Physical decellularization of fish skin utilizing electrical fields

TL;DR

A new method using electrical fields to decellularize fish skin preserves tissue structure and biocompatibility better than traditional chemical methods.

Contribution

Introduces a fast, chemical-free electrical decellularization method that preserves tissue integrity and reduces immunogenicity.

Findings

Electrical decellularization achieved DNA levels below 50 ng/mg in 2 hours without toxic reagents.

Electrically decellularized skin showed better microstructure and higher cell survival rates than chemical methods.

The method caused minimal hemolysis and elicited fewer immune responses in implantation tests.

Abstract

Decellularized tissues have attracted considerable attention in tissue engineering and regenerative medicine due to their diverse sources and excellent biocompatibility. However, current decellularization techniques often compromise the integrity of the extracellular matrix, leaving harmful chemical residues or inadequately removing immunogenic cellular components. Consequently, the biocompatibility and clinical efficacy of decellularized tissues are undermined. To address these issues, a novel decellularization technique employing an electrical field has been proposed. In the resulting decellularized tissue, the residual DNA concentration was measured at 27.44 ± 7.27 ng/mg, satisfying the evaluation criteria (<50 ng/mg) in a significantly shorter process compared to chemical/enzymatic decellularization (∼2 h vs. ∼15 h) and without the use of toxic reagents. The microstructure was better preserved in electrically decellularized skin than in chemically/enzymatically decellularized skin, maintaining an interconnected porous microstructure that facilitates neo-tissue ingrowth. Cytotoxicity assessments confirmed the non-cytotoxic nature of electrically decellularized fish skin, with a cell survival rate (115.84 ± 10.78%) higher than that of chemically/enzymatically decellularized and native skins (105.57 ± 3.43% and 118.69 ± 6.53%, respectively). The hemolysis rate of decellularized skin using electrical techniques was the lowest (0.35 ± 0.26%) compared to that of chemically/enzymatically decellularized and native skins (1.089 ± 0.03% and 1.11 ± 0.06%), indicating non-hemolytic material. Furthermore, subcutaneous implantation demonstrated that electrically decellularized skin did not elicit severe immune responses and exhibited faster tissue integration than chemically/enzymatically decellularized skin. Therefore, electrical decellularization effectively removes cellular components while preserving the natural architecture without the use of harmful reagents, establishing a secure foundation for extensive applications in tissue engineering and regenerative medicine using decellularized tissues in the future.

Figures

Scheme for preparation and characterization of decellularized fish skin utilizing an electrical field.

Flowchart depicting the process of fish skin decellularization. The concentrations and treatment time of the pretreatment solutions for decellularization are presented.

Evaluation of the efficiency of fish skin decellularization. (A) Optimization of current for decellularization (n = 3); (B) representative H&E staining images for fish skins (n = 3); (C) representative electrophoresis image of distribution of DNA fragments in non-decellularized fish skin and decellularized skin (n = 3); (D) quantification of DNA in non-decellularized and decellularized skin (n = 3). P < 0.05 was considered a significant difference (ns, no significant difference; *P < 0.05, *P < 0.001, and P < 0.0001).

Characterization of the chemical and physical properties of the skin. Concentrations of (A) collagen (n = 3), (B) sGAG (n = 3) and (C) elastin (n = 3) in skins; (D) representative SDS-PAGE patterns of fish skin and mouse tail collagen were used as controls. Molecular markers were added to the leftmost and rightmost lanes. The bands corresponding to collagen α1 and α2 chains are shown in all lanes. (E) Optical and SEM images of fish skin. (F) Representative stress–strain curves, elastic modulus (E), maximum tensile stress (σ) and strain at break (ε) of skins. (n = 5). FC, fish skin collagen; MC, mouse tail collagen. P < 0.05 was considered a significant difference (ns, no significant difference, P < 0.0001).

Biocompatibility of skin in vitro. (A) Cytotoxicity of skin (n = 3); (B) representative live/dead staining of cells (n = 3); (C) proliferation of cells cultured in skin (n = 3); (D) hemocompatibility of skin (n = 3). P < 0.05 was considered a significant difference (ns, no significant difference; P < 0.01, and P < 0.0001).

Biocompatibility of skin in vivo. (A) Representative H&E staining images of harvested subcutaneous mouse tissue surrounding implanted fish skins with different treatments after 7, 14, 21 and 28 days (n = 6); (B) H&E staining images of major organs of mice receiving various treatments (n = 6). NC, sham.

Tables

Table 1. Amino acid compositions of decellularized (CD and ED) and non-decellularized tilapia skins (ND).

Amino acids	Composition (g/100g)
	ND	CD	ED
Asp	6.02	5.86	5.69
Thr	2.90	2.86	2.86
Ser	3.51	3.56	3.50
Glu	10.26	10.43	9.97
Gly	23.23	25.16	23.92
Ala	10.22	11.12	10.03
Cys	1.72	1.65	1.71
Val	2.19	2.07	2.10
Met	1.55	1.61	1.27
Ile	1.37	1.23	1.28
Leu	3.12	2.87	2.92
Tyr	0.86	0.72	0.65
Phe	2.08	2.00	1.96
His	0.83	0.79	0.68
Lys	3.33	3.30	2.96
Arg	8.30	8.59	7.72
Pro	10.33	10.97	11.07

Full text

Introduction

Decellularization techniques used for biological scaffold preparation have been widely applied in regenerative medicine [1–3], attracting considerable attention due to their advantageous biocompatibility, biodegradability and bioinductive properties, thereby offering alternate solutions to the scarcity of donor tissues and organs [4–6]. According to the latest research report, the global decellularization therapy market had a revenue value of $8.45 billion in 2022 and is expected to exceed ∼$32.61 billion by 2030, with a compound annual growth rate of 16.21% from 2023 to 2030 (Zion Market Research). The process of decellularization entails the removal of immunogenic cellular components from donor tissues or organs while preserving the original complex ultrastructure of extracellular matrix (ECM) and bioactive components. This results in decellularized materials categorized by low immunogenicity and excellent biocompatibility, thereby enhancing regeneration of neo-tissue [2, 7, 8]. Decellularized tissues can serve as natural biomaterials for preparing tissue engineering scaffolds to promote cell attachment, proliferation and differentiation, contributing to the reconstruction of damaged tissues and organs [9–11], or can be used to prepare carriers for delivering drugs or bioactive macromolecules to treat tumor tissues or in situ recruited cells, thereby facilitating the repair of impaired tissues [12]. Among various source materials of decellularized tissues, fish skin is considered a promising candidate recently due to its abundant supply, minimal religious restrictions, waste recycling potential, and reduced risk of zoonotic disease transmission between mammals [13]. Tilapia is farmed in many countries due to its fast breeding, high protein content and high resistance to disease. The wide geographical distribution and fast maturity of tilapia allow it to be produced on a large scale and at a low cost. The skin of tilapia is rich in type I collagen, and there have been numerous studies on the extraction, purification and characterization of tilapia collagen identified as a potential biomaterial which exhibited similar bioresorption rates and inflammatory responses to porcine collagen sponges. Decellularized tilapia fish skin (DFS) is highly valued not only for its relatively high mechanical strength but also for being a natural source rich in type I collagen and other bioactive components, making it a more environmentally friendly alternative to mammalian or synthetic scaffolds [14–16]. Consequently, the applications of DFS in reconstructive surgery and regenerative medicine is becoming increasingly widespread [17, 18].

DFS is an emerging biomaterial in regenerative medicine. There is a scarcity of literature and patents specifically addressing its preparation methods, which are generally inspired by the decellularization techniques of mammalian tissues for processing fish skin [13], including physical, chemical and enzymatic methods. However, each of them possesses unique advantages and disadvantages, such as the efficacy of immunogenic component removal, residuals of decellularization reagents, the extent of ECM damage and the retention of bioactive compounds within the tissue, among others. Decellularization aims to achieve an optimal balance by removing immunogenic cellular components while retaining bioactive elements, such as collagen, glycosaminoglycans and growth factors, and preserving structural integrity [19]. Structural integrity allows the decellularized tissue to maintain its natural three-dimensional (3D) architecture, thereby providing a cell-like physical microenvironment analogous to in vivo conditions and ensuring the mechanical stability required to enhance tissue regeneration. Immunogenic cellular components, such as deoxyribonucleic acid (DNA), galactose-alpha-1,3-galactose (α-Gal) epitopes and major histocompatibility complex I, are perceived by the host as foreign substances, leading to the activation of immune-mediated rejection responses [20–22]. The content of CpG (a specific region in a DNA molecule where a cytosine (C) nucleotide is followed immediately by a guanine (G) nucleotide) and the degree of methylation in DNA vary among different species; therefore, it has been proposed that CpG motifs and cytosine methylation may enable the immune recognition system to identify foreign DNA and induce an immune response [23]. Their removal can mitigate the immune reaction of the host following the implantation of decellularized tissue, thereby enhancing its clinical efficacy. Naturally occurring bioactive components within tissues can regulate cellular states in vivo. For example, collagen participates in the regulation of cell growth through multidimensional mechanisms, including structural support, signal transduction and mechanical modulation [4]. Omega-3 fatty acids in Atlantic cod skin enhance cell migration and have anti-inflammatory properties, analgesic effects and antimicrobial activity [24]. Retaining bioactive components directly provides a unique functional bio-scaffold that combines natural structure with bioactive constituents to support tissue regeneration and improve clinical outcomes ultimately [17, 18]. Generally, the most significant advantage of the physical decellularization method is the absence of residual substances, resulting in no exogenous cytotoxicity of decellularized tissue [25]. However, its drawback is insufficient decellularization efficiency when used alone. Conversely, the chemical decellularization method is distinguished by its remarkable efficiency and thoroughness. Nonetheless, if the subsequent washing process is insufficient, chemical residues may persist in the decellularized tissue, potentially leading to exogenous cytotoxicity [26]. Enzymes can remove specific cellular components; however, solely relying on enzymes makes it challenging to fully eliminate cellular elements, and residual enzymes can disrupt the follow-up recellularization of decellularized tissue, provoking severe immune responses. Therefore, the currently used methods for decellularization of fish tissues usually involve a combination of two or three techniques to achieve optimal effects. How to remove chemical and immunogenic cell residues while preserving the structural integrity and bioactive components of natural tissues remains a challenge in decellularization technology. Innovations in technology for fish skin decellularization will strengthen the inherent advantages of DFS and improve its clinical efficacy.

In this study, we developed a clean decellularization technology that utilizes electric fields without harmful decellularization reagents to rapidly and effectively prepare DFS, as shown in Figure 1. Using this technology, the cellular components in native tissues are effectively and rapidly removed while preserving the natural microstructure and constituents. DFS demonstrated excellent biocompatibility both in vitro and in vivo. The development of clean decellularization technology for processing fish skin represents a promising advancement. This technology includes (i) a physical approach that primarily utilizes electric current to directly eliminate cellular components, thereby fundamentally avoiding the import of exogenous toxic reagents; (ii) a tremendous reduction in decellularization time from 15 h to approximately 2 h, resulting in an efficiency improvement; (iii) a DNA removal mechanism combining electroporation and electrophoretic migration was proposed, providing a theoretical foundation for the application of electric fields in the preparation of decellularized tissues. This advancement not only suggests a more rapid fabrication but also significantly mitigates the risk of matrix degradation associated with prolonged processing, thereby better preserving the native tissue activity. By leveraging the natural properties of fish skin combined with advanced processing technologies, this development will contribute to realizing the possibility of providing sustainable and biocompatible regenerative biomaterials for clinical applications, marking a significant step toward environmentally friendly practices in tissue engineering and aligning with broader sustainability objectives.

Scheme for preparation and characterization of decellularized fish skin utilizing an electrical field.

Materials and methods

Chemical and biological reagents

Sodium dodecyl sulfate (SDS), hematoxylin and eosin (H&E) staining kit, BCA protein assay kit and SDS-PAGE gel kit were procured from Beijing Solarbio Science & Technology Co., Ltd., China. Sircol insoluble collagen, Blyscan sulfated glycosaminoglycan (sGAG) and Fastin elastin assays were obtained from Biocolor Ltd (Belfast, UK). Concanavalin A (ConA), MTT cell proliferation and cytotoxicity assay kit, and cell counting kit-8 (CCK-8) were acquired from Beyotime Biotech Inc, China. 1-Ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC), N-hydroxysuccinimide (NHS), LIVE/DEAD Viability/Cytotoxicity Kit, Invitrogen Quant-iT PicoGreen dsDNA assay kits, dispase, phosphate-buffered saline (PBS), fetal bovine serum (FBS), Dulbecco’s Modified Eagle Medium (DMEM), Roswell Park Memorial Institute (RPMI) 1640 medium, penicillin-streptomycin (PS), and Pierce universal nuclease for cell lysis were sourced from Thermo Fisher Scientific Inc (Waltham, MA, USA). The TIANamp marine animal DNA kit was sourced from Tiangen Biotech Co., Ltd., China. Other reagents were obtained from Shanghai Macklin Biochemical Co., Ltd., China, unless otherwise specified.

Preparation of decellularization of fish skin

Preparation of skin

Fresh fish skin (12 cm × 8 cm) harvested from descaled Nile tilapia (Oreochromis niloticus) purchased from Maoming Aquatic Technology Co., Ltd. (Guangzhou, China). The skins were washed with deionized (DI) water, cut into small pieces (4 cm × 4 cm) and lyophilized, known as non-decellularized skin (ND), stored in a drying cabinet for later use.

Chemical/enzymatic decellularization

Decellularization was conducted in accordance with our previous studies [16], with certain modifications as shown in Figure 2. Initially, ND samples (5 cm × 5 cm, ∼500 mg each) were immersed in PBS at 4°C for a duration of 2 h. Subsequently, the samples were treated with 2.5 U/mL dispase in PBS for 3 h. Following a wash with DI water, the samples were immersed in a 1% SDS-PBS solution for 6 h. After another wash with DI water, the samples were agitated in a 25 U/mL Pierce universal nuclease PBS solution for 3 h to facilitate nucleic acid decomposition. Finally, the samples were immersed in a 1% SDS-PBS solution for 1 h. The samples were then washed with DI water and subjected to lyophilization for 72 h. The resultant product, referred to as chemically decellularized skin (CD), was stored in a dry environment at room temperature (RT) and 40% humidity.

Flowchart depicting the process of fish skin decellularization. The concentrations and treatment time of the pretreatment solutions for decellularization are presented.

Electrical decellularization

As shown in Figure 2, the lyophilized ND samples (5 cm × 5 cm, ∼500 mg each) were pre-soaked in sodium chloride (NaCl) solution (1.5 M) for 2 h and then mounted on the plate-shaped electrode (5 cm × 5 cm) surfaces. The electrodes were positioned parallel to each other in an electrolysis chamber (Ariter, C001, 500 mL, China), which consisted of a transparent glass body and a polytetrafluoroethylene cover; the distance between the electrodes was fixed at 36 mm. A sine wave alternating current with a maximum current of 6A, and a frequency of 2.5 mHz produced with a waveform generator (Vpp = 10, RIGOL, DG1022U, China) and a power amplifier (Aigtek, ATA-304, China) was used for decellularization at RT. The process was performed for 5 min. The samples were then washed with DI water and lyophilized for 72 h. The final product, known as electrically decellularized skin (ED), was stored in a dry box at RT and 40% humidity.

Crosslinking and sterilization of fish skin

The skins underwent crosslinking using NHS/EDC chemistry to enhance the mechanical properties of tissues and prevent rapid degradation, thereby facilitating long-term cell culture, as previously documented [27]. In brief, the skins were immersed in a crosslinking solution comprising 25 mM EDC and 12.5 mM NHS in an 80/20 acetone/phosphate buffer solution for a duration of 2 h. Subsequently, the skins underwent multiple washes for 2 h in 0.01M PBS, followed by an additional 2-h wash in DI water as described in previous reports [27, 28]. The cross-linked skins were then lyophilized, sterilized with ethylene oxide and aeration for 2 days. Eventually stored in a drying oven, and designated as ND-C, CD-C and ED-C, corresponding to the specific skins (ND, CD and ED) used for crosslinking.

Physical and chemical properties of skin

Morphology examination

The lyophilized ND, CD, and ED were cut into 5 mm × 5 mm square samples, sprayed with gold using a sputtering coater (50 mA, 90 s), and photographed using a scanning electron microscope (SEM) (Phenom, Phenom Pharos, Holland) with an acceleration voltage of 5 kV.

Histological examination

In vitro experiment, the ND, CD and ED were cryo-embedded and frozen in liquid nitrogen. Slices with a thickness of 5 µm were prepared using a cryostat (HM525NX, Thermo Fisher Scientific Inc., USA). In vivo experiment, tissues harvested from mice were fixed in 10% neutral buffered formalin, then paraffin‐embedded and sectioned using a microtome (Leica RM2255, Germany). The hematoxylin stains cell nuclei blue, and eosin stains the ECM and cytoplasm pink [29]. The slices were scanned and photographed using a Digital Microscope (Nexcope-NSS-6-Digital, View Solutions Inc., USA).

Quantification of residual DNA

Following lyophilization, the ND, CD and ED samples were weighed to approximately 10 mg. DNA was extracted using the TIANamp Marine Animals DNA Kit and quantified with the Quant-iT PicoGreen dsDNA Assay Kit. Fluorescence intensity at 480 nm was measured using a microplate reader (Varioskan LUX, Thermo Fisher, USA). The distribution of DNA fragments was assessed via gel electrophoresis. DNA fragments were separated on a 1% (w/v) agarose gel at 100 V, utilizing a red fluorescence nuclear stain (E1020; Solarbio, China) for visualization. A 100 bp DNA marker (BDL-036, TOROIVD, UK), ranging from 100 to 1500 bp, was employed to reveal the DNA fragments present in the ND, CD, and ED.

Quantification of collagen

To quantify the collagen content in ND, CD and ED, a Sircol insoluble collagen assay kit was employed to measure the collagen levels in each sample, following the manufacturer’s instructions. To assess the impact of decellularization on the integrity of collagen chains in ND, CD and ED, SDS-PAGE was conducted on the samples and controls (rat tail and tilapia skin collagens) as previously described, with certain modifications [30, 31]. The gels were run for 50 min at 120 V. Following electrophoresis, the gel was then stained with Coomassie brilliant blue staining solution (P1300, Solarbio, China) and photographed the distribution of each band in relation to the marker bands using a Tanon Gel Imaging System (3500BR; Tanon Science & Technology Co., China).

Quantification of elastin and sGAG

To quantify the elastin and sGAG contents in ND, CD and ED, the Fastin elastin and Blyscan sGAG assays were employed to measure elastin and sGAG levels in each skin sample, following the manufacturer’s instructions.

Quantification of amino acid

The amino acid compositions of the skin samples (ND, CD and ED) were identified and quantified using an automatic amino acid analyzer (Biochrom 30+ series, Biochrom, Cambridge, UK). In brief, 50 mg of skin was subjected to hydrolysis in 4 mL of 6 N HCl at 110°C for 22 h under a nitrogen atmosphere. The hydrolysate (2 mL) was dried. Finally, mixed well with 0.02 M HCl on a vortex mixer and passed through a 0.45-µm filter column before loading onto the analyzer.

Mechanical property characterization

The mechanical properties of ND-C, CD-C and ED-C were evaluated through tensile testing. Specimens were prepared by cutting the skins into strips (1 cm × 5 cm), and uniaxial tensile tests were conducted using a Universal Testing System (5594, Instron, USA). Prior to testing, the thickness and gauge length of the strips were documented. The strips were subjected to a uniaxial stretch at a rate of 10 mm/min until failure occurred. The maximum tensile stress, strain at break and elastic modulus, derived from the tensile testing software, were recorded and compiled. For each set of samples, the test was repeated at least 5 times.

Degradation characterization

The method for determining the skin (ND-C, CD-C and ED-C) degradation profile was based on our previous report [16] with some modifications. The fish skin was lyophilized and cut into 1 × 1 cm samples (n = 6). First, the initial weight (W_0_) was recorded, and the sample was soaked in PBS. The samples were retrieved and washed three times with DI water on D1, D3, D7 and D14, followed by lyophilization. The weights of the samples (W_t_) were recorded. The degradation profile was defined as the percentage of residual weight, calculated using the following formula:

Percentage weight remaining (%) = (W_t_/W0) × 100%,

where W0 represents the initial weight of the sample and W_t_ represents the weight of the degraded sample after the respective time points.

Biological property of skin

Cytotoxicity of skin

The cytotoxicity of the skin was evaluated using an indirect contact test in accordance with ISO-10993-5-Biological evaluation of medical devices with some modifications. In brief, following sterilization with ethylene oxide, the skins were cut into small pieces and soaked in culture medium (DMEM with 10% FBS and 1% PS) at a ratio of 6 cm^2^/mL, and then placed in a cell incubator (37°C, 5% CO_2)_ for 24 h to prepare the extract medium. The 5 × 10^3^ cells (L929, ATCC, USA)/well were seeded in 96-well plates and incubated for 24 h. The culture medium in the 96-well plates was removed, and the cells were rinsed twice with 200 μL PBS. Fish skin extract medium was added to the 96-well plates and incubated for 24 h. CCK-8 solution (10 μL) was added to each well and incubated in the incubator for 2 h. Subsequently, absorbance was measured using a microplate reader at a wavelength of 450 nm. In addition to the quantitative analysis of skin cytotoxicity, a live/dead assay was conducted to qualitatively evaluate the cytotoxicity of the skin [32]. Cells were cultured in either culture medium or extract medium for a duration of 3 days, subsequently washed with PBS, and stained with Calcein AM and ethidium homodimer. Cell viability, determined by membrane integrity, was examined using a confocal microscope (A1, Nikon, Japan).

Cell proliferation in skin

The proliferation of skin cells (HaCaT, # 339817, BeNa Culture Collection, China; HSF, FH0189, FuHeng Biology, China) cultured in skin was assessed by the amount of DNA, which is related to the number of cells in a linear proportional manner, as described in our previous report [32]. Cells were collected at different time points and lysed. The amount of DNA in the skin was examined using Invitrogen™ Quant-iT™ PicoGreen™ dsDNA assay kits as previously mentioned.

Hemolysis test

According to ISO 10993-4:2017, the hemocompatibility of the skin was assessed in vitro as previous report [15]. In brief, ND-C, CD-C and ED-C extracts were prepared by immersing them in PBS at 37°C for 72 h, with an extract ratio of 6 cm^2^/mL. PBS and DI water served as the negative control and positive control, respectively.

Hemolytic rate (%) = ((Ab of test group—Ab of negative control group)/(Ab of positive control group − Ab of negative control group)) ×100%,

where Ab represents the absorbance of the supernatant measured at 545 nm.

Lymphocyte transformation test

BALB/c mice were sacrificed and spleen was extracted aseptic, single cell suspension was prepared by grinding, the cell suspension was collected, centrifuged at 400 g for 5 min, supernatant was discarded, 5 mL red cell lysate was added, and incubated at 4°C for 5 min, RPMI1640 culture medium containing 10% FBS was added to terminate the reaction, and centrifuged at 400 g for 5 min. The supernatant was discarded, and RPMI1640 culture medium containing 10% FBS was added to adjust the cell concentration to 2 × 10^6^ cells/mL. The cell suspension was added to a 96-well plate with 200 μL per well. ConA was added to the positive control group until the final concentration was 5 μg/mL. The experimental group was three kinds of fish skin extracts. The MTT assay was performed according to the manufacturer’s instructions. The absorbances were measured using a microplate reader at a wavelength of 570 nm, with a reference wavelength of 630 nm.

Subcutaneous implantation of skin

The in vivo biocompatibility of the skin was evaluated through subcutaneous implantation, as described in references [33, 34]. The experiments were conducted in compliance with the received approval received from the Ethics Committee of the Wenzhou Institute, University of Chinese Academy of Sciences (WIUCAS24082206). A subcutaneous implantation model was developed utilizing BALB/C mice aged 6–8 weeks, which were randomly assigned to four groups based on the type of skin implanted: NC (sham control, n = 6), ND-C (n = 6), CD-C (n = 6) and ED-C (n = 6). The fish skin samples for the three groups were prepared by cutting into test specimens (10 mm × 12 mm) and sterilized using ethylene oxide prior to implantation. The mice were anesthetized with isoflurane, and their dorsal hair was shaved. A longitudinal incision approximately 1 cm in length was cut in the middle of the back of each mouse; the connective tissue was bluntly dissected, and subcutaneous pouches were created bilaterally along the spine. The test samples were inserted into the subcutaneous pouches of the mice, and the incisions were sutured with 4-0 sutures. The skin was rinsed with a disinfectant postoperatively. The tissues at the implant sites were harvested at four time points: 7, 14, 21 and 28 days post-implantation. In addition, the major organs were harvested at 14 days post-implantation. Samples were fixed, paraffin-embedded, and sectioned. H&E staining was performed for histological analysis.

Statistical analyses

All assays were performed in triplicate or more. Statistical analysis of the raw data was conducted using GraphPad Prism version 8. Results are presented as mean ± standard deviation. Student’s t-tests or one-way analysis of variance were conducted, and P < 0.05 was considered a significant difference (ns, no significant difference; *P < 0.05, **P < 0.01, ***P < 0.001 and ****P < 0.0001).

Results and discussion

Effectiveness of decellularization

Due to the excellent biocompatibility, biodegradability and bioinductivity of decellularized tissue, it has been extensively employed in the fields of regenerative medicine, providing alternative solutions to address the shortage of donor tissues and organs [35]. Decellularization is the process of obtaining decellularized tissue from animals or plants that possess low immunogenic cellular components while maximally preserving the natural ECM structure, mechanical integrity and bioactive components [13, 19]. Reduction of the residual immunogenic cellular components can mitigate the potential of immune response provoked, thereby enhancing the biocompatibility of decellularized tissue. In this study, the electrical current during the decellularization process was first optimized to achieve the desired effect. As shown in Figure 3A, with the increase of current applied during the decellularization process, the amount of residual DNA in DFS gradually decreased. When the current reached 4A, further increases did not result in a significant change in the amount of residual DNA in DFS. To evaluate the effectiveness of fish skin decellularization, H&E staining was performed to visualize the cell nucleus. As shown in Figure 3B, nuclei were observed in the ND, but no nuclei were found in either the CD or ED, indicating that both chemical/enzymatic and electrical decellularization methods effectively removed cell nuclei from tissues [36]. However, in the CD, there were many fractures and relaxations in the collagen fibers, which were not observed in the ND and were fewer in the ED, consistent with previous reports (SDS caused the rupture of collagen fibers) [26]. This indicated that the electrical decellularization process, conducted according to the established experimental parameters, did not affect the arrangement of collagen fibers within the tissue, obviously, thereby preserving the structural integrity of the skin tissue. In addition to qualitative evaluation, we also performed quantitative DNA measurements. As shown in Figure 3D, the DNA content in the skin of ND was 1018.49 ± 45.83 ng/mg, whereas it was 30.95 ± 0.79 ng/mg and 27.44 ± 7.27 ng/mg in the CD and ED, respectively. Both were conformed to the established evaluation criteria (<50 ng/mg) [6]. DNA content decreased significantly, indicating that most of the cellular components in the tissue were removed by either chemical/enzymatic decellularization or electrical decellularization, which was consistent with the H&E staining results. The analysis revealed no significant difference in DNA content between the ED and CD, suggesting that the effects of electrical and chemical/enzymatic decellularization are comparable. Li et al. developed a chemical decellularization protocol for tilapia fish skin using acetic acid, Triton X-100, sodium hydroxide, and hydrogen peroxide, resulting in a residual DNA content of 1.4 ± 0.7 ng per mg of dry decellularized skin [15]. Lau et al. employed sodium SDS combined with nucleases, reducing the residual DNA content in decellularized skin to 1.8 ± 0.9 ng/mg [16]. Bora et al. used two chemical reagents, SDS and Triton X-100, respectively, for decellularizing tilapia fish skin, achieving DNA concentrations of 8.65 ± 1.25 ng/mg and 18.05 ± 0.85 ng/mg in the decellularized skin [37]. Although their residual DNA levels were lower than those found in our study and well below the medical industry’s limit of 50 ng/mg, the chemical decellularization process is time-consuming, and there are potential risks associated with residual toxic reagents. The residual DNA fragment distributions in ND, CD and ED were revealed via electrophoresis, as depicted in Figure 3C. The gel lanes are organized as follows: the leftmost and rightmost lanes represent the DNA markers, serving as fragment length standards ranging from 100 to 1500 base pairs; the second lane contains the ND sample; the third lane contains the CD sample; and the final lane contains the ED sample. As demonstrated in the ND lane, an intensive band of DNA fragments was observed, suggesting the presence of DNA in fish skin, consistent with previous findings [38]. However, there were no obvious bands in lanes b and c, indicating that there were no detectable amounts of DNA fragments (100–1500 bp) in either CD or ED [39, 40], and revealed that there was no significant difference between the CD and ED groups on DNA removal efficiency. This result was consistent with those of H&E staining and DNA content measurement. In other words, both chemical/enzymatic and electrical decellularization ensure that the skin can be decellularized and meet the criteria for acceptable tissue decellularization [41, 42]. We hypothesized that the decellularization of tissues observed in the experiments was facilitated by two mechanisms. The first one is the increase in cell membrane permeability. Electroporation is a biophysical phenomenon that results from the application of high-magnitude electric pulses across the cell membrane, which increases cell membrane permeability by the formation of nanoscale defects in the cell membrane [43, 44]. This principle has been applied to the preparation of a few decellularized biomaterials, either alone or in combination with chemical methods, to facilitate tissue decellularization [29, 45, 46]. However, these preparations still require a time-consuming process. In addition to electroporation, as in DNA gel electrophoresis, DNA could move under electrical fields due to its negatively charged nature [47] to achieve decellularization of tissue. To verify the effect of the electric field on the DNA removal from the tissue, we prepared fish collagen membranes according to our previous report [48], and introduced DNA into the membranes by soaking them in a DNA solution. An electric field was then applied under the described electrophoretic decellularization conditions. The residual DNA in the membranes after treatment is shown in Supplementary Figure S1. It was observed that the DNA content in the membranes treated with the electric field was significantly lower than in the two control groups without electric field application (soaked with or without NaCl solution), indicating that the electric field facilitates the removal of DNA from the membranes.

*Evaluation of the efficiency of fish skin decellularization. (A) Optimization of current for decellularization (n = 3); (B) representative H&E staining images for fish skins (n = 3); (C) representative electrophoresis image of distribution of DNA fragments in non-decellularized fish skin and decellularized skin (n = 3); (D) quantification of DNA in non-decellularized and decellularized skin (n = 3). P < 0.05 was considered a significant difference (ns, no significant difference; *P < 0.05, ***P < 0.001, and ***P < 0.0001).

Chemical and physical properties of skin

In addition to cells, the primary constituents of the skin also comprise an ECM assembled from various macromolecules, including collagen, sGAG, elastin and other glycoproteins. The amounts of the three main components, collagen, elastin and sGAG, in the skin were measured and are shown in Figure 4A–C. NaCl pre-soak samples were used to isolate the salt extraction effect. The collagen content in the ND, CD and ED groups was 757.71 ± 47.77 μg/mg, 703.54 ± 10.31 μg/mg, and 752.00 ± 17.08 μg/mg, respectively, with no statistically significant difference among these three groups, indicating that neither chemical/enzymatic decellularization nor electrical decellularization significantly affected collagen content in the skin. Tilapia fish skin is abundant in type I collagen, which consists of two α_1_ chains and one α_2_ chain, thereby forming distinct collagen molecules. Hence, in Figure 4D, the 130-kDa band corresponds to the α_1_ chain, and the 110-kDa band corresponds to the α_2_ chain. Apparently, α_1_ chains formed a more intense band at 130 kDa. The presence of α_1_ and α_2_ chains indicates that the integrity of collagen chains is preserved. However, compared with the ND (4.03 ± 1.16 μg/mg), both decellularized methods resulted in varying degrees of elastin loss, with more loss in the CD (∼2.75 μg/mg) than in the ED (∼1.97 μg/mg). After decellularization, sGAG in both CD and ED also had a certain loss, and the loss in ED (∼3.04 μg/mg) was greater than that in CD (∼1.33 μg/mg). This may be due to the negative charge nature of sGAG, which stems from the presence of sulfate and carboxylate groups. The amino acid compositions of ND, CD, and ED were also measured, and the results are listed in Table 1, expressed as grams of individual amino acid per 100 g of dry skin. The table shows that skins (ND, CD and ED) are rich in Gly (23.23, 25.16 and 23.92 g/100 g, respectively), Pro (10.33, 10.97 and 11.07 g/100 g, respectively), Glu (10.26, 10.43 and 9.97 g/100 g, respectively), and Ala (10.22, 11.12 and 10.03 g/100 g, respectively), which is similar to the amino acid composition found in tilapia collagen [49]. A distinctive characteristic of collagen is the systematic arrangement of amino acids, typically adhering to the Gly-X-Y pattern, where X and Y represent various other amino acid residues distributed across the three chains [48]. Therefore, Gly is a key amino acid in collagen, constituting its primary component and accounting for 20–30% of the total amino acid content in various types of collagens, which is in line with our results (Figure 4A). Figure 4E presents the SEM images depicting the dermal and epidermal morphologies of ND, CD and ED. Additionally, it includes the camera-captured appearances of the epidermal side of the fish skin across the three groups. Consistent with the H&E staining results, both non-decellularized tissue (ND) and decellularized tissue (CD and ED) exhibited analogous multilayered fibrous and porous structures, facilitating cell ingrowth and nutrient mass transfer. The mechanical properties of the skin before and after decellularization are shown in Figure 4F. In comparison to a previous study [15], the maximum tensile strength (σ) and elastic modulus (E) of fish skin (ND-C, CD-C and ED-C) are markedly higher, likely due to crosslinking. Notably, the maximum tensile strength of CD-C surpasses that of ND-C and ED-C, which may be attributed to the chemical reagents employed in the preparation of CD-C. Furthermore, ED-C demonstrates greater extensibility than ND-C and CD-C, possibly owing to its lower content of low-molecular-weight macromolecules and a more intact collagen structure. Supplementary Figure S2 presents the degradation curves of the three groups of dehydrated fish skin immersed in PBS for 0, 1, 3, 7 and 14 days. The findings suggest that the degradation rates of CD-C and ED-C are both lower than that of ND-C, potentially due to the substantial presence of low-molecular-weight macromolecules in ND-C.

*Characterization of the chemical and physical properties of the skin. Concentrations of (A) collagen (n = 3), (B) sGAG (n = 3) and (C) elastin (n = 3) in skins; (D) representative SDS-PAGE patterns of fish skin and mouse tail collagen were used as controls. Molecular markers were added to the leftmost and rightmost lanes. The bands corresponding to collagen α1 and α2 chains are shown in all lanes. (E) Optical and SEM images of fish skin. (F) Representative stress–strain curves, elastic modulus (E), maximum tensile stress (σ) and strain at break (ε) of skins. (n = 5). FC, fish skin collagen; MC, mouse tail collagen. P < 0.05 was considered a significant difference (ns, no significant difference, ***P < 0.0001).

Biological properties of skin

Biocompatibility in vitro

The cytotoxicity of biomaterials refers to the harmful effects that materials may have on cells, potentially leading to cell damage, and is a necessary indicator for evaluating the biocompatibility of biomaterials used in regenerative medicine. The CCK-8 results (Figure 5A) showed that the L929 cells in all groups (ND-C, CD-C and ED-C) demonstrated similar viability to the negative control, indicating the excellent biocompatibility of the skins. However, the viability of cells on CD-C was the lowest among these groups, although the difference was not significant. The cells cultured on ND-C exhibited the highest viability, which may be related to the physical signals and biological signals in their natural ECM. In contrast, cells cultured on CD-C displayed the lowest viability, which may be associated with a certain degree of structural damage to their ECM and residual harmful substances used in the process. Furthermore, to qualitatively study the viability of cells in the skin based on membrane integrity and esterase activity, a live/dead assay was performed. The results are shown in Figure 5B. In alignment with the CCK-8 assay results, the cells cultured with extract culture media derived from skins demonstrated high viability, comparable to the non-cytotoxic negative control group, which was stained green to indicate live cells. The presence of red, representing dead cells, was almost undetectable. These results suggest that the extract culture media derived from both decellularized and ND skin were non-toxic to cells and demonstrate the potential application of decellularized and ND skin in the field of tissue engineering.

*Biocompatibility of skin in vitro. (A) Cytotoxicity of skin (n = 3); (B) representative live/dead staining of cells (n = 3); (C) proliferation of cells cultured in skin (n = 3); (D) hemocompatibility of skin (n = 3). P < 0.05 was considered a significant difference (ns, no significant difference; **P < 0.01, and ***P < 0.0001).

The vigorous proliferation of cells is crucial for achieving tissue regeneration and functional recovery. It is recognized that factors influencing cell growth are not solely chemical but also physical cues, such as the microstructure of the cell substrate. Therefore, in addition to the cytotoxicity of the skins, the effect of the skins on cell proliferation was assessed by directly culturing the cells in the skins as well. As shown in Figure 5C, during the 14-day cell culture, the number of HSF and HaCaT cells represented by the quantity of DNA in various skin types continued to increase, with a greater number of cells present in decellularized skin than in ND skin, which may be attributed to the more porous microstructure of the skin resulting from decellularization. Furthermore, the highest cell count was observed in ED-C (∼1.6-fold higher than ND-C and ∼1.3-fold higher than CD-C for HSF; ∼5-fold higher than ND-C and ∼2-fold higher than CD-C for HaCaT), likely due to the absence of harmful residues and the presence of a porous microstructure in ED-C.

Hemocompatibility refers to the tolerance of materials to blood, which is one aspect of biocompatibility and indicates the interactions of biomaterials with blood. As shown in Figure 5D, the material-induced hemolysis rates of the skin groups (ND-C, CD-C and ED-C) were <2% (1.11 ± 0.06%, 1.09 ± 0.03% and 0.35 ± 0.26%, respectively), indicating non-hemolytic (ASTM-F756-17). However, the hemolysis rate of the skin group (CD-C) was the highest, with values similar to those reported previously [15]. This may be due to the different decellularization protocols used. Therefore, skin derived from electrical decellularization has better hemocompatibility than skin derived from chemical/enzymatic decellularization, which is conducive to its clinical applications.

The MTT method can quantitatively assess the proliferation of lymphocytes, reflecting the impact of immune antigens present in the skin on lymphocytes. As shown in Supplementary Figure S3, lymphocytes cultured in CD-C and ED-C extracts proliferated significantly slower than those cultured in media containing ConA (positive control (PC)). However, lymphocytes cultured in ND-C extracts exhibited significantly faster proliferation than those cultured in RPMI1640 medium containing 10% FBS (NC), and their proliferation rate was comparable to that of lymphocytes in media containing ConA (PC), indicating lymphocytes cultured in decellularized skin extracts remained in an inactivated state, which might be due to lack of immunogenic substances that stimulate significant lymphocyte proliferation in decellularized skin [50].

Biocompatibility in vivo

At 1, 2, 3 and 4 weeks post-subcutaneous implantation, the in vivo biocompatibility and cellular infiltration of decellularized and ND fish skins were examined using H&E staining of tissues harvested from the implantation sites. Representative H&E-stained images of the implant sections are shown in Figure 6A. On day 7, the integrity of the fish skin structures beneath the dermal muscle layer was well maintained across all groups (ND-C, CD-C and ED-C). The implanted ND (ND-C) structures were the most compact, whereas some pores were observed in both the decellularized skins (CD-C and ED-C) structures, as indicated by arrows; the porosity in CD-C was greater than that in ED-C. In all implanted fish skin samples, intracellular components, including cytoplasmic proteins and ECM, stained pink. Notably, no cell nuclei (deep blue) were observed in decellularized skins. A fibrous layer including both immune cells and fibroblasts surrounded the implants in all groups, indicating symptoms of an immune response, consistent with previous reports on subcutaneous implantation of polycaprolactone, poly (lactic-co-glycolic acid), and fish collagen composites [33]. However, the density of surrounding cells in the ED-C was the lowest, which may be related to a lower lymphocytic response induced by the ED-C (Supplementary Figure S3). The immune response to the implanted material plays a critical role in successful tissue regeneration; a moderate immune reaction promotes tissue repair and integration, whereas excessive or chronic inflammation hinders regeneration [51]. Modulation of the immune response is critical in the modern paradigm of tissue regeneration, attenuating fibrous encapsulation and implant isolation. On day 14, cellular infiltration was observed in the ND-C group, but not in the CD-C and ED-C groups. Furthermore, the porosity of the CD-C and ED-C structures increased, making their architectures appear looser compared to day 7. On day 21, cellular infiltration was observed in both ND-C and ED-C groups, but not in the CD-C group. By day 28, cellular infiltration was evident in all groups. However, the boundary between the implant and host tissue remained clearly distinguishable in the CD-C group, while it was less distinct in both ND-C and ED-C groups. Moreover, the number of cells migrating into the CD-C implant was significantly lower than that migrating into ND-C and ED-C, likely related to the relatively higher cytotoxicity of CD-C (Figure 5A). The systemic toxicity of fish skin in vivo was evaluated by harvesting major organs from mice after 2 weeks of subcutaneous implantation for H&E staining. As illustrated in Figure 6B, the organ staining results for mice implanted with fish skins were comparable to those of the NC. The structural integrity of the heart, liver, spleen and lung tissues remained intact, with clear glomeruli and renal tubules, uniform cytoplasm and no evidence of cell necrosis, fluid accumulation or inflammatory cell responses. The results indicated that fish skin, with or without decellularization treatment, did not cause systemic toxicity in mice.

Biocompatibility of skin in vivo. (A) Representative H&E staining images of harvested subcutaneous mouse tissue surrounding implanted fish skins with different treatments after 7, 14, 21 and 28 days (n = 6); (B) H&E staining images of major organs of mice receiving various treatments (n = 6). NC, sham.

The 28-day subcutaneous implantation model established in this study offered important preliminary data on the acute biocompatibility and initial host response to the decellularized tissue; it is recognized that it does not fully capture the potential for long-term remodeling and functional integration. The mild immune response and gradual cellular infiltration observed in the ED-C group are promising indicators of biocompatibility. However, to thoroughly assess the clinical potential of ED fish skin, future studies should extend observation periods (e.g. 6 weeks or more) [6], utilize functional models (such as full-thickness skin wound healing) [15], and conduct quantitative analyses of key regenerative processes. This will involve detailed immunohistochemical characterization of immune cell populations (e.g. analysis of M1/M2 macrophage polarization using markers such as CD86/CD206), assessment of vascularization (e.g. CD31 staining to label endothelial cells) and evaluation of nerve ingrowth [52]. Such studies will provide deeper insights into the role of decellularized tissue in fostering a pro-regenerative microenvironment and its functional outcomes in tissue repair. The future direction is not just to provide mechanical support but to deliver a regenerative signal. Enhance the efficacy of medical devices such as bone implants, wound dressings and bioactive sutures by leveraging the inherent bioactivity of the ECM. In addition, analyses of sensitive DNA fragments, proteomics/growth factor profiling and immunogenicity testing will further enhance the understanding of the biocompatibility of decellularized tissues and contribute to the broad application of these tissues in tissue engineering and regenerative medicine.

Conclusion

To fulfill the demand for sustainable large-scale biocompatible decellularized materials for biomedical applications, we developed a novel electrical decellularization approach to efficiently prepare decellularized fish skins devoid of harmful chemical residues successfully. Biocompatibility assessments including cytotoxicity, proliferation, hemolysis, and in vivo experiments further confirmed that the toxicity and hemolytic activity of ED fish skin were significantly lower than those of chemically/enzymatically decellularized fish skin, and it supported cell growth and in vivo tissue integration, meeting the basic requirements of biomaterials for tissue engineering and regenerative medicine. However, this study has certain limitations. First, this study primarily focused on in vitro characterization. Despite short in vivo implantation experiments demonstrating good biocompatibility and low immunogenicity, there is a lack of systematic evaluation of the long-term fate of decellularized skin post-implantation, including its degradation rate and integration into host tissue. Such long-term results are crucial for assessing the potential for clinical translation. Second, the scope of this study was limited to a single tissue type, fish skin. Different organs and tissues, such as the heart, liver, and blood vessels, exhibit significant differences in cell density, lipid content, and matrix density. Whether electric field technology can achieve comparable decellularization efficacy, efficiency and matrix preservation across these diverse tissues remains to be investigated.

Supplementary Material

rbag005_Supplementary_Data