Kjeldahl catalyst tablets (free of Hg and Se), boric acid, sulfuric acid (≥ 98%), pepsin (P6887, ≥3,200 units/mg protein), pancreatin (P7545, 4 × USP specification), 6-hydroxy-2,5,7,8-tetramethylchromane-2-carboxylic acid (Trolox), 2′, 7′-dichlorodihydrofluorescein-diacetate (DCFH-DA) and 2,2′-azobis-(2-methylpropianomidine) dihydrochloride (AAPH), Dulbecco's Modified Eagles Medium (DMEM), penicillin/streptomycin, fetal bovine serum (FBS), horse serum, Hank' balanced salt solution (HBSS), D-glucose, cell proliferation reagent WST-1, Tris/HCl pH 7.4, sodium rthovanadate (Na₃VO₄), phenylmethanesulfonyl fluoride (PMSF), L-glutamine, aprotinin, pepstatin, the bicinchoninic acid (BCA) kit, and puromycin (MABE343 anti-puromycin, clone 12D10 mouse monoclonal) were purchased from Sigma-Aldrich (Dublin, Ireland). Acetic acid and sodium hydroxide (NaOH) were procured from Fisher Scientific (Dublin, Ireland). Hydrochloric acid (HCl) was from VWR (Dublin, Ireland). Trinitrobenzene sulfonic acid (TNBS) reagent was from Medical Supply Co Ltd. (Dublin, Ireland). The C2C12 (subclone C2/4) mouse adherent myoblast cell line was purchased from ATCC^® CRL1772, Manassas, VA (Lot number 60339292), USA. Serum-free and DMEM amino acid medium was from US Biological (Salem, MA, USA). Precision Plus ProteinTM Dual Color Standards (SM1811) and sodium pyruvate (GE Healthcare, Thermo-Fisher) were both purchased from Thermo Fisher (Dublin, Ireland). Human insulin growth factor-1 (IGF-1) (100-11) was from PEPROTECH (London, UK). SDS-PAGE precast gels (4–15%) were obtained from Accuscience Ltd (Dublin, Ireland). All primary antibodies for phosphor (p)mTOR, mTOR, p4-EBP1, 4-EBP1S6 ribosomal protein, pS6 ribosomal protein, and β-actin were from Cell Signaling (Bioke, Leiden, The Netherlands). Secondary antibodies, namely green rabbit (926-32211 IRDye 800 CV) and goat anti-rabbit IgG, were obtained from LI-COR Biosciences UK Ltd (Cambridge, UK).

BioMarine Ingredients Ireland Ltd (Lough Egish Food Park, Castleblayney, Co. Monaghan, Ireland) generated the SPH under proprietary conditions in a mode analogous to that previously reported (21).

A simulated gastrointestinal digest of SPH was generated using a modification of the method reported by Walsh et al. (22). A 2.0% (w/v) solution (on a protein equivalent basis) of SPH was changed to pH 2.0 (6 M HCL) and incubated at 37°C with pepsin at an E:S of 2.5% (w/w). Following 90 min incubation, the pH was adjusted to pH 7.0 (1 M NaOH), and pancreatin at an E:S of 1% (w/w) was added and incubated at 37°C for a further 150 min. The sample was then heated at 85°C for 15 min, then left to cool, freeze-dried, and stored at −20°C.

The protein equivalent content of the hydrolysate and GI digest was determined using the macro-Kjeldahl procedure (23) using a nitrogen-to-protein conversion factor of 5.892 (21). The total and free AA content of the samples was determined using an accredited external company. The AA score was calculated as the mg of the limiting essential AA(s) per g of sample protein divided by the mg of the same AA per g of the reference protein (21).

The DH of the samples was calculated by the TNBS method (21). Gel permeation high-performance liquid chromatography (GP-HPLC) and reversed-phase ultra-performance liquid chromatography (RP-UPLC) were used to determine the molecular mass distribution and peptide profiles of the SPH and its SGID-treated equivalent (24, 25).

The color [L^* (lightness), a^* (redness), and b^* (yellowness)] values of the SPH powder were measured using a Konica Minolta CR-400 Chroma Meter (Minolta Camera Co., Osaka, Japan) as described by Cermeno et al. (26).

The solubility of the SPH was determined at pH 2.0, 4.0, 6.0, 7.0, 8.0, 9.0, 10.0, and 12.0 in accordance with the method reported by Connolly et al. (27) with minor modifications. A 4.0% (w/v) solution (on a protein equivalent basis) of SPH was prepared in distilled water (dH₂O, 5 mL) and stirred gently at ambient temperature (Stuart-heat stir, SB162, Keison Products Chelmsford, England) for 20 min. Aliquots of this protein suspension (9 mL in each tube) were then adjusted to pH values between pH 2.0 and 12.0 using NaOH or HCl, as required, before the total volume was adjusted to 20 mL with dH₂O. This led to a final protein/protein equivalent concentration of 1.8% (w/v). The pH was readjusted, if necessary, following a 30 min equilibration under constant stirring. The solutions were centrifuged at 21,150 g for 20 min (Heittich Zentrifugen Universal 320R centrifuge, Andreas Heittich GmbH and Co., Tuttlingen, Germany). The supernatant was transferred into fresh tubes, an aliquot of the supernatant was diluted 1:100, and its protein content was estimated using the BCA assay. Protein solubility (%) was expressed as a percentage of the protein in the supernatant divided by the protein content in the starting suspension.

The heat coagulation time (HCT) was measured at 140°C for the SPH sample at pH 6.0 and 8.0 as described by Connolly et al. (27). An aqueous suspension of the SPH was prepared to a final concentration of 6.0% w/v (on a dry weight basis) and stirred for 1 h at ambient temperature. The pH was then adjusted and left to equilibrate for 1 h at room temperature. Following this, aliquots (2 mL, in triplicate) were added to glass tubes (10 mm i.d. 120 mm, AGB Scientific, Dublin, Ireland), sealed with silicone bungs, and submerged in an oil bath set at 140°C (Elbanton BV, Kerkdriel, The Netherlands). HCT was recorded as the time (in sec) required for the visible onset of coagulation of the sample to occur. All heat stability tests were performed in triplicate.

The emulsifying capacity of the hydrolysates was established at pH 2.0, 4.0, 6.0, 8.0, 10.0, and 12.0 as per Connolly et al. (27) with minor modifications. A final concentration of 0.05 g 100 mL⁻¹ hydrolysate solution (on a dry weight basis) was prepared in distilled water after pH adjustment with NaOH or HCl. A 28 g sample of the hydrolysate dispersion and 12 g of sunflower oil (obtained from a local food store) with Sudan III (40 mg L⁻¹ of oil) were placed in a 50 mL centrifuge tube, and homogenization was carried out using an Ultra-Turrax^® T25 homogenizer (IKA^® Werke GmbH and Co. KG, Staufen, Germany) at 16,000 rpm for 60 s. An 0.100 mL aliquot of the emulsion formed was immediately diluted 1: 800 with 0.1% (w/v) sodium dodecyl sulfate (SDS), and the absorbance was established at 500 nm using a UVmini-1240 spectrophotometer (Shimazu, Canby, USA), against a 0.1% (w/v) SDS blank for determination of the emulsion activity index (EAI). The remaining emulsion was stored upright and undisturbed in the 50 mL tube at room temperature. After 30 min, an aliquot (0.1 mL) from the lower portion of the tube was taken and diluted 1:800 with 0.1% (w/v) SDS to quantify the emulsion stability (ES).

The EAI was estimated as follows:

Where, Abs₅₀₀ is the absorbance at 500 nm, l the light path length (cm), Φ the oil volume fraction, and c the hydrolysate concentration (g 100 mL⁻¹).

The ES after 30 min holding was expressed as follows:

All emulsion analyses were carried out in triplicate at room temperature.

Undifferentiated C2C12 myoblasts were kept in a growth medium (DMEM), which included 10% (v/v) fetal bovine serum (FBS), 1% (v/v) penicillin/streptomycin, and 1% L-glutamine, in a humidified atmosphere containing 5% CO₂ at 37°C. As previously mentioned, differentiation was induced by placing 80% confluent cell cultures in a differentiation medium, which is DMEM-supplemented with 2% heat-inactivated horse serum (29). Before the main treatment, fully differentiated myotubes were starved of nutrients for 1 h in DMEM AA and serum-free medium containing 1 mM sodium pyruvate, 1% (v/v) penicillin/streptomycin, 1 mM L-glutamine, 6 mM D-glucose, and 34 mM NaCl (pH adjusted to 7.3). The C2C12 cells were then treated with the SGID-SPH in DMEM AA and serum-free medium at a final concentration of 1 mg protein equivalent/mL for 4 h. As a negative control for the assay, cells were incubated with DMEM AA and serum-free medium alone, while 100 ng/mL human insulin growth factor-1 (IGF-1) was used as a positive control.

The intracellular formation of ROS was determined using DCFH-DA as described by Yarnpakdee et al. (30) with some modifications. In brief, C2C12 cells were seeded in 96-well plates at a concentration of 7 × 10³ cells/well in 100 μL. DCFH-DA was initially prepared in dimethyl sulfoxide (DMSO) at a concentration of 4 mM and then further diluted to 100 μM in HBSS immediately prior to its application. The C2C12 myotube cells that were fully differentiated were rinsed with HBSS (100 μL/well, three times), followed by treatment with 1 mg protein equivalent.mL⁻¹ of SPH-SGID (100 μL/well) for 1 h. A positive control, consisting of Trolox at a final concentration of 100 μM, was conducted under the same conditions. A 100 μL aliquot of medium containing the test substances and controls was taken out and replaced with 100 μL of media containing 75 μM H₂O₂ and 100 μM DCFH-DA in HBSS. The fluorescence of the resulting 2′,7′-dichlorofluorescein (DCF) product was then measured every 10 min at 37°C for 90 min using a plate reader (BioTek Synergy HT, Waltham, MA, USA) with excitation at 485 nm and emission at 535 nm. Oxidation control wells consisted of cells in the presence of DCFH-DA and H₂O₂ without hydrolysate. The intracellular ROS level obtained in the presence of the hydrolysate was expressed as a percentage of the relative fluorescence intensity of the oxidation control cells.

Label-free, non-invasive, electric impedance measurements were taken using an xCELLigenceTM RTCA Instrument (ACEA Biosciences, Inc., San Diego, CA, USA) using a microelectronic E-16 well gold plated base sensor plate (ACEA Biosciences) (31). C2C12 myoblasts were cultured on an electrode-containing plate, as previously described (32). Throughout the cycle from myoblast proliferation through myotube formation, an automated reading of cell status, indicated as cell index (CI), was taken in real time (every 15 min during myoblast proliferation and every 2 min during myotube formation). Data were normalized to the start of the treatment phase of the experiment.

After 4 h incubation with controls and the SPH-SGID, multinucleated myotubes were counted under a phase-contrast microscope (Olympus CKX31, Tokyo, Japan). Image J software (National Institutes of Health, Baltimore, MD) was used to analyze images to quantify changes in myotube thickness. For each treatment condition, three diameter measurements were collected along each myotube, totaling at least 100 myotubes across at least six different fields. The average myotube diameter (μm) was then used to represent each treatment condition in the analysis.

Cells were seeded and differentiated in 6-well plates in a final volume of 2 mL cell culture medium/well. Trypsinization was used to harvest cells. These cells were washed three times with PBS (200 μL/well) and then lysed with cold lysis buffer (10 mM Tris/HCl pH 7.4, 150 mM NaCl, NaF, and 1% Na₄P₂O₇) containing phosphatase inhibitor [Na₃VO₄ (1 mM) and protease inhibitors (phenylmethanesulfonylfluoride fluoride (PMSF) (1 mM), pepstatin (1 μM), and aprotinin (1.5 μg.mL⁻¹)] for 30 min on ice, and then, each plate was scraped using a cell scraper. The homogenates obtained were then centrifuged at 130 g at 4°C for 10 min to remove nuclei and cellular debris. After determination of the protein content using the Bradford assay (following the suppliers' instructions), the lysates (30 μg protein/lane) were loaded on stain-free 4–15% linear gradient SDS polyacrylamide gel electrophoresis (PAGE) precast gels, and the gel was transferred to a nitrocellulose membrane using the semi-dry transfer technique (Trans-blotR TurboTM; Bio-Rad). After blocking with 5% (w/v) skimmed milk powder in 1X Tris-buffered saline (TBS) containing 0.5% Tween-20 (TBST) for 1 h at 37°C, the membrane was incubated with each antibody (1:1000 in 5% BSA in TBST: mTOR, phosphor (p)mTOR, P70S6K, pP70S6K, 4-EBP1, p4-EBP1, S6 ribosomal protein, pS6 ribosomal protein, β-actin, and puromycin) at 4°C overnight. The membrane was first treated with primary antibodies, except for puromycin which received a goat anti-mouse IgG2a-specific antibody, and then incubated with a secondary green rabbit antibody for 1 h. Images were captured using a UVITEC Cambridge Imaging system, and whole-lane band densitometry was quantified using NineAlliance UVITEC Software. After probing with phosphorylated antibodies, the membranes were stripped according to the manufacturer's instructions and then re-probed with total antibodies. Phosphorylated proteins were normalized to their respective total protein, while puromycin was normalized to the total protein density obtained from the stain-free lane for quantification purposes.

MPS was determined via the surface sensing of translation technique (SUnSET) (33). Following 1 h nutrient deprivation in AA and serum-free DMEM medium, differentiated C2C12 myotubes were treated with either 1 mg protein equivalent.mL⁻¹ SPH in AA and serum-free DMEM medium containing 1 μM puromycin (Merck Millipore Limited) and 100 ng.mL⁻¹ IGF-1 (positive control) or in amino acid and serum-free DMEM (negative control) containing 1 μM puromycin for a further 4 h. Immunoblotting was then used to assess MPS after obtaining the cellular protein lysates.

The results are presented as the mean ± standard deviation (SD) of three independent experiments. Data were analyzed using one-way or two-way ANOVA followed by Tukey's post hoc analysis. The SPSS software program (Version 27, IBM Inc., Chicago, IL, USA) was used to perform statistical analyses on the data. In all analyses, a P < 0.05 was taken to indicate statistical significance.

The protein content of the off-white SPH powder sample was 84.71 g/100 g. Following the terminology used for milk proteins, this product would correspond with an SPH80 since it contains more than 80% (w/w) protein. The DH of the SPH was 39.71 ± 0.24%, and this did not significantly change following simulated in vitro digestion of the sample (SPH-SGID) (Table 1). The molecular mass distribution of the SPH and SPH-SGID samples showed a high content of low molecular mass peptides (<1 kDa), followed by the fraction with a molecular mass between 1 and 5 kDa with very low levels for peptides >10 kDa (Table 1). The peptide profiles were similar in the SPH and SPH-SGID samples (data not shown) following reversed-phase ultra-performance liquid chromatography (RP-UPLC) analysis.

Table 2 provides the AA composition of the SPH sample and shows that the hydrolysate contains all the essential and non-essential AAs. The most abundant AAs were Glx (137 g/kg⁻¹) and Asx (91.5 g/kg⁻¹), and the lowest level was found for Cys (5.9 g/kg⁻¹). The protein quality of a dietary protein source can be estimated by comparing it to three reference AA scores of a model protein in accordance with the requirements of different age groups (34). These include the AA requirements for (1) infants < 6 months old (2) children 6–36 months old, and (3) children >36 months, adolescents, and adults. In this study, the ratios of each of the essential AA in the SPH with respect to the levels highlighted for each of the three reference cohorts were determined, and the lowest ratio was associated with the AA score. The manner in which the essential AA levels in a protein compare to those of a reference protein as well as which AA(s) may be limiting in the protein can be determined by AA score evaluation.

Table 3 shows the calculated AA scores for the SPH in different population cohorts. Tryptophan was identified as the limiting essential AA (0.49) when compared against the recommended AA requirement pattern for infants <6 months (34). Tryptophan was also identified as the limiting essential AA with an AA score of 0.97 for the cohort of children from 6 months to 3 years old.

Therefore, this information indicates that SPH may need to be supplemented, at different levels, with AA in order to meet the AA requirements of these two population cohorts. All AA scores for the older children, adolescent, and adult population cohort were above a value of 1.0 which indicates that the level of all essential AAs in the SPH met those recommended for this cohort. However, protein quality scores (which are measured in relation to the digestibility and the bioavailability of the essential AAs in a given protein) would need to be evaluated in order to confirm that SPH definitively meets the human dietary requirements for the specific population cohorts.

The solubility of proteins and their hydrolysates is an important technofunctional requirement for their applications in the food and pharmaceutical industries. Protein structure, pH, ionic strength (and salt type), and temperature all affect protein solubility (40). The aqueous solubility of the SPH was >90% across the pH range (pH 2.0–12.0) tested herein. While the mean solubility value at pH 8.0 was the highest, there were no statistically significant differences (p > 0.05) in solubility between the different pH values (Figure 1). A similar solubility profile was reported for protein hydrolysates from blue whiting (BWSPH), which had > 80% solubility between pH 2.0 and 10.0 (36). The overall high solubility is consistent with the solubility results observed for yellow stripe trevally (Selaroides leptolepis) (20) and sardinella, zebra blenny, goby, and ray protein hydrolysates (38).

The high solubility of the SPH could result from the large proportion of low molecular mass peptides (<1 kDa) contained therein. Furthermore, hydrolysis increases the number of terminal polar (NH3 COO-) groups and results in the unfolding of the protein structure revealing polar AA groups (such as serine, threonine, cysteine, tyrosine, asparagine, and glutamine) which can then interact with water molecules thereby enhancing solubility (41, 42).

Heat stability is an important parameter in the development of protein ingredients as it indicates changes in protein stability due to thermal treatments such as cooking. The HCT depends on several factors such as protein concentration, protein/peptide conformation, as well as the charge and structure of proteins. The impact of two different pHs on the HCT of 6% (w/v) SPH at 140°C was assessed herein. The HCT increased from 25 to 40 s by increasing the pH from pH 6.0 to 8.0. Enzymatic hydrolysis was shown previously to enhance protein/hydrolysate ingredient heat stability. The enzymatic hydrolysis of brewers' spent grain protein-enriched isolates showed a significant increase in heat stability when going from pH 6.0 to 8.0, i.e., increasing from 40 ± 3 s to >300 min (27). Ryan et al. (43) also reported the effect of pH on HCT at 140°C of soy protein hydrolysates where the HCT significantly increased with increasing pH (43).

The emulsion activity is a measure of the potential of an ingredient to be surface active and stabilize product formulations that contain both lipophilic and hydrophilic phases such as mayonnaise, salad dressings, sauces, milk, and sausages. Proteins and protein hydrolysates can stabilize emulsions by forming a viscoelastic film which prevents the phases from separation when absorbed to the surface of oil droplets during the emulsification process (44). Figure 2 shows the EAI and ES values for the SPH as a function of pH. Overall, the SPH showed higher EAI values at higher pH (pH 10 and 12 with EAI values of 825.59 and 1079.12 m² g⁻¹, respectively) and the lowest EAI was obtained at pH 4.0 (375.42 m² g⁻¹). Similar emulsion activity values have been reported for hydrolysates generated from marine-based protein sources, e.g., from Nemipterus japonicus (45). As can be observed in Figure 2, the ES of the SPH was between 58 and 100% across the pH range 2.0–12.0. The ES of the SPH was at its lowest at pH 6.0 (58.9%), and this was potentially related to the trend of a decrease in solubility at this pH. The ES increased with increasing pH which may be related to the trend of an increase in solubility (although non-significant in the present instance). Pacheco-Aguilar et al. (46) showed that the emulsion capacity (EC) of hydrolysates formed from Pacific whiting (Merluccius productus) muscle was significantly influenced by DH and pH, except at pH 4. Higher ES and EAI values were reported at higher pH values (46). According to Villamil et al. (47), the partial hydrolysis of proteins changes their structure and enhances the flexibility of the resulting peptides that subsequently align to the interface to form and stabilize emulsions. However, it is well recognized that extensive hydrolysis may reduce emulsifying properties (48). In addition, in an alkaline medium rich in negative charge, polypeptides unfold and expose hydrophobic groups facilitating protein/lipid interactions and improving the diffusion and stabilization at the oil/water interface (49).

Overall, the technofunctional data presented herein showed that the SPH had good solubility across a broad range of pH values. It had a light yellow color. It had an HCT of 40 s at pH 8.0. Moreover, compared to other studies, the SPH herein showed much higher emulsion activity and stability. The SPH did not display any foaming ability. These results indicate that the SPH has properties providing it with significant versatility for different product applications.

The ORAC and FRAP activities of the SPH and its simulated GI digests are presented in Table 4. Prior to SGID, an ORAC value of 587.49 ± 15.23 μmol TE/g sample was obtained, and a similar ORAC activity was observed following SGID (Table 4). As previously stated, the DH data and molecular mass distribution results indicate that limited hydrolysis occurred during SGID, which indicates that treatment with the gastrointestinal enzymes did not release more peptides with higher ORAC activity from the precursor peptides. However, it is possible that hydrolysis of peptides during SGID may have resulted in both a loss and gain of specific bioactivities, resulting in no overall change in the total bioactivity of the peptides in the SGID-treated SPH. The ORAC antioxidant activity of the SPH was higher than the ORAC activity described for other fish protein hydrolysates from channel catfish, pacific hake, blue mussel (Mytilus edulis), and blue whiting sources with ORAC values of 16, 225, 66.26, and 121.56 μmol TE/g sample, respectively (50–52). The SPH ORAC values observed herein were lower than the range described for Atlantic salmon (Salmo salar) trimming protein hydrolysates (601.47–882.58 μmol TE/g sample) (53).

FRAP values ranging from 10.93 ± 0.46 to 5.29 ± 0.15 μmol TE/g sample were obtained for the SPH prior to SGID and SPH-SGID, respectively (Table 4). The FRAP activity of the SPH was significantly decreased following SGID (Table 4). Similar findings were reported for other FPHs, bovine protein hydrolysates, and whey protein hydrolysates which had reduced FRAP level post-SGID compared to pre-SGID (50, 54, 55).

The differences observed in the antioxidant activity using the FRAP and ORAC assays may be explained due to the different modes of activity used in the analysis of these assays. The ORAC assay measures the capacity of test compounds to scavenge peroxyl radicals through hydrogen atom transfer (HAT), whereas the FRAP assay is categorized as an electron transfer (ET)-based, non-radical assay technique (56).

To further evaluate the antioxidant potential of SPH, the ability of the SPH-SGID to affect the endogenous antioxidant defense systems was assessed. The results from biochemical antioxidant assays, while useful, are in vitro tests and therefore may not be easily extrapolated to more complicated systems such as in the human body (57). In situ cellular-based assays, which may more accurately represent the target site of oxidative stress in vivo, may therefore be more relevant in the assessment of test compound antioxidant properties. The antioxidant activity of SPH-SGID was assessed in situ in C2C12 differentiated myotubes on the basis of the extent of ROS generation (Table 4) in the presence and absence of a pro-oxidant. The results showed that incubation with 1 mg protein equivalent.mL⁻¹ SPH-SGID for 1 h did protect C2C12 myotubes from the pro-oxidant effects of H₂O₂.

The application of cellular antioxidant-based assays of proteins and their derivatives using C2C12 myoblast and myotubes was reported in a number of previous studies (58–60). The cell cytotoxicity of two oxidative stress inducers (AAPH and H₂O₂) was pre-evaluated herein at different concentrations to investigate their potential toxic effects on C2C12 cells. The results showed that AAPH at concentrations ranging from 0 to 1,000 μM did not decrease cell viability by more than 85% which indicates that a higher concentration of AAPH was needed to decrease cell viability and induce more oxidative stress in C2C12 myotubes. Therefore, further studies are required to optimize the concentration of AAPH required to mediate a cytotoxic effect. There are several studies which employ H₂O₂ for the induction of oxidative stress in C2C12 cells (61, 62). Different concentrations of H₂O₂ (0–400 μM) resulted in a decrease in cell viability ranging from 35 to 85%. A toxic effect yielding <75% cell viability was found at ≥ 75 μM H₂O₂. Therefore, H₂O₂ was selected to represent the oxidative stress inducer at a concentration of 75 μM giving a mean cell viability value herein of 74%.

The cellular antioxidant assay was carried out to determine the potential antioxidative properties of the SPH-SGID against H₂O₂-induced intracellular ROS generation, as per Yarnpakdee et al. (30). The commercial antioxidant Trolox, which was used as a positive control, significantly reduced ROS generation as compared to the negative control, H₂O₂-stressed cells. Intracellular ROS generation in the SPH-SGID-treated cells (1 mg protein. mL−1 equivalent) was 67%. This demonstrated that treatment with SPH-SGID led to significantly (p < 0.05) lower levels of ROS generation compared to the negative control (Table 4). The significant reduction in ROS generation in the C2C12 cells treated with the SPH-SGID concurred with the results observed in the in vitro ORAC assays (Table 4).

In accordance with the antioxidant activity observed for the SPH herein, blue whiting (Micromesistius poutassou) protein hydrolysates and large yellow croaker (Pseudosciaena crocea) protein hydrolysates (MW < 3 kDa), which contained a high content of low MW peptides, presented O2-· and DPPH scavenging activity in vitro. Furthermore, they increased the activity of the antioxidant enzymes glutathione peroxidase (GSH-Px), superoxide dismutase (SOD), and catalase (CAT) in H₂O₂-induced oxidative stress in HepG2 cells (50, 63). In addition, numerous fish-derived peptides have demonstrated the ability to modulate oxidative stress pathways in vitro (64–67). Therefore, marine proteins are potential raw materials for the generation of antioxidant peptides with the ability to enhance health by reducing oxidative stress.

No studies appear to have been previously reported on the potential antioxidant activity of FPHs in oxidative stressed muscle cells. The results obtained for SPH herein demonstrate that this sample may have potential application as an antioxidant agent. Excessive production of ROS due to redox imbalance and impaired antioxidant defense systems leads to oxidative damage in most organs, including muscle (68–70). Increased ROS buildup causes oxidative stress, which in turn encourages proteolysis and causes muscle atrophy. ROS are crucial mediators of numerous signaling pathways that control this process (71, 72).

Therefore, the application of FPHs with potential antioxidant ability to reduce oxidative stress, thereby controlling oxidative stress and regulating the redox system, may be considered a promising approach for preserving muscle function.

C2C12 myotubes were treated with media conditioned with 1 mg protein equivalent.mL⁻¹ of SPH-SGID, AA and serum-free media, and IGF-1 to investigate the potential of SPH-SGID to stimulate myotube growth and activate MPS, as previously described (29, 32, 73). Based on the data from xCELLigence^TM analysis, the area under the curve (AUC) of cell index (CI) was significantly increased in response to treatment with SPH-SGID compared to the negative control (p < 0.05) as was the case for the IGF-1 positive control treatment (Figure 4A).

Myotube thickness was assessed after a 4 h treatment to verify changes in myotube diameter based on CI results. Myotube diameter was significantly increased (p < 0.001) compared to the negative control with SPH-SGID treatment (Figure 4B). These results indicate that SPH may have the potential to enhance myotube growth which may lead to the stimulation of MPS. These results are similar to the findings of Lees et al. (16) which previously showed that C2C12 cells demonstrate greater hypertrophy with BWPH-fed serum taken from healthy older adults compared with non-essential AA-fed serum based on myotube thickness (p = 0.028). Moreover, the current results demonstrate the potential ability and comparability of SPH with BWPHs regarding the promotion of myotube growth, proliferation, and MPS and demonstrate the potential of FPHs as a functional ingredient for the improvement of muscle health (32).

The mammalian target of rapamycin (mTOR) pathway is one of the most well-known signaling mechanisms in modulating MPS. A serine/threonine kinase called mTOR detects changes in the environment and within cells, such as the availability of nutrients and the level of energy (74). mTORC1 is known as a key regulator in controlling skeletal muscle mass following contraction and mechanical load-induced hypertrophy, synergistic ablation, myotube hypertrophy, and AA sensing, in which mTOR is involved in both skeletal muscle hypertrophy and atrophy (75). This important signaling molecule has been implicated in numerous studies as a crucial mediator in the conversion of mTOR activation to MPS activity (76, 77).

The ability of the SPH-SGID treatment to stimulate mTORC1 signaling was assessed by measuring the phosphorylation of mTORC1 and its downstream signaling molecules 4-EBP1 and rpS6. Many studies have identified this critical signaling molecule as a key mediator in translating the activation of mTORC1 to the activation of MPS (76–78).

mTOR phosphorylation on treatment with SPH-SGID was not significantly different to the negative or positive controls (p > 0.05, Figure 5A). Furthermore, activation of the downstream targets of mTOR activation, 4E-BP1 (Figure 5B), and rpS6 (Figure 5C) was not significantly different compared to negative or positive controls (p > 0.05). It is possible that these markers were not sufficiently sensitive to detect changes between conditions, and a lack of upregulation of mTOR phosphorylation on C2C12 treatment with BWPHs has been previously reported (16, 32).

It has been previously reported that BWPHs stimulated MPS by activation of phosphorylated 4E-BP1 and rpS6 in C2C12 cells (32). However, while no significant changes for these markers were observed following SPH treatment, downstream markers of protein elongation and translation responded differently to SPH, suggesting that these markers—which show higher activation of protein translation, may be more sensitive to subtle anabolic differences. Lees et al. (16) observed a similar response for both mTOR and downstream markers in response to treatment with both fish and milk protein sources. However, in their study, C2C12 cells were exposed to human serum-conditioned media. This media may have other stimulators for activating the mTOR signaling pathway, such as insulin.

The SunSET technique (33) was applied herein to quantify MPS and to verify whether mTOR, rpS6, and 4E-BP1 activation could mediate an increase in MPS in skeletal muscle cells (C2C12 myotubes) following 4 h treatment with SPH-SGID. The results obtained showed that puromycin incorporation was significantly increased in response to SPH-SGID treatment compared to both the negative and positive controls (p < 0.05, Figure 5D). These results support the data from the CI and myotube diameter analyses. As a result, when taken as a whole, SGID boosted skeletal MPS and anabolism in situ, which may be due to an unknown mechanism that increased translation initiation factors rather than boosting mTOR signaling.