Dietary chlorogenic acid supplementation enhanced the growth performance and muscle quality of Procambarus clarkii

Abstract

Chlorogenic acid (CGA) is a natural polyphenolic compound with antioxidant activity. We hypothesised that dietary CGA supplementation could improve the growth performance and selected muscle-related traits of Procambarus clarkii, possibly in association with changes in antioxidant-related indices and the expression of selected genes. To test this hypothesis, a 6-week feeding trial was conducted using five diets containing 0, 200, 400, 600, and 800 mg/kg CGA. Dietary supplementation with 400–600 mg/kg CGA improved growth performance and feed conversion ratio, and was associated with higher collagen-related indices, better texture in some parameters, lower freezing loss, and higher amino acid contents in some treatment groups. CGA supplementation was also associated with higher antioxidant enzyme activities, lower reactive oxygen species and malondialdehyde levels, and changes in the expression of genes related to antioxidant defence and muscle development. Overall, these results suggest that dietary CGA supplementation, particularly at 400–600 mg/kg, may improve growth performance and selected muscle-related traits of P. clarkii under the present experimental conditions. These findings support further evaluation of CGA as a dietary supplement in P. clarkii.

Graphical abstract

Highlights

• •Dietary CGA improved growth performance in Procambarus clarkii.
• •CGA at 400–600 mg/kg improved muscle texture and reduced freezing loss.
• •Dietary CGA increased muscle antioxidant capacity in crayfish.
• •CGA regulated antioxidant and muscle growth-related gene expression.

1. Introduction

Procambarus clarkii is an important freshwater crustacean in aquaculture, and its production value depends not only on growth performance but also on muscle-related traits that influence edible quality and processing characteristics. In aquatic animals, these traits are multidimensional and include nutritional composition, texture, collagen-related properties, and water-holding capacity, all of which contribute to consumer acceptance and product value (Johnston et al., 2006; Periago et al., 2005). However, variation in these traits cannot be explained solely by proximate composition. Increasing evidence suggests that muscle-related characteristics are also associated with broader physiological processes, including oxidative status, connective tissue organisation, and muscle growth and protein deposition (Cao et al., 2026; Hua et al., 2025; Yang et al., 2020). This broader perspective is relevant because many studies in aquaculture have described muscle quality traits individually, whereas fewer have considered how these trait domains may vary together within the same nutritional context.

Among these factors, oxidative status has received growing attention because oxidative imbalance may impair muscle structure and postmortem quality, whereas stronger antioxidant defence has often been associated with more favourable texture and compositional characteristics (Jiang et al., 2016; Li et al., 2023). Collagen is also an important determinant of muscle firmness and resilience, and changes in collagen content or remodelling may contribute to variation in texture-related traits (Astruc, 2014; Grau-Bové et al., 2015) In parallel, pathways involved in muscle development and protein accretion, including IGF-1/Akt/mTOR-related signalling and MEF2-related regulation, have been implicated in myofiber growth and muscle formation (Bodine et al., 2001; Mourkioti & Rosenthal, 2005; Naya & Olson, 1999). However, these relationships are not always resolved consistently across species or traits, and their relative contribution to variation in muscle quality in aquatic animals, especially crustaceans, remains incompletely understood. Therefore, antioxidant-related responses, collagen-related characteristics, and selected indicators of muscle development can be viewed as biologically relevant but distinct components for evaluating muscle-related responses in the present study.

Chlorogenic acid (CGA) is a plant-derived polyphenolic compound with antioxidant activity and reported effects on glucose and lipid metabolism, as well as other biological functions (Clifford et al., 2020; Yin et al., 2021; Zhao et al., 2023). In aquaculture species, dietary CGA supplementation has been associated with improvements in growth performance, antioxidant-related indices, immune responses, and some flesh-related traits in several fish and shrimp species, including grass carp (Li et al., 2014), white shrimp (Wang et al., 2015), koi carp (Bakhtiari et al., 2024), largemouth bass (Yin et al., 2021), blackspotted croaker (Zhang et al., 2024), goldfish (Ahmadifar et al., 2026), rainbow trout (Ghafarifarsani et al., 2023), and atlantic salmon (Kühn et al., 2017). Nevertheless, the reported effects of CGA are not fully consistent across species, dietary inclusion levels, and experimental conditions, and many available studies focus primarily on growth, antioxidant capacity, or immune function rather than on muscle-quality-related outcomes as an integrated set of traits. In addition, evidence remains limited in crustaceans, particularly in P. clarkii. As a result, it remains unclear whether dietary CGA supplementation in this species is associated not only with growth-related responses but also with concurrent changes in muscle composition, physicochemical properties, collagen-related traits, antioxidant-related measurements, and muscle-growth-related gene expression.

Based on these considerations, we hypothesised that dietary CGA supplementation would be associated with changes in growth performance and selected muscle-related traits in P. clarkii, particularly those related to nutritional composition, physicochemical characteristics, antioxidant-related indices, and muscle-growth-related gene expression. To test this hypothesis, we evaluated the effects of graded dietary CGA levels on growth performance, muscle nutritional composition, physicochemical properties, antioxidant-related parameters, and gene expression profiles in P. clarkii. This study was designed to clarify the relationship between dietary CGA supplementation and muscle-related responses in this species under the present experimental conditions, and to provide a basis for further evaluation of CGA as a dietary supplement in P. clarkii culture.

2. Materials and methods

2.1. Ethics statement

All experimental procedures involving P. clarkii in this study, including animal rearing, handling, sampling, and tissue collection, were reviewed and approved by the Fisheries College of Guangdong Ocean University under approval number GDOU-IACUC-2024-A1055. During the feeding trial, P. clarkii was maintained under controlled water-quality conditions to minimize environmental stress. Handling and sampling were performed as gently and rapidly as possible to reduce distress. Before dissection and tissue collection, P. clarkii was anesthetized on ice and humanely euthanized in accordance with institutional animal welfare guidelines. Death was confirmed before hepatopancreatic and intestinal tissues were collected. After the experiment, all remaining animals and biological waste were disposed of in accordance with institutional biosafety regulations.

2.2. Diets, animals and feeding trial

Experimental crayfish used in this study were obtained from local farms in Zhanjiang. After a period of temporary rearing, a total of 450 healthy crayfish with intact appendages and similar body size were randomly selected and distributed into 15 ponds (1 m × 1.5 m × 1 m), with 30 crayfish in each pond. The 15 ponds were randomly assigned to five dietary treatments (CGA0, CGA200, CGA400, CGA600, and CGA800; CGA, 98% purity, Shaanxi Tianxingjian Biotechnology, Shananxi, China), with three replicate ponds per treatment. For all analyses, the pond was considered the experimental unit. The experimental diets were formulated using fish meal, soybean meal, peanut meal, rapeseed meal, and flour as the main ingredients, and the basic feed formula was detailed in Table S1.

The experiment was conducted in Zhanjiang over a total period of six weeks. Experimental diets were provided twice daily at 07:00 and 18:00. The initial feeding allowance was set at 3–5% of crayfish body weight, and subsequent feeding was adjusted according to the actual feeding activity in each pond and daily weather conditions. Uneaten feed was checked 2 h after each meal, collected, and weighed, and the actual feed intake of each pond was recorded on a meal-by-meal basis. Water quality was monitored throughout the experiment and maintained within the following ranges: water temperature, approximately 23 ± 2 °C; salinity, 0.5–1.0 g/L; ammonia nitrogen, < 0.05 mg/L; dissolved oxygen, > 4 mg/L; and pH, 7.2–7.5 mg/kg. Two-thirds of the water volume was renewed daily to maintain stable culture conditions. To minimize the influence of environmental variation on the experimental results, all ponds were managed under the same feeding schedule, water exchange regime, and water quality control conditions throughout the trial.

2.3. Computation of growth performance parameters

The number of surviving crayfish in each pond was recorded daily throughout the experiment. After the feeding trial, P. clarkii was fasted for 24 h, and the final body weight was determined as the total body weight of crayfish in each pond at the end of the trial. Based on the initial and final data, the following growth performance indices were calculated using the methods described below:

$$\mathit{Survival}\mathit{Rates}\left(\mathit{SR},\%\right)=\frac{\mathit{survival}\mathit{number}}{\mathit{initial}\mathit{number}}\times 100$$ (1)

$$\mathit{Weight}\mathit{Gain}\mathit{Rate}\left(\mathit{WGR},\%\right)=\frac{\left(W2-W1\right)}{W1}\times 100$$ (2)

$$\mathit{Feed}C\text{onversion}R\text{atios}\left(\mathrm{FCR}\right)=\frac{F}{\left(W2-W1\right)}$$ (3)

$$\mathit{Specific}\mathit{Growth}R\mathrm{ate}\left(\mathit{SGR},\%d-1\right)=\frac{\left(\ln W2-\ln W1\right)}{t}\times 100$$ (4)

In the equation: W1 and W2 represent the initial body weight (IBW, g) and final body weight (FBW, g). t represents the experiment days. F represents the actual feed intake recorded for each pond during the experiment (g).

2.4. Sample collection

At the end of the feeding trial, crayfish were randomly sampled from each pond. Five crayfish from each pond were used for whole-body proximate composition analysis, and the collected samples were temporarily stored at 4 °C prior to analysis. In addition, 12 crayfish from each pond were randomly collected for muscle-related analyses. Muscle tissues from these crayfish were used for antioxidant enzyme activity assays, gene expression analysis, texture profile analysis, and water-holding capacity (WHC) determination. After removal of the portions used for texture profile analysis and WHC determination, the remaining muscle tissues from the same pond were pooled and divided into aliquots for antioxidant enzyme activity assays and gene expression analysis, with three aliquots prepared for each analysis per pond. Samples used for antioxidant enzyme activity assays and gene expression analysis were placed in sterile cryogenic vials, immediately frozen in liquid nitrogen, and then stored at −80 °C until analysis. In addition, muscle tissues from another five crayfish randomly collected from each pond were used for amino acid and fatty acid analyses. These samples were freeze-dried prior to measurement. For statistical analysis, the pond was considered the experimental unit. Moulting crayfish were not used for sample collection.

2.5. Water-holding capacity and textural properties determination

Muscle samples were collected from the abdominal muscle of crayfish, using the same anatomical location for all individuals. After removal of the shell, surrounding connective tissue, and visible surface moisture, muscle samples were trimmed to a uniform size before analysis. All dissections were performed following the same procedure to ensure consistency among samples. WHC measurement was conducted in triplicate for each biological replicate. WHC was determined according to the method described by Sun et al. (2019). One crayfish muscle sample was processed separately for heat treatment and freezing treatment. For the heat-treated sample, crayfish muscle (W₁) was wrapped in cheesecloth and heated in a thermostatic water bath at 90 °C for 5 min, then cooled to room temperature, blotted dry to remove surface moisture, and weighed (W₂). For the freeze-treated sample, crayfish muscle (W₁) was frozen at −20 °C for 24 h, thawed at room temperature, blotted dry, and weighed (W₂). The water loss rate was calculated using the following formula:

$$W\text{ater}L\mathrm{oss}\mathit{Rate}\left(\%\right)=\frac{\left(W1-W2\right)}{W1}\times 100$$ (5)

Muscle texture characteristics (including hardness, springiness, cohesiveness, gumminess, chewiness, and resilience) were determined using a TA.XTC-18 texture analyzer (Bosin Technology, Shanghai, China). Muscle samples were collected from the same anatomical region of the abdominal muscle and trimmed to a uniform size (approximately 1.0 × 1.0 × 0.5 cm) before texture analysis. The orientation of the muscle fibers was kept consistent during testing. Texture analysis was conducted immediately after sample collection at room. The test parameters were set as follows: cylindrical probe diameter, 25 mm; contact sensitivity, 5 gf; loading rate: 1 mm/s; compression strain: 50%. The program was set to maintain pressure for 2 s, with a 5 s interval between successive measurements.

2.6. Proximate composition determination

Moisture, crude protein, crude lipid, and crude ash contents were determined according to AOAC (AOAC, 2005) standard procedures. Moisture content was determined by oven drying at 105 °C to constant weight and was expressed on a wet-weight basis. Crude protein was determined in the dried samples by the Kjeldahl method (Hanon Advanced Technology Group Co., Ltd., Jinan, China) using a nitrogen-to-protein conversion factor of 6.25 and was expressed on a dry-weight basis. Crude lipid was determined by the Soxhlet extraction method, and crude ash was determined by combustion in a muffle furnace at 550 °C (Hefei Kejing Materials Technology Co., Ltd., Hefei, China). All proximate composition analyses were performed in triplicate for each biological replicate. Hydroxyproline was determined using a commercial assay kit (Nanjing Jiancheng Biotechnology Research Institute, Nanjing, China). According to the manufacturer, the kit is applicable to animal tissues. Collagen content was estimated from hydroxyproline content using the conversion factor provided in the kit instructions.

2.7. Amino acid determination

Amino acid analysis was conducted by Sichuan Will Test Co., Ltd. (CNAS, CMA, and CATL accredited; CNAS registration No. L3969) according to GB 5009.124–2016. A certified mixed amino acid standard solution was used for calibration. As part of routine quality control, reagent blanks, calibration standards, parallel sample determinations， quality-control sample analysis, and spike-recovery assessments were included in the analytical workflow. For amino acid determination, muscle samples were homogenized and analyzed following acid hydrolysis, and the final amino acid contents were expressed on a dry-matter basis using the measured moisture content of the samples. Briefly, 0.03–0.05 g of sample was hydrolyzed with 15 mL of 6 mol/L hydrochloric acid containing 3–4 drops of phenol in a sealed hydrolysis tube after vacuuming and nitrogen flushing, and the hydrolysis was conducted at 110 ± 1 °C for 22 h. After hydrolysis, the solution was cooled, filtered, diluted to 50 mL, and a 1.0 mL aliquot was evaporated to dryness. The residue was washed twice with water, dried to remove residual acid, dissolved in 1.0 mL sodium citrate buffer (pH 2.2), filtered through a 0.22 μm membrane, and subjected to amino acid analysis. Amino acids were quantified using a HITACHI L-8900 high-speed amino acid analyzer (Hitachi, Tokyo, Japan). The analytical conditions were as follows: sulfonic acid-type cation exchange resin column; column temperature, 60 °C; flow rate, 0.4 mL/min; post-column derivatization temperature, 135 °C; detection wavelengths, 440 nm for proline and 570 nm for the other amino acids; injection volume, 20 μL. A commercial certified mixed amino acid standard solution (0.1 mmol/L) was used for external single-point calibration.

The amino acid profile reported in this study represents the 16 amino acids quantified under routine acid hydrolysis conditions. Tryptophan was not determined, because its analysis requires a separate alkaline hydrolysis procedure. Methionine was quantified in the present study, whereas a dedicated oxidative hydrolysis procedure was not applied for sulfur amino acids. Therefore, the reported values should be interpreted as quantitative results obtained under the standard acid-hydrolysis protocol.

2.8. Fatty acid determination

Fatty acid analysis was conducted by Sichuan Will Test Co., Ltd. (CNAS, CMA, and CATL accredited; CNAS registration No. L3969) according to GB 5009.168–2016. Tridecanoic acid was used as the internal standard, and peak identification was performed using a certified mixed fatty acid standard solution. As part of routine quality control, reagent blanks, calibration standards, parallel sample determinations, quality-control sample analysis, and spike-recovery assessments were included in the analytical workflow to monitor instrument performance, peak identification, and analytical precision. The specific procedures were as follows. Fresh muscle samples were first homogenized into a fine powder. An accurately weighed 5.00 ± 0.25 g of sample, containing 100–200 mg of fat, was transferred to a 250 mL flat-bottom flask. Subsequently, 100 mg of pyrogallic acid, boiling chips, and 2 mL of 95% ethanol were added and mixed thoroughly, followed by the addition of 10 mL of 8.3 mol/L hydrochloric acid. The mixture was hydrolyzed in a water bath at 70–80 °C for 40 min. After cooling, 10 mL of 95% ethanol was added. The hydrolysate was then transferred to a separatory funnel. The flask was rinsed with 50 mL of diethyl ether-petroleum ether mixture (1:1), and the rinse solution was combined with the hydrolysate in the separatory funnel. After shaking for 5 min and standing for 10 min, the upper organic and the specific procedures were as follows. Fresh muscle samples were first homogenized into a fine powder. An accurately weighed 5.00 ± 0.25 g of sample, containing 100–200 mg of fat, was transferred to a 250 mL flat-bottom flask. Subsequently, 100 mg of pyrogallic acid, boiling chips, and 2 mL of 95% ethanol were added and mixed thoroughly, followed by the addition of 10 mL of 8.3 mol/L hydrochloric acid. The mixture was hydrolyzed in a water bath at 70–80 °C for 40 min. After cooling, 10 mL of 95% ethanol was added. The hydrolysate was then transferred to a separatory funnel. The flask was rinsed with 50 mL of diethyl ether-petroleum ether mixture (1:1), and the rinse solution was combined with the hydrolysate in the separatory funnel. After shaking for 5 min and standing for 10 min, the upper organic phase was collected. This extraction process was repeated three times, and all extracts were pooled and evaporated to dryness to obtain the fat extract. After that, 8 mL of 2% sodium hydroxide-methanol solution was added to the fat extract, and the mixture was refluxed at 80 °C ± 1 °C until the oil droplets disappeared. Thereafter, 7 mL of 15% boron trifluoride-methanol solution was added, and refluxing was continued for 2 min. After cooling, 10–30 mL of n-heptane and saturated sodium chloride solution were added, followed by shaking to allow phase separation. A 5 mL aliquot of the upper n-heptane phase was collected, mixed with anhydrous sodium sulfate for dehydration over 5 min, and finally filtered through a 0.22 μm membrane filter prior to analysis.

Fatty acid methyl esters were analyzed using an Agilent 7890 A gas chromatograph (Agilent Technologies, Santa Clara, CA, USA) equipped with a DB-FastFAME capillary column (90 m × 0.25 mm × 0.25 μm) and a flame ionization detector (FID). Split injection was performed with a split ratio of 20:1. The carrier gas was nitrogen at a constant pressure of 44 psi. The hydrogen flow rate was 30 mL/min, the air flow rate was 300 mL/min, and the makeup nitrogen flow rate was 25 mL/min. The injector and detector temperatures were both set at 250 °C. The oven temperature program was as follows: initial temperature at 75 °C for 1 min, increased to 200 °C at 35 °C/min and held for 14 min, then increased to 210 °C at 2.5 °C/min and held for 5 min, and finally increased to 230 °C at 12 °C/min and held for 20 min. Tridecanoic acid was used as the internal standard. Peaks were identified by comparison of retention times with those of a commercial certified mixed standard solution. The original laboratory data were expressed as g/100 g dry matter and were converted to mg/g dry matter for data presentation and statistical analysis in this study.

2.9. Determination of antioxidative metabolites and antioxidant enzymes

The activities of total antioxidant capacity (T-AOC), superoxide dismutase (SOD), and catalase (CAT), as well as the contents of malondialdehyde (MDA) and reactive oxygen species (ROS), were determined using commercial assay kits purchased from Shanghai Enzyme-linked Biotechnology Co., Ltd. (China), which are suitable for animal tissues. All assays were performed under standardised conditions according to the manufacturers' instructions. During sample preparation and analysis, all procedures were carried out on ice, a fixed homogenisation ratio was maintained, and repeated freeze-thaw cycles were avoided. Routine quality-control procedures included reagent blanks, standard curves, triplicate sample measurements, and batch-wise quality-control samples to monitor assay stability and repeatability. Soluble protein concentration in the same extract was quantified by the BCA method, and all antioxidant indices were normalised to protein content. The assay wavelengths for T-AOC, SOD, CAT, MDA, and ROS were 405, 550, 240, 532, and 525 nm, respectively. Results were expressed as U/g protein for enzyme activities and mmol/mg protein for metabolite levels. Each biological sample was analyzed in triplicate.

2.10. Gene expression analysis

To further investigate the antioxidant status, muscle development and protein synthesis in P. clarkii, the expression of genes related to antioxidant defence (SOD, CAT, GPX, MnSOD, Nrf2, and Keap1), muscle development (MEF2a and MEF2b), protein synthesis/collagen-related regulation (IGF-1, AKT, mTOR, S6K, and 4E-BP1) and muscle texture (LOX) were evaluated by real-time quantitative PCR (qPCR). Total RNA isolation from P. clarkii muscle was performed with Trizol reagent (Beijing TransGen Biotechnology Co., Ltd., Beijing, China). RNA concentration was measured using a Thermo Nanodrop 2000 (Thermo Fisher Scientific, Waltham, USA). Reverse transcription of total RNA into cDNA was carried out using the Evo M-MLV RT Kit with gDNA Clean for qPCR (including gDNA Wiper Kit, ACCURATE BIOTECHNOLOGY(HUNAN)CO.,LTD, ChangSha, China). The target gene sequences were retrieved from the NCBI database, and specific primers were designed based on these sequences using Primer 5 software. The primer sequences, amplicon sizes, and amplification efficiencies are listed in Table S2. All primers were synthesized by Shanghai Sangon Biotech Co., Ltd., Shanghai, China, and the amplification efficiency of all primers ranged from 0.93 to 1.05. Melt-curve analysis and standard curves are presented in Fig. S1 and Fig. S2, respectively, to further support amplification specificity and primer performance. QPCR was carried out with a Roche LightCycler 480 system (Roche Diagnostics, Basel, Switzerland) to quantify the gene expression. Each reaction was carried out in a total volume of 10 μL, containing 1 μL cDNA, 5 μL 2× SYBR Green mix, 0.4 μL of each primer, and 3.2 μL nuclease-free water. The qPCR reactions followed a standard two-step protocol. The reaction program was composed of an initial denaturation step maintained at 95 °C for 30 s, succeeded by 40 cycles of amplification at 95 °C for 10 s and the combined annealing and extension step at 60 °C for 30 s. Each sample was analyzed in technical triplicate. 18S rRNA was used as the reference gene for normalization (Xu, 2025). Relative gene expression levels were calculated using the 2^{^-ΔΔCt} method.

2.11. Data statistics and analysis

Data are presented as mean ± SD (n = 3). Standard deviations are reported with one or two significant figures as appropriate, and means are rounded to the same decimal place as the corresponding SD. For all analyses, the pond was considered the experimental unit. For each response variable, measurements obtained from crayfish sampled within the same pond were averaged to generate a single pond-level value, and these pond means were used for one-way ANOVA in SPSS 26.0. Thus, each dietary treatment included three independent pond replicates (n = 3). Prior to one-way ANOVA, normality and homogeneity of variance were assessed using the Shapiro–Wilk test and Levene's test, respectively. When significant differences were detected in the analysis (P < 0.05), Tukey's test was carried out. Pearson correlation analysis was performed using pond-level values. The variables included growth performance, muscle quality traits, nutritional composition indices, antioxidant-related parameters, and relative gene expression levels. Significant correlations were identified after FDR correction.

3. Results

3.1. Growth performance

Dietary CGA significantly affected the growth performance of P. clarkii, whereas survival rate (SR) remained comparable among all groups (P > 0.05, Table S3). Compared with the control group, the FBW and WGR values were significantly increased when dietary CGA supplementation reached 400 mg/kg or above (P < 0.05). Notably, the CGA600 and CGA800 groups showed the highest numerical values for growth-related parameters, but no significant difference was detected between these two treatments under the present experimental conditions. The SGR of P. clarkii significantly increased in all the CGA added groups relative to the control group. The FCR of P. clarkii decreased progressively with increasing CGA inclusion, and the CGA600 and CGA800 groups showed significantly lower values than the CGA0, CGA200, and CGA400 groups (P < 0.05).

3.2. Muscle texture

As shown in Fig. 1A, the muscle hardness and resilience of P. clarkii in the CGA400 and CGA600 groups were significantly higher than those in the control group (P < 0.05). For springiness, a significant increase was detected only in the CGA600 group compared to the control group (P < 0.05). Similarly, muscle gumminess was significantly increased in the CGA600 group. Although numerical differences were observed, these differences were not statistically significant between the other CGA-treated groups and the control group (P > 0.05). In addition, chewiness was significantly higher only in the CGA400 group than in the control group (P < 0.05).

Fig. 1.

The effect of chlorogenic acid on muscle texture (A) and water-holding capacity (B). Note: (A) Radar chart showing the overall variation in hardness, springiness, cohesiveness, gumminess, chewiness, and resilience among the dietary treatments. (B) Box plots showing cooking loss and freezing loss in each treatment group. Different lowercase letters indicate significant differences among groups (P < 0.05).

There was no significantly difference in cooking loss rate between the CGA-treated groups and the control group (P > 0.05). In contrast, the freezing loss rate in all CGA-treated groups was significantly lower than that in the control group (P < 0.05, Fig. 1B).

3.3. Muscle proximate components

As shown in Table 1, the CGA600 group had significantly higher muscle crude protein contents than the control group (P < 0.05). Compared with the control group, the CGA400, CGA600, and CGA800 groups significantly increased muscle hydroxyproline and collagen contents in P. clarkii (P < 0.05). Although the numerical values continued to rise slightly from CGA400 to CGA800, no statistically significant differences were detected among these three higher-dose groups (P > 0.05).

Table 1.

Muscle proximate composition of P. clarkii.

Indexes	Groups
	CGA0	CGA200	CGA400	CGA600	CGA800
Moisture (%, wet weight)	75.58 ± 0.47ab	75.71 ± 0.27ab	76.41 ± 0.51a	76.81 ± 0.55a	74.87 ± 0.67b
Crude fat (%, dry matter)	4.73 ± 0.54	4.71 ± 0.22	4.54 ± 0.80	4.37 ± 0.28	4.59 ± 0.33
Crude protein (%, dry matter)	80.67 ± 0.19b	80.13 ± 0.12b	80.71 ± 0.39b	81.58 ± 0.12a	80.21 ± 0.11b
HYP (μg/mg)	0.11 ± 0.01c	0.15 ± 0.03bc	0.20 ± 0.01ab	0.21 ± 0.01a	0.22 ± 0.02a
Collagen (μg/mg)	0.89 ± 0.08c	1.23 ± 0.26bc	1.62 ± 0.05ab	1.66 ± 0.08a	1.75 ± 0.20a

Note: Data are presented as mean ± SD (n = 3). Standard deviations are reported with one or two significant figures as appropriate, and means are rounded to the same decimal place as the corresponding SD. Those labeled with different lowercase letters indicate significant differences between groups (p < 0.05), and those labeled with the same lowercase letters indicate no significant differences between groups (p > 0.05) the same below.

3.4. Muscle amino acid composition

Relative to the control group, the CGA400 group had significantly elevated concentrations of leucine, arginine, aspartic acid, and tyrosine (P < 0.05, Table 2), while the CGA600 group showed a significant increase in the concentrations of aspartic acid, and alanine (P < 0.05). Regarding total amino acids (TAA), the content first increased and then declined as the CGA concentration increased, peaking in the CGA400 group. In contrast, TAA accumulation was significantly reduced in the CGA800 group (P < 0.05). A similar pattern was observed for TEAA, for which only the CGA400 group showed a significant increase relative to the control group (P < 0.05), whereas the other CGA-supplemented groups did not differ significantly from the control (P > 0.05). In addition, the total contents of umami amino acids and sweet amino acids tended to be higher in the CGA400 and CGA600 groups; however, these increases should be interpreted as numerical trends rather than statistically confirmed differences where significance was not detected.

Table 2.

Effects of different CGA levels on the Amino Acid content of muscle in P. clarkii (g/100 g, dry matter).

Amino Acids	Groups
	CGA0	CGA200	CGA400	CGA600	CGA800
Thr	3.07 ± 0.01	3.10 ± 0.01	3.19 ± 0.09	3.05 ± 0.06	3.02 ± 0.12
Val	3.40 ± 0.02	3.33 ± 0.01	3.55 ± 0.18	3.40 ± 0.04	3.31 ± 0.11
Met	1.95 ± 0.03	2.00 ± 0.01	2.06 ± 0.07	1.99 ± 0.05	1.94 ± 0.09
Ile	3.60 ± 0.13	3.44 ± 0.07	3.72 ± 0.21	3.47 ± 0.12	3.50 ± 0.13
Leu	6.31 ± 0.02b	6.34 ± 0.06b	6.63 ± 0.14a	6.28 ± 0.14b	6.22 ± 0.22b
Phe	3.36 ± 0.10ab	3.30 ± 0.05ab	3.48 ± 0.10a	3.28 ± 0.06ab	3.22 ± 0.07b
His	1.94 ± 0.09a	1.90 ± 0.04a	1.91 ± 0.07a	1.80 ± 0.04ab	1.72 ± 0.06b
Lys	6.70 ± 0.15	6.64 ± 0.01	6.91 ± 0.28	6.58 ± 0.11	6.45 ± 0.22
Arg	8.57 ± 0.14b	8.55 ± 0.07b	8.94 ± 0.19a	8.63 ± 0.12ab	8.43 ± 0.26b
TEAA	38.90 ± 0.60b	38.60 ± 0.10b	40.39 ± 1.27a	38.47 ± 0.65b	37.81 ± 1.24b
Asp	8.46 ± 0.04b	8.62 ± 0.04ab	8.82 ± 0.26a	8.89 ± 0.03a	8.30 ± 0.35b
Glu	12.88 ± 0.02	13.01 ± 0.03	13.44 ± 0.43	13.30 ± 0.32	12.71 ± 0.56
Gly	4.47 ± 0.06a	4.33 ± 0.04ab	4.38 ± 0.09ab	4.64 ± 0.15a	4.09 ± 0.19b
Ala	4.23 ± 0.06b	4.19 ± 0.01b	4.38 ± 0.10ab	4.50 ± 0.18a	4.17 ± 0.17b
Ser	3.00 ± 0.11	3.19 ± 0.02	3.24 ± 0.09	3.21 ± 0.05	3.07 ± 0.14
Tyr	3.20 ± 0.14b	3.18 ± 0.06b	3.41 ± 0.08a	3.21 ± 0.06b	3.15 ± 0.04b
Pro	2.22 ± 0.01ab	2.35 ± 0.02a	2.20 ± 0.10ab	2.19 ± 0.08ab	2.05 ± 0.08b
TNEAA	38.45 ± 0.22ab	38.87 ± 0.17ab	39.85 ± 1.00a	39.95 ± 0.41a	37.54 ± 1.48b
TAA	77.35 ± 0.41b	77.47 ± 0.27b	80.24 ± 2.23a	78.42 ± 0.50b	75.35 ± 2.71b
DAA	36.59 ± 0.09ab	36.63 ± 0.17ab	37.89 ± 0.94a	37.82 ± 0.34a	35.64 ± 1.37b
UAA	21.34 ± 0.02	21.63 ± 0.07	22.26 ± 0.68	22.19 ± 0.29	21.01 ± 0.91
SAA	15.25 ± 0.09ab	15.00 ± 0.13ab	15.64 ± 0.32a	15.63 ± 0.07a	14.63 ± 0.46b
EAA/TAA	0.5029 ± 0.0052	0.4983 ± 0.0005	0.5033 ± 0.0032	0.4906 ± 0.0061	0.5018 ± 0.0022
DAA/TAA	0.4730 ± 0.002b	0.4729 ± 0.0005b	0.4723 ± 0.0014b	0.4823 ± 0.0058a	0.4730 ± 0.0011b

Note: UAA (umami amino acids): Asp and Glu; SAA (sweet amino acids): Gly, Ala Tyr, and Phe.

3.5. Muscle fatty acid composition

As shown in Table 3, dietary CGA supplementation significantly affected the fatty acid profile of muscle in P. clarkii. Compared with the control group, the total saturated fatty acid (SFA) content was significantly reduced in the CGA400 and CGA600 groups, with the lowest value observed in the CGA600 group (P < 0.05). A similar pattern was found for total monounsaturated fatty acids (MUFAs) and polyunsaturated fatty acids (PUFAs), both of which were significantly decreased in the CGA400 and CGA600 groups relative to the control group (P < 0.05), whereas no significant differences were detected in the CGA200 and CGA800 groups (P > 0.05). For PUFAs, C20:2, C18:3n3, C20:5n3, and C22:6n3 were significantly reduced in the CGA400 and/or CGA600 groups (P < 0.05). Consistently, both total n-3 PUFAs and n-6 PUFAs were significantly lower in the CGA400 and CGA600 groups than in the control group (P < 0.05), and the n-3/n-6 ratio was significantly decreased only in the CGA400 group (P < 0.05).

Table 3.

Effects of different CGA levels on the Fatty Acid content of muscle in P. clarkii (mg/g, dry matter).

Indexes	Groups
	CGA0	CGA200	CGA400	CGA600	CGA800
C14:0	0.0568 ± 0.0032b	0.0599 ± 0.0003b	0.0904 ± 0.0274a	0.0505 ± 0.0068b	0.0485 ± 0.0038b
C15:0	0.0631 ± 0.0031a	0.0587 ± 0.0022ab	0.0524 ± 0.0024ab	0.0517 ± 0.0049b	0.0557 ± 0.0067ab
C16:0	1.10 ± 0.07a	1.00 ± 0.03a	0.97 ± 0.06ab	0.84 ± 0.06b	1.07 ± 0.04a
C17:0	0.08 ± 0.01	0.06 ± 0.01	0.06 ± 0.01	0.05 ± 0.01	0.07 ± 0.01
C18:0	0.60 ± 0.03a	0.52 ± 0.02ab	0.48 ± 0.03b	0.45 ± 0.03b	0.59 ± 0.03a
C20:0	0.0583 ± 0.0060	0.0620 ± 0.0080	0.0579 ± 0.0043	0.0458 ± 0.0034	0.0643 ± 0.0135
SFAs	1.95 ± 0.12a	1.76 ± 0.05ab	1.71 ± 0.06b	1.50 ± 0.09c	1.90 ± 0.05ab
C16:1n7	0.20 ± 0.02ab	0.18 ± 0.01bc	0.22 ± 0.02a	0.15 ± 0.01c	0.17 ± 0.01bc
C18:1n9c	1.32 ± 0.11a	1.13 ± 0.05ab	1.03 ± 0.08b	0.96 ± 0.08b	1.31 ± 0.08a
C22:1n9	0.04 ± 0.01b	0.05 ± 0.01b	0.04 ± 0.01b	0.09 ± 0.03a	0.07 ± 0.01ab
MUFAs	1.56 ± 0.13a	1.37 ± 0.06ab	1.29 ± 0.09b	1.20 ± 0.08b	1.55 ± 0.08a
C20:2	0.0721 ± 0.0043a	0.0478 ± 0.0011b	0.0476 ± 0.0156b	0.0395 ± 0.0042b	0.0628 ± 0.0121ab
C18:3n3	0.39 ± 0.03a	0.34 ± 0.02ab	0.29 ± 0.02b	0.29 ± 0.02b	0.36 ± 0.03a
C20:3n3	0.0396 ± 0.0128	0.0247 ± 0.0030	0.0220 ± 0.0019	0.0211 ± 0.0041	0.0248 ± 0.0068
C20:5n3	1.60 ± 0.12a	1.39 ± 0.07a	1.04 ± 0.02b	1.17 ± 0.02b	1.47 ± 0.11a
C22:6n3	0.3504 ± 0.0231a	0.3289 ± 0.0144a	0.2369 ± 0.0032b	0.2683 ± 0.0226b	0.3332 ± 0.0314a
n-3PUFAs	2.38 ± 0.18a	2.08 ± 0.09a	1.59 ± 0.01b	1.75 ± 0.06b	2.19 ± 0.18a
C18:2n6c	1.07 ± 0.08a	0.97 ± 0.04a	0.92 ± 0.07ab	0.81 ± 0.05b	1.02 ± 0.06a
C20:4n6	0.71 ± 0.05a	0.62 ± 0.03ab	0.56 ± 0.01bc	0.49 ± 0.04c	0.65 ± 0.06ab
n-6PUFAs	1.78 ± 0.12a	1.60 ± 0.07ab	1.47 ± 0.08bc	1.30 ± 0.05c	1.67 ± 0.12ab
PUFAs	4.23 ± 0.30a	3.72 ± 0.17a	3.11 ± 0.08b	3.09 ± 0.08b	3.92 ± 0.31a
n-3/n-6	1.333 ± 0.018a	1.303 ± 0.004a	1.079 ± 0.059b	1.345 ± 0.067a	1.305 ± 0.019a

3.6. Antioxidant capacity in the muscle

The T-AOC values in the muscle of P. clarkii was significantly elevated in the CGA400 and CGA600 groups, whereas no significant discrepancies were found between the other CGA-treated groups and the control (P > 0.05). All CGA-treated groups showed significantly increased muscle CAT activity. The SOD activity was also significantly increased in the CGA-treated groups, with exception of the CGA400 group (P < 0.05). In contrast, MDA levels in the CGA-treated groups exhibited an overall decreasing trend, with the most notable decrease in the CGA800 group (P < 0.05). Similarly, the levels of ROS were significantly lower in the CGA400 and CGA800 groups relative to the control group (P < 0.05, Fig. 2).

Fig. 2.

The antioxidant capacity in the muscle of P. clarkii. Note: (A) Reactive oxygen species (ROS) and malondialdehyde (MDA) contents in muscle. (B) Total antioxidant capacity (T-AOC), superoxide dismutase (SOD), and catalase (CAT) activities in muscle. Different lowercase letters indicate significant differences among groups (P < 0.05).

Correspondingly, dietary CGA supplementation significantly affected the expression of antioxidant-related genes in the muscle of P. clarkii. The expression levels of SOD, CAT, GPX and Nrf2 genes were significantly upregulated in the CGA400 and CGA600 groups (Fig. 3). Although the expression levels in the CGA800 group showed a slight decline, they remained higher than those in the control group. MnSOD expression was higher than that in the control group, with a significant increase detected only in the CGA600 group. In contrast, the expression of Keap-1 was notably downregulated in all CGA-supplemented groups except the CGA200 group compared with the control group (P < 0.05).

Fig. 3.

Effects of CGA on the expression of genes related to muscle antioxidation.

3.7. Gene expression related to muscle development and protein synthesis

Dietary CGA supplementation also significantly affected the expression of muscle development and protein synthesis related genes. The mRNA expression levels of MEF2a and MEF2b increased progressively with increasing CGA inclusion in the diet. Both genes displayed significantly higher expression in the CGA400 and CGA600 groups, with the highest levels in the CGA600 group (P < 0.05). In the CGA800 group, MEF2a expression was also significantly higher than that in the control group. The expression levels of protein synthesis-related genes (including mTOR, IGF-1, AKT, S6K, and 4E-BP1) exhibited a trend of initial increase followed by subsequent decrease, among which the expression levels in the CGA400 and CGA600 groups were significantly up-regulated compared with the control group. As dietary CGA concentrations increased, the expression levels of LOX increased, with significantly upregulated expression in the CGA400 to CGA800 groups compared with the control group (P < 0.05, Fig. 4).

Fig. 4.

Effects of CGA on the expression of genes related to muscle protein synthesis.

3.8. The correlation analysis

To further examine the relationships among growth performance, muscle quality traits, nutritional composition, antioxidant-related indices, and gene expression variables within the present dataset, Pearson correlation analysis was performed using pond-level values. Correlation analysis in this study showed that collagen was significantly positively correlated with FBW, WGR, SGR, resilience, and hardness. Several amino acids (including Ala, Ser, Glu, Asp, Arg, His, Met, and Val) showed significant associations with muscle texture characteristics, among which the flavor-related amino acids Asp, Glu, Arg and Ser showed particularly strong correlations. At the same time, some fatty acids (including C15, C17, C18:3n-3, C20:3n-3, C20:5n-3, and C20:4n-6) exhibited a significant correlation with muscle water-holding capacity in this study. In this study, certain specific fatty acids, such as C22:1n-9, were significantly positively associated with the growth of P. clarkii; C16:1n-7 was notably positively correlated with FCR. Further analysis showed that muscle antioxidant status was closely associated with growth and texture traits. The activities of T-AOC, SOD, and CAT were all positively correlated with FBW, WGR, and SGR; among them, T-AOC was also positively linked with hardness, chewiness, and gumminess, while CAT was additionally positively correlated with Springiness and Cohesiveness. In contrast, MDA was significantly negatively correlated with the growth indicators FBW, WGR, and SGR, and both MDA and ROS were negatively correlated with muscle texture parameters (e.g., resilience and chewiness). The expression of several antioxidant-related genes was also associated with growth and muscle quality traits: SOD, GPX, MnSOD, and Nrf2 were markedly positively linked with growth indicators, while Keap1 was markedly negatively associated with them; The aforementioned antioxidant genes (except Keap1) were all significantly positively correlated with muscle texture characteristics, and Hardness exhibited the most prominent correlation with these genes. Similarly, the expression of several genes related to muscle development, protein synthesis, and collagen-related regulation was positively correlated with FBW, WGR, and SGR; among them, MEF2a, mTOR, IGF-1, and LOX were also positively correlated with resilience and hardness. Overall, these results indicate close associations among collagen deposition, antioxidant status, muscle-related gene expression, and phenotypic traits (Fig. 5). Because the correlation analysis was conducted across multiple pond-level variables in a relatively small dataset, these associations should be interpreted cautiously as descriptive relationships within the present study rather than as evidence of mechanism.

Fig. 5.

Correlation analysis between muscle nutrients and various test indicators.

4. Discussion

As a polyphenolic compound, CGA is known to exert multiple biological activities in animals, including antioxidant, antimicrobial, and anti-inflammatory activities, as well as lipid metabolism regulation (Wang et al., 2022). Notably, existing studies have demonstrated that CGA promotes growth and improves muscle quality in various farmed aquatic animals (Wang et al., 2022; Yang et al., 2020; Wen et al., 2010). However, such effects have not been fully investigated or elucidated in P. clarkii. Therefore, the present study was conducted to evaluate the effects of different dietary CGA levels on the growth performance and muscle quality of P. clarkii.

In the present study, all CGA-treated groups showed significantly increased WGR and SGR in P. clarkii. Except for the CGA200 group, the FCR was significantly reduced in the other CGA-treated groups. Similar results have been reported in grass carp, in which dietary CGA at 400 mg/kg increased WGR and decreased FCR (Yang et al., 2020). CGA has also been shown to promote growth in Yellow River carp (Fu et al., 2018), Chinese soft-shelled turtle (Wen et al., 2010), and Cyprinus carpio var. Jian (Zhang & Wen, 2012). However, some studies have reported that dietary CGA exerts no significant growth-promoting effect on Litopenaeus vannamei (Wang et al., 2015) and Cyprinus carpio (Xu et al., 2022). The inconsistencies may be attributable to differences in experimental duration, environmental conditions, dosage, and species. Therefore, further studies are needed to clarify the reasons for these inconsistent findings.

An increasing number of studies indicate that CGA can affect the muscle texture of aquatic animals. In blackspotted croaker, dietary CGA increased muscle hardness and chewiness (Zhang et al., 2024). Likewise, in grass carp, CGA supplementation also improved muscle hardness (Yang et al., 2020). In the present study, dietary supplementation with 400 and 600 mg/kg CGA significantly increased muscle hardness, and resilience. In addition, 400 mg/kg CGA significantly increased muscle chewiness, whereas 600 mg/kg CGA significantly increased springiness and gumminess. These findings suggest that dietary CGA at 400–600 mg/kg was associated with improved texture-related traits in P. clarkii. Muscle texture is closely linked to collagen-related properties (Moreno et al., 2012; Periago et al., 2005; Yang et al., 2020). As a major component of connective tissue, collagen contributes to muscle structure and is relevant to firmness and resilience (Astruc, 2014). In the present study, dietary CGA significantly increased muscle collagen content, consistent with previous findings in grass carp (Li et al., 2014) and Chinese soft-shelled turtle (Wen et al., 2010). Correlation analysis also showed positive associations between collagen content and several growth and texture-related traits. Suggesting that collagen-related changes were linked to the observed variation in muscle texture within the present dataset. However, although the CGA800 group had higher collagen content than the CGA400 and CGA600 groups, its hardness was lower. This suggests that collagen quantity alone may not fully explain the textural differences among groups. Previous studies have shown that muscle texture and quality are not only affected by collagen content but are also related to collagen maturity, solubility, and degree of cross-linking (McCormick, 1999; Purslow, 2014). Because these structural properties were not directly determined in the present study, the specific basis of the texture changes should not be overinterpreted. In addition to textural properties, WHC is another important indicator of muscle quality because it reflects the retention of water and soluble nutrients in muscle (Melody et al., 2004). Similar to the findings in grass carp (Yang et al., 2020), no significant difference in cooking loss rate was found among treatment groups. However, all CGA-treated groups showed a significant reduction in freezing loss rate. This result suggests that dietary CGA supplementation may help reduce freezing-associated loss from muscle tissue.

Amino acids are the fundamental components of proteins and are also involved in the regulation of various metabolic pathways. In recent years, increasing attention has been paid to the effects of amino acids and their derivatives on the muscle quality of aquatic animals (Hu et al., 2022; Jiang et al., 2021; Yun et al., 2022). In grass carp studies, dietary CGA at 400, 600, and 800 mg/kg markedly enhanced the levels of TEAA and TAA (Sun et al., 2017). Studies have also shown that CGA supplementation increases glycine concentrations in rats (Zheng et al., 2014). Amino acid metabolism is closely associated with the formation of flavor compounds (Xu et al., 2018). Previous studies have demonstrated that Eucommia ulmoides leaf extract significantly enhanced the contents of His, Pro, Phe, Ala, DAA, and TAA in the muscle of grass carp (Leng et al., 2008). In addition, research on Mylopharyngodon piceus indicated that CGA can elevate levels of Arg, Asp, and Glu in muscle tissue. In the present study, significant increases in Leu, Arg, Tyr, DAA, TAA and TEAA were observed in the CGA400 group, whereas significant increases in Asp, Ala, and DAA were observed in the CGA600 group. Correlation analysis further showed that Glu, Arg, Asp, and Ser were significantly positively correlated with muscle texture characteristics. Prior studies have demonstrated that Glu can improve muscle quality by regulating protein synthesis and deposition (Dong et al., 2023) or by modulating lipid metabolism and myogenic regulatory factors (Zhao et al., 2019). Similarly, an appropriate level of Arg can promote skeletal muscle growth and regulate muscle development (Chen et al., 2025a, Chen et al., 2025b; Wang et al., 2022). These results suggest that dietary CGA at 400–600 mg/kg may improve muscle flavor in P. clarkii by enhancing muscle protein and amino acid contents. Similar trends have also been reported in studies on grass carp (Li et al., 2014) and carp (Luo et al., 2002).

Oxidative stress has been shown to impair muscle quality in aquatic animals (Li et al., 2023; Song et al., 2022; Yang et al., 2020), whereas improved antioxidant capacity is generally associated with better muscle status. As a herbal additive, CGA is considered a relatively safe and health-promoting antioxidant (Zhang et al., 2022). Owing to its molecular structure containing phenolic hydroxyl groups, CGA can effectively scavenge reactive free radicals (Wang et al., 2022). Previous studies have also shown that CGA can reduce oxidative damage and enhance antioxidant-related responses in different animals, including channel catfish (Zhang et al., 2023), heat-stressed broilers (Zhao et al., 2019), largemouth bass (Xia et al., 2024), Atlantic salmon (Kühn et al., 2017), grass carp (Yang et al., 2024), Cyprinus carpio (Xu et al., 2022), and Carassius auratus (Jin et al., 2023). In the present study, dietary CGA reduced ROS and MDA levels in the muscle of P. clarkii and increased CAT, SOD and T-AOC activity relative to the control group, indicating that dietary CGA supplementation was associated with enhanced antioxidant-related responses in P. clarkii under the present experimental conditions. In addition, the correlation analysis showed that T-AOC, SOD, and CAT were positively associated with growth and texture traits, whereas MDA and ROS were negatively associated with these parameters, suggesting that improved antioxidant status may help preserve muscle structural integrity and thereby contribute to better muscle quality. Research on juvenile largemouth bass (Yin et al., 2021) and white shrimp (Wang et al., 2015) has further shown that dietary CGA can enhance the expression of antioxidant-related genes. Likewise, our findings indicated that dietary CGA supplementation markedly increased the expression of SOD, CAT, GPX and MnSOD in muscle, showing a pattern consistent with the corresponding antioxidant enzyme activities. Specifically, Nrf2 interacts with antioxidant response elements and induces downstream antioxidant-related genes (Ma, 2013), whereas Keap1 negatively regulates Nrf2 by promoting its degradation (Bellezza et al., 2018). In the present study, appropriate levels of CGA significantly reduced Keap-1 expression and increased Nrf2 expression in muscle. These results suggest that the antioxidant-related responses observed in this study may be related to changes in antioxidant regulatory genes, including Keap1 and Nrf2. However, because the present study assessed enzyme activities and mRNA levels only, the data do not provide direct functional evidence that this pathway was activated or that it causally mediated the improvement in muscle quality. The positive correlations between antioxidant-related genes and growth- and texture-related traits provide supportive evidence for an association between antioxidant-related responses and muscle-related phenotypes in the present dataset.

Muscle development and protein synthesis are important determinants of muscle texture, nutritional composition, and edible quality (Hua et al., 2025; Johnston et al., 2006, 2000). Previous studies have shown that IGF-1/Akt/mTOR-related signalling is associated with muscle growth and protein deposition in animals (Bodine et al., 2001; Brunet et al., 1999; Mourkioti & Rosenthal, 2005; O'Neill et al., 2015), while LOX and MEF2 are involved in connective tissue regulation and muscle differentiation, respectively (Grau-Bové et al., 2015; Naya & Olson, 1999; Snyder et al., 2013). In the present study, dietary supplementation with 400–800 mg/kg CGA remarkably enhanced the expression levels of key genes in the IGF-1/PI3K/Akt and mTOR pathways, increased LOX expression, and upregulated MEF2a and MEF2b transcription, with the highest responses generally observed around 600 mg/kg CGA. Similar increases in LOX expression have also been reported in grass carp (Yang et al., 2022). In addition, these genes were significantly positively correlated with growth indicators and muscle texture parameters, and some were also associated with muscle resilience and hardness. Taken together, these results suggest that dietary CGA was associated with transcriptional responses related to muscle development, protein synthesis, and collagen-related regulation in P. clarkii. These molecular changes were consistent with the improved growth performance, collagen content, and texture-related traits observed in the CGA400 and CGA600 groups. Based on the known roles of IGF-1/Akt/mTOR-related signalling, MEF2, and LOX in muscle growth and connective tissue regulation, one possible explanation is that CGA may have influenced muscle quality partly through transcriptional regulation of these processes. However, because only mRNA levels were determined in the present study, these findings should be regarded as supportive molecular evidence rather than direct proof of pathway activation or causal regulation of myofiber growth and collagen cross-linking.

Muscle quality is also closely associated with fatty acid composition, which not only influences the nutritional value of muscle but also plays a crucial role in regulating muscle flavor (Zhang et al., 2024). In the present study, dietary CGA reduced the contents of several PUFAs in muscle, including DHA, EPA, linoleic acid, and arachidonic acid, and also decreased the n-3/n-6 ratio in some treatment groups. This result differs from some previous reports in broilers (Zhao et al., 2019), Atlantic salmon (Kühn et al., 2017), and grass carp (Yang et al., 2024), in which CGA supplementation increased the levels of certain n-3 or n-6 PUFAs. Such discrepancies may reflect species-specific differences in lipid metabolism, dietary lipid background, and tissue-specific fatty acid deposition. These findings suggest that dietary CGA influenced muscle fatty acid deposition in P. clarkii. Based on previous studies, one possible explanation is that CGA may affect lipid metabolism by altering the balance between lipid synthesis and fatty acid oxidation (Bhandarkar et al., 2019; Shang et al., 2024; Uemura et al., 2011; Zhou et al., 2016). This change should be interpreted cautiously, because PUFAs such as DHA and EPA are important from a human nutritional perspective. Therefore, the reduction in muscle PUFAs cannot be regarded as unconditionally beneficial. Because PUFAs are highly susceptible to lipid peroxidation, muscles with higher PUFA levels are generally more vulnerable to oxidative deterioration (Krzysztof et al., 2017). In the present study, the reduction in muscle PUFA content occurred together with lower ROS and MDA levels and higher antioxidant enzyme activities, suggesting that dietary CGA may have improved muscle oxidative stability while simultaneously modulating fatty acid composition. From this perspective, the altered PUFA profile may reflect a trade-off between nutritional value and oxidative stability, rather than a uniformly beneficial change. In addition, the higher fat content of the control diet than of the CGA-treated diets may also have contributed to the difference in PUFA deposition (Yao et al., 2022). Nevertheless, because the present study did not directly determine lipid metabolism-related genes or enzymes, the mechanism underlying the reduction in muscle PUFAs remains inferential. Further studies are needed to clarify whether CGA regulates fatty acid deposition in P. clarkii through coordinated effects on lipid synthesis, β-oxidation, and oxidative protection.

It must be acknowledged that this study has certain limitations. The relatively small number of replicates may limit the stability of the statistical estimates and the generalizability of the results. The ANOVA-derived ŋ² values indicated that CGA exerted large effect sizes on multiple growth performance and muscle quality-related parameters, suggesting that treatment groups accounted for a substantial proportion of the variance in these traits. However, caution is warranted in interpreting these estimates. With only three replicates per group and relatively low within-group variation, the ŋ² values may be inflated. In other words, while the present results clearly support statistically significant and biologically meaningful treatment effects, the precise magnitude of these effects should be further validated under conditions with larger sample sizes. In addition, some antioxidant-related and gene-expression-related measurements were based on pooled muscle samples within ponds, which provided sufficient material for the assays but reduced individual-level resolution and prevented assessment of within-pond variability. Several biochemical measurements, including antioxidant-related indices and hydroxyproline, relied on commercial assay kits without independent validation for P. clarkii tissue. In addition, although 18S rRNA showed only minor Ct variation across the analyzed samples and had been used in our previous research on P. clarkii, its stability was not formally re-evaluated under the present experimental conditions. Therefore, the qPCR results should be interpreted with appropriate caution as supportive transcriptional evidence. Nevertheless, based on the current findings, the inclusion of chlorogenic acid in the diet can be considered effective in improving both growth performance and muscle quality in P. clarkii.

5. Summary and outlook

Under the present experimental conditions, dietary CGA supplementation, particularly at 400–600 mg/kg, was associated with improved growth performance and several muscle-related traits in P. clarkii, together with changes in antioxidant-related indices and gene expression. These findings provide initial evidence that CGA supplementation may influence growth and muscle-related responses in this species. However, these findings should be interpreted cautiously and confirmed by further studies with greater replication, individual-level sampling, and direct functional analyses. Future work is also needed to clarify the biological basis and long-term implications of CGA supplementation in P. clarkii.

CRediT authorship contribution statement

Qinglin Liu: Writing – original draft, Validation, Methodology, Investigation, Formal analysis, Conceptualization. Yongkang Feng: Methodology, Investigation. Sha Huang: Methodology, Investigation. Shengrong Guo: Methodology, Investigation. Lili Shi: Resources, Project administration, Conceptualization. Beiping Tan: Resources, Project administration, Conceptualization. Baogui Tang: Supervision, Resources, Project administration, Conceptualization. Shuang Zhang: Writing – review & editing, Supervision, Project administration, Funding acquisition.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Data availability

Data will be made available on request.