Materials Science and Engineering C 33 (2013) 648–655

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Biological evaluation of human hair keratin scaffolds for skin wound repair and regeneration Songmei Xu a, Lin Sang a, Yaping Zhang b, Xiaoliang Wang a, Xudong Li a,⁎ a b

National Engineering Research Center for Biomaterials, Sichuan University, Chengdu 610064, PR China Engineering Research Center of Biomass Materials, School of Materials Science and Engineering, Southwest University of Science and Technology, Mianyang 621010, PR China

a r t i c l e

i n f o

Article history: Received 3 May 2012 Received in revised form 30 September 2012 Accepted 26 October 2012 Available online 2 November 2012 Keywords: Human hair keratins Dermal substitutes Biocompatibility Biodegradation Wound healing

a b s t r a c t The cytocompatibility, in vivo biodegradation and wound healing of keratin biomaterials were investigated. For the purposes, three groups of keratin scaffolds were fabricated by freeze-drying reduced solutions at 2 wt.%, 4 wt.% and 8 wt.% keratins extracted from human hairs. These scaffolds exhibited evenly distributed high porous structures with pore size of 120–220 μm and the porosity >90%. NIH3T3 cells proliferated well on these scaffolds in culture lasting up to 22 days. Confocal micrographs stained with AO visually revealed cell attachment and infiltration as well as scaffold architectural stability. In vivo animal experiments were conducted with 4 wt.% keratin scaffolds. Early degradation of subcutaneously implanted scaffolds occurred at 3 weeks in the outermost surface, in concomitant with inflammatory response. At 5 weeks, the overall porous structure of scaffolds severely deteriorated while the early inflammatory response in the outermost surface obviously subsided. A faster keratin biodegradation was observed in repairing full-thickness skin defects. Compared with the blank control, keratin scaffolds gave rise to more blood vessels at 2 weeks and better complete wound repair at 3 weeks with a thicker epidermis, less contraction and newly formed hair follicles. These preliminary results suggest that human hair keratin scaffolds are promising dermal substitutes for skin regeneration. © 2012 Elsevier B.V. All rights reserved.

1. Introduction Porous biomaterials with a well-interconnected pore structure are of great interest in the fields of tissue engineering and regenerative medicine [1–3]. Their porous architecture secures the diffusion of nutrients and metabolites for the cell population and permits in-growth and differentiation of cells [3,4]. Seeding and culturing of selected target cells onto these scaffolds gives rise to different tissue-engineered constructs ex vivo, and meanwhile scaffolds could be also directly used in vivo as tissue substitutes which play critical roles in repairing various tissue defects [5]. In restoring skin defects, for example, biomaterials serve as a temporary shelter to close the skin defect so as to reduce the loss of water, electrolyte and protein and also function as a vital framework for cell anchoring and differentiation during the wound healing process [2,6,7]. Nowadays, various natural and/or synthetic biomacromolecules have been investigated in order to develop various extracellular matrix (ECM) analogues for these functional purposes [1,8–12]. Owing to the presence of cell adhesion sequences, biologically-derived protein biopolymers occupy a central place in burgeoning efforts directed toward the development of biomimetic ECM scaffolds [1]. Collagen as one of the most widely used biologically-derived biomaterials has low its immunogenicity and good affinity to cells. Having been safely used in medical fields for decades, collagen-based biomaterials are now well ⁎ Corresponding author. Tel./fax: +86 28 8541 2102. E-mail address: [email protected] (X. Li). 0928-4931/$ – see front matter © 2012 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.msec.2012.10.011

documented in developing various ECM analogues [8,12–16]. The wide use of other biologically-derived proteins for this purpose also includes fibrin [12], silk fibroin [17] and elastin [18]. In contrast, much less studies have contributed to keratins-based biomaterials although the first biocompatible and biomedical study of keratins was given in 1982 [19,20]. Keratins are a group of cysteine-rich fibrous proteins found in filamentous or hard structures such as hairs, wools, nails, horns and etc. It is recently found that keratins extracted from human hair fibers contain a cell adhesion motif of leucine-aspartic acid-valine (LDV) [21] as well as some regulatory molecules capable of enhancing nerve tissue regeneration [22]. Owing to the abundance and regeneration nature of wools and hairs, these-derived keratins have now received extensive investigations as a new kind of biomaterials [10,23]. The biomedical use of keratins is based on chemical reductive degradation of the interlinked S-S bonds of keratinous materials via oxidative [22] or reductive extraction [21,24,25]. Until now, various structural forms of extracted keratins together with some chemical modifications [26] or combinations [27–29] have been fabricated for biomedical purposes, including films [24,30,31], hydrogels [22] and scaffolds [21,32,33]. The latter generally involves freeze-drying the reduced keratinous products [21,32], and the introduction of some porogen was also reported, such as granulated sodium chloride [34] and calcium alginate [35]. However, most of the publications primarily focused on extraction, physicochemical and cellular aspects of the prepared keratin biomaterials. Those studies with in vivo biological experiments were really few.

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This study focuses on the biological evaluation of human hair keratin scaffolds for skin wound repair and regeneration. Keratin scaffolds were first fabricated by lyophilizing the reduced solutions at varying concentrations of keratins extracted from human hair fibers. Then, the porous structural features and cytocompatibility of these scaffolds were assessed. Finally, the use of one group of keratin scaffolds for subcutaneous implantation and for treating full-thickness skin defects in rats were conducted to evaluate biodegradation, biocompatibility and wound healing function of keratin biomaterials. These relevant studies are expected to provide important data for supporting tissue engineering applications of keratinous biomaterials. 2. Materials and methods Human hair was obtained from a local barber. Urea, 2-mercaptoethanol, thiourea, ethanol, sodium dodecyl sulfate (SDS) and other chemicals of analytical grade were commercially available and used as received. Cell culture reagents include Dulbecco's modified eagle medium and trypsin (DMEM, Gibco, USA), fetal calf serum and penicillin– streptomycin antibiotics (Chengduhali, China), and acridine orange (AO) (Sigma, USA). The deionized water (18 MΩ cm−1) was used in the present experiments.

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axial pore architecture of keratin scaffolds. In terms of the pore size of these scaffolds, fifty apparent pores were randomly selected, then measured with computerized image analyzer in arbitrary zones. 2.4. Porosity measurement The porosities of three groups of scaffold discs were measured by a liquid displacement method with ethanol as the displacement liquid. In brief, the weighed scaffolds (Wo) were immersed in absolute ethanol at room temperature and then placed in a desiccator under a reduced pressure for 5 min to remove air bubbles. Samples were subsequently taken out, wiped gently with a filter paper to remove excess ethanol, and weighed immediately (We). Geometrical volumes (Vs) of the scaffold discs were calculated by measuring the diameters and heights, and the pore volumes (Vp) were measured by ethanol displacement method. The porosity of scaffolds was calculated according to the following equation: P ¼ V p =V s  100%:



V p is defined as ðW e −W o Þ=ρe



where ρe represents the density of ethanol (0.789 mg mL−1).

2.1. Extraction and identification of human hair keratins

2.5. Swelling and water uptake capability

The keratin extraction from human hair fibers was prepared according to the reported methods with some modifications [24,25]. Briefly, human hair was washed with ethanol and external lipids were removed using a mixture of chloroform/methanol (2:1, v/v) for 24 h. The delipidized hair (100 g) was mixed with 25 mM Tris–HCl solution (1.5 L, pH 8.5) containing 2.6 M thiourea, 5 M urea, 75 g SDS and 5% 2-mercap toethanol at 50 °C for 3 days. After the mixture was filtered and centrifuged at 15,000 ×g for 20 min at room temperature, the obtained supernatant was dialyzed against deionized water using cellophane tubing (molecular weight cutoff of about 10 kDa) and the outer water was replaced with distilled water twice a day. The extracted keratins were identified by using sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and Fourier transform infrared spectroscopy analyses.

Three groups of keratin scaffold discs (3 samples in each group) were placed separately in sealed tubes containing PBS. At different soaking intervals at 37 °C, e.g., 2, 4, 8, 16 and 32 h, the swollen samples were removed from the tube, gently blotted with a filter paper to remove the excess liquid, and immediately weighed as Wwet. The swollen samples at 32 h were lyophilized and then weighed as Wdry, as these Wwet values showed no significant change in comparison with those measured at 16 h and even at 8 h. The water uptake percentage (WA%) of these scaffolds was calculated using the formula:

2.2. Fabrication of keratin scaffolds

2.6. Cell culturing

A previously developed freeze-drying protocol was applied to fabricate keratin scaffolds [4]. For this purpose, three groups of reductive solutions at concentrations of 2 wt.%, 4 wt.% and 8 wt.% keratins were prepared by diluting the dialyzed solution which contained 9.2 wt.% keratins. These solutions were poured into the flat-bottom tubes (10 mm in diameter), and then placed into a freeze-dryer (LGJ-18S, Songyuan Huaxing Co. Ltd, Beijing). Controlled with the preset programming, the solutions were kept frozen at −60 °C for 6 h and subsequently maintained at 0 °C for drying in vacuum (1 Pa) for 16 h. The obtained products were thereafter designated as 2 wt.%, 4 wt.% and 8 wt.% keratin scaffolds. These scaffolds were washed five times with phosphate buffer saline (PBS, pH 7.4), re-lyophilized using the protocol mentioned above. Discs (2 mm in thickness) were cut with a razor and stored in a desiccator for the following experiments.

Three groups of keratin scaffolds were used to evaluate cytocompatibility and cellular behaviors. All the keratin discs (diameter: 10 mm, thickness: 2 mm) were sterilized with 70% (v/v) alcohol and then rinsed six times with sterile PBS. Before NIH3T3 cell seeding, scaffolds were conditioned with complete DMEM overnight at 37 °C. Fibroblasts of the third passage were harvested by trypsinization using a 0.05% trypsin/EDTA, counted and re-suspended in complete medium. 3000 cells in 10 μl culture medium were seeded onto the pre-wetted scaffold discs placed in 24-well culture plates. After incubation for 4 h, the cell-seeded scaffolds were supplemented with 1 mL complete medium. Then, the cell-seeded scaffolds were cultured for up to 22 days under a static condition in an incubator with the medium changing every three days.

2.3. Pore architecture examination The cross-sectional morphologies of three groups of keratin scaffolds were observed on a Hitachi S-4800 field emission scanning electron microscope (FE-SEM) at the working voltage of 3.0 kV. All the samples were coated with a thin layer of gold before SEM observations. Furthermore, these scaffold discs were also embedded in paraffin and then longitudinal sections were prepared. Randomly selected sections were observed on an optical microscope (Olympus, Japan) to examine the

  WA ð% Þ ¼ Wwet −Wdry =Wdry  100 ð% Þ

2.6.1. Cell proliferation Cell viability seeded on keratin scaffolds was examined by using MTT (3-(4, 5-dimethy) thiazol-2-yl)-2, 5-diphenyltetrazolium bromide) assay. At a selected culture interval (2, 7, 12, 17 and 22 days) each well with one cell-cultured scaffold was injected with 50 μL MTT (5 mg mL−1, Sigma) solution. After further culturing for another 4 h, the supernatant was carefully aspirated and discarded. 1 mL dimethylsulfoxide (DMSO) was then added into each well and the solution was mixed thoroughly with a vortex mixer for 15 min. Finally, the absorbance of 200 μL solutions in a 96-well plate was recorded by using a microplate reader at 570 nm on a Bio-Rad 550 spectrometer.

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3. Results

2.6.2. Cell growth within keratin scaffolds by confocal microscopy After cell culturing of 2, 12 and 22 days, the growth behaviors of NIH3T3 cells onto the scaffolds were observed by using confocal laser scanning microscopy. The cell-cultured scaffolds were taken out from the culture plate and washed with PBS for 3 times. After the cells were stained with 10 μg mL −1 acridine orange (AO) solution at room temperature for 15 min, the cell-cultured scaffolds were thoroughly washed with PBS to remove unreacted AO. Green fluorescence of stained cells was obtained on a Leica SP5 CLSM with laser lines at the excitation/emission wavelength of 500/526 nm. A series of 10 μm thick slices along the z-axis were taken from the surface and down to a depth of 250 μm, viewing as three dimensional micrographs to show cell attachment and proliferation within the scaffolds.

The SDS-PAGE showed that the extracted keratins mainly consisted of keratins with molecular weights of 40–60 kDa and matrix proteins with molecular weights of 15–30 kDa (Fig. 1a). The FT-IR spectrum of extracted keratins is shown in Fig. 1b. The adsorptions at 3300 and 3050 cm−1 are arisen from the N–H stretching vibrations of Amide A and Amide B. Amide I, II and III adsorptions of keratins are located at 1650, 1533 and 1236 cm−1 while the strong adsorptions centered at 620 cm−1 are attributed to C-S stretching vibrations, validating the presence of reduced (−SH) form of keratins [21,29,36].

2.7. In vivo animal evaluation

3.2. Porous microstructure and water uptake of keratin scaffolds

The animal experiments were performed following approval of the Animal Care and Ethics Committee of Sichuan University, Chengdu, PR China. 4 wt.% keratin scaffold discs (diameter: 10 mm, thickness: 2 mm) were chosen to evaluate biocompatibility and biodegradation through subcutaneous implantation as well as biofunctional performance of repairing full-thickness skin defects. These keratin discs (diameter: 10 mm, thickness: 2 mm) were sterilized with 70% (v/v) alcohol and then rinsed six times with sterile PBS. The selection of 4 wt.% keratin scaffolds was mainly based on the consideration that they had medium porous structural features among three groups of fabricated scaffolds as shown in 3.2.

The porous structural characteristics of keratin scaffold discs were revealed by microscopic examination of the respective cross-sections and axial sections of cylindrical samples and by pore size, porosity and water uptake measurements. The cross-sectional HE-SEM micrographs and axial sectional optical micrographs are given in Fig. 2. The cross sections for SEM were cut with a razor whereas the samples for optical observation were prepared by axially sectioning the discs embedded in paraffin wax. All the three groups of 2 wt.%, 4 wt.% and 8 wt.% keratin scaffolds present an evenly distributed open pore structure, according to SEM and optical observations, suggesting good porous interconnectivity. The measured data of pore size range, porosity and water uptake of three groups of keratin scaffolds are listed in Table 1. The measured pore sizes and porosities follow a keratin concentration-dependent fashion, i.e. lyophilizing the solution at a higher keratin concentration yields a scaffold with a smaller pore size and lower porosity. Even so, three groups of scaffolds all exhibited high porosities (>90%), confirming that a highly porous structure was fabricated in the present study. Fig. 3 is the swelling behaviours of 2 wt.%, 4 wt.% and 8 wt.% keratin scaffolds in PBS at 37 °C. A rapid swelling occurred during the early soaking of 4 h, but after 8 h the swelling tendency obviously attenuated, especially for 2 wt.% and 4 wt.% scaffolds. The water uptake percentage of 2 wt.%, 6 wt.% and 8 wt.% keratin scaffolds at 32 h is 376±39%, 421± 21% and 554±35%, respectively. The greater standard deviation recorded for the swollen samples at 32 h is probably due to the variant physical deformation during handling a specific, highly porous, swollen sample after soaking of longer intervals. In fact, the swelling tendency plotted with the measured average water uptake percentages (Fig. 3) shows that the swelling equilibrium reached after soaking of 32 h, whereas this soaking interval for the reported keratin scaffolds was 24 h [34].

2.7.1. Subcutaneous implantation Nine Wistar rats (180–220 g) were randomly divided into three groups (3 rats for each group) used to evaluate the biodegradability and biocompatibility of keratin scaffolds subcutaneously implanted for 1 week, 3 and 5 weeks. The dorsal hair of rats was clipped under anaesthesia. After disinfection of the skin with iodine, a cut was made and one sterilized keratin disc was then implanted into the subcutaneous tissues. At selected implantation intervals, the rats were sacrificed by euthanasia. The harvested scaffolds together with the surrounding tissues were fixed in 10% formaldehyde, embedded in paraffin wax, sectioned and stained with hematoxylin and eosin (H&E) staining for histological observations. 2.7.2. Repairing full-thickness skin defects Nine Wistar rats (180–220 g) were used and randomly divided into three groups (3 rats for each group) for treating full-thickness skin defect tests of 1 week, 2 and 3 weeks. Under anaesthesia with pentobarbital, the dorsal hair of rats was clipped and the exposed skin was disinfected with iodine. Two full-thickness wounds of 10 mm in diameter were prepared by excising the dorsum of a Wistar rat. One excised wound was dressed with a sterilized keratin scaffold disc while the other was not dressed as a blank control. Both wounds were covered with two layers of vaseline gauze and subsequently bounded with an stainless steel framework for protection. At the first, second and third week postoperatively, the rats were sacrificed by euthanasia. A skin wound tissue harvested from the central regions of the wound were fixed in 10% formaldehyde, embedded in paraffin wax, sectioned and stained with hematoxylin and eosin (H&E) staining for histological investigation of wound healing effects. 2.8. Statistical analysis All quantitative measurements were conducted in triplicate unless specially stated and the resulting data were expressed as mean±standard deviation (SD). The experiment for MTT assay was run in five replicates with each group of scaffold sample (n=5). Statistical differences for the proliferation assay between groups were analyzed according to a paired Student's t test, and the P valueb 0.05 was considered statistically significant.

3.1. Identification of extracted keratins

3.3. Cell growth and infiltration within the scaffolds MTT data (Fig. 4) show that NIH3T3 cells proliferated properly on keratin scaffolds (3000 cells/scaffold) in different culture intervals lasting up to 22 days under the static culturing condition. In general, a slightly higher proliferation rate was achieved in cell-seeded scaffolds fabricated with a lower keratin concentration. 2 wt.% scaffolds yielded the highest proliferation rate starting on day 7. Cells grown on this group of keratin scaffolds were at a plateau growth stage in between day 12 and day 17, but a further proliferation growth was recorded on day 22. In contrast, a steady proliferation growth was observed on other groups of keratin scaffolds. The cells grown on three groups of keratin scaffolds were also stained with AO, and then imaged on a confocal laser scanning microscope to reveal cell growth and infiltration into the porous architecture of keratin scaffolds. The three dimensional CLSM micrographs in Fig. 5 were obtained after NIH3T3 culturing of 2, 12 and 22 days onto 2 wt.% (Fig. 5a–c), 4 wt.% (Fig. 5d–f) and 8 wt.% (Fig. 5g–i) keratin scaffolds. These confocal images, viewed along the z-axis, delineated a

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Fig. 1. Identification of extracted human hair keratins. (a) SDA–PAGE: Lane 1 — molecular weight markers, Lane 2 — human hair keratins, and (b) FT-IR spectrum.

well-stacked three dimensional structure, not only visually verifying cell proliferation with prolonging culture time, but also showing the occurrence of cell infiltration under the present static culturing condition. Meanwhile, the well-stacked three dimensional porous structure imagined by AO-stained cells, especially those achieved on the day 22 day (Fig. 5c, f, i and the respectively magnified), also validates the pore architectural stability of keratin scaffolds after a long-term cell culturing. 3.4. Subcutaneous implantation The histological sections of 4 wt.% keratin scaffolds implanted subcutaneously are shown in Fig. 6 to acquire the preliminary information about the biodegradability and biocompatibility of keratin scaffolds. One week postoperatively, the inflammatory response was located mainly at the outer surface region of the scaffold (Fig. 6a). Compared with the architectural integrity of inner porous structure of this scaffold, the obvious pore deformation could be observed in its outer region. The concomitant

inflammatory cells are believed to be due to the preliminary keratin degradation (Fig. 6b). The inward infiltration of inflammatory cells proceeded further into the scaffold three weeks postoperatively (Fig. 6c). Although the overall porous structure of keratin scaffold still kept its integrity, the explicit deterioration of pore structure could be noticed in the outermost surface of the scaffold sample (see arrowed in Fig. 6d). Five weeks postoperatively, the severe deterioration of the overall pore architecture is evident (Fig. 6e). This event corresponds to the population of inflammatory cells within the deteriorated scaffold (Fig. 6e and f). However, it is noted that the early inflammatory response in the outermost surface region (Fig. 6a) significantly subsided at this moment (see arrowed in Fig. 6e). 3.5. Histological examination of repairing full-thickness skin wounds HE-stained sections in Fig. 7 are the typical of those of full-thickness skin wounds treated without (the blank control) or with 4 wt.% keratin

Fig. 2. Cross-sectional FE-SEM micrographs (upper column) and axial sectional optical micrographs (lower column) of 2 wt.%, 4 wt.% and 8 wt.% keratin scaffolds.

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Table 1 Measured porosity, pore size range and water uptake percentage of human hair keratin scaffolds. Data represent as mean± standard deviation. Human hair keratin concentration (wt.%)

Porosity (wt.%) Pore size range (μm) Water uptake percentage (%)

2

4

8

94.8 ± 2.7 200–220 376 ± 39

93.6 ± 0.8 180–200 421 ± 21

91.2 ± 1.5 120–150 554 ± 35

scaffolds one and two weeks postoperatively. An obvious inflammatory response was observed at the first week. In the blank control, inflammatory cells presented with a diffuse distribution mode (Fig. 7a). In contrast, these cells were populated and delineated a definite separation line in the wound treated with a keratin scaffold (Fig. 7b). Two weeks after surgery, compared with the blank control (Fig. 7c), a larger number of blood vessels are observed in the wound treated with a keratin scaffold (as indicated by black arrows in Fig. 7d), suggesting early vascularization. The existence of keratin-like substance could be indentified with white arrows at the upper region of the wound. However, it is hard to forge a porous structure by these arrowed objects, suggesting occurrence of the degradation of the scaffold. The green dotted rectangle in the blank control (Fig. 7c) shows the presence of obvious contraction without the use of the keratin scaffold. The histological sections three weeks postoperative are given in Fig. 8. The area dotted by white lines indicates a not completely healed wound which is smaller with a keratin scaffold (Fig. 8c) than without using it (Fig. 8a). Even so, epithelium regeneration is observed. The newly synthesized epidermis in the wound treated with a keratin scaffold (Fig. 8c, d) is more complete, thicker than that in the blank control which had more contraction (long white arrows in Fig. 8a, b). The existence of undigested keratin scaffold residues is indicated by black arrows, mainly located at the bottom of the healing wound (Fig. 8c). In addition, the short white arrows in the wound treated with a keratin scaffold (Fig. 8c) show the new formation of hair follicles, confirmed histologically in the magnified Fig. 8d. 4. Discussion Biological evaluation is indispensable in developing materials for biomedical purposes. Keratin proteins extracted from hairs, wools, nails and horns have long been considered as a new type of promising biomaterials. However, until recently, much fewer of the keratin developments have been applied in animal models of tissue regeneration, in contrast to vigorous studies of keratin extraction, structural confirmation, product fabrication and cytocompatible assay [23]. The reported in vivo applications of keratin biomaterials scarcely included as hemostat [37] and in

Fig. 3. Swelling curves of 2 wt.%, 4 wt.% and 8 wt.% keratin scaffolds in pH 7.4 PBS at 37 °C.

Fig. 4. MTT data of NIH3T3 cells cultured on three groups of keratin scaffolds for up to 22 days. Data are presented as the average ± standard deviation (n = 5, pb 0.05).

regeneration of peripheral nerves [22] and amelioration of cardiac dysfunction [38], and the applied keratins were solely extracted from human hair fibers. The present provides systematic in vitro and in vivo investigations of keratins scaffolds by lyophilizing the reduced solutions. The extracted keratins from human hairs in the present study mainly consisted of keratins with molecular weights of 40–60 kDa and matrix proteins with molecular weights of 15–30 kDa (Fig. 1), in agreement with the reported data [21,31]. Freeze-drying of reductive keratin solutions, according to our previously developed protocol [4], yielded keratin scaffolds with open pores and good porous architecture (Fig. 2). Pore size, porosity and water uptake capability were dependent upon keratin concentration (Table 1). With the increase in keratin concentration, pore size and porosity decreased but water uptake percentage increased in the case of 4 wt.% and 8 wt.% keratin scaffolds. This keratin concentrationdependent porous structural feature was due to higher protein mass per unit volume of keratin solution [3]. It is noted that three groups of keratin scaffolds had high porosities ((>90%). In contrast, 4 wt.% and 8 wt.% keratin scaffolds with a gradually smaller pore size range offered increasing surface in contact with water, thus giving rise to higher water uptake percentages. The hydrophilic character of the present keratin scaffolds was desirable for cell attachment [21]. NIH3T3 cell culturing experiments confirm good cytocompatibility of the prepared keratin scaffolds, as indicated by MTT results which recorded the continuous proliferation after seeding and culturing for as long as to 22 days (Fig. 4). A higher proliferation rate achieved onto 2 wt.% and 4 wt.% scaffolds supports the predominate role of higher porosity and larger pore for cell growth at static culturing condition [3]. Confocal microscopic examinations of the cell-cultured scaffolds visually show cell proliferation behavior through growth on the top surface as well as infiltration into the scaffolds (Fig. 5). Keratin proteolysis is achieved by specific enzymes, such as caspase or cathepsin, released from lysosomes [39]. Currently, biodegradation evaluation of keratin biomaterial primarily focused on the in vitro proteolysis in the presence of trypsin and pepsin (common proteolytic enzymes). As is known, the in vivo biodegradation of biomaterials is dependent upon the material structure, the implantation site and function. Subcutaneous implantation of keratin films and bars indicated a very slow biodegradation [24,40], e. g., between 3 and 6 weeks after surgery cavitations and fissures were only observed at the surface of the bars [40]. Different from the dense structural form of films and bars, the interconnected porous architecture of keratin hydrogels or scaffolds permitted the early cell infiltration but the detailed biodegradation was not

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Fig. 5. Confocal micrographs stained with AO showing NIH3T3 cell growth and attachment after culturing for 2, 12 and 22 days on three groups of keratin scaffolds: (a–c) 2 wt.%, (d–f) 4 wt.% and (g–i) 8 wt.% scaffolds. These are the z-axial three-dimensional micrographs constructed by 25 scanning sections with each 10 μm thick along the z-axis.

Fig. 6. Histological sections of 4 wt.% keratin scaffolds subcutaneously implanted for 1 week (a, b), 3 weeks (c, d) and 5 weeks (e, f).

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Fig. 7. Histological sections of wounds treated without (a, b) and with (c, d) 4 wt.% keratin scaffolds 1 and 2 weeks postoperatively. Green dotted rectangle shows the presence of obvious contraction. White arrows indicate the residual scaffolds. Black arrows indicate the capillary-like structures.

reported [22,37]. In the present study, the initial biodegradation sign was observed at 3 weeks in the uttermost surface of subcutaneously implanted 4 wt.% scaffolds, and at 5 weeks, the pore architecture of keratin scaffolds severely deteriorated (Fig. 6). When the keratin scaffolds were used to dress the full-thickness skin wounds, a faster biodegradation was observed. At 3 weeks the keratin residues were mainly located at the bottom of the healing wounds and densely populated with cells

(Fig. 8). This similar positioning of undigested dermal substitutes was previously reported [41]. A faster biodegradation rate achieved in the dressing wound is believed to be related to repair-triggering a cascade of robust cellular activities together with the presence of higher amount of keratinolysis enzymes. It is worthy to note that a faster degradation of keratin biomaterials in the heart mainly arose from the hemodynamic and contractile forces [38].

Fig. 8. Histological sections of wounds treated without (a, b) and with (c, d) the 4 wt.% keratin scaffolds 3 weeks after surgery. Long white arrows (a, b) present the uneven reconstruction epidermis. Black arrows indicate the keratin scaffold residues. Short white arrows (c, d) indicate the hair follicles. White dotted lines indicate not completely healed wound.

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Owing to the lack of the spontaneous healing, scar formation and contraction occur inevitably for full-thickness skin wounds. The use of various dermal substitutes on wound healing is found, to a varying extent, to enhance skin regeneration yielding less contraction, earlier vascularization and improved epidermal formation [2,6,7,41]. Keratins play important roles in skin, such as regulators of protein synthesis, epithelial cell growth and hair follicle cycling [42]. The development of keratin biomaterials for wound healing was long proposed in granted patents as reviewed in [10,23], the relevant biological performances have not been found yet, according to the best of our knowledge. In comparison with the blank control, the present study revealed that 4 wt.% scaffolds had more blood vessels at 2 weeks (black arrows in Fig. 7d) and better complete wound repair at 3 weeks with a thicker epidermis, less contraction and newly formed hair follicles (Fig. 8c and d). These enhanced skin regeneration effects seem to be due to the intervention of wound healing by using keratin scaffolds. A former study reported the neuroinductive capability of human hair keratin materials and further suggested that the regulatory molecules contained in the extracted keratins were responsible for enhancing nerve tissue regeneration [22,43]. However, further studies relevant to obtain quantitative data and even specific immunohistochemical assays together with repairing larger defects are needed to ascertain the present findings.

5. Conclusions In summary, human hair keratin scaffolds were prepared in the present work by freeze-drying reductive solutions with varying keratin concentration. These well-interconnected scaffolds were hydrophilic and had good cytocompatibility. The use of keratin scaffolds for subcutaneous implantation and for treating full-thickness skin defects in rats confirmed the biodegradation, biocompatibility and wound healing function of keratin biomaterials. Compared with the self healing process of full-thickness wounds, keratin scaffolds led to earlier vascularization and better skin repair with a thicker epidermis, less contraction and newly formed hair follicles. The promising biological performance of human hair keratin scaffolds is expected to be further investigated.

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Acknowledgements

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This work is supported by the National Basic Research Program of China (no. 2012CB933603), the National Natural Science Foundation of China (no. 30970729), the Doctoral Programs Foundation of the Ministry of Education of China (no. 20090181110067) and the Engineering Research Center of Biomass Materials (SWUST) of the Ministry of Education of China (no. 10zxbk04).

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Biological evaluation of human hair keratin scaffolds for skin wound repair and regeneration.

The cytocompatibility, in vivo biodegradation and wound healing of keratin biomaterials were investigated. For the purposes, three groups of keratin s...
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