Microbial chitinases — Production, characterization, purification and their biotechnological and therapeutic applications: An integrated review

Abstract

Graphical Abstract

Chitin is the second most abundant natural polysaccharide after cellulose and consists of N-acetyl-D-glucosamine units linked by β-1,4-glycosidic bonds. In nature, chitin does not accumulate due to the synergistic action of chitinolytic enzymes. Based on their catalytic domains, chitinases are classified into glycosyl hydrolase families GH18 and GH19. They are widely produced by bacteria and filamentous fungi. Different types of chitinolytic enzymes, including endochitinases, exo-acting enzymes, and N-acetylglucosaminidases, have been reported to exhibit antimicrobial and insecticidal activities, making them valuable tools for controlling phytopathogenic fungi and insect pests. Chitin degradation generates chitooligosaccharides (COS), which possess diverse biological properties such as antimicrobial, antioxidant, anti-inflammatory, and antitumor activities, contributing to improved human health. Microbial chitinases are also applied in several industrial and environmental processes, including protoplast formation, single-cell protein production, and dye removal. Advances in recombinant expression and genetic engineering have enhanced chitinase production, stability, and catalytic efficiency. Moreover, recombinant chitinases have been successfully utilized in biocontrol strategies and in developing transgenic plants with increased resistance to phytopathogens. This review highlights the broad agricultural, industrial, and biomedical applications of chitinases and their crucial role in promoting environmental sustainability and advancing bio-based industrial processes.

Full Article

Microbial Chitinases — Production, Characterization, Purification and their Biotechnological and Therapeutic Applications: An Integrated Review

Medhat A. Abu-Tahon,^a,* Heba I. Aboelmagd,^b Manal M. Housseiny,^c Ahmad M. Abdel‐ Mageed,^a Nada Daifalla,^b Adaugo C. Isichei,^b Sahar Algadi,^b Yahia H. Ali,^a Intisar K. Saeed,^a Howayada M. Mostafa,^d Salma Y. S. Elsheikh,^a Ali M. Ali,^e Amani A. S. Abdelaziz,^b Ishraga Izzeldin,^b Nermien H. Seddek,^f Assad A. Rezigalla,^g Elhadi Miskeen,^h Ali M. S. Eleragi,ⁱ Mohammed-Elfatih Twfieg,^j and George S. Isaac ^c

Chitin is the second most abundant natural polysaccharide after cellulose and consists of N-acetyl-D-glucosamine units linked by β-1,4-glycosidic bonds. In nature, chitin does not accumulate due to the synergistic action of chitinolytic enzymes. Based on their catalytic domains, chitinases are classified into glycosyl hydrolase families GH18 and GH19. They are widely produced by bacteria and filamentous fungi. Different types of chitinolytic enzymes, including endochitinases, exo-acting enzymes, and N-acetylglucosaminidases, have been reported to exhibit antimicrobial and insecticidal activities, making them valuable tools for controlling phytopathogenic fungi and insect pests. Chitin degradation generates chitooligosaccharides (COS), which possess diverse biological properties such as antimicrobial, antioxidant, anti-inflammatory, and antitumor activities, contributing to improved human health. Microbial chitinases are also applied in several industrial and environmental processes, including protoplast formation, single-cell protein production, and dye removal. Advances in recombinant expression and genetic engineering have enhanced chitinase production, stability, and catalytic efficiency. Moreover, recombinant chitinases have been successfully utilized in biocontrol strategies and in developing transgenic plants with increased resistance to phytopathogens. This review highlights the broad agricultural, industrial, and biomedical applications of chitinases and their crucial role in promoting environmental sustainability and advancing bio-based industrial processes.

DOI: 10.15376/biores.21.1.Abu-Tahon

Keywords: Chіtіnases; Microorganisms; Purification; Biological control; Chitooligosaccharides; Protoplasts; Transgenic plants

Contact information: a: Department of Biological Sciences, College of Science, Northern Border University, Arar, Saudi Arabia; b: Department of Basic Sciences, Deanship of Preparatory Year and Supporting Studies, Imam Abdulrahman Bin Faisal University, Dammam 34212, Saudi Arabia; c: Biological and Geological Sciences Department, Faculty of Education, Ain Shams University, Roxy, Heliopolis, P.C.11757, Cairo, Egypt; d: Department of Chemistry, College of Science, Northern Border University, Arar, Saudi Arabia; e: Department of Biological Sciences, College of Science, King Faisal University, Al-Ahsa, Kingdom of Saudi Arabia; f: Department of Respiratory Care, College of Applied Medical Sciences-Jubail 4030, Imam Abdulrahman Bin Faisal University, Al Jubail, Saudi Arabia; g: Department of Anatomy, College of Medicine, University of Bisha, Bisha 61922, Saudi Arabia, h: Department of Obstetrics and Gynecology, College of Medicine, University of Bisha, Bisha 61922, Saudi Arabia; i: Department of Microorganisms and Clinical Parasitology, College of Medicine, University of Bisha, Bisha 61922, Saudi Arabia; j: Department of Biomedical Sciences, College of Medicine, King Faisal University, Al-Ahsa 31982, Saudi Arabia;

* Corresponding author: medhatahon@gmail.com

Graphical Abstract

INTRODUCTION

Chitinases (EC 3.2.1.14) are ubiquitous endo-acting glycosyl hydrolases (GH) that catalyze the cleavage of β-1,4-glycosidic bonds within chitin, the second most abundant biopolymer. This enzymatic action produces chitooligosaccharides, which can be further hydrolyzed by N-acetyl-β-D-hexosaminidases into monomeric N-acetylglucosamine (Cohen-Kupiec and Chet 1998). These enzymes are primarily grouped into two main families, the glycosyl hydrolase family 18 (GH18) and the glycosyl hydrolase family 19 (GH19), based on their distinct active sites (Udaya Prakash et al. 2010).

From a research perspective, chitinases present a fascinating area of study due to their diverse catalytic mechanisms and intricate molecular architecture (Berini et al. 2018). Investigating their substrate specificity is paramount, as it serves as a critical avenue for elucidating the precise correlations between enzymatic function and their physiological roles across various biological systems. Furthermore, a comprehensive understanding of this specificity directly informs and optimizes strategies for the bioproduction of industrially significant compounds. Notably, these enzymes demonstrate catalytic activity across a broad spectrum of chitin polymorphs, encompassing α-, β-, and γ-chitin, in addition to their derivatives such as chitosan and various chitooligosaccharides (Horn et al. 2006).

Chitinases are primarily produced by microorganisms; although similar enzymes have been reported in plants and animals, they function in plant defense against chitin-containing pathogens (Grover, 2012) and in immune responses and inflammation in animals (Garth et al. 2018). Microbial chitinases are increasingly favored due to their superior attributes compared to chitinases from other sources. Their production is characterized by significantly higher yields and reduced operational costs, which can be primarily attributed to their capacity to efficiently utilize abundant and inexpensive raw materials, such as chitinous shellfish waste (Karthik et al. 2014), insect exoskeletons (Merzendorfer and Zimoch 2003), and fungal cell walls (Langner and Göhre 2016). Furthermore, these raw materials can be pre-processed into more bioavailable forms, such as colloidal chitin, thereby optimizing enzyme production efficiency (Deng et al. 2019). A critical advantage lies in the inherent stability of microbial chitinases across diverse environmental conditions, coupled with their amenability to genetic engineering. This allows for targeted modifications to enhance catalytic activity or tailor specific enzymatic properties, thereby expanding their biotechnological utility (Stoykov et al. 2015).

Chitinases are broadly distributed across a multitude of organisms, where they fulfill critical physiological and ecological functions (Jahromi and Barzkar 2018). In bacterial systems, these enzymes are integral to nutrient acquisition, parasitic interactions, and the essential recycling of chitin (Jahromi and Barzkar 2018). Fungi leverage chitinases for fundamental processes such as spore germination, hyphal development, morphogenesis, and nutrient assimilation (Thakur et al. 2023). In plants, chitinases are expressed during growth as pathogenesis-related proteins, conferring protection against chitin-containing pathogens (Ali et al. 2020). Human physiology includes chitinases in serum, leukocytes, and gastric secretions, contributing to innate defense mechanisms against chitinous threats (Abdelraouf et al. 2024). Crustaceans depend on chitinases for vital processes, including molting, growth, reproduction, and defense against pathogens (Zhang et al. 2014).

Numerous bacterial genera, including Aeromonas, Bacillus, Streptomyces, Paenibacillus, Serratia, and Pseudomonas have been identified as prolific chitinase producers (Itoh and Kimoto 2019). Likewise, several filamentous fungi, such as Aspergillus, Conidiobolus, Beauveria, Mucor, Neurospora, Trichoderma, and Penicillium, are known to synthesize diverse chitinases (Thakur et al. 2023; Gupta et al. 2025). Among the available approaches, submerged fermentation remains the most widely employed method for microbial chitinase production (Stoykov et al. 2015).

Chitinases, owing to their capacity to hydrolyze chitin found in insect exoskeletons, fungal cell walls, and various other chitin-containing structures, have gained substantial biotechnological importance (Avupati et al. 2017). Their applications are multifaceted, encompassing roles as antimicrobial and insecticidal agents in the biocontrol of phytopathogens (Chatterton and Punja 2009). Furthermore, chitinases are central to the bioconversion of chitin into pharmacologically valuable products, such as N-acetylmuramic acid and chitooligosaccharides (Liang et al. 2018). These products exhibit diverse bioactivities and have been explored as antimicrobial agents, modulators of host defense mechanisms, drug delivery vehicles, cholesterol-lowering agents, and food preservatives (Rameshthangam et al. 2018). In addition to these functionalities, emerging evidence highlights the potential of chitinases as diagnostic biomarkers for a range of pathological conditions, including inflammatory and autoimmune disorders, asthma, oral diseases, and both acute and chronic inflammatory states (Castañeda-Ramírez et al. 2013). Furthermore, chitinases have been extensively investigated for their utility in fungal protoplast generation (Hassan 2014) and in the development of transgenic plants with improved resistance against phytopathogens and insect pests (Kumar et al. 2018).

This review aims to provide a comprehensive and integrated perspective on microbial chitinases. It encompasses their classification, mechanisms of action, microbial producers, and the various strategies employed to enhance their production, stability, purification, recombinant expression, and genetic engineering. Additionally, this review presents an inclusive overview of their therapeutic, agricultural, and industrial applications, with a particular emphasis on recent progress and emerging innovations reported in the literature.

Chitin Production and Characterization

Chitin is structurally similar to cellulose, as both are polysaccharides composed of β-(1→4)-linked glycosidic bonds; however, chitin differs in that the hydroxyl group at the C2 position of the glucose monomer is replaced by an N-acetyl group (N-acetylglucosamine), which enhances its stability, rigidity, and mechanical strength compared to cellulose (Kobayashi et al. 2023). Chitin exists in three main crystalline forms: α-, β-, and γ-chitin, which differ in molecular organization, intermolecular interactions, and physicochemical properties (Kaya et al.,2017). α-Chitin, with antiparallel polysaccharide chains, forms extensive intra- and intermolecular hydrogen bonds, resulting in a highly ordered, densely packed structure that is chemically inert and poorly soluble; it is predominantly found in fungal cell walls, insect exoskeletons, and crustacean shells (Av et al. 2004; Langner and Göhre 2016; Merzendorfer and Zimoch 2003; Casadidio et al. 2019). In contrast, β-chitin has parallel chains with weaker intermolecular forces, producing a more loosely packed and chemically reactive structure, mainly occurring in diatoms and cephalopod skeletal structures (Gardner and Blackwell 1975). γ-Chitin exhibits a mixed chain arrangement, resulting in intermediate structural order, solubility, and reactivity, and is found in some beetles and cephalopods (Kaya et al. 2017).

Chitin’s robust molecular structure makes it highly persistent in the environment, leading to large amounts of chitin-containing waste from shrimp, crab, and other seafood processing (Zhou et al. 2018). Over 60% of this waste is improperly managed, contributing to pollution due to chitin’s resistance to natural degradation (Yadav et al. 2021). To address this problem, researchers have developed various extraction methods from chitin-containing residues, which are broadly categorized into chemical and biological approaches

Chemical extraction techniques, applied at both laboratory and industrial scales, typically necessitate the application of potent acids and bases (El Knidri et al. 2018). Chemical extraction of chitin typically involves three main steps: deproteinization, demineralization, and decolorization. Deproteinization is performed by treating shells with alkaline solutions, while demineralization (elimination of calcium carbonate) is carried out using acidic conditions, most commonly hydrochloric acid at elevated temperatures (Younes and Rinaudo 2015). Decolorization is achieved with organic solvents such as acetone, ethanol, methanol, and chloroform, or with inorganic agents such as sodium hypochlorite and hydrogen peroxide (Younes and Rinaudo 2015). The chemical approach, while effective, is risky and expensive, underscoring the demand for safer chitin extraction methods (El Knidri et al. 2018).

Fig. 1. Schematic representation of chitin structure and its enzymatic degradation

The biological extraction of chitin primarily involves two steps: deproteinization and demineralization. Deproteinization can be achieved either enzymatically or via microbial fermentation. Enzymatic methods employ proteases such as alcalase and trypsin, whereas fermentation utilizes protease-producing microorganisms such as Aspergillus and Bacillus, typically under controlled conditions. Demineralization is performed through lactic acid fermentation by Lactobacillus, which converts calcium carbonate into calcium lactate. The resulting chitin is then thoroughly washed, dried at low temperatures, and milled to preserve its structural integrity. This biological approach offers several advantages, including cost-effectiveness, environmental friendliness, and the production of chitin with desirable physicochemical properties (Karthik et al. 2014).

Mechanisms and Classification of Chitinases

Chitinolytic enzymes can be classified based on their amino acid sequences and mechanistic pathways. Broadly, they are divided into two main types according to their mode of action: endo-chitinases and exo-chitinases (Fig. 1). Endo-chitinases (EC 3.2.1.14) randomly hydrolyze chitin, producing soluble low-molecular-weight oligomers of N-acetylglucosamine (GlcNAc), such as chitotetraose, chitotriose, and di-acetylchitobiose, as well as longer oligosaccharides with a degree of polymerization greater than four (DP > 4). Exo-chitinases are further classified into chitobiosidases (EC 3.2.1.29), which release di-acetylchitobiose sequentially from the non-reducing end of the chitin chain, and 1-4-β-N-acetylglucosaminidases (EC 3.2.1.30), which hydrolyze oligomeric products into GlcNAc monomers (Cohen-Kupiec and Chet 1998). In this context, chitinases in Family GH 18 exhibit critical dual functionality encompassing both hydrolytic (bond cleavage) and transglycosylation (bond creation/synthesis) activities. While hydrolysis is the primary catabolic function, degrading chitin into smaller oligomers, transglycosylation serves as the synthetic function, creating new glycosidic bonds to synthesize longer oligosaccharides. Consequently, the equilibrium between cleavage and synthesis is critically dependent on environmental factors, notably water activity and substrate concentration (Madhuprakash et al. 2013).

Fig. 2. Classification of chitinases according to amino acids sequences and mode of action. Adapted from (Funkhouser and Aronson 2007)

Chitinases cleave the β-1,4-glycosidic bonds in chitin by means of retaining or inverting mechanisms. Most GH18 chitinases follow a two-step retaining mechanism, using aspartic acid as a nucleophile and glutamic acid as an acid/base catalyst to maintain the anomeric configuration of the product. Endochitinases, such as SmChiA (GH18), are non-processive enzymes with a shallow, exposed substrate-binding cleft that enables random internal cleavage; their catalytic site contains the conserved Asp-Glu-Asp motif (Horn et al. 2006). In contrast, exochitinases, including chitobiosidases, are processive enzymes with a tunnel-shaped active site that guides the chitin chain, allowing sequential release of disaccharide units without enzyme dissociation (Morimoto et al. 1997).

Chitinases are also classified into three glycosyl hydrolase (GH) families based on the sequences of their catalytic domains (Funkhouser and Aronson 2007): GH18, found in viruses, fungi, bacteria, insects, and mammals; GH19, present in actinobacteria, purple phototrophic bacteria, and plants; and GH20, associated with human chitinases that act on chitin degradation products rather than polymeric chitin (Fig. 2).

GH18 chitinases employ substrate-assisted catalysis, where the C2-acetamido group of GlcNAc acts as a nucleophile, forming a transient oxazolinium ion and retaining the stereochemistry of the glycosidic bond (Chen et al. 2020). GH19 chitinases use acid-base catalysis, where an acid protonates the glycosidic oxygen and a base attack the anomeric carbon, resulting in inversion of the anomeric configuration (Kawase et al. 2004). GH20 enzymes, including β-N-acetylhexosaminidases and chitobiases, hydrolyze chitobiose or N-acetylgalactos-amine from glyco-conjugates (Vaaje‐Kolstad et al. 2013).

Chitinolytic Microorganisms

Microbial chitinases are generally preferred over their plant or animal counterparts in industrial and biotechnological applications for several key reasons. First, microorganisms can produce large amounts of chitinases at low production costs, especially when inexpensive and readily available substrates, such as seafood waste, are utilized (Kuddus 2014). Second, microbial chitinases exhibit stability across a wide range of temperatures and pH, making them suitable for diverse industrial applications (Al Abboud et al. 2022; Al-Rajhi, et al. 2023). Third, microorganisms can be genetically manipulated with relative ease to enhance chitinase productivity or enzymatic properties, opening avenues for the development of tailor-made enzymes for specific applications (Stoykov et al. 2014). Moreover, the extraction and purification of chitinases from microbial sources is generally more efficient and cost-effective compared to plant or animal sources (Oyeleye and Normi 2018). Chitinolytic microorganisms inhabit diverse terrestrial and aquatic environments, and shellfish waste. A straightforward approach to screen for these microorganisms involves culturing them on agar media containing colloidal chitin as the sole carbon source and identifying colonies by the formation of clearing zones around them (Abu-Tahon and Isaac 2020).

Bacterial chitinases

Bacterial chitin degradation is essential for biogeochemical cycling and sustaining ecosystem carbon–nitrogen balance (Kumar et al. 2022). Bacteria can sense chitin and respond through mechanisms such as movement toward the chitin source (chemotaxis) or growth in its direction (chemotropism). They also adhere to chitin surfaces, secrete extracellular chitinolytic enzymes to break down the polymer, and uptake the resulting chitin-derived oligosaccharides for metabolic use (Selenius et al. 2018).

The review emphasizes the considerable potential of bacterial chitinases for various commercial applications, owing to their stability under extreme pH and temperature conditions, rapid growth, and suitability for genetic engineering (Kumar et al. 2022). Chitinases are broadly occurring in Arthrobacter, Aeromonas, Bacillus, Clostridium, Chromobacterium, Klebsiella, Pseudomonas, Serratia, Streptomyces, and Xanthomonas (Jahromi and Barzkar 2018). From a functional and structural perspective, bacterial chitinases are classified into three GH families: GH-18, GH-19, and GH-23 (Fig. 3), and the majority of bacterial chitinases belong to the GH-18 family (Udaya Prakash et al. 2010).

Fig. 3. Classification of bacterial and fungal chitinases

GH-18 bacterial chitinases are divided into three subfamilies, A, B, and C, based on sequence and structural features. Subfamily A, which is the most widespread, typically contains a chitin insertion domain (CID) that enhances binding to insoluble chitin, and generally functions as an endochitinase by randomly cleaving internal β-1,4-glycosidic bonds. Subfamily B, which lacks a CID, still contributes significantly to chitin hydrolysis and exhibits both endo- and exo-chitinase activities. Subfamily C is less well characterized and restricted to a limited number of bacterial species, indicating possible specialized adaptations.

Chitin-binding proteins (CBPs) enhance chitinase efficiency by promoting chitin degradation or binding to chitin-containing surfaces (Frederiksen et al. 2013). GH-19 chitinases are mainly distributed in actinobacteria and purple bacteria, whereas GH-23 chitinases are mainly present in peptidoglycan lyases from bacteria and bacteriophage. This group also includes goose-type (G-type) lysozymes, which are specifically active in the hydrolysis of chitin and chitooligosaccharides (Arimori et al. 2013). Several species noted for high chitinase production are listed in Table 1.

In nature, Serratia marcescens is among the most organized and efficient bacterial chitin degraders and has been extensively studied for its chitinase production (Vaaje‐Kolstad et al. 2013). Multiple genes encoding chitinase have been identified in Serratia marcescens strains, whereas the S. marcescens Nima strain exhibits nearly 43-fold higher activity compared to the others (Bhattacharya et al. 2007).

Table 1. Bacterial Chitinases and their Classification

Fungal chitinases

The chitinase enzyme is essential in the fungal life cycle, where it contributes to cell wall remodeling and plasticization, thereby regulating hyphal growth, tube extension, branching, fusion, germination, and division (Karthik et al. 2014; Bakri et al. 2022). The distribution and abundance of chitin differ among fungi; in filamentous species, it is mainly located in the inner cell wall layers adjacent to the plasma membrane, with a relatively high content of about 20%, whereas in yeasts, it is restricted to constriction rings, septa, and budding scars, where its content ranges from 0.5% to 5% (Hartl et al. 2012).

The regulation of fungal cell wall degradation, whether targeting self or non-self-structures, is thought to be governed more by substrate accessibility in healthy hyphae than by the specificity of chitinases. The susceptibility of the fungal cell wall to enzymatic hydrolysis is controlled by the balance between protection and deprotection during mycoparasitism, aging, and autolysis (Gruber and Seidl-Seiboth 2012). Fungi produce hydrophobic cell wall proteins such as QID74 and carbohydrate-binding proteins to shield their cell walls from the action of hydrolytic enzymes. Trichoderma harzianum produces a 74 kDa cell wall protein that is essential for both adherence to hydrophobic surfaces and mycelium protection (Rosado et al. 2007).

Fungal chitinases are traditionally categorized into classes III and V, based on their predominant occurrence in specific organisms (Fig. 3). Class III (plant-type) and Class V (bacterial-type) chitinases differ in their substrate-binding grooves: Class V enzymes possess deep, tunnel-shaped grooves and act as processive exo-chitinases, whereas Class III enzymes have shallow, open grooves and function as non-processive endo-chitinases (van Aalten et al. 2001; Hoell et al. 2005).

Based on the amino acid sequences of the GH18 family and the structure of their substrate-binding clefts, fungal chitinases are further divided into three subclasses: A, B, and C. Subgroups A and C belong to class V, while subgroup B belongs to class III (Gruber et al. 2011). Functionally, subgroup A chitinases are involved in fungal growth and autolysis, subgroup B chitinases are primarily nutritional enzymes in mycoparasites and insect-pathogen fungi, and subgroup C chitinases, found mainly in Trichoderma atroviride and T. virens, participate in both endogenous and exogenous chitin degradation (Berini et al. 2018; Hartl et al. 2012).

Several fungal genera, including Aspergillus, Beauveria, Conidiobolus, Metarhizium, Mucor, Neurospora, Penicillium, Trichoderma, and Verticillium, have been reported as potent producers of chitinases (Thakur et al. 2023). Some of the species reported to exhibit excellent chitinase activity are summarized in Table 2. Deng et al. (2019) reported that Chit-46 chitinase from Trichoderma harzianum suppresses the growth of the phytopathogenic fungus Botrytis cinerea, hydrolyzing chitin into GlcNAc₂ with up to 94.8% efficiency.

Optimization Parameters for Maximum Production of Microbial Chitinase

Environmental and nutritional conditions have a strong impact on microbial chitinases. These variables include the initial pH of the medium, the duration and temperature of the incubation, the various chitin sources, the impact of shaking velocity, and the effects of various carbon and nitrogen sources that were tested as salt basal media supplements (Tables 1 and 2). The primary factor influencing chitinase productivity is the form of chitin, such as crystalline chitin, shrimp or crab shell powder, or colloidal chitin. Colloidal chitin is a highly accessible form of chitin, ideal for chitinase studies. It is prepared from insoluble chitin powder, regardless of source. The process involves treating chitin powder with a concentrated strong acid (e.g., HCl) to break its crystalline structure. This acid-chitin mixture is then diluted with cold water, causing amorphous particles to precipitate, forming a colloidal suspension. Subsequent neutralization and washing remove residual acid and impurities. The final product is a milky, viscous suspension of amorphous chitin, offering increased surface area for enzymatic degradation (Abu-Tahon and Isaac 2020). Fungal chitinase production is typically performed by submerged fermentation, this method allows for better oxygen transfer, nutrient availability, and enzyme secretion into the liquid phase. It is widely used because it enables easy recovery and purification of chitinase from the culture broth (Abu-Tahon and Isaac 2020). In Trichoderma viride, maximum yields were obtained using colloidal chitin as the carbon source under optimized conditions of pH 6.5, 35 °C, and 125 rpm (Abu-Tahon and Isaac 2020).

Moreover, chitinase production is strongly influenced by incubation time, generally increasing to a peak before declining during prolonged cultivation. This pattern occurs because enzyme synthesis is closely linked to the microorganism’s growth phase and metabolic activity, with maximum production typically observed during the logarithmic or early stationary phase. In addition, the availability of chitin as a substrate and the accumulation of metabolic byproducts significantly affect enzyme output, as nutrient depletion or the buildup of inhibitory compounds can reduce secretion or promote enzyme degradation (Karthik et al. 2014).

Shaking speed is a key factor influencing enzyme productivity, as mechanical forces can induce vacuolation in older hyphal compartments, potentially weakening the hyphae or promoting fragmentation (Paul et al. 1994). Chitinase production has been shown to vary with shaking velocities ranging from 100 to 200 rpm (Table 1 & 2), as agitation directly affects oxygen transfer, nutrient distribution, and shear stress in the culture; optimal shaking promotes sufficient aeration and mixing to enhance enzyme production, whereas too little or excessive agitation can reduce yield due to stress or limited substrate availability (Alves et al. 2018).

From a physiological standpoint, solid-state fermentation (SSF) provides several advantages for chitinase production, including high volumetric yields, elevated product concentrations, reduced effluent generation, and minimal requirements for sophisticated equipment. Moreover, solid substrates bound to amino acids are chemically more stable than free substrates, making them particularly suitable for large-scale production of economically valuable compounds (Stoykov et al. 2015). In this regard, El-Beltagi et al. (2022) reported that the ideal medium for chitinase production by Talaromyces funiculosus was crab shell chitin amended with yeast extract 0.2% and beet molasses 100% at pH 6.5 for seven days.

The One-Factor-At-a-Time (OFAT) approach involves varying a single factor while keeping all others constant. Although widely used, this method has notable limitations: it requires many experiments to assess multiple factors, it cannot reveal interactions between variables, it is time-consuming and costly, and it may miss the optimal combination of conditions (Vaidya et al. 2003). To address the limitations of the OFAT method, several statistical approaches have been developed, including Plackett-Burman design (PBD), central composite design (CCD), Taguchi’s robust design (TRD), and response surface methodology (RSM) (Han et al. 2008). These methods offer simplicity, efficiency, and nutrient savings, while allowing the analysis of factor interactions, making them effective for optimizing enzyme production and media components (Mishra et al. 2012). For example, Lee and Kim (2015) optimized chitinase production in Pseudomonas fluorescens was achieved using PBD and CCD, identifying yeast extract, CaCl₂.2H₂O, and crab shell powder, as key factors. The CCD-optimized medium increased enzyme activity to 1.03 U/mL, nearly 2.9 times greater than standard conditions.

Table 2. Chitinases Produced by Fungi and their Classification

Purification and Characterization of Microbial Chitinases

Enzyme purification involves sequential steps aimed at isolating the enzyme from complex mixtures while preserving its activity. The purification and characterization of microbial chitinases are crucial for determining their specific biochemical properties, such as substrate specificity, optimal pH, and thermal stability. This knowledge is essential for harnessing their potential in various biotechnological applications, including biocontrol of plant pathogens, waste management, and the production of valuable chitooligosaccharides (Govindaraj et al. 2024). Various techniques have been employed for chitinase purification, typically including dialysis, precipitation using ammonium sulfate or organic solvents, gel filtration chromatography, and ion-exchange chromatography. The final assessment of enzyme purity and homogeneity is typically conducted using sodium dodecyl sulfate-polyacrylamide gel electrophoresis (Table 3). The final and most critical assessment of enzyme purity and homogeneity is typically conducted using Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE). This technique is considered the gold standard because it separates proteins based solely on their molecular weight, independent of their native charge or conformation. The successful outcome of the purification process is visually confirmed by the appearance of a single, sharp band on the gel, which corresponds precisely to the expected molecular weight of the target enzyme, thereby providing unambiguous evidence that all contaminating proteins have been effectively removed (Chen et al. 2013).

Table 3. Purification and Characterization of Chitinases Produced by Different Microorganisms

According to the literature, Sephadex is the most used gel filtration medium for chitinase purification, while diethylaminoethyl cellulose (DEAE) is frequently utilized for ion exchange chromatography, as shown in Table 3. In this respect, Abu Tahon and Issac (2020) reported that chitinase from Trichoderma viride was purified to homogeneity with a 73.1% yield and a 5.48-fold purification using ammonium sulfate precipitation (65%), Sephadex G-100, and DEAE-cellulose. The pure enzyme recorded a molecular mass of 62 kDa, exhibited maximal activity at pH 6.5 and 40 °C, and was activated by Ca²⁺ and Mn²⁺, while Hg²⁺, Zn²⁺, Cu²⁺, Co²⁺, dodecyl sulfate, and EDTA inhibited its activity. Colloidal chitin was the preferred substrate, with an apparent Michaelis constant (K_m) of 6.66 mg/mL and maximal velocity (V_max) of 90.8 U/mL

Applications for Chitinases

Chitinases are versatile enzymes. They have numerous applications such as agricultural applications, medical applications, as well as biotechnological applications. Those applications will be discussed in detail.

Agriculture applications

In agriculture, chitinases are being used because they exhibit a combating role against pathogenic chitin-containing organisms like fungi, insects, and the eggshells of plant-parasitic nematodes (Malik et al. 2022). It was suggested that chitinases are used as fungal antagonists, confirming their important role as biocontrol agents against fungal plant diseases. Chitinases break down the cell wall of fungi, which is composed of chitin, glucan, and wall proteins (Abdelraouf et al. 2024). They also damage pathogen conidial germination, germ tube elongation, and can damage oospores. Additionally, other effects of chitinases were observed as deformities in the fungal cellularity, damage of the protoplasm, mycelial distortion and lysis, and changing the membrane permeability leading to leakage of intracellular contents (Awad et al. 2017). Table 4 presents a list of various microbial chitinases that have been reported to demonstrate fungicidal, insecticidal, and nematocidal activities. For instance, chitinase derived from Streptomyces enissocaesilis and S. rochei showed antifungal effect against the causal agents of Fusarium wilt (Fusarium oxysporum) and damping-off disease (Rhizoctonia solani) (El-Akshar et al. 2024).

Moreover, the growth of the human (opportunistic) pathogens Candida species, Aspergillus fumigatus, and Cryptococcus neoformans was inhibited by the chitinase produced by Trichoderma viride (Abdelraouf et al. 2024). It is known that invertebrate animal species contain chitin in their exoskeleton, tracheal system, epidermal cuticle, and the eggshell of the nematodes. Hence, chitinases can be used as insecticides and pesticides.

Chitinases are indispensable for biological control because their hydrolytic activity targets chitin, the second most abundant biopolymer, which is integral to the structural integrity of various pests. Specifically, by degrading chitin—a key component of fungal cell walls, insect cuticles, and the peritrophic matrix—chitinases effectively induce the lysis of pathogenic fungi and disrupt essential insect processes like metamorphosis and gut function. This mechanism establishes chitinases as potent, environmentally sound biopesticides and biocontrol agents (Navarro-González et al. 2019). It was found that Penibacillus sp. effectively controlled Helicoverpa armigera larvae by reducing the feeding rate and body weight, which subsequently increased the rate of larval mortality (Singh et al. 2016). Additionally, Stenotrophomonas and Chromobacterium were found to suppress the cyst nematode Globodera rostochiensis (Iqbal and Anwar 2019).

Table 4. Antifungal, Insecticidal, and Nematocidal Actions of Microbial Chitinases

Pharmaceutical applications

Chitinases are characterized by their potent antifungal, antibacterial, and antioxidant properties. Hence, their dominant usages in various medical applications are apparent. Because chitin is a main component of fungal cell walls, chitinase effectively acts as a biocontrol agent by breaking down the cell wall of pathogenic fungi, whether they are active or not (Halder et al. 2013). Chitinase causes damage and deformity of the hyphae and spores by disrupting the fungal cell walls, which causes mycelial lysis (Halder et al. 2013, Al-Rajhi et al. 2022). Chitinases of microbial origin can degrade chitin into chitosan-oligosaccharides with positive charges, which enable them to attack the negatively charged bacterial cell wall, causing its damage, increasing its permeability, leaking of the bacterial cell components, and finally, death of the bacterial cell occurs (Shehata et al. 2018).

The enhanced antimicrobial efficacy of polycationic chitosan against Gram negative bacteria is fundamentally driven by structural differences in the bacterial cell wall. Chitosan is positively charged due to its free amino groups, enabling strong electrostatic interactions. The Gram negative bacteria surface, characterized by lipopolysaccharides (LPS), carries a high overall negative charge. This strong attraction between positive chitosan and negative LPS leads to the crucial disruption and permeabilization of their outer membrane. This breach allows the chitosan to penetrate the cell, resulting in the leakage of intracellular contents and subsequent cell death. In contrast, Gram +ve bacteria, while also negatively charged, possess a significantly thicker, rigid peptidoglycan layer. This robust physical barrier effectively hinders the access and disruptive action of chitosan, rendering Gram positive bacteria less susceptible to the compound (Olicón-Hernández et al., 2015). Table 5 presents the various medical applications of microbial chitinases, including their potential roles as antifungal, antibacterial, anticancer, and antioxidant agents. Chitinases produced by Bacillus haynesii exhibited antifungal activity against Fusarium oxysporum and Penicillium chrysogenum, with mean inhibition zones of 33 mm and 12 mm in diameter, respectively (Govindaraj et al. 2024).

Similarly, microbial chitinases attain their anticancer potential from their ability to interact with cancer-specific polysaccharides containing compounds such as glycoproteins or glycolipids, which are located on the surface of the tumor cells and break down their carbohydrate moieties, causing functional damage and tumor cell death (Pan et al. 2005). However, the precise mechanism of inhibiting the proliferation of cancer cells remains unknown. Some explanations attributed such effects on the proliferation of the cancer cells to the differences in the electrostatic chargers of chitosan-oligosaccharides that may lead to increased cell permeability as above-described and/or alteration of the factor expressions of tumor cells (Liaqat and Eltem 2018). Many lines of cancer cells are being influenced by microbial chitinases, such as breast, lung, colon, bladder, and melanoma (Pan 2012). IC50 values of ChiB and ChiC from Serratia marcescens were found to be 4.63 µM and 2.36 µM, respectively, for the MCF-7 cells (Shrivastava et al. 2024).

Synthesis of antioxidants with potential scavenging effects to eliminate free radicals is accompanied by adverse effects, hence the importance of applying novel biological strategies for reactive oxygen species (ROS) scavenging, chelation of transition metals, and detoxification of antioxidants for free radical elimination (Halder et al. 2013). Chitooligosaccharides, produced by enzymatic hydrolysis of chitin and chitosan, display potent antioxidant activity (Khalil et al. 2017). Their antioxidant effects are attributed to hydroxyl and amino groups, which interact with unstable free radicals, converting them into molecular radicals (Halder et al. 2013). Purified chitinase from Talaromyces funiculosus achieved maximum inhibition of DPPH and ABTS radicals at approximately 57.8% and 63.7%, respectively (El-Beltagi et al. 2022).

Table 5. Medical Applications of Microbial Chitinases

Industrial and Environmental Applications

Protoplast release

Fungal cell wall degradation and protoplast formation are primarily mediated by microbial enzymes. Although non-enzymatic and mechanical methods have also been reported, their practical applications remain limited (Sun et al. 1992). Microbial protoplasts serve as important tools in biochemical, genetic, and physiological studies (Hassan 2014) and are particularly useful for investigating enzyme localization in fungi (Sonawane et al. 2016). Advances in protoplast fusion have enabled genetic manipulation, allowing the combination of genes from different organisms, which facilitates strain improvement, enhances genetic recombination, and contributes to the development of industrially valuable strains (Patil et al. 2013).

Chitinase plays a key role in releasing protoplasts from microbial species whose cell walls contain substantial amounts of chitin. Crude chitinase extracted from Rhizopus stolonifer generated protoplasts of Aspergillus niger, Aspergillus oryzae, Fusarium moniliforme, and Trichoderma viride (Sonawane et al. 2016). In another study, Penicillium ochrochloron chitinase demonstrated high efficacy in generating protoplasts from Aspergillus niger (Patil et al. 2013). Due to the complex composition of chitin and glucans in fungal cell walls, protoplast formation necessitates a mixture of lytic enzymes, since single enzymes show limited activity (Sonawane et al. 2016). In this context, the combination of purified chitinase A from Streptomyces cyaneus with α-1,3-glucanase from Bacillus circulans KA-304 exhibited enhanced protoplast formation activity (Yano et al. 2008). Likewise, chitinase and β-glucanase enzyme complexes demonstrated great protoplast-forming efficiency between Trichoderma harzianum and T. viride (Hassan 2014).

Production of single cell proteins

A substantial amount of chitinous shellfish waste is generated. Thus, seafood waste, which is a byproduct of the shellfish processing industry, is considered a significant challenge (Nirmala 1991). Among the shell waste, Crustacean shell consists of 30 to 40% proteins, 30 to 50% calcium carbonate, and 20 to 30% chitin (Kurita 2006).

Single-cell protein (SCP) represents a valuable protein source and is considered an alternative to fish and soybean meals (Le and Yang, 2019). Chitinase plays a vital role in the production of single-cell proteins SCP by hydrolyzing chitin, which is abundant in shellfish and other chitinous waste, into soluble chitooligosaccharides (COS) and monomeric sugars, primarily N-acetylglucosamine (GlcNAc). The resulting COS serve as readily assimilable carbon and nitrogen sources for various microorganisms, including yeasts and bacteria, which further hydrolyze them into monomeric sugars. These monosaccharides are then metabolized to produce microbial biomass rich in proteins, forming SCP. This enzymatic process provides a sustainable and efficient pathway to convert chitinous waste into high-value protein, supporting the development of a circular bioeconomy (Le and Yang 2019). Chitinases from Penicillium ochrochloron hydrolyzed chitin into GlcNAc, which was then utilized as a substrate for SCP production by Yarrowia lipolytica (Le and Yang 2019). Chitin was enzymatically hydrolyzed by Serratia marcescens QMB1466 chitinase to generate a hydrolysate, later used for yeast single-cell protein production (Revah and Carroad 1981). The SCP is produced from fungal sources, for example, Candida tropicalis, Myrothecium verrucaria, Hansenula polymorpha, and Saccharomyces cerevisiae, which is a major source of chitinase production with over 60% SCP and low nucleic acid contents (Dahiya et al. 2006). Moreover, the generation of SCP using Penicillium ochrochloron was also previously reported (Patil 2014).

Production of chitooligosaccharides

Hydrolysis of chitin by chitinases produces small chitooligomers and chitooligosaccharides (COS), which have diverse applications in agriculture, medicine, pharmaceuticals, and the food industry. COS are insoluble in propanol, ethanol, acetone, ethyl acetate, and butanol, partially soluble in dimethyl sulfoxide (DMSO) and methanol, and fully soluble in water (Liang et al. 2018). In the food industry, COS are used to enhance product quality and as dietary supplements to boost immunity (Martinez et al. 2012). Incorporation of COS into chitosan films for food packaging has been shown to improve antimicrobial properties (Fernandez-de Castro et al. 2016). In the human colon, specific bacteria degrade COS into short-chain fatty acids and other beneficial metabolites, providing probiotic effects (Selenius et al. 2018), while daily administration of 100 mg/kg COS increases Bifidobacterium populations and decreases E. coli levels (Wan et al. 2017).

Pharmaceutical applications of chitinases include antihypertensive, antioxidant, antitumor, wound-healing, antiallergic, and hypocholesterolemic effects, making them suitable for drug delivery, disease treatment, and the production of implants and surgical materials (Rameshthangam et al. 2018). COS also exhibit potent antimicrobial activity against Gram-positive and Gram-negative bacteria, including Staphylococcus aureus, Staphylococcus xylosus, Bacillus cereus, Listeria monocytogenes, Klebsiella pneumoniae, and Proteus vulgaris (Castañeda-Ramírez et al. 2013).

COS shows potent antifungal effects against Aspergillus, Candida, Saccharomyces, and Trichophyton (Muanprasat and Chatsudthipong 2017). These molecules can reduce colonic mucosal inflammation through various mechanisms. They may increase malondialdehyde levels and enhance nitric oxide synthase activity, while decreasing catalase and glutathione levels, and modulating the TNF-α pathway (Bekale et al. 2015). Additionally, COS showed cytotoxic effects against A549 and HCT-116 cell lines, with in vitro IC₅₀ values of 48.6 μg/mL and 1329.9 μg/mL, respectively. In vivo studies in mice revealed a tumor inhibition percentage of up to 58.5% (Zou et al. 2016). In the agricultural field, fungicidal and bactericidal properties against phytopathogens were observed. Additionally, they were used as plant growth regulators, immune boosters, and to improve the tolerance of plant seedlings to salt, heat, and cold stress (Zhang et al. 2019). An increase in the level of the IAA hormone in Brassica napus was seen after treatment with hetero COS, resulting in enhancement in the height of the plant, the number of branches as well as plant biomass. The plant agronomic properties and upregulation of the main genes controlling the signaling pathway were improved by COS treatment (Tang et al. 2022). The germination of wheat seeds was reported to be prompted by COS (Fu et al. 2019). Similarly, the level of glutamate and proline, which contribute to powerful plant growth and cold tolerance enhancement of rice seedlings, was increased by COS treatment (Zhang et al. 2019).

Dye removal

Several chitin-synthesizing microorganisms have been investigated for their dye-removal potential. An innovative and powerful biocomposite absorbent was created by Bacillus subtilis through the bacterial biomass-mediated modification of chitosan. Such absorbents showed a high efficiency in removing the toxic dye of the textile Reactive Orange 16 in aqueous solution (Agha et al. 2025). Similarly, the efficiency of the Brilliant Blue dye removal was improved by Aspergillus niger MK981235, especially after the powder of the carb shells was involved as bioadsorbents (Abdel Wahab et al. 2023). Byproducts resulting from chitinous waste fermentation were observed to enhance the dye-removal potential of Paenibacillus mucilaginosus TKU032, which exhibits strong adsorption capabilities. In case of adding fermented powder of shrimp heads as adsorbent, such capability of adsorption achieved 99% removal of Congo Red and 97% of Red No.7 (Doan et al. 2020). Similarly, using fermented squid pen powder, Bacillus cereus TKU034 achieved up to 99.5% adsorption of various disperse dyes (Liang et al. 2015).

Enhancement Strategies for the Production, Stability, and Activity of Microbial Chitinases

Recombinant expression of microbial chitinases

Chitinase synthesis by genetic engineering and their subsequent expression in various strains of microorganisms represents a promising approach to develop recombinant strains with enhanced overexpressed chitinases and desired functional properties (Yu et al. 2022). Therefore, the genes that are responsible for the synthesis of thermostable chitinase in specific microorganisms can be easily cloned and expressed in different hosts (Sarma et al. 2013). Furthermore, the enzymes exhibit thermostable properties, maintaining correct folding under harsh conditions. They also possess resistance to host cell proteases and, therefore, are not degraded by these proteases (Sarma et al. 2013).

The shared repertoire of chitinase families between bacteria and fungi is a powerful illustration of Horizontal Gene Transfer (HGT), a key non-sexual mechanism driving microbial evolution (Goughenour et al. 2021). Evidence from phylogenetic studies consistently demonstrates that specific fungal chitinase genes exhibit a closer evolutionary relationship to bacterial counterparts, strongly indicating a bacterial origin for these clades (Gonçalves et al. 2016). This genetic exchange provides a substantial adaptive benefit, allowing the recipient organism to effectively break down chitin, which is a crucial structural component in both fungal cell walls and insect exoskeletons (Zhang et al. 2025). The cross-kingdom transfer is hypothesized to occur through various mechanisms, including conjugation-like events, the activity of transposable elements, and the intimate physical proximity within shared ecological niches (Richards et al. 2011). Moreover, the rate of HGT is closely linked to the organism’s ecology, with parasitic and saprotrophic fungi showing elevated gene acquisition due to their constant interaction with bacteria (Liu et al. 2025).

The expression of chitinase genes can be increased by using innovative methods of biotechnology, such as cloning and recombinant technologies. Thus, the production and the activities of the enzymes will be developed and increased. Furthermore, several studies have reported the industrial and agricultural applications of cloned and overexpressed microbial chitinases in heterologous hosts. E. coli BL21 (DE3) successfully expressed the Mtch509 chitinase gene from Microbulbifer thermotolerans, producing a recombinant enzyme with high stability under elevated temperatures and in the presence of high salt concentrations (5 M NaCl) (Lee et al. 2018).

Bacillus subtilis, recognized as GRAS, can synthesize and secrete recombinant proteins extracellularly, although its application is limited by the lack of suitable expression vectors (Heravi et al. 2015). Yeast systems, such as Pichia pastoris, are ideal for heterologous gene expression due to their ease of genetic manipulation; chitinase expressed in P. pastoris reached maximum activity at 50 °C, with activity decreasing to 80% at 60 °C (Kaczmarek et al. 2021).

Similarly, Saccharomyces cerevisiae can efficiently express chitinase genes while performing post-translational modifications; for instance, Thermomyces lanuginosus chitinase expressed in S. cerevisiae exhibited optimal activity at pH 6.5 and 60 °C (Prasad and Palanivelu 2012).

Table 6 summarizes the enhancement of activity of these cloned microbial chitinase genes in various hosts.

Table 6. Cloned Microbial Chitinases from Various Microorganisms for Enhanced Enzyme Stability and Activity

Genetic engineering strategies and directed evolution

To improve the level of expression and the activity of chitinase, certain gene modifications are required. However, at higher pH and temperature conditions, the enzyme stability and selectivity on the substrate may be affected and negatively changed (Okongo et al. 2019). Different methods have been established to obtain modifications in chitinase genes, such as site-directed mutagenesis, directed evolution, and the selection of desired properties (Berini et al. 2018). The process begins under extreme conditions with the isolation of the microbial enzyme, followed by rational mutagenesis and site-directed mutagenesis. For further improvement of the enzyme traits, direct evolution is applied (Sarma et al. 2013).

Table 7. Recombinant Chitinases from Various Microorganisms against Phytopathogens and Pests

The directed evolution methods can be applied to improve the thermal resilience of chitinase synthesized by the fungus Beauveria bassiana and the chitinase gene (Bbchit1) from Erwinia carotovora by DNA shuffling and screening (Fan et al. 2007). After deleting a single nucleotide in the sequence of chitinase encoding in E. coli, site-directed mutagenesis was used, resulting in a recombinant strain that exhibited activity at 90 °C and a pH ranging from 6.0 to 7.5 (Oku and Ishikawa 2006). Furthermore, the enzymatic thermostability and catalytic activities are being enhanced after the chitinase gene (ChiD) of Serratia proteamaculans was manipulated by site-directed mutagenesis (Madhuprakash et al. 2012). Several works were designed for enhancing the plant-protecting activities against plant pathogens and pests by heterologous and homologous overexpression of chitinase, Table 7.

Recombinant chitinases in biocontrol and transgenic plant development

Biocontrol materials, when added to protect plants from phytopathogens, introduce them to high temperatures over an extended period. This highlights the significant importance of recombinant thermostable chitinase (Alves et al. 2018). For example, incorporating the carbohydrate-binding module from Serratia marcescens into Trichoderma atroviride Chi42 created a modified chitinase with enhanced antifungal activity (Matroodi et al. 2013).

Table 8. Disease-Resistant Transgenic Plants by Incorporating Chitinase Genes

Moreover, E. coli BL21 expressed the BhChitA chitinase gene of Bacillus halodurans and the recombinant necrosis-suppressing enzymes produced by Botrytis cinerea on tomato leaves (Ezzine et al. 2024). Similarly, the Sschi61 chitinase gene, synthesized by Streptomyces sampsonii, was successfully expressed by E. coli BL2, whereas the recombinant chitinase inhibited the black spot pathogens of Pestalotiopsis trachicarpicola (Wang et al. 2022).

Incorporation of chitinase genes expressed in bacteria and fungi could successfully help in the production of pathogen-resistant transgenic plants using the above-mentioned techniques. The most popular method for this is plant transformation using Agrobacterium tumefaciens as a vector of chitinase genes. Chitinase genes from Trichoderma species are extensively employed to develop transgenic plants with enhanced pathogen resistance (Table 8).

A subsequent study reported that the expression of recombinant chitinase (CHI) from Phomopsis liquidambaris in Glycine max conferred transgenic plant resistance against head blight disease caused by Fusarium graminearum (Zhu et al. 2022).

Current Limitations and Challenges in Production and Application of Chitinases

Despite their promising potential, microbial chitinases face significant challenges in both research and industrial applications. The most prominent limitations include enzyme denaturation and instability under harsh processing conditions, which reduce their long-term effectiveness (Oyeleye and Normi 2018), Purifying chitinases from the fermentation broth typically involves multiple steps, which can be costly and may result in enzyme activity loss, thereby increasing the overall production expense (Singh et al. 2021). Recent studies suggest the potential of nanoparticles as effective inducers for improving the yield and catalytic efficiency of industrially relevant enzymes, including chitinases (Al-Rajhi et al. 2024). Additionally, the complexity of chitinase–substrate interactions and the need for precise reaction conditions impose constraints on scalability (Eijsink et al. 2008). The widespread use of chitinases, particularly in agriculture, raises concerns about their potential impact on non-target organisms that contain chitin, such as beneficial insects and fungi (Unuofin et al. 2024). This could lead to unintended ecological imbalances. while applications in transgenic plants raise ethical and regulatory concerns regarding environmental safety and consumer acceptance (Hasan et al. 2023).

CONCLUSIONS AND FUTURE PROSPECTS

1. Chitinases are valuable enzymes with broad applications in agriculture, biotechnology, medicine, and waste management. Future research is focused on expanding the functionality of chitinases, including their potential use as food preservatives, immunomodulators, and anti-tumor agents.
2. Advances in genetic engineering and enzyme modification are expected to enhance their stability, activity, and industrial viability. The development of thermostable chitinases through bacterial and fungal sources is a key priority, particularly for applications requiring prolonged enzyme efficiency under extreme conditions.
3. The biocontrol potential of chitinases in agriculture, as well as their medical applications in ophthalmic treatments and microbicides, highlights their diverse utility. Genetic engineering enhances chitinase stability and activity, increasing their industrial viability in extreme conditions.
4. As research progresses, the integration of biotechnology in optimizing chitinase production and function will be essential in making these enzymes more accessible and effective across various industries.

Data Availability Statement

Data is contained within the article.

Conflicts of Interest

The authors declare no conflicts of interest.

Use of Generative AI

During the preparation of this manuscript, the authors used ChatGPT to improve language and readability. After using this tool, the authors reviewed and edited the content as needed and took full responsibility for the content of the published article.

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Article submitted: October 1, 2025; Peer review completed: November 1, 2025; Revised version received: November 4, 2025; Accepted: December 18, 2025; Published: December 29, 2025.

DOI: 10.15376/biores.21.1.Abu-Tahon