Next Article in Journal
Screening of Lipid-Reducing Activity and Cytotoxicity of the Exometabolome from Cyanobacteria
Previous Article in Journal
Altechromone A Ameliorates Inflammatory Bowel Disease by Inhibiting NF-κB and NLRP3 Pathways
Previous Article in Special Issue
Functional Characterization of the First Bona Fide Phytoene Synthase in Red Algae from Pyropia yezoensis
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Marine Drugs
Volume 22
Issue 9
10.3390/md22090411
marinedrugs-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

Enzymes from Fishery and Aquaculture Waste: Research Trends in the Era of Artificial Intelligence and Circular Bio-Economy

by
Zied Khiari
National Research Council of Canada, Aquatic and Crop Resource Development Research Centre, 1411 Oxford Street, Halifax, NS B3H 3Z1, Canada
Mar. Drugs 2024, 22(9), 411; https://doi.org/10.3390/md22090411
Submission received: 28 August 2024 / Revised: 6 September 2024 / Accepted: 7 September 2024 / Published: 10 September 2024
(This article belongs to the Special Issue Enzymes from Marine By-Products and Wastes)
Browse Figures
Versions Notes

Abstract

:
In the era of the blue bio-economy, which promotes the sustainable utilization and exploitation of marine resources for economic growth and development, the fisheries and aquaculture industries still face huge sustainability issues. One of the major challenges of these industries is associated with the generation and management of wastes, which pose a serious threat to human health and the environment if not properly treated. In the best-case scenario, fisheries and aquaculture wastes are processed into low-value commodities such as fishmeal and fish oil. However, this renewable organic biomass contains a number of highly valuable bioproducts, including enzymes, bioactive peptides, as well as functional proteins and polysaccharides. Marine-derived enzymes are known to have unique physical, chemical and catalytic characteristics and are reported to be superior to those from plant and animal origins. Moreover, it has been established that enzymes from marine species possess cold-adapted properties, which makes them interesting from technological, economic and sustainability points of view. Therefore, this review centers around enzymes from fishery and aquaculture waste, with a special focus on proteases, lipases, carbohydrases, chitinases and transglutaminases. Additionally, the use of fishery and aquaculture wastes as a substrate for the production of industrially relevant microbial enzymes is discussed. The application of emerging technologies (i.e., artificial intelligence and machine learning) in microbial enzyme production is also presented.

1. Introduction

As primary industries, both fisheries and aquaculture constitute essential sectors of the economy in many parts of the world [1]. The latest report from the Food and Agricultural Organization (FAO) stated that global fisheries and aquaculture production in 2022 reached an all-time record of 185.4 million tons of aquatic animals (i.e., fish, crustaceans and mollusks) [2]. Statistical data published in the same report also indicated that the aquaculture production in 2022 represented 51% of the total production, surpassing for the first time that from capture fisheries [2]. The recovery of capture fisheries, along with the expansion of aquaculture (which is currently the fastest-growing animal food production sector [3]), is believed to have positive impacts on aquatic animal production, with the latest projections predicting a 10% increase in production by 2032 [2].
The blue bio-economy aims to promote a sustainable utilization of fisheries resources for economic growth and development, along with the preservation of the ocean’s health [4]. However, achieving all the goals of the blue bio-economy could be challenging, mostly due to climate change and unsustainable practices in the fishery and aquaculture sectors. For instance, climate change, on the one hand, has severe negative impacts on the ocean (such as ocean acidification as well as sea level and temperature rises) [5] and could constitute a hindering factor for sustainable ocean development [6]. The fishery and aquaculture industries, on the other hand, still have not addressed the pressing need to operate in a sustainable manner. In this regard, it is known that fishery and aquaculture production are not entirely used for human consumption. In fact, processing fish, shellfish and other marine animals results in significant amounts of waste, which, depending on the species, can reach as high as 50 to 70% of the total production [7,8]. Fishery and aquaculture waste are typically discarded at sea or in landfills and, in the best-case scenario, rendered into fishmeal and fish oil [9,10]. However, under the blue bio-economy concept, in which the sustainable utilization of the ocean and its resources is a priority, it is critical to minimize and/or eliminate waste generation [11]. Due to the abundance of value-added biomolecules (i.e., enzymes, proteins/peptides, polyunsaturated fatty acids, carotenoids, minerals and hydroxyapatite, as well as polysaccharides, including chitin and glycosaminoglycans [7,12,13]), fishery and aquaculture waste could be considered as a renewable source of highly valuable and industrially relevant bioproducts. The development of low-cost, green and circular processes for valorizing and/or upcycling these wastes could decrease the pressure on marine ecosystems, avoid pollution and create sustainable wealth through revenue generation and job creation [7,11].
The aim of this review is to provide an overview of the major enzymes from fishery and aquaculture waste. This review also discusses the use of fishery and aquaculture waste as a substrate for the production of bacterial and fungal enzymes. In addition, the application of emerging technologies (i.e., artificial intelligence and machine learning) with respect to enzyme production using fishery and aquaculture waste as a substrate is presented.

2. Fishery and Aquaculture Waste

During the capture, farming and processing of fish, shellfish and other marine animals, substantial quantities of waste are produced. In capture fisheries, the waste mainly consists of by-catch (i.e., unwanted species that have lower commercial value than the target catch [9,14]). By-catch remains underutilized, largely due to several negatively viewed attributes in terms of color, flavor, texture, size and fat content [15]. Some examples of by-catch in commercial fish capture include butterfish (Peprilus triacanthus), which are a non-targeted species of the squid fishery [16], and Chinook salmon (Oncorhynchus tshawytscha), which are caught in the Bering Sea pollock fishery [17]. In shellfish fisheries (i.e., tropical shrimp), species that are highly susceptible to capture by trawls include ariid catfish (Ariidae), conger eels (Congridae), wrasse (Labridae) and eeltail catfish (Plotosidae) [18]. In the aquaculture industry, mortalities and discarded species are the major sources of waste [19].
When processing fish, shellfish and other marine animals into marketable seafood products, any parts that are not typically regarded as edible (i.e., not used for human consumption) are considered to be waste; this includes the heads, frames, tails, viscera, belly flaps, shells, etc. [20,21]. Figure 1 presents the approximate percentages of waste generated during the processing of fish and shellfish. In general, the percentages of waste relative to the whole fish (based on wet weight) are in the range of 9–25% for the heads, 3–6% for the skins, 12–15% for the frames, 5–14% for the viscera and 3–7% for the belly flaps [8,22,23,24]. For shellfish such as shrimps, the proportion of waste (heads, shells and tails) varies from 40 to 46% with respect to the whole shellfish’s wet weight [25]. For shrimp waste, the heads represent 72.5%, while the shells with tails represent 27.5% of the total waste [25]. For other shellfish species, such as crabs, the shells correspond to the waste generated during processing and represent about 55% of the whole crab’s wet weight [26].
Fishery and aquaculture waste (both by-catch and the processing discards) are highly unstable due to their high organic matter and lipid content as well as their strong proteolytic enzyme activity. Recently, there has been a growing interest in valorizing fishery and aquaculture waste by recovering value-added products instead of converting them into low-value commodities (such as fishmeal, fish oil and compost/fertilizer) [27]. Numerous research studies indicated that fishery and aquaculture waste contains functional and bioactive compounds, such as a range of enzymes, proteins, lipids rich in omega-3 fatty acids, pigments, chitin, glycosaminoglycans, vitamins and other biomolecules with high market values [7,21,22]. Enzymes from fish, shellfish and other marine animals, in particular, have been reported to possess high catalytic activities at relatively low concentrations [7,22]. They also exhibit cold-adapted properties (i.e., active at low temperatures and unstable at higher temperatures) and remain stable over a wide range of pHs [7,22]. These properties make marine-derived enzymes very interesting from an economic point of view by lowering the energy requirement for enzyme-based processes [7].

3. Overview of Major Industrially Relevant Enzymes from Fishery and Aquaculture Waste

The internal organs of the fish, shellfish and other marine animals contain different types of enzymes. These marine-derived enzymes have been found to possess cold-adapted properties (i.e., active between 0 and 30 °C but unstable at temperatures higher than 50 °C [7,28]). The mechanisms of cold adaptation in marine enzymes have been recently reviewed by Khiari [7]. Briefly, cold adaptation is believed to be associated with a higher molecular flexibility that is not typically observed with mesophilic and thermophilic enzymes [7]. In addition, cold-adapted enzymes have been reported to have a smaller number of hydrogen bonds, a structure that is less densely packed, a higher surface hydrophilicity and a greater number of methionine residues [7]. It is worth noting that cold-adapted enzymes in fish species from the Northern Seas (i.e., Arctic region) are different than those from the Southern Seas (i.e., Antarctic region). In this regard, enzymes from Antarctic fish have been shown to be active at extremely low temperatures and are, in general, more stable in the presence of high salt concentrations and metal ions compared to those from Arctic fish [29,30].
Cold-adapted enzymes are generally preferred over mesophilic and thermophilic counterparts. Unlike mesophilic enzymes, cold-adapted enzymes are highly active at low temperatures (20–25 °C) and are able to preserve half of their maximum activity at even lower temperatures (0–10 °C); however, they are easily inactivated through moderate thermal treatment [30]. A number of studies investigated the kinetic properties of cold-adapted fish enzymes compared to mesophilic mammalian enzymes. Some examples of the kinetic properties of selected fish and mammalian enzymes are presented in Table 1. These kinetic studies proved that fish enzymes possess significantly higher catalytic efficiency at low temperatures compared to mammalian enzymes, as indicated by the kcat/Km values.
Owing to their unique characteristics, cold-adapted enzymes from fish, shellfish and other marine animals have promising prospects for several industrial applications. This includes the food industry (i.e., processing at low temperatures slows the growth of pathogenic microorganisms [which subsequently increases food safety] and reduces the occurrence of oxidation reactions [which subsequently improves food quality]), detergent industry (i.e., cold-adapted enzymes are suitable for laundering in cold water), bioremediation (i.e., cold-adapted enzymes could be applied to decontaminate soils and waters in cold regions) as well as green chemistry (i.e., cold-adapted enzymes could be used for the synthesis of heat-labile chemicals) [34].

3.1. Proteases

Proteases (i.e., peptide-bond hydrolases) are enzymes that catalyze the cleavage of peptide bonds in both proteins and peptides. Peptidases are types of proteases that act on peptides and break them down into amino acids [35]. Generally, peptidases can either act on peptide bonds away from the protein termini (i.e., within the protein molecule, and in this case, the protease is termed endopeptidases) or cleave peptide bonds only near the C- or N-terminus (i.e., at a free terminus and, in this case, the protease is termed exopeptidases) [35]. Depending on the amino acid residue involved in the nucleophilic attack of the peptide-bond in the substrate, proteases are classified into six catalytic types: serine proteases, cysteine proteases, glutamic proteases, aspartic proteases, threonine proteases and metalloproteases [35,36].
In fish, proteolytic enzymes are mostly found in viscera, specifically in the stomach, pancreas, pyloric caeca and intestine [37], while in shellfish and other marine animals, digestive proteases are mainly found in the hepatopancreas [38]. Four major groups of proteolytic enzymes are found in fishery and aquaculture waste: aspartyl proteases, serine proteases, cysteine proteases and metalloproteases [39]. Aspartyl proteases (or acidic proteases) are found in the stomach and include pepsin, chymosin and gastricsin [40]. Serine proteases (or alkaline proteases) are found in the pyloric ceca and small intestine and include trypsin, chymotrypsin and elastase [7,40]. Cysteine proteases (or thiol proteases) are typically found in the intestine and include cathepsin [40]. Metalloproteases are proteases that require the presence of bound divalent cations to perform proteolytic hydrolysis and are usually found in muscle tissues [40] but have also been detected in the digestive tracts of marine animals (such as sea cucumber) [41], in fish viscera [42] as well as in fish heads [43].
Figure 2 shows the structures of four representative proteases (pepsin, trypsin, chymotrypsin and elastase).
It is worth noting that the serine endopeptidases found in shellfish (i.e., crustacea) are called brachyurins [48]. Brachyurins comprise three distinctive types: (i) brachyurins Ia, which possess a wide specificity similar to that of trypsin, chymotrypsin, elastase and collagenase; (ii) brachyurins Ib, which possess a wide specificity but a reduced activity toward trypsin substrates; and (iii) brachyurins II, which are trypsin-like proteases [49].
In fishery and aquaculture waste, fish viscera [50], shellfish hepatopancreas [51] and the digestive tract in marine animals [41] represent the major sources of enzymes. Table 2 shows some examples of the proteases recovered from fishery and aquaculture waste.

3.2. Lipases

Lipases are lipolytic enzymes and comprise both triglyceride lipases and phospholipases [91]. Triglyceride lipases catalyze the hydrolysis of triglycerides, while phospholipases catalyze the hydrolysis of phospholipids [91]. In addition to lipid hydrolysis, lipases can also catalyze a number of other reactions, including esterification, transesterification, interesterification, acidolysis, aminolysis and alcoholysis [92,93]. Lipases are widely used in biotechnology due to their high specificity and milder reaction requirements [93].
During digestive lipolysis, two neutral lipid-hydrolyzing enzymes are released from the pancreas: pancreatic lipase and bile salt-dependent lipase [94]. Pancreatic lipases, also called triacylglycerol acylhydrolases, catalyze the hydrolysis of the carboxyl ester bonds of acylglycerols, converting them into diglycerides and subsequently into monoglycerides and free fatty acids [95]. Bile salt-dependent lipases, also called carboxyl ester lipase or cholesterol esterase, catalyze the hydrolysis of the carboxyl ester bonds of cholesterol esters and fat-soluble vitamin esters in addition to those of acylglycerols [94].
A number of studies have focused on lipases from fish, shellfish and other marine animals; however, marine lipases are relatively less studied compared to marine proteases. Table 3 shows some examples of the fishery and aquaculture waste used as sources of lipases.
Most recently, Liu et al. purified novel salt-tolerant, organic solvent-stable, and bile salt-activated lipase from the viscera of golden pompano (Trachinotus ovatus) [104]. Lipases have also been recovered from shellfish such as crab (Carcinus mediterraneus) [105,106], shrimp (Penaeus vannamei) [107,108] and the marine gastropod mollusk (Hexaplex trunculus) [109].
Among the phospholipases, phospholipase A2 (also called phosphatide 2-acyl-hydrolase), which catalyzes the hydrolysis of the ester bond at the sn-2 position of glycerophospholipids [110], is the most widely studied phospholipase [111]. The majority of phospholipase activity in fish has been observed in muscle, such as in pollock [112], Atlantic cod [113] and cod [111]. However, phospholipase activity has also been reported in fishery and aquaculture waste. For instance, phospholipase has been purified from the pyloric caeca of starfish (Asterina pectinifera) [114] and red sea bream (Pagrus major) [115]. Phospholipase has also been found in red sea bream’s gills [116] and hepatopancreas [117], as well as crab (Carcinus mediterraneus) digestive glands (i.e., hepatopancreas) [118].

3.3. Carbohydrases

A number of carbohydrases (i.e., carbohydrate-digesting enzymes) have been reported to be present in fish, and it is believed that the pyloric caeca, pancreas and intestinal mucosa are the main sources of these enzymes [119]. Specifically, the carbohydrate-digesting enzymes α-amylase, α-glucosidase, β-glucosidase and β-galactosidase have been detected in several fish species [120,121,122]. Amylase, the enzyme responsible for hydrolyzing starch to maltose, is by far the most studied amylolytic enzyme in fish. In this regard, Munilla-Mordn and Saborido-Rey assessed the amylase activity in the gut of sea bream (Sparus aurata), turbot (Scophthalmus maximus) and redfish (Sebastes mentella) [123]. Their findings indicated that the maximum activity of amylase from sea bream and turbot was around the neutral pH (7.0–7.5), while that from redfish was at acidic pH values (pH 4.5–5.0) [123]. The optimum temperature for the activity of the extracted amylases ranged between 35 and 45 °C for all the studied fish species [123]. Another study by Fish found that amylase was present in both perch and tilapia [124]. In tilapia, amylase was detected throughout the alimentary canal and mostly in the intestine, while in perch, amylase occurred in the pancreatic secretions [124].
Hidalgo et al. studied the amylase activity in six species of fish: rainbow trout (Oncorhynchus mykiss), sea bream (Sparus aurata), European eel (Anguilla anguilla), common carp (Cyprinus carpio), goldfish (Carassius auratus) and tench (Tinca tinca) [125]. Their results indicated that the omnivorous species presented higher amylase activity than the carnivores, and trout possessed the lowest amylase activity [125]. A similar observation of poor amylase activity in trout has been reported by Spannhof and Plantikow [126]. Skea et al. examined the amylolytic activity in three marine fish: butterfish (Odax pullus), silver drummer (Kyphosus sydneyanus) and marblefish (Aplodactylus arctidens) [127]. Their results also showed that the amylase activity was highest in the anterior gut wall tissue extracts from all three species of fish and decreased in the following order: marblefish followed by silver drummer and butterfish, respectively [127].
Amylase has also been reported in crustaceans. In this respect, amylase has been isolated from blue crab (Portunus segnis) viscera [128] and the midgut gland of three shrimp species (wild Farfantepenaeus subtilis, wild Litopenaeus schmitti, as well as farmed Litopenaeus vannamei) [129]. The results from the latter study indicated that the total amylolytic activity in farmed shrimp was three times higher than that from wild shrimp [129].

3.4. Chitinolytic Enzymes

Chitin is an abundant mucopolysaccharide in crustaceans [10]. Structurally, chitin is a β-(1,4)-linked polymer of N-acetyl-D-glucosamine (GlcNAc) [130]. Chitin-degrading enzymes hydrolyze chitin into GlcNAc through the synergistic action of endo-type chitinase (EC 3.2.1.14) and exo-type β-N-acetyl-hexosaminidase (EC 3.2.1.52) [131]. For instance, chitinases hydrolyze the β-1,4 glycosidic bonds of chitin to produce N-acetyl-chito-oligosaccharides (NACOs), which are subsequently degraded into GlcNAc units by β-N-acetyl-hexosaminidase [132]. High levels of chitinase activity were reported in fish stomachs [133,134], while low or negligible activities were observed in the pyloric caeca, intestines and livers of fish [132], whereas low levels of β-N-acetyl-hexosaminidase activity were reported in fish stomachs, but high levels were observed in the intestines and/or the pyloric caeca [132]. This suggests that chitinases start the hydrolysis of chitin in the stomach, which is further completely decomposed to soluble monomers of N-acetyl-D-glucosamine in the intestine and/or pyloric caeca by the action of β-N-acetyl-hexosaminidases [132]. The presence of both chitinase and β-N-acetyl-hexosaminidase in fish, shellfish and other marine animals could also suggest that these species are able to degrade chitin to penetrate prey exoskeletons as well as to turn them into nutrients (i.e., monosaccharide, N-acetyl-D-glucosamine) [134]. Chitinolytic activities have been observed in different parts of fish, shellfish and marine animals (stomach, intestine, pyloric caeca, liver, spleen, kidney, heart and gill [133]), which are generally considered to be waste [7]. Table 4 provides a summary of marine species that contain chitinolytic enzymes.
Chitosan, a β-1,4-linked polysaccharide consisting of glucosamine residues, is obtained through the enzymatic or chemical deacetylation of chitin [135]. To enhance its functionalities, chitosan is typically depolymerized to generate chitosan oligomers [136]. Chitosan depolymerization can be achieved through chemical (using acid) or enzymatic (using chitosanase) processes, with the enzymatic depolymerization process being preferable to the chemical depolymerization process [136]. The enzymatic depolymerization of chitosan is based on the hydrolytic action of chitosanases, which cleave the β-(1,4)-linked glycosidic bonds in chitosan. Chitosanases (EC 3.2.1.132) are largely found in microorganisms; however, they have also been detected in fish, shellfish and other marine animals. For example, Baehaki et al. [137], Affes et al. [138] and Yao et al. [139] reported the presence of chitosanases in the digestive tract of snakehead fish (Channa striata), the viscera of blue crab (Portunus segnis) and the digestive tract of sea cucumber (Stichopus japonicus), respectively.
Table 4. Examples of marine species containing chitinolytic enzymes.
Table 4. Examples of marine species containing chitinolytic enzymes.
Chitinolytic EnzymeEC NumberMarine Species
ChitinaseEC 3.2.1.14Silver croaker (Pennahia argentatus) [140,141], chub mackerel (Scomber japonicus) [140,142], Kuruma prawn (Penaeus japonicus) [143], Atlantic salmon (Salmo salar) [144], threeline grunt (Parapristipoma trilineatum) [145], fat greenling (Hexagrammos otakii) [142], marbled rockfish (Sebastiscus marmoratus) [146,147], Japanese sardine (Sardinops melanostichtus) [148], red sea bream (Pagrus major) [149], Japanese eel (Anguilla japonica) [150], Chinese black sleeper (Bostrychus sinensis) [151], red scorpionfish (Scorpaena scrofa) [152], Nile tilapia (Oreochromis niloticus) [153], coelacanth (Latimeria chalumnae) [154], red king crab (Paralithodes camtschaticus) [155], fiddler crab (Uca pugilator) [156], golden cuttlefish (Sepia esculenta) [157], squid (Ommas trephes sloani pacificus) [158], octopus (Polypus dofleini) [158], mussel (Mytilus edulis) [158] and octopus (Eledone cirrhosa) [158]
β-N-acetyl-hexosaminidaseEC 3.2.1.52Marbled rockfish (Sebastiscus marmoratus) [147], cuttlefish (Sepia esculenta) [157,158], mussel (Mytilus edulis) [158], Japanese common squid (Todarodes pacificus) [158], flying squid (Ommastrephes batrami) [158], arrow squid (Loligo bleekeri) [158], brown sole (Limanda herzensteini) [132], European carp (Cyprinus carpio) [132], Pacific saury (Cololabis saira) [132], Atlantic bluefin tuna (Thunnus thynnus) [132], Pacific cod (Gadus macrocephalus) [132], Pacific herring (Clupea pallasii) [132], threeline grunt (Parapristipoma trilineatum) [132], chub mackerel (Scomber japonicus) [132], sardine (Sardinops melanostictus) [132], fat greenling (Hexagrammos otakii) [132], Japanese amberjack (Seriola quinqueradiata) [132], sea bass (Lateolabrax japonicus) [132] and red sea bream (Pagrus major) [132]
EC number: enzyme commission number.

3.5. Transglutaminases

Transglutaminases, also called protein-glutamine gamma-glutamyl-transferase, are Ca2+-dependent enzymes that belong to a class of transferases and catalyze the acyl-transfer between the glutamine residues and a wide variety of primary amines (i.e., catalyze the formation of cross-links between proteins) [159].
Transglutaminase activity has been mostly found in the muscles of marine and freshwater fish as well as in the muscle tissues of shellfish and other marine animals [160,161]. In this regard, Nozawa et al. extracted and characterized transglutaminases from the muscles of six marine species: scallop (Patinopecten yessoensis), Botan shrimp (Pandalus nipponensis), squid (Todarodes pacificus), carp (Cyprinus carpio), rainbow trout (Oncorhynchus mykiss) and Atka mackerel (Pleurogrammus azonus) [160]. Binsi and Shamasundar isolated and characterized transglutaminases from four different fish species (bigeye snapper, Indian oil sardine, tilapia and common carp) [162]. Transglutaminases were also extracted from the tissues of Tropical tilapia (Oreochromis niloticus) [163] and Japanese oysters (Crassostrea gigas) [164]. Figure 3 shows the structure of a representative fish transglutaminase.
Presently, the majority of research on transglutaminases from marine species has focused on isolating the enzyme from muscle tissues. Only a limited number of studies investigated the recovery of transglutaminases from fishery and aquaculture waste. Examples of fishery and aquaculture waste from which transglutaminase was extracted include the red sea bream (Pagrus major) liver [166,167], walleye pollock (Theragra chalcogramma) liver [168], squid (Todarodes pacificus) gills [169] as well as horseshoe crab (Limulus polyphemus) hemolymph [170,171].

4. Utilization of Fishery and Aquaculture Waste for Production of Microbial Enzymes

The nitrogen source is by far the most expensive component of microbial growth media [172]. Fishery and aquaculture waste contain high protein levels and well-balanced amino acid profiles, making them ideal, low-cost sources of nutritional substrate for the growth of enzyme-producing microorganisms [173]. Numerous studies investigated the possibility of using fish processing waste as a substrate for bacterial and fungal enzyme production [174,175]. The available data indicate that fishery and aquaculture waste, either entirely (by-catch and mortality) or consisting of parts (heads, viscera, shells, etc.), have been successfully used as a substrate for enzyme production [174]. Many strains were reported to directly utilize fishery and aquaculture waste to meet their carbon and nitrogen requirements [176]. Examples of fishery and aquaculture waste that have been used as sources of nitrogen for bacterial and fungal enzyme production are listed in Table 5.
Microbial enzymes are typically produced through fermentation. This bioprocess can either be based on submerged fermentation (i.e., fermentation that occurs in a liquid medium with free water available for the microorganisms) or solid-state-fermentation (i.e., fermentation that occurs in a solid medium without or with very little free water available for the microorganisms) [209]. Several factors have been reported to affect the microbial enzyme production bioprocesses. This includes the microbial strain (which must be capable of producing a high yield of the target enzyme), the concentration and the type of nutrients in the culture medium, the pH, the temperature and other bioprocess parameters, such as agitation and aeration (especially for submerged-type fermentations) [209].
The global industrial enzyme market reached USD 6.1 billion in 2023 and is projected to attain USD 8.2 billion by 2030 [210]. Industries, including leather, paper, textile, biofuel, and, to a lesser degree, food industries, are among the major users of industrial enzymes [211]. Innovative research and development (R&D) efforts have been made, aiming at increasing enzyme production at a lesser cost by discovering new enzyme-producing microbial strains, improving the current available microbial strains and/or developing new microbial growth media [211]. Knowing that the culture media represent around 30% of the total cost of fermentation [212], the use of fishery and aquaculture waste as substitutes for the current commercial growth media could, therefore, significantly improve the economics of the entire enzyme production bioprocess. This approach not only upcycles fishery and aquaculture waste but also lowers enzyme production costs by offering low-cost microbial growth media (i.e., fishery and aquaculture waste-based media vs. industrially produced media). This, in turn, can boost the sustainability of these industries (i.e., seafood processing, enzyme production and enzyme biotechnology).
From the examples listed in Table 5, there is a clear indication that fishery and aquaculture waste are suitable as a substrate for microbial enzyme production. However, the utilization of microbial enzymes could face safety and regulatory concerns. For instance, as highlighted in Table 5, many enzymes are produced by microorganisms that could pose potential safety and health issues, such as being opportunistic pathogens to humans and/or non-human species, i.e., terrestrial and aquatic/marine animals. In addition, toxic substances (i.e., bacterial toxins and mycotoxins) could occur in the final enzyme preparations/isolates [213]. These drawbacks could make the approach of microbial enzyme production less attractive than the direct extraction of enzymes from the waste unless the enzyme-producing microorganisms are safe.

5. Application of Artificial Intelligence and Machine Learning in Microbial Enzyme Production Using Fishery and Aquaculture Waste as a Substrate

Artificial intelligence (AI) is a revolutionary technology in which computers have the ability to carry out intelligent tasks faster than humans can perform [214]. Machine learning (ML), a discipline of AI, relies on algorithms that receive and analyze data (input) and predict values (output) [215]. ML can be classified into four categories: supervised, unsupervised, semi-supervised and reinforcement learning. In supervised learning, the ML algorithms take direct feedback for the prediction [216]. In this case, the ML algorithm develops a function by mapping input variables from the training data into the desired output [217]. Artificial neural network (ANN), naive Bayes (NB) k-nearest neighbors (kNN), digital twin (DT), support vector machines (SVM), logistic regression (LR), etc., are among the most popular algorithms of supervised learning [216]. In unsupervised learning, ML algorithms do not take any feedback for the prediction [216], and in this case, ML algorithms discover patterns and structures in unlabeled data [218]. K-mean, as well as the self-organized model (SOM), principal component analysis (PCA), latent Dirichlet allocation (LDA), etc., are among the most well-known unsupervised learning algorithms [216]. The semi-supervised learning combines both labeled and unlabeled data to generate a function [219]. Reinforcement learning acts through a policy-based platform [217] and aims to achieve a goal by taking the appropriate measures to maximize the reward [220].
Traditionally, the optimization of microbial enzyme production was heavily based on the one-factor-at-a-time (OFAT) approach. However, OFAT can be inefficient and unreliable and can lead to false optimal conditions [221]. Recently, ML techniques have gained more relevance in the field of enzyme production and have been applied to determine the relationships between the input data (process conditions, such as the pH, temperature, substrate concentration, flow rates, etc.) and output variables (such as enzyme production yields) [215,222]. Artificial neural networks (ANN) and genetic algorithms (GAs) are among the most widely used AI-based techniques for the optimization of microbial enzyme production. ANNs are computer programs that mimic the learning ability of the human brain [223,224]. The ANN has two major advantages: the ability to work with large datasets and the ability to recognize complex patterns [225,226]. GAs, on the other hand, are random global search methods that imitate the process of mutation [226]. GAs simultaneously evaluate several parameters, and the search is based on the genetic selection principle (i.e., survival of the fittest) [227].
AI-based approaches have been applied to the production of bacterial and fungal enzymes using different agro-industrial wastes. For example, Fernández Núñez et al. investigated the production of amylases by Rhizopus oligosporus grown on wheat bran, sugarcane bagasse and soybean meal (either separately or combined) using an artificial intelligence approach [228]. Singh et al. optimized the production of glucansucrase by Leuconostoc dextranicum using an ANN and GA [229]. Bezerra et al. also optimized the production of cellulase by Trichoderma stromaticum grown on peach-palm wastes using an ANN and GA [230]. De Farias Silva et al. applied machine learning techniques (support vector machine (SVM) and ANN) to predict the yield of alginate lyase production by Cunninghamella echinulate grown on microalgae [231].
When it comes to the application of AI- and ML-based methods for the optimization of enzyme production using fishery and aquaculture waste as a substrate, only a limited number of studies have been carried out so far. For instance, Suryawanshi and Eswari optimized, through an ANN, the production of chitinase by Thermomyces lanuginosus using colloidal chitin extracted from shrimp shells [232,233]. Ekpenyong et al. optimized, through the application of the manta ray foraging optimization (MRFO) algorithm, the production of an acidic peptidase by an acidophilic Bacillus cereus strain grown using a mixture of yam peels and fish processing waste [234].
Despite the numerous advantages of AI- and ML-based techniques, the number of published studies related to applying these revolutionary approaches for improving enzyme production using fishery and aquaculture waste is still scarce. The difficulties of standardizing raw materials, as well as the lack of data due to the high diversity of marine species, could explain the scarcity of this type of study. In fact, fishery and aquaculture waste are commonly generated from multiple sources and have different properties, which creates challenges in producing standardized datasets for the ML models. Therefore, more studies are needed to fill this knowledge gap.

6. Concluding Remarks

There is an increasing need for the fishery and aquaculture industries to adopt a zero-waste approach through the complete utilization of fishery and aquaculture production, the reduction of waste generation and the valorization of the waste produced. Upcycling and/or valorizing fishery and aquaculture waste not only reduces their severe environmental impact but also increases their economic value. Within this context, the present review highlighted the major industrially relevant enzymes that are highly valuable bioproducts of fishery and aquaculture waste. The currently available scientific evidence indicates that fishery and aquaculture waste contain enzymes with unique characteristics compared to their mammalian counterparts. Specifically, marine-derived enzymes are known to possess greater enzymatic activity at lower temperatures combined with their reduced stability at higher temperatures. These properties are preferable from technological, economic and sustainability points of view since carrying out enzymatic processes at lower temperatures offers several advantages (i.e., cost reduction, energy savings, etc.). In addition, it has been confirmed that fishery and aquaculture waste represent low-cost, renewable sources of nutritional substrate for the growth of enzyme-producing microorganisms. However, enzymes produced by microorganisms could pose potential safety and health issues, making the microbial enzyme production approach unattractive unless safe enzyme-producing microorganisms are used. Extracting enzymes from fishery and aquaculture waste and/or turning the waste into a culture media for enzyme-producing microorganisms could potentially make these industries more circular, boost the bio-economy principles and further expand the field of microbial enzyme production. The application of artificial intelligence and machine learning technologies could also play a major role in advancing these fields. However, the real challenges facing the successful recovery/production of enzymes from fishery and aquaculture waste are associated with the inconsistent availability of the wastes as well as the seasonal variations in enzyme activities.

Funding

This work was internally funded by the National Research Council Canada (NRC) Oceans Program: OCN-113-1.

Acknowledgments

The financial support given by the National Research Council Canada (NRC) Oceans Program is gratefully acknowledged. The review by Joerg Behnke is highly appreciated. The author would also like to thank the anonymous reviewers for their constructive comments and valuable suggestions. This is an ACRD publication, no. 58522.

Conflicts of Interest

The author declares no conflict of interest.

References

  1. Dyck, A.J.; Sumaila, U.R. Economic Impact of Ocean Fish Populations in the Global Fishery. J. Bioeconomics 2010, 12, 227–243. [Google Scholar] [CrossRef]
  2. FAO. The State of World Fisheries and Aquaculture 2024—Blue Transformation in Action; FAO: Rome, Italy, 2024. [Google Scholar]
  3. Anderson, J.L.; Asche, F.; Garlock, T.; Chu, J. Aquaculture: Its Role in the Future of Food. In World Agricultural Resources and Food Security; Frontiers of Economics and Globalization; Emerald Publishing Limited: Bingley, UK, 2017; Volume 17, pp. 159–173. ISBN 978-1-78714-515-3/978-1-78714-516-0. [Google Scholar]
  4. Upadhyay, D.K.; Mishra, M. Blue Economy: Emerging Global Trends and India’s Multilateral Cooperation. Marit. Aff. 2020, 16, 30–45. [Google Scholar] [CrossRef]
  5. Sari, D.A.A.; Muslimah, S. Blue Economy Policy for Sustainable Fisheries in Indonesia. IOP Conf. Ser. Earth Environ. Sci. 2020, 423, 012051. [Google Scholar] [CrossRef]
  6. Daly, J.; Knott, C.; Keogh, P.; Singh, G.G. Changing Climates in a Blue Economy: Assessing the Climate-Responsiveness of Canadian Fisheries and Oceans Policy. Mar. Policy 2021, 131, 104623. [Google Scholar] [CrossRef]
  7. Khiari, Z. Sustainable Upcycling of Fisheries and Aquaculture Wastes Using Fish-Derived Cold-Adapted Proteases. Front. Nutr. 2022, 9, 875697. [Google Scholar]
  8. Välimaa, A.L.; Mäkinen, S.; Mattila, P.; Marnila, P.; Pihlanto, A.; Mäki, M.; Hiidenhovi, J. Fish and Fish Side Streams Are Valuable Sources of High-Value Components. Food Qual. Saf. 2019, 3, 209–226. [Google Scholar]
  9. Khiari, Z.; Mason, B. Comparative Dynamics of Fish By-Catch Hydrolysis through Chemical and Microbial Methods. LWT 2018, 97, 135–143. [Google Scholar] [CrossRef]
  10. Khiari, Z.; Kelloway, S.; Mason, B. Turning Invasive Green Crab (Carcinus maenas) into Opportunity: Recovery of Chitin and Protein Isolate Through Isoelectric Solubilization/Precipitation. Waste Biomass Valorization 2020, 11, 133–142. [Google Scholar] [CrossRef]
  11. Veríssimo, N.V.; Mussagy, C.U.; Oshiro, A.A.; Mendonça, C.M.N.; Santos-Ebinuma, V.D.C.; Pessoa, A.; Oliveira, R.P.D.S.; Pereira, J.F.B. From Green to Blue Economy: Marine Biorefineries for a Sustainable Ocean-Based Economy. Green Chem. 2021, 23, 9377–9400. [Google Scholar] [CrossRef]
  12. Khiari, Z.; Rico, D.; Martin-Diana, A.B.; Barry-Ryan, C. Structure Elucidation of ACE-Inhibitory and Antithrombotic Peptides Isolated from Mackerel Skin Gelatine Hydrolysates. J. Sci. Food Agric. 2014, 94, 1663–1671. [Google Scholar] [CrossRef]
  13. Singh, S.; Negi, T.; Sagar, N.A.; Kumar, Y.; Tarafdar, A.; Sirohi, R.; Sindhu, R.; Pandey, A. Sustainable Processes for Treatment and Management of Seafood Solid Waste. Sci. Total Environ. 2022, 817, 152951. [Google Scholar] [CrossRef] [PubMed]
  14. Pons, M.; Watson, J.T.; Ovando, D.; Andraka, S.; Brodie, S.; Domingo, A.; Fitchett, M.; Forselledo, R.; Hall, M.; Hazen, E.L.; et al. Trade-Offs between Bycatch and Target Catches in Static versus Dynamic Fishery Closures. Proc. Natl. Acad. Sci. USA 2022, 119, e2114508119. [Google Scholar] [CrossRef] [PubMed]
  15. Venugopa, V.; Shahidi, F. Value-Added Products from Underutilized Fish Species. Crit. Rev. Food Sci. Nutr. 1995, 35, 431–453. [Google Scholar] [CrossRef] [PubMed]
  16. Patrick, W.S.; Benaka, L.R. Estimating the Economic Impacts of Bycatch in U.S. Commercial Fisheries. Mar. Policy 2013, 38, 470–475. [Google Scholar] [CrossRef]
  17. Sugihara, G.; Criddle, K.R.; Ye, H.; Lee, A.; Pao, G.; James, C.; Saberski, E.; Giron-Nava, A. Comprehensive Incentives for Reducing Chinook Salmon Bycatch in the Bering Sea Walleye Pollock Fishery: Individual Tradable Encounter Credits. Reg. Stud. Mar. Sci. 2018, 22, 70–81. [Google Scholar] [CrossRef]
  18. Stobutzki, I.; Miller, M.; Brewer, D. Sustainability of Fishery Bycatch: A Process for Assessing Highly Diverse and Numerous Bycatch. Environ. Conserv. 2001, 28, 167–181. [Google Scholar] [CrossRef]
  19. Cristiano, S.; Baarset, H.; Bruckner, C.; Johansen, J.; Pastres, R. Innovative Options for the Reuse and Valorisation of Aquaculture Sludge and Fish Mortalities: Sustainability Evaluation through Life-Cycle Assessment. J. Clean. Prod. 2022, 352, 131613. [Google Scholar] [CrossRef]
  20. Khiari, Z. Recent Developments in Bio-Ink Formulations Using Marine-Derived Biomaterials for Three-Dimensional (3D) Bioprinting. Mar. Drugs 2024, 22, 134. [Google Scholar] [CrossRef]
  21. Caruso, G.; Floris, R.; Serangeli, C.; Di Paola, L. Fishery Wastes as a Yet Undiscovered Treasure from the Sea: Biomolecules Sources, Extraction Methods and Valorization. Mar. Drugs 2020, 18, 622. [Google Scholar] [CrossRef]
  22. Ghaly, A.E.; Ramakrishnan, V.V.; Brooks, M.S.; Budge, S.M.; Dave, D. Fish Processing Wastes as a Potential Source of Proteins, Amino Acids and Oils: A Critical Review. J. Microb. Biochem. Technol. 2013, 5, 107–129. [Google Scholar] [CrossRef]
  23. He, S.; Franco, C.; Zhang, W. Characterisation of Processing Wastes of Atlantic Salmon (Salmo salar) and Yellowtail Kingfish (Seriola lalandi) Harvested in Australia. Int. J. Food Sci. Technol. 2011, 46, 1898–1904. [Google Scholar] [CrossRef]
  24. Ketnawa, S.; Benjakul, S.; Martínez-Alvarez, O.; Rawdkuen, S. Three-Phase Partitioning and Proteins Hydrolysis Patterns of Alkaline Proteases Derived from Fish Viscera. Sep. Purif. Technol. 2014, 132, 174–181. [Google Scholar] [CrossRef]
  25. Ngoan, L.D.; Lindberg, J.E.; Ogle, B.; Thomke, S. Anatomical Proportions and Chemical and Amino Acid Composition of Common Shrimp Species in Central Vietnam. Asian-Australas. J. Anim. Sci. 2000, 13, 1422–1428. [Google Scholar] [CrossRef]
  26. Parthiban, F.; Balasundari, S.; Gopalakannan, A.; Rathnakumar, K.; Felix, S. Comparison of the Quality of Chitin and Chitosan from Shrimp, Crab and Squilla Waste. Curr. World Environ. 2017, 12, 670–677. [Google Scholar] [CrossRef]
  27. Nakkarike, S.M.; Narayan, B.; Hosokawa, M.; Miyashita, K. Value Addition to Seafood Processing Discards. In Handbook of Seafood Quality, Safety and Health Applications; Wiley: Hoboken, NJ, USA, 2010. [Google Scholar]
  28. Wang, G.x.; Gao, Y.; Hu, B.; Lu, X.l.; Liu, X.y.; Jiao, B.h. A Novel Cold-Adapted β-Galactosidase Isolated from Halomonas Sp. S62: Gene Cloning, Purification and Enzymatic Characterization. World J. Microbiol. Biotechnol. 2013, 29, 1473–1480. [Google Scholar] [CrossRef]
  29. Berthelot, C.; Clarke, J.; Desvignes, T.; William Detrich, H.; Flicek, P.; Peck, L.S.; Peters, M.; Postlethwait, J.H.; Clark, M.S. Adaptation of Proteins to the Cold in Antarctic Fish: A Role for Methionine? Genome Biol. EVolume 2019, 11, 220–231. [Google Scholar] [CrossRef]
  30. Liu, Y.; Jia, K.; Chen, H.; Wang, Z.; Zhao, W.; Zhu, L. Cold-Adapted Enzymes: Mechanisms, Engineering and Biotechnological Application. Bioprocess Biosyst. Eng. 2023, 46, 1399–1410. [Google Scholar] [CrossRef]
  31. Outzen, H.; Berglund, G.I.; Smalås, A.O.; Willassen, N.P. Temperature and PH Sensitivity of Trypsins from Atlantic Salmon (Salmo salar) in Comparison with Bovine and Porcine Trypsin. Comp. Biochem. Physiol. B Biochem. Mol. Biol. 1996, 115, 33–45. [Google Scholar] [CrossRef]
  32. Ásgeirsson, B.; Bjarnason, J.B. Properties of Elastase from Atlantic Cod a Cold-Adapted Proteinase. Biochim. Biophys. Acta BBA Protein Struct. Mol. 1993, 1164, 91–100. [Google Scholar] [CrossRef]
  33. Smalås, A.O.; Leiros, H.-K.S.; Os, V.; Willassen, N.P. Cold Adapted Enzymes. Biotechnol. Annu. Rev. 2000, 6, 1–57. [Google Scholar] [CrossRef]
  34. Lee, C.W.; Jang, S.H.; Chung, H.S. Improving the Stability of Cold-Adapted Enzymes by Immobilization. Catalysts 2017, 7, 112. [Google Scholar] [CrossRef]
  35. Philipps-Wiemann, P. Chapter 12—Proteases—General Aspects. In Enzymes in Human and Animal Nutrition; Academic Press: Cambridge, MA, USA, 2018. [Google Scholar]
  36. López-Otín, C.; Bond, J.S. Proteases: Multifunctional Enzymes in Life and Disease. J. Biol. Chem. 2008, 283, 30433–30437. [Google Scholar] [CrossRef] [PubMed]
  37. Rios-Herrera, G.D.; Osuna Ruiz, I.; Hernández, C.; Valdez-Ortiz, A.; Sandoval-Gallardo, J.M.; Martínez-Montaño, E.; Ramírez-Pérez, J.S.; Salazar-Leyva, J.A. Chihuil Sea Catfish Bagre Panamensis Viscera as a New Source of Serine Proteases: Semi-Purification, Biochemical Characterization and Application for Protein Hydrolysates Production. Waste Biomass Valorization 2020, 11, 5821–5833. [Google Scholar] [CrossRef]
  38. Kibenge, F.S.B.; Strange, R.J. Introduction to the Anatomy and Physiology of the Major Aquatic Animal Species in Aquaculture. In Aquaculture Pharmacology; Academic Press: Cambridge, MA, USA, 2021. [Google Scholar]
  39. Alarcón, F.J.; García-Carreño, F.L.; Navarrete Del Toro, M.A. Effect of Plant Protease Inhibitors on Digestive Proteases in Two Fish Species, Lutjanus Argentiventris and L. Novemfasciatus. Fish. Physiol. Biochem. 2001, 24, 179–189. [Google Scholar] [CrossRef]
  40. Haard, N.F.; Simpson, B.K. Digestive Proteinases from Marine Animals. In Seafood Enzymes; Marcel Dekker: New York, NY, USA, 2020. [Google Scholar]
  41. Fu, X.Y.; Xue, C.H.; Miao, B.C.; Li, Z.J.; Gao, X.; Yang, W.G. Characterization of Proteases from the Digestive Tract of Sea Cucumber (Stichopus Japonicus): High Alkaline Protease Activity. Aquaculture 2005, 246, 321–329. [Google Scholar] [CrossRef]
  42. Taghizadeh Andevari, G.; Rezaei, M.; Tabarsa, M.; Rustad, T. Extraction, Partial Purification and Characterization of Alkaline Protease from Rainbow Trout (Oncorhynchus mykiss) Viscera. Aquaculture 2019, 500, 458–463. [Google Scholar] [CrossRef]
  43. Michail, M.; Vasiliadou, M.; Zotos, A. Partial Purification and Comparison of Precipitation Techniques of Proteolytic Enzymes from Trout (Salmo gairdnerii) Heads. Food Chem. 2006, 97, 50–55. [Google Scholar] [CrossRef]
  44. Karlsen, S.; Hough, E.; Olsen, R.L. Structure and Proposed Amino-Acid Sequence of a Pepsin from Atlantic Cod (Gadus morhua). Acta Crystallogr. D Biol. Crystallogr. 1998, 54, 32–46. [Google Scholar] [CrossRef]
  45. Leiros, H.-K.S.; Ân, S.; Mcsweeney, M.; Smala Ês, A.O. Biological Crystallography Atomic Resolution Structures of Trypsin Provide Insight into Structural Radiation Damage. Acta Crystallogr. Sect. D Biol. Crystallogr. 2001, 57, 488–497. [Google Scholar] [CrossRef]
  46. Tsukada, H.; Blow, D.M. Structure of α-Chymotrypsin Refined at 1.68 Å Resolution. J. Mol. Biol. 1985, 184, 703–711. [Google Scholar] [CrossRef]
  47. Berglund, G.I.; Willassen, N.P.; Hordvik, A.; Smalås, A.O. Structure of Native Pancreatic Elastase from North. Atlantic Salmon at 1.61 Å Resolution. Acta Crystallogr. Sect. D Struct. Biol. 1995, 51, 925–937. [Google Scholar] [CrossRef] [PubMed]
  48. Rawlings, N.D.; Salvesen, G. Handbook of Proteolytic Enzymes; Academic Press: Cambridge, MA, USA, 2013; Volume 1–3. [Google Scholar]
  49. Perera, E.; Pons, T.; Hernandez, D.; Moyano, F.J.; Martínez-Rodríguez, G.; Mancera, J.M. New Members of the Brachyurins Family in Lobster Include a Trypsin-like Enzyme with Amino Acid Substitutions in the Substrate-Binding Pocket. FEBS J. 2010, 277, 3489–3501. [Google Scholar] [CrossRef] [PubMed]
  50. Estiasih, T.; Ahmadi, K.; Ali, D.Y.; Nisa, F.C.; Suseno, S.H.; Lestari, L.A. Valorisation of Viscera from Fish Processing for Food Industry Utilizations. IOP Conf. Ser. Earth Environ. Sci. 2021, 924, 012024. [Google Scholar] [CrossRef]
  51. Ponomareva, T.; Timchenko, M.; Filippov, M.; Lapaev, S.; Sogorin, E. Prospects of Red King Crab Hepatopancreas Processing: Fundamental and Applied Biochemistry. Recycling 2021, 6, 3. [Google Scholar] [CrossRef]
  52. Bougatef, A.; Balti, R.; Zaied, S.B.; Souissi, N.; Nasri, M. Pepsinogen and Pepsin from the Stomach of Smooth Hound (Mustelus mustelus): Purification, Characterization and Amino Acid Terminal Sequences. Food Chem. 2008, 107, 777–784. [Google Scholar] [CrossRef]
  53. Wu, T.; Sun, L.C.; Du, C.H.; Cai, Q.F.; Zhang, Q.B.; Su, W.J.; Cao, M.J. Identification of Pepsinogens and Pepsins from the Stomach of European Eel (Anguilla anguilla). Food Chem. 2009, 115, 137–142. [Google Scholar] [CrossRef]
  54. Gildberg, A. Digestive Enzyme Activities in Starved Pre-Slaughter Farmed and Wild-Captured, Atlantic Cod (Gadus morhua). Aquaculture 2004, 238, 343–353. [Google Scholar] [CrossRef]
  55. Zhou, Q.; Fu, X.P.; Zhang, L.J.; Su, W.J.; Cao, M.J. Purification and Characterization of Sea Bream (Sparus latus Houttuyn) Pepsinogens and Pepsins. Food Chem. 2007, 103, 795–801. [Google Scholar] [CrossRef]
  56. Nalinanon, S.; Benjakul, S.; Kishimura, H. Biochemical Properties of Pepsinogen and Pepsin from the Stomach of Albacore Tuna (Thunnus alalunga). Food Chem. 2010, 121, 49–55. [Google Scholar] [CrossRef]
  57. Nalinanon, S.; Benjakul, S.; Visessanguan, W.; Kishimura, H. Tuna Pepsin: Characteristics and Its Use for Collagen Extraction from the Skin of Threadfin Bream (Nemipterus Spp.). J. Food Sci. 2008, 73, C413–C419. [Google Scholar] [CrossRef]
  58. Zhou, Q.; Liu, G.M.; Huang, Y.Y.; Weng, L.; Hara, K.; Su, W.J.; Cao, M.J. Pepsinogens and Pepsins from Mandarin Fish (Siniperca chuatsi). J. Agric. Food Chem. 2008, 56, 5401–5406. [Google Scholar] [CrossRef] [PubMed]
  59. Klomklao, S.; Kishimura, H.; Yabe, M.; Benjakul, S. Purification and Characterization of Two Pepsins from the Stomach of Pectoral Rattail (Coryphaenoides pectoralis). Comp. Biochem. Physiol. B Biochem. Mol. Biol. 2007, 147. [Google Scholar] [CrossRef] [PubMed]
  60. Kishimura, H.; Klomklao, S.; Nalinanon, S.; Benjakul, S.; Chun, B.S.; Adachi, K. Comparative Study on Thermal Stability of Trypsin from the Pyloric Ceca of Threadfin Hakeling (Laemonema longipes). J. Food Biochem. 2010, 34, 50–65. [Google Scholar] [CrossRef]
  61. Zamani, A.; Benjakul, S. Trypsin from Unicorn Leatherjacket (Aluterus monoceros) Pyloric Caeca: Purification and Its Use for Preparation of Fish Protein Hydrolysate with Antioxidative Activity. J. Sci. Food Agric. 2016, 96, 962–969. [Google Scholar] [CrossRef]
  62. Klomklao, S.; Kishimura, H.; Benjakul, S. Trypsin from the Pyloric Ceca of Pectoral Rattail (Coryphaenoides pectoralis): Purification and Characterization. J. Agric. Food Chem. 2009, 57, 7097–7103. [Google Scholar] [CrossRef]
  63. Ktari, N.; Ben Khaled, H.; Nasri, R.; Jellouli, K.; Ghorbel, S.; Nasri, M. Trypsin from Zebra Blenny (Salaria basilisca) Viscera: Purification, Characterisation and Potential Application as a Detergent Additive. Food Chem. 2012, 130, 467–474. [Google Scholar] [CrossRef]
  64. Cai, Q.F.; Jiang, Y.K.; Zhou, L.G.; Sun, L.C.; Liu, G.M.; Osatomi, K.; Cao, M.J. Biochemical Characterization of Trypsins from the Hepatopancreas of Japanese Sea Bass (Lateolabrax japonicus). Comp. Biochem. Physiol. B Biochem. Mol. Biol. 2011, 159, 183–189. [Google Scholar] [CrossRef]
  65. Klomklao, S.; Benjakul, S.; Visessanguan, W.; Kishimura, H.; Simpson, B.K. Purification and Characterization of Trypsin from the Spleen of Tongol Tuna (Thunnus tonggol). J. Agric. Food Chem. 2006, 54, 5617–5622. [Google Scholar] [CrossRef]
  66. Zamani, A.; Rezaei, M.; Madani, R.; Habibi Rezaie, M. Trypsin Enzyme from Viscera of Common Kilka (Clupeonella cultriventris Caspia): Purification, Characterization, and Its Compatibility with Oxidants and Surfactants. J. Aquat. Food Prod. Technol. 2014, 23, 237–252. [Google Scholar] [CrossRef]
  67. Klomklao, S.; Benjakul, S.; Visessanguan, W.; Kishimura, H.; Simpson, B.K. Purification and Characterisation of Trypsins from the Spleen of Skipjack Tuna (Katsuwonus pelamis). Food Chem. 2007, 100, 1580–1589. [Google Scholar] [CrossRef]
  68. Kishimura, H.; Tokuda, Y.; Klomklao, S.; Benjakul, S.; Ando, S. Enzymatic Characteristics of Trypsin from Pyloric CECA of Spotted Mackerel (Scomber australasicus). J. Food Biochem. 2006, 30, 466–477. [Google Scholar] [CrossRef]
  69. Kishimura, H.; Klomklao, S.; Benjakul, S.; Chun, B.S. Characteristics of Trypsin from the Pyloric Ceca of Walleye Pollock (Theragra chalcogramma). Food Chem. 2008, 106, 194–199. [Google Scholar] [CrossRef]
  70. Klomklao, S.; Kishimura, H.; Benjakul, S.; Simpson, B.K.; Visessanguan, W. Cationic Trypsin: A Predominant Proteinase In Pacific Saury (Cololabis saira) Pyloric Ceca. J. Food Biochem. 2010, 34, 1105–1123. [Google Scholar] [CrossRef]
  71. Balti, R.; Barkia, A.; Bougatef, A.; Ktari, N.; Nasri, M. A Heat-Stable Trypsin from the Hepatopancreas of the Cuttlefish (Sepia officinalis): Purification and Characterisation. Food Chem. 2009, 113, 146–154. [Google Scholar] [CrossRef]
  72. Yang, F.; Su, W.J.; Lu, B.J.; Wu, T.; Sun, L.C.; Hara, K.; Cao, M.J. Purification and Characterization of Chymotrypsins from the Hepatopancreas of Crucian Carp (Carassius auratus). Food Chem. 2009, 116, 860–866. [Google Scholar] [CrossRef]
  73. Castillo-Yáñez, F.J.; Pacheco-Aguilar, R.; García-Carreño, F.L.; de los Ángeles Navarrete-Del Toro, M.; López, M.F. Purification and Biochemical Characterization of Chymotrypsin from the Viscera of Monterey Sardine (Sardinops sagax Caeruleus). Food Chem. 2006, 99, 252–259. [Google Scholar] [CrossRef]
  74. Castillo-Yañez, F.J.; Pacheco-Aguilar, R.; Lugo-Sanchez, M.E.; Garcia-Sanchez, G.; Quintero-Reyes, I.E. Biochemical Characterization of an Isoform of Chymotrypsin from the Viscera of Monterey Sardine (Sardinops sagax Caerulea), and Comparison with Bovine Chymotrypsin. Food Chem. 2009, 112, 634–639. [Google Scholar] [CrossRef]
  75. Daboor, S.M.; Budge, S.M.; Ghaly, A.E.; Brooks, M.S.; Dave, D. Isolation and Activation of Collagenase from Fish Processing Waste. Adv. Biosci. Biotechnol. 2012, 3, 191–203. [Google Scholar] [CrossRef]
  76. Felberg, H.S.; Hagen, L.; Slupphaug, G.; Batista, I.; Nunes, M.L.; Olsen, R.L.; Martinez, I. Partial Characterisation of Gelatinolytic Activities in Herring (Clupea harengus) and Sardine (Sardina pilchardus) Possibly Involved in Post-Mortem Autolysis of Ventral Muscle. Food Chem. 2010, 119, 675–683. [Google Scholar] [CrossRef]
  77. Oliveira, V.d.M.; Assis, C.R.D.; Silva, J.C.; Silva, Q.J.; Bezerra, R.d.S.; Porto, A.L.F. Recovery of Fibrinolytic and Collagenolytic Enzymes from Fish and Shrimp Byproducts: Potential Source for Biomedical Applications. Bol. Inst. Pesca 2019, 45, 4459–4466. [Google Scholar] [CrossRef]
  78. Aoki, H.; Ahsan, M.N.; Watabe, S. Molecular and Enzymatic Properties of a Cathepsin L-like Proteinase with Distinct Substrate Specificity from Northern Shrimp (Pandalus borealis). J. Comp. Physiol. B 2004, 174, 59–69. [Google Scholar] [CrossRef] [PubMed]
  79. Aoki, H.; Ahsan, M.N.; Watabe, S. Molecular Cloning and Characterization of Cathepsin B from the Hepatopancreas of Northern Shrimp Pandalus Borealis. Comp. Biochem. Physiol. B Biochem. Mol. Biol. 2003, 134, 681–694. [Google Scholar] [CrossRef] [PubMed]
  80. Aoki, H.; Ahsan, N.; Matsuo, K.; Hagiwara, T.; Watabe, S. Purification and Characterization of Collagenolytic Proteases from the Hepatopancreas of Northern Shrimp (Pandalus eous). J. Agric. Food Chem. 2003, 51, 777–783. [Google Scholar] [CrossRef] [PubMed]
  81. Sainz, J.C.; García-Carreño, F.L.; Hernández-Cortés, P. Penaeus Vannamei Isotrypsins: Purification and Characterization. Comp. Biochem. Physiol. B Biochem. Mol. Biol. 2004, 138, 155–162. [Google Scholar] [CrossRef] [PubMed]
  82. Hu, K.J.; Leung, P.C. Shrimp Cathepsin L Encoded by an Intronless Gene Has Predominant Expression in Hepatopancreas, and Occurs in the Nucleus of Oocyte. Comp. Biochem. Physiol. B Biochem. Mol. Biol. 2004, 137, 21–33. [Google Scholar] [CrossRef]
  83. Laycock, M.V.; Hirama, T.; Hasnain, S.; Watson, D.; Storer, A.C. Purification and Characterization of a Digestive Cysteine Proteinase from the American Lobster (Homarus americanus). Biochem. J. 1989, 263, 439–444. [Google Scholar] [CrossRef]
  84. Celis-Guerrero, L.E.; García-Carreño, F.L.; Navarrete Del Toro, M.A. Characterization of Proteases in the Digestive System of Spiny Lobster (Panulirus interruptus). Mar. Biotechnol. 2004, 6, 262–269. [Google Scholar] [CrossRef]
  85. Wang, P.A.; Stenvik, J.; Larsen, R.; Mæhre, H.; Olsen, R.L. Cathepsin D from Atlantic Cod (Gadus morhua L.) Liver. Isolation and Comparative Studies. Comp. Biochem. Physiol. B Biochem. Mol. Biol. 2007, 147, 504–511. [Google Scholar] [CrossRef]
  86. Zhou, D.Y.; Chang, X.N.; Bao, S.S.; Song, L.; Zhu, B.W.; Dong, X.P.; Zong, Y.; Li, D.M.; Zhang, M.M.; Liu, Y.X.; et al. Purification and Partial Characterisation of a Cathepsin L-like Proteinase from Sea Cucumber (Stichopus japonicus) and Its Tissue Distribution in Body Wall. Food Chem. 2014, 158, 192–199. [Google Scholar] [CrossRef]
  87. Zhu, B.W.; Zhao, L.L.; Sun, L.M.; Li, D.M.; Murata, Y.; Yu, L.; Zhang, L. Purification and Characterization of a Cathepsin L-like Enzyme from the Body Wall of the Sea Cucumber Stichopus Japonicus. Biosci. Biotechnol. Biochem. 2008, 72, 1430–1437. [Google Scholar] [CrossRef]
  88. Whang, I.; De Zoysa, M.; Nikapitiya, C.; Lee, Y.; Kim, Y.; Lee, S.; Oh, C.; Jung, S.J.; Oh, M.J.; Choi, C.Y.; et al. Molecular Characterization and Expression Analysis of Cathepsin B and L Cysteine Proteases from Rock Bream (Oplegnathus fasciatus). Fish. Shellfish. Immunol. 2011, 30, 763–772. [Google Scholar] [CrossRef] [PubMed]
  89. Szymczak, M. Recovery of Cathepsins from Marinating Brine Waste. Int. J. Food Sci. Technol. 2017, 52, 154–160. [Google Scholar] [CrossRef]
  90. Zhang, K.; Wei, R.; Song, R. Extraction of Cathepsin D-like Protease from Neon Flying Squid (Ommastrephes bartramii) Viscera and Application in Antioxidant Hydrolysate Production. Biomolecules 2019, 9, 228. [Google Scholar] [CrossRef] [PubMed]
  91. Singh, A. Hyperlipidemia in Cardiovascular Health and Digestion. In Nutrition and Functional Foods in Boosting Digestion, Metabolism and Immune Health; Academic Press: Cambridge, MA, USA, 2021. [Google Scholar]
  92. Patchimpet, J.; Sangkharak, K.; Klomklao, S. Lipolytic Activity of Viscera Extract from Three Freshwater Fish Species in Phatthalung, Thailand: Comparative Studies and Potential Use as Dishwashing Detergent Additive. Biocatal. Agric. Biotechnol. 2019, 19, 101143. [Google Scholar] [CrossRef]
  93. Mateos, P.S.; Navas, M.B.; Morcelle, S.R.; Ruscitti, C.; Matkovic, S.R.; Briand, L.E. Insights in the Biocatalyzed Hydrolysis, Esterification and Transesterification of Waste Cooking Oil with a Vegetable Lipase. Catal. Today 2021, 372, 211–219. [Google Scholar] [CrossRef]
  94. Iijima, N.; Tanaka, S.; Ota, Y. Purification and Characterization of Bile Salt-Activated Lipase from the Hepatopancreas of Red Sea Bream, Pagrus Major. Fish. Physiol. Biochem. 1998, 18, 59–69. [Google Scholar] [CrossRef]
  95. Nayak, J.; Nair, P.G.V.; Ammu, K.; Mathew, S. Lipase Activity in Different Tissues of Four Species of Fish: Rohu (Labeo rohita Hamilton), Oil Sardine (Sardinella iongiceps Linnaeus), Mullet (Liza subviridis Valenciennes) and Indian Mackerel (Rastrelliger kanagurta Cuvier). J. Sci. Food Agric. 2003, 83, 1139–1142. [Google Scholar] [CrossRef]
  96. Gjellesvik, D.R.; Lombardo, D.; Walther, B.T. Pancreatic Bile Salt Dependent Lipase from Cod (Gadus morhua): Purification and Properties. Biochim. Biophys. Acta BBA Lipids Lipid Metab. 1992, 1124, 123–134. [Google Scholar] [CrossRef]
  97. Gjellesvik, D.R.; Lorens, J.B.; Male, R. Pancreatic Carboxylester Lipase from Atlantic Salmon (Salmo salar): CDNA Sequence and Computer-Assisted Modelling of Tertiary Structure. Eur. J. Biochem. 1994, 226, 603–612. [Google Scholar] [CrossRef]
  98. Léger, C.; Ducruet, V.; Flanzy, J. Lipase et Colipase de La Truite Arc-En-Ciel. Quelques Résultats Récents. Ann. Biol. Anim. Biochim. Biophys. 1979, 19, 825–832. [Google Scholar] [CrossRef]
  99. Mukundan, M.K.; Gopakumar, K.; Nair, M.R. Purification of a Lipase from the Hepatopancreas of Oil Sardine (Sardinella longiceps Linnaeus) and Its Characteristics and Properties. J. Sci. Food Agric. 1985, 36, 191–203. [Google Scholar] [CrossRef]
  100. Taniguchi, A.; Takano, K.; Kamoi, I. Purification and Properties of Lipase from Tilapia Intestine—Digestive Enzyme of Tilapia-VI. Nippon Suisan Gakkaishi Jpn. Ed. 2001, 67, 78–84. [Google Scholar] [CrossRef]
  101. Noriega-Rodríguez, J.A.; Gámez-Meza, N.; Alanis-Villa, A.; Medina-Juárez, L.A.; Tejeda-Mansir, A.; Angulo-Guerrero, O.; García, H.S. Extraction and Fractionation of Lipolytic Enzyme from Viscera of Monterey Sardine (Sardinops sagax Caerulea). Int. J. Food Sci. Technol. 2009, 44, 1223–1228. [Google Scholar] [CrossRef]
  102. Mardina, V.; Harmawan, T.; Fitriani; Hildayani, G.M.; Yusof, F. Screening of Protease and Lipase Sources from Visceral Organs of Euthynnusaffinis. IOP Conf. Ser. Mater. Sci. Eng. 2018, 420, 012083. [Google Scholar] [CrossRef]
  103. Borlongan, I.G. Studies on the Digestive Lipases of Milkfish, Chanos Chanos. Aquaculture 1990, 89, 315–325. [Google Scholar] [CrossRef]
  104. Liu, H.; Liu, S.; Zhang, X.; Liu, Z.; Li, C.; Xia, G.; Shen, X. Novel Lipase from Golden Pompano (Trachinotus ovatus) Viscera: Purification, Characterization, and Application in the Concentrating of n-3 Polyunsaturated Fatty Acids. J. Ocean. Univ. China 2023, 22, 501–508. [Google Scholar] [CrossRef]
  105. Cherif, S.; Fendri, A.; Miled, N.; Trabelsi, H.; Mejdoub, H.; Gargouri, Y. Crab Digestive Lipase Acting at High Temperature: Purification and Biochemical Characterization. Biochimie 2007, 89, 1012–1018. [Google Scholar] [CrossRef]
  106. Cherif, S.; Gargouri, Y. Thermoactivity and Effects of Organic Solvents on Digestive Lipase from Hepatopancreas of the Green Crab. Food Chem. 2009, 116, 82–86. [Google Scholar] [CrossRef]
  107. Rivera-Pérez, C.; del Toro, M.d.l.Á.N.; García-Carreño, F. Purification and Characterization of an Intracellular Lipase from Pleopods of Whiteleg Shrimp (Litopenaeus Vannamei). Comp. Biochem. Physiol. B Biochem. Mol. Biol. 2011, 158, 99–105. [Google Scholar] [CrossRef]
  108. Rivera-Pérez, C.; García-Carreño, F.L.; Saborowski, R. Purification and Biochemical Characterization of Digestive Lipase in Whiteleg Shrimp. Mar. Biotechnol. 2011, 13, 284–295. [Google Scholar] [CrossRef]
  109. Zarai, Z.; Ali, M.B.; Fendri, A.; Louati, H.; Mejdoub, H.; Gargouri, Y. Purification and Biochemical Properties of Hexaplex Trunculus Digestive Lipase. Process. Biochem. 2012, 47, 2434–2439. [Google Scholar] [CrossRef]
  110. Uchiyama, S.; Iijima, N. Partial Purification and Characterization of Pro-Phospholipase A 2 Activating Proteases from Gill Membranes of the Red Sea Bream, Chrysophrys Major. Comp. Biochem. Physiol. B Biochem. Mol. Biol. 2005, 141, 121–127. [Google Scholar] [CrossRef] [PubMed]
  111. Aaen, B.; Jessen, F.; Jensen, B. Partial Purification and Characterization of a Cellular Acidic Phospholipase A2 from Cod (Gadus morhua) Muscle. Comp. Biochem. Physiol. Part. B Biochem. 1995, 110, 547–554. [Google Scholar] [CrossRef]
  112. Audley, M.A.; Shetty, K.J.; Kinsella, J.E. Isolation and Properties of Phospholipase A from Pollock Muscle. J. Food Sci. 1978, 43, 1771–1775. [Google Scholar] [CrossRef]
  113. Chawla, P.; Ablett, R.F. Detection of Microsomal Phospholipase Activity in Myotomal Tissue of Atlantic Cod (Gadus morhua). J. Food Sci. 1987, 52, 1194–1197. [Google Scholar] [CrossRef]
  114. Kishimura, H.; Hayashi, K. Purification and Properties of Phospholipase A2 Isozymes from Pyloric Ceca of the Starfish (Asterina pectinifera). J. Food Biochem. 2004, 28, 181–194. [Google Scholar] [CrossRef]
  115. Iijima, N.; Chosa, S.; Uematsu, K.; Goto, T.; Hoshita, T.; Kayama, M. Purification and Characterization of Phospholipase A2 from the Pyloric Caeca of Red Sea Bream, Pagrus Major. Fish. Physiol. Biochem. 1997, 16, 487–498. [Google Scholar] [CrossRef]
  116. Uchiyama, S.; Fujikawa, Y.; Uematsu, K.; Matsuda, H.; Aida, S.; Iijima, N. Localization of Group IB Phospholipase A2 Isoform in the Gills of the Red Sea Bream, Pagrus (Chrysophrys) Major. Comp. Biochem. Physiol. B Biochem. Mol. Biol. 2002, 132, 671–683. [Google Scholar] [CrossRef]
  117. Ono, H.; Iijima, N. Purification and Characterization of Phospholipase A2 Isoforms from the Hepatopancreas of Red Sea Bream, Pagrus Major. Fish. Physiol. Biochem. 1998, 18, 135–147. [Google Scholar] [CrossRef]
  118. Cherif, S.; Bacha, A.B.; Ali, Y.B.; Horchani, H.; Rekik, W.; Gargouri, Y. Crab Digestive Phospholipase: A New Invertebrate Member. Bioresour. Technol. 2010, 101, 366–371. [Google Scholar] [CrossRef]
  119. Phillips, A.M. Nutrition, Digestion, and Energy Utilization. Fish. Physiol. 1969, 1, 366–371. [Google Scholar] [CrossRef]
  120. Chiu, Y.N.; Benitez, L.V. Studies on the Carbohydrases in the Digestive Tract of the Milkfish Chanos Chanos. Mar. Biol. 1981, 61, 247–254. [Google Scholar] [CrossRef]
  121. Kawai, S.i.; Ikeda, S. Studies on Digestive Enzymes of Fishes—I. Carbohydrases in Digestive Organs of Several Fishes. Nippon Suisan Gakkaishi 1971, 37, 333–337. [Google Scholar] [CrossRef]
  122. Mehrani, H.; Storey, K.B. Characterization of α-Glucosidases from Rainbow Trout Liver. Arch. Biochem. Biophys. 1993, 306, 188–194. [Google Scholar] [CrossRef]
  123. Munilla-Morán, R.; Saborido-Rey, F. Digestive Enzymes in Marine Species. II. Amylase Activities in Gut from Seabream (Sparus aurata), Turbot (Scophthalmus maximus) and Redfish (Sebastes mentella). Comp. Biochem. Physiol. B Biochem. Mol. Biol. 1996, 113, 827–834. [Google Scholar] [CrossRef]
  124. Fish, G.R. The Comparative Activity of Some Digestive Enzymes in the Alimentary Canal of Tilapia and Perch. Hydrobiologia 1960, 15, 161–178. [Google Scholar] [CrossRef]
  125. Hidalgo, M.C.; Urea, E.; Sanz, A. Comparative Study of Digestive Enzymes in Fish with Different Nutritional Habits. Proteolytic and Amylase Activities. Aquaculture 1999, 170, 267–283. [Google Scholar] [CrossRef]
  126. Spannhof, L.; Plantikow, H. Studies on Carbohydrate Digestion in Rainbow Trout. Aquaculture 1983, 30, 95–108. [Google Scholar] [CrossRef]
  127. Skea, G.L.; Mountfort, D.O.; Clements, K.D. Gut Carbohydrases from the New Zealand Marine Herbivorous Fishes Kyphosus Sydneyanus (Kyphosidae), Aplodactylus Arctidens (Aplodactylidae) and Odax Pullus (Labridae). Comp. Biochem. Physiol. B Biochem. Mol. Biol. 2005, 140, 259–269. [Google Scholar] [CrossRef]
  128. Maalej, H.; Maalej, A.; Affes, S.; Hmidet, N.; Nasri, M. A Novel Digestive α-Amylase from Blue Crab (Portunus segnis) Viscera: Purification, Biochemical Characterization and Application for the Improvement of Antioxidant Potential of Oat Flour. Int. J. Mol. Sci. 2021, 22, 1070. [Google Scholar] [CrossRef]
  129. Castro, P.F.; Freitas, A.C.V.; Santana, W.M.; Costa, H.M.S.; Carvalho, L.B.; Bezerra, R.S. Comparative Study of Amylases from the Midgut Gland of Three Species of Penaeid Shrimp. J. Crustac. Biol. 2012, 32, 607–613. [Google Scholar] [CrossRef]
  130. Charoenpol, A.; Crespy, D.; Schulte, A.; Suginta, W. Marine Chitin Upcycling with Immobilized Chitinolytic Enzymes: Current State and Prospects. Green Chem. 2022, 25, 467–489. [Google Scholar] [CrossRef]
  131. Yang, S.; Song, S.; Yan, Q.; Fu, X.; Jiang, Z.; Yang, X. Biochemical Characterization of the First Fungal Glycoside Hydrolyase Family 3 β-N-Acetylglucosaminidase from Rhizomucor Miehei. J. Agric. Food Chem. 2014, 62, 5181–5190. [Google Scholar] [CrossRef] [PubMed]
  132. Matsumiya, M.; Mochizuki, A. Distribution of Chitinase and β-N-Acetylhexosaminidase in the Organs of Several Fishes. Fish. Sci. 1996, 62, 150–151. [Google Scholar] [CrossRef]
  133. Ikeda, M.; Kakizaki, H.; Matsumiya, M. Biochemistry of Fish Stomach Chitinase. Int. J. Biol. Macromol. 2017, 104, 1672–1681. [Google Scholar] [CrossRef]
  134. Gutowska, M.A.; Drazen, J.C.; Robison, B.H. Digestive Chitinolytic Activity in Marine Fishes of Monterey Bay, California. Comp. Biochem. Physiol. A Mol. Integr. Physiol. 2004, 139, 351–358. [Google Scholar] [CrossRef]
  135. Shinya, S.; Fukamizo, T. Interaction between Chitosan and Its Related Enzymes: A Review. Int. J. Biol. Macromol. 2017, 104, 1422–1435. [Google Scholar] [CrossRef]
  136. Kim, S. Competitive Biological Activities of Chitosan and Its Derivatives: Antimicrobial, Antioxidant, Anticancer, and Anti-Inflammatory Activities. Int. J. Polym. Sci. 2018, 2018, 1708172. [Google Scholar] [CrossRef]
  137. Baehaki, A.; Lestari, S.D.; Wahidman, Y.; Gofar, N. Characteristics of Chitinase Isolated from Different Part of Snakehead Fish (Channa striata) Digestive Tract. IOP Conf. Series: Earth Environ. Sci. 2018, 102, 012057. [Google Scholar] [CrossRef]
  138. Affes, S.; Aranaz, I.; Hamdi, M.; Acosta, N.; Ghorbel-Bellaaj, O.; Heras, Á.; Nasri, M.; Maalej, H. Preparation of a Crude Chitosanase from Blue Crab Viscera as Well as Its Application in the Production of Biologically Active Chito-Oligosaccharides from Shrimp Shells Chitosan. Int. J. Biol. Macromol. 2019, 139, 558–569. [Google Scholar] [CrossRef]
  139. Yao, D.R.; Zhou, M.Q.; Wu, S.J.; Pan, S.K. Depolymerization of Chitosan by Enzymes from the Digestive Tract of Sea Cucumber Stichopus Japonicus. Afr. J. Biotechnol. 2011, 11, 423–428. [Google Scholar] [CrossRef]
  140. Kakizaki, H.; Ikeda, M.; Fukushima, H.; Matsumiya, M. Distribution of Chitinolytic Enzymes in the Organs and CDNA Cloning of Chitinase Isozymes from the Stomach of Two Species of Fish, Chub Mackerel (Scomber japonicus) and Silver Croaker (Pennahia argentata). Open J. Mar. Sci. 2015, 5, 398–411. [Google Scholar] [CrossRef]
  141. Ikeda, M.; Miyauchi, K.; Mochizuki, A.; Matsumiya, M. Purification and Characterization of Chitinase from the Stomach of Silver Croaker Pennahia Argentatus. Protein Expr. Purif. 2009, 65, 214–222. [Google Scholar] [CrossRef] [PubMed]
  142. Matsumiya, M.; Arakane, Y.; Haga, A.; Muthukrishnan, S.; Kramer, K.J. Substrate Specificity of Chitinases from Two Species of Fish, Greenling, Hexagrammos Otakii, and Common Mackerel, Scomber japonicus, and the Insect, Tobacco Hornworm, Manduca Sexta. Biosci. Biotechnol. Biochem. 2006, 70, 971–979. [Google Scholar] [CrossRef]
  143. Kono, M.; Wilder, M.N.; Matsui, T.; Furukawa, K.; Aida, K.; Koga, D. Chitinolytic Enzyme Activities in the Hepatopancreas, Tail Fan and Hemolymph of Kuruma Prawn Penaeus Japonicus during the Molt Cycle. Fish. Sci. 1995, 61, 727–728. [Google Scholar] [CrossRef]
  144. Mengkrog Holen, M.; Tuveng, T.R.; Kent, M.P.; Vaaje-Kolstad, G. The Gastric Mucosa of Atlantic Salmon (Salmo salar) Is Abundant in Highly Active Chitinases. FEBS Open Bio 2024, 14, 23–36. [Google Scholar] [CrossRef]
  145. Ikeda, M.; Kondo, Y.; Matsumiya, M. Purification, Characterization, and Molecular Cloning of Chitinases from the Stomach of the Threeline Grunt Parapristipoma Trilineatum. Process. Biochem. 2013, 48, 1324–1334. [Google Scholar] [CrossRef]
  146. Ikeda, M.; Shirase, D.; Sato, T.; Ueda, M.; Hirabayashi, S.; Matsumiya, M. Primary Structure and Enzymatic Properties of Chitinase Isozymes Purified from the Stomach of the Marbled Rockfish Sebastiscus Marmoratus. J. Chitin Chitosan Sci. 2014, 2, 106–116. [Google Scholar] [CrossRef]
  147. Watanabe, M.; Kakizaki, H.; Tsukamoto, T.; Fujiwara, M.; Fukushima, H.; Ueda, M.; Matsumiya, M. Distribution of Chitinolytic Enzyme in the Organs and Molecular Cloning of a Novel Chitinase Gene from the Kidney of Marbled Rockfish Sebastiscus Marmoratus. Adv. Biosci. Biotechnol. 2018, 9, 36–51. [Google Scholar] [CrossRef]
  148. Kawashima, S.; Ikehata, H.; Tada, C.; Ogino, T.; Kakizaki, H.; Ikeda, M.; Fukushima, H.; Matsumiya, M. Stomach Chitinase from Japanese Sardine Sardinops Melanostictus: Purification, Characterization, and Molecular Cloning of Chitinase Isozymes with a Long Linker. Mar. Drugs 2016, 14, 22. [Google Scholar] [CrossRef]
  149. Karasuda, S.; Yamamoto, K.; Kono, M.; Sakuda, S.; Koga, D. Kinetic Analysis of a Chitinase from Red Sea Bream, Pagrus Major. Biosci. Biotechnol. Biochem. 2004, 68, 1338–1344. [Google Scholar] [CrossRef] [PubMed]
  150. Kono, M.; Matsui, T.; Shimizu, C.; Koga, D. Purification and Some Properties of Chitinase from the Stomach of Japanese Eel, Anguilla Japonica. Agric. Biol. Chem. 1990, 54, 973–978. [Google Scholar] [CrossRef]
  151. Chen, Y.; Tao, Z.; Zhang, M.; Sun, L.; Liu, G.; Cao, M. Identification of a Chitinase from the Hepatopancreas of Chinese Black Sleeper (Bostrychus sinensis). Acta Oceanol. Sin. 2021, 40, 50–60. [Google Scholar] [CrossRef]
  152. Laribi-Habchi, H.; Dziril, M.; Badis, A.; Mouhoub, S.; Mameri, N. Purification and Characterization of a Highly Thermostable Chitinase from the Stomach of the Red Scorpionfish Scorpaena Scrofa with Bioinsecticidal Activity toward Cowpea Weevil Callosobruchus Maculatus (Coleoptera: Bruchidae). Biosci. Biotechnol. Biochem. 2012, 76, 1733–1740. [Google Scholar] [CrossRef]
  153. Molinari, L.M.; Pedroso, R.B.; Scoaris, D.d.O.; Ueda-Nakamura, T.; Nakamura, C.V.; Dias Filho, B.P. Identification and Partial Characterisation of a Chitinase from Nile Tilapia, Oreochromis niloticus. Comp. Biochem. Physiol. B Biochem. Mol. Biol. 2007, 146, 81–87. [Google Scholar] [CrossRef]
  154. Matsumiya, M.; Karasuda, S.; Miyauchi, K.; Mochizuki, A. Substrate Specificity and Partial Amino Acid Sequence of a Chitinase from the Stomach of Coelacanth Latimeria Chalumnae. Fish. Sci. 2008, 74, 1360–1362. [Google Scholar] [CrossRef]
  155. Novikov, V.Y.; Mukhin, V.A. Chitosan Depolymerization by Enzymes from the Hepatopancreas of the Crab Paralithodes Camtschaticus. Appl. Biochem. Microbiol. 2003, 39, 464–468. [Google Scholar] [CrossRef]
  156. Zou, E.; Bonvillain, R. Chitinase Activity in the Epidermis of the Fiddler Crab, Uca Pugilator, as an in Vivo Screen for Molt-Interfering Xenobiotics. Comp. Biochem. Physiol. C Toxicol. Pharmacol. 2004, 139, 225–230. [Google Scholar] [CrossRef]
  157. Nishino, R.; Kakizaki, H.; Fukushima, H.; Matsumiya, M. Distribution of Chitinolytic Enzymes in the Organs and CDNA Cloning of Chitinase Isozymes from the Liver of Golden Cuttlefish Sepia Esculenta. Adv. Biosci. Biotechnol. 2017, 8, 361–377. [Google Scholar] [CrossRef]
  158. Matsumiya, M.; Miyauchi, K.; Mochizuki, A. Distribution of Chitinase and β-N-Acetylhexosaminidase in the Organs of a Few Squid and a Cuttlefish. Fish. Sci. 1998, 64, 166–167. [Google Scholar] [CrossRef]
  159. Duarte, L.; Matte, C.R.; Bizarro, C.V.; Ayub, M.A.Z. Transglutaminases: Part I—Origins, Sources, and Biotechnological Characteristics. World J. Microbiol. Biotechnol. 2020, 36, 15. [Google Scholar] [CrossRef] [PubMed]
  160. Nozawa, H.; Mamegoshi, S.I.; Seki, N. Partial Purification and Characterization of Six Transglutaminases from Ordinary Muscles of Various Fishes and Marine Invertebrates. Comp. Biochem. Physiol. B Biochem. Mol. Biol. 1997, 118, 313–317. [Google Scholar] [CrossRef] [PubMed]
  161. Seki, N.; Uno, H.; Lee, N.H.; Arai, K.I.; Kimura, I.; Toyoda, K.; Fujita, T. Transglutaminase Activity in Alaska Pollack Muscle and Surimi, and Its Reaction with Myosin B. Nippon. Suisan Gakkaishi 1990, 56, 125–132. [Google Scholar] [CrossRef]
  162. Binsi, P.K.; Shamasundar, B.A. Purification and Characterisation of Transglutaminase from Four Fish Species: Effect of Added Transglutaminase on the Viscoelastic Behaviour of Fish Mince. Food Chem. 2012, 132, 1922–1929. [Google Scholar] [CrossRef]
  163. Worratao, A.; Yongsawatdigul, J. Purification and Characterization of Transglutaminase from Tropical Tilapia (Oreochromis niloticus). Food Chem. 2005, 93, 651–658. [Google Scholar] [CrossRef]
  164. Kumazawa, Y.; Sano, K.I.; Seguro, K.; Yasueda, H.; Nio, N.; Motoki, M. Purification and Characterization of Transglutaminase from Japanese Oyster (Crassostrea gigas). J. Agric. Food Chem. 1997, 45, 604–610. [Google Scholar] [CrossRef]
  165. Noguchi, K.; Ishikawa, K.; Yokoyama, K.I.; Ohtsuka, T.; Nio, N.; Suzuki, E.I. Crystal Structure of Red Sea Bream Transglutaminase. J. Biol. Chem. 2001, 276, 12055–12059. [Google Scholar] [CrossRef]
  166. Kumazawa, Y.; Yasueda, H.; Motoki, M. Purification and Characterization of a Tissue-Type Transglutaminase from Red Sea Bream (Pagrus major). Biosci. Biotechnol. Biochem. 1994, 58, 2041–2045. [Google Scholar] [CrossRef]
  167. Yasueda, H.; Nakanishi, K.; Kumazawa, Y.; Nagase, K.; Motoki, M.; Matsui, H. Tissue-Type Transglutaminase from Red Sea Bream (Pagrus major): Sequence Analysis of the CDNA and Functional Expression in Escherichia Coli. Eur. J. Biochem. 1995, 232, 411–419. [Google Scholar] [CrossRef]
  168. Kumazawa, Y.; Nakanishi, K.; Yasueda, H.; Motoki, M. Purification and Characterization of Transglutaminase from Walleye Pollack Liver. Fish. Sci. 1996, 62, 959–964. [Google Scholar] [CrossRef]
  169. Nozawa, H.; Cho, S.Y.; Seki, N. Purification and Characterization of Transglutaminase from Squid Gill. Fish. Sci. 2001, 67, 912–919. [Google Scholar] [CrossRef]
  170. Iijima, M.; Hashimoto, T.; Matsuda, Y.; Nagai, T.; Yamano, Y.; Ichi, T.; Osaki, T.; Kawabata, S.I. Comprehensive Sequence Analysis of Horseshoe Crab Cuticular Proteins and Their Involvement in Transglutaminase-Dependent Cross-Linking. FEBS J. 2005, 272, 4774–4786. [Google Scholar] [CrossRef] [PubMed]
  171. Tokunaga, F.; Iwanaga, S. Horseshoe Crab Transglutaminase. Methods Enzymol. 1993, 223, 378–388. [Google Scholar] [CrossRef] [PubMed]
  172. Ghorbel, S.; Souissi, N.; Triki-Ellouz, Y.; Dufossé, L.; Guérard, F.; Nasri, M. Preparation and Testing of Sardinella Protein Hydrolysates as Nitrogen Source for Extracellular Lipase Production by Rhizopus Oryzae. World J. Microbiol. Biotechnol. 2005, 21, 33–38. [Google Scholar] [CrossRef]
  173. Fahmy, N.M.; El-Deeb, B. Optimization, Partial Purification, and Characterization of a Novel High Molecular Weight Alkaline Protease Produced by Halobacillus Sp. HAL1 Using Fish Wastes as a Substrate. J. Genet. Eng. Biotechnol. 2023, 21, 48. [Google Scholar] [CrossRef]
  174. Ramkumar, A.; Sivakumar, N.; Victor, R. Fish Waste-Potential Low Cost Substrate for Bacterial Protease Production: A Brief Review. Open Biotechnol. J. 2016, 10, 335–341. [Google Scholar] [CrossRef]
  175. Ben Rebah, F.; Miled, N. Fish Processing Wastes for Microbial Enzyme Production: A Review. 3 Biotech. 2013, 3, 255–265. [Google Scholar] [CrossRef]
  176. Souissi, N.; Ellouz-Triki, Y.; Bougatef, A.; Blibech, M.; Nasri, M. Preparation and Use of Media for Protease-Producing Bacterial Strains Based on by-Products from Cuttlefish (Sepia officinalis) and Wastewaters from Marine-Products Processing Factories. Microbiol. Res. 2008, 163, 473–480. [Google Scholar] [CrossRef]
  177. Triki-Ellouz, Y.; Ghorbel, B.; Souissi, N.; Kammoun, S.; Nasri, M. Biosynthesis of Protease by Pseudomonas Aeruginosa MN7 Grown on Fish Substrate. World J. Microbiol. Biotechnol. 2003, 19, 41–45. [Google Scholar] [CrossRef]
  178. Ellouz, Y.; Bayoudh, A.; Kammoun, S.; Gharsallah, N.; Nasri, M. Production of Protease by Bacillus Subtilis Grown on Sardinelle Heads and Viscera Flour. Bioresour. Technol. 2001, 80, 49–51. [Google Scholar] [CrossRef]
  179. Vázquez, J.A.; Docasal, S.F.; Mirón, J.; González, M.P.; Murado, M.A. Proteases Production by Two Vibrio Species on Residuals Marine Media. J. Ind. Microbiol. Biotechnol. 2006, 33, 661–668. [Google Scholar] [CrossRef] [PubMed]
  180. Tran, T.N.; Doan, C.T.; Nguyen, V.B.; Nguyen, A.D.; Wang, S.L. Conversion of Fishery Waste to Proteases by Streptomyces Speibonae and Their Application in Antioxidant Preparation. Fishes 2022, 7, 140. [Google Scholar] [CrossRef]
  181. Doan, C.T.; Tran, T.N.; Nguyen, M.T.; Nguyen, H.K.; Tran, T.K.T.; Nguyen, T.H.; Tran, T.P.H.; Nguyen, V.B.; Nguyen, A.D.; Wang, S.L. Potential of the Liquid Fermentation of Fishery Waste by Paenibacillus Elgii for Metalloprotease Production. Polymers 2022, 14, 2741. [Google Scholar] [CrossRef] [PubMed]
  182. Wang, S.L.; Yeh, P.Y. Production of a Surfactant- and Solvent-Stable Alkaliphilic Protease by Bioconversion of Shrimp Shell Wastes Fermented by Bacillus Subtilis TKU007. Process. Biochem. 2006, 41, 1545–1552. [Google Scholar] [CrossRef]
  183. Wang, S.L.; Hsu, W.T.; Liang, T.W.; Yen, Y.H.; Wang, C.L. Purification and Characterization of Three Novel Keratinolytic Metalloproteases Produced by Chryseobacterium Indologenes TKU014 in a Shrimp Shell Powder Medium. Bioresour. Technol. 2008, 99, 5679–5686. [Google Scholar] [CrossRef]
  184. Wang, S.L.; Yeh, P.Y. Purification and Characterization of a Chitosanase from a Nattokinase Producing Strain Bacillus Subtilis TKU007. Process. Biochem. 2008, 43, 132–138. [Google Scholar] [CrossRef]
  185. Wang, S.L.; Chen, S.J.; Wang, C.L. Purification and Characterization of Chitinases and Chitosanases from a New Species Strain Pseudomonas Sp. TKU015 Using Shrimp Shells as a Substrate. Carbohydr. Res. 2008, 343, 1171–1179. [Google Scholar] [CrossRef]
  186. Wang, S.L.; Chao, C.H.; Liang, T.W.; Chen, C.C. Purification and Characterization of Protease and Chitinase from Bacillus Cereus TKU006 and Conversion of Marine Wastes by These Enzymes. Mar. Biotechnol. 2009, 11, 334–344. [Google Scholar] [CrossRef]
  187. Liang, T.W.; Kuo, Y.H.; Wu, P.C.; Wang, C.L.; Dzung, N.A.; Wang, S.L. Purification and Characterization of a Chitosanase and a Protease by Conversion of Shrimp Shell Wastes Fermented by Serratia Marcescens Subsp. Sakuensis TKU019. J. Chin. Chem. Soc. 2010, 57, 857–863. [Google Scholar] [CrossRef]
  188. Wang, S.L.; Peng, J.H.; Liang, T.W.; Liu, K.C. Purification and Characterization of a Chitosanase from Serratia Marcescens TKU011. Carbohydr. Res. 2008, 343, 1316–1323. [Google Scholar] [CrossRef]
  189. Barghini, P.; Moscatelli, D.; Garzillo, A.M.V.; Crognale, S.; Fenice, M. High Production of Cold-Tolerant Chitinases on Shrimp Wastes in Bench-Top Bioreactor by the Antarctic Fungus Lecanicillium Muscarium CCFEE 5003: Bioprocess Optimization and Characterization of Two Main Enzymes. Enzym. Microb Technol 2013, 53, 331–338. [Google Scholar] [CrossRef] [PubMed]
  190. Doan, C.T.; Tran, T.N.; Nguyen, V.B.; Nguyen, A.D.; Wang, S.L. Production of a Thermostable Chitosanase from Shrimp Heads via Paenibacillus Mucilaginosus TKU032 Conversion and Its Application in the Preparation of Bioactive Chitosan Oligosaccharides. Mar. Drugs 2019, 17, 217. [Google Scholar] [CrossRef] [PubMed]
  191. Liang, T.W.; Lin, J.J.; Yen, Y.H.; Wang, C.L.; Wang, S.L. Purification and Characterization of a Protease Extracellularly Produced by Monascus Purpureus CCRC31499 in a Shrimp and Crab Shell Powder Medium. Enzym. Microb. Technol. 2006, 38, 74–80. [Google Scholar] [CrossRef]
  192. Annamalai, N.; Rajeswari, M.V.; Sahu, S.K.; Balasubramanian, T. Purification and Characterization of Solvent Stable, Alkaline Protease from Bacillus Firmus CAS 7 by Microbial Conversion of Marine Wastes and Molecular Mechanism Underlying Solvent Stability. Process. Biochem. 2014, 49, 1012–1019. [Google Scholar] [CrossRef]
  193. Wang, S.L.; Shih, I.L.; Liang, T.W.; Wang, C.H. Purification and Characterization of Two Antifungal Chitinases Extracellularly Produced by Bacillus Amyloliquefaciens V656 in a Shrimp and Crab Shell Powder Medium. J. Agric. Food Chem. 2002, 50, 2241–2248. [Google Scholar] [CrossRef]
  194. Wang, S.L.; Hsiao, W.J.; Chang, W.T. Purification and Characterization of an Antimicrobial Chitinase Extracellularly Produced by Monascus Purpureus CCRC31499 in a Shrimp and Crab Shell Powder Medium. J. Agric. Food Chem. 2002, 50, 2249–2255. [Google Scholar] [CrossRef]
  195. Chang, W.T.; Chen, C.S.; Wang, S.L. An Antifungal Chitinase Produced by Bacillus Cereus with Shrimp and Crab Shell Powder as a Carbon Source. Curr. Microbiol. 2003, 47, 102–108. [Google Scholar] [CrossRef]
  196. Wang, S.L.; Wang, C.Y.; Huang, T.Y. Microbial Reclamation of Squid Pen for the Production of a Novel Extracellular Serine Protease by Lactobacillus Paracasei Subsp Paracasei TKU012. Bioresour. Technol. 2008, 99, 3411–3417. [Google Scholar] [CrossRef]
  197. Wang, S.L.; Lin, C.L.; Liang, T.W.; Liu, K.C.; Kuo, Y.H. Conversion of Squid Pen by Serratia Ureilytica for the Production of Enzymes and Antioxidants. Bioresour. Technol. 2009, 100, 316–323. [Google Scholar] [CrossRef]
  198. Suyotha, W.; Srisuk, K.; Cheirsilp, B. Production, Purification, and Characterization of Acid-Active Chitosanase from Lentzea Sp. OUR-I1 Using Squid Pen Waste as Low-Cost Nutrient Source with Simultaneous Recovery of Bioactive Compounds from Microbial Fermentation. Biocatal. Agric. Biotechnol. 2024, 56, 103001. [Google Scholar] [CrossRef]
  199. Doan, C.T.; Tran, T.N.; Nguyen, V.B.; Nguyen, A.D.; Wang, S.L. Conversion of Squid Pens to Chitosanases and Proteases via Paenibacillus Sp. TKU042. Mar. Drugs 2018, 16, 83. [Google Scholar] [CrossRef] [PubMed]
  200. Doan, C.T.; Tran, T.N.; Nguyen, V.B.; Tran, T.D.; Nguyen, A.D.; Wang, S.L. Bioprocessing of Squid Pens Waste into Chitosanase by Paenibacillus Sp. TKU047 and Its Application in Low-Molecular Weight Chitosan Oligosaccharides Production. Polymers 2020, 12, 1163. [Google Scholar] [CrossRef] [PubMed]
  201. Doan, C.T.; Tran, T.N.; Tran, T.P.H.; Nguyen, T.T.; Nguyen, H.K.; Tran, T.K.T.; Vu, B.T.; Trinh, T.H.T.; Nguyen, A.D.; Wang, S.L. Chitosanase Production from the Liquid Fermentation of Squid Pens Waste by Paenibacillus Elgii. Polymers 2023, 15, 3724. [Google Scholar] [CrossRef] [PubMed]
  202. Esakkiraj, P.; Austin Jeba Dhas, G.; Palavesam, A.; Immanuel, G. Media Preparation Using Tuna-Processing Wastes for Improved Lipase Production by Shrimp Gut Isolate Staphylococcus Epidermidis CMST Pi 2. Appl. Biochem. Biotechnol. 2010, 160, 1254–1265. [Google Scholar] [CrossRef]
  203. Esakkiraj, P.; Immanuel, G.; Sowmya, S.M.; Iyapparaj, P.; Palavesam, A. Evaluation of Protease-Producing Ability of Fish Gut Isolate Bacillus Cereus for Aqua Feed. Food Bioproc Tech. 2009, 2, 383–390. [Google Scholar] [CrossRef]
  204. Ben Rebah, F.; Frikha, F.; Kamoun, W.; Belbahri, L.; Gargouri, Y.; Miled, N. Culture of Staphylococcus Xylosus in Fish Processing By-Product-Based Media for Lipase Production. Lett. Appl. Microbiol. 2008, 47, 549–554. [Google Scholar] [CrossRef]
  205. Haddar, A.; Fakhfakh-Zouari, N.; Hmidet, N.; Frikha, F.; Nasri, M.; Kamoun, A.S. Low-Cost Fermentation Medium for Alkaline Protease Production by Bacillus Mojavensis A21 Using Hulled Grain of Wheat and Sardinella Peptone. J. Biosci. Bioeng. 2010, 110, 288–294. [Google Scholar] [CrossRef]
  206. Nidheesh, T.; Pal, G.K.; Suresh, P.V. Chitooligomers Preparation by Chitosanase Produced under Solid State Fermentation Using Shrimp By-Products as Substrate. Carbohydr. Polym. 2015, 121, 1–9. [Google Scholar] [CrossRef]
  207. Doan, C.T.; Tran, T.N.; Wang, S.L. Production of Thermophilic Chitinase by Paenibacillus Sp. Tku052 by Bioprocessing of Chitinous Fishery Wastes and Its Application in n-Acetyl-d-Glucosamine Production. Polymers 2021, 13, 3048. [Google Scholar] [CrossRef]
  208. Matsumoto, Y.; Saucedo-Castañeda, G.; Revah, S.; Shirai, K. Production of β-N-Acetylhexosaminidase of Verticillium Lecanii by Solid State and Submerged Fermentations Utilizing Shrimp Waste Silage as Substrate and Inducer. Process. Biochem. 2004, 39, 665–671. [Google Scholar] [CrossRef]
  209. dos Santos Aguilar, J.G.; Sato, H.H. Microbial Proteases: Production and Application in Obtaining Protein Hydrolysates. Food Res. Int. 2018, 103, 253–262. [Google Scholar] [CrossRef] [PubMed]
  210. Research and Markets Industrial Enzymes—Global Strategic Business Report; Dublin. 2024. Available online: https://www.researchandmarkets.com/reports/338655/industrial_enzymes_global_strategic_business?srsltid=AfmBOorl1xyR_hpZObN7m58k-RClBrXom0hy_PINbtssJ2rwVYbfurcl (accessed on 5 September 2024).
  211. Rojas, L.F.; Zapata, P.; Ruiz-Tirado, L. Agro-Industrial Waste Enzymes: Perspectives in Circular Economy. Curr. Opin. Green Sustain. Chem. 2021, 34, 100585. [Google Scholar] [CrossRef]
  212. Téllez-Luis, S.J.; Ramírez, J.A.; Vázquez, M. Production of Transglutaminase by Streptoverticillium Ladakanum NRRL-3191 Using Glycerol as Carbon Source. Food Technol. Biotechnol. 2004, 42, 75–81. [Google Scholar]
  213. Srivastava, N. Production of Food-Processing Enzymes from Recombinant Microorganisms. In Enzymes in Food Biotechnology: Production, Applications, and Future Prospects; Academic Press: Cambridge, MA, USA, 2018. [Google Scholar]
  214. Elbadawi, M.; Li, H.; Basit, A.W.; Gaisford, S. The Role of Artificial Intelligence in Generating Original Scientific Research. Int. J. Pharm. 2024, 652, 123741. [Google Scholar] [CrossRef]
  215. Sampaio, P.S.; Fernandes, P. Machine Learning: A Suitable Method for Biocatalysis. Catalysts 2023, 13, 961. [Google Scholar] [CrossRef]
  216. Chauhan, T.; Rawat, S.; Malik, S.; Singh, P. Supervised and Unsupervised Machine Learning Based Review on Diabetes Care. In Proceedings of the 2021 7th International Conference on Advanced Computing and Communication Systems, ICACCS, Coimbatore, India, 19–20 March 2021. [Google Scholar]
  217. Rashidi, H.H.; Tran, N.K.; Betts, E.V.; Howell, L.P.; Green, R. Artificial Intelligence and Machine Learning in Pathology: The Present Landscape of Supervised Methods. Acad. Pathol. 2019, 6, 2374289519873088. [Google Scholar] [CrossRef]
  218. Rajoub, B. Supervised and Unsupervised Learning. In Biomedical Signal Processing and Artificial Intelligence in Healthcare; Elsevier: Amsterdam, The Netherlands, 2020; pp. 51–89. ISBN 9780128189467. [Google Scholar]
  219. Nasteski, V. An Overview of the Supervised Machine Learning Methods. Horizons B 2017, 4, 51–62. [Google Scholar] [CrossRef]
  220. Joshi, D.J.; Kale, I.; Gandewar, S.; Korate, O.; Patwari, D.; Patil, S. Reinforcement Learning: A Survey. In Proceedings of the Advances in Intelligent Systems and Computing, Hangzhou, China, 29–31 May 2021; Volume 1311 AISC. [Google Scholar]
  221. Wahid, Z.; Nadir, N. Improvement of One Factor at a Time through Design of Experiments. World Appl. Sci. J. 2013, 21, 56–61. [Google Scholar] [CrossRef]
  222. Bhardwaj, A.; Kishore, S.; Pandey, D.K. Artificial Intelligence in Biological Sciences. Life 2022, 12, 1430. [Google Scholar] [CrossRef]
  223. Bhagat, P. Introduction to Neural Nets. Chem. Eng. Prog. 1990, 86, 229–255. [Google Scholar] [CrossRef]
  224. Agatonovic-Kustrin, S.; Beresford, R. Basic Concepts of Artificial Neural Network (ANN) Modeling and Its Application in Pharmaceutical Research. J. Pharm. Biomed. Anal. 2000, 22, 717–727. [Google Scholar] [CrossRef] [PubMed]
  225. Singh, V.; Haque, S.; Niwas, R.; Srivastava, A.; Pasupuleti, M.; Tripathi, C.K.M. Strategies for Fermentation Medium Optimization: An in-Depth Review. Front. Microbiol. 2017, 7, 2087. [Google Scholar] [CrossRef] [PubMed]
  226. Kennedy, M.; Krouse, D. Strategies for Improving Fermentation Medium Performance: A Review. J. Ind. Microbiol. Biotechnol. 1999, 23, 456–475. [Google Scholar] [CrossRef]
  227. Chen, L.Z.; Nguang, S.K.; Chen, X.D.; Li, X.M. Modelling and Optimization of Fed-Batch Fermentation Processes Using Dynamic Neural Networks and Genetic Algorithms. Biochem. Eng. J. 2004, 22, 51–61. [Google Scholar] [CrossRef]
  228. Fernández Núñez, E.G.; Barchi, A.C.; Ito, S.; Escaramboni, B.; Herculano, R.D.; Mayer, C.R.M.; de Oliva Neto, P. Artificial Intelligence Approach for High Level Production of Amylase Using Rhizopus Microsporus Var. Oligosporus and Different Agro-Industrial Wastes. J. Chem. Technol. Biotechnol. 2017, 92, 684–692. [Google Scholar] [CrossRef]
  229. Singh, A.; Majumder, A.; Goyal, A. Artificial Intelligence Based Optimization of Exocellular Glucansucrase Production from Leuconostoc Dextranicum NRRL B-1146. Bioresour. Technol. 2008, 99, 8201–8206. [Google Scholar] [CrossRef]
  230. Bezerra, C.O.; Carneiro, L.L.; Carvalho, E.A.; das Chagas, T.P.; de Carvalho, L.R.; Uetanabaro, A.P.T.; da Silva, G.P.; da Silva, E.G.P.; da Costa, A.M. Artificial Intelligence as a Combinatorial Optimization Strategy for Cellulase Production by Trichoderma Stromaticum AM7 Using Peach-Palm Waste Under Solid-State Fermentation. Bioenergy Res. 2021, 14, 1161–1170. [Google Scholar] [CrossRef]
  231. De Farias Silva, C.E.; Costa, G.Y.S.C.M.; Ferro, J.V.; de Oliveira Carvalho, F.; da Gama, B.M.V.; Meili, L.; dos Santos Silva, M.C.; Almeida, R.M.R.G.; Tonholo, J. Application of Machine Learning to Predict the Yield of Alginate Lyase Solid-State Fermentation by Cunninghamella Echinulata: Artificial Neural Networks and Support Vector Machine. React. Kinet. Mech. Catal. 2022, 135, 3155–3171. [Google Scholar] [CrossRef]
  232. Suryawanshi, N.; Eswari, J.S. Chitin from Seafood Waste: Particle Swarm Optimization and Neural Network Study for the Improved Chitinase Production. J. Chem. Technol. Biotechnol. 2022, 97, 509–519. [Google Scholar] [CrossRef]
  233. Suryawanshi, N.; Eswari, J.S. Purification and Characterization of Chitinase Produced by Thermophilic Fungi Thermomyces Lanuginosus. Prep. Biochem. Biotechnol. 2022, 52, 1087–1095. [Google Scholar] [CrossRef]
  234. Ekpenyong, M.; Asitok, A.; Ben, U.; Amenaghawon, A.; Kusuma, H.; Akpan, A.; Antai, S. Application of the Novel Manta-Ray Foraging Algorithm to Optimize Acidic Peptidase Production in Solid-State Fermentation Using Binary Agro-Industrial Waste. Prep. Biochem. Biotechnol. 2024, 54, 226–238. [Google Scholar] [CrossRef]
Marinedrugs 22 00411 g001
Figure 1. Approximate percentages of waste generated during processing of fish and shellfish: (A) wastes generated during processing of fish, and (B) waste generated during processing of shellfish (i.e., shrimps and crabs).
Figure 1. Approximate percentages of waste generated during processing of fish and shellfish: (A) wastes generated during processing of fish, and (B) waste generated during processing of shellfish (i.e., shrimps and crabs).
Marinedrugs 22 00411 g001
Marinedrugs 22 00411 g002
Figure 2. Structures of representative proteases: (A) pepsin, (B) trypsin, (C) α-chymotrypsin and (D) pancreatic elastase. The enzyme structures were obtained from the Research Collaboratory for Structural Bioinformatics Protein Data Bank—RCSB PDB (RCSB.org)—using PDB ID 1AM5 for pepsin from Atlantic cod [44], PDB ID 1HJ8 for trypsin from Atlantic salmon [45], PDB ID 4CHA for bovine α-chymotrypsin [46] and PDB ID 1ELT for elastase from North Atlantic salmon [47]. The different colors in subfigure (C) represent the different chains in α-chymotrypsin.
Figure 2. Structures of representative proteases: (A) pepsin, (B) trypsin, (C) α-chymotrypsin and (D) pancreatic elastase. The enzyme structures were obtained from the Research Collaboratory for Structural Bioinformatics Protein Data Bank—RCSB PDB (RCSB.org)—using PDB ID 1AM5 for pepsin from Atlantic cod [44], PDB ID 1HJ8 for trypsin from Atlantic salmon [45], PDB ID 4CHA for bovine α-chymotrypsin [46] and PDB ID 1ELT for elastase from North Atlantic salmon [47]. The different colors in subfigure (C) represent the different chains in α-chymotrypsin.
Marinedrugs 22 00411 g002
Marinedrugs 22 00411 g003
Figure 3. Structure of a representative transglutaminase. The enzyme structure was obtained from the Research Collaboratory for Structural Bioinformatics Protein Data Bank—RCSB PDB (RCSB.org)—using PDB ID 1G0D for transglutaminase from red sea bream [165].
Figure 3. Structure of a representative transglutaminase. The enzyme structure was obtained from the Research Collaboratory for Structural Bioinformatics Protein Data Bank—RCSB PDB (RCSB.org)—using PDB ID 1G0D for transglutaminase from red sea bream [165].
Marinedrugs 22 00411 g003
Table 1. Kinetic properties of cold-adapted fish enzymes compared to mesophilic mammalian enzymes at various temperatures *.
Table 1. Kinetic properties of cold-adapted fish enzymes compared to mesophilic mammalian enzymes at various temperatures *.
EnzymeSourceTemperature
(°C)		Km
(mM)		kcat
(s−1)		kcat/Km
(s−1 mM−1)		References
TrypsinBovine40.300.210.70[31]
Porcine0.961.041.08
Fish0.0370.9024.3
Bovine200.500.711.42[31]
Porcine0.821.551.89
Fish0.0501.7735.4
Bovine370.721.351.84[31]
Porcine0.912.722.99
Fish0.0682.6939.6
ElastasePorcine100.7211.616.1[32]
Fish0.7924.631.1
Porcine250.7316.122.1[32]
Fish1.0244.143.2
* Adapted from Smalås et al. [33]. Km: Michaelis constant; Kcat: turnover number; kcat/Km: specificity constant.
Table 2. Proteases from fishery and aquaculture waste *.
Table 2. Proteases from fishery and aquaculture waste *.
ProteaseGroupMarine Species
PepsinAspartyl
protease		Smooth hound (Mustelus mustelus) [52], European eel (Anguilla anguilla) [53], Atlantic cod (Gadus morhua) [54], sea bream (Sparus latus) [55], albacore tuna (Thunnus alalunga) [56,57], skipjack tuna (Katsuwonus pelamis) [57], tongol tuna (Thunnus tonggol) [57], mandarin fish (Siniperca chuatsi) [58] and pectoral rattail (Coryphaenoides pectoralis) [59]
TrypsinSerine
protease		Threadfin hakeling (laemonema longipes) [60], unicorn leatherjacket (Aluterus monoceros) [61], pectoral rattail (Coryphaenoides pectoralis) [62], zebra blenny (Salaria basilisca) [63], Japanese sea bass (Lateolabrax japonicus) [64], tongol tuna (Thunnus tonggol) [65], common kilka (Clupeonella cultriventris caspia) [66], skipjack tuna (Katsuwonus pelamis) [67], spotted mackerel (Scomber australasicus) [68], walleye pollock (Theragra chalcogramma) [69], Pacific saury (Cololabis saira) [70] and cuttlefish (Sepia officinalis) [71]
ChymotrypsinSerine
protease		Crucian carp (Carassius auratus) [72] and Pacific sardine (Sardinops sagax caeruleus) [73,74]
Elastase and collagenaseSerine
protease		Haddock (Melanogrammus aeglefinus) [75], Atlantic herring (Clupea harengus) [75,76], flounder (Paralichthys dentatus) [75], sardine (Sardina pilchardus) [76], butterfly peacock bass (Cichla ocellaris) [77] and rough scad (Trachurus lathami) [77]
BrachyurinSerine
protease		Northern shrimp (Pandalus borealis) [78,79], Northern shrimp (Pandalus eous) [80], white shrimp (Penaeus vannamei) [81], greasyback shrimp (Metapenaeus ensis) [82], American lobster (Homarus americanus) [83], Caribbean spiny lobster (Panulirus argus) [49] and spiny lobster (Panulirus interruptus) [84]
CathepsinCysteine
protease		Atlantic cod (Gadus morhua) [85], sea cucumber (Stichopus japonicus) [86,87], rock bream (Oplegnathus fasciatus) [88], Atlantic herring (Clupea harengus) [89] and neon flying squid (Ommastrephes bartramii) [90]
Proteolytic enzymesMetallo-
protease		sea cucumber (Stichopus japonicus) [41], rainbow trout (Oncorhynchus mykiss) [42] and trout (Salmo gairdnerii) [43]
* This information (non-exhaustive) was compiled from a number of papers published during the past two decades (2004–2024).
Table 3. Examples of fishery and aquaculture waste used as sources of lipases.
Table 3. Examples of fishery and aquaculture waste used as sources of lipases.
Fishery and Aquaculture WasteMarine SpeciesReferences
Pyloric caeca and pancreascod (Gadus morhua)[96]
PancreasAtlantic salmon (Salmo salar)[97]
rainbow trout (Oncorhynchus mykiss)[98]
Hepatopancreassea bream (Pagrus major)[94]
sardine (Sardinella longiceps)[99]
Stomach, pyloric caeca, liver and intestinerohu (Labeo rohita)[95]
Indian oil sardine (Sardinella longiceps)[95]
mullet (Liza subviridis)[95]
Indian mackerel (Rastrelliger kanagurta)[95]
Stomach and intestinetilapia (Oreochromis niloticus)[100]
Visceracatfish (Clarias macrocephalus) [92]
catfish (Clarias gariepinus)[92]
snakehead (Channa stiata)[92]
Nile tilapia (Oreochromis niloticus)[92]
Pacific sardine (Sardinops sagax caerulea)[92,101]
tuna species (Euthynnusaffinis)[102]
Intestines, pancreas and pyloric caecamilkfish (Chanos chanos)[103]
Table 5. Examples of fishery and aquaculture waste used for production of microbial enzymes.
Table 5. Examples of fishery and aquaculture waste used for production of microbial enzymes.
Fishery and Aquaculture WasteMicroorganismsEnzymesReferences
Fish heads and visceraPseudomonas aeruginosa *Protease[177]
Bacillus subtilisProtease[178]
Fish visceraVibrio anguillarum *Protease[179]
Vibrio splendidus *Protease[179]
Fish headsStreptomyces speibonaeProtease[180]
Paenibacillus elgiiProtease[181]
Shrimp shellsBacillus subtilisProtease[182]
Chryseobacterium indologenes *Protease[183]
Bacillus subtilisChitosanase[184]
Pseudomonas *Chitinase[185]
Pseudomonas *Chitosanase[185]
Bacillus cereus *Protease[186]
Bacillus cereus *Chitinase[186]
Serratia marcescens *Protease[187]
Serratia marcescens *Chitosanase[187,188]
Lecanicillium muscariumβ-N-acetyl-hexosaminidase[189]
Paenibacillus mucilaginosusChitosanase[190]
Shrimp and crab shellsMonascus purpureus *Protease[191]
Bacillus firmusProtease[192]
Bacillus amyloliquefaciensChitinase[193]
Monascus purpureusChitinase[194]
Bacillus cereus *Chitinase[195]
Cuttlefish by-productsBacillus licheniformisProtease[176]
Bacillus subtilisProtease[176]
Pseudomonas aeruginosa *Protease[176]
Bacillus cereus *Protease[176]
Vibrio parahaemolyticus *Protease[176]
Squid pensLactobacillus paracaseiProtease[196]
Serratia ureilytica *Protease[197]
LentzeaChitosanase[198]
Serratia ureilytica *Chitinase[197]
Paenibacillus sp.Protease[199]
Paenibacillus sp.Chitosanase[199,200]
Paenibacillus elgiiChitosanase[201]
Fish by-productsStaphylococcus epidermidis *Lipase[202]
Bacillus cereus *Protease[203]
Fish and shellfish wastesStaphylococcus xylosusLipase[204]
Hydrolyzed fish wastesRhizopus oryzaeLipase[172]
Bacillus mojavensisProtease[205]
Shrimp by-productsPurpureocillium lilacinum *Chitosanase[206]
Chitinous fishery wastesPaenibacillus sp.Chitinase[207]
Shrimp waste silageVerticillium lecaniiβ-N-acetyl-hexosaminidase[208]
* Indicates that the microorganism could pose potential health and safety issues.
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Khiari, Z. Enzymes from Fishery and Aquaculture Waste: Research Trends in the Era of Artificial Intelligence and Circular Bio-Economy. Mar. Drugs 2024, 22, 411. https://doi.org/10.3390/md22090411

AMA Style

Khiari Z. Enzymes from Fishery and Aquaculture Waste: Research Trends in the Era of Artificial Intelligence and Circular Bio-Economy. Marine Drugs. 2024; 22(9):411. https://doi.org/10.3390/md22090411

Chicago/Turabian Style

Khiari, Zied. 2024. "Enzymes from Fishery and Aquaculture Waste: Research Trends in the Era of Artificial Intelligence and Circular Bio-Economy" Marine Drugs 22, no. 9: 411. https://doi.org/10.3390/md22090411

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Article metric data becomes available approximately 24 hours after publication online.
Back to TopTop