Abstract
Introduction:
This study aimed to systematically compare the tissue specific nutrient and protein profiles of the head, body, and viscera of farmed Hippocampus abdominalis.
Methods:
A total of 90 individuals were dissected into three anatomical regions: head, body (including trunk and tail), and viscera. All measurements were performed on three independent biological replicates (n = 3) .
Results:
The results revealed that the viscera, though often discarded as a processing by product, exhibited the most balanced essential amino acid profile and the highest protein quality indices. In contrast, the body was rich in structural proteins and collagen, while the head contained high concentrations of minerals (especially magnesium and zinc) as well as proteins associated with neural and visual functions.
Discussion:
These tissue specific functional specializations provide a molecular basis for the value added utilization of seahorse by products and offer practical insights for sustainable aquaculture practices.
1 Introduction
Hippocampus abdominalis belongs to the phylum Vertebrata, class Osteichthyes, order Gasterosteiformes, suborder Syngnathoidei, family Syngnathidae, and genus Hippocampus (). It is currently recognized as the largest species within its genus, reaching lengths of up to 30 centimeters. According to Traditional Chinese Medicine (TCM), Hippocampus has a warm nature, a salty and sweet taste, and acts on the kidney and liver meridians—energy pathways associated with vital functions and metabolism (). China is recognized as the earliest country to utilize Hippocampus for medicinal and health purposes. Today, Hippocampus remains an important medicinal material. Recent studies have reported various bioactivities of seahorse extracts. For example, identified antioxidants from a water extract of H. japonicus using an in vitro assay. showed that recombinant cystatin B from H. abdominalis exhibits anti-apoptotic effects in cell culture. demonstrated anti-fatigue activity of peptides derived from whole Hippocampus in a mouse model. In folk tradition, it is customary to use “a pair of Hippocampus” as a medicinal treatment for impotence, infertility, and similar conditions.
However, the large-scale Hippocampus trade has exerted severe pressure on wild populations (). In 2002, CITES listed all Hippocampus species in Appendix II, requiring export permits and non-detriment findings for wild catches, while encouraging captive-bred trade. To meet market demand and promote sustainability, China started artificial breeding of H. trimaculatus in 1957 (). The Australian H. abdominalis was introduced to China in 2016 because of its strong disease resistance, rapid growth, and high juvenile survival. These traits make it highly suitable for intensive aquaculture, and China has become the world’s leading producer of farmed seahorses.
Despite these advances, research on the nutritional quality of H. abdominalis remains limited. Most previous studies focused on the effects of rearing conditions and sex on nutrient composition (; ; ; ). Key knowledge gaps still exist: no proteomic study has linked protein expression to tissue function; no systematic comparison of nutrient elements across body parts has been performed; and the potential value of the discarded viscera remains unaddressed. To fully leverage the potential of farmed H. abdominalis as a food resource, a detailed understanding of its nutritional constituents, particularly proteins, is essential. The amino acid composition of proteins determines their nutritional value; well-balanced profiles improve bioavailability and utilization in humans (Yang et al., 2024; ). Among protein fractions, sarcoplasmic proteins are rich in bioactive peptides (), myofibrillar proteins contain high levels of essential amino acids (), and stromal proteins support skin, bone, and joint health ().
A systematic comparison of the nutrient and protein profiles across different body parts of H. abdominalis is therefore necessary to understand their tissue-specific biological functions. In this study, an integrated analysis of farmed H. abdominalis was conducted, quantifying the proximate composition, amino acid profile, and mineral content, and exploring the underlying molecular mechanisms via proteomics across its key tissues: head, body, and viscera.
2 Materials and methods
2.1 Materials
Specimens of H. abdominalis were purchased from Weihai Yinze Biotechnology Co., Ltd. (Weihai, China) and stored frozen at −20 °C for less than three months before thawing and analysis. A total of 90 individuals (mean body length: 10.00 ± 1.00 cm; mean weight: 4.00 ± 0.50 g) were used in this study. The 90 individuals were randomly divided into three independent biological replicates (n = 3), with 30 individuals per replicate. After thawing, a transverse incision was made at the junction of the gill openings and pectoral fins to separate the head from the rest of the body. A ventral incision was subsequently performed to excise the visceral organs. The remaining portion, which comprised both the anatomical trunk and the tail, was designated as the “body” sample. For each biological replicate, the head, body (including trunk and tail), and viscera from the 30 individuals were dissected and pooled separately to obtain one pooled sample per tissue type per replicate. Thus, for each tissue type (head, body, viscera), three independent pooled samples (biological replicates) were obtained. Whole-body samples were prepared for each biological replicate by homogenizing the 30 intact individuals and pooling the homogenates. All measurements were performed once on each biological replicate sample. Analytical-grade reagents, including boric acid, hydrochloric acid, and nitric acid, were purchased from Sinopharm Group Reagent Co., Ltd. (Shanghai, China).
2.2 Basic nutritional analysis
Moisture content was determined using the direct drying method according to GB 5009.3-2016 (). Approximately 2–5 g of homogenized sample was weighed into a pre-dried, constant-weight aluminum dish and placed in an electric forced-air drying oven (DHG-9070A, Jinghong, China) at 101–105°C with natural convection. The sample was dried for 4 h, then transferred to a desiccator, cooled to room temperature, and weighed. Drying was repeated in 1 h cycles until the difference between two consecutive weighing was ≤ 2 mg, which was defined as constant weight. The moisture content was calculated as the percentage of mass loss relative to the initial sample mass.
Crude protein content was determined by the Kjeldahl method following GB 5009.5-2016 (). Approximately 0.5–1 g of sample was digested with a catalyst mixture composed of copper sulfate (CuSO4) and potassium sulfate (K2SO4) in a 1:15 (w/w) ratio, together with 12 mL of concentrated sulfuric acid. Digestion was carried out at 420 °C until the solution became clear and colorless. After cooling, the digest was diluted and distilled using a Kjeldahl distillation unit. The liberated ammonia was collected in a boric acid solution (20 g/L) containing a mixed indicator of methyl red and bromocresol green. The distillate was titrated with 0.1 mol/L hydrochloric acid until the color changed from green to pink. A nitrogen-to-protein conversion factor of 6.25 was used to calculate crude protein content.
Crude fat content was measured by the Soxhlet extraction method according to GB/T 5009.6-2016 (). Dried sample (2–5 g) was placed into a cellulose thimble and extracted with petroleum ether (boiling point 30–60 °C) for 6–8 h at a circulation rate of 4–6 cycles per hour. The extraction was performed in a Soxhlet apparatus with a pre-weighed receiving flask. After extraction, the solvent was evaporated on a rotary evaporator, and the residue was dried to constant weight at 101–105 °C. The crude fat content was calculated as the percentage of extracted residue relative to the dry sample weight.
Ash content was determined by the muffle furnace method according to GB 5009.4-2016 (). Approximately 2–5 g of sample was placed in a pre-weighed crucible and carbonized on an electric stove until no more smoke was produced. The crucible was then transferred to a muffle furnace (SX2-4-10A, Longyue, Shanghai, China) and ashed at 550 °C ± 25 °C for 4–6 h until a white or light gray residue was obtained. A blank crucible was processed in parallel to correct for any contamination. The crucible was cooled to room temperature in a desiccator and weighed. The ash content was calculated as the percentage of residue relative to the initial sample mass.
Total carbohydrate content was determined using the phenol-sulfuric acid method as specified in GB/T 15672-2009, with results expressed as glucose equivalents (). Approximately 0.1 g of sample was hydrolyzed with 10 mL of 6 mol/L hydrochloric acid in a boiling water bath for 30 min. After hydrolysis, the solution was neutralized with 6 mol/L sodium hydroxide and diluted to a final volume of 50 mL with distilled water. An aliquot of the hydrolysate was mixed with 6% (w/v) phenol and concentrated sulfuric acid in a test tube. The mixture was then placed in an electrically heated thermostatic water bath (HH-4, Shanghai Boxun Medical Biological Instrument Corp., Shanghai, China) set to 100 °C (boiling water) and kept for 30 min and the absorbance was measured at 490 nm using a UV-Vis spectrophotometer. A standard curve was prepared using glucose solutions at concentrations of 0, 20, 40, 60, 80, and 100 μg/mL. The total carbohydrate content was calculated as glucose equivalents and expressed in g/100 g of sample.
2.3 Collagen content determination
The total hydroxyproline content was determined using a commercial colorimetric assay kit (Solarbio, Beijing, China). Approximately 0.5 g of each freeze-dried pooled sample (head, body, viscera, whole) was hydrolyzed in 10 mL of 6 mol/L HCl at 110 °C for 12 h. After hydrolysis, the solution was neutralized to pH 6–7 with 10 mol/L NaOH and then diluted to 50 mL with distilled water. An aliquot of 0.1 mL was mixed with 0.4 mL of oxidation reagent (chloramine T solution) and allowed to react at room temperature for 20 min. Subsequently, 0.4 mL of colorimetric reagent (Ehrlich’s reagent) was added, and the mixture was incubated at 60 °C for 15 min. After cooling to room temperature, the absorbance was measured at 560 nm using a UV-Vis spectrophotometer. A standard curve was prepared using hydroxyproline standards (0–10 μg/mL). The collagen content was calculated by multiplying the hydroxyproline content by a factor of 7.46.
2.4 Mineral element and heavy metal content
Mineral and heavy metal contents were determined using microwave digestion followed by inductively coupled plasma mass spectrometry (ICP-MS) according to GB 5009.268-2016 (). Approximately 0.5 g of each freeze-dried pooled sample (head, body, viscera, whole) was digested with 5 mL of concentrated HNO3 and 2 mL of H2O2 (30%) using a microwave digestion system. The digestion program was: ramp to 120 °C in 5 min and hold for 5 min, then ramp to 180 °C in 10 min and hold for 20 min. After cooling, the digests were diluted to 50 mL with ultrapure water and filtered through a 0.45 μm membrane. Blank digests were prepared in parallel. The solutions were analyzed by ICP-MS under standard operating conditions. Calibration curves were constructed using multi-element standard solutions (0–200 μg/L), and rhodium (10 μg/L) was used as an internal standard.
2.5 Purine content determination
The purine content was determined by high-performance liquid chromatography (HPLC), adapted from the method of . To determine the purine content, different parts of the H. abdominalis were freeze-dried, ground, and passed through a 20-mesh sieve. Approximately 0.5 g of the sample was accurately weighed and mixed with 20 mL of water. The mixture was subjected to ultrasonic treatment for 30 minutes, followed by centrifugation at 12,000 rpm for 5 minutes. The supernatant was collected, and the procedure was repeated for the pellet. The two supernatants were combined into a 50 mL volumetric flask, diluted to the mark with water, and mixed thoroughly. The combined solution was then filtered through a 0.22 μm membrane for subsequent analysis. The purine analytes quantified were adenine, guanine, xanthine, and hypoxanthine. Calibration curves were constructed using authentic standards (0.5–50 μg/mL) without an internal standard. The results obtained on a dry weight basis were converted to fresh weight (wet weight) using the separately determined moisture content of each sample. Final results are expressed as mg per gram of fresh weight (mg/g, wet basis). The HPLC conditions were as follows: injection volume: 20 μL; column temperature: 30°C; flow rate: 1 mL/min; mobile phase: water (containing 0.02 mol/L potassium dihydrogen phosphate, pH adjusted to 3.2): methanol = 98:2 (v/v); detection wavelength: 254 nm.
2.6 Determination of amino acid content
The content of hydrolyzed amino acids in different parts of H. abdominalis was determined using the acid hydrolysis method according to GB 5009.124–2016 (). Specifically, 0.2–0.4 g of sample was weighed into a hydrolysis tube, and 10–15 mL of 6 mol/L hydrochloric acid (containing 0.5% mercaptoethanol) was added. The mixture was hydrolyzed at 110°C for 22 h. After being diluted to 50 mL in a volumetric flask, 1 mL of the solution was withdrawn, dried under a nitrogen stream, and then reconstituted. The amino acid content was analyzed using a fully automated amino acid analyzer (Hitachi L-8900, Japan).
2.7 Amino acid nutritional evaluation
This study is based on the amino acid scoring patterns recommended by the Food and Agriculture Organization of the United Nations (FAO) in 2013 for different age groups () and the essential amino acid content of the commonly used reference protein, egg. Protein indices, including the Essential Amino Acid Index (EAAI) (), Amino Acid Score (AAS) (), Chemical Score (CS), Nutritional Index (NI) (), Predicted Biological Value (P-BV), and Protein Efficiency Ratio (P-PER) (), were calculated for different regions of the H. abdominalis to evaluate the nutritional value of its amino acids.
2.8 Protein fraction extraction from H. abdominalis
The extraction of sarcoplasmic, myofibrillar, and matrix proteins from different regions of H. abdominalis was performed following the method described by with modifications. Specifically, 10 g of tissue was precisely weighed, ground, and homogenized with 6 volumes of 0.03 mol/L phosphate-buffered saline (PBS, pH 7.4). The homogenate was centrifuged at 10,000×g for 20 min, and the supernatant was collected. This extraction procedure was repeated twice. The combined supernatants constituted the sarcoplasmic protein fraction. To the remaining sediment, six volumes of PBS (0.1 mol/L, pH 7.4) containing 1.1 mol/L KI were added. The mixture was homogenized and centrifuged, and the supernatant was collected. This process was repeated twice. The combined supernatants represented the myofibrillar protein fraction. After degreasing with an ethyl acetate-ethanol mixture (3:1, v/v), the precipitate was divided into two equal portions. One portion was extracted with 5 volumes of 0.1 mol/L acetic acid at 25°C for 24 h, followed by centrifugation at 7000×g for 20 min. The supernatant was collected as the acid-soluble matrix proteins. Another portion was extracted with 5 volumes of 0.1 mol/L NaOH at 25°C for 10 h. After centrifugation, the supernatant constituted the alkaline-soluble matrix proteins. The remaining precipitate was neutralized with HCl, washed with water, and then treated with 5 volumes of 5% (w/v) sodium dodecyl sulfate (SDS) at 85°C for 1 h in a water bath to obtain the alkaline-insoluble matrix proteins. The protein concentration of each component was determined using the bicinchoninic acid (BCA) assay.
2.9 SDS-PAGE
The extracted proteins from different fractions were mixed with 5×SDS loading buffer at a ratio of 4:1 (v/v) and incubated in a boiling water bath (HH-4, Shanghai Boxun Medical Biological Instrument Corp., Shanghai, China) at 100°C for 5 min. The SDS-polyacrylamide gel was prepared using a commercial rapid preparation kit (Epizyme, Shanghai). A 4 μL aliquot of a pre-stained protein molecular weight marker was loaded into one lane of the gel. For each sample, 10–20 μL was loaded into individual lanes. Any unloaded wells were filled with 10 μL of 1×SDS loading buffer. Electrophoresis was performed initially at 80 V through the stacking gel, followed by 110 V through the separating gel. After electrophoresis, the gel was stained with a solution containing 0.5% (w/v) Coomassie Brilliant Blue R-250. Subsequently, the gel was destained with an ice-cold solution of acetic acid, methanol, and water (1:3:6, v/v/v) until the background became clear.
2.10 Proteomics analysis
Total protein was extracted from the freeze-dried pooled samples of each tissue type (head, body, viscera, and whole) using a phenol-based method. For each tissue type, a single processing was performed. Proteins were precipitated, washed, and redissolved. After reduction, alkylation, and acetone precipitation, the protein was digested with trypsin (1:50, w/w) overnight at 37 °C.
Peptides were desalted using SOLAμ™ SPE plates and separated by reverse-phase HPLC for spectral library generation. For data-independent acquisition (DIA), peptides were analyzed on a nanoElute UPLC coupled to a timsTOF Pro mass spectrometer.
DIA data were processed using DIA-NN software (version 1.8) against the Hippocampus genus UniProt database (Taxonomy ID: 72046). Differentially expressed proteins were identified with thresholds of fold change ≥ 2.0 or ≤ 0.5 and p < 0.05 (Student’s t-test).
2.11 Data processing and analysis
A total of 90 individuals were randomly divided into three independent biological replicates (n = 3), with 30 individuals per replicate. For each biological replicate, the head, body, and viscera from the 30 individuals were dissected and pooled separately to obtain one pooled sample per tissue type per replicate. Thus, for each tissue type (head, body, viscera), three independent pooled samples (biological replicates) were obtained. Whole-body samples were prepared for each biological replicate by homogenizing the 30 intact individuals and pooling the homogenates. All measurements were performed once on each biological replicate sample. Percentage data satisfied normality and homogeneity of variance (Shapiro–Wilk and Levene’s tests, both p > 0.05) and were analyzed without transformation. One-way ANOVA using SPSS version 26.0 followed by Duncan’s test was used to compare sample types (head, body, viscera, whole) with a significance level of p < 0.05. All graphs were generated using Origin 2024.
3 Results and discussion
3.1 Evaluation of essential nutrients
The nutritional value of fish primarily depends on its protein and fat composition (). As shown in Figure 1, H. abdominalis exhibits a high-protein, low-fat profile. The crude protein content in the body was the highest, reaching 19.66 g/100 g, exceeding that of grass carp, bighead carp, Siberian sturgeon, and wels catfish (15.69–18.25 g/100 g) (), cultured sea bass (18.00 g/100 g) (). Moreover, the body constituted the largest portion of H. abdominalis, making it a promising source for extracting proteins and peptides. The highest concentrations of crude fat and total carbohydrates were found in the viscera, at 5.34 and 2.03 g/100 g, respectively.
Figure 1
This inverse distribution—high protein in the body versus high lipid and carbohydrate in the viscera—reflects tissue-specific metabolic functions. In fish, skeletal muscle (body) is primarily composed of contractile proteins and supports locomotion (; ). In contrast, the viscera serve as the central metabolic hub for nutrient digestion, absorption, and energy storage (). Lipids are the most energy−dense fuel, and visceral adipose tissue acts as the primary energy reservoir (; ). Thus, allocating lipids to visceral organs while sparing protein for structural functions represents an adaptive energy trade-off ().
Marine organism lipids have been reported to contain unique fatty acids and phospholipids (). Therefore, the lipid-rich viscera warrant further investigation for the characterization of specific lipid compounds. As the primary center for energy metabolism, visceral organs benefit from high fat content for rapid energy supply, while high sugar content supports short-term energy demands (). Thus, the viscera of H. abdominalis represent a potential source of bioactive lipids that warrant further study.
3.2 Collagen content analysis
Differences in collagen content reflect variations in physiological functions and metabolic demands across distinct regions of the H. abdominalis. As shown in Figure 1-f, the body region exhibited the highest collagen content among the three anatomical parts, reaching 44.47 mg/g. Collagen is primarily localized in the skin, periosteal bone fragments, and skeletal structures of the H. abdominalis (). Collagen is a key component of muscles and other connective tissues, providing structural support and elasticity. These properties are essential for the body region because it undergoes repeated bending and relaxation during swimming; collagen fibrils in connective tissues store elastic energy and improve swimming efficiency (Zhu et al., 2023). In contrast, the viscera function primarily in energy supply and metabolic regulation, requiring less structural support; thus, their collagen content is lower.
3.3 Mineral element content analysis
The mineral content in different body parts reflects their specific physiological functions and metabolic requirements. In animals, the liver is the primary site for iron storage, with the majority stored as heme iron, a key component for hemoglobin synthesis (). As shown in Table 1, the highest iron content was found in the viscera of H. abdominalis (302.15 mg/kg), which was significantly higher than that in the head and body (p < 0.05).Magnesium serves as an essential cofactor for enzyme activity, neuronal signaling, and muscle contraction (). Consistent with these roles, the head contained significantly higher magnesium (2396.52 mg/kg) than the viscera and body (p < 0.05).
Table 1
| Mineral contents (mg/kg) | Head | Viscera | Body | Whole |
|---|---|---|---|---|
| Natrium(Na) | 2796.86 ± 10.53a | 2486.11 ± 33.99a | 2548.56 ± 25.47a | 2615.93 ± 23.85a |
| Potassium (K) | 952.35 ± 7.58a | 963.69 ± 10.19a | 1011.77 ± 2.06a | 1003.82 ± 4.25a |
| Magnesium (Mg) | 561.51 ± 10.29c | 323.96 ± 4.50a | 456.20 ± 24.78b | 464.64 ± 19.48b |
| Calcium (Ca) | 13906.46 ± 864.97c | 416.72 ± 9.64b | 9797.79 ± 735.64b | 9494.64 ± 674.15b |
| Zinc (Zn) | 37.37 ± 1.98a | 28.65 ± 0.23a | 39.72 ± 8.37a | 38.21 ± 6.11a |
| Iron (Fe) | 35.18 ± 4.85b | 58.62 ± 4.12c | 19.64 ± 0.85a | 28.04 ± 2.09ab |
| Copper (Cu) | 1.02 ± 0.15a | 1.35 ± 0.06a | 1.11 ± 0.45a | 1.14 ± 0.34a |
| Phosphorus (P) | 23836.50 ± 1323.97c | 3265.10 ± 103.93a | 17609.26 ± 1173.15b | 17161.25 ± 1075.56b |
| Selenium (Se) | 1.48 ± 0.05b | 2.10 ± 0.04c | 1.40 ± 0.02a | 1.52 ± 0.02ab |
| Manganese (Mn) | 13.80 ± 0.62c | 5.53 ± 0.78a | 9.68 ± 0.43b | 10.06 ± 0.52b |
The content of mineral elements in different parts of the H. abdominalis.
a–cMeans with different superscript letters in the same row indicate significant differences (p < 0.05). Data are expressed as means ± SD (n = 3 biological replicates).
Zinc is an essential trace element involved in enzymatic reactions, immune function, and protein synthesis. Among the three anatomical parts, the body exhibited the highest zinc concentration (39.72 mg/kg). However, no statistically significant differences were detected among the parts (Table 1, p > 0.05). In comparison to another seahorse species, H. erectus, the zinc levels in H. abdominalis were significantly higher, suggesting species-specific differences in trace element accumulation ().
Selenium is a component of antioxidant enzymes such as glutathione peroxidases and also contributes to thyroid hormone metabolism (). Its content was highest in the viscera (2.10 mg/kg), followed by the head (1.48 mg/kg) and body (1.40 mg/kg), with significant differences among all three parts (Table 1, p < 0.05). The high selenium level in the viscera may support antioxidant defense against oxidative damage in metabolically active organs such as the liver, pancreas, and kidney, where selenium−dependent glutathione peroxidases detoxify peroxides generated during intense metabolism (). Elevated selenium in H. abdominalis viscera likely supports high oxidative metabolism and detoxification, consistent with its role in redox balance ().
3.4 Heavy metal content analysis
Table 2 shows the heavy metal content in different regions of H. abdominalis. The head region exhibited a greater tendency for heavy metal accumulation, particularly lead. Among the three anatomical parts, the head contained the highest lead concentration. In contrast, heavy metal concentrations were lower in the body. The viscera contained relatively high levels of arsenic, reaching 2.12 mg/kg. However, studies indicate that most arsenic in fish exists as organic arsenic, with inorganic arsenic levels being lower ().
Table 2
| Heavy metal content (mg/kg) | Head | Viscera | Body | Whole | Limited standard |
|---|---|---|---|---|---|
| Cadmium (Cd) | 0.027 ± 0.0068b | 0.016 ± 0.0014b | 0.0076 ± 0.0018ab | 0.012 ± 0.0024a | ≤0.1 |
| Chromium (Cr) | 1.75 ± 0.43b | 0.93 ± 0.015a | 1.00 ± 0.13ab | 1.14 ± 0.0057ab | ≤2.0 |
| Arsenic (As) | 0.71 ± 0.13b | 2.12 ± 0.12c | 0.38 ± 0.031a | 0.64 ± 0.03ab | ≤0.5 |
| Plumbum (Pb) | 1.33 ± 0.58b | 0.27 ± 0.057a | 0.24 ± 0.056a | 0.46 ± 0.11ab | ≤0.5 |
The heavy metal content in different parts of the H. abdominalis.
a–cMeans with different superscript letters in the same row indicate significant differences (p < 0.05). Data are expressed as means ± SD (n = 3 biological replicates). Limited standard values refer to the maximum allowable levels for aquatic products (fish) according to the Chinese National Food Safety Standard GB 2762-2022.
Lead in the head exceeded the GB 2762–2022 limit for aquatic products (0.5 mg/kg). Therefore, consumption of the head alone should be limited. This elevation can be attributed to lead being a persistent, bioaccumulative, and toxic (PBT) metal that tends to accumulate in lipid-rich tissues, bones, and neural tissues (). In fish, lead uptake occurs primarily via the gills, where it rapidly binds and accumulates; gill lead concentrations can be up to four-fold higher than in the skeleton or skin (). Moreover, the head contains dense neural tissues, extensive blood supply, and lead-binding proteins, all of which facilitate lead sequestration (). However, the overall heavy metal levels of the whole H. abdominalis are below the regulatory thresholds, indicating that whole consumption of this H. abdominalis is safe.
3.5 Purine content analysis
Purines (adenine, guanine, xanthine, hypoxanthine) are intermediates in nucleotide metabolism. However, high intake of purine-rich foods may increase serum uric acid levels (). The purine content in different parts of H. abdominalis is shown in Figure 2. The viscera contained the highest levels of adenine (0.43 mg/g), guanine (0.96 mg/g), and xanthine (0.17 mg/g), significantly higher than in the head and body (p < 0.05). In contrast, hypoxanthine showed the highest concentration in the body (0.47 mg/g), which was significantly higher than in the head and viscera (p < 0.05).
Figure 2
High purine levels in the viscera reflect active nucleotide turnover in metabolically active organs such as the liver and intestine, consistent with their roles in energy production and detoxification (). Meanwhile, hypoxanthine accumulation in the body arises from ATP breakdown during muscular contraction, as adenosine nucleotides are catabolized to hypoxanthine under high energy demand (Yin et al., 2021).
3.6 Amino acid content analysis
As shown in Table 3, The total amino acid content in the body section of the H. abdominalis is significantly higher than in other parts (p < 0.05), reaching 131.94 g/kg. In the viscera of H. abdominalis, essential amino acids constitute over 45% of the total amino acids. This exceeds the ideal protein pattern suggested by FAO/WHO (), which is more than 35.38%. The ΣEAA/ΣAA ratio in the head and body of the H. abdominalis also approaches this pattern.
Table 3
| (mg/g protein) | Head | Viscera | Body | Whole |
|---|---|---|---|---|
| Ile | 24.03 ± 0.20c | 38.03 ± 2.41d | 15.45 ± 0.13a | 18.21 ± 0.35b |
| Leu | 46.95 ± 0.44c | 76.80 ± 3.74d | 30.35 ± 0.05a | 35.83 ± 2.42b |
| Lys | 47.24 ± 0.13c | 65.08 ± 3.55d | 41.34 ± 1.13a | 44.06 ± 2.14b |
| Met+Cys | 23.06 ± 0.35c | 29.47 ± 3.36d | 18.77 ± 0.43a | 20.25 ± 1.45b |
| Phe+Tyr | 51.42 ± 0.12c | 76.94 ± 2.91d | 38.12 ± 0.20a | 42.77 ± 0.24b |
| Thr | 35.50 ± 0.11c | 44.65 ± 2.61d | 24.30 ± 1.71a | 27.47 ± 1.14b |
| Val | 35.54 ± 0.28c | 46.98 ± 2.87d | 28.22 ± 1.67a | 30.74 ± 0.86b |
| His | 21.19 ± 0.16b | 26.07 ± 1.14c | 17.91 ± 2.50a | 30.74 ± 0.35d |
| Arg | 65.57 ± 0.49d | 63.88 ± 4.85c | 54.05 ± 1.61a | 57.09 ± 1.23b |
| Ala | 70.19 ± 0.97d | 62.08 ± 5.20b | 59.81 ± 0.66a | 62.35 ± 0.46c |
| Asp | 73.06 ± 0.32c | 89.21 ± 5.25d | 52.73 ± 0.62a | 58.56 ± 1.35b |
| Glu | 110.89 ± 0.32c | 125.27 ± 6.92d | 81.61 ± 2.11a | 89.61 ± 5.78b |
| Gly | 130.47 ± 3.45d | 66.57 ± 7.31a | 81.61 ± 15.89b | 127.79 ± 13.35c |
| Pro | 57.61 ± 1.00d | 37.80 ± 3.57a | 46.33 ± 6.25b | 48.27 ± 2.24c |
| Ser | 44.71 ± 0.54c | 48.67 ± 3.41d | 32.87 ± 1.59a | 36.00 ± 0.35b |
| Hypro | 44.92 ± 1.01d | 12.24 ± 2.41a | 41.47 ± 0.16c | 41.00 ± 0.35b |
| BCAA | 106.52 | 161.81 | 74.02 | 84.77 |
| AAA | 51.42 | 76.94 | 38.12 | 42.77 |
| ∑EAA/∑AA (mg/g) | 34.02% | 45.02% | 31.96% | 33.20% |
| ∑EAA/∑NEAA (mg/g) | 51.57% | 81.87% | 46.96% | 49.70% |
| EAAI | 58.84 | 81.8 | 44.16 | 49.16 |
| NI | 7.96 | 6.81 | 8.68 | 8.35 |
| AAS | Leu, 76.96 | Val, 117.46 | Leu, 49.76 | Leu, 58.74 |
| CS | Met+Cys, 40.46 | Met+Cys, 51.70 | Ile, 28.62 | Ile, 33.71 |
| P-BV | 52.44 | 77.46 | 36.43 | 41.88 |
| P-PER1 | 1.25 | 2.76 | 0.51 | 0.76 |
| P-PER2 | 1.44 | 2.79 | 0.71 | 0.95 |
Amino acid composition and evaluation of amino acid nutrition of different parts of the H. abdominalis.
Different superscript letters in the same row indicate significant differences between data points (p < 0.05). Data are means ± SD (n = 3 biological replicates). Specific metrics include: EAAI (Essential Amino Acid Index); AAS (Amino Acid Score); CS (Chemical Score); P-BV (Protein Biological Value); P-PER1 (Protein Performance Ratio1) and P-PER2 (Protein Performance Ratio2).
Glutamic acid and glycine were the predominant amino acids. Based on known taste−active amino acids, the high levels of Glu (savory/umami) and Gly (sweet) suggest that H. abdominalis may have a savory-sweet taste profile. The amino acids with the lowest concentrations were His and Ile. Histidine is important for children, as it promotes growth, improves sleep quality, and reduces fatigue (). Therefore, combining H. abdominalis with other histidine-rich foods can improve overall amino acid balance. Research indicates that restricting dietary isoleucine promotes metabolic health and extends lifespan in mice (). The low isoleucine content in H. abdominalis is consistent with this dietary pattern.
3.7 Amino acid nutritional evaluation
The amino acid nutritional evaluation results are presented in Table 3. Leucine and Isoleucine were identified as the first and second limiting amino acids for the head, body, and whole body of H. abdominalis. Compared with the FAO/WHO scoring pattern () and egg protein, the viscera exhibited the highest AAS (117.46 for valine) and CS (51.70 for Met+Cys), while the body had the lowest values (AAS 49.76 for leucine; CS 28.62 for isoleucine). The viscera also showed the highest P-PER values (2.76 and 2.79, well above the high−quality threshold of 2.0) and the highest EAAI (81.8). The body had the highest NI due to its protein content, whereas the viscera possessed a more balanced EAA profile despite a lower NI. The head contained higher levels of aromatic amino acids (AAAs), likely related to neural signaling (). High concentrations of branched−chain amino acids (BCAAs) in the head and viscera may support energy metabolism and protein synthesis (), while lower levels in the body reflect its lower metabolic demands.
Collectively, the protein quality indices (AAS, CS, P-PER, EAAI, and EAA profile) indicate that the viscera of H. abdominalis has favorable protein quality. Although often discarded as a by−product, the viscera warrants consideration as a valuable protein source for future applications.
3.8 Protein composition analysis
Figure 3-a shows regional differences in the protein composition of H. abdominalis. Water-soluble sarcoplasmic proteins constituted 42–75% of the total protein, salt-soluble myofibrillar proteins accounted for approximately 11–33%, and alkali-insoluble matrix proteins comprised 5–8%. Specifically, the body exhibited the highest myofibrillar protein content, accounting for 33% of the total protein. The visceral region contained the least myofibrillar protein. In most marine fish, the distribution differs markedly: sarcoplasmic protein constitutes about 20%, myofibrillar protein about 75%, and insoluble matrix proteins approximately 5% (). Myofibrillar proteins are crucial for the texture and quality of muscle foods, as their intra- and intermolecular interactions significantly influence these attributes (Zhu et al., 2025). Consequently, the muscular tissue of the body is likely to possess superior mechanical and chemical stability. In contrast, myofibrillar proteins were least abundant in the viscera, whereas sarcoplasmic proteins predominated. Sarcoplasmic proteins comprise approximately 100 distinct proteins, most of which are enzymes involved in glycolytic energy metabolism (). This composition may contribute to the relative instability and susceptibility to breakdown of the visceral tissue. Matrix proteins primarily constitute the connective tissue, including collagen, elastin, and reticulin, which are key determinants of meat tenderness and elasticity (). The distribution pattern of matrix proteins aligned with the regional variations in collagen content observed in this study.
Figure 3
3.9 SDS-PAGE
As shown in Figure 3, SDS-PAGE analysis revealed distinct regional differences in protein composition within the H. abdominalis. The major protein classes (myofibrillar, sarcoplasmic, and matrix) exhibited unique molecular weight distributions and banding patterns. Several prominent bands were tentatively identified as specific metabolic or structural proteins. In different regions of H. abdominalis, the molecular weight of myosin is primarily below 70 kDa, while that of myofibrillar proteins ranges mainly from 35 to 250 kDa. In the myosin proteins of the body and head, distinct bands were observed at 63 kDa and 35 kDa. In the acid-soluble matrix proteins of the viscera, bands were observed at molecular weights above 250 kDa and between 70 and 100 kDa. The molecular weight distribution of alkaline-insoluble matrix proteins in the body primarily ranged from 250 to 70 kDa. In contrast, the distribution in the viscera and head exhibited a broader range (250–25 kDa).
The prominent bands at 63 kDa and 35 kDa in the sarcoplasmic proteins of the body and head are tentatively identified as phosphoglucose isomerase and glyceraldehyde-3-phosphate dehydrogenase, respectively. The characteristic bands of myosin heavy chain, myosin light chain, actin, and troponin were most distinctly visible in the myofibrillar proteins of the body, followed by those in the head. In contrast, the viscera exhibited lower myofibrillar protein content, with characteristic bands such as myosin heavy chain and myosin light chain often absent. In the acid-soluble matrix proteins of the viscera, intense bands were observed at molecular weights above 250 kDa and between 70 and 100 kDa, likely representing acid-soluble collagen subunits. However, no distinct bands were observed in the head or body. The bands corresponding to alkaline-soluble matrix proteins in the body and viscera were faint or indistinct. This may be attributed to the presence of numerous low-molecular-weight proteins within the alkaline-soluble fraction, which have very similar molecular weights and therefore cannot be resolved. The molecular weight distribution of alkaline-insoluble matrix proteins in the body primarily ranged from 250 to 70 kDa. In contrast, that in the viscera and head exhibited a broader range, from 250 to 25 kDa.
3.10 Proteomics analysis
3.10.1 Proteomics data profiling and differential expression analysis
The results (Figure 4) showed that 6,268 proteins and 33,943 peptides were identified in different parts of H. abdominalis. The majority of peptides ranged from 7 to 20 amino acids in length. The number of identified peptides decreased with increasing length. This pattern indicates an appropriate distribution of peptide lengths and successful enzymatic digestion. In the three comparisons, 2,316, 1,688, and 1,295 proteins were upregulated, while 978, 472, and 1,314 proteins were downregulated. The Venn diagram shows that 795 proteins exhibited differential expression across different parts of the H. abdominalis.
Figure 4
3.10.2 Biological function analysis of tissue-specific proteins
To elucidate the biological functions of proteins in different parts of H. abdominalis, the unique proteins in each region were identified. Supplementary Table 1 lists the top 10 proteins with the highest expression levels. In the head, numerous lens-related proteins and myosin isoforms were identified. These proteins are crucial for maintaining lens clarity and facilitating muscle contraction (). The unique proteins in the viscera were primarily involved in carbohydrate metabolism, lipid metabolism, reproductive processes, and immune responses (). The body region contained unique proteins mainly related to signaling and cell cycle regulation. These proteins play crucial roles in protein modification, signal transduction, cell cycle regulation, and maintaining intracellular calcium balance.
3.10.3 Gene ontology analysis
GO analysis of different anatomical parts in H. abdominalis revealed highly tissue-specific functional divisions (Figure 5). Specifically, the body, as the main site of movement, is significantly enriched in biological processes related to myofibrillar assembly, muscle development, myocardial contraction, and sarcomere organization. Key protein complexes include the myosin-troponin complex, Z-disc structures, and numerous actin-binding proteins. The activation of these contraction-related pathways closely matches their high protein content and collagen-rich characteristics. This reflects a functional specialization for movement and structural support. The head exhibits unique features of visual perception and neural regulation. It is primarily enriched in pathways related to lens development and visual perception. Key components include those constituting the ocular lens, highlighting its complexity as a sensory hub for information processing and metabolic regulation. This functional profile is consistent with the high mineral content in the head.
Figure 5
The viscera, acting as the metabolic hub, showed significant enrichment in energy-yielding pathways. These include organic acid metabolism, fatty acid β-oxidation, the tricarboxylic acid cycle, and carbohydrate metabolism. Key cellular components and complexes include the proton-coupled ATP synthase complex, mitochondrial matrix components, the U2 snRNP spliceosome, lysosomal proteins, and several oxidoreductase enzymes. The significant enrichment of these metabolic pathways aligns with the chemical composition profile characterized by high fat, carbohydrate, and purine content, confirming the viscera’s role as a core energy supplier for physiological activities such as reproduction and immunity. Comparative proteomics showed that metabolic enzymes were up-regulated and muscle contraction proteins were down-regulated in the viscera compared to the body, which confirms their distinct functional specializations.
Overall, the body (including trunk and tail) is specialized for motility, supported by structural proteins. The head is specialized for visual perception, dependent on lens proteins. The viscera is specialized for energy metabolism and conversion, driven by metabolic enzymes. Collectively, these findings elucidate the molecular basis for the specialized processing of H. abdominalis. This knowledge facilitates the development of targeted strategies to optimize the utilization of its high-value protein resources.
3.10.4 Kyoto encyclopedia of genes and genomes analysis
KEGG pathway analysis revealed functionally distinct biological pathways enriched in different parts of H. abdominalis, aligning with their tissue-specific physiological roles (Figure 6). The body was significantly enriched in pathways related to muscle contraction and energy metabolism, underpinning its core functions in locomotion and energetics. The head region showed enrichment in pathways associated with neuroactive ligand-receptor interaction and synaptic signaling, highlighting its specialization in sensory perception and neural regulation. In contrast, the viscera exhibited dominant enrichment in comprehensive metabolic pathways (e.g., carbohydrate, lipid, and amino acid metabolism) as well as detoxification pathways (e.g., chemical carcinogenesis-reactive oxygen species), consistent with its role as the body’s primary metabolic and homeostatic hub.
Figure 6
This integrative analysis demonstrates that the tissue-specific nutrient profiles are not merely compositional facts but are direct consequences of these underlying, proteomically-defined physiological functions. The viscera’s enrichment in metabolic and detoxification pathways molecularly explains its high content of energy substrates (fats, carbohydrates) and purines, reflecting an investment in catabolism, anabolism, and xenobiotic processing. The body’s dominance of contractile and structural pathways directly accounts for its high myofibrillar protein and collagen content, representing an investment in locomotion and mechanical support. Similarly, the head’s neural and sensory protein enrichment, coupled with its high concentration of essential mineral cofactors like zinc and magnesium, points to a concerted investment in high-order information processing and regulation. This multi-omics perspective transcends mere correlation, providing a mechanistic understanding that both validates the traditional whole-organism use of seahorse and offers a compelling scientific rationale for its future targeted, tissue-specific valorization in nutraceutical and functional food applications.
4 Conclusion
In summary, the head, body, and viscera of H. abdominalis exhibit distinct functional specializations: The body represents the main edible portion, with the highest crude protein content and rich structural proteins that support locomotion. The body is particularly rich in myofibrillar proteins and collagen, making it suitable for direct consumption or as a source of gelatin and collagen peptides. The viscera are nutrient-dense, characterized by a balanced essential amino acid profile, high protein quality indices, and elevated metabolic enzyme activities. The head is abundant in essential minerals (notably magnesium and zinc) and contains proteins associated with visual perception and neural regulation.
The viscera and head are largely discarded as processing by-products, yet they possess considerable nutritional value. To align with circular economy principles and sustainable aquaculture, future research should focus on valorizing these tissues. Potential applications include their use as fish feed ingredients, the production of protein hydrolysates and bioactive peptides with antioxidant, antihypertensive, or immunomodulatory activities, and the development of nutraceuticals or functional food components. The body is also suitable for producing gelatin or collagen peptides.
The present study has several limitations, including the use of samples obtained from a single farm under specific rearing conditions. Consequently, future investigations should encompass multiple farms, controlled dietary interventions, and larger sample sizes to better capture nutritional variability. Despite these limitations, our findings provide a foundation for the comprehensive and sustainable utilization of H. abdominalis, transforming currently discarded by-products into valuable resources while maintaining a high-quality protein supply from the edible muscle tissue.
Statements
Data availability statement
The original contributions presented in the study are included in the article/Supplementary Material, further inquiries can be directed to the corresponding author/s.
Ethics statement
Ethical review and approval were waived for this study, as it exclusively utilized frozen tissue samples obtained from commercially farmed Hippocampus abdominalis. The specimens were donated by Weihai Yinze Biotechnology Co., Ltd. (Weihai, China), a facility that is licensed for large-scale artificial seahorse breeding and strictly complies with the Convention on International Trade in Endangered Species of Wild Fauna and Flora (CITES) and national aquatic wildlife protection laws. All animals were humanely euthanized prior to tissue collection. Consequently, this study involved no live animal experimentation, which is in full compliance with the ethical guidelines for animal research. Therefore, specific approval from an Animal Ethics Committee was not required.
Author contributions
FL: Writing – review & editing, Resources, Writing – original draft, Conceptualization, Formal Analysis. JR: Writing – review & editing, Formal Analysis, Conceptualization, Resources. HL: Writing – review & editing, Investigation, Methodology, Data curation. JY: Methodology, Writing – review & editing, Supervision. YW: Writing – review & editing, Investigation, Data curation. TZ: Project administration, Writing – original draft, Funding acquisition. YX: Writing – review & editing, Conceptualization, Supervision, Funding acquisition. GY: Writing – review & editing, Funding acquisition, Conceptualization.
Funding
The author(s) declared that financial support was received for this work and/or its publication. This work was financially supported by grants from the National Key Research and Development Program of China (2023YFF1103901) and the Hainan Provincial Key Research and Development Program (grant No. ZDYF2024XDNY188).
Acknowledgments
The authors acknowledge the assistance of Analysis and Testing Center, College of Food Science and Engineering, Ocean University of China.
Conflict of interest
HL was employed by company Qingdao Haoran Marine Technology Co., Ltd.
The remaining author(s) declared that this work was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.
Generative AI statement
The author(s) declared that generative AI was not used in the creation of this manuscript.
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Supplementary material
The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmars.2026.1847931/full#supplementary-material
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Summary
Keywords
amino acids, by-product utilization, Hippocampus abdominalis, nutritional evaluation, proteomics, tissue specialization
Citation
Li F, Ren J, Li H, Yu J, Wang Y, Zhang T, Xue Y and Yu G (2026) Nutrient and protein profiling of different parts of Hippocampus abdominalis: high value utilization of by-products. Front. Mar. Sci. 13:1847931. doi: 10.3389/fmars.2026.1847931
Received
05 April 2026
Revised
17 June 2026
Accepted
22 June 2026
Published
08 July 2026
Volume
13 - 2026
Edited by
Mohammed A. E. Naiel, Zagazig University, Egypt
Reviewed by
Jorge Palma, University of Algarve, Portugal
Vladimir G. Dvoretsky, Murmansk Marine Biological Institute, Russia
Updates
Copyright
© 2026 Li, Ren, Li, Yu, Wang, Zhang, Xue and Yu.
This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.
*Correspondence: Yong Xue, xueyong@ouc.edu.cn; Gang Yu, gyu0928@163.com
†These authors have contributed equally to this work
Disclaimer
All claims expressed in this article are solely those of the authors and do not necessarily represent those of their affiliated organizations, or those of the publisher, the editors and the reviewers. Any product that may be evaluated in this article or claim that may be made by its manufacturer is not guaranteed or endorsed by the publisher.