Submitted Successfully!
To reward your contribution, here is a gift for you: A free trial for our video production service.
Thank you for your contribution! You can also upload a video entry or images related to this topic.
Version Summary Created by Modification Content Size Created at Operation
1 -- 2759 2023-12-18 15:56:07 |
2 Format correct -1 word(s) 2758 2023-12-19 14:02:33 |

Video Upload Options

Do you have a full video?


Are you sure to Delete?
If you have any further questions, please contact Encyclopedia Editorial Office.
Hasan, I.; Gai, F.; Cirrincione, S.; Rimoldi, S.; Saroglia, G.; Terova, G. Enhanced Digestibility and Nutrient Utilization with Chitinase. Encyclopedia. Available online: (accessed on 20 June 2024).
Hasan I, Gai F, Cirrincione S, Rimoldi S, Saroglia G, Terova G. Enhanced Digestibility and Nutrient Utilization with Chitinase. Encyclopedia. Available at: Accessed June 20, 2024.
Hasan, Imam, Francesco Gai, Simona Cirrincione, Simona Rimoldi, Giulio Saroglia, Genciana Terova. "Enhanced Digestibility and Nutrient Utilization with Chitinase" Encyclopedia, (accessed June 20, 2024).
Hasan, I., Gai, F., Cirrincione, S., Rimoldi, S., Saroglia, G., & Terova, G. (2023, December 18). Enhanced Digestibility and Nutrient Utilization with Chitinase. In Encyclopedia.
Hasan, Imam, et al. "Enhanced Digestibility and Nutrient Utilization with Chitinase." Encyclopedia. Web. 18 December, 2023.
Enhanced Digestibility and Nutrient Utilization with Chitinase

The aquaculture industry is looking for sustainable alternatives to conventional fish meals in fish feed, and insect-based meals are proving to be a promising solution. These meals are nutritionally optimal as they have a high protein content and an ideal amino acid profile. However, the presence of chitin, a component of the insect exoskeleton in these meals presents both an opportunity and a challenge. Chitosan, a derivative of chitin, is known to improve the physiological functions of fish, including growth, immunity, and disease resistance. While chitin and its derivative chitosan offer several physiological benefits, their presence can affect the digestibility of feed in some fish species, making the inclusion of insect-based meals in aquafeeds complex. Chitinase, an enzyme that breaks down chitin, is being investigated as a potential solution to improve the nutritional value of insect meals in aquafeed.

chitinase insect meal aquaculture chitosan digestibility nutrient

1. Introduction

The aquaculture industry is currently facing a crucial challenge: the search for sustainable and economically viable alternatives to conventional fishmeal (FM) in fish feeds. One promising solution on the horizon is the use of insect-based meals as an alternative protein source due to their optimal nutritional properties, especially their high protein content and ideal amino acid profile [1][2], but also due to their potential to meet the growing demand for alternative protein sources in aquaculture feeds. Their potential is further enhanced by recent advances in processing, economic viability, and scalability [3]. Insect meals (IMs) are increasingly proving their suitability as a partial or complete replacement for FM in aquaculture feeds. Numerous feeding trials conducted by different research groups [1][4][5][6][7][8][9][10] have produced promising results in this respect. These studies highlight the potential of IM as a sustainable and effective alternative to conventional FM in aquaculture feed formulations. In recent studies, it has been found that encouraging results with the incorporation of black soldier fly (Hermetia illucens) and yellow mealworm (Tenebrio molitor) meals into the feeds of carnivorous marine and freshwater fish [6][7][8][9][10][11]. These promising results are an important step towards the use of IMs as a sustainable alternative in aquaculture feeds and offer the aquaculture industry a viable way to overcome the challenges of sustainability and economic feasibility.
The use of insect meal in aqua feed can vary depending on the specific circumstances and the type of fish. Research indicates that IM can replace a significant proportion of FM in aquaculture feeds. For example, mealworms and housefly larvae meals can replace up to 40–80% and 75% of FM in the diet of Nile tilapia/standard catfish, respectively [1]. The use of IM as a sustainable alternative to FM in aquaculture has also shown promising results [2]. However, some difficult issues still need to be resolved, including the cost and expansion of use [2]. Chitin, a major component of the insect exoskeleton, can pose a challenge for digestibility. Monitoring the chitin content in insect meal and its impact on nutrient digestibility is critical for determining appropriate incorporation rates.
Insects do indeed offer a unique nutritional profile for aquaculture feeds, and a notable component of them is chitin. Chitin is a polysaccharide, the second most abundant on earth after cellulose, consisting of N-acetyl-2-amino-2-deoxyglucose (GlcNAc) units linked by β-(1→4) bonds [11][12][13]. Chitin and its deacetylated derivative chitosan represent a significant source of insoluble “animal” fibers that may have potential utility as functional ingredients, or bioactive compounds in aquaculture feeds [12][14][15]. This component adds an interesting dimension to the nutritional value of IMs, and ongoing research is investigating the various ways in which chitin can positively impact the health and performance of fish in aquaculture systems.
Numerous studies have shown that chitosan plays an important role in improving various physiological functions in fish. These effects include improved growth performance, enhanced immunity, and improved antimicrobial capabilities, which underlines the multiple benefits of chitosan in fish farming [12]. The inclusion of chitin and chitosan in fish feed has various positive effects on different fish species in aquaculture. These effects include increased growth rates, improved feed efficiency, and increased disease resistance in species such as rainbow trout, Nile tilapia, grey mullet, red sea bream, Japanese eel, and yellowtail [16][17][18][19][20][21]. While chitosan and chitin in IMs provide numerous benefits, it is important to recognize that the presence of chitin can be a limiting factor in the use of insect-based feeds in aquaculture. The presence of chitin in fish feed can reduce the digestibility of the feed, as most fish species cannot efficiently digest and absorb chitin, as found in the study by [22]. However, the ability of fish to digest chitin is controversial as other fish species, such as cod (Gadus morhua) [23], juvenile cobia (Rachycentron canadum) [24], channel rockcod (Sebastolobus alascanus), splitnose rockfish (Sebastes diploproa), and black cod (Anoplopoma fimbria) [25] have shown activity of chitinase, the enzyme responsible for the degradation of chitin. These differences in the ability of fish species to digest chitin make the incorporation of IMs into aquafeeds even more complex.
Similar to cellulose, chitin can potentially serve as a prebiotic in fish diets. This is because it can increase bacterial diversity in the gut by promoting the proliferation of beneficial and chitin-degrading bacteria. As a result, chitin can stimulate fermentation in the gut, leading to the production of essential short-chain fatty acids, with acetate, propionate, and butyrate being the main end products of bacterial fermentation. This mechanism contributes to the improvement of intestinal health and the general well-being of fish [22][26].
Numerous studies have shown that insect-derived ingredients can effectively influence the microbial communities in the fish gut. Feeding insects to fish often results in a remarkable increase in the abundance of bacterial families, such as Bacillaceae, Lactobacillaceae, and Actinobacteria [6][8][9][10][27][28][29][30]. Similarly, adding chitosan to the feed has been found to promote the growth of beneficial bacteria in the fish gut while reducing the presence of potentially harmful pathogens [16]. In contrast to the potential benefits of chitin in fish feed, some studies have reported a reduced growth rate in salmon fed a chitin-rich diet. It has been hypothesized that chitin may act as an energy reservoir when fish are unable to efficiently digest and utilize this polysaccharide, suggesting that improper chitin digestion may affect fish growth and overall performance [31][32].
Chitinase and IM are becoming important components in aquaculture nutrition. The unique properties of insects and their suitability for use in aquafeeds as a substitute for FM have become a focus of recent research in aquaculture [33]. The use of the enzyme chitinase could enable the inclusion of higher amounts of IM in aquaculture feeds and thus increase the usefulness of IM [34]. Chitinase is an enzyme that breaks down chitin, a major component of the insect exoskeleton, into digestible carbohydrates. This process can increase the nutritional value of IM and make it more digestible for aquatic species. These enzymes have been detected in the digestive tracts of a variety of fish species, as demonstrated by several studies [35][36][37][38]. Fish species in which these enzymes have been detected include rainbow trout (Oncorhynchus mykiss) [39]. The growth of rainbow trout fed diets containing different amounts of chitin and the relationship between chitinolytic enzymes and chitin digestibility showed that the growth was significantly reduced when the diets contained, 4, 10 and 25% chitin compared to that of the controls fed starchy diets. There was no difference in growth rate between control fish and fish fed 10% N-acetylglucosamine (GlcNAc). A relatively high chitinase activity was found in the stomachs and a relatively high chitobiase activity in the intestines. These enzyme activities were similar in all trout, regardless of the amount of chitin in their diet, except that chitobiase in the gut of fish fed a diet containing GlcNAc had higher activity levels than controls. The study also found that chitin was not significantly digested when fed at levels of 10 and 30% of the diet and that the presence of antibiotics or live chitinolytic bacteria (Vibrio alginolyticus) in the diet had no effect on the digestibility of chitin. This suggests an endogenous origin of the chitinolytic enzymes in the gastrointestinal tract of trout [39]. Most of these chitin-degrading enzymes are found in the gastrointestinal tract of fish and exhibit acid-resistant activities when it comes to the degradation of insoluble chitin substrates. This resistance to acidic conditions suggests that these enzymes have the potential to effectively digest chitin-rich feeds, as shown in the study by [36]. Other studies have suggested that chitinase activity in fish is not sufficient to digest chitin [40]. However, some studies have hypothesized that fish are able to digest chitin to some extent and that the chitin metabolites are absorbed, suggesting that chitin may act as both a nutrient source and an antinutrient in fish [39]. The primary function of fish chitinases appears to be to chemically disrupt the exoskeleton or other chitin-containing external structures of prey so that the internal nutrients can be reached by the enzymes or to prevent blockage of the gut by these structures [41].

2. Fungal Chitinases: A Key to Improved Digestibility

The extraction of chitinases from other insects or microbial organisms is a promising way to enable the use of chitinases in fish feed. Fungi are the dominant group of chitinase producers among microorganisms, and many efficient chitinolytic fungi with potential applications have been identified in a variety of environments, including soil, water, marine litter, and plants. This suggests that fungal chitinases may be a sustainable option for chitin degradation in aquaculture [42]. Recent research has shown that the use of fungal chitinases in IM containing feeds may be a promising strategy to improve nutrient uptake in marine fish such as European seabass (Dicentrarchus labrax), which is one of the most commonly farmed species in Mediterranean aquaculture [43][44].
Aeromonas veronii B565, a bacterium isolated from the sediment of aquaculture ponds, produces chitinases, enzymes that can degrade chitin, a component of the cell walls of fungi and the exoskeletons of insects and crustaceans. These chitinases have been proposed as a potential means of controlling Myxozoa-related or fungal diseases in fish [45][46][47]. Myxozoa are a class of microscopic parasites that can cause severe damage in aquaculture, resulting in significant economic losses. Currently, there are no effective treatments for Myxozoa infections, so the development of new control strategies is a high priority [46].
Chitinolytic bacteria capable of producing chitinases were isolated from the gut microbiota of European seabass fed with different IMs [43]. These bacteria can help to degrade chitin, a major component of the exoskeleton of many marine organisms, which is generally indigestible for several economically valuable fish species [24][43]. This may improve the use of feeds containing a high proportion of chitin-rich IM. The degradation of chitin by chitinases can improve nutrient uptake in seabass. By degrading chitin, chitinases can improve feed digestibility and facilitate the access of digestive enzymes to entrapped proteins or lipids, thereby improving overall digestive health and nutrient absorption in seabass [43][44].
Chitin derivatives produced by chitinase-producing bacteria have been shown to modulate the immunological response of fish and improve disease resistance [44]. In particular, the study showed that red blood cell and white blood cell counts increased significantly in sea bass fed chitin derivatives, indicating an improved immune response. Bioconversion of chitin waste by Stenotrophomonas maltophilia to produce chitin derivatives has been shown to have a significant effect on the non-specific immune response of seabass [44]. In addition, fungal polysaccharides, which include chitin, can regulate the growth and immune response of fish, enhancing immune response and disease resistance [48]. This suggests that chitin derivatives produced by fungal chitinases may have immunomodulatory effects in fish.
A study on juvenile cobia (Rachycentron canadum) found that high levels of chitinase in the digestive tract are associated with fish that lack the mechanical structures to break down chitinous material [24]. This suggests that the use of fungal chitinases in fish feed may be particularly beneficial for species with limited ability to digest chitin. The use of fungal chitinases in fish feed containing IM has the potential to improve nutrient uptake and increase the nutritional value of the feed for marine fish, such as sea bass. Several fungal chitinases have been already investigated for use in aquaculture (Table 1).
Table 1. Studies evaluating the effects of different fungal chitinases used in aquaculture.
Fungal Chitinases Major Finding(s) References
Aspergillus flavus Fungal chitinases from Aspergillus flavus have been shown to have an anti-fungal effect against many plant pathogenic fungi. This indicates that Aspergillus flavus chitinases can be used to control pests and diseases in aquaculture. [49]
Mucor Chitin-degrading fungi such as Mucor have been identified in the aquatic environment. This indicates that Mucor chitinases can be used for chitin degradation in aquaculture. [50]
Trichoderma Fungal chitinases from Trichoderma facilitate mycoparasitism of other fungi. This suggests that Trichoderma chitinases can be used to control fungal infections in aquaculture. [42]
Aspergillus sp. S1–13 Aspergillus sp. S1–13 has been shown to synthesize chitinolytic enzymes when grown on a medium containing shrimp waste. This suggests that the chitinases of Aspergillus sp. S1–13 can be used to convert chitin waste from fisheries into simpler, useful components to reduce water pollution. [50]
Fungal chitinases can help break down chitin, a major component of the exoskeleton of many marine organisms. This process can improve nutrient uptake in fish, making feed more efficient and beneficial for the fish [49][50]. Chitinases from fungi have been shown to have immunomodulatory effects. For example, fungal polysaccharides, which include chitin, can regulate the growth and immune response of fish, enhancing immune response and disease resistance [48]. Fungi are the dominant group of chitinase producers among microorganisms, and many efficient chitinolytic fungi with potential applications have been identified in a variety of environments, including soil, water, marine litter, and plants [49]. This suggests that fungal chitinases may be a sustainable option for mediating chitin degradation in aquaculture. In addition, fungal chitinases can be used to convert chitin-containing waste from fisheries into simpler, useful components, thereby reducing water pollution and contributing to waste management in aquaculture [51].

3. Enhanced Nutrient Utilization with Chitinase

The enzyme chitinase hydrolyses chitin into digestible carbohydrates that can be used in fish feed, thus increasing the nutritional value of the feed [47][52]. One of the ways in which chitinase improves nutrient uptake is by breaking down chitin-containing biomass, such as the waste of marine organisms, into simpler, useful components [52]. Once chitin is broken down into simpler oligosaccharides, these molecules become more bioavailable. This means that they can be more easily absorbed in the animal’s digestive tract, contributing to overall nutrient uptake. This process effectively converts chitin waste into a usable form that can be added to fish feed, contributing to improved nutrient utilization [53].
Preliminary studies have shown that chitin oligosaccharides, products of chitinase activity, could serve as prebiotics. They may promote the growth of beneficial gut bacteria, which in turn play a role in nutrient absorption and overall gut health. Studies have shown that dietary chitin or its derivative chitosan acts as a prebiotic, modulating microbial communities in the gut of fish [11]. This modulation can lead to improved nutrient uptake and the general health of the fish. In addition, chitinase can be used in the biocontrol of pathogenic fungi and harmful insects as it acts on chitin, a major component of their structures. This function can contribute to disease management in aquaculture as it can help in the control of pathogenic fungi and harmful insects that have chitin structures [53].
Furthermore, the introduction of chitinase into the feed of fish can lead to an increase in chitin digestibility and consequently to an improvement in nutrient digestibility. Studies have shown that the inclusion of chitinase in the diet can improve chitin digestibility in fish such as juvenile cobia and Nile tilapia, resulting in higher growth rates and more efficient nutrient utilization [24][34][53]. The impact on aquaculture productivity is significant. By improving nutrient utilization, chitinase can contribute to the growth and health of aquatic organisms, leading to higher productivity in aquaculture. However, it is important to note that the digestibility of chitin may decrease with higher levels of chitin in the diet [53]. Therefore, while the inclusion of chitinase in the diet may improve digestibility to some extent, the specific increase to be expected may vary depending on chitin intake and fish species.
By adding chitinase, chitin-rich feed ingredients can be utilized more effectively. This can lead to a reduction in feed waste and improved feed conversion, which in turn leads to more sustainable and cost-effective farming. Improved nutrient absorption often correlates with better overall health. Furthermore, the use of chitinase can help to avoid chitin waste, which is a significant environmental problem in aquaculture. Despite these potential benefits, it is difficult to increase microbial chitinase production due to the inducibility of the enzyme, low titer, high production costs, and susceptibility to challenging environmental conditions. Therefore, further research is needed to optimize the production and application of chitinase in aquaculture [52].


  1. Nogales-Mérida, S.; Gobbi, P.; Józefiak, D.; Mazurkiewicz, J.; Dudek, K.; Rawski, M.; Kierończyk, B.; Józefiak, A. Insect Meals in Fish Nutrition. Rev. Aquac. 2019, 11, 1080–1103.
  2. Hasan, I.; Rimoldi, S.; Saroglia, G.; Terova, G. Sustainable Fish Feeds with Insects and Probiotics Positively Affect Freshwater and Marine Fish Gut Microbiota. Animals 2023, 13, 1633.
  3. Hua, K.; Cobcroft, J.M.; Cole, A.; Condon, K.; Jerry, D.R.; Mangott, A.; Praeger, C.; Vucko, M.J.; Zeng, C.; Zenger, K.; et al. The Future of Aquatic Protein: Implications for Protein Sources in Aquaculture Diets. One Earth 2019, 1, 316–329.
  4. Gasco, L.; Acuti, G.; Bani, P.; Dalle Zotte, A.; Danieli, P.P.; De Angelis, A.; Fortina, R.; Marino, R.; Parisi, G.; Piccolo, G.; et al. Insect and Fish By-Products as Sustainable Alternatives to Conventional Animal Proteins in Animal Nutrition. Ital. J. Anim. Sci. 2020, 19, 360–372.
  5. Lock, E.-J.; Biancarosa, I.; Gasco, L. Insects as Raw Materials in Compound Feed for Aquaculture. In Edible Insects in Sustainable Food Systems; Springer: Berlin/Heidelberg, Germany, 2018; pp. 263–276. ISBN 978-3-319-74011-9.
  6. Rimoldi, S.; Antonini, M.; Gasco, L.; Moroni, F.; Terova, G. Intestinal Microbial Communities of Rainbow Trout (Oncorhynchus mykiss) May Be Improved by Feeding a Hermetia illucens Meal/Low-Fishmeal Diet. Fish Physiol. Biochem. 2021, 47, 365–380.
  7. Rimoldi, S.; Gini, E.; Iannini, F.; Gasco, L.; Terova, G. The Effects of Dietary Insect Meal from Hermetia illucens Prepupae on Autochthonous Gut Microbiota of Rainbow Trout (Oncorhynchus mykiss). Animals 2019, 9, 143.
  8. Terova, G.; Gini, E.; Gasco, L.; Moroni, F.; Antonini, M.; Rimoldi, S. Effects of Full Replacement of Dietary Fishmeal with Insect Meal from Tenebrio molitor on Rainbow Trout Gut and Skin Microbiota. J. Anim. Sci. Biotechnol. 2021, 12, 30.
  9. Terova, G.; Ceccotti, C.; Ascione, C.; Gasco, L.; Rimoldi, S. Effects of Partially Defatted Hermetia illucens Meal in Rainbow Trout Diet on Hepatic Methionine Metabolism. Animals 2020, 10, 1059.
  10. Terova, G.; Rimoldi, S.; Ascione, C.; Gini, E.; Ceccotti, C.; Gasco, L. Rainbow Trout (Oncorhynchus mykiss) Gut Microbiota Is Modulated by Insect Meal from Hermetia illucens Prepupae in the Diet. Rev. Fish Biol. Fish 2019, 29, 465–486.
  11. Rimoldi, S.; Ceccotti, C.; Brambilla, F.; Faccenda, F.; Antonini, M.; Terova, G. Potential of Shrimp Waste Meal and Insect Exuviae as Sustainable Sources of Chitin for Fish Feeds. Aquaculture 2023, 567, 739256.
  12. Abdel-Ghany, H.M.; Salem, M.E.-S. Effects of Dietary Chitosan Supplementation on Farmed Fish: A Review. Rev. Aquac. 2020, 12, 438–452.
  13. Jiménez-Gómez, C.P.; Cecilia, J.A. Chitosan: A Natural Biopolymer with a Wide and Varied Range of Applications. Molecules 2020, 25, 3981.
  14. Alishahi, A.; Aïder, M. Applications of Chitosan in the Seafood Industry and Aquaculture: A Review. Food Bioprocess. Technol. 2012, 5, 817–830.
  15. Ringø, E.; Zhou, Z.; Olsen, R.E.; Song, S.K. Use of Chitin and Krill in Aquaculture—The Effect on Gut Microbiota and the Immune System: A Review. Aquac. Nutr. 2012, 18, 117–131.
  16. Ahmed, F.; Soliman, F.M.; Adly, M.A.; Soliman, H.A.M.; El-Matbouli, M.; Saleh, M. Dietary Chitosan Nanoparticles: Potential Role in Modulation of Rainbow Trout (Oncorhynchus mykiss) Antibacterial Defense and Intestinal Immunity against Enteric Redmouth Disease. Mar. Drugs 2021, 19, 72.
  17. Dawood, M.A.O.; Gewaily, M.S.; Soliman, A.A.; Shukry, M.; Amer, A.A.; Younis, E.M.; Abdel-Warith, A.-W.A.; Van Doan, H.; Saad, A.H.; Aboubakr, M.; et al. Marine-Derived Chitosan Nanoparticles Improved the Intestinal Histo-Morphometrical Features in Association with the Health and Immune Response of Grey Mullet (Liza ramada). Mar. Drugs 2020, 18, 611.
  18. Kono, M.; Matsui, T.; Shimizu, C. Effect of Chitin, Chitosan, and Cellulose as Diet Supplements on the Growth of Cultured Fish. Nippon. Suisan Gakkaishi 1987, 53, 125–129.
  19. Qin, C.; Zhang, Y.; Liu, W.; Xu, L.; Yang, Y.; Zhou, Z. Effects of Chito-Oligosaccharides Supplementation on Growth Performance, Intestinal Cytokine Expression, Autochthonous Gut Bacteria and Disease Resistance in Hybrid Tilapia Oreochromis niloticus ♀ × Oreochromis aureus ♂. Fish Shellfish Immunol. 2014, 40, 267–274.
  20. Elserafy, S.S.; Abdel-Hameid, N.-A.H.; Abdel-Salam, H.A.; Dakrouni, A.M. Effect of Shrimp Waste Extracted Chitin on Growth and Some Biochemical Parameters of the Nile Tilapia. Egypt. J. Aquat. Biol. Fish 2021, 25, 313–329.
  21. Shi, F.; Qiu, X.; Nie, L.; Hu, L.; Babu, V.S.; Lin, Q.; Zhang, Y.; Chen, L.; Li, J.; Lin, L.; et al. Effects of Oligochitosan on the Growth, Immune Responses and Gut Microbes of Tilapia (Oreochromis niloticus). Fish Shellfish Immunol. 2020, 106, 563–573.
  22. Gasco, L.; Biasato, I.; Dabbou, S.; Schiavone, A.; Gai, F. Animals Fed Insect-Based Diets: State-of-the-Art on Digestibility, Performance and Product Quality. Animals 2019, 9, 170.
  23. Danulat, E. The Effects of Various Diets on Chitinase and SS-Glucosidase Activities and the Condition of Cod, Gadus morhua (L.). J. Fish Biol. 1986, 28, 191–197.
  24. Fines, B.C.; Holt, G.J. Chitinase and Apparent Digestibility of Chitin in the Digestive Tract of Juvenile Cobia, Rachycentron canadum. Aquaculture 2010, 303, 34–39.
  25. 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.
  26. Jozefiak, A.; Engberg, R. Insect Proteins as a Potential Source of Antimicrobial Peptides in Livestock Production. A Review. J. Anim. Feed Sci. 2017, 26, 69998.
  27. Bruni, L.; Belghit, I.; Lock, E.-J.; Secci, G.; Taiti, C.; Parisi, G. Total Replacement of Dietary Fish Meal with Black Soldier Fly (Hermetia illucens) Larvae Does Not Impair Physical, Chemical or Volatile Composition of Farmed Atlantic Salmon (Salmo salar L.). J. Sci. Food Agric. 2020, 100, 1038–1047.
  28. Foysal, M.J.; Gupta, S.K. A Systematic Meta-Analysis Reveals Enrichment of Actinobacteria and Firmicutes in the Fish Gut in Response to Black Soldier Fly (Hermetica illucens) Meal-Based Diets. Aquaculture 2022, 549, 737760.
  29. Gaudioso, G.; Marzorati, G.; Faccenda, F.; Weil, T.; Lunelli, F.; Cardinaletti, G.; Marino, G.; Olivotto, I.; Parisi, G.; Tibaldi, E.; et al. Processed Animal Proteins from Insect and Poultry By-Products in a Fish Meal-Free Diet for Rainbow Trout: Impact on Intestinal Microbiota and Inflammatory Markers. Int. J. Mol. Sci. 2021, 22, 5454.
  30. Huyben, D.; Vidaković, A.; Werner Hallgren, S.; Langeland, M. High-Throughput Sequencing of Gut Microbiota in Rainbow Trout (Oncorhynchus mykiss) Fed Larval and Pre-Pupae Stages of Black Soldier Fly (Hermetia illucens). Aquaculture 2019, 500, 485–491.
  31. Karlsen, Ø.; Amlund, H.; Berg, A.; Olsen, R.E. The Effect of Dietary Chitin on Growth and Nutrient Digestibility in Farmed Atlantic Cod, Atlantic Salmon and Atlantic Halibut. Aquac. Res. 2017, 48, 123–133.
  32. Renna, M.; Schiavone, A.; Gai, F.; Dabbou, S.; Lussiana, C.; Malfatto, V.; Prearo, M.; Capucchio, M.T.; Biasato, I.; Biasibetti, E.; et al. Evaluation of the Suitability of a Partially Defatted Black Soldier Fly (Hermetia illucens L.) Larvae Meal as Ingredient for Rainbow Trout (Oncorhynchus mykiss Walbaum) Diets. J. Anim. Sci. Biotechnol. 2017, 8, 57.
  33. Maulu, S.; Langi, S.; Hasimuna, O.J.; Missinhoun, D.; Munganga, B.P.; Hampuwo, B.M.; Gabriel, N.N.; Elsabagh, M.; Doan, H.V.; Kari, Z.A.; et al. Recent Advances in the Utilization of Insects as an Ingredient in Aquafeeds: A Review. Anim. Nutr. 2022, 11, 334.
  34. Rangel, F.; Monteiro, M.; Santos, R.A.; Ferreira-Martins, D.; Cortinhas, R.; Gasco, L.; Gai, F.; Pousão-Ferreira, P.; Couto, A.; Oliva-Teles, A.; et al. Novel Chitinolytic Bacillus spp. Increase Feed Efficiency, Feed Digestibility, and Survivability to Vibrio anguillarum in European Seabass Fed with Diets Containing Hermetia illucens Larvae Meal. Aquaculture 2024, 579, 740258.
  35. Gao, C.; Cai, X.; Zhang, Y.; Su, B.; Song, H.; Wenqi, W.; Li, C. Characterization and Expression Analysis of Chitinase Genes (CHIT1, CHIT2 and CHIT3) in Turbot (Scophthalmus maximus L.) Following Bacterial Challenge. Fish Shellfish Immunol. 2017, 64, 357–366.
  36. Ikeda, M.; Kakizaki, H.; Matsumiya, M. Biochemistry of Fish Stomach Chitinase. Int. J. Biol. Macromol. 2017, 104, 1672–1681.
  37. 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, 05, 398.
  38. Pohls, P.; González-Dávalos, L.; Mora, O.; Shimada, A.; Varela-Echavarria, A.; Toledo-Cuevas, E.M.; Martínez-Palacios, C.A. A Complete Chitinolytic System in the Atherinopsid Pike Silverside Chirostoma Estor: Gene Expression and Activities. J. Fish Biol. 2016, 88, 2130–2143.
  39. Lindsay, G.J.H.; Walton, M.J.; Adron, J.W.; Fletcher, T.C.; Cho, C.Y.; Cowey, C.B. The Growth of Rainbow Trout (Salmo gairdneri) given Diets Containing Chitin and Its Relationship to Chitinolytic Enzymes and Chitin Digestibility. Aquaculture 1984, 37, 315–334.
  40. LeCleir, G.R.; Buchan, A.; Hollibaugh, J.T. Chitinase Gene Sequences Retrieved from Diverse Aquatic Habitats Reveal Environment-Specific Distributions. Appl. Environ. Microbiol. 2004, 70, 6977–6983.
  41. Hamid, R.; Khan, M.A.; Ahmad, M.; Ahmad, M.M.; Abdin, M.Z.; Musarrat, J.; Javed, S. Chitinases: An Update. J. Pharm. Bioallied Sci. 2013, 5, 21–29.
  42. Thakur, D.; Bairwa, A.; Dipta, B.; Jhilta, P.; Chauhan, A. An Overview of Fungal Chitinases and Their Potential Applications. Protoplasma 2023, 260, 1031–1046.
  43. Rangel, F.; Santos, R.A.; Monteiro, M.; Lavrador, A.S.; Gasco, L.; Gai, F.; Oliva-Teles, A.; Enes, P.; Serra, C.R. Isolation of Chitinolytic Bacteria from European Sea Bass Gut Microbiota Fed Diets with Distinct Insect Meals. Biology 2022, 11, 964.
  44. Subramanian, K.; Balaraman, D.; Panangal, M.; Nageswara Rao, T.; Perumal, E.; Kumarappan, R.A.A.; Sampath Renuga, P.; Arumugam, S.; Thirunavukkarasu, R.; Aruni, W.; et al. Bioconversion of Chitin Waste through Stenotrophomonas maltophilia for Production of Chitin Derivatives as a Seabass Enrichment Diet. Sci. Rep. 2022, 12, 4792.
  45. Agbohessou, P.S.; Mandiki, S.N.M.; Mbondo Biyong, S.R.; Cornet, V.; Nguyen, T.M.; Lambert, J.; Jauniaux, T.; Lalèyè, P.A.; Kestemont, P. Intestinal Histopathology and Immune Responses Following Escherichia coli Lipopolysaccharide Challenge in Nile Tilapia Fed Enriched Black Soldier Fly Larval (BSF) Meal Supplemented with Chitinase. Fish Shellfish Immunol. 2022, 128, 620–633.
  46. Liu, Y.; Zhou, Z.; Miao, W.; Zhang, Y.; Cao, Y.; He, S.; Bai, D.; Yao, B. A Chitinase from Aeromonas veronii CD3 with the Potential to Control Myxozoan Disease. PLoS ONE 2011, 6, e29091.
  47. Zhang, Y.; Zhou, Z.; Liu, Y.; Cao, Y.; He, S.; Huo, F.; Qin, C.; Yao, B.; Ringø, E. High-Yield Production of a Chitinase from Aeromonas veronii B565 as a Potential Feed Supplement for Warm-Water Aquaculture. Appl. Microbiol. Biotechnol. 2014, 98, 1651–1662.
  48. Mohan, K.; Ravichandran, S.; Muralisankar, T.; Uthayakumar, V.; Chandirasekar, R.; Seedevi, P.; Rajan, D.K. Potential Uses of Fungal Polysaccharides as Immunostimulants in Fish and Shrimp Aquaculture: A Review. Aquaculture 2019, 500, 250–263.
  49. Poria, V.; Rana, A.; Kumari, A.; Grewal, J.; Pranaw, K.; Singh, S. Current Perspectives on Chitinolytic Enzymes and Their Agro-Industrial Applications. Biology 2021, 10, 1319.
  50. Rathore, A.S.; Gupta, R.D. Chitinases from Bacteria to Human: Properties, Applications, and Future Perspectives. Enzyme Res. 2015, 2015, 791907.
  51. Swiontek Brzezinska, M.; Jankiewicz, U.; Burkowska, A.; Walczak, M. Chitinolytic Microorganisms and Their Possible Application in Environmental Protection. Curr. Microbiol. 2014, 68, 71–81.
  52. Gomaa, E.Z. Microbial Chitinases: Properties, Enhancement and Potential Applications. Protoplasma 2021, 258, 695–710.
  53. Eggink, K.M.; Pedersen, P.B.; Lund, I.; Dalsgaard, J. Chitin Digestibility and Intestinal Exochitinase Activity in Nile Tilapia and Rainbow Trout Fed Different Black Soldier Fly Larvae Meal Size Fractions. Aquac. Res. 2022, 53, 5536–5546.
Subjects: Fisheries
Contributors MDPI registered users' name will be linked to their SciProfiles pages. To register with us, please refer to : , , , , ,
View Times: 165
Revisions: 2 times (View History)
Update Date: 19 Dec 2023
Video Production Service