Evaluation of Partially Reduced Keratins Extracted from Wool Fibers as a Hydrogel Forming Biomaterial

Abstract

In this study, it was aimed to prepare low-cost hydrogel from reduced keratin. Keratin proteins were obtained from Merino wool via three extraction methods. In the first method, keratins were reduced using sodium sulfide. In the second method, keratins extracted with the first method were precipitated with HCl. Urea, EDTA, and sodium sulfide were used in the third method. Extraction yields of method 1, method 2, and method 3 were determined as 44 ± 2, 27 ± 1, and 42 ± 2 %, respectively. For all extraction methods, the average value of the free thiol amounts was obtained as 0.06 ± 0.02 mmol SH/g keratin. A considerable portion of the highly polydisperse keratins was separated between ~40 kDa and ~60 kDa in the SDS-PAGE gel, and this fraction corresponds to α-keratin proteins with low sulfur content. A strong band at ~1654 ± 1 cm^-1 detected in the FTIR spectra of the keratins confirms mainly α-helical secondary structure. The self-standing hydrogel was obtained upon incubating 15 wt. % keratin solution at 37^oC. Storage modulus and loss modulus of the hydrogel were determined as 1.3 ± 0.08 kPa and 0.1 ± 0.015 kPa, respectively. The keratin hydrogel is not cytotoxic to L929 mouse fibroblast cells, suggesting that this affordable hydrogel can be applied as a drug delivery/encapsulation system and in wound healing.

Keywords:
keratin; sulfitolysis; hydrogel; rheology; biocompatibility

HIGHLIGHTS

Keratin was extracted from wool fibers using a simple sulfitolysis method with an acceptable yield.

A self-standing hydrogel was obtained upon incubating the keratin solution in phosphate buffer saline (PBS) buffer at 37^oC without addition of any crosslinking agent.

The study of the interaction of the hydrogel and L929 cells indicates that this low-cost and sustainable keratin hydrogel is biocompatible.

INTRODUCTION

Sheep wool fibers are valuable raw materials in the textile industry due to their outstanding hygroscopicity, elasticity, warmth, affinity for dyestuffs, and softness [¹-⁴]. Raw wool varies in fiber diameter from one breed to another, and only a small portion of it, defined as fine grades, is suitable for the textile market [³]. The number of sheep worldwide is estimated to rise from 1.7 billion to 2.7 billion between 2000 and 2050 [¹]. Thus, large quantities of wool waste have been generated, necessitating to manage its proper disposal and efficient reuse for environmental concerns and sustainable development [², ⁴, ⁵].

Waste wool can be valorized as an insulating material in building construction, in the reinforcement of composites, and as an adsorbent to remedy wastewater [¹, ²]. Additionally, keratinous proteins extracted from wool waste can be utilized as value-added products such as nutrients, biomaterials, and ingredients in cosmetic formulations [², ⁶-⁸].

About 40% of raw wool comprises impurities such as lanolin, suint, vegetable matter, and small amount of insecticides, and clean wool fibers are obtained after scouring processes [⁹, ¹⁰]. Pure wool fibers can contain up to 95% keratin, a fibrous protein with remarkable tensile properties, toughness, and chemical resistance stabilized by disulfide crosslinks and polar and nonpolar forces within its structure [¹¹-¹⁴]. Keratins can be solubilized by various extraction methods, which are classified as physical, biological, and chemical methods, and these methods are well documented elsewhere [⁷, ¹⁵-¹⁷]. Physical extraction methods such as steam explosion and microwave treatment generally give rise to low molecular mass keratin proteins. Likewise, keratin hydrolysis/degradation is also possible by the biological methods with proteolytic enzymes such as keratinases, which can be produced by bacteria, actinomycetes, or fungi [¹⁵, ¹⁶]. On the other hand, most chemical keratin extraction methods predominantly cleave inter-molecular disulfide bonds rather than peptide bonds, preserving the intact protein chains. In the oxidative processes where peracetic acid is widely used, cysteine sulfur atoms are converted to sulfonic acid, and hence reformation of disulfide crosslinks is prevented. The resultant oxidized keratin is named keratose. Reduced keratins or kerateines, on the other hand, are obtained via the reduction of disulfide bonds generating free thiols, which can reform disulfide linkages [¹³, ¹⁸-²⁰]. A commonly used reducing agent, 2-mercaptoethanol (2-ME), gives a high yield without damaging the keratin structure, but its high cost and toxicity limit its industrial use. However, wool and chicken feather keratins extractions revealed that sodium sulfide can be considered a safe, low-cost, and effective reducing agent [²¹, ²²].

Once solubilized, the keratins can be processed into different forms, such as fibers, films, sponges, and hydrogels for end uses [²³-²⁶]. Inherent biocompatibility, biodegradability, and the presence of cell attachment peptide sequences such as RGD, LDV, LDS, and EDS make the keratins appealing biomaterials [¹⁴, ²⁷, ²⁸]. Especially self-assembling nature and the reactive functional groups of the keratins allow the preparation of keratin-based hydrogels easily. For example, wool keratin was solubilized using a guanidine and 2-ME solution. A porous hydrogel was readily prepared by dialysis via the aggregation of the keratin chains and reformation of the disulfide bonds. [²⁶]. A similar gelation mechanism was used to form hydrogels of wool keratins extracted using urea, thiourea, and sodium metabisulfite [²⁹]. In a different study, Nakata and coauthors (2015) prepared hydrogels from carboxymethylated, acetamidated, aminoethylated, and reduced wool keratin. The reaction between the free thiols of cysteines and iodoacetic acid, iodoacetamide, and bromoethylamine was used to obtain these modified keratins [³⁰]. Wool keratoses were also evaluated in the hydrogel formulations. Sando and coauthors (2010) obtained mechanically stable hydrogel via photocrosslinked wool keratoses [³¹]. Our research group exploited wool keratoses to form self-assembled and chemically crosslinked hydrogels with varying viscoelastic properties [²⁷, ³²].

In the current study, keratin proteins were extracted from Karacabey Merino wool using three different sulfitolysis methods based on sodium sulfide as a reducing agent. Characterization of these partially reduced keratins formed was performed. A self-standing hydrogel was obtained upon incubating the keratin solution in phosphate buffer saline (PBS) buffer at 37^oC. Viscoelastic, swelling, and morphological characteristics of the keratin hydrogel were investigated. The cytocompatibility of the hydrogel was confirmed using L929 mouse fibroblast cells.

MATERIAL AND METHODS

Materials

Karacabey (Turkey) Merino sheep wool fibers were used to extract keratins. The reagents employed in extracting the keratins from the wool samples were purchased from Sigma-Aldrich. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) analysis and cell culture experiments were performed with the previously described chemicals [³²].

Keratin extraction methods

The wool fibers were washed and defatted using previously reported procedures [²⁷, ³²]. Keratins were extracted using the following methods, Method 1, Method 2, and Method 3. After extraction, the keratin solutions were dialyzed against deionized water, and the dialyzed solution was freeze-dried to isolate keratin proteins.

Extraction of the keratin proteins with sodium sulfide (Method 1)

1 g defatted wool in 50 ml 125 mM sodium sulfide solution was shaken at 40^oC and 150 rpm for 4 h by avoiding exposure to the light. Extracted keratins were separated from the undissolved solid by using a ceramic filter.

Extraction of the keratin proteins with sodium sulfide and HCl (Method 2)

1 M HCl solution was added to the keratins extracted with sodium sulfide as described above until the pH of the solution was adjusted to ~4.0. Precipitated keratins were isolated by centrifugation and dissolved in deionized water at pH 8.0.

Extraction of the keratin proteins with sodium sulfide, urea, and EDTA (Method 3)

1 g defatted wool was treated with 50 ml solution containing 8 M urea, 3 mM EDTA, and 125 mM sodium sulfide at 40°C by shaking at 150 rpm for 4 h in the dark. Solubilized keratin proteins were filtered.

Characterization of the keratin proteins

The yield of keratin extraction methods was determined using the equation given below:

$Yield\%=\left(\frac{Amountofthe\ker atinproteinsextracted}{Amountofwoolusedfortheextraction}\right)\times 100\%$ (1)

Free thiol contents of the keratin proteins were determined via the DTNB (5,5-dithiobis(2-nitrobenzoic acid) assay method described by Aitken and Learmonth (1996) [³³]. Absorbance measurements were taken on Shimadzu UV-2450 model UV-Vis spectrophotometer. The concentration of free thiols was calculated using the extinction coefficient of 14150 M^-1 cm^-1 at 412 nm [³⁴].

X-ray diffraction (XRD) patterns of the samples were obtained with a Philips PANalytical X'Pert Pro model instrument using a CuK( source. The XRD data were recorded using 2θ values between 5° and 80° and a scan rate of 0.08°/s.

FTIR spectra of the extracted proteins were taken using a Shimadzu 8400 spectrophotometer using a resolution of 2 cm^-1. The KBr pellet technique was employed for sample preparation.

Preparation and characterization of the keratin hydrogels

Aqueous solutions of the keratin proteins (extracted using Method 1) were prepared at 15 wt. % concentration in deionized water or phosphate buffer saline (PBS) at pH 7.4. Gelation was induced by incubating the keratin solution at 37^oC overnight. The rheology of hydrogels was investigated using a HAAKE MARS model rheometer. A protocol similar to the one described by Chen et al. (2017) was followed [³⁵]. Scanning electron microscopy (SEM) was used to monitor the pore structure of the keratin hydrogel. Before the observation, the sample was frozen in liquid nitrogen, freeze-dried at -80°C, and coated with gold. SEM analysis was conducted on an FEI Quanta 250 FEG instrument. ImageJ software was used to determine the average pore size of the freeze-dried hydrogel [³⁶].

For the swelling experiments, freeze-dried hydrogel samples were placed in excess buffer, and the weight of the swollen hydrogel was measured at various time intervals. The swelling ratio (SR) was calculated using the equation below:

$SR=\frac{W_t-W_i}{W_i}$ (2)

where W_i represents weight the freeze-dried hydrogel measured initially, and W_t corresponds to swollen hydrogel weight measured at time t.

The cytocompatibility of the keratin hydrogel was assessed using a CCK-8 assay following the procedure reported previously [²⁷, ³²]. L929 mouse fibroblast cells were used. Cell viability was determined by measuring the absorbances at 450 nm wavelength, A₄₅₀. The measurements were performed using a Varioskan Flash model microplate reader. The following formula was used to calculate the relative proliferation rate:

$\mathrm{Re}lativecellproliferationrate=\frac{A_{450}ofthehydrogel}{A_{450}ofTCPSforDay1}$ (3)

Empty wells (TCPS) were used as the control samples. In the statistical analyses, p-values were determined with Minitab software.

RESULTS AND DISCUSSION

Extraction and characterization of the wool keratins

Wool keratins were extracted using the sulfitolysis and the modified sulfitolysis methods given in Table 1. A comparison of different keratin extraction methods indicates that sulfitolysis with sodium sulfites is quite an effective method with high yield and low cost [¹⁶]. Thus, the sulfitolysis procedure was performed using sodium sulfide only in the first method. The second method is involved in sulfitolysis, followed by HCl precipitation. In the third procedure, sulfitolysis was carried out using urea, EDTA, and sodium sulfide solution. The extraction yields and the free thiol amounts of the solubilized keratins are summarized in Table 1. Not surprisingly, the lowest yield was obtained in Method 2, as the keratins with an isoelectric point of ( 4 were isolated during the HCl precipitation. Although it was expected that using urea in the extraction would increase the yield by making the disulfide bonds in the intra-molecular structure of the keratin proteins accessible, the yields of Method 1 and Method 3 are similar. In water, sodium sulfide forms hydrosulfide and hydroxyl ions, resulting in an extraction solution with a high pH [²²]. High solution pH can also be considered a denaturing agent as it can break the hydrogen bonds and cleave the disulfide bonds [²², ³⁷]. However, the use of urea lowered the pH of the sodium sulfide solution from (11 to (8. Thus, in Method 1, the extraction solution's high pH facilitated the keratins' dissolution without necessitating an additional denaturing agent. Method 1 and Method 3's yields are significantly higher than that of a keratin obtained from a sheep hair, which employed a sodium sulfide reduction procedure with a lower liquor ratio [³⁸].

Table 1
The yields and the free thiol amounts obtained in different keratin extraction methods.

Method	Reagents	Yield - (%)	Free Thiol Amounts [mmol/g keratin]
1	Sodium sulfide	44 ± 2	0.07 ± 0.02
2	Sodium sulfide + HCl	27 ± 1	0.06 ± 0.01
3	Sodium sulfide + Urea + EDTA	42 ± 2	0.05 ± 0.01

The average value of the free thiol amounts was determined as 0.06 ± 0.02 mmol SH/g keratin for all extraction methods. The free thiol amount of keratin obtained from human hair extracted using a similar technique was < 0.01 mmol SH/g keratin. The low free thiol content is due to the reformation of disulfide bonds during the dialysis step [³⁹].

X-ray diffraction (XRD) patterns of the extracted keratin proteins are given in Figure 1 and were used to compare the crystallinity of these samples. The keratins obtained by different extraction methods exhibit a similar broad diffraction peak at 2Θ = 20°. The diffraction peaks at 2θ = 17.8° (5.1 Å) and 2θ = 19° (4.65 Å) indicate the presence of α-helix and β-sheet secondary structures, respectively. However, the overlapping of α-helix and β-sheet peaks hampers the assessment of the contribution of these secondary structures to the peak observed [⁴⁰, ⁴¹].

In the XRD spectrum of the parent defatted Karacabey Merino wool, both characteristic diffraction peaks at 2( values of 9^o and 20^o were observed. Therefore, the extraction methods employed in this study reduced the crystallinity of the keratin. Similarly, compared to the parent wools, the crystallinity of the wool keratin proteins regenerated from ionic liquids and deep eutectic solvents decreased [⁴⁰, ⁴²].

Figure 2 presents the FTIR spectra of the extracted samples and identifies the functional groups of these solubilized keratin proteins. Typical peptide bond vibrations corresponding to amide I, amide II, amide III, and amide A bands were observed for all samples [⁴³-⁴⁵]. A sharp band at (1654 ± 1 cm^-1 in the amide I region indicates that the keratins have a predominantly (-helical secondary structure [⁴⁶]. Wool keratins oxidized by peracetic acid (keratoses) were also reported to be rich in (-helical conformation [²⁰, ²⁷, ³²].

Figure 1
XRD patterns of the keratins extracted using (a) Method 1, (b) Method 2, and (c) Method 3.

Figure 2
FTIR spectra of the keratins extracted using (a) Method 1, (b) Method 2, and (c) Method 3.

Figure 3
SDS-PAGE results of the keratins; from left to right, lane 1 = Method 3, lane 2 = Method 1, lane 3 = Method 2, and lane 4 = protein molecular weight marker.

SDS-PAGE analysis results of the keratins are given in Figure 3. Molar mass distributions of the keratins are revealed by the diffusive fractions from (20 kDa to > 170 kDa and a distinct band at (10 kDa similar to those of the Merino wool keratoses [³¹, ³²]. The protein fractions between (40 kDa and (60 kDa correspond to α-keratins having low sulfur content. The bands resolved at higher molar masses indicate protein aggregates or proteins having uncleaved/reformed disulfide bonds. (-keratins with high sulfur content have molar masses between 11 and 28 kDa. Proteins rich in glycine and tyrosine are associated with the fractions having molar masses between 7.4 kDa and 12.3 kDa. [¹⁵] However, the molar mass of γ-keratin was reported to be 10.4 kDa for oxidized keratin extracted from Merino wool [³¹].

Characterization of the keratin hydrogel

The keratin hydrogel readily forms by incubating 15 wt. % of keratin solution in deionized water or PBS buffer at 37°C overnight (Figure 4). Since the Ellman test indicates a small but detectable amount of free thiols, the hydrogel formation is triggered by self-assembly and disulfide bond reformation.

Viscoelastic properties of the hydrogel were determined via oscillatory rheology measurements. The strain sweep and the frequency sweep data plots are shown in Figure 5a and Figure 5b, respectively. The strain sweep curve represents the linear viscoelastic region (LVR) where the storage modulus (G’) and the loss modulus (G”) are independent of strain. LVR of the keratin hydrogel spans up to (20% strain. The yielding (irreversible deformations) occurs above this critical strain value (the limiting value of LVR) [⁴⁷]. The self-assembled keratose hydrogel prepared at 37^oC exhibited a yield point above (1% strain [²⁷]. A much higher yield strain of the keratin hydrogel can be attributed to the higher flexibility of the keratin network held together by the disulfide linkages. The frequency sweep test was used to determine the elastic and viscous properties of the keratin hydrogel under a constant strain of 0.2%. Figure 5b indicates that the storage modulus (G’) value (1.3 ± 0.08 kPa) is higher than the loss modulus (G”) value (0.1 ± 0.015 kPa) confirming the rigid network structure of the keratin hydrogel. G’ values of the self-assembled keratose hydrogel and the tetrakis (hydroxymethyl) phosphonium chloride (THPC) crosslinked keratose hydrogel prepared by our group are 0.17 ± 0.03 kPa and 0.8 ± 0.05 kPa, respectively [²⁷, ³²]. The stronger gel structure of the keratin hydrogel can be explained by its higher biopolymer content and the contribution of the disulfide crosslinking to the network.

Figure 4
Picture of the keratin hydrogel.

Figure 5
(a) Strain sweep and, (b) frequency sweep curves of the keratin hydrogel.

The swelling kinetic data of the keratin hydrogel is given in Figure 6 and represents the water absorption property of the freeze-dried hydrogel. The inset graph indicates the initial stage of the swelling process. The freeze-dried hydrogel absorbed the PBS buffer very quickly (less than 15 min) and retained the buffer for 4 days until its disintegration. The average swelling ratio of the hydrogel in the PBS buffer was determined as 6.4 ± 0.4. Swelling ratios of the chemically crosslinked hydrogels prepared at 7.5% keratose concentration were obtained between ~20 and ~70 [³²]. The much lower swelling ratio obtained in this study can be attributed to a higher number of crosslinking points of the hydrogel network, which restricts the mobility of the keratin chains [⁴⁸].

SEM image of the keratin hydrogel is presented in Figure 7. The pore structure of the hydrogel is highly irregular, with a pore size distribution between (10 (m and (150 (m. The average pore size was measured as ( 35 ± 20 (m, which is suitable for the regeneration of mammalian skin [⁴⁹].

Figure 6
Change in the swelling ratio of the hydrogel as a function of time.

Figure 7
SEM picture of the freeze-dried hydrogel. Scale bar = 300 (m.

The cytocompatibility of the keratin hydrogel was tested and compared with the control sample (TCPS) using CCK-8 assay. The results are shown in Figure 8. At the end of Day 1, no significant difference (p-value = 0.77) was observed in the cell proliferation rate of the hydrogel and the control sample. After Day 4, The keratin hydrogel supports the proliferation of the cells but with a lower rate compared to the TCPS (p-value < 0.05 for Day 4 and Day 7), similar to the other keratinous hydrogels. [³², ⁵⁰] This result can be attributed to the high polymer concentration of the hydrogel limiting the space for cell proliferation. Between Day 4 and Day 7, cell proliferation on the hydrogel increased slightly, which can be due to the dissolution of the hydrogel as observed during the swelling measurements. Nevertheless, the hydrogel is cytocompatible, so it can be valuable in the encapsulation of drugs and wound healing.

Figure 8
Cell proliferation results of the keratin hydrogel.

CONCLUSION

In this study, we developed a low-cost hydrogel from reduced keratin extracted from Merino wool using different sulfitolysis methods. The simplest method (Method 1), where only Na₂S was used, offered an acceptable yield. In all three processes, partially reduced keratins were obtained due to the oxidation of the thiols during the dialysis process. These keratins exhibited predominantly (-helical secondary structure and similar molar mass distribution. Mechanically robust keratin hydrogel formed effortlessly under physiological conditions without the addition of external crosslinking agents. The keratin network formation is triggered by self-assembly and reformation of disulfide bonds, providing self-standing hydrogel. The study of the interaction of the hydrogel and L929 cells indicates that this cheap and sustainable keratin hydrogel is biocompatible and may have potential in drug delivery and wound healing applications. In conclusion, we showed the possibility of developing a value-added product, a keratinous hydrogel, from wool.

Acknowledgments

We acknowledge the Sheep Breeding Research Institute (Balıkesir, Turkey) for the wool samples and Prof. Muhsin Çiftçioğlu for the rheology experiments. We also thank the Integrated Research Centers at İzmir Institute of Technology for the other characterizations.

REFERENCES

• ¹ Allafi F, Hossain MS, Lalung J, Shaah M, Salehabadi A, Ahmad MI, et al. Advancements in applications of natural wool fiber. J Nat Fibers. 2022;19(2):497-512.
• ² Sun Y, Li B, Zhang Y, Dou H, Fan W, Wang S. The progress and prospect for sustainable development of waste wool resources. Text Res J. 2023;93(1-2):468-85.
• ³ Rajabinejad H, Bucişcanu II, Maier SS. Current approaches for raw wool wool waste management and unconventional valorization: A review. Environ Eng Manag J. 2019;18(7):1439-56.
• ⁴ Zhang C, Xia L, Zhang J, Liu X, Xu W. Utilization of waste wool fibers for fabrication of wool powders and keratin: A review. J Leather Sci Eng. 2020;2:1-15.
• ⁵ Su C, Gong JS, Qin J, He JM, Zhou ZC, Jiang M, et al. Glutathione enables full utilization of wool wastes for keratin production and wastewater decolorization. J Clean Prod. 2020;270:122092.
• ⁶ Giteru SG, Ramsey DH, Hou Y, Cong L, Mohan A, Bekhit AEDA. Wool keratin as a novel alternative protein: A comprehensive review of extraction, purification, nutrition, safety, and food applications. Compr Rev Food Sci Food Saf. 2023;22(1):643-87.
• ⁷ Chilakamarry CR, Mahmood S, Saffe SNBM, Arifin MAB, Gupta A, Sikkandar MY, et al. Extraction and application of keratin from natural resources: a review. 3 Biotech. 2021;11:1-12.
• ⁸ Sharma S, Gupta A. Sustainable management of keratin waste biomass: Applications and future perspectives. Braz Arch Biol Technol. 2016;59:e16150684.
• ⁹ Allafi FA, Hossain MS, Shaah M, Lalung J, Ab Kadir MO, Ahmad MI. A review on characterization of sheep wool impurities and existing techniques of cleaning: Industrial and environmental challenges. J Nat Fibers. 2022;19(14):8669-87.
• ¹⁰ Petek B, Marinšek Logar R. Management of waste sheep wool as valuable organic substrate in European Union countries. J Mater Cycles Waste Manag. 2021;23:44-54.
• ¹¹ Wang K, Li R, Ma JH, Jian YK, Che JN. Extracting keratin from wool by using L-cysteine. Green Chem. 2016;18(2):476-81.
• ¹² Cardamone JM, Nuñez A, Garcia RA, Aldema-Ramos M. Characterizing wool keratin. Adv Mater Sci Eng. 2009;2009(1):147175.
• ¹³ Cardamone JM. Investigating the microstructure of keratin extracted from wool: Peptide sequence (MALDI-TOF/TOF) and protein conformation (FTIR). J Mol Struct. 2010;969(1-3):97-105.
• ¹⁴ Ranjit E, Hamlet S, George R, Sharma A, Love RM. Biofunctional approaches of wool-based keratin for tissue engineering. J Sci Adv Mater Dev. 2022;7(1):100398.
• ¹⁵ Shavandi A, Silva TH, Bekhit AA, Bekhit AE-DA. Keratin: dissolution, extraction and biomedical application. Biomater Sci. 2017;5(9):1699-735.
• ¹⁶ Perța-Crișan S, Ursachi CȘ, Gavrilaș S, Oancea F, Munteanu FD. Closing the loop with keratin-rich fibrous materials. Polymers. 2021;13(11):1896.
• ¹⁷ Reddy CC, Khilji IA, Gupta A, Bhuyar P, Mahmood S, AL-Japairai KAS, et al. Valorization of keratin waste biomass and its potential applications. J Water Process Eng. 2021;40:101707.
• ¹⁸ Ham TR, Lee RT, Han S, Haque S, Vodovotz Y, Gu J, et al. Tunable keratin hydrogels for controlled erosion and growth factor delivery. Biomacromolecules. 2016;17(1):225-36.
• ¹⁹ Yang G, Yao Y, Wang X. Comparative study of kerateine and keratose based composite nanofibers for biomedical applications. Mater Sci Eng C. 2018;83:1-8.
• ²⁰ Shavandi A, Carne A, Bekhit AA, Bekhit AE-DA. An improved method for solubilisation of wool keratin using peracetic acid. J Environ Chem Eng. 2017;5(2):1977-84.
• ²¹ Brown EM, Pandya K, Taylor MM, Liu CK. Comparison of methods for extraction of keratin from waste wool. Agric Sci. 2016;7(10):670.
• ²² Poole AJ, Lyons RE, Church JS. Dissolving feather keratin using sodium sulfide for bio-polymer applications. J Polym Environ. 2011;19:995-1004.
• ²³ Zhu J, Ma N, Li S, Zhang L, Tong X, Shao Y, et al. Reinforced wool keratin fibers via dithiol chain re‐bonding. Adv Funct Mater. 2023;33(14):2213644.
• ²⁴ Ramirez DOS, Carletto RA, Tonetti C, Giachet FT, Varesano A, Vineis C. Wool keratin film plasticized by citric acid for food packaging. Food Packag Shelf Life. 2017;12:100-6.
• ²⁵ Tachibana A, Furuta Y, Takeshima H, Tanabe T, Yamauchi K. Fabrication of wool keratin sponge scaffolds for long-term cell cultivation. J Biotechnol. 2002;93(2):165-70.
• ²⁶ Ozaki Y, Takagi Y, Mori H, Hara M. Porous hydrogel of wool keratin prepared by a novel method: An extraction with guanidine/2-mercaptoethanol solution followed by a dialysis. Mater Sci Eng C. 2014;42:146-54.
• ²⁷ Pakkaner E, Yalçın D, Uysal B, Top A. Self-assembly behavior of the keratose proteins extracted from oxidized Ovis aries wool fibers. Int J Biol Macromol. 2019;125:1008-15.
• ²⁸ de Guzman RC, Merrill MR, Richter JR, Hamzi RI, Greengauz-Roberts OK, Van Dyke ME. Mechanical and biological properties of keratose biomaterials. Biomaterials. 2011;32(32):8205-17.
• ²⁹ Esparza Y, Bandara N, Ullah A, Wu J. Hydrogels from feather keratin show higher viscoelastic properties and cell proliferation than those from hair and wool keratins. Mater Sci Eng C. 2018;90:446-53.
• ³⁰ Nakata R, Osumi Y, Miyagawa S, Tachibana A, Tanabe T. Preparation of keratin and chemically modified keratin hydrogels and their evaluation as cell substrate with drug releasing ability. J Biosci Bioeng. 2015;120(1):111-6.
• ³¹ Sando L, Kim M, Colgrave ML, Ramshaw JA, Werkmeister JA, Elvin CM. Photochemical crosslinking of soluble wool keratins produces a mechanically stable biomaterial that supports cell adhesion and proliferation. J Biomed Mater Res A. 2010;95(3):901-11.
• ³² Yalçın D, Top A. Novel biopolymer-based hydrogels obtained through crosslinking of keratose proteins using tetrakis (hydroxymethyl) phosphonium chloride. Iran Polym J. 2022;31(9):1057-67.
• ³³ Aitken A, Learmonth M. Estimation of disulfide bonds using Ellman’s reagent. In: Walker JM, editor. The Protein Protocols Handbook. Totowa, NJ: Humana Press; 2009. p. 1053-5.
• ³⁴ Riddles PW, Blakeley RL, Zerner B. [8] Reassessment of Ellman's reagent. Methods in Enzymology. 91: Academic Press; 1983. p. 49-60.
• ³⁵ Chen MH, Wang LL, Chung JJ, Kim YH, Atluri P, Burdick JA. Methods to assess shear-thinning hydrogels for application as injectable biomaterials. ACS Biomater Sci Eng. 2017;3(12):3146-60.
• ³⁶ Schneider CA, Rasband WS, Eliceiri KW. NIH Image to ImageJ: 25 years of image analysis. Nat Methods. 2012;9(7):671-5.
• ³⁷ Schrooyen PM, Dijkstra PJ, Oberthür RC, Bantjes A, Feijen J. Partially carboxymethylated feather keratins. 1. Properties in aqueous systems. J Agric Food Chem. 2000;48(9):4326-34.
• ³⁸ Ramya KR, Thangam R, Madhan B. Comparative analysis of the chemical treatments used in keratin extraction from red sheep’s hair and the cell viability evaluations of this keratin for tissue engineering applications. Process Biochem. 2020;90:223-32.
• ³⁹ Yue K, Liu Y, Byambaa B, Singh V, Liu W, Li X, et al. Visible light crosslinkable human hair keratin hydrogels. Bioeng Transl Med. 2018;3(1):37-48.
• ⁴⁰ Nuutinen E-M, Willberg-Keyriläinen P, Virtanen T, Mija A, Kuutti L, Lantto R, et al. Green process to regenerate keratin from feathers with an aqueous deep eutectic solvent. RSC Adv. 2019;9(34):19720-8.
• ⁴¹ Jiang Z, Zhang N, Wang Q, Wang P, Yu Y, Zhou M, et al. Can thiourea dioxide regenerate keratin from waste wool? J Nat Fibers. 2022;19(13):5991-9.
• ⁴² Idris A, Vijayaraghavan R, Rana UA, Patti AF, Macfarlane DR. Dissolution and regeneration of wool keratin in ionic liquids. Green Chem. 2014;16(5):2857-64.
• ⁴³ Kong J, Yu S. Fourier transform infrared spectroscopic analysis of protein secondary structures. Acta Biochim Biophys Sin. 2007;39(8):549-59.
• ⁴⁴ Gazioglu Ruzgar D, Altun Kurtoglu S, Bhullar SK. A study on extraction and characterization of keratin films and nanofibers from waste wool fiber. J Nat Fibers. 2020;17(3):427-36.
• ⁴⁵ Çelik MG, Morcali MH, Ziba CA, Dolaz M. Valorization of chicken feather waste: fabrication of keratin‐chitosan biofilms. ChemistrySelect. 2021;6(9):2189-97.
• ⁴⁶ Jackson M, Mantsch HH. The use and misuse of FTIR spectroscopy in the determination of protein structure. Crit Rev Biochem Mol Biol. 1995;30(2):95-120.
• ⁴⁷ Dabbaghi M, Namjoshi S, Panchal B, Grice JE, Prakash S, Roberts MS, et al. Viscoelastic and deformation characteristics of structurally different commercial topical systems. Pharmaceutics. 2021;13(9):1351.
• ⁴⁸ Suvarnapathaki S, Nguyen MA, Wu X, Nukavarapu SP, Camci-Unal G. Synthesis and characterization of photocrosslinkable hydrogels from bovine skin gelatin. RSC Adv. 2019;9(23):13016-25.
• ⁴⁹ Annabi N, Nichol JW, Zhong X, Ji C, Koshy S, Khademhosseini A, et al. Controlling the porosity and microarchitecture of hydrogels for tissue engineering. Tissue Eng Part B Rev. 2010;16(4):371-83.
• ⁵⁰ Wang S, Taraballi F, Tan LP, Ng KW. Human keratin hydrogels support fibroblast attachment and proliferation in vitro. Cell Tissue Res. 2012;347(3):795-802.

• Funding:
  This research was funded by IZMIR INSTITUTE OF TECHNOLOGY RESEARCH FOUNDATION, grant number 2020IYTE0096.

Edited by

• Editor-in-Chief:
  Alexandre Rasi Aoki

• Associate Editor:
  Alexandre Rasi Aoki

Publication Dates

• Publication in this collection
  15 Nov 2024
• Date of issue
  2024

History

• Received
  08 Oct 2023
• Accepted
  16 Sept 2024