Recent Progress in Sustainable Fish Byproduct Utilization: Unveiling Fish Collagen as a Potential Wound Healing Agent

Collagen extracted with the acidic solution, is commonly known as acid-solubilised collagen (ASC) (Figure 2). The use of acids (organic and inorganic acids) during collagen production can cleave bonds in collagen molecules to enable extraction of the tissues (Skierka and Sadowska, 2007). In an acidic condition, the collagen molecules undergo a net positive change and the resulting electrostatic repulsive force between them facilitates molecular separation. There are many acid compounds used to obtain collagen from the marine fish tissues, including organic acids (acetic, lactic, and citric acids) and inorganic acid (hydrochloric acid) (Tan and Chang, 2018). Among these acids, organic acids are apparently more effective to break some inter-stand crosslinks of collagen molecules, resulting in higher extractability of collagen yield, as compared to the mineral acid. In addition to this, organic acids are capable of solubilising non-crosslinked collagens. However, acetic acid (AcOH) is mostly employed in extracting collagen, in particular collagens from marine fish sources. Collagen extraction from marine fish derived byproducts has been effectively achieved by using various parameters such as time, concentration of acetic acid and temperature. The reported ASCs extraction by using various concentrations of acetic acids are from skin, bones, scales, and swim bladders of spotted golden goatfish (P. heptacanthus) (Matmaroh et al., 2011), purple-spotted bigeye (P. tayenus) (Oslan et al., 2022), barracuda (Sphyraena sp.) (Matarsim et al., 2023), bigeye tuna (T. obesus) (Ahmed et al., 2019) starry triggerfish (A. stellatus) (Ahmad et al., 2016), unicorn fish (N. reticulatus) (Fatiroi et al., 2023), sin croaker (J. sina) (Normah and Afiqah, 2018), seabass (L. calcalifer) (Chuaychan et al., 2015), grouper (Epinephelus sp.) (Dong and Dai, 2022), shortfin scad (D. macrosoma) (Sulaiman and Sarbon, 2020), grey mullet (M. cephalis) (Minh et al., 2014), needle fish (T. melanotus) (Ramle et al., 2022), and sharpone stingray (D. zugei) (Ong et al., 2021).

Figure 2.

ASC and PSC extraction process

In general, collagen extraction involves the use of various acids, such as acetic, lactic, citric, and hydrochloric acids, without the addition of protease enzymes. However, this method often yields incomplete dissolution of collagens from different sources, resulting in suboptimal extraction yields (Jaziri et al., 2022 b; Skierka and Sadowska, 2007). To address this limitation and achieve maximum extractability, enzymatic extraction methods have been developed. These methods employ enzymes like pepsin, trypsin, bromelain, papain, and various collagenases under specific environmental conditions (i.e., concentration, pH, temperature, and liquid-solid ratio). Among these enzymes, pepsin is a prominent choice for extracting collagen from marine fish sources (Table 1). Pepsin can be applied either independently or in conjunction with different concentrations of acetic acid. Literally, pepsin can effectively cleave the telopeptide regions within the triple helical structure of collagen molecules, thereby aiding in the release of collagen peptides into the solution and consequently enhancing extraction yields (Benjakul et al., 2010). Collagen obtained through pepsin-assisted extraction are referred to as pepsin solubilised collagen (PSC) (Figure 2). Notably, researchers have found that utilizing limited concentrations of enzymes, particularly pepsin, along with acids, significantly enhances the overall yield of collagen. This underscores the importance of enzymatic extraction methods in optimizing collagen extraction efficiency (Arumugam et al., 2018). Enzymatic approach is a promising technique for collagen extraction, driven by some reasons. For instance, improving extraction efficiency by facilitating the enhanced solubilisation of collagen in acetic medium, reducing collagen antigenicity, and preserving triple helical structure of collagen. Overall, these factors contribute to the increasing applications of enzyme-extracted collagen in both the food and pharmaceutical industries (Ahmed et al., 2020).

Optimization is a critical aspect of extraction processes, involving the systematic exploration of various factors to determine the conditions that yield optimal results (Arumugam et al., 2018). Particularly in complex processes where the targeted response is influenced by multiple factors, the selection of an appropriate experimental design becomes imperative. In the context of collagen extraction from marine fish byproducts, efficiency is notably influenced by factors such as concentration, extraction time, solid-liquid ratio, and temperature. Thus, a well-designed experiment capable of analysing these multiple factors is essential. Once the region of optimal response is identified through preliminary studies, such as one-factor-at-a-time (OFAT), further characterization of the response within that region is often necessary. To achieve this, experimental designs such as the Box-Behnken design (BBD) and central composite design (CCD) are commonly applied in response surface methodology (RSM). These designs facilitate the approximation of a second-order polynomial estimation for the response within the identified optimal region (Mohammadi et al., 2016).

RSM employs mathematical and statistical techniques to model situations where variables impact a response of interest, with the primary objective of achieving an optimized response. When the relationship between independent variables and the response is linear, the first-order model is employed as an approximate function. However, if the response surface exhibits curvature, a second-order polynomial, or a higher degree polynomial, is used as the model. Response surface design is a specific approach for fitting response surfaces, and two common types are the one-factor-at-a-time design and factorial design. Factorial design is particularly useful when studying two or more levels, including both full factorial and fractional factorial designs. In cases where the number of runs for a full factorial design becomes impractical, fractional designs offer a viable alternative. Notably, BBD and CCD are examples of widely used fractional factorial designs employed for optimization purposes (Mohammadi et al., 2016). RSM offers several advantages, including a reduction in the number of experimental attempts required to assess multiple parameters and the capacity of the statistical tool to identify interactions. RSM provides the added benefit of generating a comprehensive mathematical model that captures the entirety of the process. Besides that, it proves to be a time-saving method that minimizes errors in assessing parameter effects. Mostly used statistical approaches in RSM are CCD and BBD, both of which have proven successful in optimizing marine collagen extractions, as illustrated in Table 2.

For instance, the effects of NaOH concentration, pretreatment time, pepsin concentration, and hydrolysis time on the extraction yield, a four-factor, five-level CCD for RSM was employed. The CCD encompassed 28 experimental runs, including 8 axial points, 16 factorial points, and 6 replicates of the central point. Using the desirability function approach, the estimation of the R² value, multi-factor ANOVA, lack of fit value, and second-order model prediction determinants was employed to attain optimal conditions. Response surface plots, including surface plots and contour plots, were generated to elucidate the impacts of the four factors on the extraction yield. The ANOVA of the estimated model led to the conclusion that the effects of NaOH concentration, pretreatment time, pepsin concentration, and hydrolysis time showed interactive effects on the collagen yield. Consequently, with the assistance of the regression model, the optimal conditions were determined to be the NaOH concentration of 0.92 N, pretreatment time of 24 h, pepsin concentration of 0.98%, and hydrolysis time of 23.5 h (Woo et al., 2008).

Colour is one of the most important parameters of collagen in industrial applications (viz. foods, cosmetics, pharmacy, and medicine). Collagen with bright colour is more acceptable because it does not change the original colour of final products, and it does not influence the functional properties of collagen (Liu et al., 2019). Some works related to the tropical marine fish collagen examined the attributes of colour in the final products using CIELAB colour space, which represents the values of L* (lightness), a* (red/green), b* (blue/yellow) described by the International Commission on Illumination (CIE). For instance, the barracuda (Sphyraena sp.) and needle-fish (T. melanotus) skin collagens extracted with different organic acids had a low value of L* (54.34–56.88 and 57.14–69.77, respectively) (Matarsim et al., 2023; Ramle et al., 2022). In addition to this, the ASC and PSC derived from barramundi (L. calcalifer) skin showed the L* values of 65.41 and 61.33, respectively (Bakar et al., 2013). In contrast, collagens isolated from the bones of unicorn fish (N. reticulatus) and parrotfish (S. sordidus) exhibited the L* values of 881.55 and 74.81, respectively (Fatiroi et al., 2023; Jaziri et al., 2023). Besides that, the lightness property observed in the threadfin bream (N. japonicus) scale and fins collagen was considered higher (L* = 93.70–94.82) (Normah and Afiqah, 2018). In can be stated that the byproducts of bones, scale and fins are more effective raw materials in producing collagen rather than fish skins. As described by Sadowska et al. (2003), the existence of pigments in fish skins causes the difficulty of preparing fish collagen entirely devoid of colour. Thus, the colour of collagen depends on the raw materials used and method of extraction. As confirmed by Liu et al. (2019), the L* value of extracted collagens generated from snakehead (Channa argus) skins was 89.49, indicating higher than the aforementioned fish skin collagens due to addition of hydrogen peroxide (H₂O₂) during pretreatment process. The use of H₂O₂ solution might be effective to remove the skin sample’s pigment.

Ultraviolet visible (UV-vis) absorption is a common method to evaluate the characteristic of extracted collagen from several sources. It is considered as a simple approach by scanning the collagen sample from 200 nm to 400 nm, because triple helical collagen has maximum peak at 230 nm and a negative peak near 204 nm (Piez and Gross, 1960). In addition, collagen contains a low concentration of chromophore amino acids, such as phenylalanine, tryptophan, and tyrosine, in which these amino acids typically absorb UV light at 250 nm and 280 nm (Liao et al., 2018). The number of studies on the ASC and PSC produced from various tropical marine fish byproducts had different prominent absorption peaks, such as purple-spotted bigeye (P. tayenus) skin collagens (230–240 nm) (Oslan et al., 2022), barracuda (Sphyraena sp.) skin collagens (230.5 nm) (Matarsim et al., 2023), puffer fish (L. inermis) skin collagens (230 nm) (Iswariya et al., 2018), unicorn fish (N. reticulatus) bone collagens (229.8–231.2 nm) (Fatiroi et al., 2023), parrotfish (S. sordidus) bone collagens (230–232 nm) (Jaziri et al., 2023), and needlefish (T. melanotus) skin collagens (231.5 nm) (Ramle et al., 2022). Overall, the mentioned peaks are closely related to the functional groups of carbonyl (C=O), carboxyl (−COOH) and amide (−CO-NH₂) belonging to the polypeptide chains of collagen (Edwards and O’Brien, 1980).

So far, type 1 collagen is mostly explored and applied in industrial fields. It is characterized by possessing at least two different alpha chains (α1 and α2) and the dimer beta (β) chain (governed by intramolecular crosslinking) (Matmaroh et al., 2011). Using the sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE), the electrophoretic bands of collagen can be confirmed, and the molecular weights (MW) of collagen may also be determined. Table 1 shows all tropical marine collagens isolated from the skin, bone, scale, fin, and swim bladder byproducts were classified as a type 1 due to the presence of two identical subunits of α1 chains and one subunit of α2. However, the obtained collagens from the cartilage and head bone of silvertip shark (Carcharhinus albimarginatus) have been identified as a type II collagen, indicating one α chain and its dimer (β chains) (Jeevithan et al., 2014). Although most of cartilage-derived collagens are identical to the type II collagen, Kittiphattanabawon et al. (2010) reported that the collagens produced from the cartilages of brownbanded bamboo shark (Chiloscyllium punctatum) and blacktip shark (Carcharhinus limbatus) were categorized as both type I and type II, as similar to that of Carcharius acutus cartilage collagen documented by Rama and Chandrakasan (1984). In terms of MW prediction, the α1 and α2 of type 1 marine fish collagens were comparable, and almost above 100 kDa, such as starry triggerfish (A. stellatus) skin collagen had α1 of 119–121 kDa and α2 of 112–117 kDa (Ahmad et al., 2016), while the purple-spotted bigeye (P. tayenus) skin collagens were 118 kDa and 106 kDa (Oslan et al., 2022). Other collagen sources, the spotted golden goat-fish (P. heptacanthus) and parrotfish (S. sordidus) scales showed a similar MW of α1 and α2 (118 kDa and 107 kDa, respectively) (Jaziri et al., 2023; Matmaroh et al., 2011). On the other hand, higher MWs were found in the unicorn fish (N. reticulatus) bone collagen (α1 = 138 kDa and α2 = 118.3 kDa and the grouper (Epinephelus sp.) swim bladder collagen (α1 = 133 kDa and α2 = 123 kDa) (Dong and Dai, 2022; Fatiroi et al., 2023), indicating that different fish species and extraction methods resulted in different molecular weight distributions. Besides the different MW distributions, the band intensities of α1 chain in the extracted marine fish collagen are generally twice those of α2 chain. The β- and γ-components of all tropical marine collagens had higher MWs of 200 kDa (Matarsim et al., 2023; Ramle et al., 2022).

Since collagen is protein molecules made up of amino acids, understanding amino acid composition of collagen is an essential perspective, particularly type 1 collagen. In general, the quantification of amino acids is carried out by using chromatographic methods (Jafari et al., 2020). It is well recognized that collagen predominantly contains glycine (about a third of the total amino acid composition), proline and hydroxyproline amino acids. As presented in Table 3, the major amino acids found in the tropical marine fish collagens are glycine, alanine, proline, glutamic acid, hydroxyproline, and arginine, thus describing the presence of type I. These amino acids are also documented in other sources of collagen like calf skin, porcine skin, and human type I (Yu et al., 2018). Among these amino acids, as aforementioned, both proline and hydroxyproline are called an imino acid due to presence of a secondary amine group, and they are often used to identify collagen properties from different sources. Additionally, hydroxyproline content represents the amount of collagen extracted in the final product, and its composition is diverse depending on the raw material of extracted collagen. Based on the existing literature related to collagen isolated from several sources, the imino acid contents are higher in mammalian collagens than those found in fish (e.g., warm-water fish and cold-water fish) collagens. As can be seen in Table 3, the contents of imino acid obtained in different collagen sources are approximately 22% for mammals, 17–20% for tropical marine fish, and 15% for tempered fish (Ahmad and Benjakul, 2010; Ding et al., 2019; Matmaroh et al., 2011; Yu et al., 2018). Moreover, the diverse amino acid compositions of collagen influence the collagen properties. For instance, the thermal stability of mammals’ collagen is greater compared to that of collagen derived from fish species, because the sterical side groups of imino acid restrict the conformation of the polypeptide chain, and the hydroxyl group of hydroxyproline plays a major role in intramolecular hydrogen bonds.

Fourier transform infrared (FTIR) spectroscopy is widely used to examine the collagen extracted from organisms’ tissues, including tropical marine fish species. Using this tool, the chemical composition and type classification of collagen can be determined after being isolated with various extraction conditions. As illustrated in Table 4, the characteristic peaks of extracted marine collagen samples were amide a, amide b, amide I, amide II, and amide III (Ahmed et al., 2019; Fatiroi et al., 2023; Matarsim et al., 2023). The absorption parameter of amide A is closely related to N-H stretch vibration, and normally located at the wavenumber of 3400–3440 cm⁻¹ (Sai and Babu, 2001), while amide B is associated with CH₂ asymmetric stretching (Abe et al., 1972). For amide I, it is found in the wave-number range 1600–1700 cm⁻¹ and representing the C=O stretching/hydrogen bond coupled with COO− (Payne and Veis, 1988). Amide II is commonly responsible for N-H bend coupled with C-N stretching (Krimm and Bandekart, 1986), and the rest of amide III is regarded as N-H bend coupled with C-H stretching (Payne and Veis, 1988). By assessing these amides as recognized in the absorption peaks of collagens (Table 4), all extracted collagens were consistent with the wavenumber of amides. Furthermore, especially in the amide I–III, these peaks are involved in the formation of a triple-helical structure of collagen molecules. Other studies by Nikoo et al. (2013) and Plepis et al. (1996), revealed that the triple-helical regions can be examined by measuring the difference in the wavenumber (cm⁻¹) between amide I and II bands using the formula Δv (vI − vII), where values <100, indicating that the collagen structure is preserved. Besides that, the triple helix can be also verified by using the absorption ratio (~1.0) of the amide III to the 1450 cm⁻¹ band (AIII/A1450) (Plepis et al., 1996). Through these approaches, for the resultant collagens from tropical fish species prepared by using acids and enzymatic processes it could be confirmed that the functional groups present in the triple-helical structure were not damaged, suggesting that the treatment conditions during extraction would not influence the final collagen products.

X-ray diffraction (XRD) method is often used in aquatic and marine sources to evaluate the fibrillar structure of collagen after being extracted by different approaches. XRD uses the Bragg law with an equation: 2dsinθ = λ, and was employed to determine the minimum of repeated interval (d), where λ is the wavelength of x-ray (1.5405 Å), and θ is the Bragg diffraction angle (Wang et al., 2009). The peaks in the 2θ curves plotted against intensity correspond to features like the spacing between collagen fibres, bonds within polypeptide chains, and/or the diameter of the collagen triple helix. These characteristics could be influenced by the method used for extraction and solubilisation, ultimately indicating the integrity of the extracted collagen (Chen et al., 2016). Basically, XRD pattern of collagen has three prominent peaks (e.g., peak A₁, A₂ and A₃) with the diffraction angle usually located in 5–10°, near 20° and 30–35°, respectively (Jaziri et al., 2022 c). Peak A₁, first peak in XRD, reflects the distance between the molecular diffraction pattern of collagen. Second peak (A₂) is obtained by diffuse scattering, and the last peak of A₃ describes the unit height, typical of the triple helical structure of collagen molecules (Giraud-Guille et al., 2000). Comparably, some researchers have reported the presence of two significant peaks i.e, sharp and broad peaks in the assessment of tropical marine fish collagens isolated from the skin of barracuda (Sphyraena sp.) (ASC: 11.6–12.2 Å and 4.4–4.6 Å) (Matarsim et al., 2023), the bones of unicorn fish (A. monocerous) (ASC: 11.3–11.4 Å and 3.3–3.4 Å) (Fatiroi et al., 2023), barramundi (L. calcalifer) (PSC: 12.5 Å in the first peak) (Liao et al., 2018), and the skin and bone of golden pompano (T. ovatus) (PSC: 11.1 Å and 11.7 Å in the peak 1) (Cao et al., 2019). According to Chen et al. (2016), if the spacing between fibres is higher at first peak, collagen is more capable of carrying drugs, making it a preferred choice for the pharmaceutical company.

Differential scanning calorimeter (DSC) assesses collagen enthalpy (ΔH) variations using a thermally inert reference material (usually indium). Under identical temperature conditions, the sample and reference material produce positive and negative peaks, indicating energy removal (exothermic peak) or addition (endothermic peak). The endothermic peak in the sample reflects collagen denaturation, with T_max representing the maximum transition temperature and correlating directly with enthalpy (ΔH) and denaturation resistance. Pepsin-soluble collagen (PSC) exhibits lower thermostability than acid-soluble collagen (ASC) due to telopeptide region removal by pepsin. The study establishes a positive correlation between collagen thermostability and imino acids (proline and hydroxyproline) content, indicating enhanced hydrogen bond stabilization (Matmaroh et al., 2011). Many studies have been conducted on the tropical marine fish sources with having a variety of thermal stability, such as the ASC and PSC from spotted golden goatfish (P. heptacanthus) scale (T_max = 41.58°C, ΔH = 12.38 J/g and T_max = 41.01°C, ΔH = 15.39 J/g) (Matmaroh et al., 2011), starry triggerfish (A. stellatus) skin (T_max = 35.9°C, ΔH = 2.4 J/g and T_max = 33.6°C, ΔH = 2.1 J/g) (Ahmad et al., 2016), and brownbanded bamboo shark (C. punctatum) (T_max = 36.73°C, ΔH = 1.55 J/g and T_max = 35.98°C, ΔH = 0.85 J/g) (Kittiphattanabawon et al., 2010).

Microstructural property and surface area of collagen are important to evaluate its potential applications in biomedicine and biomedical engineering. Its characteristics are normally observed under scanning electron microscopy (SEM) or field emission scanning electron microscopy (FESEM) (Bhuimbar et al., 2019). Typically, fish collagens extracted by using acid medium and pepsin-added process show irregular dense sheet-like film linked by random-coiled filaments and their surface is partially wrinkled due to dehydration during freeze-drying. Moreover, with a certain magnification employed, lyophilized fish collagens present some fibrillar structures. Ogawa et al. (2004) stated that collagens with interconnectivity, fibrillary and sheet-like film structures have the potentiality to be used in new tissue formation, cell seeding, growth, wound healing and mass transport and migration. In general, the microstructural analysis of collagens obtained from the tropical marine sources, such as barracuda (Sphyraena sp.) skin (Matarsim et al., 2023), puffer fish (L. inermis) skin (Iswariya et al., 2018), unicorn fish (N. reticulatus) bone (Fatiroi et al., 2023), giant croaker (N. japonica) swim bladder (Chen et al., 2019), silvertip shark (C. albimarginatus) skeletal and head bone (Jeevithan et al., 2014), indicated the microstructural characteristics conducive to various medical applications. This cumulative evidence suggests that collagens from these tropical marine sources may indeed serve as promising biomaterials for a range of medical applications.

Solubility of collagen is extensively treated with various pH levels (1–12), and with various salt (NaCl) concentrations (0–60 g/L and 0–6%), in acetic acid medium, usually initial concentrations used are between 3 mg/mL and 6 mg/mL (Jaziri et al., 2022 d). In general, the protein content in the supernatant (solubilised samples) is determined by the Lowry procedure using the standard curve of bovine serum albumin (BSA). Furthermore, the solubility of collagen can be measured in comparison to that obtained at the pH and/or NaCl concentration yielding the highest solubility, referred to as the relative solubility (%) (Ahmed et al., 2019). The relative solubility range of collagen varies based on the extraction method applied. Table 1 shows that the collagens isolated from the skin, bones, scales and swim bladder of tropical marine fish were soluble within the pH range of 1–4, 1–5, 1–5, and 1–4, respectively. Meanwhile, for the NaCl treatment, the mentioned samples were almost soluble at the salt concentration of below 30 g/mL (Cao et al., 2019; Chen et al., 2019; Fatiroi et al., 2023; Jaziri et al., 2023). The solubility of collagen is affected by its structure and amino acid composition, particularly when subjected to elevated concentrations of NaCl (Ahmed et al., 2019).

Proteases (also known as proteinase, peptidase, or proteolytic enzymes) are enzymes that catalyse proteolysis, breaking down proteins into smaller polypeptides or single amino acids, and facilitating the formation of new protein products (López-Otín and Bond, 2008). They belong to a certain group of enzymes that can hydrolyse by catalysing the reaction of hydrolysis of different bonds and they do that with the participation of a water molecule. Each enzyme has its own optimal conditions in relation to temperature, pH and ionic strength that differ between enzymes. The order in which enzymes are added to the reaction mixture can change the effects of each individual enzyme. When the first enzyme has brought on their reaction it becomes the substrate of the second enzyme. The order of the enzyme reactions can influence the degree of hydrolysis (León-López et al., 2019 b). In general, proteolytic enzymes can be divided into two major groups: endopeptidases, which cleave internal peptide bonds, and exopeptidases, which cleave C- or N-terminal peptide bonds, and those enzymes were widely used in industrial applications, accounting for more than 60% of total global enzyme sales with the market size estimated at USD 3.54 billion in 2024.

In terms of marine fish collagen hydrolysates, researchers have employed a variety of proteases in producing collagen hydrolysates, such as papain, alkaline protease, collagenase, pepsin, Alcalase, Neutrase, trypsin, and Flavourzyme (Felician et al., 2019; Hema et al., 2017; Wang et al., 2020). Those proteolytic enzymes were used individually and/or in combination in generating fish collagen hydrolysates, and their properties (physicochemical, functional, and antioxidant) have been evaluated. Among the mentioned proteinases, papain and alkaline protease are often used in hydrolysing collagens from the byproducts of tropical marine fish source, as reported by Chotphruethipong et al. (2020), Felician et al. (2019), Hema et al. (2017), and Wang et al. (2020) for collagen hydrolysates derived from the grouper (Epinephelus malabaricus) skin, jellyfish (R. esculentum) filaments, redlip croaker (Pseudosciaena polyactis), and barramundi (L. calcarifer), respectively.

The process of wound healing can be divided into four stages: haemostasis, inflammation, proliferation, and remodelling, as depicted in Figure 3.

Many collagens from the byproducts of marine fish sources were studied by researchers in terms of their capability of enhancing wound healing under in vitro evaluations. As tabulated in Table 6, collagen from the byproducts of bigeye tuna (T. obesus) (Lin et al., 2019), unicorn leatherjacket (A. monoceros) (Kumar et al., 2019), Nile tilapia (O. niloticus) (Zhou et al., 2017), narrow-barred Spanish mackerel (S. commerson) (Naderi Gharahgheshlagh et al., 2023), pirapitinga (Piaractus brachypomus) (Manjushree et al., 2023), flounder fish (Paralichthys sp.) (Sousa et al., 2023), giant croaker (N. japonica) (Zheng et al., 2020), mrigal fish (Cirrhinus cirrhosus) (Pal et al., 2016), jellyfish (Rhopilema esculentum) (Felician et al., 2019), gurijuba (H. parkeri) (Ferreira et al., 2022) increased the growth of normal skin cells (i.e., fibroblasts, keratinocytes, and endothelial cells) during viability and/or scratch tests. Among them, the type I collagen derived from the skin of bigeye tuna (T. obesus) showed the increment of NIH-3T3 fibroblasts’ growth even though there were no significant differences (P>0.05) on all treated groups; however, the scratch closure rates were better compared to the control (Lin et al., 2019). Other examples of collagen originated from the skins of unicorn leatherjacket (A. monoceros) and Nile tilapia (O. niloticus) exhibited that at concentration of 0.2 mg/mL, a greater migration of cells was noted in the collagen compared to other treated samples and a higher scratch closure presented in the hydrolysed collagen (50.0 μg/mL) than other treatments, respectively (Hu et al., 2017; Kumar et al., 2019).

Manjushree et al. (2023) investigated that at the highest concentration of the hydrolysed collagen from pirapitinga (P. brachypomus) skin could increase the percentage of L292 mouse fibroblast cells, and its hydrolysate became non-toxic when observed in the MTT assay. In addition to this, a remarkable wound closure ability of more than 80% at 12 h and 100% within 24 h was recorded during the scratch evaluation. Besides that, Felician et al. (2019) confirmed that collagen from jellyfish (R. esculentum) had significant effects on the scratch closure on human umbilical vein endothelial cells treated with collagen (6.25 mg/mL for 48 h), as compared to the vehicle treated cells, and even reported by Zheng et al. (2020) that the collagen from the swim bladders of giant croaker (N. japonica) could significantly minimize the oxidative stress damage caused by H₂O₂ in HUVEC.

Fish collagen incorporated with other components also supported the growth of skin cells, as documented from Afzali and Boateng (2022), Ferreira et al. (2022), Muthukumar et al. (2014 a), Ramanathan et al. (2017 b), and Selvaraj et al. (2023). Barramundi (L. calcarifer) scale collagen incorporated with mupirocin and extracts of M. uniflorum exhibited a great biocompatibility agent against the NIH-3T3 fibroblast cells and human keratinocyte cells (HaCaT) (Muthukumar et al., 2014 b). In line with previous example, a high biocompatibility with NIH-3T3 murine fibroblast cells was observed in the swim bladder collagen of gurijuba (H. parkeri) incorporated with chitosan (Ferreira et al., 2022), and a good biocompatibility of the scaffolds (Arothron stellatus skin collagen loaded with extract of Coccinia grandis and drug ciprofloxacin) was noted owing to the presence of grooved pattern of the collagen matrix with cellular growth and adhesion (Ramanathan et al., 2017 b). Furthermore, collagen-coated silk fibroin nanofibers increased production of collagen shown in the BALB/3T3 clone A31 fibroblast cells (Selvaraj et al., 2023).

Figure 4.

Schematic diagram of four phases in the wound healing process

The existing investigations documented that the collagens from the byproducts of aquatic fish sources had potential wound healing agents, as presented in Table 7. The collagen products were widely treated via oral/intragastrical administration in rodent models (rats and mice) in a dose-dependent manner. Zhang et al. (2011) initially observed the collagen from the skin of chum salmon (Oncorhynchus keta) administered orally started 1 day before wounding (2 g/kg/day) in male Sprague-Dawley rats (230–250 g) and the results showed twelve percent (p<0.01) increase in the % coverage of wound found in the treated group as compared to the control group (excision wound type), increased fibroblast infiltration, vascularization, epithelialisation and collagen deposition in the wounds treated with collagen peptides (incision type), and strongest upregulation in the VEGF and FGF-2 genes expression. Another finding was reported by Yang et al. (2018), wherein collagen peptides with low-molecular weight from Alaska pollock (Theragra chalcogramma) gavaged with collagen samples daily (0.5–2.0 g/kg) in male Sprague-Dawley rats (160–170 g) had faster wound healing rate in the treated group with collagen peptides on day 4–12 post-surgery with a near-normal epidermis structure, increased rates of EGF, bFGF, TGF-β1 and TβRII, and decreased levels of Smad7 in TGF/Smad signaling pathway. Mei et al. (2020), moreover, updated the collagen from Salmo salar and tilapia skins could accelerate significant wound healing, with a complete healing on day 12 post-injury, decrease TNF-α, IL-6 and IL-8, upregulate BD14, NOD2, IL-10, VEGF, and β-FGF, alter cutaneous microbiome colonization through upregulated NOD2 and BD14 genes expression. The collagen samples were previously prepared through daily oral administration of collagen peptides (2 g/kg body weight) to male Sprague-Dawley rats.

Besides that, some researchers evaluated the wound healing process treated intragastrically by the collagen derived from fish byproducts in mice model. Felician et al. (2019) revealed that smaller wound closure on mice (excisional wound = 8.5 mm²) after 6 days post-injury was obtained from the collagen from jellyfish (R. esculentum) administered intragastrically every morning for 6 days (0.3–0.9 g/kg b.w.), resulting in remarkable sign of reepithelialisation, tissue regeneration and increased collagen deposition. Another study from Xiong et al. (2020) reported that tilapia collagen peptide administered via drinking water (15–45 g/L) could enhance wound healing rate after 5 days post-wounding on male C57BL/6 mice, and it could increase collagen deposition, hydroxyproline level, as well as improve IGF-1, FGF2, mRNA expression of growth factors, serum cytokine, NO, SOD and CAT. Yang et al. (2019) also reported that the active peptides with low dose (0.5 g/kg) exhibited a shortened epithelialisation time by inhibiting inflammatory response, enhanced the proliferation of FGF, CD31 and EGF, and increased collagen synthesis via the TGF-β/Smad signalling pathway. Furthermore, latest research by Hou and Chen (2023) confirmed that sturgeon fish (Acipenser baerii × Huso huso) skin collagen peptide-based nanoemulsion was the most effective in decreasing fasting blood glucose (46.75%) obtained from the treated group with low-MW and high-dose nanoemulsion, as well as the low-MW and high-dose nanoemulsion also enhanced wound healing area (95.53%) compared to other treated groups.

Unless oral gavage approach, collagens from the byproducts of aquatic species were also applied in the form of wound dressings, including sponge, film, nanofiber, and hydrogel dressings. For instance, Lin et al. (2019) observed collagen sponges from the swim bladders of giant croaker (Nibea japonica) had quicker wound closure in the wounds compared to the control group (untreated sample), and the sponges significantly decreased the levels of IL-1β, IL-6 and TNF-α in male ICR mice (6–8 weeks old, 22–24 g). Similarly, Li et al. (2023) used swim bladder collagen from sea eel (Muraenesox cinereus) as a sponge dressing in the wounded male ICR mice (excision wound model = 10 mm²), and the results show rapid wound healing was exhibited in the treated group and prevented scar formation in the later stage. Another example using type II collagen from the cartilage of Acipenser baerii, the findings showed that the wounded male C57BL/6J mice (8–10 weeks old) were repaired after treated by the sponge dressing. This treatment was associated with reduced inflammation, increasing granulation, tissue formation and collagen deposition, upregulating the production of growth factors (Lai et al., 2020). In terms of film dressing, Ramanathan et al. (2017 b) explored starry puffer (Arothron stellatus) skin collagen film incorporated with Coccinia grandis had a faster healing of the wounded Wistar albino male rats (2×2 cm²) that was detected in the treated group after 8th day up to the complete healing (at day 18). Besides film dressing, Fatemi et al. (2021) studied the collagen-chitosan from Scomberomorus guttatus and shrimp exhibited a significant reduction in wound size of wounded Sprague-Dawley rats compared to silver sulfadiazine treated group on days 15 and 25.