On chemistry of γ-chitin

Accepted Manuscript Title: On Chemistry of ␥-chitin Authors: Murat Kaya, Muhammad Mujtaba, Hermann Ehrlich, Asier M. Salaberria, Talat Baran, Chris T. Amemiya, Roberta Galli, Lalehan Akyuz, Idris Sargin, Jalel Labidi PII: DOI: Reference:

S0144-8617(17)30952-9 http://dx.doi.org/10.1016/j.carbpol.2017.08.076 CARP 12683

To appear in: Received date: Revised date: Accepted date:

15-7-2017 15-8-2017 17-8-2017

Please cite this article as: Kaya, Murat., Mujtaba, Muhammad., Ehrlich, Hermann., Salaberria, Asier M., Baran, Talat., Amemiya, Chris T., Galli, Roberta., Akyuz, Lalehan., Sargin, Idris., & Labidi, Jalel., On Chemistry of ␥-chitin.Carbohydrate Polymers http://dx.doi.org/10.1016/j.carbpol.2017.08.076 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

On Chemistry of γ-chitin Murat Kayaa*, Muhammad Mujtabaa, Hermann Ehrlichb, Asier M. Salaberriac, Talat Barand, Chris T. Amemiyae, Roberta Gallif, Lalehan Akyuzg, Idris Sargina, Jalel Labidic

a

Aksaray University, Faculty of Science and Letters, Department of Biotechnology and

Molecular Biology, 68100 Aksaray, Turkey b

Institute of Experimental Physics, TU Bergakademie Freiberg, Leipziger str. 23, 09599

Freiberg, Germany. c

Biorefinery Processes Research Group, Department of Chemical and Environmental

Engineering, University of the Basque Country (UPV/EHU), Plaza Europa 1, 20018 Donostia-San Sebastian, Spain d

Department of Chemistry, Faculty of Science and Letters, Aksaray University, Aksaray,

Turkey. e

Benaroya Research Institute at Virginia Mason, 1201 Ninth Avenue, Seattle, WA 98101,

USA f

Clinical Sensoring and Monitoring, Department of Anesthesiology and Intensive Care

Medicine, Faculty of Medicine, TU Dresden, Fetscher str. 74, D-01307 Dresden, Germany g

Aksaray University, Technical Vocational School, Department of Chemistry Technology,

68100, Aksaray, Turkey

* Corresponding Author Address: Aksaray University, Faculty of Science and Letters, Department of Biotechnology and Molecular Biology, 68100, Aksaray, Turkey. E-Mail: [email protected] Tel.: +90-382-288-2184 Fax: +90-382-288-2125

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Highlights 

First detailed physicochemical characterization of γ-chitin was carried-out



The isolated γ-chitin was compared with α and β forms



Chemical structure of the chitins was expressed through Quantum chemical calculations



γ-chitin was found much closer to α-chitin than β-chitin

Abstract The biological material, chitin, is present in nature in three allomorphic forms: α, β and γ. Whereas most studies have dealt with α- and β-chitin, only few investigations have focused on γ-chitin, whose structural and physicochemical properties have not been well delineated. In this study, chitin obtained for the first time from the cocoon of the moth (Orgyia dubia) was subjected to extensive physicochemical analyses and examined, in parallel, with α-chitin from exoskeleton of a freshwater crab and β-chitin from cuttlebone of the common cuttlefish. Our results, which are supported by13C CP-MAS NMR, XRD, FT-IR, Raman spectroscopy, TGA, DSC, SEM, AFM, chitinase digestive test and elemental analysis, verify the authenticity of γchitin. Further, quantum chemical calculations were conducted on all three allomorphic forms, and, together with our physicochemical analyses, demonstrate that γ-chitin is distinct, yet closer in structure to α-chitin than β-chitin.

Keywords: chitin; alpha; beta; gamma; characterization; molecular modelling

1. Introduction According to “Web of Science,” over the past 10 years, more than 9,000 studies have been conducted on chitin and more than 40,000 studies have been conducted on the most commonly known chitin derivative, i.e., chitosan. This number of studies has been constantly increasing over the past few years, confirming an abiding interest in this field. These studies suggest that chitin exhibits highly ordered, crystalline structures comprising three allomorphic forms designated as α, β and γ (Carlström, 1957; Hackman & Goldberg, 1965; Roberts, 1992). In α-chitin, the catenaries are arranged in layers (the catenaries of each end having the same direction or sense). In β-chitin, the adjacent layers are parallel and have the same 2

direction, while in α-chitin the adjacent layers have different directions and are anti-parallel. In γ–chitin, every 3rd layer has an opposing direction relative to the two preceding ones (Azuma et al., 2015). According to the currently held view, α-chitin is the most abundant allomorph, and is found in hard structures, whereas β- and γ-chitin are found in flexible structures (Jang, Kong, Jeong, Lee, & Nah, 2004). Myriad studies have been published with respect to detailed characterizations of α- and β-chitin forms. However, with regard to γchitin, despite its detection in various organisms (Kumirska et al., 2010; No, 1987; RamírezWong et al., 2016; Rudall & Kenchington, 1973), very is little known about its natural distribution and structure. (Rudall, 1962; Rudall (1963)) reported the presence of gamma chitin in cocoon threads of larvae of the spider beetle, Ptinus tectus, larval and adult peritrophic membranes (PM) of adult locust, cockroach, mantis and dragonfly silkworm larva (Antheraea pernyi) and in a sawfly larva (Phymatocera aterrima). Rudall and Kenchington (1973) described the use of XRD in order to delineate γ-chitin from the other chitin forms with respect to the configuration of parallel and antiparallel chains. Chitin is a biologically-important biopolymer which can be found in more than 70% of all living organisms in the world. α-chitin is the most common form found in the phylum Arthropoda, Porifera, Bryozoa and in the cell walls of fungi (Ehrlich et al., 2010; Ehrlich et al., 2013; Ehrlich et al., 2007; Muzzarelli, 2011). β-chitin is present within the cell walls of diatoms (Brunner et al., 2009), and skeletal structures of cephalopods (Hamed, Özogul, & Regenstein, 2016; Park & Kim, 2010; Rinaudo, 2006). γ-chitin was reported in Ptinus beetle cocoon fibers and stomach of Loligo sp (squid) (Jollès & Muzzarelli, 1999; Rudall, 1963; Rudall & Kenchington, 1973). Recently, chitin and its cognate chitin synthase genes have also been discovered in the fishes and amphibians, which comprise over half of all vertebrates (Tang, Fernandez, Sohn, & Amemiya, 2015). These chitin synthase genes have been shown to synthesize chitin at sites such as the skin and gut, though the amount of chitin produced is comparatively low relative to that in the arthropods, likely contributing to the long-held tenet that chitin was not produced in vertebrates. The low yield of chitin extracted from scale epithelia of salmon necessitated use of microscope FT-IR, which showed that it could be of the α or γ form. However, while the chitin was detectable via molecular, histochemical and chemical means, its definitive structure awaits additional, more sensitive, analyses. The role of chitin in the vertebrates is still unclear and is an area of active investigation. Chitin is known to be nontoxic, biodegradable, edible, biocompatible, antioxidative, antimicrobial, thermally stable, anti-oncogenic, and possesses a nanofibrous and porous surface. These characteristics enable chitin and its derivatives to be employed for myriad 3

economically-important applications in wide ranging fields including agriculture, medicine, food industry, textiles, cosmetics, extreme biomimetics and bioremediation, etc. (Anitha et al., 2014; Fernando, Poblete, Ongkiko, & Diaz, 2016; Hamed et al., 2016; Jayakumar, Prabaharan, Kumar, Nair, & Tamura, 2011; Jeon, Shahidi, & KIM, 2000; Merzendorfer; Petrenko et al., 2017; Rinaudo, 2006; Wysokowski et al., 2015). It is not surprising that for all of these applications, α- and occasionally, β-chitin (and their derivatives) have been employed (Wysokowski, Motylenko, et al., 2013). Due to limited available knowledge regarding its sources and physicochemical nature, γ-chitin has not yet been used in any application, emphasizing the need for procuring more information on this third allomorph. The characterization of γ-chitin has received scant attention in the literature, these studies primarily utilizing XRD in order to differentiate it from the α and β forms (Jollès & Muzzarelli, 1999; Kumirska et al., 2010; No, 1987; Ramírez-Wong et al., 2016; Rudall & Kenchington, 1973). Further, it is suggested in these reports that the arthropod cocoon is, in addition to being a rich source of silk, a bona fide source of gamma chitin (the silk proteins readily being removed during the chitin extraction process) (Davies, Knight, & Vollrath, 2013; Rudall, 1962, 1963; Rudall & Kenchington, 1973) More recently, Jang et al. (2004) described presumptive γ-chitin using a variety of complementary methods such as FT-IR, TGA, DSC, XRD and 13C CP-MAS NMR and made comparisons with well-characterized alpha and beta forms. The study, though pioneering in characterizing this novel allomorphic form of chitin, lacked certain details, including the source of the tissue for the γ-chitin isolate. Our present study was undertaken with the goal of better understanding the chemical nature of γ-chitin, and establishing a set of defined reference criteria for this third allomorph. Here, we isolated chitin from the cocoon of the moth, Orgyia dubia (Noctuidae: Lepidoptera), and analyzed it structurally and physicochemically. Based on these analyses we could determine that our moth cocoon chitin was definitively of the γ form and that our analyses are generally in agreement with those of Jang et al. (2004). In addition to carrying out the same analyses used in this previous study, we further characterized our γ-chitin specimen using additional techniques that included SEM, AFM, Raman spectroscopy, elemental analysis and chitinolytic assay. Our extensive analyses and molecular modelling collectively provide a much clearer picture of the characteristics of γ-chitin, particularly with reference to the alpha and beta allomorphs to which it is compared. This new information and the better overall definition of the diagnostic properties of γ-chitin will provide increased insight in terms of its biological roles and potential areas of application. 4

2. Materials and Methods 2.1 Chemicals All the chemicals (HCl, NaOH, chloroform, methanol, ethanol, NaH2PO4, Na2HPO4 and Na2CO3) were purchased from Sigma-Aldrich with the exception of potassium hexacyanoferrate(III) salt (Merck). Enzymatic hydrolysis of respective chitin isolates was performed with Streptomyces griseus chitinase (EC. 3.2.1.14; Sigma-Aldrich).

2.2 Extraction of chitin isolates α-, β- and putative γ-chitin isolates were respectively obtained from the exoskeleton of freshwater crab Potamon ibericum (Alanya, Turkey, 12.07.2015), cuttlebone of common cuttlefish (Sepia sp) (bought from a pet shop) and the cocoon of the moth Orgyia dubia (Figure 1). The cocoons were collected on bushes by hand in Lalebaglari, Aksaray (Turkey) on 24.06.2016. The same extraction procedure was followed for each chitin isolation by starting 15 g material from each source. Briefly, the selected samples were crushed by mortar into powder form and rinsed in deionized water. Then the samples were kept in an oven to dry for 2 days at 50° C. For demineralization the specimens of the crab exoskeletons, cuttlebones and cocoons were stirred in 2 M HCl at 50° C for 4 h using a magnetic stirrer. After HCl treatment all the samples were filtered using a filter paper (400x400 mm, FILTROSN ANOIA, S.A.) and washed extensively up to pH 7 with distilled water. The samples were refluxed for deproteinization in 2 M NaOH solution at 110° C for 16 h. After that the samples were washed extensively (with distilled water) and filtered as above using filter paper. Then, for depigmentation the isolates were treated with a water-methanol-chloroform mixture (4:2:1, by volume). Finally, the isolates were washed and filtered again up to neutral pH. All the samples were air-dried at room temperature for one week.

2.3 Fourier transform infrared spectroscopy (FT-IR) Infra-red spectra for chitin isolates were recorded using a Perkin Elmer FT-IR spectrometer over the range of 4000-650 cm−1. For each sample, about 10 mg of purified chitin was used.

2.4 Raman spectroscopy

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Raman spectra were obtained using a Raman spectrometer (RamanRxn1™, Kaiser Optical Systems Inc., Ann Arbor, USA), which was coupled to a light microscope (DM2500 P, Leica Microsystems GmbH, Wetzlar, Germany). The excitation of Raman scattering was taken with a diode laser emitting at a wavelength of 785 nm, propagated to the microscope with a 100 µm optical fibre and focused on the samples by means of a 100x / 0.75 microscope objective, leading to a focal spot of about 20 µm. The Raman signal was collected in reflection configuration and sent to the f/1.8 holographic imaging spectrograph by using a 100 µm core optical fibre. The spectral resolution in the range 150-3250 cm-1 was about 4 cm-1. Raman spectra of crab (α-) and cuttlebone (β-) chitin were recorded with a laser power of 200 mW, using an integration time of 1 s and averaging of 40 spectra in order to improve the signal-to-noise ratio. The spectrum of moth cocoon chitin was recorded with a laser power of 50 mW and an integration time of 0.5 s. As this sample was slightly pigmented, the laser power was reduced in order to avoid thermal damage and the integration time was reduced to comply with intense fluorescence. 250 spectra were averaged in order to improve the signalto-noise ratio. Moreover, 15 min optical bleaching with laser power of 50 mW was performed prior to the measurement in order to reduce the fluorescence. Spectroscopic data were processed with MATLAB toolboxes (MathWorks Inc., Natick, USA). A variable baseline was calculated applying the function “msbackadj”, estimating the baseline within multiple windows of 200 cm-1 width, shifted with 100 cm-1 step; a linear interpolation method was chosen. For better comparison, the spectra were normalized to the total area.

2.5 Thermogravimetric analysis (TGA) TGA assays were carried out using a TGA Q500-TA instrument. Respective chitin isolates were heated at a constant temperature rate of 10 ºC min-1 from 30 to 600º C under a nitrogenous atmosphere.

2.6 Differential scanning calorimetry (DSC) measurements DSC measurements for chitin isolates were performed on a Mettler Toledo DSC822e Differential Scanning Calorimeter (DSC) (Switzerland). The DSC curves were obtained under dynamic N2 atmosphere. The chitin samples (5 mg) were heated (from −50 to 420° C) with at a heating rate of 5º C min−1 after placing in hermetic aluminum pans.

2.7 X-ray diffraction (XRD) 6

X-ray diffraction patterns of the chitin allomorphs were analyzed via a Philips X’pert Pro automatic diffractometer using Cu-Kα radiation (operating at 40 kV and 40 mA) over the angular range of 5–70° 2θ (step size = 0.04 and time per step =353 s) at room temperature. The crystalline index (CrI) of three chitin allomorphs was calculated (Focher, Beltrame, Naggi, & Torri, 1990); CrI110 (%) = [(I110 – Iam) / I110] x 100

(1)

where Iam is the amorphous peak and I110 is the maximum intensity at 2θ ≅ 19°.

2.8 Elemental analysis The C, N, H and O contents of the respective chitin isolates were determined using Euro EA Elemental Analyzer (CHNS, EuroVector). For analysis, 15 mg of chitin for each sample was used. The analysis was carried out following complete and instantaneous oxidation via combustion with oxygen at an approximate temperature of 1020° C. The separation was carried out by transporting the combusted samples using the gas carrier to a chromatographic column. The degree of acetylation (DA) of α-, β- and γ-chitin isolates was calculated by following formula (Xu, McCarthy, Gross, & Kaplan, 1996). DA=[(C/N-5.14)/1.72]∗100

(2)

where C:N is ratio of carbon to nitrogen (w/w). 2.9 Nuclear magnetic resonance (13C CP-MAS NMR) The 13C solid state cross-polarized magic angle spinning nuclear magnetic resonance (13C CP-MAS NMR) spectra were obtained for chitin isolates on a Bruker 400WB Plus spectrometer. Spectra were constructed by using a 4mm CP-MAS probe with a sample spinning rate of 10,000 Hz. 13C CP-MAS spectra at 100.6 MHz of the solid state samples were obtained using 12 h spectral accumulation time, a time domain 2K points, a spectra width of 29 kHz, and contact time of 1.5 ms and an inter-pulse delay of 5 s. The DA of the chitin samples was calculated according to the method reported by Kasaai (2010).

2.10 Scanning electron microscope (SEM) 7

The surface characteristics of respective chitin isolates were inspected on a QUANTA FEG 250 scanning electron microscope. A Sputter Coater (Gatan Precision Etching Coating System) was used for gold coating of the chitin samples.

2.11 Atomic force microscopy (AFM) AFM measurements of chitin isolates were taken via a Dimension 3100 NanoScope IV (Veeco, USA). The samples were scanned in tapping mode under normal conditions through silicon nitride cantilevers having a tip nominal radius of 10 nm.

2.12 Colloidal solutions of chitin isolates for chitinase digestive test Colloidal solutions of chitin were prepared by incubating 100 mg of each chitin isolate in 2 mL of cold concentrated HCl solution (37%) for 24 h (M. Kaya et al., 2015). After the incubation, 5 mL of water-ethanol solution was poured into the chitin solutions and stirred on a magnetic stirrer. Each chitin solution was then placed in dialysis tubing (Viskase Sales Crop, Seamless Cellulose Tubing, Size: 16/32, 100ft, Lot: 208001) and dialyzed against water until reaching neutral pH. Colloidal solutions were subsequently dialyzed against phosphate buffer solution (pH 7.0, 100 mM). Finally, colloidal chitin solutions in the dialysis bags were transferred into test tubes and kept at 4° C.

2.12.1 Chitinase digestive test on chitin isolates Hydrolytic activity of S. griseus chitinase on respective chitin isolates was assayed following the same procedure reported elsewhere (M. Kaya et al., 2015). Chitin isolates were treated with concentrated HCl in an ice-bath to give colloidal chitin solutions. Subsequently, the chitin solutions were incubated at room temperature. After 3-day incubation, hydrolytic activity of the enzyme was stopped by heating in hot water bath. The solution of the digestion products was separated by centrifugation. The supernatant solutions were treated with the reagent potassium hexacyanoferrate(III) in Na2CO3 solution in boiling water bath. The debris was removed by centrifugation and the supernatants were assayed at 420 nm with a UV-vis spectrophotometer (Shimadzu, Tokyo, Japan, Model 1601). As a result of chitinolytic activity, chitin chains are digested and oligosaccharide residues with reducing end are released into the solution. The reducing end has hemiacetal functionality and therefore can reduce metal cations. Digestion products were quantified employing the spectrophotometric potassium

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ferricyanide assay (Horn & Eijsink, 2004), where the amount of K4[Fe(CN)6] present in the solution is proportional to the amount of reducing sugar content. Freshly prepared chitinase solutions (5 mg of enzyme in 10 mL of phosphate buffer solution 100 mM, pH: 7.0) were used throughout the experiments. The reagent for the ferricyanide assay was prepared by dissolving 0.025 g of potassium hexacyanoferrate(III) salt in 0.5 L of 0.5 M Na2CO3 solution and kept in the dark. 2.13 Investigations of chemical structure of α-, β- and γ-chitin using quantum chemical calculations The subtle differences of the H bonds in molecule structure can be detected via changes in the FT-IR spectrum. In the present study, to explain the differences in the FT-IR spectra of the respective chitin isolates (α-, β- and moth cocoon or γ-chitin), inter- and intrasheet H bonds were investigated for the first time via quantum chemical calculations using Spartan packet software. The semi empirical PM3 method was used for quantum chemical calculations. Also, heat of formation values and H bond distances of the molecules were determined in order to account for stability. Semiempirical Methods which used a part of experimental results (AM1, PM3, MNDO) can made calculations with pre-calculated data for organic molecules. The PM3 method is more accurate than the AM1 and MNDO method especially for large molecules. It gives more accurate results about some properties such as heat of formation. For this reason, PM3 method was chosen for the quantum chemical calculations for the identification of α-, β- and γ-chitin.

3. Results and Discussions 3.1 FT-IR Three different allomorphic forms of chitin, as α, β and γ are known from the literature. FT-IR spectroscopy is one of the fastest and simplest techniques used to determine allomorphic form of chitin (Darmon & Rudall, 1950). As known from many reported studies regarding FT-IR spectrum of α- and β-chitin, the amide I band of the α form gives two sharp bands at 1660 and 1620 cm-1 whereas the β form only presents a single band around 1640 cm1

due to the hydrogen bonds between the molecules. However, except for a single study (Jang

et al., 2004), there is limited information about the FT-IR spectrum of the γ-chitin. In the present study, the FT-IR spectra of respective α, β and moth cocoon chitin were obtained and are depicted in Figure 2.

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. When the chitin spectrum of the crab exoskeleton was examined, it was observed that the amide I band was clearly divided into two sharp peaks, 1652 and 1620 cm-1 (Figure 2b) confirming its status as α-chitin. For chitin from the cuttlebone, the amide I was observed as undivided single peak at 1640 cm-1 in agreement with its status as β-chitin (Jang et al., 2004) (Figure 2d). When the FT-IR spectrum of the chitin from moth cocoon was examined, we observed that the amide I band was partially divided into two bands, one weak and unclear at 1654 cm-1 and one sharper peak at 1621 cm-1 (Figure 2f). Table S1 also lists other important peaks belonging to the spectra of the respective chitin isolates. The observations of the moth cocoon chitin are largely consistent with its identification as γ-chitin with the possible exception that the previous FT-IR of γ-chitin by Jang et al. (2004) showed an amide I band that had a completely divided peak that was split into two sharp sub-peaks at 1660 and 1620 cm-1. Given the overall similarities (further discussed below), we shall henceforth refer to the moth cocoon chitin as γ-chitin.

3.2 Raman spectroscopy Raman spectra of α- and β-chitin have already been reported (Bo et al., 2012; Focher, Naggi, Torri, Cosani, & Terbojevich, 1992; Wysokowski, Bazhenov, et al., 2013) and our analysis of chitin from crab exoskeleton and cuttlebone, respectively, are largely corroborative (Figure 3). The amide I vibration of α- chitin shows a splitting pattern in two separate bands at 1622 and 1658 cm-1, while in the spectrum of β-chitin only one band at 1662 cm-1 is visible. These findings were also observed to be in good agreement with the IR spectra described for FT-IR (above Section 3.1). For the moth cocoon (γ-) chitin, the Amide I vibration produced a broad asymmetric band at 1618 cm-1. The amide I band at higher wavelength is found at 1657 cm-1, but is barely separated. This is in very good agreement with the results of FT-IR spectroscopy, and confirms the existence of non-equivalent populations of C=O groups with different degrees of hydrogen bonding among the three chitin allomorphs. The main Raman bands are reported and assigned in Table S2. Overall, the spectral profile of γ-chitin appears more similar to that of α-chitin.

3.3 TGA

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TG and DTG curves were constructed for α-, β- and γ-chitin to evaluate and compare their thermogravimetric profiles with respect to thermal stability and degree of degradation (Figure 4). In accordance with previous literature reports concerning degradation stages of chitin, here we also observed the degradation of α-, β- and γ-chitin in two different stages (Al Sagheer, Al-Sughayer, Muslim, & Elsabee, 2009; Shinsuke Ifuku et al., 2010). First weight loss was due to the evaporation of water from hydrophilic groups in chitin chains (Jang et al., 2004). For α-, β- and γ-chitin, the first weight loss was recorded at temperature range of ≈ 205 °C (5.20%), ≈ 205 °C (7.93%) and ≈ 130 °C (4.95%) respectively. No notable differences were observed in first degradation stage for all three of the respective chitin samples. The second weight loss corresponds to the degradation of polymeric structure i.e., saccharide ring dehydration and deterioration of acetylated chitin units (Paulino, Simionato, Garcia, & Nozaki, 2006). DTGmax values were recorded for all three chitin isolates as 391.09 °C (74.37 %) for α-chitin, 348.77 °C (72.56 %) for β-chitin and 381.99° C (69.87 %) for γ-chitin. The results revealed that DTGmax for α-chitin was slightly higher than γ-chitin, however, it was much higher than β-chitin. The γ-chitin revealed a low degradation % at its DTGmax temperature compared to α-chitin and β-chitin. This can be attributed to the structural organization of γ-chitin as it is comprised of thick rod shaped microfibers (see SEM section below, 3.8). In previous reports the DTGmax values for α-chitin was recorded as 350-400 ° C (Abdou, Nagy, & Elsabee, 2008; Shinsuke Ifuku et al., 2010; S. Ifuku & Saimoto, 2012; Paulino et al., 2006; Sajomsang & Gonil, 2010). M. Kaya and Baran (2015) reported the DTGmax values for Periplaneta americana wing chitin as 389° C. Sajomsang and Gonil (2010) observed the DTGmax values as 372° C and 369° C for crab shell and Cicada sloughs respectively. For β-chitin the thermal stability was reported in range of 250-350° C (Chen, Wang, & Ou, 2004; Maeda, Jayakumar, Nagahama, Furuike, & Tamura, 2008; Peesan, Rujiravanit, & Supaphol, 2003). The thermal stability of γ-chitin was given as 380.7° C in the sole report on its characterization (Jang et al., 2004). Taking into account the previous report, DTGmax of γ-chitin in current study were found in a close accordance with this earlier report. DTGmax of γ-chitin was observed intermediate between that of α- and β-chitin.

3.4 DSC An alternative metric of the thermal stability of chitin alloforms can be obtained using differential scanning calorimetry. DSC analyses of α-, β- and γ-chitin isolates are shown in Figure 5. Two endothermic peaks were observed for all the chitin forms. For all the chitin 11

isolates, the first degradation was because of water evaporation. The maximum degradation temperature (Tmax) for α-, β- and γ-chitin was recorded as 65.25, 72.59 and 66.05 °C respectively. The second degradation step was due to degradation of chitin. Tmax values were determined as 375.34 °C for α-chitin, 336.00 °C for β-chitin and 373.38 °C for γ-chitin. According to the DSC data, thermal stability of γ-chitin was recorded very close to α-chitin, however, the thermal stability of β-chitin was observed to be much lower.

Comparing the DSC mass losses with Jang et al. (2004), the same degradation steps (two steps) were recorded for the newly isolated α-, β- and γ-chitin in the present study. However, thermal stabilities of our newly isolated α-, β- and γ-chitin were collectively higher than those reported by Jang et al. (2004). In both studies, the thermal stability of γ-chitin was found closer to that of α-chitin rather than β-chitin. Additionally, in the current study the maximum degradation temperature of γ-chitin (373.38 °C) was recorded comparatively higher than that of γ-chitin (300 °C) reported by Jang et al. (2004).

3.5 XRD XRD analysis is the gold standard for assessing the overall crystallinity of isolated chitin and for differentiating the three different allomorphic forms (α-, β- and γ-chitin) (Cárdenas, Cabrera, Taboada, & Miranda, 2004). The XRD patterns of α-, β- and γ-chitin here showed a total of six crystalline reflections at 2θ range, two of which were observed as relatively sharper peaks while the other four peaks were less pronounced (Figure 6). For α-, βand γ-chitin the I020 crystalline reflection values were observed at 9.46, 8.59° and 9.35° respectively. For α- and γ-chitin the second reflection peak was observed sharper at 12.74° while for β-chitin the same peak was recorded slightly weaker at 12.29°. As expected the CrI values were recorded slightly higher for α-chitin, while lower for the β-chitin. The CrI value of the γ-chitin was observed as a value (68.6°) in the middle of both α- and β-chitin (Table S3).

In accordance with previous reports on the XRD of the different allomorphic forms of chitin we observe nearly the same reflection peaks in the current study (Al Sagheer et al., 2009; Hajji et al., 2014; Zhang, Haga, Sekiguchi, & Hirano, 2000). Jang et al. (2004) revealed crystalline reflections at 9.6, 19.6, 21.1 and 23.7° for α-chitin, 9.1° and 20.3° for β-chitin, and 12

at 9.6 and 19.8° for γ-chitin. Fan, Saito, and Isogai (2008) reported the crystalline peaks for βchitin extracted from squid pen at 9.8° and 19.3°. Cárdenas et al. (2004) reported the main XRD reflection peaks for α-chitin (shrimp, lobster, prawn and king crab) at 9.28° ± 0.1, 19.36° ± 0.3, 26.20° ± 0.1 and for β-chitin (squid) as 8.64°, 18.78° and 26.24°.

3.6 Elemental analysis Theoretically the percent nitrogen in chitin is calculated to be 6.89% when completely acetylated, however, it is impossible to observe these theoretical values in practice because chitin in powdered form absorbs water from the atmosphere under normal conditions thus resulting in anywhere from 1-8 % water. For all intents and purposes, it can be stated that chitin samples with nitrogen content close to 6.89 % and acetylation value close to 100% are in high purity. In the present study, N % values for α-, β- and γ-chitin were recorded as 6.55, 5.79 and 6.13 %, respectively. The acetylation values of α-, β- and γ-chitin were calculated as 90.1%, 115.2% and 88.6%, respectively. The carbon (C), hydrogen (H) and oxygen (O) contents for all the three chitin isolates are given in Table S4. Looking at some previous studies, the N % and DA values has been recorded as 6.24 % and 112.2 % for shrimp shell chitin, 6.45 % and 101.2 % for beetle (Liu et al., 2012); 5.92 % and 132 % for the bumblebee chitin, 4.85 % and 245 % for the shrimp chitin (Majtan et al., 2007); 5.92 % and 102.3 % for cicada slough chitin (Sajomsang & Gonil, 2010); 6.20 % and 118 % for the purified crab chitin (Yen, Yang, & Mau, 2009); commercial chitin from SigmaAldrich 6.13 % and 112 % respectively (Murat Kaya, Baran, & Karaarslan, 2015). All of the other analyses carried out in these studies suggested the chitin obtained was of high purity. As a general summary of these studies, the N % values were between 4.85-6.45 % and the DA values were in the range of 101.2-245 %. In the present study, the N % and DA values measured for α-, β- and γ-chitin were very similar to those of the commercially available chitin used in literature (Murat Kaya, Evaldas Lelešius, et al., 2015). Compared to α- and γchitin, the N % value for β-chitin was observed to be slightly lower, and accordingly, the DA value of the β-chitin was also slightly higher than that of other α- and γ-chitin. These phenomena can be attributed to the presence of water content, comparatively higher than the rest of other two chitin forms which was also observed in the TGA analysis of β-chitin. 3.7 13C CP-MAS NMR 13

C CP-MAS NMR spectra and peaks of carbon atoms of the isolated α-, β- and γ-

chitin structures are given in Table S5 and Figure 7. When the 13C CP-MAS NMR spectrum 13

of α-chitin was examined, it was observed that the peaks of C3 and C5 were split into doublets. When the 13C CP-MAS NMR spectrum of β-chitin was examined, it was observed that, unlike α-chitin, the C3 and C5 peaks gave a single peak in singlet form. The observation of this variation in the 13C CP-MAS NMR and FT-IR spectra of the two allomorphs indicate that the isolated α- and β-chitin were of good purity. On the other hand, it has been reported in the literature that γ-chitin is a combination of α and β chitin in terms of characteristics and that it resembles the α-chitin structure (Jang et al., 2004). When the 13C CP-MAS NMR spectrum of γ-chitin obtained in the present study was examined, it was observed that C3 and C5 peaks were split into doublets in a similar manner as in the NMR spectrum of α-chitin. Other important resonance values are listed in Table S5. Also, the degree of acetylation (DA) was calculated using 13C CP-MAS NMR resonance peaks and listed in Table S5. DA results calculated by 13C CP-MAS NMR supported the DA values calculated by elemental analysis.

3.8 SEM Surface morphologies of α, β and γ chitin samples obtained from different sources are shown in Figure 8. As is seen from Figure 8a,b, surface of α-chitin had clearly visible nanofibers but only slightly visible and smaller nanofibers were present in the β-chitin (Figure 8e,f). In contrast to α and β chitin, γ chitin had microfibers on the surface not having any nanofiber (Figure 8i,j). In earlier papers, it was stated that α-chitin consisted of nanofibers and sometimes pores between the fibers (Al Sagheer et al., 2009; Shinsuke Ifuku, Nomura, Morimoto, & Saimoto, 2011; Mushi, Butchosa, Salajkova, Zhou, & Berglund, 2014). Slightly visible fibers were reported for β-chitin (Murat Kaya et al., 2016; Lavall, Assis, & Campana-Filho, 2007). But surface morphology of γ-chitin was described here for the first time and it was observed that γ-chitin from the cocoon had long microfibers.

3.9 AFM AFM is used as a complementary method to SEM for higher resolution analysis of macromolecular structures. After our SEM analyses, clear nanofibers were observed for αchitin, smaller and less defined nanofibers were seen for β-chitin. On the other hand, strong and visible microfibers were recorded for γ-chitin. In order to visualize the chitin structure in more detail, sonication and homogenization were used to separate possible nanofibers from

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each other and the resultant products were analyzed using AFM. As was expected, AFM gave better and clear view than SEM for each chitin sample (Figure 8c,d,g,h,k,l). For α- and β-chitin, very clear nanofibers were observed; but for γ-chitin, rod shaped microfibers were seen and these γ-chitin rods were comprised of tightly aligned nanofibers (Figure8k,l). A vastly different surface morphology was observed for γ-chitin compared to αand β-chitin despite applying the same isolation procedure and equipment, and we attribute these differences to the inherent macromolecular organization of the respective allomorphs. 3.10 Chitinase hydrolytic activity on α-, β- and γ-chitin isolates The extent of the hydrolysis of α, β and γ chitin isolates was determined by the percentage of hydrolysis after chitinase treatment. The amount of reducing sugars present in the supernatant was determined calorimetrically from the decrease in the absorbance of ferricyanide. The chitin isolates were hydrolysed by chitinase under the same incubation conditions. After 72 h incubation, the highest percentage of digestion was recorded for βchitin closely followed by α-chitin, 94% and 93%, respectively. The lowest chitinolytic activity, 65%, was observed for γ-chitin. This comparative low level of hydrolysis was unexpected but perhaps attributable to the macromolecular organization of microfibrils in γchitin from cocoon. As indicated in SEM and AFM images, γ-chitin exists in the moth cocoon as higher-order microfibers, which could prevent accessibility of enzyme molecules to internal glycosidic bonds. 3.11 Chemical structure analysis of α-, β- and γ-chitin Quantum chemical calculations of α-, β- and γ-chitin are summarized in the Figure S1. According to results of the quantum chemical calculations, it was determined that 7 intersheet H bonds were formed in the α- and γ-chitin, whereas 5 inter-sheet H bonds were formed in the β-chitin. Additionally, when intra-sheet H bonds were examined for α-, β- and γ-chitin, it was observed that α-chitin has 9 intra-sheet H bonds. In contrast, as seen from the Figure S1, 6 intra-sheet H bonds were formed for β-chitin and 3 intra-sheet H bonds for γ-chitin. The increase in the number of H bonds especially those involved in intermolecular bonding increases the stability of the molecules due to an increase of strong interactions. Although, the numbers of inter-sheet H bonds α- and γ-chitin are equal, they do not exhibit the same stability according to the TGA results. This situation can be explained by changing of the Hbond distances in the molecules. H bonds formed in the α-chitin were shorter than those of the γ-chitin (Table S6). It means that the intermolecular interactions were stronger in the α-chitin 15

than β and γ forms. Also its degradation temperature could be higher. These results were agreement with the TGA results. In the OH and NH stretching region in the FT-IR spectra of α- and γ-chitin, two peaks were observed. The strong OH stretching peaks of α-chitin which are assigned the intra and inter-sheet H bonds were recorded in the range of 3200-3500 cm-1. The NH stretching peak with the amide I overtone was observed at nearly 3180 cm-1 for α-chitin. In the same way, γchitin showed in this region partially divided OH stretching peaks centered in the range of 3200 and 3400 cm-1. According to the results our quantum chemical calculations, H bonds formed in the γ-chitin molecule structure are different from those of the α-chitin. For this reason, OH stretching peak of the γ-chitin may be partially divided. However, a single broad band was observed for OH and NH stretching in the β-chitin spectrum. It can, in fact, be that the number of intra and inter-sheet H bonds formed between CH2OH---OH and CH2OH--CH2OH groups was greater than that for α-and γ-chitin (Figure S1). When the OH stretching band was broader, it was overlapping with the NH stretching band. Additionally, it affected the aliphatic stretching peaks. The C-O stretching observed at around 1100 cm-1 was split in to two peaks for α-and γ-chitin but less sharp and partially divided peak was recorded for β-chitin. When the data (Figure S1) were examined, it was observed that there were 2 H bonds between the C-O and O-H groups for β-chitin. However, the numbers of H bonds between C-O and O-H groups were 4 and 1 for α-and γ-chitin, respectively. In the β-chitin structure, H bonds formed between C-O and O-H groups may be expected to different direction according to those of the α- and γ-chitin, thereby it was possible that C-O absorption band may be observed as different. Heat of formation values obtained from the quantum chemical calculations were calculated as -7499,55 kJ/mol, -7096,98 kJ/mol and -7425,74 kJ/mol for α-, β- and γ-chitin, respectively. These values are close to one-another for α-and γ-chitin, but lower for β-chitin. This value indicates that the β-chitin is more unstable than the α- and γ-chitin. Also, it can explain why β-chitin has a higher reactivity for reactions and higher activity for solvents than α-chitin, as mentioned by Jang et al. (2004).

Conclusions In this study, γ-chitin was isolated from a bona fide γ-chitin source and characterized comprehensively with respect to α- and β-chitin. In accordance with previous reports, here γchitin was found to be structurally closer to α-chitin. Additionally, in order to better 16

understand the structural properties of the γ-chitin, extra characterizations were carried out for the first time. The obtained results of these extra characterizations are summarized as: (i) Raman spectroscopy showed amide I band of γ-chitin broadened and partially divided in accordance with FT-IR spectra. (ii) SEM exhibited the surface morphology of γ-chitin to be comprised of microfibers however the surface structure of α- and β-chitin are comprised of nanofibers. (iii) After carrying out sonication and homogenization, AFM revealed that rod shaped microfibers in γ-chitin consisted of tightly bonded nanofibers. (iv) Chitinase digestive rate of γ-chitin was recorded comparatively lower than α- and β-chitin which can be ascribed to the rod shaped tightly packed microfibers in the structure of γ-chitin. (v) By evaluating the DA values and nitrogen contents, elemental analysis proved that all the chitin isolates were of high purity. (vi) Molecular structure of α-, β- and γ-chitin were investigated theoretically and these theoretical results supported the physicochemical characterizations. However, some differences were also observed from an only available report by Jang et al. (2004). These differences are as follow: (i) According to Jang et al. (2004), amide I band of FT-IR spectra was observed as fully divided for α- and γ-chitin but unlike previous report by Jang et al. (2004), in current study the amide I band was recorded fully divided for α-chitin but partially divided for γ-chitin. (ii) Considering DSC results, thermal stability in the present study was recorded relatively higher than earlier report by Jang et al. (2004) for all of the three chitin forms. These alterations in FT-IR and DSC results for γ-chitin can be ascribed to source. In this study, chitin extracted from moth cocoon was deemed to be γ-chitin due to overt structural differences from α- and β-chitin and because of general agreement with the physicochemical information on γ-chitin based on previous available reports. Our analyses provide a more detailed insight into the physicochemical nature of γ-chitin in comparison with alpha and beta forms. Our findings should prove useful for establishing a set of reference criteria for γ-chitin, for better understanding its distribution and structure-function relationships in the natural world, and as a prelude to its use in biotechnological applications.

Acknowledgments We are thankful for the following financial support – DFG Project HE 394/3-2 and the BHMZ Programme of Dr.-Erich-Krüger-Foundation in Germany.

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Figure 1. Animals and material used for chitin extraction: a, b) freshwater crab (Potamon ibericum), c, d) cuttlebone of common cuttlefish (Sepia sp.) and e) caterpillar and f) cocoon of the moth (Orgyia dubia).

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Figure 2. FT-IR spectra of a,b) α-chitin, c,d) β-chitin and e,f) γ-chitin studied (Amide I and amide II bands of b) α-chitin, d) β-chitin and f) γ-chitin).

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Figure 3. Raman spectra of α-, β- and γ-chitin studied.

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Figure 4. TGA/DTG of α-, β- and γ-chitin studied, a) TGA and b) DTG

Figure 5. DSC thermograms of α-, β- and γ-chitin studied.

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Figure 6. X-ray diffractograms α-, β- and γ-chitin studied.

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Figure 7. 13C CP/MAS NMR spectra of α-, β- and γ-chitin studied.

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Figure 8. SEM and AFM images of α-chitin (a,b,c,d), β-chitin (e,f,g,h) and γ-chitin (i,j,k,l) studied (SEM: a,b,e,f,i,j; AFM: c,d,g,h,k,l).

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