Fish Digestive Capacity: Definition and Methods

Abstract

The nutritional value of a diet and its bioavailability in fish depend on three primary capacities: (a) ingestion, (b) digestion, and (c) absorption. Among these, digestive capacity, defined here as the total enzyme activity available to hydrolyze the bonds of dietary macromolecules to obtain hydrolysis products that are ultimately converted into absorbable micromolecular units, establishes the upper limit for the bioaccessibility of nutrients. To clarify usage and measurement, we conducted a systematic SCOPUS survey (January 2020–June 2024; 62 relevant articles). Most studies either omit a clear definition of digestive capacity or conflate it with digestive organ morphology or isolated enzyme activities. We compared indicators and assay conditions (substrate type, pH, temperature, and expression of units), revealing significant inter-study variability. Based on this synthesis, we propose four operational definitions: (a) Extract Theoretical Volume (ETV)—calculated volume of extract, considering both the solvent volume (SV) used for tissue homogenization and the tissue’s water content; (b) digestive capacity (U)—the total catalytic activity present in the digestive tract at the moment of sampling, where 1 U is the amount of enzyme catalyzing the formation of 1 µmol of product per minute under species-specific physiological pH, ionic strength, and temperature, with the total activity expressed as U fish⁻¹, U organ⁻¹, or U g⁻¹ fish or U g⁻¹ organ, enabling direct comparisons across studies; (c) Digestive Processing (DP)—the total number of bonds hydrolyzed during a given digestion time, whether instantaneous or over a defined period; and (d) Digestive Processing Index (DPI, U-min or U-h), which integrates digestive capacity over time. This framework provides a harmonized checklist for assay standardization and advances comparative studies in fish digestive physiology.

1. Introduction

Nutrient Origin and Early Digestive Ontogeny in Teleosts

Before exogenous feeding, teleost larvae subsist on yolk-sac reserves rich in glycogen, free amino acids, protein, and lipids, which support development until the digestive tract becomes functional [1,2]. In Atlantic halibut (Hippoglossus hippoglossus), Gawlicka et al. followed the ontogeny of key digestive enzymes from 161 to 276 degree-days (dd; water temperature °C × days post-hatch) [3]. Specific activities of lipase, α-amylase, trypsin, and alkaline phosphatase all rose during this interval. Trypsin peaked at 230 dd (16.7 ± 0.7 U mg⁻¹ protein; 0.4 ± 0.0 U mg⁻¹ tissue), whereas lipase (37.3 ± 0.7 U mg⁻¹ protein; 1.7 ± 0.2 U mg⁻¹ tissue) and alkaline phosphatase (111.9 ± 7.2 U mg⁻¹ protein; 5.0 ± 0.5 U mg⁻¹ tissue) reached maxima at 276 dd. α-Amylase activity emerged only between 230 and 276 dd, increasing from 27 ± 6 to 1634 ± 401 U mg⁻¹ protein (0.5 ± 0.1 to 74 ± 20 U mg⁻¹ tissue). On the basis of these trajectories, the authors recommended commencing first feeding shortly after 230 dd and no later than 276 dd to avoid starvation. All assays were run at 0.2 M ionic strength (physiological for marine fish, pH 7.0–7.8, with BAPNA (trypsin), soluble starch (α-amylase), p-nitrophenyl myristate + 0.5% Triton X-100 (lipase), and p-nitrophenyl phosphate (alkaline phosphatase)); activities were reported as nmol product min⁻¹. Nevertheless, this is only an example of an ontogenetic pattern in relation to feeding and digestive capacities because interspecific diversity is probably the main characteristic of digestive function in fish. In fact, different patterns can be observed during larval development, with each fish species giving preference to the appearance of proteases or carbohydrases according to diet habits. [4]. In species that develop a stomach, gastric differentiation is the final milestone in gut ontogeny and coincides with a marked rise in digestive capacity [5]. The timing of the appearance of gastric glands and pepsinogen mRNA expression and enzymatic activity is also a species-specific characteristic and may vary between hours and weeks after the mouth opening [6]. As an example, in the carnivorous sparid Sparidentex hasta, rudimentary gastric glands appear at 12 days post-hatch (dph), a functional stomach at 20 dph, and full maturation at 25 dph; pepsin activity is first detected two days after pepsinogen mRNA expression and increases steeply during weaning [7]. However, in the beluga (Huso huso), pepsin activity has been detected from the first feeding [8].

Once yolk reserves are depleted, indirectly developing larvae—and species hatching as miniature juveniles—depend entirely on external food sources. Growth and survival, therefore, hinge on four sequential capacities: ingestion, digestion, absorption, and assimilation [9].

The hydrolysis of dietary macromolecules is mediated by pepsin-like acid proteases (in stomach-bearing species), a suite of alkaline proteases (trypsin, chymotrypsin, amino- and carboxypeptidases), amylases, lipases, and acid/alkaline phosphatases. According to the authors, trophic habit shapes enzyme prominence: carnivorous taxa exhibit higher protease activities, whereas herbivorous and omnivorous fish display comparatively elevated carbohydrase levels [10]. Temperature serves as a master modulator, influencing feeding motivation, gastric and intestinal evacuation, catalytic efficiency, mucosal tissue remodeling, and absorptive transport [11,12]. Each species operates within a narrow thermal window beyond which digestive efficiency, health, and performance decline [10].

Recent method-oriented reviews on alkaline phosphatase [13], pepsin-like acid proteases [14], total alkaline proteases [15], α-amylase [16], and lipase [17,18] reveal marked variability in substrate choice, assay pH, temperature, molar extinction coefficients, and activity unit definitions. This heterogeneity hampers meaningful cross-study comparisons, particularly when targeting digestive capacity, here defined as the physiological enzymatic potential for dietary bond hydrolysis.

Sampling time is another critical aspect that differs across studies and deserves consideration. Digestion is a dynamic process led by ingestion and the transit of digesta throughout the digestive tract. The complex mechanisms regulating the synthesis and activation of digestive enzymes induce the predominance of different enzymes at different moments of the digestion of a meal [19]. Therefore, a single sampling point may underestimate the activity of some given enzymes.

Because nutrient utilization in fish is governed by three interacting capacities—ingestion, digestion, and absorption—this review focuses specifically on digestion (digestive capacity). The objectives are as follows:

• Catalog historical and current uses of the term digestive capacity.
• Synthesize the analytical approaches employed to quantify it.
• Propose a physiologically grounded definition based on hydrolytic potential.
• Recommend harmonized assay procedures applicable across teleost species (and other taxa) and their environmental contexts.

2. Digestive Capacity in Fish

2.1. Systematic Search

A SCOPUS query for “digestive capacity” AND fish (January 2020–June 2024) retrieved 70 records. In total, 8 non-fish papers were excluded, leaving 62 relevant studies (Supplementary Table S1). Additional references cited within these articles were reviewed for context but not included in the quantitative synthesis. References cited within tables are described in Supplementary material (Table S1).

2.2. Operational Definitions: Digestive Capacity

Among the 62 studies, nearly half (30) used the term digestive capacity without providing an explicit definition (

Table 1. Digestive capacity definitions and the type of indicators used in studies.

Digestive Capacity Definition	Studies	References	%
The activity of digestive enzymes.	19	4, 5, 7, 15, 16, 17, 18, 26, 33, 34, 42, 43, 49, 51, 52, 57, 61, 64, 69	30.6
No definition, but the activity of digestive enzymes.	18	6, 8, 10, 19, 27, 40, 44, 48, 54, 55, 56, 58, 59, 60, 63, 65, 66, 70	29.0
The anatomical properties.	3	1, 9, 29	4.8
No definition, but anatomical properties.	4	2, 30, 35, 37	6.5
The anatomical properties and the activity of digestive enzymes.	6	13, 22, 23, 45, 46, 68,	9.7
No definition, but the anatomical properties and the activity of digestive enzymes.	4	14, 24, 39, 47	6.5
No definition, but the intestine upregulated the interactome.	1	20	1.6
The in vivo digestibility (apparent digestibility of diet protein) a digestive capacity assay.	1	36	1.6
No definition, but in vivo digestibility (apparent digestibility coefficient).	1	3	1.6
The gene expression of digestive enzymes (pepsin, trypsin, chymotrypsin, amylase, and phospholipase A2).	1	50	1.6
No definition, but gene expression (genes related to protein, fat, and carbohydrate digestion, absorption pathways, and pancreatic secretion).	1	11	1.6
No definition, but viscerosomatic index, hepatosomatic index, and the expression of intestine transport genes.	1	25	1.6
The gastric evacuation rate (the amount of meal remaining in the stomach) as a measure of digestive capacity.	1	28	1.6
The use of residual nutrients on their biological performance as a digestive capacity.	1	38	1.6

). Nine equated it with morphometric traits such as villus length, goblet cell density, foregut fold area, or the mass of the stomach, pyloric caeca, and intestine. Three defined it through transcriptomic or interactomic profiles of digestive enzymes and transporters. The remaining 20 explicitly linked digestive capacity to enzymatic activity and/or in vitro digestibility assays. Overall, 47 studies measured at least one digestive enzyme.

2.3. Taxonomic and Ontogenetic Coverage

The dataset encompasses 36 teleost species (35 genera) plus 1 hybrid, comprising 20 freshwater and 16 marine taxa (

Table 2. Studied fish species.

Species	Environment: Milieu	Preferred Temperature	References	%
Acanthopagrus latus	Marine; freshwater; brackish; demersal	21.7–28.5, mean 27.4 °C	48, 65	3.2
Acipenser baerii	Freshwater; brackish; demersal;	1 °C–19 °C (Environm.)	69	1.6
Amphiprion ocellaris	Marine; reef-associated; non-migratory	26.2–29.3, mean 28.7 °C	22	1.6
Argyrosomus regius	Marine; brackish; benthopelagic; oceanodromous	13.3–19.4, mean 15.3 °C	58	1.6
Carassius auratus gibelio	Freshwater; brackish; benthopelagic	10 °C–20 °C	54	1.6
Centropomus viridis	Marine; demersal. Tropical	22.3–29.1, mean 26.3 °C	61	1.6
Channa argus	Freshwater; benthopelagic. Subtropical	4 °C–22 °C	39	1.6
Ctenopharyngodon idella	Freshwater; brackish; benthopelagic; potamodromous	0 °C–35 °C	42	1.6
Cyprinus carpio	Freshwater; brackish; benthopelagic	3 °C–35 °C	4, 6, 8, 17, 37, 49, 57	11.3
Danio rerio	Freshwater; benthopelagic	18 °C–24 °C	11	1.6
Epinephelus coioides	Marine; brackish; reef-associated	24.4–29.1, mean 28.1 °C	47	1.6
Epinephelus fuscoguttatus♀×lanceolatus♂	Marine; reef-associated	25.4–29.1, mean 28.2 °C (E. fuscoguttatus), 24.3–29.1, mean 28.1 °C (E. lanceolatus)	63, 66	3.2
Gadus morhua	Marine; brackish; benthopelagic; oceanodromous	0.5–10.3, mean 6.6 °C	45	1.6
Gymnocorymbus ternetzi	Freshwater; pelagic	20 °C–26 °C	68	1.6
Ictalurus punctatus	Freshwater; demersal	10 °C–32 °C	29	1.6
Larimichthys crocea	Marine; brackish; benthopelagic; oceanodromous	20.8–24.7, mean 22.8 °C	23	1.6
Lateolabrax maculatus	Marine; freshwater; brackish; reef-associated; catadromous	12.7–26.3, mean 22.4 °C	24, 25	3.2
Megalobrama amblycephala	Freshwater; benthopelagic	10 °C–20 °C	34	1.6
Micropterus salmoides	Freshwater; benthopelagic	10 °C–32 °C	1, 15, 40, 55	6.5
Mugil cephalus	Marine; freshwater; brackish; benthopelagic; catadromous	11.3–27.9, mean 23.2 °C	38	1.6
Oncorhynchus mykiss	Marine; freshwater; brackish; benthopelagic; anadromous	10 °C–24 °C	26, 30, 36	4.8
Oreochromis niloticuss	Freshwater; brackish; benthopelagic; potamodromous	14 °C–33 °C	33, 64	3.2
Paracheirodon innesi	Freshwater; pelagic	20 °C–26 °C	13	1.6
Pelteobagrus fulvidraco	Freshwater; demersal	16 °C–25 °C	5, 19	3.2
Pseudoplatystoma punctifer	Freshwater; demersal. Tropical	Nind	50	1.6
Puntigrus tetrazona	Freshwater; benthopelagic	20 °C–26 °C	44	1.6
Salmo salar	Marine; freshwater; brackish; benthopelagic; anadromous	2 °C–9 °C	20	1.6
Sander lucioperca	Freshwater; brackish; pelagic; potamodromous	6 °C–22 °C	46	1.6
Scophthalmus maximus	Marine; brackish; demersal; oceanodromous	5.9–11.9, mean 9.4 °C	18, 51	3.2
Seriola dumerili	Marine; reef-associated; oceanodromous	16.9–29, mean 27.1 °C	9	1.6
Silurus meridionalis	Freshwater; demersal. Subtropical	NInd	28	1.6
Siniperca chuatsi	Freshwater; benthopelagic	4 °C–22 °C	16, 27	3.2
Sparus aurata	Marine; brackish; demersal	12.1–21, mean 17.8 °C	2, 35, 43, 52, 56, 60, 70	11.3
Tachysurus fulvidraco	Freshwater; demersal	16 °C–25 °C	10	1.6
Totoaba macdonaldi	Marine; brackish; benthopelagic	19.8–26.7, mean 22.5 °C	3, 7, 14	4.8
Vieja melanurus, V. bifasciata	Freshwater; brackish; benthopelagic/freshwater; benthopelagic. Tropical	24 °C–30 °C (V. melanurus), 26 °C–30 °C (V. bifasciata)	59	1.6

Environment: milieu (https://www.fishbase.se/); preferred temperature (https://www.fishbase.se/), accessed on 22 June 2025.

). Cyprinus carpio, Micropterus salmoides, and Sparus aurata were the most frequently investigated models. Larval and juvenile stages dominate the dataset (~94%;

Table 3. Fish stage in studies.

Stage	References	%
Embryo	11	1.6
Larvae	9, 13, 22, 23, 33, 44, 46, 48, 50, 52, 56, 68	19.4
Postlarvae	15, 35	3.2
Juvenile	1, 2, 3, 4, 5, 6, 6, 7, 8, 10, 14, 16, 17, 18, 19, 20, 24, 25, 26, 27, 28, 29, 30, 34, 36, 37, 38, 39, 40, 42, 43, 47, 49, 51, 54, 55, 57, 58, 59, 60, 61, 63, 64, 65, 66, 69, 70	74.2
Adult	45	1.6

).

2.4. Enzymes Assessed

Across the 62 studies, the enzymes most frequently quantified were lipase (39 studies), trypsin (36), α-amylase (27), pepsin-like acid protease (18), total alkaline protease (18), alkaline phosphatase (10), chymotrypsin (7), leucine aminopeptidase (6), general aminopeptidase (4), leucine–alanine peptidase (1), and acid phosphatase (1) (

Table 4. Digestive enzymes in studies.

Enzyme Types	References	%
Amylase, lipase, pepsin (acid protease), and phosphatase (alkaline)	1	1.6
Amylase, lipase, pepsin, and trypsin	65, 66	3.2
Amylase, phosphatase (alkaline), and trypsin	6	1.6
Leucine-aminopeptidase, protease (alkaline), and trypsin	14	1.6
Aminopeptidase (N), amylase, chymotrypsin, lipase, phosphatase (alkaline), and trypsin	56	1.6
Aminopeptidase (N), phosphatase (alkaline), pepsin, and trypsin,	46	1.6
Aminopeptidase (N), amylase, lipase, phosphatase (alkaline), protease (alkaline), and trypsin	43	1.6
Aminopeptidase, amylase, pepsin, phosphatase (acid and alkaline), and trypsin	52	1.6
Amylase, chymotrypsin, lipase, pepsin, and trypsin	10	1.6
Amylase, chymotrypsin, lipase, protease (alkaline), and trypsin	38, 60	3.2
Amylase, leucine-amino peptidase, lipase, and phosphatase (alkaline)	48	1.6
Amylase and lipase	18, 37	3.2
Amylase, lipase, and pepsin	69	1.6
Amylase, lipase, pepsin, and trypsin	27	1.6
Amylase, lipase, phosphatase (alkaline), and protease (alkaline)	34	1.6
Amylase, lipase, and protease (alkaline)	17, 40, 42, 54, 64	8.1
Amylase, lipase, and trypsin	3, 4, 5, 7, 24, 26, 49, 55, 57, 63	16.1
Amylase, lipase, trypsin, and pepsin	51	1.6
Amylase, pepsin, and trypsin,	39	1.6
Amylase, leucine-aminopeptidase, lipase, and trypsin	23	1.6
Chymotrypsin, lipase, pepsin, protease (alkaline), and trypsin	61	1.6
Chymotrypsin, pepsin, protease (alkaline), and trypsin	8	1.6
Chymotrypsin, trypsin, and chymotrypsin/trypsin ratio	15	1.6
Leucine-alanine peptidase and phosphatase (alkaline)	35	1.6
Lipase, pepsin, and trypsin	13, 68, 70	4.8
Lipase and protease (alkaline)	16, 70	3.2
Lipase, protease (alkaline), and trypsin	58	1.6
Lipase and trypsin	19, 44	3.2
Pepsin (not measured, just mentioned)	45	1.6
Phosphatase (alkaline) and trypsin	25	1.6
Protease (acid and alkaline)	33, 59	3.2
Protease (alkaline) and trypsin	47	1.6
Not determined	2, 9, 11, 20, 28, 29, 30, 36, 50	14.5

).

2.4.1. α-Amylase

α-Amylase was analyzed in 37 studies. Four reported catalytic rates in μmol/min, but only three provided total activity (U fish⁻¹, U organ⁻¹, or U g⁻¹). Commercial diagnostic kits were commonly used (

Table 5. Amylase methods.

Substrate or Method (Number of Studies)	Unit	Activity Expression
ELISA kits (Shanghai Jianglai Industry Co., Ltd., Shanghai, China) (1)	Nind	Specific activity
EPS, 37 °C (1)	Nind	Specific activity
Kit (Jiancheng Biotech., Nanjing, China) (19)	Nind	Specific activity
Kit (Molecular Probes, Eugene, OR, USA), pH 6–9 (1)	RFU (Relative Fluorescence Units)	Total activity (RFU/mg larva dry weight)
Kit (NInd) (1)	Nind	Specific activity
Kit (Spinreact, Girona, Spain) (2)	1 μmol product/min at 37 °C	Specific activity
Starch (25 °C) (2)	0.1 Abs U/min	Specific activity
Starch (1)	1 Abs U/min	U/Organ
Starch (1)	1 Abs U	U/g weight? Or U/mg wet fish?
Starch (1)	1 mg starch hydrolyzed/30 min	Specific activity
Starch (1)	1 μmol/min	Specific activity
Starch, pH 7.4 (1)	1 mg starch hydrolyzed/30 min	Specific activity
Starch-DNS (2)	Nind	Specific activity
Starch-Iodine (1)	1 mg starch hydrolyzed/30 min	Specific activity
Starch-Iodine, 25 °C (1)	1 mg starch hydrolyzed/30 min	u/Fish, U/mg protein
Starch-Somogyi-Nelson (1)	1 μmol/min	U/L of serum
Nind: not indicated, EPS (4,6-Ethylidene-(G7)-1,4-nitrophenyl-(G1)-D-maltoheptaoside)
Specific activity (U/mg protein or U/g protein)

).

2.4.2. Chymotrypsin

Seven studies assessed chymotrypsin activity. One quantified catalytic rate, while three reported total activity values (

Table 6. Chymotrypsin methods.

Substrate or Method (Number of Studies)	Unit	Activity Expression
BTEE (1)	1 Abs U/min	U/Organ
BTEE (1)	1 Abs U	U/g weight? Or U/mg wet fish?
BTEE, pH 7.8 (1)	Nind	Specific activity
BTEE, pH 7.9 (1)	1 μmol product/min at 25 °C	Specific activity
Kit (Jiancheng Biotech.) (1)	Nind	Specific activity
N-Succinyl-Ala-Ala-Phe-7-amido-4-methylcoumarin (1)	RFU (Relative Fluorescence Units) per mg larvae dry weight	Total activity (RFU/mg larva dry weight)
SAPNA (1)	Nind	ND
Specific activity (U/mg protein or U/g protein)
SAPNA: N-succinil-Ala-Ala-Pro-Phe p-nitroanilide

).

2.4.3. Leucine–Alanine Peptidase and Leucine Aminopeptidase

Two studies quantified leucine-alanine peptidase and four quantified leucine-aminopeptidase. Only one defined the enzyme unit by absorbance change, and one gave total activity (

Table 7. Leucine-amino peptidase methods.

Substrate or Method (Number of Studies)	Unit	Activity Expression
Kit (Jiancheng Biotech.) (1)	Nind	Specific activity, U/dL (amylase)
Kit (NInd) (1)	Nind	Specific activity
L-leucine p-nitroanalide (1)	1 Abs U/min	U/Organ
L-leucine p-nitroanalide (1)	Nind	Specific activity
Specific activity (U/mg protein or U/g protein)

).

2.4.4. Lipase

Lipase was the most frequently studied enzyme, with 41 reports. Eleven studies measured ester-bond hydrolysis rates, and six quantified total activity. p-Nitrophenyl esters—especially medium-to-long-chain derivatives—were the preferred substrates, and commercial assay kits were commonly used (

Table 8. Lipase methods.

Substrate or Method (Number of Studies)	Unit	Activity Expression
4-methylumbelliferyl butyrate, pH 7.0 (1)	RFU (Relative Fluorescence Units)	Total activity (RFU/mg larva dry weight)
DGGR (1,2-o-dilauryl-rac-glycero-3-glutaric acid-(6′-methylresorufin) ester) (1)	1 μmol FFA/min	Specific activity
ELISA kits (Shanghai Jianglai Industry Co., Ltd.) (1)	Nind	Specific activity
Kit (Jiancheng Biotech.) (18)	Nind	Specific activity
Kit (NInd) (1)	Nind	Specific activity
Kit (Spinreact) (1)	Nind	Specific activity
Kit (Spinreact), 1-2-O-dilauryl-rac-glycero-3-glutaric acid-(6′-methylresorufin)-ester (1)	1 μmol product/min at 37 °C	Specific activity
pNPCaproate (1)	Nind	U/Fish
pNPCaproate (1)	1 μmol s/min at 30 °C	U/Larvae
pNPCaproate, pH 7.4 (1)	1 μmol product/min at 25 °C	Specific activity
PNPCaproate, pH 7.4, 30 °C (1)	1 μmol product/min	U/Fish
pNPCaproate (1)	1 μmol product/min	U/Fish
pNPMyristate (1)	1 Abs U/min	U/Organ
pNPMyristate (2)	Nind	Specific activity
pNPMyristate (4)	1 μmol product/min	U/L of serum
pNPMyristate (25 °C, pH 8.5) (2)	0.1 Abs U/min	Specific activity
pNPMyristate, 25 °C, pH 9.0 (1)	1 μmol pNPM/min	u/Fish, U/mg protein
pNPPalmitate (1)	Nind	Specific activity
Nind: not indicated; NA: not available
Specific activity (U/mg protein or U/g protein)

).

2.4.5. Acid and Alkaline Phosphatases

Acid phosphatase was analyzed in three studies and alkaline phosphatase was analyzed in thirteen. Four studies reported catalytic rates in moles/min, and two supplied total activity (

Table 9. Acid and alkaline phosphatase methods.

Substrate or Method (Number of Studies)	Unit	Activity Expression
Acid phosphatase
Kit (Jiancheng Biotech.) (2)	Nind	Specific activity
pNP-Phosphate, pH 4.8	1 μmol/min	Specific activity
Alkaline phosphatase
4-methylumbelliferyl phosphate	RFU (Relative Fluorescence Units)	Total activity (RFU/mg larva dry weight)
Kit (Jiancheng Biotech.) (4)	Nind	Specific activity
Kit (Sigma), pNP-Phosp	1 μmol product/min at 37 °C	Specific activity
pNP-Phosphate	Nind	Specific activity
pNP-Phosphate	Nind	Specific activity
pNP-Phosphate	Nind	Specific activity
pNP-Phosphate, 37 °C, pH 8.0	1 nmol/min	Total activity (U/mg dry weight)
pNP-Phosphate, pH 9.8	1 μg/min	Specific activity
pNP-Phosphate, pH 9.8	1 μmol/min	Specific activity
NInd	Nind	Specific activity
NInd: not indicated; NA: not available
Specific activity (U/mg protein or U/g protein)

).

2.4.6. Pepsin-like Acid Protease

Twenty-one studies examined pepsin-like proteolytic activity, almost exclusively using hemoglobin as the substrate. No one calculated the number of peptide bonds hydrolyzed per unit of time, although five reported total activity (

Table 10. Acid protease methods.

Substrate or Method (Number of Studies)	Unit	Activity Expression
Hemoglobin (1)	1 Abs U/min	U/Organ
Hemoglobin (3)	Nind	Specific activity
Hemoglobin (1)	Nind	U/Fish
Hemoglobin (1)	Nind	Specific activity
Hemoglobin (1)	1 μmol/min	U/Fish
Hemoglobin, 60 mM HCl, 37 °C (1)	1 μmol tyrosine/min	U/Fish
Hemoglobin, pH 2.0 (1)	1 μmol tyrosine/min	Specific activity
Hemoglobin, pH 2.0 (2)	1 μg tyrosine/min	U/Fish and U/mg protein
Hemoglobin, pH 3.0 (1)	1 μg tyrosine/min at 37 °C	Specific activity
Hemoglobin, pH 3.0 (1)	1 μmol product/min at 25 °C	Specific activity
Kit (Jiancheng Biotech.) (7)	Nind	Specific activity
Hemoglobin? (Lowry method) (1)	Nind	Specific activity
Nind	Nind	Specific activity

).

2.4.7. Total Alkaline Protease

Seventeen papers evaluated total alkaline protease, generally using azocasein or casein as substrates. No one calculated the number of peptide bonds hydrolyzed per unit of time, and only two quantified total activity (

Table 11. Alkaline protease methods.

Substrate or Method (Number of Studies)	Unit	Activity Expression
Azocasein (1)	Abs/min	Specific activity
Azocasein (1)	1 μmol azo/min	Specific activity
Azocasein, 37 °C (1)	Nind	Specific activity
Azocasein, pH 9.0 (1)	NInd	Specific activity
Casein (1)	1 Abs U	U/g weight? Or U/mg wet fish?
Casein, pH 9.0 (1)	1 μmol product/min at 37 °C	Specific activity
Casein, pH 9.0 (1)	1 μg tyrosine/min at 37 °C	Specific activity
Casein, pH 9.0 (1)	1 μmol product/min at 25 °C	Specific activity
Casein? Or azocasein? (1)	1 Abs U/min	U/Organ
Kit (Jiancheng Biotech.) (5)	Nind	Specific activity
Kit (Spinreact) (1)	Nind	Specific activity
Lowry method (1)	Nind	Specific activity
Nind	Nind	Specific activity
Nind: not indicated; NA: not available
Specific activity (U/mg protein or U/g protein)

).

2.4.8. Trypsin

Trypsin activity was assessed in 33 studies, nearly always with BAPNA as the substrate. Seven studies calculated peptide-bond hydrolysis rates (in moles/min), while seven reported total activity (

Table 12. Trypsin methods.

Substrate or Method (Number of Studies)	Unit	Activity Expression
BAPNA (1)	1 Abs U/min	U/Organ
BAPNA (3)	Nind	Specific activity
BAPNA (1)	Nind	U/Fish
BAPNA (1)	1 Abs U	U/g weight? Or U/mg wet fish?
BAPNA, 37 °C (1)	1 μmol/min	Specific activity
BAPNA, 25 °C, pH 8.2 (1)	0.1 Abs U/min	Specific activity
BAPNA, 25 °C, pH 8.2 (1)	1 μmol/min	Specific activity
BAPNA, pH 7.4 (1)	1 μmol/min	Specific activity
BAPNA, pH 7.5, 30 °C (3)	1 μmol/min	U/Fish
BAPNA, pH 8.2 (1)	Nind	Specific activity
BAPNA, 25 °C, pH 9.0 (1)	1 μmol/min	Specific activity
Boc-Gln-Ala-Arg-7- methylcoumarin hydrochloride	RFU (Relative Fluorescence Units)	RFU/mg larva dry weight
BTEE or BAPNA? (2)	Nind	Specific activity
ELISA kits (Shanghai Jianglai Industry Co., Ltd.) (1)	Nind	Specific activity
Kit (Jiancheng Biotech.) (12)	Nind	Specific activity
Kit (NInd)	Nind	Specific activity
Nind: not indicated; NA: not available
Specific activity (U/mg protein or U/g protein)

).

3. Discussion—Interpreting Digestive Capacity Measurements

Of the 62 studies reviewed, 47 (75.8%; Table 1) quantified digestive capacity solely through enzyme assays, thereby operationally defining the concept as the aggregate catalytic potential present in gut tissues. This approach is pragmatic: it reflects the biochemical ceiling for nutrient hydrolysis. However, its comparative value hinges on the units and expressions used to report activity.

3.1. Toward a Standard Unit

To harmonize reporting, we recommend adopting the International Union of Biochemistry and Molecular Biology (IUBMB) definition of the enzyme unit (U): one unit is the amount of enzyme that catalyzes the conversion of 1 μmol of substrate per minute under specified assay conditions (1 U = 16.67 nkat) [20]. Expressing total activity in U fish⁻¹, U organ⁻¹, U fish g⁻¹, or U organ g⁻¹ enables direct scaling from in vitro catalytic rates to whole-animal enzymatic potential, improving cross-study comparability and ecological relevance.

3.2. Ionic Environment Matters

Digestive capacity studies encompass both freshwater and marine teleosts. Because salinity determines the ionic composition of digestive fluids, enzyme kinetics are substantially influenced by Na⁺ and Ca²⁺ concentrations [21]. Seawater typically contains ≈0.46 M Na⁺ and ≈20 mM Ca²⁺ [22], whereas freshwater holds ≤2.4 mM Na⁺ and ≈0.10 mM Ca²⁺ [23]. We therefore recommend the following: (i) measuring Na⁺ and Ca²⁺ directly in gastric and intestinal fluids or sourcing species-specific data from the literature, and (ii) adjusting reaction buffers to reflect those physiological levels. Where such information is unavailable, in vitro analysis of ionic optima for the target enzymes offers a robust alternative.

3.3. Temperature—The Most Neglected Variable

Each species operates within a specific thermal window, yet assay temperatures are rarely reported and often default to 37 °C, a practice inherited from mammalian biochemistry. Because catalytic velocity increases with temperature, assays conducted at 37 °C routinely overestimate in vivo activity [24,25,26,27,28]. We urge investigators to conduct enzyme assays at the physiological temperature—defined either as (a) the ambient temperature of the species’ natural habitat or (b) the rearing temperature used in experimental culture.

3.4. pH—An Equally Critical Parameter

Fewer than one-quarter of the reviewed studies reported assay pH (Table 5, Table 6, Table 7, Table 8, Table 9, Table 10, Table 11 and Table 12). Given the strong pH dependence of the catalytic rate, activity values are not directly comparable—even within a single species—without this information [29,30,31,32]. Whenever possible, stomach and intestinal pH should be measured in situ and replicated in reaction buffers. If direct measurement is impractical, published values for the focal or a closely related species can be used. In larval studies, culture-water pH may serve as a provisional proxy. However, in vitro assays that employ enzyme pH optima outside the physiological range should be avoided.

3.5. Gaps in Enzyme Coverage

The current dataset encompasses major digestive hydrolases—pepsin-like acid protease, trypsin, chymotrypsin, amino- and leucine-peptidases, phosphatases, α-amylase, and lipase. Two notable gaps remain: (i) carboxypeptidases have not been assayed in any reviewed study, and (ii) chymotrypsin appears in only seven. This underrepresentation is consequential: in Dicentrarchus labrax larvae, the trypsin-to-chymotrypsin activity ratio ranges from 0.24 to 0.41 during early weaning, indicating that chymotrypsin may be at least as influential as trypsin in vivo [33].

3.6. Closing the Gaps

Implementing standardized enzyme units, ion-adjusted buffers, physiological temperatures, and in situ pH measurements will significantly improve cross-study comparability and help close remaining knowledge gaps in teleost digestive physiology.

3.7. Digestive Capacity Determination Incorporating Extract Theoretical Volume (ETV)

Most studies report only specific activity (U mg⁻¹ or U g⁻¹ protein), a metric that does not reflect the organism’s total catalytic potential. We recommend that authors provide total activity—expressed as U fish⁻¹, U organ⁻¹, U in organ g⁻¹ fish body mass, or U g⁻¹ organ—by following the workflow below:

1.

Sampling Through a Standardized Method

The sampling strategy must be standardized. In principle, procedures must be followed in accordance with the official Guidelines for the Use and Experimentation of Laboratory Animals.

The effect of the circadian cycle on enzyme activity must be considered, so fish in experimental treatments must be sampled and sacrificed at the same time of day. Generally, a 12 to 24 h fasting period is used before sampling. Any sampling time must be accompanied by sampling in the control experiment.

If a specific experiment is to compare the enzymatic capacity of fed and fasted fish, the recommendation (for greater digestive activity) would be to sample and sacrifice the fish two hours after the established feeding time (for example, if feeding is at 8:00 a.m., sample at 10:00 a.m. for both fed and fasted fish).

Digestive tract content washing procedures are not applicable for quantifying digestive capacity and should therefore not be performed.

If, for some experimental reason, the digesta content is considered to contribute significant weight, this contribution can be calculated by using additional fish from the same experimental treatment, only for weight calculation purposes.

Fish body mass_{with food} = Fish body mass_{without food} + Residual ingesta

Digestive organ mass_{with food} = Digestive organ mass_{without food} + Residual ingesta

2.

Identify and Isolate Digestive Organs

Use pressure forceps to reduce fluid transfer between organs, then weigh each organ.

3.

Prepare Enzymatic Extracts

Keep samples on ice during processing. Homogenize tissues using an appropriate method—ballistic homogenization (e.g., FASTPREP) or shear homogenization (e.g., ULTRATURRAX)—with a standard solvent. Clarify by centrifugation (13,000× g, 15 min), recover the supernatant, adjust the pH to the physiological lumen value for the digestive organ, centrifuge again, and store aliquots (0.5–1 mL) at –80 °C until use.

4.

Measure Crude Extract Activity (U mL⁻¹)

• Determine the change in absorbance per minute.
• Correct to a 1 cm light path.
• Convert absorbance to μmol of product released, using the molar extinction (absorption) coefficient, derived from a standard curve of the reaction product.
• One unit is the amount of enzyme that catalyzes the conversion of 1 μmol of substrate per minute under specified assay conditions.
• Calculate unit mL⁻¹.

5.

Calculate Extract Theoretical Volume (ETV)

$$\mathit{ETV}=\mathit{SV}+(\mathit{TM} \times {\displaystyle \frac{TH}{100}})$$

where

• SV = solvent volume (mL) used for homogenization;
• TM = tissue mass (g);
• %TH = (${\displaystyle \frac{\mathbf{T}\mathbf{H}}{\mathbf{100}}}$) = tissue moisture content (%).

Example: For 2.5 g of tissue homogenized in 3 volumes of Milli-Q water, with 71% tissue moisture:

ETV = 7.5 + (2.5 × 0.71) = 9.275 mL

6.

Compute Total Units (TU)

TU = Activity (U mL⁻¹) × ETV

where 1 U = enzyme amount releasing 1 μmol product min⁻¹ under physiological assay conditions.

7.

Calculate Digestive Capacity (DC)

Express DC as U fish⁻¹, U organ⁻¹, U in organ g⁻¹ fish body mass, and U g⁻¹ organ. For comparative purposes, specific enzyme activity (U mg⁻¹ protein) can also be reported.

4. Proposal for an Approach Framework to Quantify Digestive Capacity (DC)

Digestive efficiency in fishes is a multifactorial trait, reflecting the organism’s ability to convert ingested food into absorbable nutrients. Conventional metrics—such as arbitrary enzyme units or specific enzyme activities—provide valuable biochemical insights (related to the organ enzyme extract) but fail to integrate the dynamic interplay of enzymatic hydrolysis with physiological, developmental, and environmental contexts related to the digestive capacity of the fish. To address this gap, we propose a two-tiered conceptual framework for quantifying digestive efficiency, moving from simplified biochemical proxies toward integrative physiological indices.

4.1. Extract Theoretical Volume (ETV)

The Extract Theoretical Volume (ETV) was calculated by the formula: ETV = SV + (TW × H/100), where SV is the solvent volume (mL) of water or buffer used for extraction, TW is the weight (g) of organ tissue, and H is the moisture content of the digestive organ. According to Monge-Ortíz et al. (2020) [34], for Seriola dumerili, a 71.3% moisture tissue content (%TH) was considered. If the volume of NaOH or HCl used for pH adjustment is 5% or higher, it must be added to SV.

4.2. Digestive Capacity (DC)

Digestive capacity (CD) is the total number of active enzyme units in the fish’s digestive tract (ST, PC, AI, MI, PI) at time t. It was calculated using the formula CD = EA × ETV, where EA is the extract activity in U/mL, where 1 U is the enzyme required to hydrolyze bonds to release 1 μmol of product (amino group, reducing sugar, fatty acid, etc.) per minute.

4.3. Enzyme Activity Expression

Enzymatic activities as DC are expressed as total U Fish⁻¹, U Organ⁻¹, U g⁻¹ fish, U g⁻¹ organ, and additionally as specific enzyme activity (U mg⁻¹ protein).

4.4. Digestive Processing (DP)

Digestive Processing (DP) is the number of bonds hydrolyzed by the active enzymes (E), where E are the U of total acid protease (stomach), total alkaline protease, total amylase, total lipase, etc. (pyloric caeca and intestine), according to DC, present in the digestive tract at time t, which generate the μmoles of free products (amino group, reducing sugar, fatty acid, etc.) in the digestive tract during the studied digestion time (DC vs. time). The average DPI can be expressed as U-min or U-hour, respectively. For example, if the acid digestion capacity is 14 U/fish and the digestion time under those conditions is 50 min, the DPI for that time interval will be 700 U-min.

5. Kit-Based Versus Custom Assays—Strengths, Pitfalls, and Recommendations

Commercial kits dominate recent digestive enzyme research: aminopeptidase (4 studies), amylase (24), chymotrypsin (1), leucine-aminopeptidase (1), lipase (22), acid phosphatase (2), alkaline phosphatase (4), pepsin-like protease (7), alkaline protease (6), and trypsin (14). Kits are convenient—reagents arrive pre-formulated and protocols are turnkey—but they are primarily designed for mammalian (often human) samples. As a result, buffer composition seldom reflects fish physiology. Standard formulations typically omit or under-represent NaCl and CaCl₂, ions that modulate many fish digestive enzymes, and nearly all are calibrated for pH ≈ 7.4 and 37 °C—at the least, temperature conditions rarely encountered by aquatic ectotherms. Since suppliers withhold full reagent compositions, researchers cannot readily adjust the ionic strength or pH, and unless otherwise reported, incubations must be assumed to occur at the kit’s default 37 °C.

Another limitation is unit reporting. Some kits fail to yield activity in µmol substrate hydrolyzed min⁻¹. A widely used amylase kit, for instance, quantifies residual starch after 30 min at 37 °C and reports activity as “units per 100 mL extract,” a metric that cannot be converted to true catalytic rates without additional calibration.

Natural macromolecular substrates—e.g., hemoglobin for acid protease or casein for alkaline proteases—better capture total proteolytic digestive potential. However, reporting only the absorbance increase at 280 nm min⁻¹ provides no stoichiometric link to peptide bond hydrolysis, even using a standard tyrosine curve. This can be remedied by quantifying free amino groups with the OPA assay, where 1 µmol of released amines equals 1 µmol of peptide bonds cleaved [14,15]. Similarly, azocasein, though widely used, yields azo-pigments that cannot be stoichiometrically related to bond cleavage. There is no information for any other secondary calibration.

For amylase, using starch as a substrate permits accurate stoichiometry if reducing sugars are quantified. This can be achieved at the microplate scale by the Somogyi–Nelson method [35] or the DNS method [16], with hydrolysis rates expressed in µmol glycosidic bonds cleaved min⁻¹.

Synthetic chromogenic and fluorogenic substrates (e.g., p-nitroanilides, amido-4-methylcoumarin (AMC) derivatives, p-nitrophenyl esters) are ideally suited for 96-well assays. The released chromophores (p-nitroaniline, AMC, p-nitrophenol) follow Beer–Lambert kinetics, allowing stoichiometric rate calculations if—and only if—a standard curve is prepared in the identical buffer used for the enzyme test (matching pH, NaCl, CaCl₂, bile salts, substrate concentration, wavelength, etc.). Any mismatch alters extinction coefficients and leads to biased results.

6. Integrative Technical–Biological Insights Leading to Recommendations

Kit-based assays are useful for rapid screening, but physiological accuracy requires either (i) custom buffer adjustments with ion and pH corrections or (ii) stoichiometric calibration of chromogenic/fluorogenic outputs. For comparative enzymology in fish, explicitly reporting assay temperature, ionic composition, and pH is indispensable.

Practical recommendations

• Match ionic conditions. Adjust assay buffers with NaCl and CaCl₂ concentrations that reproduce the species-specific ionic environment of the digestive tract.
• Calibrate in the identical buffer. Construct chromophore standard curves in the same ion-balanced buffer, using the same microplate, wavelength, and incubation temperature as the enzymatic reaction.
• Report absolute units. Express activity as µmol bonds hydrolyzed per minute. Scale results beyond specific activity (U mg⁻¹ protein) to total activity (U fish⁻¹, U organ⁻¹, or U g⁻¹ organ), ensuring biological relevance and enabling robust cross-study comparisons.

By following these steps, microplate assays with natural or synthetic substrates yield enzyme-rate data that are chemically accurate, physiologically relevant, and directly comparable across studies. This alignment strengthens the link between in vitro enzymatic assays and in vivo digestive performance, reducing artifacts introduced by kit defaults or non-physiological conditions.

7. Conclusions

7.1. Operational Definition

Digestive capacity can now be defined in operational terms that unify enzyme kinetics with organismal physiology:

Digestive capacity is the total number of active enzyme units present in a fish’s digestive system at the moment of sampling, where one enzyme unit (U) equals the amount of enzyme that hydrolyzes one µmol of substrate bonds per minute under the assay’s specified ionic, pH, and thermal conditions, assayed under a physiologically relevant pH, ion balance and temperature, and expressed as total active enzyme units. Digestive capacity may then be normalized as U fish⁻¹, U g⁻¹ fish body mass, U organ⁻¹, or U g⁻¹ organ; ST = stomach, PC = pyloric caeca, AI = anterior intestine (foregut), MI = medium intestine (midgut), and PI = posterior intestine (hindgut) (Figure 1).

7.2. Generalized Concept

For all organisms, digestive capacity is the total number of active enzyme units present in the digestive milieu (organs, lumen, or secreted phase) at the time of sampling. One enzyme unit corresponds to the hydrolysis of 1 µmol of substrate bonds per minute, under defined assay conditions. To ensure comparability, digestive capacity should always be normalized to an appropriate scale—per individual, per organ, or per gram of tissue.

7.3. Minimum Reporting Checklist

To render digestive capacity data reproducible, physiologically meaningful, and cross-comparable, authors should report, at minimum, the following:

• Biological context:
  • Species and developmental stage;
  • Habitat or culture conditions (water temperature, pH, salinity);
  • Sample size, mean body mass, and recent feeding status (including feed information).
• Sampling details:
  • Time of slaughter relative to last feeding;
  • Organs collected, residual feed status, organ wet mass.
• Extract preparation:
  • Homogenization equipment and conditions—type of water (distilled or mili-Q) or buffer composition (including physiological NaCl and CaCl₂);
  • Solvent volume and tissue moisture used to calculate theoretical extract volume.
• Enzyme assay specifics:
  • Substrate and concentration;
  • Ionic additives (the proper bile and concentration for lipases) and final buffer pH;
  • Incubation temperature (physiological) and duration;
  • Detection wavelength (absorbance/fluorescence)—indicates whether the method is kinetic or endpoint;
  • Chromophore/fluorescent standard curve and molar extinction coefficient;
  • Explicit unit definition (µmol product min⁻¹);
  • Indicate the units’ calculation formulas.

7.4. General Guidance

• How to assess the digestive capacity of fish

To determine the digestive capacity of a fish, the following factors must be considered: (a) Species. (b) Origin of the fish (aquaculture or wild). (c) Physiological conditions of its environment of origin, such as temperature, pH (environmental and in the digestive tract), and optimal concentration of NaCl and CaCl₂ for the activity of digestive enzymes. (d) Feeding and fasting history prior to analysis. (e) Weight of the fish. (f) Weight of the digestive organs or their regions (foregut, midgut, hindgut). (g) Preparation of enzyme extracts preferably in milli-Q water at digestive pH. (h) Optimization of enzymatic methods based on the amount of enzyme extract, extract dilution, digestion time, pH, and physiological temperature, optimal or physiological NaCl and CaCl₂, and quantification of enzyme units (U) as μmoles of substrate hydrolyzed per minute. (i) Calculation of U of enzyme fish⁻¹, U of enzyme g⁻¹ of fish, U of enzyme in the digestive organ g⁻¹ of fish, and U of enzyme g⁻¹ of digestive organ.

• Which enzyme activities need to be measured

The basic enzymes to be measured are those involved in the hydrolysis of the main food polymers. (a) For proteins and peptides: acid protease (fish with a true stomach), alkaline protease, trypsin, chymotrypsin, and leucine aminopeptidase. (b) For polysaccharides, oligosaccharides, and disaccharides (mainly starches, glycogen, maltose): amylases and maltase. (c) For lipids: lipases. (d) For phosphorus-containing polymers: acid phosphatase and alkaline phosphatase. However, there may be specific studies where it is required to measure the digestive capacity of other polymers (chitin, cellulose, nucleic acids, etc.), including the specific enzymes involved [36].

• Which digestive enzyme in which tissue is being examined

(a). True stomach: Pepsin-like. (b) Pancreas: alkaline proteases, trypsin, chymotrypsin, amylases, alkaline phosphatase, and lipases. (c) Pyloric caeca: alkaline proteases, trypsin, chymotrypsin, amylases, and lipases. (d) Intestine: alkaline proteases, trypsin, chymotrypsin, leucine-alanine aminopeptidase, leucine aminopeptidase (brush border), amylases, maltase (brush border), alkaline phosphatases (brush border), and lipases [37].

• What the patterns of activity variation in these digestive enzymes in different tissues are

The patterns of variation in enzyme activity across different organs depend on several factors, with the most important being ontogenetic development, the circadian cycle, and dietary stimulation. For fish species with true stomachs, the onset of pepsin activity occurs when the stomach is developed and functional. In general, digestive capacity (proteases, amylases, and lipases) is expected to increase with the development of the fish [38].

8. Way Forward

Adopting this operational definition and harmonized checklist will accomplish the following:

• Establish a universal benchmark for digestive enzyme assays across taxa and laboratories.
• Bridge the gap between in vitro enzymatic rates and whole-animal catalytic potential.
• Advance toward reliable estimations of digestive capacity during the complete process of digestion (from ingestion to egestion) of a meal.
• Strengthen physiological relevance, enabling more robust applications in nutritional physiology, ecological comparisons, and aquaculture innovation.

Teleost fishes, though morphologically simpler than birds or mammals, exhibit extraordinary functional diversity and environmental plasticity. Their enzyme systems dynamically adjust to fluctuations in salinity, temperature, bile–salt composition, and nutrient inputs, while maintaining distinctive ω-3/ω-6 lipid biochemistry. These traits position teleosts as powerful comparative models for probing vertebrate digestion, lipid metabolism, and adaptive evolution.

In conclusion, digestive capacity in teleosts reflects the interplay of developmental constraints and ecological adaptation. By treating them not as simplified systems, but as finely tuned evolutionary models, future research can reveal general principles of digestive physiology that transcend aquatic contexts—informing aquafeed design, biomedical nutrition, and our broader understanding of vertebrate evolution.