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Carboxymethyl cellulose-human hair keratin hydrogel with controlled clindamycin release as antibacterial wound dressing
Journal Pre-proofs Carboxymethyl cellulose-human hair keratin hydrogel with controlled clindamycin release as antibacterial wound dressing Soodeh Sadeghi, Jhamak Nourmohammadi, Azadeh Ghaee, Neda Soleimani PII: DOI: Reference:
S0141-8130(19)34286-2 https://doi.org/10.1016/j.ijbiomac.2019.09.251 BIOMAC 13588
To appear in:
International Journal of Biological Macromolecules
Received Date: Revised Date: Accepted Date:
9 June 2019 17 September 2019 30 September 2019
Please cite this article as: S. Sadeghi, J. Nourmohammadi, A. Ghaee, N. Soleimani, Carboxymethyl cellulosehuman hair keratin hydrogel with controlled clindamycin release as antibacterial wound dressing, International Journal of Biological Macromolecules (2019), doi: https://doi.org/10.1016/j.ijbiomac.2019.09.251
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© 2019 Published by Elsevier B.V.
Carboxymethyl cellulose-human hair keratin hydrogel with controlled clindamycin release as antibacterial wound dressing Soodeh Sadeghi1, Jhamak Nourmohammadi1, *, Azadeh Ghaee1, Neda Soleimani2
1-Division of Biomedical Engineering, Department of Life Science Engineering, Faculty of New Sciences and Technologies, University of Tehran, Tehran, Iran 2- Departments of Microbiology and Microbial Biotechnology and Nanobiotechnology, Faculty of Life Sciences and Biotechnology, Shahid Beheshti University, Tehran, Iran
*Corresponding Author: Jhamak Nourmohammadi; PhD. Department of Life Science Engineering, University of Tehran, P.O. Box: 143951561, Tel: +98-21- 86093264 Email: [email protected]
Abstract This study offers a new antibacterial wound dressing from carboxymethyl cellulose (CMC)-human hair keratin with topical clindamycin delivery. Keratin was successfully extracted from human hair. Different sponges fabricated by changing CMC to keratin ratio were characterized and compared. Halloysite nanotubes were used as carriers to control the clindamycin release. Various characterization techniques were used to determine the effects of keratin addition on the structure, morphology, physical properties, drug release, antibacterial activity, and cellular behavior of CMC hydrogels. As proved by SEM and EDS, porous structure with interconnected pores was successfully formed and clindamycin-loaded HNTs were uniformly dispersed within the porous structures. Increasing the keratin in CMC hydrogel not only lowered its water vapor transmission rate to a suitable range for wound healing but also improved the water stability of CMC hydrogel. The in vitro release study indicated that clindamycin was released slower in samples containing higher keratin and the Fickian diffusion mechanism controlled their release profile. The fabricated dressing effectively inhibits S. aureus bacterial colonies growth after 24 h. Fibroblast culturing on the fabricated sponges indicated that cellular attachment, proliferation, and spreading were significantly enhanced with increasing the keratin amount.
Keywords: Carboxymethyl cellulose; Keratin; Clindamycin; Wound dressing; Fibroblast attachment.
Introduction The skin has a key role in protecting one’s body against external invasions [1]. Being the first and the largest layer in contact with the surrounding environment, the skin is more likely to be injured. Thus, using an effective wound dressing has been recognized as an effective approach toward accelerating the healing process. The suitable wound dressing should be able to provide an optimum condition around the wound, which best mimics the extracellular matrix (ECM) of skin to help to permit epithelial cell movements, passing oxygen properly, allowing a moist environment, draining wound exudates and considerably preventing the wound from being infected [2]. In this regard, carboxymethyl cellulose (CMC)-based hydrogels come to special consideration because of their abundance, transparency and low cost [3, 4]. The high hydrophilicity of CMC leads to the high absorption of wound exudates; it also provides a moist environment around the wound and prevents the tissue from losing water which is of great importance in burn and diabetic wounds [5-8]. Regardless of such improvements, the relatively poor cell adhesion of CMC hydrogels has been reported in previous studies. Moreover, the less antibacterial activity and water stability limit their practical applicability as a wound dressing [9, 10]. To make up for this shortcoming, the blending of CMC with other polymers has been reported previously [11, 12]. Choosing a nature-based protein beside CMC in fabricating the scaffolds can be a step closer to making a construction similar to the natural ECM. Keratin is a cysteine-rich intracellular cytoskeleton protein which exists in hair, epidermis, nails, feather, wool, hoof and horn [13]. In recent years human-derived keratin has gained much attention in the various biomedical engineering applications because of its low cost, excellent biocompatibility, availability, biodegradability, hemostatic, and less immune reactions. Moreover, it encourages a more powerful cell-matrix response because of the existing cell-binding motifs like glutamic acid-aspartic acid-serine (EDS), leucine-aspartic acid-valine (LDV) and arginine–glycine– aspartic acid (RGD) similar to ECM proteins such as collagen and fibronectin [14]. Yamauchi et al. [15] proposed that keratin-coated substrate has shown better L929 fibroblasts attachment and growth compared to the ones coated with collagen. Park et. al [16] have also reported that keratin extracted from human hair is more effective in healing wounds due to the almost complete regeneration of epithelial cells. Thus, it seems that the combination of keratin and CMC not only could resemble ECM composition but also could improve the cellular response of CMC. However, the aforementioned dressing is not effective in killing the bacteria at the wound sites. Infection is very probable in severe skin damages which can prolong the healing process [17]. Currently, multiple wound dressings have been studied for the localized delivery of antibiotics to the infected wounds in a controlled manner.
Halloysite nanotubes (HNTs) have been introduced as a potential substance for target delivery of various bioagents and drugs at a defect site [18]. HNTs (Al2Si2O5(OH)4.nH2O) are considered as biocompatible one-dimensional nanotubular aluminosilicate clays that have been widely studied in various biomedical applications [19]. Due to the high aspect ratio, low density, and hydrophilicity, HNTs can easily disperse within hydrophilic biopolymers without the need for surface pre-treatment [20-22]. Moreover, their high surface area provides a very good absorption property which allows drugs to be loaded easily into and on them[23]. In this regard, this study introduces and characterizes a novel antibacterial wound dressing from CMC, keratin containing clindamycin using HNTs as a drug carrier. Clindamycin is an efficient antibiotic for healing serious skin and soft tissue infections especially those caused by staphylococcus aureus (s. aureus) such as burn wound infection [24]. Different dressings were fabricated by changing the CMC to keratin ratio and citric acid was used as a crosslinker. The effect of keratin addition on the morphology, porosity, water uptake, water vapor transmission rate, and clindamycin release profile of the composites was studied and then compared to those of the pure CMC sample. In the end, the antibacterial activity, as well as the behavior of seeded fibroblast on different nanocomposites, were studied. 2. Materials and methods 2.1. Materials Human hair was gained from local Barber’s. Besides, Tris-HCL (PH 8.5, Merck, Germany), thiourea (Merck, Germany), Urea (Merck, Germany), 2-mercaptoethanol (Merck, Germany), absolute ethanol (Merck, Germany), methanol (Merck, Germany), chloroform (Merck, Germany), carboxymethyl cellulose (Sigma, USA), halloysite nanotubes (Sigma, USA), cellulose dialysis tube (12KDa, MWCO; Sigma, USA), and Clindamycin (sepia Co., Iran) were purchased. All used chemicals were of analytical grade. 2.2. Isolation of keratin from human hair Keratin was extracted from non-colored hair based on the Shindai method as described previously by Lee et al. [25]. In brief, chopped hair was washed with ethanol and then soaked in methanol/chloroform mixture (1:2 (v/v). After 24h soaking, the delipidated hair was washed thoroughly with deionized water 3 to 4 times and then dried. The dried hair was immersed in the Shindai solution (which is composed of 25 mM Tris-HCl, 2.6 M thiourea, 5 M urea, and 5% (v/v) 2-mercaptoethanol) and incubated at 50℃ for 72 h. Next, the mixture was passed through the filter paper and centrifuged 10000 rpm for 20 min to separate hair residues from the liquid. The obtained supernatant was dialyzed in
cellulose tubing against distilled water for 3 days. The concentration of the keratin in the obtained solution was about 8 wt.%, which obtained by weighing the keratin powder after lyophilizing corresponding to a known volume. The dialyzed solution was diluted and then kept at 4oC for further use. The extracted keratin was analyzed using Fourier transform infrared spectroscopy (FTIR, Perkin Elmer, Frontier) over the range of 400-4000 cm-1. 2.3. Loading of clindamycin with HNTs and characterization Clindamycin (2 g) was first dissolved in deionized water (40 mL) with the assistance of stirrer for about 20 min at room temperature. Then HNTs (2 g) was added to the drug solution and stirred for 10 min under vacuum. After that, the suspension was removed from the vacuum, stirred vigorously for 24 h and then centrifuged (7000 rpm) for 30 min. The supernatant was thrown away and the obtained precipitate was dried at 50 oC for 24 h. The loading efficiency of the drug was measured using UV-Vis spectroscopy at the wavelength of 210 nm from the following equation [26]. A calibration curve was plotted using different clindamycin concentrations ranging from 5 to 30 μg/mL. 𝐶𝑙𝑖𝑛𝑑𝑎𝑚𝑎𝑐𝑦𝑛 𝑙𝑜𝑎𝑑𝑖𝑛𝑔 𝑒𝑓𝑓𝑖𝑐𝑖𝑒𝑛𝑐𝑦 (%) =
𝐶0 𝑉 − 𝐶𝑠 𝑉 × 100 𝐶0 𝑉
C0 is the initial concentration of clindamycin, Cs is the concentration of clindamycin in the supernatant, and V is the volume of the solution. The morphology of HNTs was studied using SEM (Mira 3-XMU, Czech Republic)). Moreover, the structural changes of HNTs after clindamycin loading was assessed using Fourier transform infrared spectroscopy (FTIR; Bruker, Germany). 2.4. Preparation of clindamycin-loaded HNTs-CMC-keratin nanocomposites For pure CMCs sample, CMC powder (2 g) was added gradually to 100 mL deionized water and then stirred gently until completely dissolved. In the case of CMC-keratin composites, different weight ratios of keratin and CMC solutions (0:1, 1:2, 1:1, and 2:1) were mixed thoroughly while the Keratin–CMC concentration in the final mixture was kept constant (2% w/v). Besides, 5 wt.% (weight of polymers) citric acid as a cross-linker and 3 wt. % glycerol (weight of polymers) as a crosslinking extender was also added to each solution [27] . After mixing for 3h, clindamycin-loaded HNTs (2 wt. % of total polymers) were added to the different keratin-CMC solutions and stirred for about 15 min to disperse the nanoparticles evenly all over it, after that the mixture was poured into the 48-well tissue culture polystyrene plates. Subsequently, the plates were stored in -70oC overnight and then freeze-dried. Next, the samples were crosslinked at 70oC for 72 h. The code and the composition of the constructed scaffolds are outlined in Table 1. The fabricated dressings were coded as KC (0:1), KC (1:2), KC (1:1), and KC (2:1) based on the changing
keratin/CMC volume ratios (as noticed above). The Photographs of the fabricated dressings are shown in Fig.1. 2.5. Characterization of fabricated nanocomposites The morphology of the studied nanocomposites was assessed using SEM (Mira 3-XMU, Czech Republic). The mean pore size and their distributions were calculated by Image J software (National Institutes of Health, Bethesda, MD, USA). Moreover, the porosity percentages were calculated for each SEM image as follows: 𝑃𝑜𝑟𝑜𝑠𝑖𝑡𝑦 (%) =
𝐴𝑝 × 100 𝐴𝑇
Where, AP is the total area of pores in each cross-section, AT is the total area of the image in the same cross-section. To verify the distribution of HNTs in the scaffolds energy dispersive spectroscopy (EDS) elemental mapping of Al and Si was used to demonstrate the uniformity of HNT’s dispersion throughout the scaffolds. FTIR was performed to study the structural properties of the fabricated nanocomposites. Water absorption is another characteristic of biomaterials that should be considered especially when used as wound dressings. For obtaining the initial weight of samples (Wo), scaffolds were weighed before immersion in deionized water. After immersing and being incubated at 37°C for about 24 h, swelled scaffolds were weighed again (W s) while the excess water wiped off gently with a filter paper. The amount of equilibrium water uptake was obtained by the following equation [28]: Eqiulibrium water uptake (g/g) =
𝑊𝑆 − 𝑊0 𝑊0
The ability of the fabricated nanocomposites to control water loss was evaluated by measuring their water vapor transmission rate (WVTR) based on our previous study [29]. Briefly, the sample was fixed on top of a glass vial (A= 1.5 cm2) filled with deionized water. The assembly sealed completely using Teflon tape, weighed and then placed into an incubator (T= 32 ± 1 ºC) with a relative humidity of 37%. At each hour, the weight loss of the assembly was measured and plotted versus time. WVTR was calculated according to the following equation: WVTR (g/𝑚2 /day) =
Slope × 24 A
The weight loss of each sample was measured by soaking the sample with the initial weight (W 0) in phosphate buffer saline (PBS; pH = 7.4) at 37 ºC. At predetermined times, the samples were removed, completely dried and weighed (Wd). The weight loss (%) was measured as follows [30]: Weight loss(%) =
𝑊0 – 𝑊𝑑 × 100 𝑊𝑑
The Proteins adsorption on each sample was performed to demonstrate the blood compatibility of the fabricated nanocomposite dressings. Thus, each sample with known weight of 20 mg were inserted in 2 mL of Bovine Serum o
Albumin (BSA) solution in PBS (2 mg/mL, pH=7.4). After incubation at 37 C for 2h, the samples were removed from the solution and washed thoroughly with known volume of PBS. Bradford assay was used to measure the amounts of free protein in the remaining BSA solution and washing solution [31]. The BSA adsorption on each sample was calculated as follows [32, 33]: Protein adsorption (mg/g) =
C1 V1 − ( C2 V1 + C3 V2 ) weight
Where C1 is the initial concentration of BSA (here is 2 mg/mL), C2 is the concentration of BSA in the solution after 2h soaking of the nanocomposites, and C3 is the concentration of BSA in the washing solution. Moreover, V1 and V2 are the volumes of the BSA solution and washing solutions, respectively. The coagulant activity of the fabricated nanocomposites was investigated by the Prothrombin time (PT) coagulation assay [34]. Thus, 0.1 mL citrated normal human plasma was dropped onto the surface of the samples and kept at 37 oC for 3 min. Next, 0.2 mL PT reagent was added to each sample and the clotting time was measured. The compression tests of the cylindrical samples (d = 5 mm, h = 10 mm) were performed using Zwick/Roell Z050 -1
with a crosshead speed of 1 mm.min . The compressive strength and modulus were calculated from the stress-strain curve [35]. 2.6. In vitro release study of clindamycin from the nanocomposites The amount of clindamycin released from the composites was measured using UV-Vis spectroscopy (model,) at the wavelength of 210 nm[36]. Thus, each sample was soaked in 5 mL of phosphate buffer saline (PBS; pH=7.4) and kept in a shaker incubator (90 rpm; T= at 37 ºC). At each time point, 2 mL of PBS was removed and an equal volume of fresh PBS was added to each container. The amount of clindamycin released from the composites was measured using a clindamycin standard curve of different concentrations (5 - 30 μg/mL) The clindamycin release kinetics were also assessed with fitting the first 60 % of the release data with KorsmeyerPeppas model [37]: Mt = Kt n M∞ Mt is the amount of drug released at time t, M∞is the amount of released drug at an infinite time, K is the constant
value, and n is the release exponent that indicates the release mechanisms. 2.7. Antibacterial activity The antibacterial activity of each nanocomposite was evaluated based on disk diffusion and colony-forming units (CFU) assay Staphylococcus aureus (S. aureus; gram-positive; ATCC 12600) bacteria [38]. The sterilized sample was immersed in 500 µL of the bacterial solution containing 106CFU/mL and then placed into a shaker incubator (200 rpm) at 37oC. After 24 h incubation, the samples removed from the container and the remaining solution was diluted 10-fold with PBS. Afterward, 20 µL of the diluted solution was taken out and spread on the agar plate with a sterilized loop. The number of viable bacteria on the agar plate was counted after 24 h incubation at 37 oC (Ns) and then compared with the control (Nc) to calculate the bacteria reduction (%): 𝐵𝑎𝑐𝑡𝑒𝑟𝑖𝑎 𝑟𝑒𝑑𝑢𝑐𝑡𝑖𝑜𝑛 (%) =
[(𝑁𝐶 − 𝑁𝑆 )] × 100 𝑁𝐶
2.8. Fibroblast attachment and proliferation L929 fibroblast cells from National Cell Bank of Iran (NCBI; Pasteur institute) were cultured in Dulbecco’s Modification of Eagles Medium (DMEM; Gibco) containing 10 (% v/v) fetal bovine serum (FBS; Gibco) and (1% v/v) Pen-Strep in an incubator at 37 ºC under 5% CO2. The nanocomposites were sterilized using ethanol 70% and PBS. The attachment and morphology of seeded fibroblast cells on each sample were studied using SEM. Accordingly, 1 × 104 cells seeded on each sterilized nanocomposite and then incubated in a humidified incubator (T=37 oC, 5% CO2). After 3 days, fibroblasts were fixed with 4 (%v/v) glutaraldehyde solution followed by dehydration with graded ethanol and then coated with a thin gold layer before imaging. Viability and proliferation of fibroblast cells were evaluated using the MTT assay kit (Bioidea, Iran) [39]. Briefly, fibroblast cells (1 × 104) were first seeded on each sterilized sample and then incubated in an incubator at 37˚C and 5% CO2 for 3 and 5 days. After each incubation time, the culture medium was completely removed and replaced with ready-to-use RPMI 1640 in the kit. Next, 10 μL MTT stock solution (12 mM) was added to each well and the plate kept in the incubator for 3h. After that, 50 μL DMSO was added to each well, pipetting thoroughly, incubated in an incubator for 10 min to dissolve the created formazan crystals. After pipetting, the absorbance was read at 570 nm using an ELISA Reader (Stat Fax-2100, USA). The culture medium without the sample was chosen as a control group.
2.9. Statistical analysis All measurements were done three times for each composition and the results were reported as the mean ± standard deviation (SD). Student’s t-test was used to understand the difference between the studied composition and a p-value less than 0.05 defined as a significant difference. 3. Result and discussion 3.1. IR analysis of extracted keratin Fig. S1 (supplementary files) shows the FTIR spectrum of extracted hair keratin. As expected, the peaks at 3299 cm 1
, 1654 cm-1, 1534 cm−1, and 1244 cm-1 are related to the amide A, amide I, amide II, and amide III bands, respectively
[40]. The presence of these peaks indicated that keratin protein was successfully obtained from the hair. 3.1. Characterization of HNTs before and after clindamycin loading Fig. 2 shows the SEM image of the HNTs, which have shown tubular structures with a length of 380–1100 nm and diameters of 35–102 nm. The FTIR spectra of HNTs before and after clindamycin addition are shown in Fig. 3. As expected, HNTs shows typical peaks at 471 cm-1 (Si-O deformation of Si-O-Si), 550 cm-1 (Al-O deformation of AlO-Si), 751 cm-1 and 795 cm-1 (Si–O stretching), 910 cm-1 (OH deformation of Al-OH groups), 1015 cm-1 (Si-O stretching), 1640 cm-1 (OH deformation of water), 3623 cm-1 and 3695 cm-1 (OH stretching of inner and outer hydroxyl groups) [41]. After clindamycin addition, the clindamycin peaks overlapped with the peaks of HNTs. However, the new peaks at 1560 cm-1, 1690 cm-1, and 2965 cm-1 are related to the C=C, C=O, and C-H stretching vibration of clindamycin [42]. The presence of such peaks approved the successful loading of clindamycin in HNTs. 3.2. Characterization of nanocomposites Fig. 4 shows the cross-section views of different composites. Besides, the mean pore size and the porosity percentage of the scaffolds are represented in Table 1. As expected, all compositions have shown the open and interconnected pores. As summarized in Table 1, pure CMC shows the mean pore sizes of 98 ± 5 mm and porosity percentages of 68± 5%. As expected, there are no significant changes in the mean pore sizes and porosity percentages with increasing keratin content (p>0.05). To identify the presence and distribution of HNTs in each sample, an EDS elemental mapping of Al and Si was done. As shown in Fig. 4, both ions were well-dispersed throughout the pore walls, suggesting that clindamycin-loaded HNTs were present and uniformly distributed in the nanocomposite dressings. Fig. 5 depicts the FT-IR spectra of different nanocomposite dressings. In K0C1, a broad peak centered at 3402 cm -1 is attributed to OH vibration and the peaks at 2931 and 2871 cm-1 are related to C-H vibrations [30]. As expected, the
peaks at 1735 cm-1(carbonyl ester), 1602 cm-1, 1424 cm-1, and 1326 cm-1 (carboxylate), 1235 cm-1 (C-O of ester bonds), and 1120 cm−1 (C-O-C stretching) were observed. The coexistence of C=O and C-O ester bands with carboxylate (COO-) in CMC structure confirms that citric acid crosslinked the CMC structures [43-45]. Besides, the vibrations of Al-O and Si-O of HNTs were also observed at 1041 cm-1, 573 cm-1, and 491 cm-1 [46], respectively, suggesting that HNTs were successfully loaded in CMC structure. After keratin addition, a new shoulder appeared at 1655 cm-1, which is related to the C=O stretching of amide I bonds [40]. Interestingly, the N-H vibrations of amide II bonds (1542 cm1
) [40] were also detected in samples containing a higher amount of keratin (K2C1). The presence of these peaks
indicated the successful crosslinking of keratin and CMC with citric acid. This result is in good agreement with the previous study by H. Xu et.al [47] showing that new amide groups were formed by nucleophilic interaction between carboxyl and amine group when citric acid was used as a cross-linker. Table 1 shows the results of the equilibrium water uptake of each sample after being soaked in deionized water. As summarized in this table, K0C1 has the highest amount of water uptake (3.9± 0.2 g/g), with increasing the amount of keratin the water uptake equilibrium decreased to 2.8 ± 0.09 (g/g) in K1C2, 2.1 ± 0.1 (g/g) in K1C1, and 1.5 ± 0.05 (g/g) in K2C1. As expected, higher CMC weight ratios in a keratin-CMC sample enhanced the water uptake ability, indicating increased hydrophilicity as the CMC content rose. This could be due to the presence of hydrophobic amino acids in the keratin structure. It is generally held that keratin consisted of 60% hydrophobic and 40% hydrophilic amino acids [48]. A similar result was also reported by P. Hartrianti et.al [49] showing that higher amounts of keratin in alginate-keratin sponges reduced the water uptake capacities. Fig. 6 shows the weight-loss percentages of nanocomposites during 5 days soaking in PBS at 37 oC. All samples show an obvious increase in weight loss with time. It was seen that the maximum mass loss was the observed in K0C1 (59.36 ±3.5%), whereas in K1C2, K1C1, and K2C1 it was 41.43 ±3.1%, 32 ± 1.7%, and 20 ± 4.03%, respectively. Such a reduction could be due to the lowering of the water absorption of samples with the addition of keratin (Table 1). Reduction of water uptake impede the penetration of the water into the composites and consequently slow down the hydrolysis process. This finding was in good agreement with the previous study by Mandal et al. [50]. showing that the degradation rate of gelatin scaffolds reduced with the addition of silk fibroin. The results of WVRT for different dressings are summarized in Table 1. The WVRT is reported as a significant factor of the dressing and shows the ability of the dressing to maintain moisture at the wound site [51]. The high WVTR may cause wound dehydration and consequently lead to dermal necrosis occurrence. However, its low value leads to
the accumulation of wound exudates and consequent occurrence of infection. Therefore, a proper WVTR of dressing ranging between 500 to 2500 g/m2/day is needed to regulate the moist environment for obtaining the best healing [31]. As expected, the highest value is related to the K0C1 sample (3200 ± 196 g/m2/day). This could be due to the more hydrophilic nature of CMC than keratin and consequently its higher water uptake ability compared to the other samples. As expected, increasing the keratin content reduced the water uptake of the studied dressing, which reduced the WVRT to a suitable range for wound healing. It has been reported that an appropriate moist environment shortens the inflammatory phase, enhanced fibroblasts, and keratinocyte proliferation, collagen secretion, angiogenesis, and wound contraction [52]. A similar result was also reported previously by others that WVRT capacity has a direct relation with the hydrophilicity of materials [49, 53]. Protein adsorption is reported to be an important parameter in determining blood compatibility since it is an initial event happening when biomaterials contact with blood [54]. Table 1 represents the BSA adsorption of different dressings. As expected, protein adsorption was the highest in K2C1 (36.85 ± 2.3 mg/g), while such value decreased with increasing CMC. This could be more due to the hydrophobic nature of keratin than CMC. It has been reported that 40% of the keratin structure consists of hydrophilic amino acid and the rest (about 60%) are hydrophobic chemical complexes [48]. This is in good agreement with previous reports that hydrophobic surfaces adsorb more proteins than hydrophilic ones [55]. The blood clotting time (PT) of different nanocomposites is shown in Table 1. It can be seen that the clotting times of the K2C1 sample were almost similar to the human plasma, whereas the clotting time became longer as the CMC content rose. This suggests that the addition of keratin to CMC accelerates the blood clotting formation. The hemostatic properties of keratins derived from human hair have been widely reported previously [56]. Fig. 7 and Table 2 show the compressive mechanical properties of the fabricated composite dressings. As expected, the K0C1 sample shows the highest compressive modulus and strength (E= 2475 ± 54 KPa and =207 ± 8 KPa). The compressive strength decreased with the addition of keratin, however, such value was not significantly changed with increasing keratin amount (p>0.05). As shown in Fig. 7 and Table 2, the compressive moduli of the fabricated dressings were also reduced with increasing keratin content, starting at 1326 ± 74 KPa in K1C2, 814 ± 31 KPa in K1C1, and reaching 179 ± 21 KPa in K2C1. In spite of such reduction, the measured compressive mechanical properties are suitable for skin regeneration applications, since the human skin compressive modulus is less than 35 KPa [57].
3.3. In vitro clindamycin release Fig. 8 depicts the cumulative release profiles of clindamycin from different dressings as a function of time. It can be observed that the release profiles were started with burst release during first 4 h of soaking in PBS, which was 35.08 ±1.09% in K0C1, 28.5 ± 1.3% in K1C2, 22.01 ±1.9 % in K1C1, and 13.2 ± 2.03 % in K2C1. With increasing the incubation time, the release profile slowed down. The total amount of released clindamycin after 168 h (7 days) of soaking in PBS was the highest in K0C1 (91.5 ± 3.1%), however, this value decreased with addition of keratin to 76.15 ± 2.03 % in K1C2, 67.1 ± 1.9 % in K1C1, and 56.5 ± 4.03 % in K0C1. This could be related to the lowering of water uptake value and hydrophilicity with increasing the keratin. Thus, fewer amounts of water could penetrate inside the samples, which led to the release of fewer amounts of water-soluble clindamycin. This result is in line with previous studies proposed that the water uptake of the drug delivery system affects the released profile of watersoluble drugs [58]. Table 3 shows the kinetic analysis results of the clindamycin release from the fabricated nanocomposites. As revealed in this table, the R2 value is 1, 0.9908, 0.9803, and 0.9790 in K0C1, K1C2, K1C1, and K2C1 samples, respectively, which showed the best fit with Peppas model. Besides, the n values for all samples were less than 0.5, which were 0.376 in K0C1, 0.239 in K1C2, 0.236 in K1C1, and 0.277 in K2C1. Based on the n-value results, Fickian diffusion is the main mechanism for clindamycin release from the studied nanocomposites [59]. 3.4. Antibacterial activity The antimicrobial activity of different nanocomposites against Staphylococcus aureus (S. aureus; Gram-positive) bacteria is shown in Fig. 9. As expected, all compositions reduced the number of bacterial colonies on the LB agar plates in comparison with the control group. Interestingly, the K0C1 samples exhibited the highest antibacterial activity (99.66 ± 0.7%), while this value reduced to 97.66 ± 0.9 % in K1C2, 90.3 ± 1.03 % in K1C1, and 78.66 ± 3.4 % in K2C1. This could be due to the hydrophobic nature of keratin than CMC which, resulted in the release of less clindamycin as shown in Fig. 8. Overall, all samples show acceptable antibacterial activity against S. aureus. Previous studies also confirm that clindamycin has a bacteriostatic effect on S. aureus and consequently could effectively treat various skin tissue infections [60]. It has been reported that clindamycin prevents bacterial protein synthesis through binding with the large 50S bacterial ribosomal subunits and consequently disrupt bacterial cell membranes [60, 61]. 3.5. Fibroblast attachment and viability
Fig. 10 represents the attachment of 3 day-seeded fibroblasts on each dressing. It can be observed from the SEM images that fibroblasts were attached well on all samples. However, the cells are more rounded and prone to agglomerate in CMC compared to other samples. As observed from the SEM image the interactions between cells are stronger than a cell with a matrix in CMC samples, which caused poor cellular attachment to the surface and cell’s cluster formation. These results could be related to the highly hydrophilic surface of CMC, which is in good agreement with previous studies [9, 31]. As expected, greater cellular spreading was observed in dressings containing more keratin. Like fibronectin, keratin has leucine–aspartic acid–valine (LDV) cell-binding peptides in its structure that enhances cell-matrix interaction and cellular spreading [62]. The results of cell viability and proliferation of different nanocomposites during 5 days of culture are shown in Fig. 11. As observed from the MTT results, all samples showed the cellular viability more than 90%, suggesting that all sample were biocompatible. Moreover, the amounts of clindamycin release didn’t show any toxicity for fibroblasts. Similar to the SEM images, the cellular proliferation enhanced as the keratin content rose (p<0.01). This is consistent with the results of the study reported by Wang et. al [63], showing that keratin hydrogel significantly enhanced L929 mouse fibroblasts attachment and proliferation and might be an interesting template for skin regeneration applications. Conclusion The 3-D antibacterial wound dressing with a distinctly interconnected porous structure, which could mimic extracellular matrix structure was successfully fabricated from keratin, CMC and clindamycin loaded HNTs through freeze-drying. The effect of keratin addition on the morphology, physicochemical, drug release, antibacterial, and cellular response of the CMC hydrogel was studied. As revealed by SEM images, no obvious changes were observed on the pore sizes of the hydrogels. Higher keratin content reduced the WVRT of CMC from 3200±196 to 1921±92 g/m2/day, which is in the proper range for wound treatment. The addition of keratin enhanced the water stability of the CMC hydrogel and consequently improved its applicability. The in vitro release data indicated that the clindamycin release profile from the composite dressing was controlled via a Fickian diffusion mechanism and was slower in samples containing more keratin (K2C1). All samples showed obvious antibacterial activity against S. aureus, although a small number of colonies were still alive after 24 h in samples consisting of higher keratin. These CMC-keratin hydrogels were cytocompatible considering the in vitro cell viability response of more than 90% towards fibroblasts. Higher keratin content led to better cellular attachment, proliferation, and spreading. Taken together, our results indicated that the fabricated bioactive dressing has a great potential for speeding up the healing process in
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Figure captions:
Fig. 1. Photographs of the prepared dressings (a) K0C1, (b) K1C2, (c) K1C1, and (d) K2C1. Fig. 2: The morphology of HNTs Fig. 3. The FTIR spectra of HNTs before and after clindamycin loading. Fig. 4. The morphology and EDS mapping of Al and Si ions in different composites (a, b) K0C1, (c, d) K1C2, (e, f) K1C1, and (g, h) K2C1. Fig. 5. The FT-IR spectra of different nanocomposite dressings. o
Fig. 6. Weight-loss percentages of different samples during 5 days soaking in PBS at 37 C. Fig. 7. The compressive mechanical properties of the fabricated composite dressings. Fig. 8. Cumulative release profiles of clindamycin from different dressings as a function of time. Fig. 9. Changes in the number of colonies and bacterial reduction (%) cultured for 24 h against S. aureus bacteria. Fig. 10. The morphologies of fibroblasts cultured for 3 days on different samples (a) K0C1, (b) K1C2, (c) K1C1, and (d) K2C1. Fig. 11. MTT results of different composites over 5 days of culture (*p<0.05, **p<0.01, and ***p<0.001).
Tables Table 1. The code, composition, and properties of different dressings
Samples
Keratin
Mean
to
pore
CMC
diameter
weight
(m)
Porosity (%)
Water uptake
WVRT (g/m2/day)
Protein
Clotting
adsorption
Time
(mg/g)
(PT) *
(g/g)
(s)
ratio K0C1
0:1
98 5
68± 5
3.9 0.2
3200196
17.56 ±1.2
16 ± 0.1
K1C2
1:2
103 9
65± 7
2.8 0.09
2895105
21.53 ± 0.9
15.6 ± 0.1
K1C1
1:1
108 8
64± 4
2.10.1
2423183
27.67 ± 1.6
15 ± 0.2
K2C1
2:1
104 7
67± 4
1.50.05
192192
36.85 ± 2.3
13 ± 1
*The clotting time (PT) of human plasma: 12.5± 1
Table 2. The compressive mechanical properties of nanocomposites Samples K0C1 K1C2 K1C1 K2C1
Compressive Strength (KPa) 207 ± 8 171 ± 16 160 ± 14 166± 12
Compressive Modulus (KPa) 2475 ± 54 1326 ± 74 814 ± 31 179 ± 21
Table. 3: Clindamycin release kinetic parameters of different samples fitted with Korsmeyer- Peppas equation. Samples K0C1 K1C2 K1C1 K2C1
R2 1 0.9908 0.9803 0.9790
n 0.376 0.239 0.236 0.277
S0141-8130(19)34286-2 https://doi.org/10.1016/j.ijbiomac.2019.09.251 BIOMAC 13588
To appear in:
International Journal of Biological Macromolecules
Received Date: Revised Date: Accepted Date:
9 June 2019 17 September 2019 30 September 2019
Please cite this article as: S. Sadeghi, J. Nourmohammadi, A. Ghaee, N. Soleimani, Carboxymethyl cellulosehuman hair keratin hydrogel with controlled clindamycin release as antibacterial wound dressing, International Journal of Biological Macromolecules (2019), doi: https://doi.org/10.1016/j.ijbiomac.2019.09.251
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© 2019 Published by Elsevier B.V.
Carboxymethyl cellulose-human hair keratin hydrogel with controlled clindamycin release as antibacterial wound dressing Soodeh Sadeghi1, Jhamak Nourmohammadi1, *, Azadeh Ghaee1, Neda Soleimani2
1-Division of Biomedical Engineering, Department of Life Science Engineering, Faculty of New Sciences and Technologies, University of Tehran, Tehran, Iran 2- Departments of Microbiology and Microbial Biotechnology and Nanobiotechnology, Faculty of Life Sciences and Biotechnology, Shahid Beheshti University, Tehran, Iran
*Corresponding Author: Jhamak Nourmohammadi; PhD. Department of Life Science Engineering, University of Tehran, P.O. Box: 143951561, Tel: +98-21- 86093264 Email: [email protected]
Abstract This study offers a new antibacterial wound dressing from carboxymethyl cellulose (CMC)-human hair keratin with topical clindamycin delivery. Keratin was successfully extracted from human hair. Different sponges fabricated by changing CMC to keratin ratio were characterized and compared. Halloysite nanotubes were used as carriers to control the clindamycin release. Various characterization techniques were used to determine the effects of keratin addition on the structure, morphology, physical properties, drug release, antibacterial activity, and cellular behavior of CMC hydrogels. As proved by SEM and EDS, porous structure with interconnected pores was successfully formed and clindamycin-loaded HNTs were uniformly dispersed within the porous structures. Increasing the keratin in CMC hydrogel not only lowered its water vapor transmission rate to a suitable range for wound healing but also improved the water stability of CMC hydrogel. The in vitro release study indicated that clindamycin was released slower in samples containing higher keratin and the Fickian diffusion mechanism controlled their release profile. The fabricated dressing effectively inhibits S. aureus bacterial colonies growth after 24 h. Fibroblast culturing on the fabricated sponges indicated that cellular attachment, proliferation, and spreading were significantly enhanced with increasing the keratin amount.
Keywords: Carboxymethyl cellulose; Keratin; Clindamycin; Wound dressing; Fibroblast attachment.
Introduction The skin has a key role in protecting one’s body against external invasions [1]. Being the first and the largest layer in contact with the surrounding environment, the skin is more likely to be injured. Thus, using an effective wound dressing has been recognized as an effective approach toward accelerating the healing process. The suitable wound dressing should be able to provide an optimum condition around the wound, which best mimics the extracellular matrix (ECM) of skin to help to permit epithelial cell movements, passing oxygen properly, allowing a moist environment, draining wound exudates and considerably preventing the wound from being infected [2]. In this regard, carboxymethyl cellulose (CMC)-based hydrogels come to special consideration because of their abundance, transparency and low cost [3, 4]. The high hydrophilicity of CMC leads to the high absorption of wound exudates; it also provides a moist environment around the wound and prevents the tissue from losing water which is of great importance in burn and diabetic wounds [5-8]. Regardless of such improvements, the relatively poor cell adhesion of CMC hydrogels has been reported in previous studies. Moreover, the less antibacterial activity and water stability limit their practical applicability as a wound dressing [9, 10]. To make up for this shortcoming, the blending of CMC with other polymers has been reported previously [11, 12]. Choosing a nature-based protein beside CMC in fabricating the scaffolds can be a step closer to making a construction similar to the natural ECM. Keratin is a cysteine-rich intracellular cytoskeleton protein which exists in hair, epidermis, nails, feather, wool, hoof and horn [13]. In recent years human-derived keratin has gained much attention in the various biomedical engineering applications because of its low cost, excellent biocompatibility, availability, biodegradability, hemostatic, and less immune reactions. Moreover, it encourages a more powerful cell-matrix response because of the existing cell-binding motifs like glutamic acid-aspartic acid-serine (EDS), leucine-aspartic acid-valine (LDV) and arginine–glycine– aspartic acid (RGD) similar to ECM proteins such as collagen and fibronectin [14]. Yamauchi et al. [15] proposed that keratin-coated substrate has shown better L929 fibroblasts attachment and growth compared to the ones coated with collagen. Park et. al [16] have also reported that keratin extracted from human hair is more effective in healing wounds due to the almost complete regeneration of epithelial cells. Thus, it seems that the combination of keratin and CMC not only could resemble ECM composition but also could improve the cellular response of CMC. However, the aforementioned dressing is not effective in killing the bacteria at the wound sites. Infection is very probable in severe skin damages which can prolong the healing process [17]. Currently, multiple wound dressings have been studied for the localized delivery of antibiotics to the infected wounds in a controlled manner.
Halloysite nanotubes (HNTs) have been introduced as a potential substance for target delivery of various bioagents and drugs at a defect site [18]. HNTs (Al2Si2O5(OH)4.nH2O) are considered as biocompatible one-dimensional nanotubular aluminosilicate clays that have been widely studied in various biomedical applications [19]. Due to the high aspect ratio, low density, and hydrophilicity, HNTs can easily disperse within hydrophilic biopolymers without the need for surface pre-treatment [20-22]. Moreover, their high surface area provides a very good absorption property which allows drugs to be loaded easily into and on them[23]. In this regard, this study introduces and characterizes a novel antibacterial wound dressing from CMC, keratin containing clindamycin using HNTs as a drug carrier. Clindamycin is an efficient antibiotic for healing serious skin and soft tissue infections especially those caused by staphylococcus aureus (s. aureus) such as burn wound infection [24]. Different dressings were fabricated by changing the CMC to keratin ratio and citric acid was used as a crosslinker. The effect of keratin addition on the morphology, porosity, water uptake, water vapor transmission rate, and clindamycin release profile of the composites was studied and then compared to those of the pure CMC sample. In the end, the antibacterial activity, as well as the behavior of seeded fibroblast on different nanocomposites, were studied. 2. Materials and methods 2.1. Materials Human hair was gained from local Barber’s. Besides, Tris-HCL (PH 8.5, Merck, Germany), thiourea (Merck, Germany), Urea (Merck, Germany), 2-mercaptoethanol (Merck, Germany), absolute ethanol (Merck, Germany), methanol (Merck, Germany), chloroform (Merck, Germany), carboxymethyl cellulose (Sigma, USA), halloysite nanotubes (Sigma, USA), cellulose dialysis tube (12KDa, MWCO; Sigma, USA), and Clindamycin (sepia Co., Iran) were purchased. All used chemicals were of analytical grade. 2.2. Isolation of keratin from human hair Keratin was extracted from non-colored hair based on the Shindai method as described previously by Lee et al. [25]. In brief, chopped hair was washed with ethanol and then soaked in methanol/chloroform mixture (1:2 (v/v). After 24h soaking, the delipidated hair was washed thoroughly with deionized water 3 to 4 times and then dried. The dried hair was immersed in the Shindai solution (which is composed of 25 mM Tris-HCl, 2.6 M thiourea, 5 M urea, and 5% (v/v) 2-mercaptoethanol) and incubated at 50℃ for 72 h. Next, the mixture was passed through the filter paper and centrifuged 10000 rpm for 20 min to separate hair residues from the liquid. The obtained supernatant was dialyzed in
cellulose tubing against distilled water for 3 days. The concentration of the keratin in the obtained solution was about 8 wt.%, which obtained by weighing the keratin powder after lyophilizing corresponding to a known volume. The dialyzed solution was diluted and then kept at 4oC for further use. The extracted keratin was analyzed using Fourier transform infrared spectroscopy (FTIR, Perkin Elmer, Frontier) over the range of 400-4000 cm-1. 2.3. Loading of clindamycin with HNTs and characterization Clindamycin (2 g) was first dissolved in deionized water (40 mL) with the assistance of stirrer for about 20 min at room temperature. Then HNTs (2 g) was added to the drug solution and stirred for 10 min under vacuum. After that, the suspension was removed from the vacuum, stirred vigorously for 24 h and then centrifuged (7000 rpm) for 30 min. The supernatant was thrown away and the obtained precipitate was dried at 50 oC for 24 h. The loading efficiency of the drug was measured using UV-Vis spectroscopy at the wavelength of 210 nm from the following equation [26]. A calibration curve was plotted using different clindamycin concentrations ranging from 5 to 30 μg/mL. 𝐶𝑙𝑖𝑛𝑑𝑎𝑚𝑎𝑐𝑦𝑛 𝑙𝑜𝑎𝑑𝑖𝑛𝑔 𝑒𝑓𝑓𝑖𝑐𝑖𝑒𝑛𝑐𝑦 (%) =
𝐶0 𝑉 − 𝐶𝑠 𝑉 × 100 𝐶0 𝑉
C0 is the initial concentration of clindamycin, Cs is the concentration of clindamycin in the supernatant, and V is the volume of the solution. The morphology of HNTs was studied using SEM (Mira 3-XMU, Czech Republic)). Moreover, the structural changes of HNTs after clindamycin loading was assessed using Fourier transform infrared spectroscopy (FTIR; Bruker, Germany). 2.4. Preparation of clindamycin-loaded HNTs-CMC-keratin nanocomposites For pure CMCs sample, CMC powder (2 g) was added gradually to 100 mL deionized water and then stirred gently until completely dissolved. In the case of CMC-keratin composites, different weight ratios of keratin and CMC solutions (0:1, 1:2, 1:1, and 2:1) were mixed thoroughly while the Keratin–CMC concentration in the final mixture was kept constant (2% w/v). Besides, 5 wt.% (weight of polymers) citric acid as a cross-linker and 3 wt. % glycerol (weight of polymers) as a crosslinking extender was also added to each solution [27] . After mixing for 3h, clindamycin-loaded HNTs (2 wt. % of total polymers) were added to the different keratin-CMC solutions and stirred for about 15 min to disperse the nanoparticles evenly all over it, after that the mixture was poured into the 48-well tissue culture polystyrene plates. Subsequently, the plates were stored in -70oC overnight and then freeze-dried. Next, the samples were crosslinked at 70oC for 72 h. The code and the composition of the constructed scaffolds are outlined in Table 1. The fabricated dressings were coded as KC (0:1), KC (1:2), KC (1:1), and KC (2:1) based on the changing
keratin/CMC volume ratios (as noticed above). The Photographs of the fabricated dressings are shown in Fig.1. 2.5. Characterization of fabricated nanocomposites The morphology of the studied nanocomposites was assessed using SEM (Mira 3-XMU, Czech Republic). The mean pore size and their distributions were calculated by Image J software (National Institutes of Health, Bethesda, MD, USA). Moreover, the porosity percentages were calculated for each SEM image as follows: 𝑃𝑜𝑟𝑜𝑠𝑖𝑡𝑦 (%) =
𝐴𝑝 × 100 𝐴𝑇
Where, AP is the total area of pores in each cross-section, AT is the total area of the image in the same cross-section. To verify the distribution of HNTs in the scaffolds energy dispersive spectroscopy (EDS) elemental mapping of Al and Si was used to demonstrate the uniformity of HNT’s dispersion throughout the scaffolds. FTIR was performed to study the structural properties of the fabricated nanocomposites. Water absorption is another characteristic of biomaterials that should be considered especially when used as wound dressings. For obtaining the initial weight of samples (Wo), scaffolds were weighed before immersion in deionized water. After immersing and being incubated at 37°C for about 24 h, swelled scaffolds were weighed again (W s) while the excess water wiped off gently with a filter paper. The amount of equilibrium water uptake was obtained by the following equation [28]: Eqiulibrium water uptake (g/g) =
𝑊𝑆 − 𝑊0 𝑊0
The ability of the fabricated nanocomposites to control water loss was evaluated by measuring their water vapor transmission rate (WVTR) based on our previous study [29]. Briefly, the sample was fixed on top of a glass vial (A= 1.5 cm2) filled with deionized water. The assembly sealed completely using Teflon tape, weighed and then placed into an incubator (T= 32 ± 1 ºC) with a relative humidity of 37%. At each hour, the weight loss of the assembly was measured and plotted versus time. WVTR was calculated according to the following equation: WVTR (g/𝑚2 /day) =
Slope × 24 A
The weight loss of each sample was measured by soaking the sample with the initial weight (W 0) in phosphate buffer saline (PBS; pH = 7.4) at 37 ºC. At predetermined times, the samples were removed, completely dried and weighed (Wd). The weight loss (%) was measured as follows [30]: Weight loss(%) =
𝑊0 – 𝑊𝑑 × 100 𝑊𝑑
The Proteins adsorption on each sample was performed to demonstrate the blood compatibility of the fabricated nanocomposite dressings. Thus, each sample with known weight of 20 mg were inserted in 2 mL of Bovine Serum o
Albumin (BSA) solution in PBS (2 mg/mL, pH=7.4). After incubation at 37 C for 2h, the samples were removed from the solution and washed thoroughly with known volume of PBS. Bradford assay was used to measure the amounts of free protein in the remaining BSA solution and washing solution [31]. The BSA adsorption on each sample was calculated as follows [32, 33]: Protein adsorption (mg/g) =
C1 V1 − ( C2 V1 + C3 V2 ) weight
Where C1 is the initial concentration of BSA (here is 2 mg/mL), C2 is the concentration of BSA in the solution after 2h soaking of the nanocomposites, and C3 is the concentration of BSA in the washing solution. Moreover, V1 and V2 are the volumes of the BSA solution and washing solutions, respectively. The coagulant activity of the fabricated nanocomposites was investigated by the Prothrombin time (PT) coagulation assay [34]. Thus, 0.1 mL citrated normal human plasma was dropped onto the surface of the samples and kept at 37 oC for 3 min. Next, 0.2 mL PT reagent was added to each sample and the clotting time was measured. The compression tests of the cylindrical samples (d = 5 mm, h = 10 mm) were performed using Zwick/Roell Z050 -1
with a crosshead speed of 1 mm.min . The compressive strength and modulus were calculated from the stress-strain curve [35]. 2.6. In vitro release study of clindamycin from the nanocomposites The amount of clindamycin released from the composites was measured using UV-Vis spectroscopy (model,) at the wavelength of 210 nm[36]. Thus, each sample was soaked in 5 mL of phosphate buffer saline (PBS; pH=7.4) and kept in a shaker incubator (90 rpm; T= at 37 ºC). At each time point, 2 mL of PBS was removed and an equal volume of fresh PBS was added to each container. The amount of clindamycin released from the composites was measured using a clindamycin standard curve of different concentrations (5 - 30 μg/mL) The clindamycin release kinetics were also assessed with fitting the first 60 % of the release data with KorsmeyerPeppas model [37]: Mt = Kt n M∞ Mt is the amount of drug released at time t, M∞is the amount of released drug at an infinite time, K is the constant
value, and n is the release exponent that indicates the release mechanisms. 2.7. Antibacterial activity The antibacterial activity of each nanocomposite was evaluated based on disk diffusion and colony-forming units (CFU) assay Staphylococcus aureus (S. aureus; gram-positive; ATCC 12600) bacteria [38]. The sterilized sample was immersed in 500 µL of the bacterial solution containing 106CFU/mL and then placed into a shaker incubator (200 rpm) at 37oC. After 24 h incubation, the samples removed from the container and the remaining solution was diluted 10-fold with PBS. Afterward, 20 µL of the diluted solution was taken out and spread on the agar plate with a sterilized loop. The number of viable bacteria on the agar plate was counted after 24 h incubation at 37 oC (Ns) and then compared with the control (Nc) to calculate the bacteria reduction (%): 𝐵𝑎𝑐𝑡𝑒𝑟𝑖𝑎 𝑟𝑒𝑑𝑢𝑐𝑡𝑖𝑜𝑛 (%) =
[(𝑁𝐶 − 𝑁𝑆 )] × 100 𝑁𝐶
2.8. Fibroblast attachment and proliferation L929 fibroblast cells from National Cell Bank of Iran (NCBI; Pasteur institute) were cultured in Dulbecco’s Modification of Eagles Medium (DMEM; Gibco) containing 10 (% v/v) fetal bovine serum (FBS; Gibco) and (1% v/v) Pen-Strep in an incubator at 37 ºC under 5% CO2. The nanocomposites were sterilized using ethanol 70% and PBS. The attachment and morphology of seeded fibroblast cells on each sample were studied using SEM. Accordingly, 1 × 104 cells seeded on each sterilized nanocomposite and then incubated in a humidified incubator (T=37 oC, 5% CO2). After 3 days, fibroblasts were fixed with 4 (%v/v) glutaraldehyde solution followed by dehydration with graded ethanol and then coated with a thin gold layer before imaging. Viability and proliferation of fibroblast cells were evaluated using the MTT assay kit (Bioidea, Iran) [39]. Briefly, fibroblast cells (1 × 104) were first seeded on each sterilized sample and then incubated in an incubator at 37˚C and 5% CO2 for 3 and 5 days. After each incubation time, the culture medium was completely removed and replaced with ready-to-use RPMI 1640 in the kit. Next, 10 μL MTT stock solution (12 mM) was added to each well and the plate kept in the incubator for 3h. After that, 50 μL DMSO was added to each well, pipetting thoroughly, incubated in an incubator for 10 min to dissolve the created formazan crystals. After pipetting, the absorbance was read at 570 nm using an ELISA Reader (Stat Fax-2100, USA). The culture medium without the sample was chosen as a control group.
2.9. Statistical analysis All measurements were done three times for each composition and the results were reported as the mean ± standard deviation (SD). Student’s t-test was used to understand the difference between the studied composition and a p-value less than 0.05 defined as a significant difference. 3. Result and discussion 3.1. IR analysis of extracted keratin Fig. S1 (supplementary files) shows the FTIR spectrum of extracted hair keratin. As expected, the peaks at 3299 cm 1
, 1654 cm-1, 1534 cm−1, and 1244 cm-1 are related to the amide A, amide I, amide II, and amide III bands, respectively
[40]. The presence of these peaks indicated that keratin protein was successfully obtained from the hair. 3.1. Characterization of HNTs before and after clindamycin loading Fig. 2 shows the SEM image of the HNTs, which have shown tubular structures with a length of 380–1100 nm and diameters of 35–102 nm. The FTIR spectra of HNTs before and after clindamycin addition are shown in Fig. 3. As expected, HNTs shows typical peaks at 471 cm-1 (Si-O deformation of Si-O-Si), 550 cm-1 (Al-O deformation of AlO-Si), 751 cm-1 and 795 cm-1 (Si–O stretching), 910 cm-1 (OH deformation of Al-OH groups), 1015 cm-1 (Si-O stretching), 1640 cm-1 (OH deformation of water), 3623 cm-1 and 3695 cm-1 (OH stretching of inner and outer hydroxyl groups) [41]. After clindamycin addition, the clindamycin peaks overlapped with the peaks of HNTs. However, the new peaks at 1560 cm-1, 1690 cm-1, and 2965 cm-1 are related to the C=C, C=O, and C-H stretching vibration of clindamycin [42]. The presence of such peaks approved the successful loading of clindamycin in HNTs. 3.2. Characterization of nanocomposites Fig. 4 shows the cross-section views of different composites. Besides, the mean pore size and the porosity percentage of the scaffolds are represented in Table 1. As expected, all compositions have shown the open and interconnected pores. As summarized in Table 1, pure CMC shows the mean pore sizes of 98 ± 5 mm and porosity percentages of 68± 5%. As expected, there are no significant changes in the mean pore sizes and porosity percentages with increasing keratin content (p>0.05). To identify the presence and distribution of HNTs in each sample, an EDS elemental mapping of Al and Si was done. As shown in Fig. 4, both ions were well-dispersed throughout the pore walls, suggesting that clindamycin-loaded HNTs were present and uniformly distributed in the nanocomposite dressings. Fig. 5 depicts the FT-IR spectra of different nanocomposite dressings. In K0C1, a broad peak centered at 3402 cm -1 is attributed to OH vibration and the peaks at 2931 and 2871 cm-1 are related to C-H vibrations [30]. As expected, the
peaks at 1735 cm-1(carbonyl ester), 1602 cm-1, 1424 cm-1, and 1326 cm-1 (carboxylate), 1235 cm-1 (C-O of ester bonds), and 1120 cm−1 (C-O-C stretching) were observed. The coexistence of C=O and C-O ester bands with carboxylate (COO-) in CMC structure confirms that citric acid crosslinked the CMC structures [43-45]. Besides, the vibrations of Al-O and Si-O of HNTs were also observed at 1041 cm-1, 573 cm-1, and 491 cm-1 [46], respectively, suggesting that HNTs were successfully loaded in CMC structure. After keratin addition, a new shoulder appeared at 1655 cm-1, which is related to the C=O stretching of amide I bonds [40]. Interestingly, the N-H vibrations of amide II bonds (1542 cm1
) [40] were also detected in samples containing a higher amount of keratin (K2C1). The presence of these peaks
indicated the successful crosslinking of keratin and CMC with citric acid. This result is in good agreement with the previous study by H. Xu et.al [47] showing that new amide groups were formed by nucleophilic interaction between carboxyl and amine group when citric acid was used as a cross-linker. Table 1 shows the results of the equilibrium water uptake of each sample after being soaked in deionized water. As summarized in this table, K0C1 has the highest amount of water uptake (3.9± 0.2 g/g), with increasing the amount of keratin the water uptake equilibrium decreased to 2.8 ± 0.09 (g/g) in K1C2, 2.1 ± 0.1 (g/g) in K1C1, and 1.5 ± 0.05 (g/g) in K2C1. As expected, higher CMC weight ratios in a keratin-CMC sample enhanced the water uptake ability, indicating increased hydrophilicity as the CMC content rose. This could be due to the presence of hydrophobic amino acids in the keratin structure. It is generally held that keratin consisted of 60% hydrophobic and 40% hydrophilic amino acids [48]. A similar result was also reported by P. Hartrianti et.al [49] showing that higher amounts of keratin in alginate-keratin sponges reduced the water uptake capacities. Fig. 6 shows the weight-loss percentages of nanocomposites during 5 days soaking in PBS at 37 oC. All samples show an obvious increase in weight loss with time. It was seen that the maximum mass loss was the observed in K0C1 (59.36 ±3.5%), whereas in K1C2, K1C1, and K2C1 it was 41.43 ±3.1%, 32 ± 1.7%, and 20 ± 4.03%, respectively. Such a reduction could be due to the lowering of the water absorption of samples with the addition of keratin (Table 1). Reduction of water uptake impede the penetration of the water into the composites and consequently slow down the hydrolysis process. This finding was in good agreement with the previous study by Mandal et al. [50]. showing that the degradation rate of gelatin scaffolds reduced with the addition of silk fibroin. The results of WVRT for different dressings are summarized in Table 1. The WVRT is reported as a significant factor of the dressing and shows the ability of the dressing to maintain moisture at the wound site [51]. The high WVTR may cause wound dehydration and consequently lead to dermal necrosis occurrence. However, its low value leads to
the accumulation of wound exudates and consequent occurrence of infection. Therefore, a proper WVTR of dressing ranging between 500 to 2500 g/m2/day is needed to regulate the moist environment for obtaining the best healing [31]. As expected, the highest value is related to the K0C1 sample (3200 ± 196 g/m2/day). This could be due to the more hydrophilic nature of CMC than keratin and consequently its higher water uptake ability compared to the other samples. As expected, increasing the keratin content reduced the water uptake of the studied dressing, which reduced the WVRT to a suitable range for wound healing. It has been reported that an appropriate moist environment shortens the inflammatory phase, enhanced fibroblasts, and keratinocyte proliferation, collagen secretion, angiogenesis, and wound contraction [52]. A similar result was also reported previously by others that WVRT capacity has a direct relation with the hydrophilicity of materials [49, 53]. Protein adsorption is reported to be an important parameter in determining blood compatibility since it is an initial event happening when biomaterials contact with blood [54]. Table 1 represents the BSA adsorption of different dressings. As expected, protein adsorption was the highest in K2C1 (36.85 ± 2.3 mg/g), while such value decreased with increasing CMC. This could be more due to the hydrophobic nature of keratin than CMC. It has been reported that 40% of the keratin structure consists of hydrophilic amino acid and the rest (about 60%) are hydrophobic chemical complexes [48]. This is in good agreement with previous reports that hydrophobic surfaces adsorb more proteins than hydrophilic ones [55]. The blood clotting time (PT) of different nanocomposites is shown in Table 1. It can be seen that the clotting times of the K2C1 sample were almost similar to the human plasma, whereas the clotting time became longer as the CMC content rose. This suggests that the addition of keratin to CMC accelerates the blood clotting formation. The hemostatic properties of keratins derived from human hair have been widely reported previously [56]. Fig. 7 and Table 2 show the compressive mechanical properties of the fabricated composite dressings. As expected, the K0C1 sample shows the highest compressive modulus and strength (E= 2475 ± 54 KPa and =207 ± 8 KPa). The compressive strength decreased with the addition of keratin, however, such value was not significantly changed with increasing keratin amount (p>0.05). As shown in Fig. 7 and Table 2, the compressive moduli of the fabricated dressings were also reduced with increasing keratin content, starting at 1326 ± 74 KPa in K1C2, 814 ± 31 KPa in K1C1, and reaching 179 ± 21 KPa in K2C1. In spite of such reduction, the measured compressive mechanical properties are suitable for skin regeneration applications, since the human skin compressive modulus is less than 35 KPa [57].
3.3. In vitro clindamycin release Fig. 8 depicts the cumulative release profiles of clindamycin from different dressings as a function of time. It can be observed that the release profiles were started with burst release during first 4 h of soaking in PBS, which was 35.08 ±1.09% in K0C1, 28.5 ± 1.3% in K1C2, 22.01 ±1.9 % in K1C1, and 13.2 ± 2.03 % in K2C1. With increasing the incubation time, the release profile slowed down. The total amount of released clindamycin after 168 h (7 days) of soaking in PBS was the highest in K0C1 (91.5 ± 3.1%), however, this value decreased with addition of keratin to 76.15 ± 2.03 % in K1C2, 67.1 ± 1.9 % in K1C1, and 56.5 ± 4.03 % in K0C1. This could be related to the lowering of water uptake value and hydrophilicity with increasing the keratin. Thus, fewer amounts of water could penetrate inside the samples, which led to the release of fewer amounts of water-soluble clindamycin. This result is in line with previous studies proposed that the water uptake of the drug delivery system affects the released profile of watersoluble drugs [58]. Table 3 shows the kinetic analysis results of the clindamycin release from the fabricated nanocomposites. As revealed in this table, the R2 value is 1, 0.9908, 0.9803, and 0.9790 in K0C1, K1C2, K1C1, and K2C1 samples, respectively, which showed the best fit with Peppas model. Besides, the n values for all samples were less than 0.5, which were 0.376 in K0C1, 0.239 in K1C2, 0.236 in K1C1, and 0.277 in K2C1. Based on the n-value results, Fickian diffusion is the main mechanism for clindamycin release from the studied nanocomposites [59]. 3.4. Antibacterial activity The antimicrobial activity of different nanocomposites against Staphylococcus aureus (S. aureus; Gram-positive) bacteria is shown in Fig. 9. As expected, all compositions reduced the number of bacterial colonies on the LB agar plates in comparison with the control group. Interestingly, the K0C1 samples exhibited the highest antibacterial activity (99.66 ± 0.7%), while this value reduced to 97.66 ± 0.9 % in K1C2, 90.3 ± 1.03 % in K1C1, and 78.66 ± 3.4 % in K2C1. This could be due to the hydrophobic nature of keratin than CMC which, resulted in the release of less clindamycin as shown in Fig. 8. Overall, all samples show acceptable antibacterial activity against S. aureus. Previous studies also confirm that clindamycin has a bacteriostatic effect on S. aureus and consequently could effectively treat various skin tissue infections [60]. It has been reported that clindamycin prevents bacterial protein synthesis through binding with the large 50S bacterial ribosomal subunits and consequently disrupt bacterial cell membranes [60, 61]. 3.5. Fibroblast attachment and viability
Fig. 10 represents the attachment of 3 day-seeded fibroblasts on each dressing. It can be observed from the SEM images that fibroblasts were attached well on all samples. However, the cells are more rounded and prone to agglomerate in CMC compared to other samples. As observed from the SEM image the interactions between cells are stronger than a cell with a matrix in CMC samples, which caused poor cellular attachment to the surface and cell’s cluster formation. These results could be related to the highly hydrophilic surface of CMC, which is in good agreement with previous studies [9, 31]. As expected, greater cellular spreading was observed in dressings containing more keratin. Like fibronectin, keratin has leucine–aspartic acid–valine (LDV) cell-binding peptides in its structure that enhances cell-matrix interaction and cellular spreading [62]. The results of cell viability and proliferation of different nanocomposites during 5 days of culture are shown in Fig. 11. As observed from the MTT results, all samples showed the cellular viability more than 90%, suggesting that all sample were biocompatible. Moreover, the amounts of clindamycin release didn’t show any toxicity for fibroblasts. Similar to the SEM images, the cellular proliferation enhanced as the keratin content rose (p<0.01). This is consistent with the results of the study reported by Wang et. al [63], showing that keratin hydrogel significantly enhanced L929 mouse fibroblasts attachment and proliferation and might be an interesting template for skin regeneration applications. Conclusion The 3-D antibacterial wound dressing with a distinctly interconnected porous structure, which could mimic extracellular matrix structure was successfully fabricated from keratin, CMC and clindamycin loaded HNTs through freeze-drying. The effect of keratin addition on the morphology, physicochemical, drug release, antibacterial, and cellular response of the CMC hydrogel was studied. As revealed by SEM images, no obvious changes were observed on the pore sizes of the hydrogels. Higher keratin content reduced the WVRT of CMC from 3200±196 to 1921±92 g/m2/day, which is in the proper range for wound treatment. The addition of keratin enhanced the water stability of the CMC hydrogel and consequently improved its applicability. The in vitro release data indicated that the clindamycin release profile from the composite dressing was controlled via a Fickian diffusion mechanism and was slower in samples containing more keratin (K2C1). All samples showed obvious antibacterial activity against S. aureus, although a small number of colonies were still alive after 24 h in samples consisting of higher keratin. These CMC-keratin hydrogels were cytocompatible considering the in vitro cell viability response of more than 90% towards fibroblasts. Higher keratin content led to better cellular attachment, proliferation, and spreading. Taken together, our results indicated that the fabricated bioactive dressing has a great potential for speeding up the healing process in
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Figure captions:
Fig. 1. Photographs of the prepared dressings (a) K0C1, (b) K1C2, (c) K1C1, and (d) K2C1. Fig. 2: The morphology of HNTs Fig. 3. The FTIR spectra of HNTs before and after clindamycin loading. Fig. 4. The morphology and EDS mapping of Al and Si ions in different composites (a, b) K0C1, (c, d) K1C2, (e, f) K1C1, and (g, h) K2C1. Fig. 5. The FT-IR spectra of different nanocomposite dressings. o
Fig. 6. Weight-loss percentages of different samples during 5 days soaking in PBS at 37 C. Fig. 7. The compressive mechanical properties of the fabricated composite dressings. Fig. 8. Cumulative release profiles of clindamycin from different dressings as a function of time. Fig. 9. Changes in the number of colonies and bacterial reduction (%) cultured for 24 h against S. aureus bacteria. Fig. 10. The morphologies of fibroblasts cultured for 3 days on different samples (a) K0C1, (b) K1C2, (c) K1C1, and (d) K2C1. Fig. 11. MTT results of different composites over 5 days of culture (*p<0.05, **p<0.01, and ***p<0.001).
Tables Table 1. The code, composition, and properties of different dressings
Samples
Keratin
Mean
to
pore
CMC
diameter
weight
(m)
Porosity (%)
Water uptake
WVRT (g/m2/day)
Protein
Clotting
adsorption
Time
(mg/g)
(PT) *
(g/g)
(s)
ratio K0C1
0:1
98 5
68± 5
3.9 0.2
3200196
17.56 ±1.2
16 ± 0.1
K1C2
1:2
103 9
65± 7
2.8 0.09
2895105
21.53 ± 0.9
15.6 ± 0.1
K1C1
1:1
108 8
64± 4
2.10.1
2423183
27.67 ± 1.6
15 ± 0.2
K2C1
2:1
104 7
67± 4
1.50.05
192192
36.85 ± 2.3
13 ± 1
*The clotting time (PT) of human plasma: 12.5± 1
Table 2. The compressive mechanical properties of nanocomposites Samples K0C1 K1C2 K1C1 K2C1
Compressive Strength (KPa) 207 ± 8 171 ± 16 160 ± 14 166± 12
Compressive Modulus (KPa) 2475 ± 54 1326 ± 74 814 ± 31 179 ± 21
Table. 3: Clindamycin release kinetic parameters of different samples fitted with Korsmeyer- Peppas equation. Samples K0C1 K1C2 K1C1 K2C1
R2 1 0.9908 0.9803 0.9790
n 0.376 0.239 0.236 0.277