The skin is the largest organ in mammalian organisms. It acts as the main barrier and plays several vital roles in immunity, fluid homeostasis, thermoregulation, excretion, sensation, and metabolic functions of the body [1
]. When the skin is wounded, the tissue undergoes healing in the ordered steps of homeostasis, inflammation, proliferation, and remodeling, to promote wound closure and eventually restore the normal tissues [1
]. This physiological healing process is highly coordinated and involves interactions between various cellular and molecular networks in the wound [2
]. In general, the acute wound usually heals completely within weeks with normal healing phases and has normal wound physiology. However, chronic wounds are characterized by a persistent inflammatory phase, which are difficult to heal within months or a year [4
]. Moreover, chronic wounds are medical and healthcare burdens that result in a poor quality of life in patients, causing pain, loss of mobility, and increased mortality. Therefore, effective medical strategies and appropriate care are crucial for the management of skin wounds and the improvement of wound healing.
To facilitate wound healing and also prevent microbial infection, a number of dressing products for wound treatment and care have been developed by targeting different aspects of the healing process [5
]. In recent years, the use of biological materials as wound dressings has gained momentum, including the use of polysaccharides (chitosan, alginate, fucoidan, etc.) and proteins (keratin, fibrin, collagen, etc.) owing to their biocompatibility, biodegradability, and non-toxic nature [6
]. Collagens are the most abundant structural proteins, existing as a fibril with the basic triple helical structure in the skin, bone, and other connective tissue in mammals. Among the 28 members of collagen, type I collagen is the most abundant in the body. It demonstrates excellent physical and mechanical properties and is considered as a unique dressing material in wound treatment and care [7
Fish collagen is produced from by-products of processing including skin, scales, fins, and bone, which display various biological properties in skin wound healing, engineering, and regeneration [8
]. Collagen from the tilapia facilitates skin regeneration and wound healing by stimulating adhesion, proliferation, migration, angiogenesis, and epidermal differentiation in keratinocytes, fibroblasts, and wounded animals [10
]. This is type I collagen and is comparable to that derived from terrestrial sources, despite being the predominant collagen in skin and scales. Other common types such as types II, III, and IV are distributed in other tissues of fish and may present different beneficial and biochemical properties in wound healing. Type II collagen is also a fibril-forming collagen, with a different polypeptide chain composition compared to type I collagen [13
]. As the main component of cartilage, type II collagen is important for providing tensile strength to the tissue [14
]. Type II collagen from marine materials such as tilapia scale, chum salmon, Rohu, Catla, shark skin, and cartilage has been used as a biomaterial or in combination with other functional scaffolds that are widely applied to bone and cartilage tissue engineering and regeneration [9
]; however, it is less known for its effect on skin wound healing. Sturgeon is one of the most valuable fish worldwide, serving as an important source of meat and caviar, and with considerable processing wastes including the cartilage [16
]. We extracted and identified type II collagen from the cartilage of a sturgeon, Acipenser baerii
, the biological effect of which had not been reported. Here, we aimed to investigate the effect of the collagen extracted from Acipenser baerii
on wound healing in vitro and in vivo. We also included tilapia skin collagen (TSC) as a positive control for the wound healing effect, because it is typical of type I collagen and its wound healing properties are well documented. We demonstrated that collagen from Acipenser baerii
and tilapia skin collagen (TSC) both stimulated human fibroblast proliferation, migration, and wound healing in mice, but with different targeted cell types in the skin tissue.
In the current study, we found that despite SCC sharing a similar amino acid profile to the type II collagen from the cartilage of Amur sturgeon and Siberian sturgeon [17
], the denaturation temperature of SCC is lower than that of these two sturgeons. This may contribute to the difference in amino acid composition and sequence, the number and the nature of covalent crosslinks, as well as the molecular conformation of type II collagen. In addition, in our study, TSC was identified as being composed of one α1 chain and one α2 chain, similar to the result by Li et al. [19
]. The result of amino acid profile showed the content of Pro, Hyp, Gly, Ala, and Lys of TSC was slightly different than that in the reports of Li et al. and Song et al. [19
We further investigated the effect of type II collagen extracted from the cartilage of Acipenser baerii
on wound healing in vitro and in vivo. For the first time, we found coating SCC significantly increased proliferation, migration, and invasion of HDFa. At the molecular level, we demonstrated that SCC regulated HDFa invasion through Akt and MAPK signaling. In addition, coating SCC stimulated HDFa to increase gene expressions involved in ECM production and remodeling, including Col-Iα1, Col-IIIα1, elastin, Has 2, and MMP-1. It is suggested that the stiffness and composition of ECM is involved in regulating cell behaviors of fibroblast, including morphology, proliferation, differentiation, and migration [31
]. Xie et al. demonstrated a lower proliferative rate of human mesenchymal stem cells on collagen with stiff fibers than that with soft fibers [33
]. In addition, Stylianou et al. reported that an increase of collagen concentration elevated fiber density, stiffness, and contributed to an increase in cell spreading of fibroblasts [34
]. In our study, we found the proliferation of HDFa was decreased by coating SCC at high concentrations, whereas the migration and invasion ability was increased, maybe due to the substrate stiffening.
Numerous studies have suggested the effect of tilapia type I collagen on the modulation of fibroblast proliferation and function as well as wound healing. Song et al. demonstrated that collagen from tilapia skin stimulated L929 fibroblast proliferation [26
]. Electrospun collagen nanofibers were developed from tilapia skin and were shown to stimulate adhesion, proliferation, migration, and epidermal differentiation in human keratinocytes, as well as to facilitate wound healing in rats [11
]. Other studies also reported the in vivo wound accelerating effect of tilapia type I collagen-derived peptide and tilapia type I collagen incorporated with other polymers [11
], but the molecular mechanism of action remains unclear. Here we showed coating TSC significantly promoted proliferation, migration, and invasion of HDFa, in agreement with results of previous studies. The mechanism by which TSC promoted the invasion of HDFa was similar to that of SCC. Notably, TSC dramatically upregulated MMP-1 in HDFa, compared to coating SCC. MMP-1 is shown to cleave types I, II, and II collagen, and type I is the preferential substrate [37
]. Coating TSC (type I collagen) may stimulate the upregulation of MMP-1 in HDFa to further ECM degradation during tissue repair. Moreover, our in vivo study showed that the dorsal wound administration of SCC or TSC facilitated the wound healing process in mice, with reduced inflammation, increased fibroblasts proliferation, ECM deposition, and re-epithelialization. The application of both SCC and TSC enhanced elastin expression at day 5 to day 10, as well as the deposition of collagen in the wound of mouse skin, indicating their ability to recover the function of wounded skin [38
A number of growth factors, cytokines, and chemokines, such as VEGF, bFGF (also known as FGF-2), TGF-β, and CTGF, have been reported to regulate wound healing by targeting various cell types including endothelial cells, keratinocytes, and fibroblasts. They form a complex signaling network to influence cellular behaviors and play multiple roles during the healing process [39
]. Chen et al. reported that the application of type I collagen from tilapia skin on dorsal wounds of SD rats accelerated wound healing by reducing inflammatory infiltration, promoting fibroblasts proliferation, collagen synthesis, and re-epithelialization via upregulation of EGF, FGF, and CD31 in the skin tissue [10
]. In rabbits with burn wounds applied with tilapia skin, collagen-derived peptides, combined with chitosan, demonstrated healing by promoting re-epithelialization, collagen fiber deposition, and the upregulation of FGF2 and VEGF in the skin tissue [40
]. In our current study, we observed that the topical application of SCC and TSC significantly increased the levels of the above four growth factors in the skin wound, with a clearly different expressing pattern. TSC application resulted in abundant growth factor expression, mostly in the epidermis (rich in keratinocytes) and partially in the dermis (rich in fibroblasts). In contrast, a high expression of growth factors was found in the dermis, dermal, and subcutaneous white adipose tissue (consisting of preadipocytes and adipocytes) following SCC administration, and also in the epidermal layer. It is known that collagens and their interaction with other ECM components such as elastin, laminin, fibronectin, proteoglycans, and glycosaminoglycans form 3D fibrous networks that influence cell growth, attachment, migration, and differentiation [41
]. The differences in amino acid composition, sequence, covalent cross-links, structural assemblies between type I (TSC) and type II (SCC) collagen may influence their structural and biological features by interacting with different ECM constituents, cellular receptors, growth factors, and cytokines, thus leading to create individual proper environments for adhesion, proliferation, and migration of various cell types participating in wound healing. It has been suggested that type I collagen forms super-twisted microfibrils and links to neighboring microfibrils, creating the quasi-hexagonal crystalline structure [43
]. Type II collagen displayed the conformation of well-ordered cross-linking of nonhelical telopeptides. The wide cross-linking between type II collagen fibrils seems to be more stable than type I collagen [44
]. The distinct structural features of type I and type II collagen may provide different biological properties for the cellular behaviors in wound healing. In fact, the different nature of surface morphology and topography is found to influence a wide range of behaviors in various cells [45
]. However, this hypothesis requires further investigation.
The major challenge of the present study is the different conditions of collagen preparation for SCC and TSC. SCC was prepared by pepsin digestion that resulted in missing telopeptide regions of tropocollagen, but not in the case of TSC. Studies have suggested telopeptides are critical for the assembly and stabilization of collagen fibrils [47
]. In future work, we will prepare pepsin-solubilized TSC and compare it with SCC to identify specific cell and molecular target for accelerating wound healing.
4. Materials and Methods
4.1. Extraction and Coating of Collagen
The SCC and TSC were prepared with a slightly modified method of Liang et al. [17
] and Li et al. [19
]. The cartilage was washed, chopped, and stirred in 0.1 M NaOH solution and 0.5 M ethylenediaminetetraacetic acid (EDTA) (pH 7.4) at 4 °C for 1 day, respectively. The cartilage samples were then soaked in 0.01 M HCl solution including 0.1% pepsin (w
) at 4 °C for 12 h with continuous stirring followed by filtration through 5B filter paper. The supernatant was salted-out by adding NaCl (final concentration of 2 M) at 4 °C and centrifuged at 2500× g
for 30 min, then we collected the precipitate which was collagen. The cartilage tissue on the filter paper was repeatedly extracted according to the above method until it no longer contained residue. For preparing of TSC, tilapia skin was stirred in 0.1 M NaOH at 4 °C for 60 min and then was immersed in 0.5 M acetic acid with a solid/liquid ratio of 1:50 (w
) for 24 h. After centrifugation, the supernatant of tilapia skin was salted-out as described above. Collagen samples were dialyzed with deionized water in a MWCO 20 KDa dialysis membrane at 4 °C for five days then freeze-dried to obtain SCC and TSC for subsequent use. In the in vitro study, the indicated concentration of collagen was added in the culture dishes or well until the dish contained no water; then the pre-coated culture dishes or well could be used.
4.2. Characterization of SCC and TSC
The amino acid content of the collagen was determined using high-performance liquid chromatography (HPLC) (Agilent 1260). The denaturation temperature of the collagen was measured with differential scanning calorimetry (DSC) (200 F3, NETZSCH, Selb, Germany). The surface morphology of the collagen was determined with scanning electron microscopy (S3000N, HITACHI Ltd., Tokyo, Japan). For sodium-dodecyl-sulfate gel electrophoresis, collagen samples were dissolved in 0.5 M acetic acid. Electrophoresis was performed on an 8% resolving gel and 5% stacking gel. Proteins were stained with Coomassie brilliant blue R-250, then destained using a solution containing water, methanol, and acetic acid (13:5:2, v/v/v).
4.3. Cell Culture
Mouse embryo fibroblast NIH-3T3 cells and human keratinocytes HaCaT cells were purchased from American Type Culture Collection (Rockville, MD, USA) and grown in DMEM including 2 mM glutamine (Gibco BRL, Grand Island, NY, USA), 10% (v/v) fetal bovine serum, and 1% penicillin/streptomycin (10,000 units of penicillin/mL and 10 mg streptomycin/mL). The human dermal fibroblast adult cell (HDFa cells; a kind gift from Dr. Hsing-Chun Kuo) were cultured in fibroblast medium (FM) (ScienCell, no. 2301) supplemented with 2% (v/v) fetal bovine serum, 1% penicillin/streptomycin, and 1% fibroblast growth supplement. These three cells all incubated at 37 °C in a 5% CO2 humidified atmosphere.
4.4. Trypan Blue Assay
HaCaT cells, NIH-3T3 cells, and HDFa cells were seeded on SCC- or TSC-coated 24-well plates at a density of 1 × 105 cells/well, 2.5 × 104 cells/well, and 2.5 × 104 cells/well, respectively. After being incubated at 37 °C in a 5% CO2 humidified atmosphere for 36 h, cells were washed with sterile PBS, treated with 0.25% trypsin–EDTA, and harvested. Cells were stained with 0.4% trypan blue and then counted under a light microscope.
4.5. Cell Invasion Assay
To investigate the effect of SCC and TSC on cell invasion in HDFa, the insert of transwell chambers (Corning Inc., Corning, NY, USA) were pre-coated with indicating concentration of SCC or TSC. HDFa (1 × 105 cells/mL) in 100 µL serum-free medium were added to the upper compartment, and the lower chamber was added 600 µL FM containing 2% FBS. Then the cells were incubated for 16 h at 37 °C with 5% CO2. To observe whether MAPKs and Akt signaling were involved in TSC and SCC-mediated HDFa invasion, cells were placed on the insert which was pre-coated with indicating concentration of SCC or TSC for 1 h, and then inhibitors were added and incubated for a further 15 h. The inserts were fixed in 4% paraformaldehyde for 5 min. The cotton swab was used to remove the non-invaded cells in the upper compartment and stained with 0.1 µg/mL DAPI for 30 min. Under the microscope, the cells were counted in five random sights.
4.6. Scratch Wound Healing Assay
To investigate the wound healing effect of SCC and TSC on HDFa, the cells were placed in indicating concentration SCC-coated or TSC-coated 12-well plates at a density of 1.5 × 106 cells/well for 12 h. After cells formed a confluent monolayer, they were scratched using a sterile pipette tip to create a wound and washed by PBS to remove cell debris, and then incubated with fresh medium for 12 h. Images of the migrated cells were taken with an inverted microscope (Olympus) at 0, 4, 8, and 12 h. The wound area was quantified by Cellsens software in each group. Wound healing closer (%) = (original size of the wound area – wound area)/original size of the wound area × 100%.
4.7. Real-Time Polymerase Chain Reaction (qRT-PCR)
Total RNA was isolated from the HDFa cells using Quick-RNA™ MiniPrep kit (ZYMO RESEARCH Inc, Irvine CA, USA) following the manufacturer’s protocol. The gene expression of the cell migration marker (N-cadherin, Snail, and MMP1) and ECM deposition (has2, collagen Iα1, collagen IIIα1, and elastin) in HDFa was detected using real-time PCR. Glyceraldehyde 3-phosphate dehydrogenase (GAPDH) was used as an internal control. Primers for gene analysis are listed in Table S4
. RNA was reverse transcribed using SuperScript™ III First-Strand Synthesis SuperMix for qRTPCR in accordance with the manufacturer’s instructions. Sample cDNA was subjected to qPCR reactions with an StepOnePlus™ real time system (Applied Biosystems, Foster City, CA, USA) using a fast protocol with universal cycling conditions (5 min at 95 °C, followed by 40 cycles of 20 s at 94 °C, 30 s at 60 °C, and 30 s at 72 °C). Real-time PCR was performed in a final reaction volume of 25 µL containing 2 µL; cDNA, 12.5 µL Fast SYBRTM
green master mix (Roche, Rotkreuz, Switzerland), 0.4 µL F primer, and 0.4 µL R primer. Primers were used at a final concentration of 10 mM in the reaction mix. Data was analyzed using the Applied Biosystems 7500 software suite with differential expression determined by the Comparative CT method (ΔΔ
4.8. Excisional Wound Model
Forty male C57BL/6J mice (8–10 weeks old) were purchased from the BioLASCO Taiwan Co., Ltd. (Taipei, Taiwan). The mice were housed and maintained at controlled conditions with a 50% relative humidity, 12 h light/ dark cycle, and 22 ± 1 °C. All animals were free access to water and a standard diet. The animal study was approved by the Institutional Animal Care and Use Committee of Chiayi Chang Gung Memorial Hospital (Affidavit of Approval of Animal Use Protocol No. 2018062505; date of approval, 08 December 2019). After one-week acclimation, mice were randomly divided into the following five groups (n = 8): Control, SCC, and TSC (100 or 500 μg/cm2). Mice were anesthetized by intramuscular injection of Zoletil 50 (Virbac Taiwan Co., Ltd, Taipei, Taiwan) mixed with Rompun (Bayer, Leverkusen, Germany) (4:1), and two 8-mm wounds were made in their dorsal skin by biopsy punch. The wound area was treated with the indicated concentration of SCC or TSC then covered with the 3M™ Tegaderm™, and the dressings were changed every two days. The area of the skin wound was calculated by daily digital photography. The animals were sacrificed at 5 and 10 days after surgery via an overdose of isoflurane, and the skin tissues were collected and fixed in formalin, embedded in paraffin, and sectioned for histopathological examination and immunohistochemistry.
4.9. Histopathological Examination and Immunohistochemistry
The wounded skin samples were paraffin-embedded and cut into 3-μm-thick sections, stained with hematoxylin and eosin (H&E) or Masson’s trichrome stain and examined under a light microscopy (Olympus, Tokyo, Japan). For immunohistochemical staining, the wound skin sections were heated in a retrieval buffer (citrate buffer, pH 6.0) for 15 min to increase immunoreactivity. The sections were then incubated in a peroxidase-blocking solution for 10 min. The ECM deposition and growth factor in wounded skin were detected by incubating primary elastin, VEGF, bFGF, TGF-β, and CTGF antibodies for 1 h and subsequently polydetector HRP for 30 min. The immunostaining was performed by using a Mouse/Rabbit PolyDetector HRP/DAB Detection kit (BioSB, Santa Barbara, CA, USA). The ImageJ 1.46v software (US National Institutes of Health, Bethesda, United States) was used to quantify the immunostaining in each section.
4.10. Statistical Analysis
Statistical Analysis System (SAS) v 9.1 software was used for statistical analysis. Results were presented as means ± SE. The significant differences among the groups were performed using one-way ANOVA test and further with post-hoc Duncan’s multiple-range test. A p-value < 0.05 was considered statistically significant.