Nonspecific enzymatic hydrolysis of a highly ordered chitopolysaccharide substrate

Journal Pre-proof Nonspecific enzymatic hydrolysis of a highly ordered chitopolysaccharide substrate Daria N. Poshina, Sergei V. Raik, Arina A. Sukhova, Irina V. Tyshkunova, Dmitry P. Romanov, Elena V. Eneyskaya, Anna A. Kulminskaya, Yury A. Skorik PII:

S0008-6215(20)30562-0

DOI:

https://doi.org/10.1016/j.carres.2020.108191

Reference:

CAR 108191

To appear in:

Carbohydrate Research

Received Date: 19 August 2020 Revised Date:

26 October 2020

Accepted Date: 29 October 2020

Please cite this article as: D.N. Poshina, S.V. Raik, A.A. Sukhova, I.V. Tyshkunova, D.P. Romanov, E.V. Eneyskaya, A.A. Kulminskaya, Y.A. Skorik, Nonspecific enzymatic hydrolysis of a highly ordered chitopolysaccharide substrate, Carbohydrate Research, https://doi.org/10.1016/j.carres.2020.108191. This is a PDF file of an article that has undergone enhancements after acceptance, such as the addition of a cover page and metadata, and formatting for readability, but it is not yet the definitive version of record. This version will undergo additional copyediting, typesetting and review before it is published in its final form, but we are providing this version to give early visibility of the article. 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. © 2020 Elsevier Ltd. All rights reserved.

1

NONSPECIFIC ENZYMATIC HYDROLYSIS OF A HIGHLY ORDERED

2

CHITOPOLYSACCHARIDE SUBSTRATE

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Daria N. Poshina1, Sergei V. Raik1, Arina A. Sukhova1, Irina V. Tyshkunova1,

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Dmitry P. Romanov2, Elena V. Eneyskaya3, Anna A. Kulminskaya3, Yury A. Skorik1,*

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31, 199004 St. Petersburg, Russian Federation

Institute of Macromolecular Compounds of the Russian Academy of Sciences, Bolshoy pr. V.O.

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199034 St. Petersburg, Russian Federation

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«Kurchatov Institute», 188300 Gatchina, Orlova roscha 1, Russian Federation

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Institute of Silicate Chemistry of the Russian Academy of Sciences, Adm. Makarova emb. 2,

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Petersburg Nuclear Physics Institute named by B.P. Konstantinov of National Research Center

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*Corresponding author at the Institute of Macromolecular Compounds of the Russian Academy of

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Sciences. E-mail address: [email protected] ORCHID: 0000-0002-9731-6399

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17 Abstract

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Chitin and chitosan can undergo nonspecific enzymatic hydrolysis by several different

20

hydrolases. This susceptibility to nonspecific enzymes opens up many opportunities for producing

21

chitooligosaccharides and low molecular weight chitopolysaccharides, since specific chitinases and

22

chitosanases are rare and not commercially available. In this study, chitosan and chitin were

23

hydrolyzed using several commercially available hydrolases. Among them, cellulases with the

24

highest specific activity demonstrated the best activity, as indicated by the rapid decrease in

25

viscosity of a chitosan solution. The hydrolysis of chitosan by nonspecific enzymes generated a

26

sugar release that corresponded to the decrease in the degree of polymerization. This decrease

27

reached a maximum of 3.3-fold upon hydrolysis of 10% of the sample. Cellulases were better than

28

lysozyme or amylases at hydrolyzing chitosan and chitin. Analysis of 13C CP/MAS NMR and FTIR

29

spectra of chitin after cellulase treatment revealed changes in the chitin crystal structure related to

30

rearrangement of inter- and intramolecular H-bonds. The structural changes and decreases in

31

crystallinity allowed solubilization of chitin molecules of high molecular weight and enhanced the

32

dissolution of chitin in alkali by 10–12% compared to untreated chitin.

33

Keywords: chitin; chitosan; cellulases; cellulose-binding domain; crystalline structure

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34 35 1

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1. Introduction

37

The enzymatic modification of polysaccharides has attracted much research attention, as it

38

raises the possibility of using the highly selective reactions performed by naturally designed

39

proteins for practical purposes. In nature, chitin and chitosan are hydrolyzed by specific chitinase

40

and chitosanase enzymes, but proteases, lipases, and a large number of glycosyl hydrolases have

41

been reported to degrade these polysaccharides in a nonspecific manner [1]. Some of these enzymes

42

are even more active than actual chitinases and chitosanases in the hydrolysis of soluble chitosan

43

[2]. This feature opens up many opportunities for wide-scale and cost-effective enzymolysis of

44

chitopolysaccharides. A large number of nonspecific enzymes have been proposed for reducing the degree of

46

polymerization (DP) of chitin and chitosan and for production of chitooligosaccharides (COS) and

47

low molecular weight chitosan (LMWC). LMWC with MW of 5–10 × 103 shows higher biological

48

activity compared to chitosans of higher MWs [3, 4]. The DP of COS fragments varies from 2 to 20

49

units, and each fragment differs in the sequences of deacetylated and acetylated residues. COS have

50

been demonstrated to possess several biological activities, including anti-microbial, anti-oxidant,

51

anti-inflammation, immunostimulatory, anti-tumor, tissue regenerative, and drug and DNA delivery

52

enhancement effects [5]. The mechanisms of these COS actions involve the modulation of several

53

important metabolic pathways. For example, COS have been demonstrated to inhibit the

54

inflammatory responses in animal models [6]. In a cell model, the increase in the degree of

55

deacetylation of COS was correlated with the increase in their anti-inflammatory activities [7]. COS

56

also inhibit the activation of basophils and neutrophils, which are the major granulocytes involved

57

in the acute phase of inflammation [8]. COS induced the death of several cancer cell types in vitro,

58

and the anti-invasive and anti-metastatic activities of COS have also been demonstrated in several

59

types of in vitro and in vivo cancer models [9]. Introducing COS into wound dressings can enhance

60

the hemostatic and inflammatory healing phases to accelerate wound healing and it can promote

61

nerve regeneration [10].

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The use of nonspecific enzymes opens up many opportunities for cost-effective production of

63

COS. Nonspecific enzymes that can hydrolyze chitopolysaccharides include lysozyme, amylases,

64

hyaluronases, pectinases, cellulases, hemicellulases, proteases (pepsin, papain), and lipases. The

65

binding patterns and cleavage abilities of both the specific chitinases/chitosanases and the

66

nonspecific hydrolase enzymes are affected by the number and distribution of acetamide groups in

67

the chitopolysaccharides [1].

68

Achieving high hydrolytic performance remains challenging when using enzymes to degrade

69

insoluble natural polysaccharides like cellulose and chitin that have highly ordered structures. Most

70

studies on enzymatic hydrolysis of polysaccharides have focused on cellulose, which is a 2

71

recalcitrant substrate, and the research has confirmed the importance of cellulose-binding domains

72

(CBDs) on the hydrolytic enzymes for facilitating enzyme action on this insoluble substrate [11,

73

12]. Like cellulose, chitin has highly ordered crystalline structure, which makes it a poorly

74

dissolving polymer in common solvents, thereby limiting its broad application in biomedicine [13].

75

Greater perspectives could be opened up by exploiting either CBD-containing enzymes or separated

76

CBDs, as these domains can modify the highly ordered structures of polysaccharides [14]. From a

77

practical perspective, cellulose-binding modules are more suitable for research than are chitin-

78

binding modules, as they can be obtained in relatively large amounts in the laboratory [15]. Inexpensive commercial enzymes are available for cellulose treatment and usually consist of

80

highly active cellulolytic complexes containing endoglucanases and cellobiohydrolases that are all

81

active toward chitopolysaccharides [16]. Most researchers recommend a focus on the more

82

amorphous chitins that are regenerated from solutions for the effective production of

83

chitooligosaccharides [17]. However, no information is available regarding the effects of enzymatic

84

treatment on chitin crystallinity. The aim of the present work was therefore to evaluate the changes

85

in the crystalline structure of chitin after enzymatic treatment with cellulases. Chitosan hydrolysis

86

with cellulases, lysozyme, amylases, and papain was also analyzed to evaluate the action of

87

nonspecific enzymes on chitopolysaccharides.

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2. Materials and methods

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2.1. Materials

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The crab chitosan (Qingdao Honghai bio-tech Co., LTD, Qingdao City, China) used in this

92

study had a viscosity average molecular weight (Mη) of 1.35×105 and a degree of acetylation (DA)

93

of 0.29, while the shrimp shell chitin (Murmansk, Russia) had a Mη of 2.75×105 and a DA of 0.92–

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1.01. The cellulases were characterized using the following cellulosic substrates: microcrystalline

95

cellulose with particle size below 20 μm (MCC, Sigma Aldrich); carboxymethyl cellulose sodium

96

salt (CMC, Fluka) with viscosity 400–800 mPa·s (4% in water) and degree of substitution 0.70–

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0.85; and filter paper (Whatman No. 1). Nitrophenol-labeled acetyl-β-D-glucosamine (pNP-

98

GlcNAc) and cellobiose (pNP-(Glc)2) and para-hydroxybenzoic acid hydrazide were purchased

99

from Sigma Aldrich.

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100 101 102

2.2. Characterization of chitin and chitosan The Mη values of the chitin samples before and after enzymatic treatment were obtained using .

103

the Mark–Houwink equation η = 0.0024M

104

determined at 25 °C in dimethylacetamide with 5% LiCl using an Ubbelohde viscometer. The Mη

[18]. The intrinsic viscosity [η] of chitin was

3

105

of chitosan was determined similarly, in 2% acetic acid with the addition of 0.3 M NaCl, and

106

calculated from the equation η = 0.00341M

107

.

[19].

The DA of chitin was calculated using both

13

C CP-MAS NMR spectrometry and FTIR

108

spectroscopy. The 13C CP-MAS NMR spectra of chitin were recorded on a Bruker AVANCE II-500

109

WB instrument (Bruker, Billerica, MA, USA). The DA (1.01) was determined from the ratio of the

110

integral intensities of the methyl group (22.9 ppm) and the C1–C6 signals (50–110 ppm) [20]. The

111

FTIR spectra of chitin were obtained using a Vertex-70 FTIR spectrometer (Bruker, Billerica, MA,

112

USA) equipped with a ZnSe-attenuated total reflectance accessory (PIKE Technologies). The DA

113

(0.92) was determined from the ratio of intensities at 1655 and 3450 cm–1 [21]. The DA of chitosan was determined from its 1Н NMR spectrum recorded on a Bruker Avance

115

400 MHz spectrometer (Bruker, Billerica, MA, USA) in D2O/CF3COOH at 70°C using the ratio of

116

the integral intensities of the acetyl group protons (2.08 ppm) and H-1 protons (4.5–5.1 ppm).

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117 2.3. Characterization of enzymes

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The enzymes used in this work and their sources and manufacturers are listed in Table 1.

120

According to electrophoresis data, lysozyme and amylase A were monocomponent enzymes, while

121

cellulases and amylase D contained mixtures of enzymes.

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Table 1 Enzymes used in the study

lysozyme papain

Source

Molecular weight, kDa

chicken egg

14.5

ur

Enzyme

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papaya

23.0

Aspergillus niger

57.3; 44.5; 15.0

cellulase F

Trichoderma reesei

67.8; 59.4; 24.7; 30.2

cellulase C

Trichoderma reesei

67.8; 59.4; 24.7

amylase A

Baсillus liсhеnifоrmis

60.1

amylase D

-

164.4; 83.7; 57.6

cellulase A

Manufacturer

Sigma Aldrich

Novozymes (Fiber Care D) Novozymes (Cellic) Novozymes (Aquazyme) Novozymes (Duozyme)

124 125

The protein content in each enzyme preparation was determined by the Bradford method [22],

126

with measurements performed in a 1.00 cm quartz cuvette at wavelength of 595 nm in a UV 1700

127

spectrophotometer (Shimadzu, Japan). Bovine serum albumin was used as a standard protein for 4

128

calibration. The MW of each enzyme was determined by electrophoresis on 10% SDS-PAGE gels

129

[23]. The gels were photographed with a VE-10 camera (Helicon) and the electropherograms were

130

processed using GelDoc EZ equipment (BioRad) and Image Lab software. Cellulases were characterized by their main activities toward cellulosic substrates using a

132

reducing sugar determination method with picric acid [24] and glucose as a standard. The

133

measurements were performed in a quartz 1.00 cm cuvette at a wavelength of 475 nm in a UV 1700

134

spectrophotometer (Shimadzu, Japan). Activities were determined under equal conditions at 50 °C

135

and pH 5.5 (0.1 M acetate buffer solution), according to the procedures described previously [25].

136

The enzymatic activity toward chitosan was determined under heterogeneous conditions by the

137

action on chitosan dispersed in acetate buffer (pH 5.5). Reducing sugars were determined with the

138

picric acid method using glucosamine as a standard.

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CBD-containing fractions were derived from cellulase F by purification of the enzyme sample

140

on a Sephadex G25 sorbent, followed by affinity chromatography in a stream of acetate buffer (pH

141

4.5) eluent on a column containing microcrystalline cellulose, according to the procedure described

142

previously [26]. The purity and protein component contents were determined by SDS-PAGE

143

electrophoresis. Nitrophenol-labeled acetyl-β-D-glucosamine (pNP-GlcNAc) and cellobiose (pNP-

144

(Glc)2) were used to determine the native specific activities of the fractions of cellulase F. Reducing

145

sugars released by the action of cellulase F fractions on chitosan and cellulose were determined

146

with para-hydroxybenzoic acid hydrazide reagent using glucose as a standard.

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2.4. Enzymatic treatment of chitosan

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Chitosan at 2% concentration in solution or suspension was treated with enzymes at room

150

temperature for 24 h under constant stirring. The pH was adjusted to the enzyme optimum using

151

acetate buffer. Enzyme dosages were 70–100 mg per g of chitosan. At the end of the treatment time,

152

the mixture was heated to 95 °C for 10 min to inactivate the enzyme. The mixture was centrifuged

153

and the supernatant was analyzed for reducing sugars using the picric acid method. The sediment

154

was dialyzed (MWCO 12-14 kDa) for several days against distilled water and freeze-dried. The

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resulting chitosan samples were subsequently used for determination of their MW.

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The decrease in viscosity of chitosan solutions during enzymatic treatment was determined

157

using a Viscolead Pro M rotational viscometer (Fungilab S.A., Spain). A calculated amount of the

158

enzyme, corresponding to a dosage of 50 mg per 1 g of chitosan, was added to the prepared 2%

159

chitosan solution and mixed thoroughly. The solution was placed in a measuring cell of the

160

viscometer and measured at 30 °C.

161 162

2.5. Enzymatic treatment of chitin 5

163

Chitin was treated with enzymes at 40 °C (for lysozyme) and 50 °C (for cellulases) at pH 5.5

164

(0.1 M acetate buffer) for 24 h under constant stirring. The enzyme dosages were 80 mg per g of

165

chitin. At the end of the treatment time, the mixture was heated to 95 °C for 10 min to inactivate the

166

enzyme. The precipitate was separated by centrifugation, washed several times with distilled water,

167

and freeze-dried. The resulting chitin samples were subsequently used to characterize their crystal

168

structure and solubility.

169 170 171

2.6. Structural studies of enzymatically treated chitin The structure of hydrogen bonds in chitin molecules was evaluated by

13

C CP-MAS NMR

spectra, as previously described [20]. In brief, the structure was determined based on the ratio of the

173

signals of the carbonyl carbon atom corresponding to acetamide groups involved in intermolecular

174

hydrogen bonds (173.5 ppm) and the intra- and intermolecular hydrogen bonds (175.8 ppm). Signal

175

decomposition into Lorentzians was performed using the MestReNova software (Mestrelab

176

Research, Barcelona, Spain). The H-bond intensity was evaluated using FTIR spectra according to

177

procedure described in [27].

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The degree of crystallinity of the samples before and after enzymatic treatment was

179

determined using a DRON-3M instrument (Burevestnik, St. Petersburg, Russia) at a radiation

180

wavelength of Cu Kα λ = 1.54 Å. The crystallinity of the chitin samples was calculated as the ratio

181

of the intensity of the (110) reflex to the amorphous halo in the range 2θ = 22° [28].

183

2.7. Evaluation of chitin dissolution in alkali

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The solubility of two fractions of chitin powder (one with a size of 90–200 μm and the other

185

sized under 90 μm) was determined by suspending the powder in an 8% NaOH/4% urea solution

186

and storing at –20 °C for 4 days, with periodic shaking to ensure dissolution. Non-dissolved residue

187

was separated by centrifugation, washed with distilled water, lyophilized, and weighed to determine

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the degree of dissolution.

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The hydrodynamic radii (Rh) of chitin molecules dissolved in alkali were determined using

190

dynamic light scattering (DLS) on a Photocor Compact-Z instrument (Photocor, Moscow, Russia)

191

at a laser wavelength of 654 nm and a detection angle of 90°.

192 193

3. Results and discussion

194

3.1. Chitosan hydrolysis with different enzymes

195

We used chitosan hydrolysis to identify the enzymes with the most activity toward

196

chitopolysaccharides because chitosan is more available to enzymes and can be easily dissolved in

197

dilute acids. These characteristics allow chitosan hydrolysis in both homogeneous (solution) and 6

198

heterogeneous (suspension) conditions. The enzymatic activity toward chitosan was determined

199

under heterogeneous conditions, as all other polymeric substrates were also used in the undissolved

200

state. In this case, the enzyme activity was affected by the ability of the enzymes to bind to the

201

substrate.

202

Cellulases have cellulose-binding domains that provide effective sorption onto insoluble

203

substrates, thereby imparting advantages over the other tested enzymes. We observed that cellulases

204

were more active against chitosan than were amylases, lysozyme, or papain. Previous studies have

205

highlighted a high activity of papain enzyme toward chitosan [29, 30]. Despite the numerous reports

206

[31, 32] available on protease depolymerization of chitosan through the hydrolysis of glycosidic

207

bonds, the mechanism of protease action is still poorly understood. The activities of cellulases toward cellulosic substrates and chitopolysaccharides are

209

presented in Table 2. The activities of the other studied enzymes were significantly lower toward

210

chitosan, and their activities toward chitin were undetectable. In an earlier study, the activity of

211

enzymes toward chitosan (DD 0.91) were determined to decrease the following order: pepsin >

212

chitosanase > α-amylase [33]. However, all the nonspecific enzymes demonstrated significant

213

activity comparable to that of specific chitosanases. Another study [2] reported that chitinase was

214

more active toward chitosan (DD 0.85), with the following order given: chitinase > hemicellulase >

215

lysozyme > lipase. In the present work, activity was measured by both the viscosity decrease and

216

the reducing sugar release, and these two values corresponded well with each other. Activity toward

217

chitin had the same tendency as observed for chitosan.

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Table 2 Activity of cellulases toward different polysaccharides Enzyme

Protein

Activity, U/g

content, mg/g

CMC

MCC

FP

chitosan

chitin

cellulase F

180

325±10

15±2

25±3

60±5

0.30±0.10

cellulase C

180

350±12

20±2

45±5

100±5

0.40±0.09

cellulase A

930

115±9

10±2

30±5

30±5

0.25±0.07

220 221

The commercial cellulases demonstrated different cellulase activities (Table 2). All cellulases

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used were cellulolytic complexes that contain several types of endo- and exo-enzymes acting

223

synergistically.

224

The activity of endo-enzymes leads to a rapid decrease in the degree of polymerization (DP)

225

of cellulose, with a low level of monosaccharide release. The endo-activity is determined by the

226

enzyme action on soluble substrates, e.g. on carboxymethyl cellulose (CMC). By contrast, the

227

activity of exo-enzymes is determined by their effect on microcrystalline cellulose (MCC), since 7

228

they are active in the hydrolysis of recalcitrant crystalline areas. Traditionally, the overall cellulase

229

activity is evaluated using filter paper (FP), while taking into account the synergism possible

230

between the various components of the enzymes in the cellulase mixture. Three cellulases

231

demonstrated different cellulolytic activities: cellulase C appeared to have the highest cellulase

232

activity, showing slightly higher activity toward CMC and MCC and almost 1.6-fold higher general

233

cellulase activity toward filter paper compared to the other tested cellulases. Cellulase C

234

demonstrated a 3-fold higher activity toward chitosan than was observed with cellulase A, and it

235

was 1.5-fold more active than cellulase F. The activity of cellulases toward chitin was significantly

236

lower compared to their activity toward chitosan, which probably reflected the lower accessibility

237

of chitin to the enzymes. All the tested enzymes hydrolyzed chitosan and produced a sharp decrease in the viscosity of

239

the chitosan solutions. Figure 1 shows a representative curve of the viscosity decrease of 1%

240

chitosan solution treated with cellulase F. A rapid decrease occurred during the first 10 min of

241

hydrolysis, followed by a slower continuous decrease during the subsequent 2 h. After 3–4 h, the

242

viscosity remained unchanged. The first 10 min of cellulase treatment resulted in a 50% decrease in

243

the initial solution viscosity, while the maximum viscosity decrease after 4 h was 64%. Thus, the

244

formation of the main part of the LMWC occurred during the first 1–2 h. This enzyme behavior

245

indicates an endo-type action on chitosan. Many published sources have reported that this type of

246

action is typical of nonspecific enzymes [34, 35].

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Viscosity, mPa·s

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140 120 100 80 60 40 20 0

0

247 248

1

2 Hydrolysis time, h

3

4

Fig. 1. The viscosity of 1% chitosan solution treated with cellulase F.

249 250

The reducing sugar release corresponds to the viscosity decrease and reduction of the DP

251

(Table 3.) For 24 h, the reducing sugar release did not exceed 10–13% of the chitosan weight. The

252

low sugar yield also confirms the endo-type action of the nonspecific enzymes. Thus, the cellulase 8

253

treatment for COS and glucosamine production should be longer than that for LMWC production.

254

Chitosan was more effectively hydrolyzed by cellulases and lysozyme than by amylases. Amylase

255

D decreased the DP of chitosan by 1.3-fold, while lysozyme decreased it by 2.1-fold and cellulases

256

by 2.4–3.3-fold.

257 258

Table 3 Chitosan hydrolysis with different enzymes (2% chitosan solution, room temperature, 24 h)

259

*chitosan in suspension; nd – not detected [η],

Мη,

mg/g

g/dL

kDa

no enzyme

-

5.7

135

780

nd

amylase А

100

4.0

-

-

-

nd

amylase D

100

4.0

4.4

103

595

25±4

lysozyme

70

5.0

2.8

378

45±5

cellulase F

70

6.0

cellulase A

100*

5.5

cellulase A

100

5.0

DP

mg/g

of ro 65.4

Reducing sugars,

-p

рН

2.4

56.1

324

85±4

2.2

51.6

298

100±5

1.7

40.6

234

130±4

re

Enzyme dosage,

lP

Enzyme

260

The performance of cellulase A in heterogeneous conditions did not differ much from that

262

observed for homogenous hydrolysis. Additional experiments were conducted to determine the

263

effect of hydrolysis conditions and stirring on the performance of cellulase on chitosan (Fig. 2).

264

Stirring did not significantly affect the efficiency of hydrolysis of chitosan, either in solution or

265

suspension. This may be due to the presence in the cellulase molecules of a CBD that would allow

266

the effective sorption of the enzyme onto the substrate, thereby promoting hydrolysis under

267

heterogeneous conditions. A partial deactivation of cellulases at the interfaces between air bubbles

268

formed with strong stirring has been reported previously [36]. An increase in ionic strength (0.1 M

269

acetate buffer) slightly increased the degree of hydrolysis of the chitosan suspension, whereas an

270

increase in ionic strength was reported previously to decrease the efficiency of cellulose hydrolysis

271

with cellulases [37, 38]. The observed increase in the degree of hydrolysis in our case could be

272

associated with a partial dissolution of chitosan in the presence of acetate buffer.

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9

chitosan in suspension

Reducing sugars, mg/g of chitosan

160 140 120 100 80 60 40 20 0

stirring

stirring, ion strength

no stirring

stirring

of

no stirring

Fig. 2. The influence of stirring and ionic strength on chitosan hydrolysis by cellulase A.

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273 274

chitosan in solution

275

The presence of a CBD in the enzyme structure determines the efficiency of binding to

277

cellulose (or chitin/chitosan, due to their structural similarity) and presumably also determined the

278

ability of the enzyme to disrupt the highly ordered structure of polysaccharides. The electrophoresis

279

results indicated that sorption on cellulose concentrated two components of cellulase F with

280

molecular weights of approximately 3.0×104 and 3.5×104 with high sorption activity . The obtained

281

fractions showed activity toward the p-nitrophenol-labeled chromogenic substrates pNP-GlcNAc

282

and pNP-(Glc)2 (Table 4), indicating chitobiase and cellobiohydrolase activities. The activity was

283

significantly higher for the fraction with high sorption activity than with low sorption activity.

284

However, evaluation of the enzyme activities only with the chromogenic substrates may not be

285

sufficient to determine the effect on the polymer substrate. Hydrolytic reactions carried out on MCC

286

and chitosan substrates also revealed higher activity in the case of the fraction with high sorption

287

activity. Both chitosan and MCC were hydrolyzed efficiently and to a similar extent. Thus, the

288

increasing content of efficiently binding enzymes in a cellulase mixture increased the performance

289

toward cellulosic substrates and toward chitosan.

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290 291

Table 4 The activity of cellulase F fractions separated on cellulose Enzyme fraction High sorption activity Low sorption activity

Activity, U/mL

Protein

pNP-GlcNAc

pNP-(Glc)2

MCC

chitosan

content, mg/L

2.9

40.0

14.1

10.3

1

0.7

12.8

0.5

0.4

30

10

292 293 294

3.2. Enzymatic action on chitin The most active cellulase C and cellulase F were used for enzymatic treatment of chitin. The 13

295

FTIR and

C NMR spectra indicated that the sample of chitin was almost completely acetylated,

296

and the enzymatic treatment with hydrolases did not reduce the DA of chitin. Despite the low

297

degree of chitin hydrolysis after enzymatic treatment (Table 5), the solubility of the treated chitin

298

increased by 10–12% compared with untreated chitin. The chitin solutions were further characterized by DLS. The hydrodynamic radii (Rh) of the

300

unimers and aggregates of chitin macromolecules in an alkaline solution are presented in Table 5.

301

The weight average molecular weight (Mw) of the dissolved chitin is reflected by the Rh of the

302

unimers. The preliminary refinement of chitin facilitated the conversion of chitin molecules with a

303

greater molecular weight into the soluble state, as treatment with cellulase F solubilized the chitin

304

molecules of higher molecular weight. When compared to cellulase C, the highest molecular weight

305

of chitin molecules in solution was achieved with the simultaneous action of both cellulase

306

preparations. The dependences described previously [39] were originally intended for use in

307

determining the Mw of dissolved chitin from its Rh; however, the Mw of chitin in an alkaline

308

solution is overestimated by the DLS method due to the formation of aggregates. This feature was

309

taken into account later [40], and strong dilution was proposed to prevent this aggregation.

310

However, in our case, the formation of aggregates could not be prevented even with a strong

311

dilution of the solutions.

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Table 5 Chitin properties after enzymatic treatment and the hydrodynamic radii (Rh) of chitin

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molecules dissolved in alkali Enzyme

Chitin

Degree of

Crystallinity,

Solubility

Unimer

Aggregate

fraction

hydrolysis,

%

in alkali,

Rh, nm

Rh, nm

size, μm

%

No enzyme

90-200

–

94

25

61±7

270±17

cellulase F

90-200

1.0

91

28

65±6

293±23

cellulase C

90-200

0.4

95

28

55±8

240±13

No enzyme

<90

–

94

31

69±10

352±24

cellulase F +

<90

1.8

89

34

78±11

408±8

%

cellulase C 315 316

FTIR spectra have been used previously to assess the changes in the intensity of cellulose

317

hydrogen bonds after enzymatic treatment [27]. The ratio of the intensity of bands at 3433 and 3258 11

318

cm–1 (corresponding to the vibrations of some intra- and intermolecular bonds) relative to the band

319

at 2900 cm–1 has been proposed as a marker of the changes in cellulose inter- and intramolecular H-

320

bond rearrangements. Cellulose undergoing sorption onto a CBD showed a decrease of 7.6% in the

321

relative intensity of hydrogen bonds, which corresponds to a decrease of 10.6% in the intensity of

322

the intermolecular bonds and of 4.5% in the intensity of the intramolecular bonds [27]. Treatment of

323

chitin (90–200 μm) with cellulases increased the intensity of the bands at 3433 and 3258 cm-1 by

324

8% and 25% relative to the band at 2900 cm-1 under the action of cellulase C and cellulase F,

325

respectively. Therefore, both enzymes induce changes in this region of the IR spectrum, indicating

326

rearrangement of inter- and intramolecular H-bonds. The structural changes in chitin were determined by recording the 13C CP-MAS NMR spectra

328

(Fig. 3a) before and after the enzyme treatment. As discussed previously [20], the peaks at 175.8

329

and 173.5 ppm can be assigned to the carbonyl group involved in double H-bond and to the single

330

H-bond, respectively (Fig. 3c). The decomposition of the carbonyl carbon signal into Lorentzians

331

was carried out using the MestreNova program (Fig. 3b). For the untreated chitin sample, the ratio

332

of the integral intensity of the peak at 175.8 ppm and the summary intensities of both peaks (175.8

333

and 173.5 ppm) was 30%; consequently, double hydrogen bonds in the initial chitin form about

334

30% of the carbonyl groups. Enzyme treatment of chitin increased the proportion of double

335

hydrogen bonds to 33–35%.

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336 12

337

Fig. 3. 13C CP-MAS spectra of chitin (a), decomposition of carbonyl atom signal (b), two types of

338

chitin acetyl group H-bond (c) and comparison of spectra of untreated chitin (C) and enzymatically

339

treated (E) in the region of the signal of a carbonyl group.

340 341

4. Conclusion Overall, among the enzymes studied (lysozyme, papain, cellulases, and amylases), cellulases

343

were the most active in chitosan hydrolysis. One reason why cellulases show the best performance

344

could be their better binding to substrate. The concentration of efficiently binding cellulases

345

significantly increased the hydrolytic activity of the cellulase mixture toward cellulosic substrates

346

and chitosan. The enzymes showing more activity toward chitosan had also the highest activity

347

toward chitin. A 10–12% improvement in chitin dissolution in alkali was achieved by enzymatic

348

treatment. Since the sugar release was low in chitin hydrolysis, the structural changes occurring in

349

chitin after cellulase treatment were not related to the hydrolysis of glycosidic bonds; rather, they

350

reflected a slight rearrangement of inter- and intramolecular H-bonds by the enzymatic action.

351

Further studies should investigate the mechanism of action of nonspecific enzymes on

352

chitopolysaccharides, as well as the role of the degree of deacetylation on the rate of hydrolysis by

353

nonspecific enzymes.

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Funding: This work was financially supported by the Russian Foundation for Basic Research

356

(grants 18-38-00374 and 19-33-60014).

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Acknowledgements: The authors are grateful to E.N. Vlasova for performing the FTIR

359

spectroscopy analysis and to Dr. A.V. Dobrodumov for conducting the NMR studies.

360 361

Conflicts of Interest: The authors declare no conflict of interest. The funders had no role in the

362

design of the study; in the collection, analyses, or interpretation of data; in the writing of the

363

manuscript, or in the decision to publish the results.

364 365

References

366

[1] D.N. Poshina, S.V. Raik, A.N. Poshin, Y.A. Skorik, Polymer Degradation and Stability, 156

367

(2018) 269-278.

368

[2] M. Yalpani, D. Pantaleone, Carbohydrate Research, 256 (1994) 159-175.

369

[3] J. Li, Y. Du, J. Yang, T. Feng, A. Li, P. Chen, Polymer Degradation and Stability, 87 (2005)

370

441-448.

371

[4] S.-K. Kim, N. Rajapakse, Carbohydrate Polymers, 62 (2005) 357-368. 13

[5] F. Liaqat, R. Eltem, Carbohydr Polym, 184 (2018) 243-259.

373

[6] M. Yousef, R. Pichyangkura, S. Soodvilai, V. Chatsudthipong, C. Muanprasat, Pharmacol Res,

374

66 (2012) 66-79.

375

[7] S.H. Lee, M. Senevirathne, C.B. Ahn, S.K. Kim, J.Y. Je, Bioorg Med Chem Lett, 19 (2009)

376

6655-6658.

377

[8] T.-S. Vo, C.-S. Kong, S.-K. Kim, Carbohydrate Polymers, 84 (2011) 649-655.

378

[9] C. Muanprasat, V. Chatsudthipong, Pharmacol Ther, 170 (2017) 80-97.

379

[10] Y. Wang, Y. Zhao, C. Sun, W. Hu, J. Zhao, G. Li, L. Zhang, M. Liu, Y. Liu, F. Ding, Y. Yang,

380

X. Gu, Mol Neurobiol, 53 (2016) 28-39.

381

[11] V.G.H. Eijsink, G. Vaaje-Kolstad, K.M. Vårum, S.J. Horn, Trends in Biotechnology, 26

382

(2008) 228-235.

383

[12] M. Sørlie, S.J. Horn, G. Vaaje-Kolstad, V.G.H. Eijsink, Reactive and Functional Polymers,

384

148 (2020) 104488.

385

[13] C.K.S. Pillai, W. Paul, C.P. Sharma, Progress in Polymer Science, 34 (2009) 641-678.

386

[14] I. Levy, O. Shoseyov, Biotechnology Advances, 20 (2002) 191-213.

387

[15] R. Pinto, J. Carvalho, M. Mota, M. Gama, Cellulose, 13 (2006) 557-569.

388

[16] G. Tegl, C. Öhlknecht, R. Vielnascher, A. Rollett, A. Hofinger-Horvath, P. Kosma, G.M.

389

Guebitz, Biomacromolecules, 17 (2016) 2284-2292.

390

[17] M. Sukwattanasinitt, H. Zhu, H. Sashiwa, S. Aiba, Carbohydrate Research, 337 (2002) 133-

391

137.

392

[18] M. Terbojevich, C. Carraro, A. Cosani, Carbohydrate Research, 180 (1988) 73-86.

393

[19] N.V. Pogodina, G.M. Pavlov, S.V. Bushin, A.B. Mel'nikov, Y.B. Lysenko, L.A. Nud'ga, V.N.

394

Marsheva, G.N. Marchenko, V.N. Tsvetkov, Polymer Science U.S.S.R., 28 (1986) 251-259.

395

[20] T. Kameda, M. Miyazawa, H. Ono, M. Yoshida, Macromolecular Bioscience, 5 (2005) 103-

396

106.

397

[21] A. Baxter, M. Dillon, K.D. Anthony Taylor, G.A.F. Roberts, International Journal of

398

Biological Macromolecules, 14 (1992) 166-169.

399

[22] J.M. Walker, N.J. Kruger, in: Protein Protocols Handbook, The, Humana Press, New Jersey,

400

2002, pp. 15-22.

401

[23] U.K. Laemmli, Nature, 227 (1970) 680-685.

402

[24] M. DuBois, K.A. Gilles, J.K. Hamilton, P.A. Rebers, F. Smith, Analytical Chemistry, 28

403

(1956) 350-356.

404

[25] T.K. Ghose, Pure and Applied Chemistry, 59 (1987) 257-268.

Jo

ur

na

lP

re

-p

ro

of

372

14

[26] A.S. Borisova, E.V. Eneyskaya, K.S. Bobrov, S. Jana, A. Logachev, D.E. Polev, A.L. Lapidus,

406

F.M. Ibatullin, U. Saleem, M. Sandgren, C.M. Payne, A.A. Kulminskaya, J. Ståhlberg, The FEBS

407

journal, 282 (2015) 4515-4537.

408

[27] L. Wang, Y. Zhang, P. Gao, Science in China Series C: Life Sciences, 51 (2008) 620-629.

409

[28] M. Ioelovich, Research and Reviews: Journal of Chemistry, 3 (2014) 7–14.

410

[29] H. Lin, H. Wang, C. Xue, M. Ye, Enzyme and Microbial Technology, 31 (2002) 588-592.

411

[30] R.A.A. Muzzarelli, M. Tomasetti, P. Ilari, Enzyme and Microbial Technology, 16 (1994) 110-

412

114.

413

[31] T. Roncal, A. Oviedo, I. Lopez de Armentia, L. Fernandez, M.C. Villaran, Carbohydr Res, 342

414

(2007) 2750-2756.

415

[32] B.A. Kumar, M.C. Varadaraj, R.N. Tharanathan, Biomacromolecules, 8 (2007) 566-572.

416

[33] B.F.C.A. Gohi, H.-Y. Zeng, A.D. Pan, J. Han, J. Yuan, Polymers, 9 (2017) 174.

417

[34] W. Xia, P. Liu, J. Liu, Bioresource Technology, 99 (2008) 6751-6762.

418

[35] M.B. Kaczmarek, K. Struszczyk-Swita, X. Li, M. Szczęsna-Antczak, M. Daroch, Frontiers in

419

Bioengineering and Biotechnology, 7 (2019) 243.

420

[36] S. Bhagia, C.E. Wyman, R. Kumar, Biotechnology for Biofuels, 12 (2019) 96

421

[37] M. Yu, J. Li, S. Chang, L. Zhang, Y. Mao, T. Cui, Z. Yan, C. Luo, S. Li, Fuel, 175 (2016) 20-

422

25.

423

[38] E. Palmqvist, B. Hahn-Hägerdal, Bioresource Technology, 74 (2000) 25-33.

424

[39] G. Li, Y. Du, Y. Tao, Y. Liu, S. Li, X. Hu, J. Yang, Carbohydrate Polymers, 80 (2010) 970-

425

976.

426

[40] Y. Fang, B. Duan, A. Lu, M. Liu, H. Liu, X. Xu, L. Zhang, Biomacromolecules, 16 (2015)

427

1410-1417.

Jo

ur

na

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re

-p

ro

of

405

428

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-p

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Among the enzymes studied, cellulases were the most active in chitosan hydrolysis. The enzymes more active toward chitosan had also the highest activity toward chitin.

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10–12% improvement in chitin dissolution in alkali was achieved by enzymatic treatment.

Declaration of interests ☒ The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

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☐The authors declare the following financial interests/personal relationships which may be considered as potential competing interests: