The optimum pHs for amylase and alkaline protease activities were 7. A pepsin-like enzyme was detected in all three carnivorous fishes Ompok bimaculatus , Kryptopterus geminus , and Hemibagrus spilopterus with optimum reaction pH of 2.

In optimum reaction conditions, the amylase and alkaline protease from Puntioplites proctozyron showed the highest activities. Lower activities of all enzymes were observed at temperature 29 °C of Lam Nam Choen swamp than at the optimum reaction temperatures.

The fish species contained one to three and five to eight isoforms of amylase and alkaline protease, respectively, with molecular weights from Both the alkaline proteases and amylases were stable in wide pH and temperature ranges. Amylase and protease are important enzymes in cellular metabolism in plants, animals, and microorganisms.

Both enzymes function to catalyze degradation of macromolecules into small building blocks, which are subsequently used to produce energy or to synthesize other biomolecules inside cells. Extracellular digestive enzymes present in the digestive tract of animals or in the digestive system of carnivorous or insectivorous plants 1 , 2 also have a crucial role in digestion of macromolecules from food.

Proteases catalyze the breakdown of proteins into small peptides and amino acids. They are found in all living organisms. Based on the optimum pH for enzymatic activity, these enzymes are classified into two groups—acidic and alkaline proteases.

The former enzymes, such as pepsin, function in acidic conditions with optimum pH around 2. Alkaline proteases are present in the small intestine of humans and the digestive system of animals.

This group includes chymotrypsin and trypsin, which have optimum reaction pH values ranging from 8. Amylases catalyze the hydrolysis of polysaccharides such as starch and glycogen into short-chain sugars.

These enzymes are ubiquitous and play an important metabolic role. According to catalytic function and enzyme structure, amylases are classified into three types: α-amylase EC 3. α-Amylase is a metalloenzyme requiring calcium ions for catalytic function.

This enzyme randomly hydrolyzes α-1,4-glycosidic bonds, releasing maltose and glucose 3. β-Amylase is a calcium-independent enzyme that catalyzes the hydrolysis of α-1,4-glycosidic linkages at the nonreducing end of starch molecules, yielding a maltose unit.

γ-Amylase catalyzes the hydrolysis of the last α-1,4-glycosidic bond at the nonreducing end of starch yielding one glucose unit; γ-amylase is mostly active in acidic conditions.

Due to the rapid growth of microorganisms, which enables high-yield enzyme production, microbial enzymes are widely used in various industries such as the manufacture of detergent, paper and pulp, textiles, and food and animal feed 4.

Nevertheless, there are drawbacks of some bacterial enzymes, such as low stability at high temperature except for thermophilic bacteria or a narrow pH range for activity. Therefore, novel enzymes with high activity and high stability at broad ranges of pH and temperature are still required for a wide range of uses.

Studies of protease and amylase have been reported in plants 5 , 6 , 7 , insects 8 , 9 , microorganisms 10 , 11 and fish 12 , 13 , 14 , 15 , 16 , 17 , The data report various enzyme activities, biochemical properties, and molecular weights of amylase and protease.

The optimum pH values for catalytic activity of acidic proteases, alkaline protease, and amylase are in the range 2.

Molecular weights range from 18 to kDa and 18 to The number of isoenzymes the enzymes differ in amino acid sequences and molecular sizes but catalyze the same reaction varies from one to eight, dependent on the species from which the enzymes are derived.

Noticeably, almost all bacteria and plants contain only one isoform of amylase and protease, but fishes and shrimps contain many isoforms. In addition, different species have isoenzymes of different molecular weight 16 , However, almost all recently available data on fish digestive enzymes are from omnivorous fish.

Little information has been reported about these enzymes in carnivorous or herbivorous fish. Cyprinidae, Siluridae, Bagridae, and Cichlidae are fish families widely distributed in North America, Africa, and Southeast Asia All six species are important for food, and economically. The types and number of fish species in a locality depend on various factors such as the abundance of the ecosystem and the local management of water resources.

The growth rate of each fish species affects the abundance and fish diversity. Little is known on the digestive systems of freshwater fishes, therefore this study aimed to investigate the biochemical properties of digestive enzymes—amylase and protease—from six freshwater fish species.

The enzyme activities in optimum and environmental conditions were determined and compared. Moreover, the isoenzyme patterns and their molecular weights, as well as the enzyme stabilities, were analyzed. This study will help understanding of the digestive enzymes of each fish species, not only for sustainable fish management, but also for improvement of fish feed formula, in order to increase the growth rate of fish and reduce feed cost and cultivation time in aquaculture.

Carnivorous fishes Om. bimaculatus , K. geminus , H. spilopterus ; herbivorous fishes Puntius gonionotus and Puntioplites proctozysron ; and omnivorous fish Or. niloticus were selected for this study. Details of body length, body weight, digestive tract weight, and calculated digestive somatic index DSI values are indicated in Table 1.

A high DSI value indicates high digestive capability. gonionotus , Or. niloticus , and P. proctozysron exhibited high digestive enzyme activity; the two former species displayed a high DSI value, but the latter did not. Time course assays showed a linear relationship between enzyme activity and incubation time when using 20 or 25 μl of crude enzyme preparation containing mixtures of isoenzymes extracted from the digestive tract of each fish species for the detection of amylase or protease activity data not shown.

Therefore, an incubation time between 5 and 20 min was chosen for testing enzyme activity. The enzymes from the different fish species showed various optimum pHs and temperatures for enzymatic activity. Almost all of the amylases showed the same optimum pH for activity, pH 8. geminus amylase, for which the optimum pH was 7.

The maximum alkaline protease activity was observed at different pH values for the enzymes from different fishes Fig. The optimum temperatures for enzyme activity in this study were in the range 45—60 °C, 50—55 °C, and 50—55 °C for amylase, alkaline protease, and pepsin-like enzyme, respectively Fig.

No correlation between fish families and optimum temperature for enzyme activity was observed. Interestingly, we found thermostable enzymes in P. gonionotus and P. This result suggests that these enzymes are potential candidates for use in broad applications. Study of enzyme activity at different temperatures helps us to understand food digestion capacity of fishes.

Fish are cold-blooded—their temperature depends on the water temperature. However, determination of digestive enzyme activity at the water temperature of the swamp, pond, or aquarium has rarely been reported.

In this study, digestive enzyme activities determined in optimum and environmental conditions were compared among six fish species. niloticus , Puntius gonionotus , and H.

Low activities of both enzymes were observed in K. geminus and Om. bimaculatus [specific activities less than 1. Pepsin-like enzymes were apparent in the carnivorous fishes H. spilopterus , K. The digestive enzyme activities were also investigated at swamp temperature 29 °C , to evaluate food digestion capability.

Moreover, we found that the carnivorous fishes displayed lower amylase and alkaline protease activities than the omnivorous and herbivorous fishes in both temperature conditions. Amylases, alkaline proteases, and pepsin-like enzyme were separated by SDS-PAGE and the molecular weights of these enzymes were analyzed.

In this zymographic analysis, clear bands on the dark background of the stained gel indicate the position of an enzyme that can hydrolyze substrate. The number of isoforms of alkaline protease ranged from five to eight in the six fish species Fig. Zymographic results corresponded to enzyme activity analysis data; enzymes with high activity exhibited strong, clear bands, whereas enzymes with low activity showed weak bands on the gels.

In the case of pepsin-like enzyme, no clear enzyme bands were observed with μg of total protein for samples from Om. bimaculatus or K. One clear band was detected for H. spilopterus pepsin-like enzyme, with molecular weight 24 kDa data not shown.

SDS gels stained with Coomassie brilliant blue R and zymograms of amylase a , alkaline proteases b Lane M1; Pink plus prestained protein ladder GeneDireX , lane M2; unstained protein molecular weight marker Thermo Scientific , lanes 1—3; Ompok bimaculatus n6, n7, n8 , lanes 4—6; Kryptopterus geminus n6, n7, n8 , lanes 7—9; Hemibagrus spilopterus n1, n2, n4 , lanes 10—12; Puntius gonionotus n5, n7, n8 , lanes 13—15; Oreochromis niloticus n5, n7, n8 , lanes 16—18; Puntioplites proctozysron n1, n2, n3.

The effects of incubation time, pH, and temperature on enzyme activity were determined using crude enzymes samples to help understand more biochemical properties of the enzymes that will be useful for further study and application.

Enzymes from three fish species P. gonionotus , P. proctozysron , and Or. niloticus exhibiting high activities of both amylase and alkaline protease in broad pH and temperature ranges were used for this experiment.

Figure 4 a shows the enzyme activities at different times of incubation in 0. All three species exhibited the same pattern of enzyme stability. Dashed lines represent the relative activity of amylase, whereas solid lines represent the relative activity of alkaline proteases. Considering the effect of pH, most of the enzymes from the tested fish species revealed high stability in a broad pH range from 5.

However, in the case of P. At pH 2—5, acid-induced denaturation of protein often occurs. Decreasing pH by adding acids buffer pH 5. The P. gonionotus amylase seems not able to refold after adding of an optimum buffer for activity assay.

Figure 4 c reveals the effect of temperature after incubation of enzyme for 1 h at 25—60 °C. gonionotus amylase.

This research studied the biochemical properties of digestive enzymes from some freshwater fishes. The enzymes from the different fish species had various optimum pHs and temperatures for enzymatic activity.

These data correlated with results from seven cyprinid fishes 16 and Or. niloticus These results suggest that the enzymes may share conserved amino acid residues at the active site 22 , In other organisms such as bacteria Bacillus cereus , Bacillus amyloliquefaciens BH, and Aeribacillus pallidus C10 , shrimp Cherax albidus , Penaeus californiensis Holmes , Penaeus vannamei Boone , and Penaeus stylirostris Stimpson , and other fish species Osteochilus hasselti , Labiobarbus spilopleura , Osteochilus lini , Cyclocheilichthys repasson , Cyclocheilichthys apogon , Puntius brevis, Symphysodon aequifasciata , Polyodon spathula, Limia vittata , and Gambusia punctata , the amylases exhibit different optimum pH values, 6.

In the present study, different optimum pH values were observed for alkaline protease, with the values varying from 8. Moreover, acidic proteases were also observed. Low activity of this pepsin-like enzyme was observed in H. spilopterus , Om.

bimaculatus , and K. However, the maximum activity of the enzyme was detected at pH 2. Fish are cold-blooded; thus, the water temperature influences their body temperature, growth rate, and food consumption 29 , 30 , The growth rate is associated with digestive capability, which is derived from digestive enzyme activity.

Study of the digestive enzyme activity at different temperatures will help with sustainable development of aquaculture. In addition, the carnivorous fishes displayed lower amylase and alkaline protease activities than the omnivorous and herbivorous fishes.

Isoenzyme analysis and molecular weight determination of amylase and protease enzymes have been performed in bacteria, shrimp, and some species of fishes. Noticeably, bacterial enzymes display only one isoform with molecular weight ranging from 28 to 68 kDa, whereas fish and shrimp contain many isoforms 10 , 12 , 13 , 16 , 19 , 21 , 24 , 25 , 26 , 27 , The several isoforms of digestive enzymes seen in digestive tracts of fish may derive from the floral bacteria However, more experiments have to further investigate bacterial enzymes such as the expression level, biochemical properties, and molecular weights compared to enzyme extracts from digestive tracts of fish.

In the present study, the number of isoenzymes of proteolytic and amylolytic enzymes ranged from one to eight and two to eight with molecular weights between 12 and kDa and 26 and kDa, respectively.

spilopterus could be detected on a zymographic gel, and the band was very faint. pH and temperature stabilities have been reported for bacterial α-amylases 10 , 11 , 33 , 34 and bacterial proteases 33 , Amylases from Bacillus showed high stability at pHs between 5.

For marine fish, the digestive enzymes from Atlantic salmon Salmo salar 36 and thornback ray Raja clavata 37 have been reported. However, little information has been reported on stability of digestive enzymes from freshwater fish.

Seven species of fishes including of P. gonionotus have been reported the biochemical properties of digestive enzyme However, molecular weight and enzyme stabilities have not been determined. Chaijareon and Thongruang determined the stability of amylase and protease from Or. In contrast, our experiments showed that both amylase and alkaline protease from Or.

niloticus and also those from P. proctozysron and P. gonionotus exhibited high stability at a wide range of pH 6. Moreover, the enzymes were stable on incubation in buffer pH 8.

High stability of amylase and alkaline protease, digestive enzymes from freshwater fishes, was found in this study and is good evidence for further study and application.

Casein sodium salt, from bovine milk, cat. C and starch from rice, cat. S were purchased from Sigma-Aldrich, USA. Acrylamide cat. Sodium dodecylsulfate cat. DB , soluble starch cat. SB and β-mercaptoethanol cat.

MB were from Bio Basic Canada Inc. Chemical reagents for preparation of buffers were purchased from Carlo Erba France , Scharlau Barcelona, Spain , and Ajax Finechem MA, USA.

Samples were collected as part of a fish faunal survey in Lam Nam Choen swamp, Nongruea district, Khon Kaen province, Thailand, by using nets. Fish were killed in ice without using any chemical agent.

The condition did not severely distress or cause lasting harm to sentient fish. The body weight and body length were recorded. The digestive tracts were subsequently rinsed with distilled water and then homogenized on ice by uses of pestle and mortar with a buffer 0.

Determination of protein content was performed by using Bradford reagent Bovine serum albumin was used for the calibration curve of protein concentration. All methods were carried out in accordance with relevant guidelines and regulations. All experimental protocols and the care and use of experimental animals complied with animal welfare laws of Thailand, and guidelines and policies approved by ThaiIACUC permit reference number U, To determine the initial velocity of enzymatic reaction, the enzyme activity was observed after incubation of enzyme with substrate for 5 to 30 min.

One reaction for one time point of amylase assay contained 20 μl of supernatant containing digestive enzymes or crude enzyme extract, μl of 0.

The reactions were incubated at room temperature 23 °C for 5, 10, 15, 20, 25, or 30 min. Supernatants containing the catalytic products of amylase and protease were taken to measure absorbance at and nm, respectively. The absorbance values were plotted against the incubation time.

The activities of amylase and protease were observed at pH 5—11 and pH 1—11, respectively. Buffers used 0. One milliliter of reaction for amylase assay contained 20 μl of crude enzyme extract, μl of 0.

Absorbance of the supernatants was measured at nm. The reactions were incubated on ice for 30 min then centrifuged to separate denatured casein and enzyme. Absorbance of the resulting supernatants was measured at nm.

Amylase and protease reactions were monitored at 29, 35, 40, 45, 50, 55, 60, and 70 °C. The reactions were incubated for 7 min at various temperatures listed above. The supernatants were measured at and nm to determine the catalytic products of amylase and protease, respectively.

Five to eight specimens of each fish species were determined for both amylase and protease activities. One milliliter of amylase reaction contained 20 μl of crude enzyme extract, μl of 0. The reactions were incubated for 10 min at the optimum temperature for each enzyme.

The resulting supernatants were taken to measure the absorbance at and nm, respectively. The contents of product released from substrate were calculated from tyrosine and maltose standard curves, respectively.

The enzyme activity and specific activity were subsequently analyzed and compared between fish species. One unit of protease activity was defined as the amount of enzyme required to generate one nmol of tyrosine equivalents in 1 min; one unit of amylase activity was defined as the amount of enzyme required to produce one mmol of maltose in 1 min.

Specific activity is the enzyme activity nmol tyrosine equivalents min -1 for proteases or mmol maltose min -1 for amylase per mg of total protein and therefore it was reported as units mg -1 protein.

The activities of both digestive enzymes were also determined at swamp temperature; the reaction components and procedure were similar to reactions in optimum conditions, but the assay temperature was changed to 29 °C for all samples. Control reactions were also conducted for all samples.

The obtained absorbance values were subtracted from those for catalytic reactions. Data were analyzed by one-way analysis of variance IBM SPSS statistics software v. Total crude enzymes 25 and μg respectively were used to analyze amylase and protease isoenzymes, by using sodium dodecyl sulfate SDS -polyacrylamide gel electrophoresis and zymography.

Crude enzyme samples were mixed with SDS sample buffer without any reducing agent Electrophoresis was performed at V and 4 °C using Mini-PROTEAN Tetra Cell apparatus Bio-Rad.

After electrophoresis, two gels with a similar pattern of sample loading were separately examined. The first gel was stained with Coomassie Brilliant Blue [0. The protein bands were dark blue. For in-gel analysis of amylase activity, the second gel was soaked for 20 min in 0.

The enzyme was allowed to react in-gel for 1 h. The starch solution was discarded and the gel was rinsed with water. Iodine solution was used to stain the gel. The presence of starch produced a brownish—purple color on the gel. A clear band thus showed the position of amylase i.

Protein markers with different molecular weights were used in this study. M1 was Pink Plus-prestained protein ladder from GeneDirex and contained 11 proteins in the range 10— kDa. The manufacturer does not provide any detail about the specific proteins in this marker mixture.

M2 was unstained protein molecular weight marker from Thermo Scientific, containing chicken egg white lysozyme coli β-galactosidase To detect protease activity in-gel, the procedure was similar to that described above for amylase.

SDS was removed by using 0. Gels were then incubated with gentle shaking at 45 °C for 1 h. Casein solution was removed and the gels were rinsed with water before staining with Coomassie Brilliant Blue R Protein staining was performed at room temperature for 1 to 2 h.

Destaining was carried out for 30 min or until a clear band was visualized against the dark blue background. The molecular weight of proteases was analyzed as described above for amylases.

Enzyme activity was determined after incubation of crude enzyme in 0. At each time point, enzyme mixture was aliquoted and substrate solution was added 0. The reaction was incubated at room temperature 23 °C.

The enzymatic product was detected by measuring the absorbance as described above. The effect of pH on enzyme activity was investigated. Crude enzyme was incubated for 1 h at room temperature 23 °C in buffer at pH 5— Enzyme solution was aliquoted and mixed with the optimum buffer and substrate for activity testing.

To determine the effect of temperature, crude enzyme mixed with 0. Substrate and the optimum buffer for reaction were mixed with the enzyme solution before assay at the optimum reaction temperature. Tokes, Z. Digestive enzymes secreted by the carnivorous plant Nepenthes macferlanei L.. Planta Ber.

However, studies have had contradicting results. Digestive enzymes are substances that help break down macronutrients into smaller compounds to promote their absorption. Some test-tube and animal studies show that they could improve the health of your gut microbiome , which may affect weight control.

On the other hand, digestive enzyme inhibitors have been shown to reduce food intake and increase weight loss and fat loss. While digestive enzyme supplements may or may not directly boost weight loss, they could promote healthy digestion and regularity, especially for those with certain gastrointestinal conditions.

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Here's why. A Quiz for Teens Are You a Workaholic? How Well Do You Sleep? Health Conditions Discover Plan Connect. Nutrition Evidence Based Do Digestive Enzymes Promote Weight Loss? Medically reviewed by Grant Tinsley, Ph. What they are Gut bacteria Lipase Best types Weight loss Bottom line Digestive enzymes are often used to support healthy digestion and increase nutrient absorption.

Share on Pinterest. What are digestive enzymes? May affect gut bacteria. Effects of lipase. Best types. Enzyme inhibitors could support weight loss. The bottom line. Share this article. Read this next. Understanding Digestive Enzymes: Why Are They Important?

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In the stomach, pepsin is the main digestive enzyme attacking proteins. Several other pancreatic enzymes go to work when protein molecules reach the small intestine.

Lipase is produced in the pancreas and small intestine. A type of lipase is also found in breast milk to help a baby more easily digest fat molecules when nursing. Lipids play many roles, including long-term energy storage and supporting cellular health.

Enzymes, and especially digestive enzymes, can be sensitive to changes in the body. Some of these changes can create an environment that makes it difficult for enzymes to work properly. Certain health conditions that negatively impact your pancreas can reduce the number and effectiveness of digestive enzymes.

Some of these conditions include:. These conditions can lead to pancreatic exocrine insufficiency PEI , which is a chronic condition that can affect nutrient absorption.

Inhibitors can occur naturally. They can also be manufactured and produced as medications. Antibiotics are a good example. They inhibit or prevent certain enzymes from helping bacterial infections spread.

Eating highly processed or high-calorie foods, drinking a lot of alcohol, living a sedentary lifestyle, and not getting proper nutrients can all have a negative impact on your pancreas, and therefore, a negative impact on the enzymes it produces.

Digestive enzyme supplements are available over-the-counter, and may also be prescribed for specific conditions that can create enzyme insufficiency. Conditions that affect your pancreas, such as pancreatitis, cystic fibrosis , or pancreatic cancer , can all reduce the number of important enzymes your body produces.

As a result, you may not get enough enzymes to thoroughly digest your food and obtain the nutritional value from what you eat. If you have these conditions — or others in which your enzyme levels are below a normal or healthy range — talk with your doctor about treatment options.

For example, individuals living with cystic fibrosis may have to take enzymes with every meal. The only FDA-regulated enzyme replacement therapy is pancreatic enzyme replacement therapy PERT. Many digestive enzymes are sold over-the-counter OTC to help people treat various digestive issues on their own, such as :.

Lifestyle changes, including focusing on your diet and physical activity level, are typically the best bet for improving digestive enzyme function. Enzymes create chemical reactions in the body, and are crucial for a variety of processes, including digestion.

Digestive enzymes are mostly produced in the pancreas, and help your body break down foods and extract nutrients. For individuals living with a health condition that may cause pancreatic exocrine insufficiency, such as cystic fibrosis, pancreatic cancer, or type 1 diabetes, digestive enzyme supplementation may be necessary.

Your doctor will help you decide if enzyme supplementation is right for you. If you are having recurring digestive issues, talk to a doctor. There could be an underlying cause that needs more than digestive enzyme treatment. Our experts continually monitor the health and wellness space, and we update our articles when new information becomes available.

For example, in green catfish Mystus nemurus larvae, lipase activity showed a basal activity level from day 1 to day 25, and subsequently an increase in lipase activity between 30—40 dph Srichanun et al. In leopard groupers Mycteroperca rosacea larvae, lipase activity was detected before mouth opening and during feeding, with a significant peak at 30 dph Martínez-Lagos et al.

In Chinese perch Siniperca chuatsi larvae, a low lipase activity was detected before the mouth opened. In contrast to other carnivorous fish, lipase activity did not increase suddenly after the first feeding, but gradually increased until 15 dph, then suddenly decreased at 25 dph, and continued declining until 30 dph Tang et al.

Lipids are common nutrient substrates for farmed fish, and lipases are secreted in the intestine for their digestion. Subsequently, fish lipases can be used as enzymatic reagents to determine the in vitro digestibility of food oils according to the methodology of Espinosa-Chaurand and Nolasco-Soria, to select the most digestible ones and allow the possible formulation of alternative diets for aquaculture.

According to Yúfera et al. Emulsions of water-insoluble short-chain triacylglycerols e. The methods used by the authors differ in the type of substrate used.

Therefore, the hydrolysis products generated what is measured are different, making it necessary to use the standard curves or the specific molar extinction coefficients MEC to measure the hydrolyzed ester bonds per unit of time.

If the measurements are made at different pHs, temperatures, ion concentrations, or types or concentrations of bile salt, this will directly affect the lipase activity. If the definitions of the lipase unit differ, the comparison of results becomes more complex Yúfera et al.

After removing the studies that did not focus on the lipase of aquaculture fish, a final database of articles was used for their review Supplementary Material 1 , and additional references used by authors were reviewed. Lipase activity has been studied in at least 95 fish genera species and 10 fish hybrids Table S1 , Supplementary Material 2 , with the most studied genera being Oreochromis , Cyprinus , Oncorhynchus , Ctenopharyngodon , and Carassius The results indicated Table S2 , Supplementary Material 2 that the studied tissues belonged to either intestine [whole, anterior, medium, posterior This system generally comprises the mouth—esophagus, stomach or pseudostomach , and intestine.

The pancreas or hepatopancreas is connected to the first portion of the intestine. Likewise, the pyloric cecum in fish that have it is identified as a cluster of lobes. The number of larvae was from 1 to per sample, and the weights of tissue for lipase crude extract preparation were from 40 mg to 30 g.

Unfortunately, In general, the authors have used 1 to 66 volumes of extraction solvent: 1—2 1. In the specific case of larvae, the authors used 5 to 66 volumes of extraction solution.

Regarding the extraction solution, Enzymatic extracts clarification has been performed at relative centrifugal force RCF values that ranged from to 33, × g and centrifugation times that varied from 3 min to 60 min at 4°C. Some authors reported the RCF applied but not the centrifugation time; others reported the rpm applied but did not give the data of the rotor used or their radius r , which makes it impossible to calculate the RCF applied.

Only For enzyme extract preparation, the method proposed by Nolasco-Soria is recommended, also considering the time of sacrifice to avoid the effect of the circadian cycle Yúfera et al. In the case of larvae, the size i. In addition, samples should be kept cold while they are being handled, and long freezing times before their enzymatic analysis should be avoided Solovyev and Gisbert, According to differences in the fish intestinal pH Solovyev and Izvekova, and the working physiological pH of fish lipases pH that gives stability to the enzymes , as previously reported, the recommendation is to perform lipase extraction with a buffer at pH 7.

Table 1 shows the methods used by the authors. The reader who knows the rationale of the methods may recognize the difficulty of comparing lipase values reported in fish studies.

Table 1 Reference protocols type of method and their frequency of use for assessing the activity of lipase in larvae and digestive tract of fish. One of the oldest published procedures for quantifying lipase activity used by some authors 3. The titrimetric protocol described by these authors is as follows:.

These authors also suggested that the term lipase should be reserved for the enzyme capable of hydrolyzing ester bonds on true lipids fats and oils and that esterase be used for the enzyme acting upon other esters Cherry and Crandall, However, this method, indisputably valuable in determining the presence of true lipases, is tedious because samples need to be individually measured in glass flasks.

In addition, this procedure does not permit the miniaturization of volumes to be carried out at a plastic microplate level. Similar methods cited by the authors include Bier [including Furné et al.

A more straightforward, turbidimetric method using an olive oil emulsion was developed by Shihabi and Bishop A modified method using olive oil as a substrate and a cupric acetate pyridine reagent for colorimetric free-fatty-acid determination was used by Mustafa et al.

In contrast, at the microplate level, the method most used by the authors is that of a commercial kit. Unfortunately, for obvious reasons, the actual composition and concentration of the reactants are still being determined, which limits their practical utility for example, if the study requires that the concentrations of substrate or developer be varied.

This also applies to all other commercial kits used by the authors Table 1. The methods used to quantify fish digestive tract lipase were revised in the studies retrieved from the systematic bibliographic search Table 1.

Without considering commercial kits, the method that was most used by the authors is that of Iijima et al. Iijima et al. The original method was explicitly described by Albro et al.

Each assay tube received 5 ml of this mixture and was equilibrated at 37°C. Enzyme solution µl as appropriate was added and the solution was swirled at rpm for from 5 to 15 min at 37°C. In the reaction mixture used by Albro et al. In contrast, Iijima et al. However, they increased the pH of the reaction and used a stopper-extractor to separate the colored phase of the reaction by centrifugation.

The Iijima et al. method was explicitly described as follows:. Typically, μl of enzyme solution was added to the substrate solution. Incubation was carried out for 15 min at 30°C, and the reaction was terminated by adding 0. The reaction mixture was vigorously mixed and centrifuged at 6, × g for 2 min.

The absorbance at nm in the resulting lower aqueous layer 0. The extinction coefficient of p-nitrophenol was 16, M —1 cm —1 per liter at pH 9.

The p -nitrophenyl substrates with short fatty acids, such as acetate, should be avoided due to their non-specific hydrolysis De Caro et al. As reported by Nolasco-Soria et al.

Therefore, these authors are part of the most cited group. Winkler and Stuckmann previously developed a similar method using p -nitrophenyl palmitate as a substrate after it was modified by Markweg et al. Metin and Akpinar Metin and Akpinar, , who cited Winkler and Stuckmann, used p -nitrophenyl acetate as a substrate, as did Bülow and Mosbach The main disadvantage of some of the abovementioned methods is the requirement for organic solvents toxic for users or artificial detergents such as Triton X lipase inhibitor, in accordance with Aryee et al.

Faulk et al. The basis of the above methods is the quantification of the moles of p -nitrophenol released by the lipase hydrolysis of the ester bond the synthetic substrate. The second most used method by the authors is that of Bier Bier, , including Furné et al. According to Bier , the assay protocol is as follows:.

The mixture is shaken gently and incubated for 4 hours at 37° with constant shaking. At the end of the incubation time, 30 ml of a alcohol-acetone solution is added to stop the reaction and break the emulsion.

Phenolphthalein indicator is added, and the solution is titrated with 0. The basis of the method is the quantification of hydroxide moles, which are required to neutralize the protons released by the hydrolysis of ester bonds on olive oil substrate and alkalize the reaction to the change of the phenolphthalein color.

The third most used method by the authors is that of Versaw et al. The original method was explicitly described by Versaw et al. Then 0.

After incubation, the color reaction was produced by the addition of 0. The reaction was stopped with 0. At this juncture in our modification, 2. The addition of the mixed solvent produced a clarified sample which was read for absorbance. Previously, Seligman and Nachlas reported an end point method using β-naphthyl laurate.

Furthermore, more recently, Nolasco-Soria et al. The basis of this method is quantifying the moles of β-naphthol released by the lipase hydrolysis of the ester bond of the synthetic substrate β-naphthyl caprylate.

The incomplete definition of the methods and the use of concatenated citations with other references make it difficult to know how lipase activity was measured. In addition, the variations or adjustments made by different authors to a lipase method cited as a reference make comparison difficult, even when the same method and author are cited, which requires a standardization effort, considering the following.

The volumes among the only 55 studies By type of method, the assay volumes were between µl and 7, µl for colorimetric, between µl and 3, µl for fluorometric, and between 2, µl and 31, µl for titrimetric procedures.

The type of substrate used to measure lipase activity is highly relevant, mainly when synthetic substrates are used with a single associated fatty acid and different chain lengths Nolasco-Soria et al. Thus, regarding lipase substrate, studies Only 67 studies For p -nitrophenyl substrates butyrate, caproate, and myristate , 0.

For 4-methylumbelliferyl substrates butyrate, heptanoate, octanoate, and oleate , 0. For β-naphthyl caprylate, 0. In addition, an excess of substrate in the reaction mixture must be present during the entire reaction time according to the Michaelis—Menten law to determine lipase activity.

Regarding the reaction buffer, different solutions and concentrations were reported Table 4. In particular, most authors who reported this information used a Tris buffer Most buffers have been used correctly depending on their working pH Mohan, , and the experiments in most of the reported studies were performed at an alkaline pH of between 7 and 9.

If the working pH is between 7 and 9, the use of Tris-HCl buffer at a concentration between 20 mM and 50 mM is recommended. Regardless of the use of ions reported explicitly by authors, only 1. As reported by Nolasco-Soria , the requirement for NaCl and divalent ions, such as CaCl 2 , has to be previously determined, in this case for lipase activity in fish.

As is the case for all enzymes, the reaction temperature directly influences the reaction rate of lipases. Therefore, if there are differences in the incubation temperature Table 5 in the methods used by the authors, it is complicated to compare the lipase units reported among the studies, even using the same method.

In addition, it is advisable to measure lipase activity under the physiological or culture temperature conditions of the fish under study. Notably, However, if we consider the temperature used by references cited by the authors, the percentage of studies using temperatures from 35°C to 60°C, which are unusual for fish, rises to Likewise, the optimal temperature determined in the laboratory in vitro tests should not be used to measure lipase activity in fish.

It is recommended to use a temperature of 25°C, which is closer to the ecophysiological temperature, at least for temperate-water fish Gisbert et al. Incubation time varies from 1 min to 60 min for synthetic substrates and 2 h to 24 h for natural substrates oils Table 5.

It is recommended that the incubation time for lipase determination should be from 5 min to 30 min Nolasco-Soria et al.

For end point protocols regarding the method for stopping lipase reaction, the used chemicals are shown in Table 6. The foundation of the stopper is considered adequate. However, those based on acidification acetic acid or alkalinization Na 2 CO 3 , NaOH, and Tris of the medium have their limitations if the enzymes are active at those pH values, particularly if they are alkaline.

It is recommended not to use organic solvents to avoid user exposure. One way to avoid stoppers is to use kinetic methods to measure lipase activity. Only studies The wavelengths used are shown in Table 7.

In the case of colorimetric methods that use p -Nitrophenyl series substrates, the wavelengths reported nm— nm by the authors to measure p -nitrophenol p -NP are close to the maximum absorption peak Iijima et al.

However, to correctly calculate the p -NP moles released by lipase activity, it is necessary to make a p -NP standard curve at the corresponding wavelength and pH.

The same situation occurs in methods using β-naphthyl series substrates, such as β-naphthyl caprylate Versaw et al. In the case of the quantification of MU fluorometric method , the wavelengths reported by the authors are close to those recommended Roberts, for excitation and emission.

Nevertheless, the fluorometric unit calculation requires the construction of a standard curve for MU under the same experimental conditions. Unfortunately, many studies did not provide specific information on the wavelength for measuring absorbance or fluorescence of the assay mixture, so it is assumed that they used the wavelength reported in the reference cited by the authors see Table 7.

Only 3. Another 3. The remaining studies did not report the MEC used, so it is assumed that they used the MEC or calculation procedure reported in the references cited by the authors. The construction method of the standard curve including pH must be reported, in addition to the equation of the curve obtained, or the MEC used, for the calculation of the lipase units.

Finally, only studies Of the studies that used p -nitrophenyl series substrates, only four used absorbance units at nm or nm. Fortunately, 54 studies used unit expressions relative to moles of hydrolyzed substrate or generated product. Regarding the studies that used β-naphthyl series substrates, eight expressed the units in weight, and only two used unit expressions relative to moles of product generated per unit of time.

The 36 studies that used lipid substrates expressed the lipase units in moles of product fatty acids or glycerol generated per unit of time Table 9. The remaining studies The high variability or lack thereof, including the lipase unit definition, makes any comparison difficult. As examples of the variability of the reported results on lipase activity, Table 10 compares lipase units for similar samples and methods.

Table 10 Comparison of lipase in digestive tract and larvae, reported by authors, in fish. According to Brockman , lipases are water-soluble proteins, as ester hydrolases act on non-polar, water-insoluble ester substrates.

According to BRENDA , the lipases are classified with EC 3. Pirahanchi and Sharma report that lipases break triglycerides into free fatty acids and glycerol by hydrolyzing the bonds of the latter.

Lipases are present in pancreatic secretions for the digestion of dietary fats in the intestine. The natural foods preys that fish consume have high lipid contents in oils rich in triacylglycerols.

For example, lipids can account for two-thirds of the total weight of high-latitude zooplankton. In the case of artificial feeds used in aquaculture, fish oil was the most commonly used ingredient Tocher, ; Qiu et al.

The digestive capacity of fish will depend on the lipase units that are synthesized in the pancreas and secreted into the intestine to digest dietary lipids. Considering the lipase definition, the crude extract hydrolytic activity on triglycerides preferably natural ones must be demonstrated, in agreement with Cherry and Crandall The titrimetric methods Cherry and Crandall, or pH stat Kurtovic et al.

Because of that, the use of more practical spectrophotometric colorimetric methods has been preferred, as these require a smaller reaction volume and can measure the hydrolytic activity on the synthetic substrates such as p -NPM or β-NC of several samples simultaneously.

However, as evidenced in this review, the variability of the methods makes comparative studies of lipase activity in fish very difficult to conduct. Given that the methods with p -NPM Iijima et al. However, applying the hydrolysis method of natural oils is also recommended as definitive proof of the presence of lipase activity.

In all cases, adjustments must be proposed for a practical standardized method that allows comparative studies for the same or different species. The result of the review of the articles taken as a sample reports reaction volumes ranging from 70 µL Moro et al.

The proposal is to use practical volumes of µL approximately 0. Due to the insoluble nature of lipase substrates, lipase measurement methods incorporated a detergent into the reaction mixture to facilitate substrate solubility or the formation of micelles to multiply the lipid—water surface or an organic solvent.

However, due to the toxic nature of organic solvents, it is recommended not to use them. Triton-X must also be avoided due to its potential inhibitory action Aryee et al.

Replacing artificial detergents with natural ones, such as bile salts, is recommended. Only 34 of the studies analyzed reported the use of bile salts, including Na cholate Kenari and Naderi, , Na taurocholate Frias-Quintana et al. The concentration of bile salt used and reported by the authors fluctuated between 1.

It is possible that, in the case of the mM concentration, this refers to the concentration of the reagent used and not to the final concentration in the reaction mixture. Only four studies reported using 0.

The working pH for determining lipase activity varied from pH 7 Blanco et al. It is proposed to use an intermediate pH between these values pH 8 to determine lipase activity, which brings the working pH closer to the physiological pH of the fish intestine average pH 7.

Tris-HCl buffer is recommended at the final concentration in the reaction mixture, which was about 20 mM—30 mM. The working temperature for determining lipase activity varied from 4°C Jayant et al.

It is proposed to use a temperature of 25°C, which is closer to the ecophysiological temperature for temperate-water fish Gisbert et al. The working wavelength used to measure reaction mixture absorbance for the lipase determination with p -NP substrates ranged from nm Rueda-Lopez et al.

Although the absorbance difference within the nm to nm range is slight, the recommendation is to measure the absorbance at the wavelength according to the absorption spectra of the final lipase reaction mixture.

In the case of substrates from the β-N group, the wavelengths to measure the absorbance of the reaction mixture were nm Pujante et al. All the authors used the method of Versaw et al. The β-naphthyl caprylate method is recommended for the first analysis of samples with low activity because of its high sensitivity and noticeable purple color.

In the case of fluorometric methods, the wavelength used was nm— nm emission and nm— nm excitation for the MBU substrate. The molar extinction coefficient MEC of p -NP will vary depending on the wavelength used to measure its absorbance.

The few values reported by the authors were 16, at nm Thompson et al. The recommendation is to build a p -NP standard curve pH 8. Something similar occurs with the β-N MEC.

Only two studies present the value of 0. Because the MEC value used by authors is a low number, the expression units should differ from M -1 cm The proposal is that lipase units should be expressed in micromoles of the product p -NP or β-N or fatty acid released per minute.

Each mole would represent the hydrolysis of one mole of ester bonds. The significant differences between the reported lipase units see Table 10 could be due to methodological differences.

The proposal is to standardize the temperature, type of substrate, standard curve or MEC, and calculation formulas. If lipase activity is measured at a mildly acidic pH, extreme care must be taken, as the color generated by either β-N or p -NP is negatively affected color decrease.

In contrast, at an alkaline pH, the color is not significantly affected. This forces, where appropriate, the building of a specific standard curve for the working pH.

The measurement of lipase activity using emulsified NaT-olive oil as a substrate in pH stat is a practical method Nolasco, ; Nolasco-Soria et al. If it is supposed that a pH stat laboratory equipment is unavailable because it is relatively expensive , one option would be to use a high-precision potentiometer with three or four decimal places or a standard potentiometer which is available in most laboratories instead.

The pH of the reaction mixture containing the oil emulsion is adjusted to the working pH for example, 8.

The consumption of NaOH per minute is recorded, and the micromoles per min of NaOH are calculated to have the micromole of ester bonds hydrolyzed per minute.

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