Food Chemistry 150 (2014) 392–399

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Food Chemistry journal homepage: www.elsevier.com/locate/foodchem

Review

Recent developments in chitosanase research and its biotechnological applications: A review Nidheesh Thadathil, Suresh Puthanveetil Velappan ⇑ Academy of Scientific and Innovative Research, CSIR-Central Food Technological Research Institute, Mysore 570020, India Department of Meat and Marine Sciences, CSIR-Central Food Technological Research Institute, Mysore 570020, India

a r t i c l e

i n f o

Article history: Received 9 July 2013 Received in revised form 1 October 2013 Accepted 4 October 2013 Available online 26 October 2013 Keywords: Chitosanase Chitosan N-Acetyl-D-glucosamine D-Glucosamine Chitosan oligosaccharides

a b s t r a c t Chitosanases (EC 3.2.1.132) are glycosyl hydrolases that catalyse the endohydrolysis of b-1,4-glycosidic bonds of partially acetylated chitosan to release chitosan oligosaccharides (COS). Chitosanases are isolated, purified and characterised from different sources mainly from bacteria and fungi. Chitosanases have received much attention due to their wide range of applications including the preparation of bioactive COS and fungal protoplasts, as biocontrol agent against pathogenic fungi and insects, the bioconversion of chitinous bio waste associated with seafood processing, etc. Bioactive COS produced by the enzymatic hydrolysis of chitosan have finds numerous health benefits as well as other biological activities. This review summarizes the recent advances in chitosanases research, the enzyme production processes, characterization, genetic improvement and their applications. Ó 2013 Elsevier Ltd. All rights reserved.

Contents 1. 2.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Occurrence and distribution of chitosanases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Bacterial chitosanase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Fungal chitosanases. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. Cyanobacterial chitosanase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Plant chitosanases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Definition and classification of chitosanases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Fermentative production of chitosanase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Measurement of chitosanase activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6. Purification of microbial chitosanases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7. Basic biochemical properties of chitosanases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1. Molecular mass . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2. pH and temperature optima . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.3. Enzymatic hydrolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4. Inducers of chitosanase synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.5. Effect of metal ions on chitosanase activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.6. Enzyme structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8. Immobilised chitosanase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9. Cloning and genetic improvement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10. Biological roles of chitosanase. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Potential applications. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12. Futuristic considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

393 393 393 393 394 394 394 394 395 395 395 395 396 396 396 397 397 397 397 398 398 399 399 399

⇑ Corresponding author at: Department of Meat and Marine Sciences, CSIR-Central Food Technological Research Institute, Mysore, 570020, India. Tel.: +91 821 2514840; fax: +91 821 2517233. E-mail addresses: [email protected] (N. Thadathil), [email protected], [email protected] (S.P. Velappan). 0308-8146/$ - see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.foodchem.2013.10.083

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1. Introduction Chitosan is a poly cationic natural polymer with a unique structure and functional properties. It is an unbranched copolymer consisting of b-(1?4)-2-acetamido-D-glucose (N-acetyl-D-glucosamine, GlcNAc) and b-(1?4)-2-amino-D-glucose (D-glucosamine, GlcN) units with the latter usually exceeding 80% and the GlcNAc and GlcN residues are randomly distributed and not blocked together (Zitouni et al., 2012) (Fig. 1). Chitosan is found in nature as a structural component mainly in the cell wall of Zygomycetes fungi and also found in the cell wall of Chlorophycean algae Chlorella sp. and in insect cuticle (Hsu, Chung, Chang, & Sung, 2012). Chitosan is the principal N-deacetylated derivative of chitin (a b-1,4 linked polymer of GlcNAc), although the N-deacetylation is almost never complete. Currently chitosan is produced commercially from crustacean’s (shrimp and crab) chitin by N-deacetylation, to different degree using hot concentrated alkali. Thus, there is no clear distinction between chitin and chitosan based on the degree of N-deacetylation; the term chitosan represents a collective name for a family of deacetylated chitins at different degrees (Fig. 1). The studies of chitosan with respect to their preparation, structure, functional properties and applications have been extensively reviewed (Zitouni et al., 2012). Chitosan is usually susceptible to a number of enzymes; these include specific (chitosanases) and non-specific (carbohydrases, proteases, lipases etc.) chitosan hydrolysing enzymes. Chitosanases have been generally recognised as enzymes that specifically hydrolyse chitosan but not chitin (Kim & Rajapakse, 2005). In 2004, the Enzyme nomenclature committee had defined chitosanases (EC 3.2.1.132, Chitosan N-acetylglucosaminohydrolase) as the enzyme capable of performing endohydrolysis of b-1,4-linkages between GlcN residues in a partly acetylated chitosan, from the reducing end (Fig. 2a). Later, in 2008, the committee created a new (second) class of enzyme, exo-b-D-glucosaminidase (EC 3.2.1.165) that attack chitosan from its non reducing end (Fig. 2b). In addition, some other non specific enzymes such as common carbohydrases, proteases and lipases have also shown their hydrolytic ability on chitosan (Kim & Rajapakse, 2005). As on date a large number of specific chitosanolytic enzymes have been reported from different microorganisms including bacteria, fungi and cyanobacteria and plants. Recent researches on chitosanases have received much attention due to their wide range of applications in various fields. Practical applications of chitosanase include the preparation of bioactive COSs (Kim & Rajapakse, 2005; Ming, Kuroiwa, Ichikawa, Sato, & Mukataka, 2006), preparation of fungal protoplasts particularly for Zygomycetes, a biocontrol agent to increase the resistance of plants against pathogenic fungi (Hsu et al., 2012),

Fig. 2. Mode of action of chitosanase (a) and exo-b-D-glucosaminidase (b).

chitosan mediated gene delivery and the bioconversion of marine crustacean chitinous bio waste (Wang, Chao, Liang, & Chen, 2009; Wang, Tseng, & Liang, 2011). Chitosanase mediated hydrolysis has advantages over the chemical/physical mediated hydrolytic production of COS, in which chitosanases can catalyse the hydrolysis under mild reaction conditions and do not produce monosaccharides (Kim & Rajapakse, 2005; Ming et al., 2006). Reviews on chitosanses with respect to its production and application (Somashekar & Joseph, 1996) have been reported earlier. The present review will focus not only on the current knowledge of chitosanases, their occurrence and classification but also on microbial production, biochemical properties, genetic improvement and applications based on research specifically carried out in the last ten years.

2. Occurrence and distribution of chitosanases Chitosanase is produced by microbes and plants, where they play an important role in nutrition and defence. Chitosanase was first described in 1973 from different soil microorganisms. Over the past 40 years, several research papers have been published on the occurrence, production, purification and characterization of chitosanase from different microorganisms including bacteria, fungi and cyanobacteria; and plants. Detailed literature is available on various microbial sources of chitosanase (Somashekar & Joseph, 1996). 2.1. Bacterial chitosanase Extracellular chitosanases has been reported from several bacteria including Bacillus sp. (Gao, Ju, Jung, & Park, 2008; Wang et al., 2009; Zitouni et al., 2012), Serratia sp. (Wang, Peng, Liang, & Liu, 2008b), Janthinobacterium sp. (Johnsen, Hansen, & Stougaard, 2010), Paenibacillus sp. (Zitouni et al., 2012), Acinetobacter sp. (Wang et al., 2011) and Streptomyces sp. (Jiang, Chen, Chen, Yang, & Zou, 2012). Among these bacteria, species of Bacillus and Streptomyces were studied most extensively as a chitosanase source even up to structural and molecular level. 2.2. Fungal chitosanases

Fig. 1. Chemical structure of chitosan (a). Distribution of GlcNAc and GlcN in chitin (b), partially N-deacetylated chitosan (c) and fully N-deacetylated chitosan (d).

As compared to bacteria, there have been few reports on chitosanases from fungi. Chitosanase production has been reported in different fungi including Aspergillus sp. (Zhang, Sang, & Zhang, 2012), Gongronella sp. (Wang, Zhou, Yuan, & Wang, 2008a; Zhou,

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Yuan, Wang, & Yao, 2008), Trichoderma sp. (da Silva, Honorato, Franco, & Rodrigues, 2012) etc. 2.3. Cyanobacterial chitosanase Cyanobacteria constitute an extremely diverse group of photosynthetic prokaryotes exhibiting variability in physiological, morphological and developmental characteristics. In a recent report, an antifungal chitosanase from Anabaena fertilissima RPAN1 was characterised as a prelude to its use as a biocontrol agent (Gupta, Prasanna, Srivastava, & Sharma, 2012). 2.4. 4 Plant chitosanases Chitosanases are mostly found in microorganisms (bacteria and fungi), however, very few of them have been reported in plants. Chitosanase activity has been detected in vesicular–arbuscular mycorrhizal colonised leek and onion root as well as chemical or pathogen stressed leaves of various plant species. In addition, chitosanase activity has been detected in different parts of healthy plants. Chitosan and COS can induce defence reactions in plants including the induction of chitosanase and 1,3-glucanase isoforms. The endogenous function of plant chitosanases, however, has not yet been elucidated (Hsu et al., 2012). Recently Hsu et al. (2012) reported two thermally stable chitosanase isoforms from the sheaths of chitosan treated bamboo shoots. 3. Definition and classification of chitosanases Chitosanases have been generally recognised as enzymes that specifically attack chitosan but not chitin. In addition, some non specific enzymes such as common carbohydrases, proteases and lipases also have shown their hydrolytic ability on chitosan (Gupta et al., 2012; Kim & Rajapakse, 2005). The definition of chitosanase (EC 3.2.1.132, Chitosan N-acetyl glucosaminohydrolase) created in 1990 by the Enzyme Nomenclature Commission and has been amended in 2004 (http://www.chem.qmul.ac.uk/iubmb/enzyme/). Chitosanase is an enzyme which endohydrolyse the b-1,4-linkages

between GlcN residues in a partly acetylated chitosan. It acts on the reducing end of chitosan molecule and produces COS exclusively as the end product (Fig. 2a). They can hydrolyse various types of links in the chitosan molecule (Kim & Rajapakse, 2005). Based on the specificity of the cleavage positions for the partially acetylated chitosan, chitosanases are classified into three distinct subclasses viz. Subclass I chitosanases, Subclass II chitosanases and Subclass III chitosanases. Subclass I chitosanases split both GlcN–GlcN and GlcNAc–GlcN linkages. Subclass II chitosanases can cleave only GlcN–GlcN linkages. On the other hand, subclass III chitosanases split both GlcN–GlcN and GlcN–GlcNAc linkages (Fig. 3). Endohydrolysis of GlcN–GlcN links in chitosan is the common and the only constant property to all the known chitosanases. Similarly, they also do not recognise GlcNAc–GlcNAc links in partly acetylated chitosan. This selectivity at the cleavage position of the substrates might be controlled by rigid substrate recognition by the chitosanases belonging to the different subclasses. The specificity of chitosanases with respect to the cleavage of four different glycosidic linkages in partially N-deacetylated chitosan is determined by the identity of the reducing and non-reducing ends and degree of deacetylation (DD) of chitosan (Kim & Rajapakse, 2005). The difference between chitinase (EC 3.2.1.14) and chitosanase is narrow, both are chitosan hydrolysing enzymes, can act on different degrees of deacetylated chitosan, but chitosanase prefers highly deacetylated chitosan, whereas chitinase prefer highly acetylated chitosan (Somashekar & Joseph, 1996). Chitosanases belong to five glycoside hydrolase (GH) families: GH-5, GH-8, GH-46, GH-75, and GH-80, based on their amino acid sequences. Among these families, GH-46 chitosanases, especially those from Bacillus and Streptomyces have been extensively studied in terms of their catalytic features, enzymatic mechanisms and protein structures. GH-75 chitosanases are another major chitosanase family; members belonging to this family are mainly of fungal origin (Wang et al., 2008a). New types of enzyme exo-b-D-glucosaminidase, which attack chitosan and/ or COS from the non-reducing termini to remove successive GlcN residues, have been reported from different microorganisms (Fukamizo, Fleury, Côté, Mitsutomi, & Brzezinski, 2006). In 2008, the Enzyme Nomenclature Committee created a new enzyme class (EC 3.2.1.165, chitosan exo-(1?4)-b-D-glucosaminidase) for exo-b-D-glucosaminidase (Fig. 2b). This enzyme is also known as exochitosanase and GlcNase. These exo-hydrolytic enzymes can degrade GlcN–GlcNAc but not GlcNAc–GlcNAc linkages (Fukamizo et al., 2006).

4. Fermentative production of chitosanase

Fig. 3. Subclasses of chitosanase and cleavage sites.

Table 1 presents data available in the literature regarding the microbial production of extracellular chitosanses, using both submerged fermentation (SmF) and solid-state fermentation (SSF) systems. Most of the chitosanases reported so far have been inducible. The use of colloidal chitosan as a supplement for culture media in production of chitosanses is a common strategy and in the majority

Table 1 Production of microbial chitosanases. Organism

Activity

Production method

Reference

Aspergillus sp. QD-2 Bacillus sp. TKU004 Bacillus cereus D-11 Gongronella sp. JG Serratia marcescens TKU011 Recombinant chitosanase Recombinant chitosanase Trichoderma koningii

85.816 U/ml 0.14–0.16 U/ml 4.85 U/ml 800 lmol/min/l 0.024 U/ml 140 U/ml 6.2 g/l 4.84 IU/gds

SmF SmF SmF SmF SmF SmF SmF SSF

Zhang et al. (2012) Wang et al. (2009) Gao et al. (2008) Zhou et al. (2008) Wang et al. (2008b) Liu et al. (2009) Kilani-Feki et al. (2012) da Silva et al. (2012)

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of cases, its inductor effect has been established. Other substrates such as chitosan powder, squid pen and shrimp shell for SmF culture media have also been reported as inducer as well as sole carbon/nitrogen sources (Wang et al., 2009; Wang et al., 2011; Zhou et al., 2008). Optimization of various parameters and manipulation of media is one of the most important techniques used for the overproduction of enzymes in large quantities to meet its commercial requirement. Culture media formulation and physicochemical bioprocess conditions are also reported as significant for the enhancement of chitosanase production (Zhang et al., 2012). To optimise the production condition such as pH, temperature, carbon and nitrogen source, metal ions etc. classical one variable at a time experiments was employed by Zhou et al. (2008). These multiple growth parameter poses a challenge to understating and controlling the out come of the fermentation process. Response Surface methodology (RSM) is a very useful statistical technique for optimization of complex chemical, biochemical and food process and has received much attention in the investigation of process optimization for the production of microbial enzymes (Suresh, 2012). The statistical optimization of chitosanase production by Bacillus sp. RKY3 and Aspergillus sp. QD-2 (Zhang et al., 2012) using SmF have been previously reported. The results observed based on statistical experimental designs showed improvement of chitosanase activity (U/ ml) from 26.5 to 85.8 (Zhang et al., 2012). SSF is generally defined as the growth of microorganisms on moist solid substrates in the absence of free water. The solid substrates provide both support and nutrition. SSF constitutes an interesting alternative to SmF since the metabolites so produced are concentrated and purification procedures are less costly. SSF technique is generally confined to the processes involving fungi. Recently, da Silva et al. (2012) reported the production and optimization of extracellular chitosanase by an entomopathogenic fungus Trichoderma koningi under SSF. The maximum chitosanase yield of 4.84 U/gram dry substrate (gds) was achieved using a mixture of 3.0 g of wheat bran and 1.5 g of chitosan as solid substrate. This study showed the feasibility of SSF for the microbial chitosanase production.

5. Measurement of chitosanase activity Most of the chitosanases cannot act on solid/crystalline chitosan, so colloidal chitosan or glycol chitosan are generally used as substrates for the chitosanase assays (Somashekar & Joseph, 1996). Chitosanase activity is influenced by various factors such as the degree of deacetylation of chitosan, viscosity and concentration of chitosan solution, kind of acid used for dissolving the chitosan, the amount of enzyme and other reaction conditions such as pH, temperature, and agitation. Chitosanase activity can be determined by measuring the reduction in viscosity of the substrate, though it is an insensitive method (Somashekar & Joseph, 1996). Quantitative estimation of reducing sugars or amino sugar produced from chitosan substrates is the commonly used method to determine most chitosanase activity. Fukamizo et al. (2006) reported, quantitative chitosanase assay by high-pressure liquid chromatography (HPLC) separation with refractometric detection using COS as substrate. Based on the time-course of enzymatic hydrolysis of the COS substrates, the mode of action, the turnover number and the subsite affinities of the enzymes were estimated. However, high quantities of substrate are usually needed to obtain reliable time-course data, due to low sensitivity of refractometric detection. The detection of chitosanase-catalysed hydrolysis of COSs (GlcNn, n = 2–6) by realtime electrospray ionisation-mass spectrometry (ESI-MS) has reported by Dennhart, Fukamizo, Brzezinski, Lacombe-Harvey, and

395

Letzel (2008). The enzymatic reaction mixture was analysed using a continuous-flow injection system coupled with ESI-MS. They analysed the enzymatic reactions of an exo-glucosaminidase/ exochitosanase from Amycolatopsis orientalis and endochitosanase from Streptomyces sp. N174 with this analytical technique. The analytical approach of real-time MS is an enormous time-saving method. Furthermore, the high sensitivity of the mass spectrometric detection allows the determination of the reaction time-courses with very low quantities of substrate. Chitosanase activity can also be detected in gel after electrophoresis by zymography (chitosanase activity) staining method by incorporating an assay substrate (glycol chitosan) into the gel during casting. After electrophoresis, the chitosanase activity band can be visualised by staining gel with 0.1% Congo red or florescent brightener. 6. Purification of microbial chitosanases Industrial enzymes usually prepared in crude form for their commercial applications. The use of chitosanases in COS preparation usually does not require purification of the enzyme, but chitosanses in their purified form is required to study the biochemical properties, structure–function relationships and their biotechnological applications. The purification of chitosanases from microbial sources in most cases has involved classical enzyme purification methods. These methods involve removal of microbial biomass from the culture broth, selective precipitation/concentration by (NH4)2SO4 or solvents or polyethylene glycol. The concentrated chitosanase is further subjected to chromatography, commonly gel filtration, ion exchange, affinity techniques etc. The purification and characterisation of chitosanses from culture filtrates of different bacteria (Zitouni et al., 2012) and fungi (Zhou et al., 2008) have been reported. Wang et al. (2008b) have purified a chitosanase from culture supernatant using shrimp shell waste as the sole carbon/nitrogen source. The purification of Bacillus subtilis TKU002 chitosanase includes (NH4)2SO4 precipitation, DAAESepharose chromatography, a second stage of (NH4)2SO4 precipitation and the final purification by Sephacryl S-100 chromatography. A 17.2-fold purification with 48.0% activity recovery was achieved (Wang et al., 2009). Recently, Zitouni et al. (2012) purified a thermostable chitosanase from the culture filtrate of Paenibacillus sp. 1794 through a series of steps involving solvent (ice cold acetone) precipitation, Q-Sepharose FF ion exchange chromatography, concentration using Hi-Trap Q (GE Healthcare) mini-column, hydroxyapatite chromatography and finally dialysis; achieved a specific activity of 122 U/mg of protein in the purified chitosanase. 7. Basic biochemical properties of chitosanases The basic biochemical properties of chitosanases vary depending on the source of enzymes. The biochemical properties of chitosanases from different microorganisms have been thoroughly studied and reported. Table 2 represents the data available in the literature regarding the basic properties of chitosanses from various microorganisms. 7.1. Molecular mass Most chitosanases have a low apparent molecular mass within the range of 20–75 kDa (Table 2). The molecular mass of chitosanase from Aspergillus fumigatus KH-94 is 108 kDa, the highest among the known chitosanases. Chitosanase also exists in multiple isoforms. Two isoforms of chitosanases with 25 kDa and 33 kDa chitosanase isoforms from B. subtilis TKU007 (Wang et al., 2009) and 30 kDa and 31 kDa chitosanase isoforms from Penicillium

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Table 2 The biochemical properties of chitosanases. Organism

pH optima

Tem. optima (°C)

Molecular mass (kDa)

Isoforms

km

Reference

Bambusa oldhami Paenibacillus sp. 1794 Aspergillus QD-2 Anabaena fertilissima Streptomyces roseolus Acinetobacter calcoaceticus TKU024 Acinetobacter calcoaceticus TKU024 Janthinobacterium sp. 4239 Gongronella sp. JG Serratia marcescens TKU011 Bacillus cereus D-11

3–4 4.8 5.6 7.5 5 6 7 5–7 5.6 5 6

70, 60 80–85 55 27 50 50 60 45 55–60 50 60

24.5, 16.4 40 ND ND 41 27 66 33 28 21 41

A, B – – – – CHSA1 CHSA2 – – – –

0.539 ND ND 0.89 ND ND ND ND 8.86 ND 7.5

Hsu et al. (2012) Zitouni et al. (2012) Zhang et al. (2012) Gupta et al. (2012) Jiang et al. (2012) Wang et al. (2011) Wang et al. (2011) Johnsen et al. (2010) Wang et al. (2008a) Wang et al. (2008b) Gao et al. (2008)

chrysogenum AS51D (Zitouni et al., 2012) have been described in earlier reports. 7.2. pH and temperature optima The optimum pH of microbial chitosanases is found to be in the range of 4 to 8 (Table 2). Chitosanase from Gongronella sp. JG (Wang et al., 2008a) and Aspergillus sp. QD-2 (Zhang et al., 2012) have been reported to show acidic pH optima of 5.6. Chitosanase from Janthinobacterium sp. 4239 have pH optima in the acidic to neutral (5–7) range (Johnsen et al., 2010). An alkaline chitosanase with pH optima of 7.5 was observed in A. fertilissima (Gupta et al., 2012). The optimum temperature for the activity of chitosanase is associated with the growth of the microorganism and found to be in the range of 30–60 °C (Table 2). A cold active chitosanase, showing 30–70% of activity at 10 and 30 °C from Janthinobacterium sp. 4239 was reported (Johnsen et al., 2010). Most of the reported chitosanases are found to be thermolabile in nature; however, few reports are available on thermostable chitosanases. It is well known that thermostable enzymes have several advantages in industrial application because high temperature can accelerate reactions; decrease the viscosity of the liquid and increase the solubility of raw material as well as decrease microbial contamination. Recently Zitouni et al. (2012) reported a thermostable chitosanase (Csn) from Paenibacillus sp. 1794 isolated from shrimp shell and peat-based compost. Chen et al. (2012) reported a thermostable chitosanase with half-life of 205 h at 80 °C, 1 h at 90 °C and 32 min at 100 °C from Pichia pastoris GS115 containing endochitosanase gene of A. fumigatus. Thermostable chitosanase has an advantage in enzymatic hydrolysis at higher temperatures thus allowing chitosan to be dissolved at higher concentrations (Zitouni et al., 2012).

to attack the anomeric carbon of the glucosamine residue in substrate (Fukamizo et al., 2000). In the five GH families containing chitosanase, the retaining configuration of the catalytic mechanism of GH-5 is derived from glucanase. The inverting configurations have been verified experimentally for GH-8 and GH-46. GH-80 is also inferred to be an inverting enzyme. However, the catalytic stereochemistry of GH-75 enzyme remains unknown (Fukamizo et al., 2000).

7.4. Inducers of chitosanase synthesis Microbial chitosanases have been reported as inducible enzymes mainly by colloidal chitosan. The use of colloidal chitosan as a supplement for culture media in chitosanses production is a common strategy and, in the majority of cases, its inducing effect has established. Microbial chitosanase is also induced by different other substrates like chitosan powder (Zitouni et al., 2012), chitin (Jiang et al., 2012), squid pen powder (Wang et al., 2011) and shrimp shell powder (Wang et al., 2008b). However, microorganisms that produce chitosanases without chitosan as an inducer have technical advantages during fermentation because chitosan has high viscosity and reacts with other compounds in the medium

7.3. Enzymatic hydrolysis Based on the anomeric configuration of the C1 proton of the reducing end sugar obtained from the enzymatic products, chitosanases from different families may involve in different types of catalysis: either the retaining (Fig. 4a) or the inverting (Fig. 4b) mechanism. The retaining glycosidases catalyse the hydrolysis via a two-step, double-displacement mechanism with one of the two essential amino acid residues functioning as a nucleophile and the other as a general acid/base. In contrast, the inverting glycosidases follow a one-step, single displacement mechanism with the assistance of a general acid and a general base. The general base polarises a water molecule to develop a stronger nucleophile for attacking the anomeric carbon, whereas the general acid protonates the glycosidic oxygen to accelerate the reaction. In this particular enzyme, Glu22 was found to act as a proton donor, in cooperation with Asp40, thereby activating the water molecule

Fig. 4. The two major mechanisms of enzymatic glycosidic bond hydrolysis by chitosanases. Retaining mechanism (a) in which the glycosidic oxygen is protonated by Glu22-H and nucleophilic assistance to aglycon departure is provided by Asp40 (vice versa), and inverting mechanism (b) in which protonation of the glycosidic oxygen and aglycon departure are accompanied by a concomitant attack of a water molecule that is activated by general base residue B .

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N. Thadathil, S.P. Velappan / Food Chemistry 150 (2014) 392–399 Table 3 Effect of different metal ions on the activity of chitosanases. Source of enzyme Bambusa oldhami Streptomyces roseolus Anabaena fertilissima Acinetobacter calcoaceticus TKU024 Bacillus sp. TKU004 Gongronella sp. JG Serratia marcescens TKU011

Inhibition +

Ag Cu2+, Co2+, Mn2+, Zn2+ Ag+, Hg2+, Fe3+ Mn2+, Zn2+, Cu2+ Zn2+ – Mn2+, Fe2+

Fig. 5. Overall structure of chitosanase from Streptomyces sp. N174 (a) and Bacillus circulans MH-K1 (b) chitosanases are illustrated as ribbon diagrams (Saito et al., 1999).

during heat sterilization. Constitutive chitosanase from Bacillus sp. MET 1299 has also been reported. 7.5. Effect of metal ions on chitosanase activity The effect of different metal ions on the activity of different chitosanases is given in Table 3. Chitosanase enzyme activity was influenced by different metal ions like Mg2+, Cu2+, Ca2+, Zn2+, Mn2+, Ba2+, Fe2+, Hg2+, Co2+, and Ag+. Cu2+ and Mn2+ could be either an activator or an inhibitor to different chitosanases. Chitosanase inhibition by Hg2+ is a general characteristic of the chitosanase group. Like Hg2+, Ag+ also reported to decrease the activity of chitosanase (Wang et al., 2008b). 7.6. Enzyme structure Reports are not available on the structure of chitosanase during the last decade. We indent to describe the molecular structure of chitosanase from Streptomyces sp. N174 (Fig. 5a) and Bacillus circulans MH-K1 (Fig. 5b) whose molecular structures were extensively studied (Saito et al., 1999). Marcotte, Monzingo, Ernst, Brzezinski, and Robertus (1996) first described about the X-ray structure of an anti-fungal chitosanase from Streptomyces sp. N174. This chitosanase is dumbbell shaped and 55 Å long containing two globular domains connected through a bent 27-residue backbone helix. The domain forms a pronounced cleft, 10 Å wide and 12 Å deep, spanning the width of the molecule between the two domains. Secondary structure reveals the enzyme contains 10 a-helices and three b-sheets, a-helices accounts around 64% of the structure. The cleft is bordered on one face by a three stranded b-sheet, The surface of chitosanase is dominated by the electronegative substrate binding cleft, appropriate for binding a positively charged substrate, but the analogous cleft of chitinase is considerably neutral. Chitosan binding with the active site suggests that, substrate binding and catalytic mechanisms may be similar to other glycohydrolases. Six sugar binding sites (A–F) were identified in chitosanase, with

Activation +

+

Reference 2+

K , Na , Cu Mg2+ Cu2+, Zn2+ – – Mn2+, Ca2+, Sr2+ –

Hsu et al. (2012) Jiang et al. (2012) Gupta et al. (2012) Wang et al. (2011) Wang et al. (2009) Wang et al. (2008a) Wang et al. 2008b

cleavage probably occurring at the glycosidic bond between D and E sugars. The chitosanase preference for chitosan is mediated by bonding to the amines of sugar bounds in sites A–D. The X-ray structure also suggested that Glu 22 and Asp 40 are important for the catalytic function. Chitosanase from Bacillus circulans MH-K1 (Saito et al., 1999) is a 29-kDa extracellular protein composed of 259 amino acids. The overall molecular structure shows 14 a helices and 5 b strands. The enzyme has two globular upper and lower domains, which generate the active site cleft for the substrate binding. The overall molecular folding is similar to chitosanase from Streptomyces sp. N174, although there is only 20% identity at the amino acid sequence level between both chitosanases. However, there are three regions in which the topology is remarkably different. In addition, the disulfide bridge between Cys50 and Cys124 joins the b1 strand and the a7 helix, which is not conserved among other chitosanases. The orientation of two backbone helices, which connect the two domains, is also different and is responsible for the differences in size and shape of the active site cleft in these two chitosanases. This structural difference in the active site cleft is the reason why the enzymes specifically recognises different substrates and catalyse different types of chitosan degradation (Saito et al., 1999). 8. Immobilised chitosanase Immobilisation of enzymes is one of the method for protecting and stabilizing the enzymes, thereby enhancing their properties and their repetitive utilisation either in batch, or continuous mode. Immobilisation of enzymes prevents their deactivation by various physical and chemical denaturing agents and thereby enhancing their operational stability. Furthermore, utilisation of immobilised enzymes permits control of the progress of the hydrolysis reaction (Ming et al., 2006). Particularly in the case of chitosanase, immobilization is highly beneficial from the viewpoint of operating cost because chitosanase is expensive. Ming et al. (2006) has reported the immobilization of chitosanases by direct immobilisation on an agar gel-coated multidisc impeller The main applications of immobilized chitosanase are in the use of COS preparation, and a high yield of COS can be produced because the immobilised enzyme can be removed from the reaction mixture when the yield of the target COS reaches a maximum (Ming et al., 2006). However, long-term uses for stable immobilized chitosanase have yet to be developed using high active chitosanase by cost effective techniques. 9. Cloning and genetic improvement Cloning of genes has been carried out extensively for the molecular study of proteins, hyper production and protein engineering. Cloning of chitosanase genes has been carried for studying their sequence, characteristics, hyper production and expression, change in enzyme induction pattern and for enzyme engineering. Chitosanase genes from different bacteria and fungi have been cloned and expressed mainly in Escherichia coli or in heterologous host

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Pichia pistoris. Many chitosanase genes have been cloned from different bacteria including Bacillus sp. S65 (Kang, Chen, Fu, & Ma, 2012) and Pseudomonas sp. OUC1 (Liu, Li, Zhou, Chi, & Madzak, 2012). Recently, a chitosanase gene (chi) from the marine bacterium Pseudomonas sp. OUC1 was cloned, and its over-expression in Yarrowia lipolytica was reported (Liu et al., 2012). The transformants obtained could secrete much more chitosanase (9.0 U/ml) than non transformed cells (4.3 U/ml). A bacillus-derived GH8 family chitosanase with a 6xHis tag fused at its N-terminal was expressed in the E. coli strain BL21 (DE3) as a soluble and active form (Liu et al., 2009). Its expression level could be as high as 500 mg/l. Enzymatic activity could reach approximately 140 U/ml under assay conditions. The enzyme could efficiently convert chitosan into monomer-free COS: 1 g of enzyme could hydrolyze about 100 kg of chitosan. Heterologous expression and secretion of a thermostable and antifungal B. subtilis chitosanase (CSNV26) in E. coli has been reported (Kilani-Feki, Frikha, Zouari, & Jaoua, 2012). The gene csnV26 encoding for chitosanase was amplified and cloned in the pBAD vector then expressed in E. coli (Top10). The SDS–PAGE and zymogram analysis of the recombinant protein showed that it has two active forms sized 27 and 31 kDa, corresponding to the protein with and without signal peptide. This protein gets secreted by Top10-pBAD-csnV26 with a high yield of 6.2 g/l. Fungal chitosanases are less reported as compared to bacterial chitosanases in genetic improvement due to the inaccessibility of eukaryotic genes. The chitosanase gene (csn) from Fusarium solani 0114, is expressed in an industrial strain Saccharomyces cerevisiae, for the large-scale production of this enzyme was reported (Liu & Bao, 2009). Chen et al. (2012) has reported a thermostable chitosanase with half-lives of 205 h at 80 °C, 1 h at 90 °C and 32 min at 100 °C in Pichia pastoris GS115 containing endochitosanase gene from A. fumigatus. A chitosanase gene, csn, was cloned from Penicillium sp. D-1 by inverse PCR. The cDNA sequence analysis revealed that csn had no intron. The deduced CSN protein consists of 250 amino acids including a 20-amino acid signal peptide and shared 83.6% identity with the GH 75 chitosanase from Talaromyces stipitatus B8M2R4. The mature protein was over expressed in E. coli (Zhu et al., 2012). In addition to gene cloning to achieve over production of chitosanase from microorganisms, cloning of chitosanase gene in plants was also reported (Kouzai et al., 2012). Cloning and expression of chitosanase gene in rice plants have increased resistance against pathogenic fungi (Kouzai et al., 2012). 10. Biological roles of chitosanase Microbial chitosanases with different biological roles have been found in nature. Chitosan degrading microorganisms are widely distributed in nature and microorganisms secrete chitosanase extracellularly to degrade chitosan for their nutritional purpose (Somashekar & Joseph, 1996). Chitosanase together with chitinase, chitin deacetylase and glucosaminidase, involve in the decomposition and recycling of enormous quantity of crustaceans shell produced in nature. Since chitosan is the major structural component in cell wall of Zygomycetes fungi, chitosanases contributed significantly in the determination of the degradation of chitosan and morphogenesis of this class of fungi. In addition to this, plants synthesise chitosanase as a defensive mechanism against phytopathogens especially against Zygomycetes (Hsu et al., 2012). 11. Potential applications Chitosanases have industrial, as well as biotechnological applications that require different types of formulations. One of the

most important applications of chitosanase is the preparation of COS from chitosan (Ming et al., 2006; Zhang et al., 2012). Enzymatic hydrolysis of chitosan has some advantages for the production of COS, the chitosanases can catalyse the hydrolysis under mild conditions and do not produce monosaccharides (Liu et al., 2009). COS produced by enzymatic hydrolysis of chitosan typically differ from native chitosan by having a molecular weight of 10 kDa or less and are readily soluble in water due to their shorter chain lengths and their free amino groups in GlcN units (Kim & Rajapakse, 2005; Liu et al., 2009). In addition to the low molecular weight and solubility in water, COS possess higher biofunctional properties than chitosan such as antimicrobial, antioxidant, lowering of blood cholesterol, lowering high blood pressure, protective effects against infections, controlling arthritis and enhancing antitumor properties (Kim & Rajapakse, 2005). There is a growing demand for the potential bioactive COSs specifically in food and biomedical fields. Recently Chen et al. (2012) reported the potential of a thermostable chitosanase produced by gene cloning of a chitosanase gene from A. fumigatus in Pichia pastoris for large scale production of COS. According to their reports 3 g enzyme converted 200 kg chitosan into COS in 24 h at 60 °C. In addition to the free enzyme process, application of immobilized chitosanase on COS preparation have been reported using the reactor with immobilized chitosanase on agar gel-coated multidisc impeller (Ming et al., 2006). Enzymatic production of COS from chitosan in different reactor such as batch reactors, column reactors with immobilized enzymes; ultrafiltration (UF) membrane reactors; continuous production of COS by dual reactor system as well as various biological activities of COS have been reported (Kim & Rajapakse, 2005). Seafood processing industries all over the world generate a huge amount of chitionous biowaste/byproducts and disposal of this biomaterial create severe environmental pollution and related issues (Suresh, 2012). Chitosanase-producing microorganisms also find an application in the bioconversion/valorisation of marine crustacean biomaterials and production of bioactive molecules such as enzymes and antioxidants (Wang et al., 2009; Wang et al., 2011). Fungal protoplasts have gained significant importance in mycological research as well as in strain improvement program for biotechnological and industrial applications. Another major application of chitosanase is based on its property of hydrolysis of chitosan containing cell wall for protoplast preparation. Chitosanase is one of the major components of the fungal cell wall lysing enzyme complex particularly for Zygomycetes fungi. Chitosanase also finds an application in chitosan mediated gene delivery. Chitosanase which could degrade chitosan in a specific mode was used in chitosan mediated gene delivery; it improves gene expression by endo-cellular degradation (Liang et al., 2006). Biological control using microorganisms or its component to repress plant disease offers an alternative to chemical fungicide and also it is an eco-friendly approach for controlling agricultural pathogens. Several research groups reported the in vitro antifungal activity of chitosanases, they can be used to improve the resistance of plants against different phyto pathogenic fungi (Gao, Ju, Jung, & Park, 2008; Kouzai et al., 2012). A chitosanse from Bacillus cereus D11 inhibiting the hyphal growth of Rhizotonia solani (Gao et al., 2008); 27 kDa chitosanase from Amycolatopsis sp. CsO-2 inhibiting hyphal growth of Rhizopus oryzae (Kouzai et al., 2012) has reported. Recently Kouzai et al. (2012) reported the molecular mechanism of disease resistance of plant expressing chitosanase activity. Phyto pathogenic fungi change their cell wall components during the infection process to avoid degradation by host lytic enzymes and conversion of the cell wall chitin to chitosan is likely to be one of the infection strategies of pathogens. Thus, introduction of chitosan-degradation activity into plants is expected to improve fungal

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disease resistance. Cho1, from Bacillus circulans MH-K1 has antifungal activity against the rice blast fungus Magnaporthe oryzae. Introduction of the Cho1 gene conferred chitosanase activity to rice cells. Transgenic rice plants expressing Cho1 designed to be localised in the apoplast showed increased resistance to M. oryzae accompanied by increased generation of hydrogen peroxide in the infected epidermal cells. These results strongly suggest that chitosan exists in the enzyme-accessible surface of M. oryzae during the infection process and that the enhancement of disease resistance is attributable to the antifungal activity of the secreted Cho1 and to increased elicitation of the host defence response (Kouzai et al., 2012).

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