hitachi) at the end of each incubation period. The concentration (μm) of the released EGCG was obtained from a standard calibration curve of an EGCG solution. The stress–strain curves of the matrices were measured using a tabletop tensile tester (LRX Plus Series, Ametek Lloyd Instruments Ltd., Fareham, UK) equipped with a 5 kN load cell under a cross-head speed of 10 mm min−1. Prior to measuring, the matrices were cut into a rectangular shape with dimensions of 40 × 10 mm2.

Design of Animal Experiment: The animal protocol used in this study was reviewed and approved based on ethical procedures and scientific care by the Pusan National University-Institutional Animal Care and Use Committee (PNU-IACUC; Approval Number PNU-2013-0475). Adult SD rats were purchased from SamTako Inc. (Osan, Korea) and given a standard irradiated chow diet (Purina Mills, Seongnam, Korea) ad libitum. All animals were handled at the Pusan National University Laboratory Animal Resources Center accredited by the Korea FAD (Accredited Unit Number; 00231) according to Laboratory Animals Act and AAALAC International (Accredited Unit Number; 001525) according to the National Institutes of Health guidelines. All rats were maintained in a specific pathogen-free state under a strict light cycle (light on at 06:00 h and off at 18:00 h) at a temperature of 22 ± 2 °C and at 50 ± 10% relative humidity. For the experiment, six-week-old SD rats (n = 42) were assigned to either a nondiabetes group (n = 12) or a diabetes group (n = 30). Diabetes was induced in the SD rats by an intraperitoneal injection of STZ (70 mg kg−1 weight) in 0.1 m citrate buffer, as described elsewhere.[32] The nondiabetes group was subdivided into a control group (n = 6) and HA/PLGA-E(4) group (n = 6). The diabetes group was subdivided into a control group (n = 6), PLGA group (n = 6), HA/PLGA group (n = 6), HA/PLGA-E(4) group (n = 6), and Rapiderm group (n = 6). The Rapiderm (Dalim Tissen Medical Co., Seoul, Korea) is a commercial wound dressing composed of a bilayered porcine collagen membrane. First, the animals were anesthetized by an intraperitoneal injection with Zoletile (50 mg kg−1 body weight) and Rompun (5 mg kg−1 body weight). The rats were shaved with electrical clippers before the application of 70% ethanol. A round wound, 8 mm in diameter and 2–4 mm in depth, was formed by removing the skin from the shoulder region of the back skin using a Biopsy Punch (Kasco com, Sialkot, Pakistan). Two full-thickness wounds were made on the back skin of each rat (n = 6 rats per group, 12 wounds per group). After making an incision, the wound skins of the rats in the first and second group were covered with the PLGA and HA/PLGA matrices, while those in the third and fourth group were covered with the HA/PLGA-E(4) matrices and Rapiderm, respectively. The fifth group was left untreated as a control. Each incision wound in the back skin of the SD rats was covered with PLGA matrices, HA/PLGA matrices, HA/PLGA-E(4) matrices, and Rapiderm with dimensions 5 × 5 × 0.3 mm after sterilization with 70% ethanol, and the boundary between the matrices and the skin around the wound was sutured. Subsequently, all wounds were covered with sterilized gauze to prevent the wound from contamination and possible infection. During the replacement process, the repair state of the wound skin was observed and an image was taken. After two weeks, all the rats were euthanized using carbon dioxide, and samples of damaged skin were collected from the rats for histological analysis by western blot analysis.

Histological Analysis: The wound tissue was removed from the SD rats of each group, fixed with 10% formalin, embedded in paraffin wax, routinely processed, and then sectioned into 4 μm thick slices. The skin sections were then stained with hematoxylin and eosin

(H&E, Sigma-Aldrich Co.) and examined by optical microscopy (Leica Microsystems, Switzerland) for the change in skin structure. In addition, the diameter of the wound and thickness of the epidermis were measured using a Leica Application Suite (Leica Microsystems, Wetzlar, Germany). IHC analysis was also performed, as previously described.[33] Briefly, the distribution of CD31 protein was observed by optical microscopy after fixing the tissue samples in 10% formalin for 48 h, the tissues were embedded in paraffin, and sections, 4 μm in thickness, were acquired. Each section was de-paraffinized with xylene, rehydrated, and pretreated for 30 min at RT with a DPBS-based blocking buffer containing 10% bovine serum albumin (BSA, GenDEPOT, Barker, TX). The samples were then incubated with the anti-CD31 antibody (Abcam Inc., Cambridge, MA) diluted 1:1000 in BSA-blocking buffer. The Polink-2 plus polymer horseradish peroxidase (HRP) detection system (GBI Laboratories, Mukilteo, WA) and a DAB reagent kit (GBI Laboratories) were used for IHC staining.

Western Blot: Samples of wound skin from a subset of the groups (n = 6 per group) were homogenized using a PRO-PREP Solution Kit (iNtRON Biotechnology, Sungnam, Korea) supplemented with 1/2 of a protein inhibitor cocktail tablet (Roche, Penzberg, Germany), followed by centrifugation at 13 000 rpm for 5 min. The prepared proteins were then electrophoresed through a 10% SDS-PAGE gel. The proteins were then transferred to a nitrocellulose membrane (Amersham Biosciences, Corston, UK) for 2 h at 40 V in the transfer buffer (25 × 10–3 m Trizma-base, 192 × 10–3 m glycine, and 20% methanol). The efficiency of the transfer and equal protein loading were determined by staining the gel with Coomassie Blue (Sigma-Aldrich Co.). The appropriate dilutions of the primary antibodies, anticollagen antibody (Abcam Inc.), anti-MMP-1 antibody (Santacruz Biotechnology, Santa Cruz, CA), anti-CD31 antibody (Abcam Inc.), and anti-β-actin (Sigma-Aldrich Co.) were added to the membranes and allowed to hybridize overnight at 4 °C. After the antibodies were removed, the membranes were washed three times with a solution composed of 10 × 10–3 m Trizma-base (pH 7.6), 150 × 10–3 m NaCl, and 0.05% Tween-20 for 10 min. This was followed by incubation with the HRP-conjugated antisecondary antibody for 1 h at RT. The membrane was washed again, as described above, and developed using an enhanced chemiluminescence detection system (Amersham Bioscience). Finally, the results were quantified using an Image Analyzer System (Estman Kodak 2000MM, Rochester, NY) and expressed as the fold-increase over the values of the control group. β-actin expression was regarded as an internal control. All the results were confirmed by two independent researchers who performed the experiments at least twice.

Statistical Analysis: All variables were tested in three independent cultures for each experiment in vitro, which was repeated twice (n = 6). The quantitative data are expressed as the mean ± standard deviation. The data were tested for the homogeneity of the variances using the test of Levene prior to statistical analysis. Multiple comparisons were performed to detect the cellular behaviors of nHDFs on the HA/PLGA-E matrices using one-way analysis of the variance (ANOVA, SAS Institute, Cary, NC), which was followed by a Bonferroni test when variances were homogeneous and a Tamhane test when the variances were not. Statistical analysis for the animal study was performed using a Kruskal–Wallis one-way ANOVA and Mann–Whitney U-test. A p value < 0.05 was considered significant. One-way ANOVA test (SPSS for Windows, Release 10.10, Standard Version, Chicago, IL) were performed to determine the variance and significance between the nondiabetes and diabetes groups. In addition, the tests for significance between the control- and other groups on the wound skin were performed using a Post-Hoc test (SPSS for Windows, Release 10.10, Standard Version) of the variance and the significance levels are given in the text. All values are reported as the mean ± standard deviation. A p value < 0.05 was considered significant.

Supporting InformationSupporting Information is available from the Wiley Online Library or from the author.



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10 wileyonlinelibrary.com © 2016 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim Adv. Healthcare Mater. 2016, DOI: 10.1002/adhm.201600658


AcknowledgementsY.C.S. and D.-M.S. contributed equally to this work. This study was supported by the WC 300 R&D Program funded by the Small and Medium Business Administration (SMBA) in Korea (No. S2318267) and the Bio & Medical Technology Development Program of the National Research Foundation (NRF) funded by the Korean government (MEST) (No. 2015M3A9E2028643).

Received: June 21, 2016Revised: September 2, 2016

Published online:

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