Extraction of keratin from waste chicken feathers using sodium sulfide and l -cysteine

Process Biochemistry 82 (2019) 205–214

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Process Biochemistry journal homepage: www.elsevier.com/locate/procbio

Extraction of keratin from waste chicken feathers using sodium sulfide and Lcysteine

T

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Firoozeh Pourjavaheria, Saeideh Ostovar Poura, , Oliver A.H. Jonesb, Peter M. Smookera, Robert Brkljačaa, Frank Sherkata, Ewan W. Blancha, Arun Guptac, Robert A. Shanksa a

School of Science, RMIT University, Melbourne, Victoria, 3001, Australia Australian Centre for Research on Separation Science (ACROSS), School of Science, RMIT University, Melbourne, Victoria 3001, Australia c Faculty of Chemical & Natural Resources Engineering, Universiti Malaysia Pahang, 25100, Kuantan, Pahang, Malaysia b

A R T I C LE I N FO

A B S T R A C T

Keywords: Feathers Green chemistry Protein Polymers

Keratin was extracted from different segments of disposable waste chicken feathers (CF) including the whole feathers, calamus/rachis (β-sheet) and barbs/barbules (α-helix), using sodium sulfide and L-cysteine. The yield of extracted keratin from sodium sulfide and L-cysteine was ˜88% and ˜66% respectively. The mass ratio of feathers to reducing agent was 1:20 and the reaction temperature was 40 °C for 6 h. Concentration of keratin extracted by each method was measured using the Bradford assay. The protein extracted from each feather section was characterised using sodium dodecyl sulfate-polyacrylamide gel electrophoresis, vibrational spectroscopy including FTIR and Raman, nuclear magnetic resonance, and thermogravimetry. These results confirmed the keratin structures after each extraction methods. The study showed that α-helix and ß-sheet based keratin could be extracted from CF using sodium sulfide and L-cysteine with high yields. This is the first report of CF keratin extraction using L-cysteine.

1. Introduction Keratin is a tough, fibrous protein and, being the main component of hair, feathers, nails, wool, hooves and horns of mammals, reptiles and birds, it is the third most abundant polymer in the environment after cellulose and chitin. It has unique biodegradability and biocompatibility properties and is non-toxic. It can be modified and developed in various forms such as gels, films, beads and nano/micro-particles [1]. As such it represents an important source of renewable and sustainable raw material for many applications [2]. Indeed, keratin has numerous applications in green chemistry, food science, the pharmaceutical, biomedical and cosmetic industries, and composite materials [1,3]. Millions of tons of feathers are generated annually worldwide as a by-product of the poultry industry. The amount of this waste is increasing concomitant with an increase in fowl meat production. This causes an environmentally difficult disposal problem leading to pollution and it can cause human health issues [4,5]. Therefore, from both economic and environmental viewpoints, it is desirable and important to develop effective and profitable processes to use these resources and transform waste feathers into new materials. Feather keratins are small proteins, uniform in size, with a molar mass reported to be ˜10–36 kDa [1,3,6–10]. The structure of keratin ⁎

confers insolubility, mechanical stability and resistance of feathers to common proteolytic enzymes and chemicals [11]. Keratins are stabilized by many intra- and intermolecular disulfide cross-links as well as other structural features. Its high strength and stiffness are due to the high proportion of cysteine residues in the polypeptide backbone, bonded by disulfide links [1]. These features cause insolubility in polar solvents like water, weak acids and bases, as well as in a polar solvents [12]. Keratin remains reactive however, because the cysteine units can be reduced, oxidized and hydrolysed [1,13,14]. The keratin structure is stabilised by a range of non-covalent interactions (electrostatic forces, hydrogen bonds, hydrophobic forces) and covalent interactions (disulfide bonds), which must be disrupted to facilitate dissolution of feathers [13]. Keratin is a complex biopolymer, composed of 19 amino acids linked together in ladder-like polypeptide chains by peptide bonds [15]. Its molecular chains consist of tight packing of alpha (α) helix or beta (β) sheet structures [3], which further stabilize its structure. Reduction processes are generally used for keratin extraction, due to their high efficiency [3]. The function of reducing agents is to decrease the stability of keratin fibres by disassociating disulfide bonds, hydrogen bonds, thus salt linkages, and allowing proteases access to the polypeptide backbone to dissolve the protein into solution [2,16].

Corresponding author. E-mail address: [email protected] (S. Ostovar Pour).

https://doi.org/10.1016/j.procbio.2019.04.010 Received 6 December 2018; Received in revised form 11 April 2019; Accepted 11 April 2019 Available online 12 April 2019 1359-5113/ © 2019 Elsevier Ltd. All rights reserved.

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electrophoresis from Bio-Rad Laboratories Pty., Ltd. (Sydney NSW, Australia). Phosphate buffer saline (PBS) tablets (0.14 M NaCl, 0.0027 M KCl, 0.01 < Phosphate buffer pH 7.4) and dithiothreitol (DTT) (5 g ultra-pure, CAS NO. 27565-41-9) were obtained from Astral Scientific Pty Ltd (Sydney NSW, Australia). The consumed water was distilled and all the chemicals utilised in the experiments were of analytical grade, used as received without further purification.

However, many of the reductive or oxidative agents used for reducing the disulfide bonds, such as thiols and peroxides, cannot be recycled, and they are harmful, often toxic and/or difficult to handle [13,17]. Physical and chemical keratin extraction methods require considerable energy investments [18]. Research has focussed on finding effiecient and eco-friendly processing methods to dissolve keratins [13]. From an environmental point of view, enzymatic partial hydrolysis is the most attractive method, due to relatively mild treatment conditions and the preservation of functional properties of the products [16]. This study will quantify that chicken feathers consist of 50% (w/w) fibre (barbs and barbules) and 50% (w/w) quill (calamus and rachis), in agreement with the literature [19–22]. The quill fraction is composed of more ß-sheets than α-helices while the feather fibre is the reverse [23]. Various attempts have been made to extract keratin from CFF, including hydrothermal [16,24], chemical methods (i.e. oxidative and reductive chemistry) [1,9,25–29], ionic liquids [13,30,31], physical methods [2,12] and enzymatic hydrolysis [2,3,16,32]. Few studies have reported the characterization of chicken feather fibre (CFF) α-helix keratin and ß-sheet keratin separately and/or compared them with the whole CFF keratin, which is the aim of this study. An objective is to find a practical and effective appropriate procedure to extract keratin from CFF with less harm to the environment [16]. A sodium sulfide method will be compared with the dissolution of CFF keratin in L-cysteine/urea solution, as simple and green chemical processing methods. The structures and properties of the regenerated keratin were characterized by SDS gel electrophoresis, LC–MS, Fourier transform infrared (FTIR) spectroscopy with attenuated total reflection FTIR (ATR-FTIR), Raman spectroscopy, solid and liquid state nuclear magnetic resonance (NMR), and thermogravimetry (TGA). Advances in the extraction, purification, and characterization of keratins will be developed that may lead to new derivatives and applications that can open new directions for valueadding to what is currently a waste material.

2.2. Chicken feather preparation The CFFs were purified from stains, oil, dirt, and pathogens according to the ‘ethanol-extraction purification’ method detailed in Pourjavaheri et al. [33–35]. All treated CFFs were dried in an incubator at 34 ± 1 °C for 3 days and conditioned at 20 ± 2 °C and 60 ± 2% relative humidity for 72 h [33]. Barbs and barbules (α-helix structure) of CFFs were stripped off the calamus and rachis (β-sheet structure) fibers. 2.3. Extraction of keratin Whole feather, calamus, rachis, barbs and barbules were ground from CFFs and added separately with ratios of 1:20 (to completely immerse the CFFs in solution) to 100 mL of aqueous solutions containing 0.5 mol/L sodium sulfide (Na2S) solution (solution A) and 8 mol/L urea (NH2CONH2) and 0.165 mol/L of L-cysteine (solution B) which was then adjusted to pH 10.5 using NaOH (2 mol/L) (solvation step). The A and B solutions were the optimal condition based on previous studies performed by Gupta et al. [36] and Xu and Yang [37], respectively. The solutions were then heated and kept at 40 °C while continuously stirring using a magnetic stirrer at 10 g for 6 h prior to centrifuging at 11,648g for 20 min at 10 °C. The supernatant was collected, and the contained particles were discarded. Hydrochloric acid (7 mol/L) was added to the solution until a pH of 4 [38,39] (the isoelectric-point (pI) of keratin) was obtained (precipitation step). The solution was then left without heating or stirring for 2 h. Given the solution comprised deionized (DI) water and keratin and precipitation occurred, it is likely that the precipitate was protein. This hypothesis was checked using the Biuret test (see supplementary information) [36]. The aqueous phase was tested for soluble protein after precipitate was removed and the Biuret test confirmed existence of protein in the solution. Precipitated keratin was washed three times with DI water, filtered and the surface water was removed using a clean napkin. The solid precipitated keratin particles were transferred to a 40 °C oven to obtain keratin powders, then stored in sealed light-sensitive glass containers, at 4 °C before analysed, characterised and compared. The process condition reported here has been optimized to avoid severe peptide bonds scission and obtain high-quality keratin with suitable molecular weight. Each experiment was carried out in triplicate and the reproducibility of each one is above 90%. It should be noted that sodium sulfide can be dangerous at levels as low as 100 ppm, as such, the safe handling procedures for sodium sulfide solution are included in the supplementary information to this manuscript.

2. Experimental 2.1. Material White chicken feathers (ca. 3 cm–20 cm in length) of freshly slaughtered adult Leghorn chickens were supplied by Baiada Poultry Pty Ltd (Melbourne VIC, Australia). Sodium sulfide (AR hydrated, Na2S·xH2O, CAS No. 1313-82-2); Copper (II) sulfate ≥99.0% (LR, CuSO4·5H2O, CAS No. 7758-99-8); Phosphoric acid (85%·w/w, H3PO4, CAS. NO. 7664-38-2 for phosphoric acid and 7732-18-5 for water); Methanol (AR, CH3OH, CAS NO. 67-56-1) and glycerol 99.5% (AR, C3H8O3) were obtained from Chem-Supply Pty Ltd (Adelaide SA, Australia). Hydrochloric acid 32%·w/w (AR, HCl, CAS No. 7647-01-0); Bromophenol blue (C19H10BR4O5S, CAS No. 115-39-9); Ammonium persulfate ≥98%, (H8N2O8S2, CAS No. 7727-54-0); 2-mercaptoethanol ≥99.0% (HSCH2CH2OH, CAS No. 60-24-2); Bovine serum albumin (BSA) (≥98%, CAS NO. 9048-46-8); and Coomassie brilliant blue G250 (pure, CAS. NO. 6104-58-1, C47H48N3NaO7S2) were obtained from Sigma-Aldrich (St Louis MO, United States). Sodium hydroxide 97–100.5% (LR, NaOH, CAS No. 1310-73-2), potassium hydroxide 85–100.5% (LR, KOH, CAS No. 1310-58-3) and urea 99.0% (LR, NH2CONH2, CAS No. 57-13-6) were obtained from Ajax Finchem Pty Ltd (Auckland, New Zeland). L-Cysteine 99% (BR, C3H7NO2S) was purchased from BDH Laboratory Supplies (Poole, Dorset, England). Dodecyl sulfate sodium (SDS) ≥95.0% (C12H25Na4S, CAS No. 15 1-2 13) was obtained from Merck-Millipore (Darmstadt, Germany). Tris (C4H11NO3, CAS No. 77-86-1) and Tris−HCl (C4H11NO3HCl, CAS No. 1185-53-1, were obtained from Astral Scientific (Gymea NSW, Australia). TEMED ≥ 99.0% (C6H16N2, CAS No. 110-18-9) was obtained from Invitrogen (Auckland, New Zealand). Bis-acrylamide mix ≥99.0% (C7H10O2N2, CAS No. 110-26-9) was obtained from Amresco (Solon OH, United States). Coomassie Brilliant Blue R-250 (C45H44N3NaO7S2, CAS No. 6104-59-2) was used as the resolving gel in

2.4. Yield calculation The keratin yield and weight loss were determined according to the following formulae: Keratin yield (%) = (Wfk / Wik) × 100 Keratin loss (%) = ((Wik − Wfk)/Wik) × 100 Total weight loss (%) = ((Wi − Wf)/Wi) × 100 where Wik and Wfk refer to keratin weight initially and after extraction, respectively, and Wi and Wf are the initial CFF weight (before keratin extraction) and final weight of the extracted keratin (after keratin 206

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spectra were obtained and compared with those of the purified CFFs of superior ‘ethanol-extraction purification treatment [33] from previous studies.

extraction), respectively [40]. 2.5. Characterisation of keratin

2.5.5. Vibrational spectroscopic analysis Raman spectra were measured using an excitation wavelength of 785 nm. Raman spectra were collected using a Raman Xplora Plus (Horiba LabRAM HR Evolution confocal micro-Raman system, Horiba Scientific, Lille, France) equipped with a confocal microscope and motorized stage. Spectra were collected through a 50× objective lens (Olympus, Melville, NY, USA) and a numerical aperture of 0.75. The laser power on the sample was approximately 25–50 mW. Each spectrum was a co-addition of 10 scans with illumination time of 2 s across 0–1800 cm−1 (2.6–4.9 cm−1 resolution). Magnification, aperture, and laser power were optimized between analyses to minimise spectral noise and to maximize spectral signal from samples with a range of energy tolerances.

2.5.1. Keratin concentration The concentration of the keratin solutions was analysed by UV–vis absorbance using a BioPhotometer plus (Eppendorf AG, Hamburg, Germany) spectrophotometer at 280 nm. The Keratin content was measured using the Bradford assay [41]. 2.5.2. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis The molecular mass of the extracted keratins was estimated using sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDSPAGE). Approximately 1 mg of the experimental keratins were dissolved separately in a solution containing 8 mol/L urea, 50 mM Tris, and 0.1 M β-mercaptoethanol, at pH 8.4. Keratin solution concentrations were measured using UV–vis and Bradford assay. Then ca. 1 μg was mixed with a 5x loading buffer (containing 10% (v/v) SDS, 250 mM Tris−HCl buffer (pH 6.8), 50% (v/v) glycerol, 0.5 M dithiothreitol (DTT), 0.02% (w/v) bromophenol blue) at a ratio of 4:1. The solutions were heated in a dry bath heat block at 95 °C for 5 min. The stacking and the separating gels were 4% and 15% polyacrylamide, respectively. Electrophoresis voltage used was 60 V for 30 min, followed by 180 V for 50 min. After electrophoresis, the gel was washed with water and stained with a staining solution (40% (v/v) methanol, 10% (v/v) acetic acid, and 0.05% (w/v) Coomassie brilliant blue R250). The specimens were de-stained with a de-staining solution (10% (v/v) ethanol and 10% (v/v) glacial acetic acid). The protein standard (Precision plus protein standards, unstained, BIO-RAD) was used for calibration.

2.5.6. Nuclear magnetic resonance spectroscopy (liquid state) Chicken feather keratin samples and human epidermis keratin were dissolved in d6-DMSO and in an 8 mol/L urea, 50 mM Tris, 0.1 M βmercaptoethanol and 0.1% sodium azide (pH 8) solution at a concentration of 100 mg keratin / 700 μL solvent. NMR spectra were referenced to solvent signals using DMSO (δH 2.50) or using a D2O capillary placed within the NMR tube (δH 4.64). The NMR spectra were acquired on a 500 MHz Agilent DD2 console (Santa Clara CA, USA). Spectra for the feather keratin samples dissolved in d6-DMSO acquired include proton (256 scans) and gCOSY (16 scans, 512 increments) experiments. Proton spectra (256 scans) for the human epidermis keratin and two feather samples were dissolved in an 8 mol/L urea, 50 mM Tris, 0.1 M β-mercaptoethanol and 0.1% sodium azide (pH 8) solution and were acquired using the PRESAT sequence (4-step purge) with suppression of both the water and urea signals. gCOSY (4 scans, 400 increments) and HSQCAD (16 scans, 512 increments) NMR spectra were additionally acquired for these samples with suppression using the PRESAT sequence.

2.5.3. Proteomic analysis and LC–MS/MS One microgram of keratin extracted from whole CFF under L-cysteine was processed by SDS-PAGE using a 15% gel and the Tris-glycine buffer system. The 10 kg/mol (kDa) protein band observed in the Coomassie-stained gel was excised and processed for proteomic analysis. The destained gel piece was reduced with 10 mM DTT in 50 mM triethylammonium bicarbonate (TEAB) (55 °C/45 min) and alkylated with 55 mM iodoacetamide made up in 50 mM TEAB (incubated at room temperature (in the dark/30 min). The keratin was then incubated with 200 μg/μL sequencing grade trypsin (Sigma Aldrich) (incubated at 37 °C / overnight). The digestion was stopped by the addition of formic acid (FA) to a final concentration of 1% and dried in a vacuum. The digested keratin was analysed by liquid chromatography/ mass spectroscopy (LC–MS/MS, Thermo Scientific) using a Q-Exactive mass spectrometer (Thermo Scientific).

2.5.7. Nuclear magnetic resonance spectroscopy (solid state) Chicken feather samples were separated into three groups; α- and βkeratin (whole feathers), α-keratin enriched (barbs/barbules only), and β-keratin enriched (calamus/rachis only). The samples were ground into a powder using a Rocklabs Ringmill Grinder (with a zirconia mill head) for 3 min. NMR spectra were externally referenced to adamantine (δC 29.2). NMR spectra were acquired on an Agilent DD2 500 MHz NMR spectrometer equipped with a 4 mm MAS solid-state triple resonance probe. Spectra for the feather keratin samples were acquired at a spin rate of 10 kHz, 4 (proton) or 4000 (carbon) scans, and a delay time of 5 s.

2.5.3.1. MS data analysis. Data were searched against the Chicken (Gallus gallus) UniProt database using MASCOT (Matrix Science Ltd., London, UK). The search was restricted using the following parameters: assuming trypsin enzyme with two missed cleavages, fixed modifications of carbamidomethyl©, variable modifications of Oxidation (M), a fragment ion mass tolerance of 0.20 g/mol, and peptide mass tolerance of 20 ppm. Only proteins with at least two peptides (filter by ion score ≥ 20) uniquely assigned to the respective sequence were considered identified.

2.5.8. Thermogravimetry analysis A PerkinElmer TGA (Pyris 1) Thermogravimetric Analyser (Melbourne VIC, Australia) was employed to evaluate thermal degradation, mass loss, and derivative as a function of temperature, remaining char ratio, and the changes in degradation behaviour associated with CFFs keratin. The mass loss curve was recorded between 30 °C and 750 °C under nitrogen purge (20 mL/min) and between 750 °C and 850 °C under oxygen purge (20 mL/min) at a heating rate of 20 K min−1. In order to minimize the effect of thermal lag, a small sample mass of ca. 2 mg was chosen.

2.5.4. Fourier-transform infrared spectroscopy The chemical structures of keratin powders were analysed by Fourier-transform infrared (FTIR) spectroscopy, using a PerkinElmer spectrum 100/Universal diamond attenuated total reflectance (ATR) (Beaconsfield, Buckinghamshire, England). The spectra were collected within the wavenumber range of 4000 cm−1 to 650 cm−1 with transmission mode and recorded with 32 scans and 4 cm−1 resolution. Spectra were produced using infrared radiation. Keratin wavelength

3. Results and discussion 3.1. Reduction of disulfide bonds Keratin is insoluble in water with low chemical reactivity but its solubility increases at high temperature and in the presence of reducing 207

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Fig. 1. Illustration of the mechanism by which disulfide bonds in chicken feather keratin can be reduced by sodium sulfide and L-cysteine.

additionally to the 91% keratin mentioned above, therefore, only 12% and 44% of keratin has been lost during the extraction process, which can be due to not precipitating totally or during changing containers or in the washing process. However, 88% yield (the maximum useful keratin extracted) under the sodium sulfide method is considered high and an acceptable yield. The yield under sodium sulfide agent gave 22% higher yield than the L-cysteine method. This is due to the fact that less keratin was precipitated by HCl due to the presence of urea.

agents [1]. On reduction, the disulfide (eSeSe) crosslinks in hydrophilic Solution A were broken into free thiol (eSH) besides protonation of some eNH2 and other groups in keratin making its surface positive and therefore solubilisation took place [1,42], by chemical reduction using sodium sulfide solution as a reducing agent. To break disulfide bonds of keratin in Solution B, a swelling agent, urea, was used to denature the compact crystal structure of keratin, the crystal became amorphous and the disulfide bonds were exposed to the L-cysteine, therefore the bonds were broken. Reduced keratin from both A and B Solutions were therefore dissolved in water or urea-water solution, respectively. After addition of HCl, keratins were precipitated out of solution. The salt produced using solution A and the urea in Solution B were washed out in the last stage of extraction (i.e. washing stage). These reactions are shown in Fig. 1.

3.3. Characterisation of the extracted chicken feather keratin 3.3.1. Keratin concentration The concentration of the keratins was estimated as shown in Table 2. The results of both protein concentration determined using absorbance of protein at 280 nm and Bradford assay methods were compared. The amount of keratin was estimated to be higher for the extracted methods using sodium sulfide than L-cysteine, due to the solubility of keratin, which was higher in sodium sulfide. This resulted in

3.2. Yield analysis The yield was calculated based on total available keratin in chicken feathers which is known to be ca. 91% [12,43,44] of total feather mass. The keratin yields were ca. 88% and 66% and the weight loss were determined as shown in Table 1. The 20% under sodium sulfide and 40% under L-cysteine weight loss from initial weight of keratin incurred by the extraction method was due to the fact that feathers contain 1% lipid and 7% water [12,43,44]

Table 2 Extracted keratin concentrations by sodium sulfide and L-cysteine as estimated using the Bradford assay and protein absorbance at 280 nm. Protein absorbance at 280 nm (mg/ mL)

Table 1 Analysis of keratin yield by sodium sulfide and L-cysteine extraction methods. Yield and loss (%)

Extracted with sodium sulfide

Extracted with L-cysteine

Keratin yield Keratin loss Total weight loss

88 12 20

66 44 40

208

Bradford assay (mg/ mL)

Sodium sulfide extraction method whole CFFs 1.08 β-sheet CFFs 3.78 α-helix CFFs 1.98

1.48 4.81 1.87

L-cysteine extraction method whole CFFs β-sheet CFFs α-helix CFFs

0.32 0.45 0.16

0.13 0.15 0.02

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Fig. 2. SDS-PAGE of keratin analysis. Columns a–d show the keratin standard and keratin extracted from α-helix, β-sheet and whole chicken feathers using Na2S solution respectively. Columns e–g show α-helix, β-sheet, and whole chicken feather keratin extracted using L-cysteine respectively.

more disulfide band formation therefore, more keratin has been extracted. The results for both extraction methods suggest that the extraction from β-sheet CFFs is more efficient than α-helix CFFs as their concentration is higher according to both assays. This fact proves that β-sheet CFFs reacted faster than α-helix CFFs. 3.3.2. Keratin molecular mass SDS-PAGE gel electrophoresis was used to estimate the average molecular mass values of the isolated protein homologs, the patterns, and the purity of the keratin extracted via Na2S and L-cysteine are shown in Fig. 2. The molecular mass of keratin was calculated by reference to standard markers [45]. The SDS-PAGE data revealed that our extracted keratins have a major, diffuse cluster of bands ca. 11 kg/mol which represented the molecular mass of keratins concentrated. The intensity of bands at 11 kg/mol is higher in sodium sulfide extraction methods for all three parts of CFFs and again suggests that more protein was extracted using this technique. Although, the molecular mass does not change during the different keratin extraction processes (i.e. via sodium sulfide or L-cysteine methods) as they were all ca. 11 kg/mol, the change in intensity of bands from L-cysteine extraction suggest that the chemical process is very different for each case. The molecular weight measurment using SDS-PAGE was in agreement with LC–MS/MS outcomes of 11 kg/mol. According to Gregg et al. [46], amino acid sequences, the predicted molecular weight for chicken feather keratin was 10.032 kg/mol, using ExPASy Compute pI/Mw tool, which is close to our results. The SDS-PAGE results (Fig. 2) show a large keratin band and little evidence of Coomassie stainable impurities. This implies that the protein extracted is relatively pure, however further testing would be needed to confirm this. This analysis demonstrated that keratin has been purified from chicken feathers to near homogeneity.

Fig. 3. FTIR spectra of feather sections; (a) purified feathers, (b) 6-months old keratin extracted from whole feathers via Na2S, (c) keratin extracted from whole feathers via Na2S, (d) keratin extracted from β-sheets parts via Na2S, (e) keratin extracted from α-helix parts via Na2S, (f) pure Na2S, (g) 6-month old keratin extracted from whole feathers via L-cysteine., (h) keratin extracted from whole feathers via L-cysteine, (i) keratin extracted from β-sheets parts via Lcysteine, (j) keratin extracted from α-helix parts via L-cysteine and k) pure Lcysteine.

previously [33]. Comparison of the spectra of the four keratin samples extracted via solution A (Na2S as shown in Fig. 3(b–e) shows that their characteristic peaks were similar to the spectrum of purified CFF that contains 91% keratin [12,43,44]. On the other hand, the processing methods have some effect on the chemical structure of protein as revealed by the spectra of three keratin samples extracted via solution B (L-cysteine as shown in Fig. 3(g–i)), since shows different features to the keratin extracted from solution A. Therefore, we can hypothesise that the keratin sample itself is in different conformation/form based upon the extraction method. The spectra of both extracted solutions A and B are presented in Fig. 3(f and j). As expected, the characteristic absorption bands are mainly assigned to the peptide bonds (eCONH) and the vibrations in the peptide bonds originate bands known as amides A and I–III [13,47,48]. The broad absorption band region from 3500 cm−1 to 3200 cm−1 (amide A) indicates α-helix structure [3,49–51], and the sharper band at 3281 cm−1 is attributed to the stretching vibrations of OeH and NeH (amide A). Bands that fall in the 3000 cm−1 to 2800 cm−1 range are related to CeH stretching modes [16]. The band at 2923 cm−1 is attributed to the symmetrical CH3 stretching vibration [52]. The split peaks from 1700–1600 cm−1 [12,53,54] (Amide I region) is connected mainly with C]O stretching vibrations and originates from a combination of α-helix and β-sheet motifs [3,49,50], while the strong band at 1630 cm−1 is

3.3.3. LC–MS/MS In the ca. 11 kg/mol gel piece, there were three proteins, namely type II keratin, which was the most abundant, histone and chicken hemoglobin subunit alpha A-protein. The molecular mass of extracted keratin in LC–MS/MS technique confirmed that peptides match the SDS-PAGE result. 3.3.4. Infrared spectroscopy In order to examine the effects of different extraction processes on CFK, FTIR spectra of the extracted keratins produced from whole, βsheet and α-helix parts of CFFs, using both A and B solutions were obtained in the range of 4000 cm−1 to 600 cm−1 and compared with the spectra of the purified CFF as shown in Fig. 3. Typical infrared spectra of purified CFFs (Fig. 3(a)) have been described in detail 209

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Fig. 4. FTIR spectra of fresh and 6-month-old keratin extracted from whole feathers via Na2S.

related to C]O stretching [48]. The Amide II region, originating from NeH bending and CeH stretching [13,48], presents bands at 1580 cm−1 and 1480 cm−1 related to β-sheet structure [3,49,50]. A weak band in the range of 1300−1220 cm−1 in the Amide III region, which is due to the combination of CeN stretching and NeH in-plane bending motions [55,56] as well as some contribution from CeC stretching and CeO bending [15,57]. It can be seen from Fig. 3 that these bands exist in all extracted keratins. The Amide I–III bands provide critical information on protein conformation and backbone structure [3]. The NeH out-of-plane bending vibration typically occurs in the range between 750 cm−1 and 600 cm−1 [51]. There is a significant change in the FTIR spectra from the fresh to the 6 months aged keratin extracted from whole CFF via Na2S as shown in Fig. 4. This observation is probably explained by the fact that some of the constituent amino acid side chains are more prone to oxidation over time via this extraction method. There is a possibility of conformational change from more crystalline to less crystalline forms of extracted keratin via sodium sulfide solution. The FTIR spectra were used to evaluate changes in α-helix and βsheet structure resulting from the urea treatment and showed clearly distinguishable peaks in the region of 1550–1750 cm−1 (Fig. 3(g–i)), particularly those assigned to helical (1650 cm−1 and 1656 cm−1) and β-sheet (1632 cm−1, 1641 cm−1, and 1695 cm−1) structures, corresponding to the model by Fraser et al. [8]. The aging effect on keratin extraction via L-cysteine is less obvious as the bands for the 6 months old sample get only sharper and no shift in position is observed. This could be due to the degree of crystallinity of keratin has being changed over time, but no oxidation has taken place.

Fig. 5. Raman spectra of feather sections; (a) β-sheet keratin, (b) α-helix keratin and (c) 6-month old keratin extracted from whole feathers via Na2S, (d) keratin extracted from whole feathers via Na2S, (e) keratin extracted from βsheets via Na2S, (f) keratin extracted from α-helix via Na2S, (g) pure Na2S, (h) 6-month old keratin extracted from whole feathers via L-cysteine, (i) keratin extracted from whole feathers via L-cysteine, (j) keratin extracted from β-sheets via L-cysteine, (k) keratin extracted from α-helix via L-cysteine and l) pure Lcysteine.

1248 cm−1 (Fig. 5(b)), which is in agreement with Rizzo et al [61]. The strong amide III band at 1241 cm−1 is indicative of βsheet [60]. Raman spectroscopy is able to monitor disulfide linkages in keratin [25,52,58]. The SeS bonds appear as weak peaks in the 500–550 cm−1 range of the spectrum. Additionally, the disulfide stretching [58] vibration occurs from 510 to 526 cm−1, and can be observed in Fig. 5(a) and (b). Fig. 5(c) and (d) represent fresh and 6 months old keratin sample that has been extracted by Na2S. The aging of keratin has resulted in previously observed bands at 1080 and 1514 cm−1 for fresh keratin disappearing. The band at 542 cm−1 has shifted to 562 cm−1 which indicates that the local environments of sulfur groups changes over time. However, the extracted keratin from L-cysteine did not show any significant changes as a result of aging as is shown in Fig. 5(h) and (i). Therefore, the structure of keratin did not alter over time and was more strongly preserved by L-cysteine method. Since sodium sulfide is a stronger reducing agents than L-cysteine, there is a possibility of air oxidation of molecular sites that had been reduced by the redox reagent via this extraction method. Raman spectra from both Na2S (Fig. 5(e) and (f)) and L-cysteine (Fig. (j)) extracted keratins showed strong peaks at 563 cm−1 and 588 cm−1 whereas the rest of the spectra showed no significant differences. These peaks disappear for the keratin extracted from αhelix segments of chicken feathers (Fig. 5(k)) via L-cysteine and leads to an identical spectrum as for Solution B (Fig. 5(l)), which demonstrated the existence of urea residues in the keratin (the strong peak at 1088 cm−1). These results from both Raman and Infrared spectra indicate that keratin extracted by the L-cysteine method was not further affected by aging, which was not the case when using the Na2S method.

3.3.5. Raman spectroscopic analysis Our Raman spectra support the chemical disparity between the keratins extracted via Na2S and L-cysteine using whole, β-sheet and αhelix parts of CFFs and when compared with 6-month aged keratins (Fig. 5). These Raman spectra were recorded from rachis and calamus (β-sheet), barbs and barbules (αhelix) of chicken feathers as shown in Fig. 5(a) and (b), respectively. It was found that the spectra obtained from the β-sheet enriched quill fraction showed intense peaks at 1004, 1242, 1461, 1665 cm−1; which can be observed to some degree for αhelix, however, the intensity of these latter peaks varies as expected for a natural material. The typical spectra bands obtained from the βsheet component was in agreement with that obtained from the quill by Akhtar and Edwards [58] and from rachis and calamus by Church et al [59]. The strong peak at 1665 cm−1 in the Amide I region indicates a dominant β-sheet structure [59,60]. The α-helix part of CFFs shows weak bands at 1605–1610 cm−1 and 1580 cm−1 (Fig. 5(a)), and possibly arising from C]C groups in the aromatic rings of tyrosine and phenylalanine, which comprise 1.4 and 3.1%·w/w of feather keratin, respectively [58]. Another α-helix structure-related band appears at 210

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Fig. 6. 1H SSNMR spectra of chicken feather keratin extracted from (a) α-helix, (b) β-sheet, (c) whole feathers.

SSNMR could potentially be used as an analytical tool to distinguish between α-helix and β-sheet structures of keratin in different samples. As the quill and feather fibre fractions are not pure α-helix or β-sheet keratin, but rather are mixtures of both, it is difficult to characterise the α-helix or β-sheet keratin independently. The 1H SSNMR results suggest that the hydrogen environments in pure α-helix keratin are predominantly observed at ˜−2.4 ppm whilst the hydrogen environments in pure β-sheet keratin are predominantly observed at ˜0.6 ppm. This indicates that the connectivity of protons in solid α-helix keratin are of one abundant type of environment, which differs from connectivity/ environments observed in solid β-sheet keratin.

3.3.6. Solid-state NMR studies The unextracted chicken feathers were analyzed with solid-state NMR in an attempt to further characterize α-helix and β-sheet keratin and to identify potential markers to distinguish between the two structures of keratin. 13C SSNMR results revealed characteristic NMR signals for amide carbonyl carbons of keratin protein (˜170 ppm), aromatic group-containing amino acids in keratin (˜129 ppm), along with leucine residues (˜55 ppm, ˜40 ppm, and ˜24 ppm) [2]. Comparison of the samples showed no significant difference in the 13C SSNMR spectra between the α- and β-keratin enriched samples. Fig. 6 shows the 1H SSNMR spectra for the feather samples. Whilst the 13C SSNMR was unable to distinguish between the different samples, it appears that 1H SSNMR was able to reveal subtle differences between α-helix and β-sheet. Fig. 6(a) shows the 1H SSNMR spectrum for α-helix enriched feather fiber. Two main signals were observed at ˜0.6 and ˜−2.4 ppm which appear as broad signals, however, the signal at ˜−2.4 ppm appears slightly sharper and with greater intensity. The β-sheet enriched quill fraction (Fig. 6(b)) displays these two signals, however, the signal at ˜0.6 ppm appeared with higher intensity than the signal at ˜−2.4 ppm. The sample containing both α- and β-keratin (Fig. 6 (c)) shows the presence of both signals in a more equal proportion to one another, as would be expected for the whole chicken feather. This slight difference in the 1H SSNMR spectra suggests that 1H

3.3.6.1. Liquid state NMR studies. The extracted keratin from both extraction methods (Na2S and L-cysteine) was compared to an authentic human epidermis keratin sample. The authentic human epidermis keratin sample has been used as a standard to identify and compare diagnostic keratin NMR signals. The proton spectra of the extracted samples, along with the authentic human epidermis keratin, showed a series of broad resonances in the downfield region (6–8 ppm), confirming that both extraction methods retain the molecular structure of the keratin after extraction. Based on the proton NMR data, no difference in characteristic signals could be observed between α-helix and β-sheet enriched samples. The biggest difference between the 211

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Fig. 7. a) TGA, and b) DTG thermograms (mass loss) of L-cysteine, urea and purified CFFs compared with keratin extracted from whole CFF, β-sheet and α-helix parts via Na2S and L-cysteine.

signals between α-helix and β-sheet for the extracted chicken feather keratin samples. Due to poor signal-to-noise, HSQCAD spectra could not be acquired for the extracted keratin samples. These results show that while keratin can be observed in the samples extracted from chicken feathers, it is not possible to use solution state NMR to distinguish between α- and β-keratin. The proton environments do not differ substantially which makes it difficult to find a potential marker. It may be possible that differences in the carbon chemical shifts occur between α-helix and β-sheet keratin, however, it was not possible to get enough of the extracted keratin into the solution to obtain HSQCAD NMR data. If in future work enough keratin can be dissolved, it may be possible to re-acquire HSQCAD NMR data and determine if there are any differences in the carbon chemical shifts of αand β-keratin. The solvent effect from dissolving CFF could itself significantly disrupt the molecular order of an alpha-helix and/or a betasheet. Further, since the solubility of α-helix and β-sheet keratin in DMSO need not be equal, their ratio in the precipitate at the bottom of the NMR tube of CFF versus in solution need not be equal either.

Table 3 Thermal data of L-cysteine, urea, purified CFFs and keratin extracted from whole CFF, and the β-sheet and α-helix sections via sodium sulfide and L-cysteine. Materials

L-cysteine

Urea Purified CFFs CFK, whole, Na2S CFK, β-sheet, Na2S CFK, α-helix, Na2S CFK, whole, L-cysteine CFK, β-sheet, Lcysteine CFK, α-helix, Lcysteine

Temp. at 5 % Mass Loss (°C)

Mass Loss (%) at 300 °C

Char Levels (%) at 500 °C

231 184 194 165 195 171 236 231

15 24 21 22 23 25 82 78

96 99 17 21 23 22 10 9

188

79

10

extracted samples to the authentic human epidermis sample was the signal-to-noise. It appears that the keratin extracted from the feathers was not as soluble as the human sample, and the NMR tubes contained precipitate at the bottom indicating that the samples were already saturated. gCOSY and HSQCAD NMR spectra of the authentic human epidermis keratin revealed that the downfield proton signals of keratin correlate to neighboring hydrogens and were linked to distinct carbon signals. The gCOSY spectra showed no difference or characteristic

3.3.7. Thermal behaviour The thermogravimetry of the purified CFF segments and extracted keratins are presented in Fig. 7 and Table 3. A three-step decomposition process was observed in all keratin cases including purified chicken feathers. The first was a minor mass loss over the range of ˜30 °C–˜230 °C. The weight loss of purified chicken fibers and keratin extracted via both sodium sulfide and L-cysteine were approximately 212

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purposes such as anti-aging cream, shampoo, conditioner for medical and biomedical purposes such as bone replacement, bone graft and tissue engineering.

1–2% that could be attributed to the evaporation of incorporated water near 100 °C. Presence of ˜2% moisture substantiates that the purity of the keratin sample could be greater than 98% based on the composition of nitrogen and water [62], which was in agreement with the SDS-PAGE outcomes. The second mass loss step was much larger, around 70%, and occurs from ˜230 °C to ˜280 °C, for keratin extracted via L-cysteine, which corresponds to the decomposition of the urea (considering the urea curve in Fig. 7(a), rather than Lcysteine with respect to the small amount of L-cysteine used in extraction process). This observation indicates the presence of urea as a consequence of the reaction of peptides with amide groups. The TGA curves for keratin extracted via sodium sulfide demonstrated similar decomposition as the pure chicken feather curve, in the temperature range of ˜230 °C to ˜400 °C, which was mainly caused by the degradation of the helix structure of keratin materials [7,50,57]. The third mass loss stage was from ˜400 °C to 750 °C, which was 85% for keratin extracted via sodium sulfide and 96% for keratin extracted via L-cysteine. Derivative thermogravimetry (DTG) curves were calculated, which showed the temperature at the minimum corresponds to the maximum weight loss at that particular temperature [7]. This shows that the temperature difference between keratin extracted via L-cysteine versus sodium sulfide was ˜85 °C. The temperature difference between these two extraction methods is most likely due to the keratin extracted via L-cysteine having a more crystalline form. The absent of β-sheet keratin in SDS-PAGE results for L-cysteine extraction method could be another factor that affects the thermal properties. This could be explained by the fact that the amide bands for β-sheet are thermally differently stable than for α-helix. When keratin is extruded α-helical becomes β-sheet keratin and using a plasticizer lowers the temperature at which the phase transition occurs. The keratin extracted from β-sheet (green line) via sodium sulfide looks like a mixture of α -helix and βsheet. On the other hand, the keratin extracted from β-sheet via L-cysteine (orange line) shows the crystalline α -helical forms are losing urea molecules from 230 to 270 °C, and some of the α -helix is being converted to a β -sheet form. The β -sheet and/or the more amorphous mixture of α -helix and β -sheet degrades more slowly. At 430 °C, β -sheet keratin extracted by both methods (the orange and green lines) have nearly identical slopes.

Acknowledgments The authors acknowledge Ms. Ibukun Aibinu, Ms. Wanlapa Chaibangyang and Mr. Andrew Sujecki for their contribution in helping with the SDS-PAGE study, Baiada Poultry Pty Ltd for supplying the chicken feathers and Dr. Michael Czajka for helpful comments on the manuscript. The first author (FP) is grateful for a Research Training Program (RTP) scholarship administered by RMIT University. Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at doi:https://doi.org/10.1016/j.procbio.2019.04.010. References [1] M. Khosa, A. Ullah, A sustainable role of keratin biopolymer in green chemistry: a review, J. Food Process. Beverage 1 (2013) 1–8. [2] K. Wang, et al., Extracting keratin from wool by using L-cysteine, Green Chem. 18 (2) (2016) 476–481. [3] B. Ma, et al., Pure keratin membrane and fibers from chicken feather, Int. J. Biol. Macromol. 89 (2016) 614–621. [4] R. Endo, et al., Dimensional stability of waterlogged wood treated with hydrolyzed feather keratin, J. Archaeol. Sci. 35 (5) (2008) 1240–1246. [5] A.J. Poole, J.S. Church, The effects of physical and chemical treatments on Na 2 S produced feather keratin films, Int. J. Biol. Macromol. 73 (2015) 99–108. [6] K.M. Arai, et al., Amino‐acid sequence of feather keratin from fowl, Eur. J. Biochem. 132 (3) (1983) 501–507. [7] A. Ullah, et al., Bioplastics from feather quill, Biomacromolecules 12 (10) (2011) 3826–3832. [8] R. Fraser, T. MacRae, G.E. Rogers, Keratins: Their Composition, Structure, and Biosynthesis, Charles C. Thomas, 1972. [9] X.-C. Yin, et al., Study on effective extraction of chicken feather keratins and their films for controlling drug release, Biomater. Sci. 1 (5) (2013) 528–536. [10] S.I.N. Ayutthaya, S. Tanpichai, J. Wootthikanokkhan, Keratin extracted from chicken feather waste: extraction, preparation, and structural characterization of the keratin and keratin/biopolymer films and electrospuns, J. Polym. Environ. 23 (4) (2015) 506–516. [11] D.A. Parry, A. North, Hard α-keratin intermediate filament chains: substructure of the N-and C-terminal domains and the predicted structure and function of the Cterminal domains of type I and type II chains, J. Struct. Biol. 122 (1) (1998) 67–75. [12] W. Zhao, et al., Sustainable and practical utilization of feather keratin by an innovative physicochemical pretreatment: high density steam flash-explosion, Green Chem. 14 (12) (2012) 3352–3360. [13] Y.-X. Wang, X.-J. Cao, Extracting keratin from chicken feathers by using a hydrophobic ionic liquid, Process. Biochem. 47 (5) (2012) 896–899. [14] T.W. Thannhauser, Y. Konishi, H.A. Scheraga, Sensitive quantitative analysis of disulfide bonds in polypeptides and proteins, Anal. Biochem. 138 (1) (1984) 181–188. [15] J. Zhang, et al., Isolation and characterization of biofunctional keratin particles extracted from wool wastes, Powder Technol. 246 (2013) 356–362. [16] N. Eslahi, F. Dadashian, N.H. Nejad, An investigation on keratin extraction from wool and feather waste by enzymatic hydrolysis, Prep. Biochem. Biotechnol. 43 (7) (2013) 624–648. [17] C. Tonin, et al., Study on the conversion of wool keratin by steam explosion, Biomacromolecules 7 (12) (2006) 3499–3504. [18] T. Korniłłowicz-Kowalska, J. Bohacz, Biodegradation of keratin waste: theory and practical aspects, Waste Manag. 31 (8) (2011) 1689–1701. [19] W. Schmidt, Microcrystalline keratin: from feathers to composite products, MRS Symp. Procced. (2001) 702: 25. 2002. [20] N. Reddy, Y. Yang, Structure and properties of chicken feather barbs as natural protein fibers, J. Polym. Environ. 15 (2) (2007) 81–87. [21] A. Ullah, J. Wu, Feather fiber-based thermoplastics: effects of different plasticizers on material properties, Macromol. Mater. Eng. 298 (2) (2013) 153–162. [22] J.E. Winandy, et al., Potential of chicken feather fiber in wood MDF composites, Proceedings EcoComp vol. 20, (2003) 1–6. [23] F.T. Wallenberger, N. Weston, Natural fibers, plastics and composites natural, Materials Source Book, (2004). [24] W. Lamoolphak, W. De-Eknamkul, A. Shotipruk, Hydrothermal production and characterization of protein and amino acids from silk waste, Bioresour. Technol. 99 (16) (2008) 7678–7685. [25] J.R. Barone, W.F. Schmidt, N. Gregoire, Extrusion of feather keratin, J. Appl. Polym. Sci. 100 (2) (2006) 1432–1442. [26] P. Hill, H. Brantley, M. Van Dyke, Some properties of keratin biomaterials: kerateines, Biomaterials 31 (4) (2010) 585–593. [27] M.W. Donner, et al., Unravelled keratin-derived biopolymers as novel biosorbents

4. Conclusions In this study, L-cysteine was used for the dissolution of chicken feather keratin and compared with sodium sulfide extraction. The extracted keratin from L-cysteine exhibits a similar molecular mass of ca. 11 kg/mol based on SDS-PAGE, LCeMS/MS, and TGA compared to the extracted keratin from sodium sulfide. Both FTIR and Raman spectra analysis confirmed the changes in aged keratin extracted via sodium sulfide. This variation was less for extraction via L-cysteine as no significant changes were observed in the Raman spectra and only an increase in intensity of bands observed in FTIR data that could be due to crystallinity of keratin over time. Solid and liquid state NMR confirmed that the structure of keratin was retained following extraction via both methods and that 1H SSNMR could potentially be used as an analytical tool to differentiate between α- and β-keratin. The TGA and DTG curves for pure chicken feather and keratin extracted via sodium sulfide showed virtually identical behavior of decomposition, which proves the purity of the extracted keratin. However, the keratin extracted via Lcysteine showed a more crystalline structure. From this research, it can be concluded that protein can be extracted from chicken feathers using both sodium sulfide and L-cysteine. Although extraction with sodium sulfide poses environmental problems, the obtained yield (88%) was higher in comparison to the ecofriendlier L-cysteine method (66% total yield). Additionally, the structure of keratin is better preserved via L-cysteine extraction as changes in oxidation of amino acids was not evident in the IR spectra. The resultant keratin protein solutions could potentially be used for several 213

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