1 Introduction

Enzymes are biocatalysts that play a significant role in biochemical and metabolic activities in all living cells. Enzymes from different source has been widely investigated and increasingly explored in the commercial production of various commodities such as food, chemicals, pharmaceuticals, etc. (Chandrasekharan 2015; Liu and Kokare 2017; Gurpilhares et al. 2019). Over the time, enzymes and enzymatic processing have developed as a crucial and central part of the technologies used by different industries to manufacture a wide range of products for numerous applications due to their stereospecific catalytic nature and requisite of mild catalytic conditions (Jung and Park 2014; Suresh 2019; Trincone 2019). Further benefits of enzymes and green enzymatic processing are their low energy requirements, high efficiency, safety, low cost and sustainable nature compared to chemical catalysts (Chandrasekharan 2015; Suresh et al. 2015; Fernandes 2016; Venugopal 2021). Enzymatic digestion/depolymerization of organic compounds, particularly proteins and carbohydrates, provides novel products with promising applications in various fields, including foods, nutrition, agriculture, biomedicine, pharmaceutical, chemical and other industries. In addition, enzymatic biotransformation of organic molecules can offer novel compounds with enhanced properties for specific applications in different industries.

Exploration of polysaccharides, particularly chitin and chitosan from renewable biomass, is receiving much interest in recent years due to their unique functional and biological properties. These amino polysaccharides and their hydrolytic derivatives are playing a progressively vital role in the advancement of nutrition, functional foods, cosmetics, medicines, pharmaceuticals, agriculture, etc. owing to their diverse biofunctional capacities, biocompatibility, non-toxicity, biodegradability, etc. and these promising uses have vast commercial and societal benefits (Kim 2010; Zargar et al. 2015; Ahsan et al. 2018; Shang et al. 2018; Sun et al. 2020; Joseph et al. 2021). Specific, commercially viable and simple enzymatic approaches using specific glycoside hydrolases and other enzymes are evolving as green chemistry/technologies for chitin and chitosan processing (Suresh et al. 2015; Suresh 2019; Sun et al. 2020).

Chitin is an amino polysaccharide that extensively occurs in Nature. It is a linear homopolymer of β-(1 → 4)-linked N-Acetyl-D-glucosamine (N-acetyl-2-amino-2-deoxy-D-glucose, GlcNAc) units (Figs. 1 and 2). Chitin is distributed ubiquitously in Nature as a structural polymer in the exoskeletons of insects and crustaceans and the cell walls of most fungi and some algae (Synowiecki and Al-Khateeb 2003; Feisal and Montarop 2010; Muzzarelli et al. 2012; Suresh 2019). It is the second most abundant and renewable biopolymer/polysaccharide on Earth, next to cellulose (Kurita 2006; Muzzarelli et al. 2012). Despite the widespread occurrence of chitin, currently, the principal sources of commercial chitin constitute marine crustaceans (e.g. shrimp, crab, lobster, etc.) processing discards/by-products (Rinaudo 2006; Harish and Tharanathan 2007; Aranaz et al. 2009; Nidheesh and Suresh 2015a; Younes and Rinaudo 2015; Suresh et al. 2018). Production of chitin from fungal biomass was reported (Kasongo et al. 2020). Chitin is a semi-crystalline/amorphous polysaccharide. Chitin can occur in three different semi-crystalline allomorphic forms in Nature: (1) α form, consists of antiparallel (↑↓↑) GlcNAc chains, (2) β form, contains parallel (↑↑↑) chains of GlcNAc and (3) γ form consists a set of two parallel chains alternating with a single antiparallel (↑↑↓) chain. Generally, all forms of chitin are comparatively resistant to depolymerization. Among the three forms of chitin, α and β forms are the most widely distributed and abundant in Nature (Rinaudo 2006).

Fig. 1
figure 1

Structure of N-Acetyl-D-glucosamine and chitin

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Fig. 2
figure 2

Schematic presentation of chitin (a), chitosan with full D-glucosamine residues (b), partially acetylated chitosan (c), partially acetylated chitosan with N-Acetyl-D-glucosamine residue at reducing end (d) and partially acetylated chitosan with N-Acetyl-D-glucosamine residue at non-reducing end (e)

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Chitosan is also a natural amino polysaccharide. But in Nature, its distribution is restricted to few organisms. It is a copolymer of β-(1 → 4)-GlcNAc and β-(1 → 4)-D-glucosamine (2-amino-D-glucose, GlcN) (Figs. 2 and 3). Usually, chitosan represents a collective name for a family of heterogeneous compounds composed of GlcNAc and GlcN units, with the latter must exceed 70% (Rinaudo 2006; Harish and Tharanathan 2007). Further, the GlcNAc and GlcN units are arbitrarily dispersed in chitosan and do not stand jointly in the polymer molecules (Thadathil and Velappan 2014). Hence, chitosan is generally considered as the derivative of chitin. In contrast to chitin, native chitosan occurs in a meagre quantity in nature. It is mainly found as a structural element in the cell wall of Zygomycetes fungi and also appears in smaller amounts in the cell wall of green algae (Chlorella sp.) and some insect exoskeleton in Nature (Muzzarelli et al. 2012; Thadathil and Velappan 2014). However, chitosan can be prepared by de-N-acetylation of chitin by chemical or enzymatic methods. Therefore, most factories produce chitosan, despite chitin from marine crustaceans processing discards due to its extensive commercial and biological importance higher than chitin (Rinaudo 2006; Harish and Tharanathan 2007; Aranaz et al. 2009; Suresh et al. 2018, Morin-Crini et al. 2019a, b). Production of chitosan from fungal biomass was reported (Abdel-Gawad 2017; Ahmad et al. 2020).

Fig. 3
figure 3

Structure of D-glucosamine and chitosan

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All the above-described allomorphic forms of chitin are insoluble in most common solvents, while chitosan moderately dissolves in aqueous acid solutions. Albeit chitin and chitosan possess many biofunctional characteristics, their potential application is restricted in many fields due to some reasons, particularly their low solubility and poor bioavailability (Hoell et al. 2010; Zargar et al. 2015). Therefore, the depolymerized/hydrolyzed derivatives of chitin and chitosan, particularly oligosaccharides/oligomers (chitooligosaccharides), have received considerable attention owing to their high solubility and remarkable biofunctional properties than their parent polymer (Harish and Tharanathan 2007; Shahidi and Abuzaytoun 2005; Kim and Rajapakse 2005; Rinaudo 2006; Jung and Park 2014; Naidheesh et al. 2015a, b; Nidheesh and Suresh 2015a, b; Shang et al. 2018; Suresh 2019; Kaczmarek et al. 2019; Sun et al. 2020). Source of different chitin and chitosan substrate and chitooligosaccharides is illustrated in Fig. 4.

Fig. 4
figure 4

Source of chitin, chitosan and chitooligosaccharides

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Chitooligosaccharides are the partially depolymerized derivatives of chitin [(GlcNAc)2–20] or chitosan [(GlcN)2–20]. The depolymerization of chitin or chitosan and production of chitooligosaccharides can be achieved by mechanical, chemical and enzymatic (biocatalytic) methods or a combination of them. The mechanical methods for chitin or chitosan hydrolysis include ultrasonication (Baxter et al. 2005), hydrothermal treatment, hydrodynamic cavitation (Wu et al. 2014) and electromagnetic, gamma rays or microwave irradiation (Zainol et al. 2009). However, these mechanical methods have been scarcely investigated and are not well-suited for producing well-defined chitooligosaccharides (Kaczmarek et al. 2019; Noa Miguez et al. 2021). Chemical procedures for chitooligosaccharides preparation comprise degradative methods, such as chitin or chitosan depolymerization by strong acid hydrolysis or oxidation, and chemical synthesis strategies (Noa Miguez et al. 2021). Chemical methods for the synthesis of chitooligosaccharides have progressed substantially in the last decades. Still, they usually result in low yields. They may require protection/deprotection steps, the use of hazardous chemicals, the formation of hazardous waste, high production cost, formation of a large amount of monomers, etc. (Je and Kim 2012; Suresh and Kumar 2012; Thadathil et al. 2014; Kaczmarek et al. 2019).

As compared to mechanical and chemical methods, enzymatic production of chitooligosaccharides from chitin or chitosan substrate is very attractive since it requires milder reaction conditions (relatively low temperatures and slightly acidic pH) and does not generate harmful by-products or wastes (Sinha et al. 2016), which makes it safer and more environmentally friendly approach (Sheldon and Woodley 2018; Kaczmarek et al. 2019; Suresh 2019). In addition, strategies based on biocatalysis present a higher efficiency and allow easier control of the product composition due to inherent enzyme stereospecificity (Kim and Rajapakse 2005; Ilankovan et al. 2006; Wee et al. 2009; Jung and Park 2014; Naidheesh et al. 2015a, b). However, the physicochemical properties of the chitin or chitosan substrate substantially affect the chemical composition of the final products (Noa Miguez et al. 2021). In principle, it should be possible to develop enzymes and enzymatic processes that enable the production of partially depolymerized derivatives (chitooligosaccharides) of chitin or chitosan with consistent quality and biofunctional properties (Hoell et al. 2010). Though, synthesis of desired chitooligosaccharides in terms of the degree of polymerization (DP), degree of de-N-acetylation (DD), the pattern of acetylation (PA) by enzymatic depolymerization approaches is not straightforward (Jung and Park 2014).

Source of different substrates and enzymatic production of chitooligosaccharides are illustrated schematically in Figs. 4 and 5.

Fig. 5
figure 5

Schematic presentation of enzymatic production defined chitooligosaccharides

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The water-soluble low molecular mass chitooligosaccharides have high commercial value due to their versatile biofunctional capacity and wide applications in various fields (Muzzarelli et al. 2012; Thadathil and Velappan 2014; Sun et al. 2020; Noa Miguez et al. 2021). Consequently, in the recent decade, there has been an increasing demand for the production of potential chitooligosaccharides for specific applications (Wee et al. 2009; Xia et al. 2011; Nguyen et al. 2014; Pechsrichuang et al. 2013; Thadathil and Velappan 2014; Naidheesh et al. 2015a; Nidheesh and Suresh 2015b). Along this line of discussion, few reviews have been published that report different aspects of chitinase, chitosanase and enzymatic synthesis of chitooligosaccharides and applications of chitooligosaccharides (Jung and Park 2014; Thadathil and Velappan 2014; Suresh 2019; Sun et al. 2020; Noa Miguez et al. 2021). This chapter is focused mainly on the various technological approaches for the enzymatic depolymerization of chitin and chitosan and the prospective production of different types of chitooligosaccharides. The different bioactivities and various applications of chitin, chitosan chitooligosaccharides are not subjects of discussion in this chapter.

2 Enzymes Acting on Chitin and Chitosan

Enzymatic hydrolysis of chitin and chitosan can be catalyzed by several glycoside hydrolases (GHs) (Jung and Park 2014; Suresh 2019), consisting of specific enzymes, like chitinases (Oyeleye and Normi 2018; Singh et al. 2021) and chitosanases (Thadathil and Velappan 2014; Li et al. 2015; Suresh 2019). In addition, non-specific enzymes such as other glycosidases (e.g. cellulases, glycanases, pectinases, muramidase, etc.), proteases (e.g. pepsin) and lipases have also been applied in chitin and chitosan depolymerization (Kittur et al. 2005; Xia et al. 2008; Montilla et al. 2013). A large number of factors can influence the outcome of an enzymatic depolymerization reaction of any polysaccharides. In chitooligosaccharides production from chitin or chitosan, both the choice of enzyme and the selection of substrate apply an extensive control on the outcome (Noa Miguez et al. 2021). Various specific and non-specific enzymes involved in chitin or chitosan depolymerizations for chitooligosaccharides formation are briefly discussed below.

2.1 Specific Enzymes in the Formation of Chitooligosaccharides

2.1.1 Chitinases

Chitinases are a group of enzymes, can catalyze the hydrolysis of β-(1 → 4) glycosidic bond of GlcNAc-GlcNAc linkages of chitin polymer (Stoykov et al. 2015; Oyeleye and Normi 2018). According to the recommendations of the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology (IUBMB) in 1992, the chitinases are grouped into two classes (Class 1 and Class 2) based on the substrate specificity and mode of action (Reetarani et al. 2000; Beygmoradi et al. 2018) (https://www.qmul.ac.uk/sbcs/iubmb/enzy me/). Class 1: Endochitinases [EC 3.2.1.14; (1 → 4)-2-acetamido-2-deoxy-β-D-glucan glycanohydrolase], and Class 2: β-N-acetylhexosaminidases (EC 3.2.1.52). The current IUBMB enzyme nomenclature has included former EC 3.2.1.29 and EC 3.2.1.30 exo-chitinases enzyme classes under a single enzyme class, known as β-N-acetylhexosaminidases (Reetarani et al. 2000; Hoell et al. 2010; Suresh 2019). Nevertheless, the existing terminology for chitinolytic enzymes is confusing to some authors (Sun et al. 2020), owing in part to the point that the previous classification system has not been fully rejected (Palanivelu and Vaishna 2013; Vincenzi et al. 2014). The endochitinases are the specific enzymes acting on chitin polymer at the internal sites and randomly cleave the GlcNAc-GlcNAc linkages, thus exclusively release low molecular mass oligosaccharides of GlcNAc with different DP as the end product (Fig. 6). Whereas the β-N-acetylhexosaminidases act on chitin oligosaccharides [(GlcNAc)2–20] from the reducing end and produce monomeric GlcNAc as the sole end product (Reetarani et al. 2000; Oyeleye and Normi 2018; Suresh 2019; Sun et al. 2020). In this chapter, we follow the terminology suggested by IUBMB. In addition, we discuss only the endochitinases that catalyze the formation of oligosaccharides/oligomers from chitin molecules/biomass.

Fig. 6
figure 6

Schematic presentation of endochitinase activity on chitin polymer and formation of N-acetyl-oligosaccharides [(GlcNAc)2–20]

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Chitinases activities are widely distributed in Nature in various groups of living organisms. Chitinases are reported from various organisms, including those that lack chitin like plants, bacteria, viruses and vertebrates, similarly in chitin containing organisms like fungi, insects and crustaceans (Oyeleye and Normi 2018). These organisms synthesize them for different physiological roles. Along this subject line, many reviews have been published by various research teams (Kasprzewska 2003; Guan et al. 2009; Hoell et al. 2010; Stoykov et al. 2015; Oyeleye and Normi 2018; Malik and Preety 2019; Sun et al. 2020). The structure and action mechanism of endochitinases have been reported (Cantarel et al. 2009; Hoell et al. 2010; Liu et al. 2015; Kidibule et al. 2018; Noa Miguez et al. 2021). Based on the similarity of amino acid sequence, the endochitinases belongs to two-glycoside hydrolase (GH) families, GH-18 and GH-19 (Cantarel et al. 2009; Liu et al. 2015), which displayed different catalytic mechanisms such as retaining and inverting processes. Family GH18 comprises bacterial and fungal chitinases (Horn et al. 2006; Huang et al. 2012), while family GH19 mainly contains plant chitinases (Malik and Preety 2019; Suresh 2019). Family GH18 enzymes use the substrate-assisted retaining double displacement mechanism, whereas family GH19 enzymes employ the inverting direct displacement mechanism (Hoell et al. 2010). Chitinases are also competent for hydrolyzing β-(1 → 4) glycosidic linkage of GlcNAc-GlcN and GlcN-GlcNAc units, allowing degradation of partly acetylated chitosan, but their degree of activity is significantly lower than chitosanases (Noa Miguez et al. 2021). Further, chitinases never cleave the β-1,4-glycoside GlcN-GlcN bonds of chitosan chains, differentiating these enzymes from chitosanases (Jung and Park 2014). Chitinases belonging to family GH-18 need a GlcNAc moiety at subsite-1 (Kidibule et al. 2018), while family GH-19 chitinases do not require GlcNAc moiety at this subsite. Due to this cause, family GH-18 chitinases produce oligosaccharides with GlcNAc residues at their reducing end (Stoykov et al. 2015; Lienemann et al. 2009). Conversely, oligosaccharides formed by family GH-19 chitinases have N-acetylated or de-N-acetylated reducing ends (Noa Miguez et al. 2021). Both types of chitinases have been explored to synthesize N-acetyl chitooligosaccharides [(GlcNAc) 2–20] from chitin bioresource.

2.1.2 Chitosanases

Chitosanases (EC 3.2.1.132, chitosan N-acetylglucosaminohydrolase) catalyze the hydrolysis of β-(1 → 4)-glycosidic linkages of chitosan polymer. Chitosanases are generally endo-acting enzymes, which randomly cleave the chitosan polymer at the internal sites and release a mixture of oligosaccharides, exclusively as the end product (Thadathil and Velappan 2014; Suresh 2019) (Figs. 7, 8 and 9). Chitosanase are widely distributed in Nature and have been produced, purified and characterized by several groups of organisms, including viruses (Sun et al. 1999), bacteria (Oh et al. 2011), fungi (Nidheesh et al. 2015a) and plants (Osswald et al. 1994). The biochemical characteristics, structure functional relationship, etc., of chitosanases produced from various sources, have been reviewed (Thadathil and Velappan 2014; Singh et al. 2019; Sun et al. 2020; Noa Miguez et al. 2021). Shinya and Fukamizo (2017) have studied the interaction between chitosan and its related enzymes.

Fig. 7
figure 7

Schematic presentation of chitosanase subclass I activity on chitosan polymers and formation of chitosan oligosaccharides [(GlcN)2–20]. Chitosan polymer with full GlcN units (a), partially acetylated chitosan (b), [(GlcN)2–20] with full GlcN units (c), partly acetylated [(GlcN)2–20] (e), partly acetylated [(GlcN)2–20] with GlcNAc units at reducing end (d, f)

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Fig. 8
figure 8

Schematic presentation of chitosanase subclass II activity on chitosan polymers and formation of chitosan oligosaccharides [(GlcN)2–20]. Chitosan polymer with full GlcN units (a), partially acetylated chitosan (b), [(GlcN)2–20] with full GlcN units (c), partly acetylated [(GlcN)2–20] (d)

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Fig. 9
figure 9

Schematic presentation of chitosanase subclass III activity on chitosan polymers and formation of chitosan oligosaccharides [GlcN)2–20]. Chitosan polymer with full GlcN units (a), partially acetylated chitosan (b), [(GlcN)2–20] with full GlcN units (c), partly acetylated [(GlcN)2–20] (e), partly acetylated [(GlcN)2–20] with GlcNAc units at non-reducing end (d, f)

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According to the comparison of amino acid sequence, chitosanases belongs to five GH families, viz: GH-5, GH-8, GH-46, GH-75 and GH-80 (Wang et al. 2008; Hoell et al. 2010; Shinya and Fukamizo 2017) and are capable of hydrolyzing different types of β-(1 → 4)-glycosidic linkages in the chitosan molecules. Most bacterial chitosanases belong to the GH-46 family (Saito et al. 1999), and chitosanases from fungi include in the GH-75 family (Zhu et al. 2012). Chitosanases from GH-46, GH-75, and GH-80 families are thought to be a substrate (chitosan) specific, though GH-5, GH-7 and GH-8 chitosanases also showed other enzyme actions, like cellulase and transglycosylation activities (Sun et al. 2020). Chitosanases are grouped into three subclasses based on the specificity of cleavage sites, viz: (1) subclass I chitosanases, which cleave both GlcN-GlcN and GlcNAc-GlcN linkages (Fig. 7), (2) subclass II chitosanases, which cleave only GlcN-GlcN linkages (Fig. 8) and (3) subclass III chitosanases, which cleave both GlcN-GlcN and GlcN-GlcNAc linkages (Fig. 9) in the chitosan molecules (Jung and Park 2014; Thadathil and Velappan 2014; Hoell et al. 2010; Weikert et al. 2017) The hydrolysis of GlcN-GlcN linkages is the common property of all these chitosanases (Figs. 7, 8 and 9). While, chitosanases are unable to hydrolyze GlcNAc-GlcNAc linkages in the partly acetylated chitosan, and this property distinguishes chitosanases from chitinases (Hoell et al. 2010). A new classification system for chitosanases employing their uniqueness and inclinations for subsites (−2) to (+2) have been proposed (Weikert et al. 2017).

The above-described specific enzymes acting on chitin and chitosan are synthesized by various organisms, including microorganisms, plants and animals and found abundantly in Nature (Hoell et al. 2010; Suresh 2019; Sun et al. 2020). The molecular structure and function as well as other aspects of chitinase (Oyeleye and Normi 2018; Sun et al. 2020; Noa Miguez et al. 2021), human chitinases (Guan et al. 2009), plant chitinases (Kasprzewska 2003) and microbial chitinases (Hoell et al. 2010; Stoykov et al. 2015) have been reviewed. The chitosanases, particularly of microbial chitosanases, have been reviewed by the various research team (Fukamizo et al. 2000; Lukas et al. 2012; Thadathil and Velappan 2014; Weikert et al. 2017; Langner and Göhre 2016; Shinya and Fukamizo 2017; Singh et al. 2019; Sun et al. 2020; Tzelepis and Karlsson 2021). Among the different groups of organisms, which produce specific enzymes acting on chitin and chitosan, the microorganisms, mainly bacteria and fungi, have commercial significance concerning the production of these enzymes as well as their application in chitooligosaccharides synthesis from different chitin or chitosan substrate/biomass (Thadathil and Velappan 2014; Naidheesh et al. 2015a, b; Santos-Moriano et al. 2018).

2.2 Other Enzymes Involved in the Synthesis of Chitooligosaccharides

2.2.1 Non-specific Enzymes Acting on Chitin and Chitosan

As well as the specific enzymes such as chitinases and chitosanases, few non-specific enzymes have exhibited some potential to identify the β-(1 → 4)-glycosidic linkages of chitin and chitosan to depolymerize them (Muzzarelli 1997; Pantaleone et al. 1992; Jung and Park 2014). The non-specific enzymes, which exhibited chitin or chitosan depolymerization activity was reported to few glycosidases (e.g. cellulases, lysozyme, pectinases, amylases, etc.) (Amano and Ito 1978; Kittur et al. 2005; Santos-Moriano et al. 2016), proteases (e.g. papain, pronase) (Pan et al. 2016), lipases (Lee et al. 2008) and chitin-active Lytic Polysaccharide Monooxygenases (LPMOs) (Vaaje-Kolstad et al. 2010; Mutahir et al. 2018). However, the rate of chitin or chitosan depolymerization by these non-specific enzymes can be far below the rate exhibited by the specific enzymes (Aktuganov et al. 2019; Noa Miguez et al. 2021).

2.2.2 Chitin Deacetylase in the Synthesis of Chitooligosaccharides

Chitin deacetylases (EC 3.5.1.41; Chitin amidohydrolase) catalyze the hydrolysis of N-acetamido bonds of chitin, converting it to chitosan (Tsigos et al. 2000; Zhao et al. 2010; Jung and Park 2014; Suresh 2019). Chitin deacetylases activities have been reported in various microorganisms, particularly fungi (Aranaz et al. 2009, 2014; Suresh et al. 2011a). However, the activity of these chitin deacetylases is severely restricted by the insolubility of the chitin polymer. Chitin deacetylases have exhibited potential in synthesizing specific chitooligosaccharides (Tokuyasu et al. 1997, 2000). However, presently, there is no commercially viable method existing to produce chitin deacetylases and their uses in the synthesis of specific chitooligosaccharides (Suresh et al. 2011a; Zhao et al. 2010; Suresh 2019).

2.2.3 Chitinases and Chitosanases with Transglycosylation Activity

The endoglycosidase-catalyzed transglycosylation has emerged recently as a competent technique for creating novel oligosaccharides with potential bio-functionalities (Mangas-Sánchez and Adlercreutz 2015; Li and Wang 2016). Endochitinases belonging to the GH-18 family and a few chitosanases have exhibited some amount of transglycosylation capacity (Sirimontree et al. 2014; Noa Miguez et al. 2021). These specific enzymes act on chitin or chitosan molecules with transglycosylation capacity that can move the cleaved oligosaccharide molecules to a proper acceptor forming new glycosidic bonds and generating precise chitooligosaccharides (Tanabe et al. 2003; Purushotham and Podile 2012; Mallakuntla et al. 2020). It was reported that chitinases with transglycosylation activity are more significant than chitinases with hydrolytic activity for application in food and biomedical fields owing to their capacity to change crystalline chitin to soluble long-chain oligosaccharide (Oyeleye and Normi 2018).

2.2.4 Chitin Active Lytic Polysaccharide Monooxygenases (LPMOs)

LPMOs (EC 1.14.99.53–56) are enzymes capable of breaking glycolic linkages in the polysaccharides with crystalline nature via oxidizing either C1 or C4 of the glucopyranose moiety (Vaaje-Kolstad et al. 2017; Mutahir et al. 2018; Kaczmarek et al. 2019 LPMOs are comparatively new enzymes that have been discovered in 2010 and an extensively studied enzyme in current years (Gaber et al. 2020). LPMOs have been allotted to auxiliary activity (AA) families AA9, AA10, AA11, AA13, AA14 and AA15 in the Carbohydrate-Active Enzymes (CAZy) database (Tandrup et al. 2018). These enzymes act a leading role in the degradation of sugar-based biopolymers (such as cellulose, hemicellulose, chitin and starch) and have an encouraging implication for biomass transformation (Wang et al. 2021). LPMOs are potent tools in the degradation of various types of biomass, including marine biomasses such as chitins (Vaaje-Kolstad et al. 2010; Mekasha et al. 2017 Nakagawa et al. 2020). In distinction to GHs, such as chitinases and chitosanases, LPMOs are potent for directly breaking glycolic linkages in extremely crystallized chitin, enhancing its accessibility for succeeding enzymatic processing (Mutahir et al. 2018). Consequently, considerable attention has been received in chitin active LPMOs research. So far, LPMOs with chitinolytic activities have been recognized in carbohydrate AA families of AA10, AA11 and AA15 (Hemsworth et al. 2015). The chitin-active LPMO was first reported in Serratia marcescens AA10 by Vaaje-Kolstad et al. (2010). Currently, the chitinase active LPMO has been reported in various groups of organisms, including bacteria, fungi, viruses and arthropods (Chiu et al. 2015; Sabbadin et al. 2018). Among these organisms’ fungi and bacteria are the most important producer of chitinolytic LPMOs (Kaczmarek et al. 2019). Nakagawa et al. (2013) investigated the enzymatic hydrolysis of α-chitin with varying particle size and crystallinity produced by mechanical pre-treatment using a cockatiel of chitinase and LPMO (from S. marcescens). Mekasha et al. (2017) studied the percentage of chitinases, an LMPO and a β-N-acetylhexosaminidase from S. marcescens to depolymerize crustacean (crab and shrimp) chitins.

3 Chitooligosaccharides and Types of Chitooligosaccharides

Chitooligosaccharides can be defined as the partially hydrolyzed/depolymerized products/derivatives of chitin or chitosan. It can be β-(1 → 4) linked homo or hetero-oligomers/oligosaccharides of GlcNAc and/or GlcN (Yang and Yu 2014; Sun et al. 2020). Usually, chitooligosaccharides have a DP up to 20 and an average molecular mass of  <4.0 kDa. Chitooligosaccharides are highly soluble in water. They can exhibit different DP, DD and PA depending on the type of chitin or chitosan substrates used in their production and the extent of their depolymerization. These characteristics of chitooligosaccharides significantly affect their different biofunctional properties (Hamer et al. 2015). The chitooligosaccharides can be classified into three distinct types based on their composition (Santos-Moriano et al. 2019; Noa Miguez et al. 2021) viz: (1) fully acetylated chitin oligosaccharides (FA-COS or N-acetyl-COS) (composed exclusively of GlcNAc residues (Fig. 6), (2) partly acetylated chitosan oligosaccharides (PA-COS) (formed mainly by GlcN residues with few GlcNAc residues) (Figs. 7, 8 and 9) and (3) fully deacetylated chitosan oligosaccharides (FD-COS) (composed of GlcN residues only) (Figs. 7, 8 and 9). These chitooligosaccharides are very high-value natural products with numerous promising bioactivities and potential applications in various fields, such as nutrition, functional foods, agriculture, cosmetics, pharmaceutical and biomedicine and so on (Moon et al. 2017; Marmouzi et al. 2019; Aktuganov et al. 2019; Ismail et al. 2020). The reader can refer to other chapters of this book for more information on these aspects of chitooligosaccharides. In general, these chitooligosaccharides synthesized by depolymerization of chitin or chitosan are heterogeneous blends of different types of oligosaccharides, still with extended hydrolysis time (Jung and Park 2014) (Figs. 7, 8 and 9), which describes the low reproducibility rate and conflicting results when their biofunctional capacities are checked. Consequently, several methods have been studied for the separation and purification of chitooligosaccharides, such as metal affinity (Le Dévédec et al. 2008); gel filtration (Sørbotten et al. 2005) and ion exchange (Haebel et al. 2007), chromatography and ultrafiltration (Lopatin et al. 2009). Preparative scale separation of chitooligosaccharides is usually based on their mass/size by gel filtration chromatography (Jung and Park 2014). The following sections briefly describe the development of enzymatic processes towards the synthesizing of specific type of chitooligosaccharides. As detailed above, the three types of chitooligosaccharides (Figs. 6, 7, 8 and 9) can be produced by enzymatic approaches using different enzymes.

Generally, high-performance liquid chromatography (HPLC) alone or in combination with matrix-assisted laser desorption ionization-time of flight (MALDI-TOF) mass spectrometer was used to analyze the oligosaccharides obtained from the hydrolysis of chitin or chitosan substrate. In addition to HPLC and MALDI-TOF, silica gel thin layer (TLC) was also widely used in the separation and identification of oligosaccharides liberated by the hydrolysis of chitin or chitosan substrate.

4 Enzymatic Production of Chitooligosaccharides

4.1 Enzymatic Production of Chitooligosaccharides by Specific Enzymes

4.1.1 Enzymatic Production of N-Acetyl-COS/FA-COS from Chitin

As described above, N-Acetyl-COS/FA-COS is the partly depolymerized derivative of chitin (Fig. 6). N-Acetyl-COS/FA-COS and its derivatives prepared from chitin have exhibited good solubility in water and various biological properties and can be explored in a wide variety of chemical and biological industries (Synowiecki and Al-Khateeb 2003; Oyeleye and Normi 2018). Endochitinases are the specific enzymes investigated in chitin depolymerization and synthesis of N-acetyl-COS/FA-COS (Fig. 6). As noted previously, the specific enzymes acting on chitin molecules are usually produced as a chitinolytic enzyme complex. Hence, to develop a high yield of N-acetyl-COS/FA-COS from chitin biomass, the enzyme preparation must comprise higher endochitinase activity and relatively low or nil β-N-acetylhexosaminidases activity (Aloise et al. 1996; Suresh 2019). Chitinases preparations from various groups of microorganisms, mainly from bacteria and fungi, have been explored in the manufacture of N-acetyl-COS/FA-COS (GlcNAc)2–20 from diverse chitin substrates (Takiguchi and Shimahara 1988, 2003; Woo and Park 2003; Tanaka et al. 1999; Kuk et al. 2006; Laura et al. 2006; Jung et al. 2007; Suresh et al. 2011b; Wang et al. 2012; Thadathil et al. 2014). Because of the strong crystalline nature and insolubility in most common solvents, in general, chitin is subjected to pre-treatment with strong acids (such as HCl/H2SO4 for colloidal chitin or H2PO4 for swollen chitin) before enzymatic digestion to enhance the accessibility of the substrate to the enzyme (Jung and Park 2014).

Preparation of N-acetyl-COS/FA-COS (GlcNAc)2–20 using bacterial chitinases from Paenibacillus illiosensis KJA424 (Jung et al. 2007), Bacillus sp. WY22 (Woo and Park 2003), Pyrococcus kodakaraensis KOD1 (Tanaka et al. 1999), Aeromonas sp. GJ-18 (Kuk et al. 2006), Bacillus cereus TKU027 (Wang et al. 2012), Vibrio anguillarum E-383a (Takiguchi and Shimahara 1988), and hyperthermophilic archaeon Thermococcus kodakaraensis KOD1 (Tanaka et al. 2003) have reported. Jung et al. (2007) was reported the highest N-acetyl-COS/FA-COS formation of 2.21 ± 0.13 mg/l from swollen chitin after 24 h of hydrolysis with a chitinase preparation of P. illiosensis KJA-424. Production of N-acetyl chitotriose as a primary product from a colloidal and glycol chitin employing the chitinase prepared from Bacillus sp. WY22 has been reported (Woo and Park 2003). N-acetyl chitobiose [(GlcNAc)2] was prepared by a chitinase from a marine V. anguillarum E-383a from colloidal chitin (Takiguchi and Shimahara 1988). Production of N-acetyl chitobiose [(GlcNAc)2] from colloidal chitin by using a chitinase from the hyperthermophilic archaeon T. kodakaraensis KOD1 was reported (Tanaka et al. 2003). Kuk et al. (2006) reported the production of N-acetyl chitobiose [(GlcNAc)2] after 7 days of hydrolysis of swollen α-chitin and powdered β-chitin with a yield of 78.9% and 56.6%, respectively, using a crude chitinase preparation of Aeromonas sp. GJ-18. N-acetyl-COS [(GlcNAc)2–5] was produced from partially deacetylated chitin (60% DD) using the culture supernatant prepared by a chitinolytic B. cereus TKU027 (Wang et al. 2012).

Preparation of N-acetyl-COS/FA-COS [(GlcNAc)2–20] using fungal chitinases from Lecanicillium fungicola (Laura et al. 2006); Penicillium monoverticillium CFR 2 (Suresh et al. 2011b) and Aspergillus flavus CFR 10 and Fusarrium oxysporum CFR 8 (Thadathil et al. 2014) have been reported. Recently, a chemo-enzymatic synthesis of N-acetyl-COS using the fungal chitinase from Thermomyces lanuginosus ITCC 8895 have been reported (Kumar et al. 2021). The N-acetyl-CO/FA-COS was produced from different chitin substrates employing the chitinases obtained from L. fungicola (Laura et al. 2006). The highest yield (mmol/l) of N-acetyl COS obtained was 0.17 from a crystalline α-chitin, 2.77L from a deacetylated α-chitin and 4.44 from a deacetylated β-chitin (Laura et al. 2006). In a study, Thadathil et al. (2014) reported the production of N-acetyl-COS using the chitinases preparation from A. flavus CFR 10 and F. oxysporum CFR 8 with the highest yield (mmol/l) of 10.4 ± 0.28 after 6 h hydrolysis and 10.2 ± 0.01 after 6 h hydrolysis 30 h, respectively. Suresh et al. (2011b) reported a maximum N-acetyl-COS production (mmol/l) of 6.81 ± 0.03 from colloidal chitin, 0.36 from a deproteinized shrimp shell and 0.60 ± 0.02 from a demineralized crab shell using the crude chitinase obtained from P. monoverticillium CFR 2. These authors observed that N-acetyl chitohexose (~62.43%) was the primary product of the hydrolysate from colloidal chitin by HPLC analysis. Production of N-acetyl-COS by the chitinase from a novel Streptomyces chilikensis strain RC1830 and its various bioactivities was reported (Behera et al. 2020).

Recently, a chemo-enzymatic method for synthesizing short-chain N-acetyl-COS using ionic liquids and a chitinase from T. lanuginosus ITCC 8895 has been reported (Kumar et al. 2021). The authors produced 1.10 ± 0.89 mg/ml of N-acetyl chitobiose [(GlcNAc)2] and 1.07 ± 0.92 mg/mL of N-acetyl chitotriose [(GlcNAc)3] from the Trihexyltetradecylphosphonium bis(2,4,4-trimethylpentyl) phosphinate treated chitin substrate.

4.1.2 Enzymatic Depolymerization of Chitosan and Production of Chitosan Oligosaccharides

Chitosan oligosaccharides [(GlcN)2–20] are the partly degraded derivative of chitosan, and the only positively charged oligosaccharides occur in Nature (Thadathil and Velappan, 2014; Sun et al. 2020). They possess exceptional biological functions, such as antioxidant, antimicrobial, antitumor, immunoenhancing, etc., activities for a wide range of potential applications in various fields (Sun et al. 2020). Consequently, there has been increasing attention in converting chitosan to specific chitosan oligosaccharides (Thadathil and Velappan 2014; Suresh 2019). As described above (Figs. 7, 8 and 9), the partial depolymerization of chitosan can develop two types of chitosan oligosaccharides, such as (1) FD-COS and (2) PD-COS, depending on the type of chitosan substrates employed. The enzymatic production of FD-COS or PD-COS from chitosan substrates can be carried out effectively using specific chitosanases (Kim and Rajapakse 2005; Naidheesh et al. 2015a, b; Nidheesh 2016; Suresh 2019). It was well reported that the viscosity of chitosan declines quickly when added with chitosanase, in accord with depolymerization and production of the oligosaccharides (Jung and Park 2014). Chitosanase prepared from a different group of fungi and bacteria has been explored in producing oligosaccharides [(GlcN)2–20] from various chitosan substrates.

Enzymatic production of oligosaccharides [(GlcN)2–20] from different chitosan substrates employing microbial chitosanase has been reported by many research teams (Shimosaka et al. 1995; Kim et al. 1998; Kurakake et al. 2000; Cheng and Li 2000; Choi et al. 2004; Gao et al. 2009; Pechsrichuang et al. 2013; Nguyen et al. 2014; Naidheesh et al. 2015a, b; Nidheesh 2016). Shimosaka et al. (1995) reported the production of chitobiose [(GlcN)2] and chitotriose [(GlcN)3] from chitopentose [(GlcN)5] and chitosan using crude chitosanase from Acinetobacter sp. CHB10. Production of chitobiose [(GlcN)2] was reported from chitosan mixture with 75%–85% DD using a chitosanase obtained from B. cereus S1 (Kurakake et al. 2000). A purified chitosanase (22.5 kDa) from A. fumigatus KH-94 was used for the production of chitobiose [(GlcN)2], chitotriose [(GlcN)3] and chitotetraose [(GlcN)4] from chitohexaose [(GlcN)6] (Kim et al. 1998). In addition to the production of chitosan oligosaccharides with DP 3–6 (50% yields), the 22.5 kDa chitosanase from A. fumigatus KH-94 has produced oligosaccharides with higher DP (> DP 7, 50% yield) from chitosan (Kim et al. 1998). Choi et al. (2004) reported the reaction pattern and production of different oligosaccharides from chitohexose [(GlcN)6] using the chitosanase produced from Bacillus sp. KCTC 0377BP. A purified chitosanase produced by B. cereus D-11 hydrolyzed chito-oligomers [(GlcN)5–7] into chitobiose [(GlcN)2], chitotriose [(GlcN)3] and chitotetraose [(GlcN)4] as the end products (Gao et al. 2009). Production of oligosaccharides [(GlcN)2–5] from chitosan using a chitosanase obtained from Aspergillus sp. Y2K has been reported (Cheng and Li 2000). In this study, a chitosan-degrading soil fungus (Aspergillus sp. Y2K) was grown in a minimal medium containing chitosan as the sole carbon source. Further, the purified chitosanase were applied for the hydrolysis of chitosan substrate and production of chitosan oligosaccharides. They reported that chitotriose [(GlcN)3], chitotetraose [(GlcN)4] and chitopentaose [(GlcN)5], were the main products in the chitosan hydrolysate (Cheng and Li 2000). Naidheesh et al. (2015a; b) reported chitosan oligosaccharides production employing crude chitosanase from a soil fungi Purpureocillium lilacinum CFRNT12. The chitosanase was prepared from P. lilacinum CFRNT12 utilizing solid-state fermentation (SSF) using shrimp (Penaeus sp.) by-products as the substrate. Later the crude chitosanase was concentrated using the dialysis method employing polyethylene glycol (mol. wt. 20,000). The chitosanase (169.92 unit/ml) was tested on two different chitosan substrates [crystalline (>22 mesh size) and colloidal chitosan]. The highest yield of oligosaccharides (4.43 mmol/l from colloidal chitosan and 1.7 mmol/l from crystalline chitosan) was observed after 24 h of digestion. Chitotriose [(GlcN)3] and chitotetraose [(GlcN)4] were the main products of hydrolysis (Naidheesh et al. 2015a; b; Nidheesh 2016).

A purified chitosanase (46 kDa) was obtained from Bacillus sp. HW-002 produced chitobiose [(GlcN)2] as a main end product from chitosan (Lee et al. 1996). Park et al. (1999) have reported the digestion of chitopentose [(GlcN)5] and chitohexose [(GlcN)6] into chitobiose [(GlcN)2] and chitotriose [(GlcN)3] by a purified chitosanase (34 kDa) produced from Matsuebacter chitosanotabidus 3001. Sáncheza et al. (2017) investigated the chitooligosaccharides production by two different enzymatic processes. The first process used chitosanase enzymatic hydrolysis using a commercial chitosanase from Streptomyces griseus, and the second consisted of a two-step procedure based on chemical hydrolysis followed by chitosanase hydrolysis. It was observed that chitooligosaccharides produced in the second process were composed of 63% of fully deacetylated sequences (Sáncheza et al. 2017).

In addition, the potential of recombinant chitosanase in the depolymerization of different chitosan substrates and production of oligosaccharides have been reported (Chen et al. 2012; Pechsrichuang et al. 2013; Cheng et al. 2021). Preparation of chitosan oligosaccharides using a thermostable chitosanase produced by recombination of a gene responsible for chitosanase synthesis from A. fumigatus in Pichia pastoris (Chen et al. 2012). The authors reported that 3 g of enzyme preparation hydrolyzed 200 kg of chitosan into oligosaccharides after 24 h at 60 °C. Pechsrichuang et al. (2013) reported the preparation of oligosaccharides [(GlcN)2–6] from different chitosan substrates using a recombinant B. subtilis chitosanase. In a recent study, Cheng et al. (2021) has reported a pilot-scale production of chitosan oligosaccharides using a recombinant chitosanase. In this work, a Vector pET20b-csn containing the B. circulans MH-K1 chitosanase gene was transformed into Escherichia coli strain BL21 (DE3) for recombinant protein (chitosanase) expression. Further, the recombinant chitosanase was prepared by growing recombinant E. coli strain BL21 (DE3) in a Luria–Bertani (LB) medium. The intracellular recombinant chitosanase was extracted from the cell pellet and purified. Further, the semi-purified recombinant chitosanase was used in the pilot-scale production of chitosan oligosaccharides by mixing ~2500 units to a 10 litter chitosan solution (5%) for 72 h, at ambient temperature. The author reported the formation of a digestion product having molar masses of 833 ± 222 g/mol with multiple DP [chitotriose (GlcN)3 to chitotetrose (GlcN)4] (Cheng et al. 2021).

All the above-described studies revealed that application of specific enzymes (chitosanases) for the partial depolymerization of chitosan are desirable enzymatic processes for the preparation of oligosaccharides [(GlcN)2–20]. Though, the application of chitosanase in chitosan depolymerization is restricted due to various reasons, including the high cost of commercially available chitosanase, lack of availability in bulk volume and so on (Thadathil and Velappan 2014; Suresh 2019; Cheng et al. 2021). Hence, some alternative approaches have been reported on the production of chitosan oligosaccharides from different chitosan substrates, such as enzyme immobilization recycling methods (Kim and Rajapakse 2005; Ming et al. 2006; Kuroiwa et al. 2008), increasing the enzymatic concentration by stepwise substrate mixing (Ming et al. 2006) and continual production in a dual-reactor system (Jeon and Kim 2000; Santos-Moriano et al. 2016). Ming et al. (2006) have used a reactor with an immobilized chitosanase on agar gel-coated multidisc impeller for the production of chitosan oligosaccharides and attained yields of 20–45% for the addition of chitopentaose [(GlcN)5] and chitohexaose [(GlcNAc)6], dependent on the surface activity of the enzyme. Preparation of chitosan oligosaccharides from chitosan using different immobilized enzyme reactors like batch and column reactor and ultrafiltration membrane reactor and continuous production of oligosaccharides by a dual-reactor system have been reported (Kim and Rajapakse 2005). Kuroiwa et al. (2008) studied the development of a chitosanase immobilized on amylose-coated magnetic nanoparticles for the production of chitosan oligosaccharides. The authors have reported that in the digestion reaction employing the immobilized chitosanases, pentamers [(GlcN)5] and hexamers [(GlcNAc)6], of chitosan oligosaccharide were formed at a high yield (40%), which is higher than that of conventional methods for producing these oligosaccharides (Kuroiwa et al. 2008). Jeon and Kim (2000) developed a dual-reactor system with a packed-bed reactor (PBR) to minimize the viscosity of chitosan, followed by an ultrafiltration membrane reactor to form chitosan oligosaccharide of the preferred DP.

Fungal biomasses are an excellent source for chitosan extraction and oligosaccharides production (Mahata et al. 2014; Abdel-Gawad et al. 2017; Ahmad et al. 2020). In a study, Mahata and co-workers (2014) were hydrolyzed intact mycelium of Rhizopus oligosporus NRRL2710 with a chitosanase preparation from Streptomyces sp. N174 and produced a mixture of functionally active chitooligosaccharides. This study shows the potential of bioconversion of abundant fungal chitin or chitosan biomass to high-value biomolecules with no ecological issues.

4.2 Chitooligosaccharides Production by Non-specific Enzymatic Depolymerization

Because of the high cost of commercial chitinase and chitosanase, the use of cheap non-specific enzymes such as glucanase, lipase, lysozyme, cellulase, amylase, papain and pectinase (derived from bacteria, fungi, mammal and plant sources) have been explored for chitin or chitosan depolymerization and synthesis of their oligosaccharides (Pantaleone et al. 1992; Terbojevich et al. 1996; Muzzarelli 1997; Kittur et al. 2005; Ilankovan et al. 2006; Lee et al. 2008; Pan et al. 2016; Santos-Moriano et al. 2016). A pectinase preparation from A. niger has been explored for the hydrolysis of chitosan and production of a mixture of low molecular weight chitosan (LMWC) (86%) and oligosaccharides (4.8%) (Kittur et al. 2005). Production of N-acetyl COS (~81% yield) from de-crystallized chitin using the commercial bovine pepsin has been reported (Ilankovan et al. 2006). Chitosan with 80% DD was digested using the cellulase prepared from A. niger to produce chitooligosaccharides with DP 3–11 (Xie et al. 2009). Zhang et al. (1999) effectively applied a blend of cellulase, α-amylase and proteinase to produce chitooligosaccharides with DP 5–17. Wu (2012) has reported the production of chitosan oligosaccharides (DP 2–8) using the commercial α-amylase from a chitosan substrate prepared from the skin of Clanis bilineata larvae. Enzymatic preparation of chitosan oligosaccharides (DP 1–6) by a commercial lipase yield 93.8% after 24 h digestion at 37 °C was reported (Lee et al. 2008). Among these non-specific enzymes, papain (a cysteine protease) was very attractive because of its low cost and wide acceptance in the food industry and the ability to depolymerize chitosan efficiently (Pan et al. 2016).

In addition to free non-specific enzyme processes, immobilized non-specific enzymes such as papain (Lin et al. 2002), lysozyme (Aiba 1994), pronase (Kumar et al. 2004) and α-amylase (Santos-Moriano et al. 2016) have been tested for chitooligosaccharides production. In a work, chitosanlytic activity present in a commercial α-amylase (from B. amylolyquefaciens, BAN) was covalently immobilized onto glyoxal agarose beads and used for the continuous production of chitooligosaccharides in a dual-reactor system (Santos-Moriano et al. 2016). All the studies confer its potential in the commercial preparation of chitooligosaccharides since these enzymes are commercially available in bulk quantity, inexpensive than chitinase and chitosanase and very simple to manage the digestion processes (Jung and Park 2014; Suresh 2019).

4.3 Enzymatic Synthesis of Chitooligosaccharides by the Transglycosylation Activity of Chitinase and Chitosanase

As detailed above, chitin and chitosan oligosaccharides can be prepared using various techniques, including mechanical, chemical and enzymatic methods. However, all these methods, including enzymatic hydrolysis (either specific or nonspecific) of chitin and chitosan, generally produce a complex oligosaccharides mixture (N-acetyl COS/FA-COS, FD-COS, PA-COS), which has been grouped according to the DA or DD, DP or molecular weight, and the molecular weight distribution (poly-disparity, PD) (Li et al. 2016). Therefore, the bioactivities of these oligosaccharides are habitually analyzed using heterogeneous and/or relatively poorly characterized oligosaccharides mixtures. Hence, it is difficult to determine which molecules are responsible for the detected biofunctional properties. To overcome these problems, the enzymatic synthesis of specific chitooligosaccharides has been investigated by transglycosylation (Akiyama et al. 1995; Tokuyasu et al. 1997, 2000; Kobayashi et al. 1996, 2006; Li et al. 2016). The transglycosylation methods allow regioselective synthesis under mild conditions and without complicated protection (Li et al. 2016). GlcNAc-(1 → 4)-GlcN and β-GlcNAc-(1 → 4)-β-GlcNAc(1 → 4)-β-GlcNAc-(1 → 4)-GlcN was synthesized using the chitin deacetylase from Colletotrichum lindemuthianum (Tokuyasu et al. 1997, 2000).

The synthesis of chitin or chitosan oligosaccharides (N-acetyl COS or FD-COS) using a transglycosylation reaction catalyzed by lysozyme and an N-acetyl COS or FD-COS with DP4-12 have reported utilizing N, N′, N′′-tri(monochloro)acetylchitotriose and N-acetyl chitotriose [(GlcNAc)3] as glycosyl donors followed by N-demonochloroacetylation (Akiyama et al. 1995). The transglycosylation reaction activity of a chitosanase from B. cereus was used to produce high-DP chitosan oligosaccharides (DP2-7) (Hsiao et al. 2008).

The synthesis of single oligosaccharides with higher DP such as N-acetyl chitopentaose [(GlcNAc)5] using a recombinant E. coli harbouring the nodC gene (encoding chitooligosaccharide synthase) has been reported (Zhang et al. 2007). A one-pot chemo-enzymatic technique for regio- and stereospecific production of N-acetyl chitoheptaose [(GlcNAc)7] utilizing 1, 2-oxazoline derivative of [(GlcNAc)5] and [(GlcNAc)2] as starting materials by defined transglycosylation reaction by a mutant chitinase has also been described (Yoshida et al. 2012).

4.4 Synthesis of Chitooligosaccharides by Whole-Cell Biocatalytic Approaches

Most chitin and chitosanase synthesizing microorganisms require chitin or chitosan or their derivates as source nutrients (for growth) and an inducer for production of respective enzymes (chitinase or chitosanase) (Jung and Park 2014; Suresh 2019). In a study, Liang et al. (2012) have reported the direct production of N-acetyl COS [(GlcNAc)2, (GlcNAc)4, (GlcNAc)5 and (GlcNAc)6] using the fermentation approaches by growing a chitosanase producing B. cereus TKU022 in a medium containing 1.5% shrimp head powder after 48 h of incubation. Chitooligosaccharides production was reported in the culture supernatant of Acinetobacter calcoaceticus TKU024 from shellfish by-products along with chitosanases after 96 h fermentation (Wang et al. 2011). Even though the quantity of end products of this approach is meagre, these types of research have significance to the development processes for the direct production of chitooligosaccharides from the abundant chitin bio-resources.

5 Conclusion and Future Prospects

Chitooligosaccharides, the partially degraded derivative of polymer chitin or chitosan, possess numerous unique biological activities that describe these biomolecules’ significance in various fields. However, the biological properties of these chitooligosaccharides have been significantly influenced by many factors, including their chemical composition, molecular mass, the extent of deacetylation and acetylation pattern. Therefore, the production of well-defined selective chitooligosaccharides has immense value to understand their bioactive effect more profound and, therefore, their upcoming applications in nutrition and biomedicine. The enzymatic processes of chitin and chitosan depolymerization, employing specific GH enzymes such as endochitinase and chitosanase, has achieved considerable progress in chitooligosaccharides production. Further, current advances in endochitinases and chitosanases with transglycosylation activity and chitin active LPMOs research have improved the enzymatic production of defined chitooligosaccharides. Today, all these enzymes are progressively emerged as powerful tools for the enzymatic preparation of chitooligosaccharides, specifically when a precise and distinct method is essential. Regardless of much progress in the past decade, currently, commercial-scale productions of pure chitooligosaccharides with defined characteristics by enzymatic approaches is still a task. The main limitations of the commercial-scale enzymatic processes recline on the high cost of commercially available specific enzymes, non-availability in bulk volume and so on, even though these specific enzymes are copious in Nature (Jung and Park 2014; Thadathil and Velappan 2014; Suresh 2019; Cheng et al. 2021). Nevertheless, numerous recent investigations report the significance of the enzymatic methodologies for chitooligosaccharides production and might soon advance it into economical to conventional chemical methods. Hence, future research must focus on developing highly efficient and appropriate technologies by properly selecting the chitin or chitosan substrate and the enzyme source towards making highly defined chitooligosaccharides for wide application in biomedicine and other fields.