3D Skeletal Scaffolds of Marine Keratosan Demosponges Origin as Renewable Sources for Bioinspiration in Modern Structural Biomimetics and Tissue Engineering

Abstract

This experimental review discusses evolutionarily approved, naturally pre-designed skeletal architectures of marine keratosan sponges in the form of 3D scaffolds, which have garnered increasing interest in the fields of structural and functional biomimetics as well as in tissue engineering. It has been demonstrated that these renewable, ready-to-use natural scaffolds can undergo further modifications through specialized treatments such as metallization and carbonization, enabling the creation of functional biomaterials while maintaining the species-specific hierarchical 3D structure. The study presented remarkable findings, including the demonstration of the unique shape-memory behavior of these scaffolds even after two months of exposure to high mechanical pressure at temperatures exceeding 100 °C. Additionally, the cytocompatibility and biological performance of natural and carbonized (1200 °C) spongin scaffolds, derived from selected bath sponges, were comparatively investigated with respect to growth and proliferation of human MG-63 osteoblastic cells. Understanding whether carbonization universally enhances osteogenic capabilities or selectively amplifies the inherent architectural advantages remains to be critical for the rational design of sponge-derived scaffolds in bone and structural tissue engineering applications.

1. Introduction

According to the modern view, scaffolding strategies include two key methods: the fabrication of requested 3D constructs from corresponding natural and/or synthetic precursors using technological tools as “bottom-up” approach. Another is the use of natural decellularized biomaterials as prefabricated scaffolds originating from cultivated plant and animal sources, which is a “top-down” approach [1].

Combining biomimetic ideas with the principles of porous 3D structural design continues to be an effective approach for creating a wide variety of sophisticated artificial [2], hybrid [3,4], and biopolymer-based [5] scaffolds. Developing greener alternatives for all materials that could be potential sources of microplastic contamination in the environment is increasingly important in modern society [5]. As a result, the use of various microplastic-free raw materials derived from industrial [6] and biological waste [7,8] has gained attention among experts in bioinspired materials science. Currently, the fundamentals of scaffolding [9,10] and recent advances in sustainable scaffolding methods [11], including additive manufacturing (AM) techniques such as stereolithography (SLA), digital light processing (DLP), selective laser sintering (SLS), and fused filament fabrication (FFF) [12], remain highly relevant. Additionally, developing 3D printing methods for porous scaffolds with specific architectures continues to be a popular focus, especially in tissue engineering [13]. The recent progress in sustainable scaffolds for regenerative medicine is effectively documented in the book titled “Sustainable Scaffolds-based Strategies in Tissue Engineering and Regenerative Medicine” (see [14]).

Thus, a classic example of cellular bioarchitecture is the physical form called “a sponge,” which is derived from living nature. Since ancient times, humans have known about plant sponges (such as Luffa plants) and marine organisms called sponges (phylum Porifera). Currently, the cellulose-based Luffa plant sponge continues to attract researchers as a natural cellular material that is ready to use [15] in fields like cushioning and energy absorption [16], mechanical design [17], tissue engineering [18], and as inspiration for developing similar structures for biomedicine [19,20,21,22].

Unlike the spongy skeletons of plant origin, their morphological counterpart in marine sponges only shares an external resemblance and differs fundamentally in chemical composition, shape diversity, pores, and sizes. As the first multicellular organisms on Earth [23], sponges have evolved over more than 800 million years [24] and have produced more than a hundred morphotypes of their three-dimensional, porous skeletons composed of either biominerals (biosilica, calcium carbonates) or biopolymers (spongin, collagen, chitin) (see overview [25,26,27,28]). The functional morphology of sponges [29], characterized by being mechanically robust and porous as “living pumps” and “living filters” [30], is strictly determined by the hydrodynamics laws of their aquatic environment, shaped by a sedentary lifestyle. Both chitin- and spongin-based skeletons provide structural stability to the sponge body. It is reasonable to believe that marine sponges, torn from their natural substrate during storms, were washed ashore, weathered naturally, and discovered as three-dimensional skeletons by ancient humans, who began exploring their uses. Primarily, this refers to the so-called bath, horny, or commercial sponges [25,31], belonging to the Spongididae family (Order Dictyoceratida, Subclass Keratosa, Class Demospongiae), which have biomineral-free, proteinaceous spongin skeletons (see Figure 1). The variety of shapes, sizes, porosity, and hierarchical structural ornamentation of their interconnected micro-fiber three-dimensional networks remains a focus of modern biomimetics [32,33,34], bioinspired materials science [35], and biomedicine [36,37,38]. A similar scenario developed around chitin-based three-dimensional scaffolds derived from other keratosan sponges of the Verongiidae family (Subclass Keratosa, Class Demospongiae) (see overview by [39,40]), which are widely used in extreme biomimetics [41], biomedicine, and tissue engineering (see [42,43,44]).

Figure 1.

Synthetic sponges, in all their chemical diversity, are a product of engineering thought, inspired by the fundamental structural motif of the evolutionarily proven porous bioarchitecture of the skeletons of marine bath sponges.

Considering that the ultimate goal of using 3D scaffolds in biomedicine is the real possibility of their application in the clinic, it is worth noting the undeniable advantages of scaffolds derived from marine sponges. These poriferan scaffolds were actively and effectively used both in ancient medicine [45] and in clinical studies, especially in the 19th century, with detailed descriptions available in scientific literature. Briefly, protein-based 3D scaffolds made from commercial marine sponges have been used, for example, as implants [46], sponge-tents [47,48,49], sponge-grafts [50,51,52,53,54,55,56,57], dressings, and hemostatic materials in treating both civilians and military victims [58]. Naturally, at that time, there were no legal restrictions on using commercial sponges for medical purposes. Furthermore, the scientific insights from these unique works seem invaluable today.

These historical medical applications implicitly demonstrate that bath sponge-derived spongin scaffolds can interact favorably with human tissues; however, their biological performance was never evaluated using modern cell-based biological assays. Despite the extensive biomimetic and structural relevance of marine bath sponge-derived scaffolds, surprisingly little is known about how species-specific skeletal architectures translate into functional osteoblast responses under biologically relevant conditions. In particular, systematic comparisons addressing osteoblast viability, migration, 3D infiltration, and metabolic activity on spongin scaffolds of different natural origin—and how these responses are modulated by high-temperature carbonization—remain scarce. Given that carbonization profoundly alters surface chemistry and mechanical properties while preserving native 3D morphology, understanding whether it universally enhances biological performance or selectively amplifies intrinsic architectural advantages is essential for tissue engineering applications.

The third example of natural scaffolds includes the so-called decellularized matrices, such as decellularized extracellular matrix scaffolds for soft tissue augmentation [59,60], decellularized human and non-human tissues [61], as well as arteries or veins [62]. However, significant challenges remain in the clinical implementation of these bioscaffolds [63].

From an economic viewpoint, the two main scaffolding strategies described above differ significantly in terms of overall costs. It is one thing to extract and purify natural renewable biomaterials into ready-to-use 3D matrices, but it is quite another to develop cost-intensive technologies for creating advanced functional 3D scaffolds using modern computer-assisted tools. The practicality of using natural scaffolds or those created through technology ultimately depends on their intended application. Therefore, if the density and porosity of the material used for the scaffold can be controlled in artificially created matrices (Figure 2), then in the case of similar structures extracted from different types of collected or cultivated sponges, appropriate screening and subsequent selection will be necessary.

Figure 2.

Comparative digital microscopy images of a 3D spongin scaffold isolated from Spongia lamella keratosan demosponge (A) and three selected and differently coloured (see Figure 1) synthetic sponges (B–D).

It is hoped that marine sponge economics [64] will continue to develop as an independent and highly important discipline. Only then will we be able to obtain key information about the economic benefits of the main directions in modern scaffolding strategies, as briefly outlined above. Our review paper titled “Marine Spongin: Naturally Prefabricated 3D Scaffold-Based Biomaterial” published in 2018 [31] already analyzed comprehensive aspects of spongin, including its history, mechanical and physicochemical properties, the commercial sponge fishery, diversity of marine sponge ranching strategies, and multifaceted applications of this unique natural cellular material.

Importantly, high-temperature carbonization of biological scaffolds is often implicitly assumed to enhance their biological performance by increasing stiffness, surface roughness, or electrical conductivity. However, such assumptions are rarely verified in a comparative, species-resolved biological context. In the case of naturally evolved sponge skeletons, carbonization may preserve macro- and micro-architecture while simultaneously amplifying intrinsic architectural differences, potentially leading to divergent cellular responses.

This work represents the first experimental review combining structural, physicochemical, and biological characterization of selected renewable spongin- and chitin-based 3D scaffolds derived from marine keratosan sponges. By integrating advanced materials modification strategies with systematic biological evaluation, this study highlights how species-specific skeletal architecture governs osteoblast behavior in both natural and carbonized scaffolds, providing design-relevant insights for future biomimetic and tissue engineering applications.

2. Materials and Methods

2.1. Materials

In the study, we used spongin scaffolds isolated from various keratosan demosponges such as Hippospongia communis Lamark, 1814, H. communis covered with lepidocrocite, Spongia tampa de Laubenfels & Storr, 1958, Spongia lamella Schulze, 1879, Spongia tubilifera Lamark, 1814, as well as from the marine verongiid demosponge Ianthella basta Pallas, 1766, which were purchased from INTIB GmbH (Freiberg, Germany). Hydrochloric acid (35–38%), acetic acid, and sodium hydroxide used for treating and purifying the demosponge skeletons were purchased from Stanlab (Lublin, Poland), while hydrogen fluoride (40%) was obtained from Chempur, Piekary Śląskie, Poland.

2.2. Staining Test with a Protein-Specific Dye (Roti^®Blue/Coomassie Blue)

To visualize the distribution of protein components within the spongin framework, small fragments of H. communis spongin skeletons were immersed in an aqueous solution of Roti^®Blue (Coomassie Brilliant Blue–based stain; Carl Roth GmbH, Karlsruhe, Germany) and incubated overnight at room temperature without agitation. After staining, the samples were gently rinsed with deionized water to remove excess dye and examined in the wet state under a high-resolution digital optical microscope (VHX-7000, Keyence International, Osaka, Japan) [26]. The Coomassie-based dye selectively stained the protein matrix, allowing for a clear distinction between the spongin fibres and the mineral inclusions embedded within them.

2.3. Protein Matrix Dissolving

To partially dissolve the protein matrix before MXRF measurements, spongin samples were exposed to a 1% NaOH solution (Sigma Aldrich, GBMH, Hamburg, Germany) for 24 h at 40 °C, then washed three times with demineralized water.

2.4. Liquid Absorption Capacity of Spongin

The liquid absorption capacity test was conducted on spongin scaffolds from demosponge S. lamella. Fifteen different liquids were used in this experiment: (1) tap water, (2) distilled water, (3) simulated seawater obtained by dissolving a commercial salt mixture, Instant Ocean, in distilled water (Aquarium Systems, Blacksburg, VA,USA), (4) alcohol-free and colorant-free dental water Elmex (Colgate-Palmolive Manufacturing, Świdnica, Poland), (5) syrup—50% sugar solution in hot water (Diamant, Miejska Górka, Poland), (6) pork blood from a butcher’s shop—ingredients: pork blood, vinegar 10%, sodium citrate (P.H.U. AGA Paweł Geisler, Oborniki, Poland), (7) physiological saline—0.9% NaCl solution (Eurochem, Tarnów, Poland) in distilled water, (8) human plasma—Defibrinated, delipidated human plasma (Seqens, Limoges, France), (9) serum—fetal bovine serum (FBS) (Sigma-Aldrich, St. Louis, MO USA), (10) cell culture medium—Dulbecco’s Modified Eagle’s Medium (DMEM) with high glucose (Sigma-Aldrich, St. Louis, MO, USA), (11) 10% spirit vinegar (ZPS JAMAR Szczepaniak, Albertów, Poland), (12) red wine (Rotglut, Mainz-Hechtsheim, Germany), (13) beer—Łomża Pils (Van Pur S.A, Łomża, Poland), (14) glycerin-analytical grade (Eurochem, Tarnów, Poland), and (15) hyaluronic acid—a 1% solution of hyaluronic acid with an average Mw of 000 (Sigma-Aldrich, St. Louis, MO, USA) in distilled water.

Before the experiment, the spongin scaffolds from S. lamella were cut into cubes with sides of about 15 mm, washed in distilled water in an ultrasound bath (Bandelin electronic GmbH & Co. KG, Berlin, Germany) for 3 h to remove solid precipitate, and then dried. These prepared spongin skeleton samples were weighed, immersed in the subsequent solutions, and left for 15 min to absorb the liquid. Afterwards, the samples were carefully removed from the solution with tweezers without pressing their skeletons, and weighed again. For solutions of the highest viscosity—(6) pork blood and (14) glycerin—the weighing was performed after the excess solution on the surface of the sample had finished dripping. For other solutions, no drops were observed after removing the sample. Measurements for each solution were repeated 3 times. The experiments were conducted at room temperature. The density of each solution under the experimental conditions was determined by weighing 1 mL of the solution, measured using a precise automatic pipette (Transferpette S, 100–1000 μL, Brand GMBH, Wertheim, Germany). Sample weighing was performed with the laboratory balance PR Series: PR202, d = 0.01 g (Ohaus, Nänikon, Switzerland).

2.5. Surface Wettability of Spongin Microfibres and Capillary Effects

The molding silicone XIAMETER RTV-4234-T4 (POLYCORE, Warszawa, Poland) and polyurethane (PUR) (SikaBiresin^® UR5805/UR5895; Sika Deutschland GmbH, Stuttgart, Germany) were used to coat the spongins under study. Selected spongin samples were immersed in silicone or PUR solutions at room temperature to enable the coating of the biomaterial due to penetration into the fibrous skeletal network. Excess liquid was subsequently drained off. The prepared specimens were left to fully cure under laboratory conditions before being subjected to further analyses of surface properties.

2.6. Creation of Spongin–Titanium 3D Composites Using Ion–Plasma (Vacuum Arc) Deposition Method (VAD)

A schematic representation of the spongin metallization process is shown in Figure 3. The vacuum chamber of the ion–plasma deposition setup (pos. I) is equipped with a special rotating disc-type device inclined at a 45° angle. Spongin samples were secured in three sections of this device (pos. II), periodically exposed to the metallic plasma flow. Two types of spongin samples were used: one with a less dense structure (pos. III, type 1) and another with a denser structure (pos. VI, type 2). The type 2 samples measured 30 × 30 mm with thicknesses of 10 and 20 mm, while the type 1 samples had a rounded shape with approximately 60 mm diameter and 30 mm thickness.

Figure 3.

Schematic representation of the spongin metallization process.

The spongin metallization process was conducted in an argon atmosphere following the conditions outlined in Table 1.

Table 1.

Modes of Spongin Metallization.

Working Pressure, PAr, Pa	Arc Current, A	Arc Voltage, V	Bias Voltage, V	Deposition Time, min
2.66 × 10−1	90…100	60…70	50…80	5…10

2.7. Chromium Tanning of Spongin Scaffolds

Spongin scaffolds of H. communis containing lepidocrocite were first treated with 3 M HCl for 24 h at room temperature. After removing the acid by rinsing with distilled water, the material was dried at 40 °C (using a Binder FD-S 056) and weighed. The cleaned samples were then transferred into a 50% solution of basic chromium sulfate (Cr(OH)SO₄) (Chemat, Gdańsk, Poland). They were kept at 40 °C for 48 h. The volume of the chromium solution was adjusted to the sample’s mass, using 15 mL per 100 mg. After the treatment, excess liquid was removed on 615 filter paper for 2 min, and the samples were again oven-dried at 40 °C and weighed [65,66,67]. To remove non-bound chromium compounds, the scaffolds underwent ultrasonic treatment in a Bandelin Sonorex DT 52H bath for 4.5 h, with the temperature kept below 40 °C. After subsequent drying and weighing, the samples were placed in graphite crucibles (68 mm diameter) and carbonized in a Czylok PT-1/220/GR furnace at 1450 °C for 90 min. The thermal program involved heating at 20 °C per minute to 950 °C, then at 10 °C per minute to the final temperature, under argon flow (180 L/min during heating, 120 L/min during carbonization, and 60 L/min during cooling; with one chamber purge before the process). After carbonization, the material was reweighed and subjected to an additional 30-min ultrasonic bath to assess the stability of the resulting structures.

2.8. Carbonization of 3D Spongin Scaffolds

Digital microscopy image of the carbonized H. communis spongin fragment at 1200 °C, selected for pore size and pore diameter calculations. These calculations were performed using the original software of the VHX 6000 digital microscopy system (Keyence, Osaka, Japan). First, the selected spongin scaffolds were washed with running water and treated with 3M HCl solution for 24 h to dissolve residual calcium carbonates. The carbonization of selected fragments of purified spongin scaffolds has been carried out during 1–1.5 h using a special high-temperature furnace Czylok PT-1/220/GR (Jastrzębie - Zdrój, Poland) under oxygen-free conditions. After the system was pumped and purged with argon once, the temperature was ramped at 20 °C/min up to 950 °C, 10 °C/min up to 1950 °C, and 5 °C/min up to 2200 °C with argon fed at 100 standard cubic centimeters per minute at ambient pressure [32]. A schematic of the procedure is shown in Figure 4.

Figure 4.

Schematic representation of sample preparation and their carbonization on example of spongin scaffolds derived from H. communis demosponge.

2.9. 3D Spongin-Based Scaffolds and the Shape-Memory Phenomenon

The spongin isolated from S. tubilifera demosponge was placed between two steel plates (400 × 400 × 4 mm). A layer of silicone paper was additionally inserted between the spongin scaffold and the plates. The pressing was performed using a hydraulic press (type PH40T, ReMi-Plast, Czerwonak, Poland) equipped with a heating/cooling table and a maximum pressing force of 400 kN. The temperature of both the upper and lower heating plates was set to 100 °C, with a pressing time of 30 s and a force of 200 kN. The mold height was 6 mm, while the spongin was compressed to 2 mm during pressing. Spongin was stored at room temperature for 2 months before being immersed in water. The molding silicone XIAMETER RTV-4234-T4 was purchased from POLYCORE, Poland.

2.10. Preparation of Ag-Spongin Scaffolds

All samples of spongin scaffolds from S. lamella, before Ag-spongin preparation, were treated with 3M HCl for 3 h using an ultrasonic bath (UST) (Bandelin Sonorex DT 52H, Berlin, Germany), then with 40% HF for 2 h without UST treatment to achieve complete demineralization of the samples.

2.10.1. Ag-Spongin Preparation—Method I

A sample of purified S. lamella scaffold (587.2 mg) was used in this experiment. 17 g of AgNO₃ (Stanlab, Lublin, Poland) was dissolved in 1000 mL of distilled water to prepare a 0.1 M AgNO₃ solution. 16 g of NaOH was dissolved in 500 mL of distilled water to prepare 0.8 M NaOH. To make the reduction agent, 92.05 g of fructose (F0127, Sigma-Aldrich) and 35 g of citric acid (C0759, Sigma-Aldrich) were dissolved in distilled water to make 450 mL of solution (1.03 M fructose + 0.40 M citric acid). The spongin scaffold sample was immersed in 68.2 mL of the prepared AgNO₃ solution (the volume was adjusted based on the sample mass: 30 mL of solution per 258.2 mg of sample) for 1 h in a glass bottle. Next, 34.1 mL of 0.8 M NaOH was added (calculation: 15 mL per 258.2 mg of sample). Then, 25% NH₄OH was added until the precipitate dissolved. Following this, 2.27 mL of 96% ethanol (Stanlab, Lublin, Poland) was added (calculation: 1 mL per 258.2 mg of sample). Then, 22.7 mL of reduction agent was added (calculation: 10 mL per 258.2 mg of sample). During reagent addition, the solution was stirred with a rod. After completing the additions, the sample was left at room temperature in a dark place for 24 h [68]. Then, it was dried in an oven at 50 °C for 24 h. The dried sample was then placed in an ultrasonic bath for 2.5 h (maximum distilled water temperature in the bath was 60 °C). After that, the sample was dried again in an oven at 50 °C for 24 h.

The Ag-spongin sample was pressed using a laboratory press (ReMi-Plast s.c., PH40T) at 40 tons for 30 s, resulting in a 2 mm thick sample (mold height: 6 mm; pressure force: 150 kN; temperature of upper and lower heating plates: 120 °C). Next, 15% of the pressed sample was immersed in distilled water for 19 h. Finally, the sample was dried in an oven at 40 °C for 24 h.

2.10.2. Ag-Spongin Preparation—Method II

To prepare a 1% solution of AgNO₃, 11 g of AgNO₃ (Stanlab, Lublin, Poland) was dissolved in 1100 mL of distilled water. Next, the pH was raised to 11.2 using 12.4 mL of 25% NH₄OH (Chempur, Piekary Śląskie, Poland)—the base was added until the precipitate dissolved. Then, a sample of S. lamella (850.1 mg) was immersed in the prepared solution (51.1 mL), with the solution volume adjusted based on the sample mass: 100 mL of AgNO₃ solution per 1664 mg of the sample. After 10 min, the closed glass bottle containing the spongin sample was placed in an oven (Binder FD-S 056) at 40 °C for 24 h [69]. The sample was then dried in an oven at 40 °C for another 24 h. To achieve optimal results, the process was repeated three times.

The resulting Ag-spongin sample was pressed using a 10-ton tablet press (13 mm diameter, 150 mbar, for 5 min). A 25% fragment of the spongin sample with silver and 25% control spongin sample were prepared. Next, 25% of the pressed samples were immersed in distilled water for 19 h, then dried in an oven at 40 °C for 24 h.

2.11. Gentle Method for Isolation of 3D Flat Chitinous Scaffolds from Ianthella basta Demosponge

The dried 40 × 50 cm skeleton of I. basta marine verongiid demosponge was immersed in distilled water for 30 min at room temperature (RT) to soften it and remove salts. After that, it was cut into 4 cm × 4 cm pieces using scissors. These skeletal fragments were placed in a 2L bottle with distilled water and incubated at 50 °C in an oven (Solid.Line FD-S 56, BINDER GmbH, Tuttlingen, Germany) for 7 days. Then, the skeletal fragments were separated from the aqueous extract using a paper filter. Finally, the brownish-colored aqueous extract was dried under a fume hood at room temperature for 15 days. Corresponding bromotyrosines were isolated from the resulting powder-like residue by column chromatography on silica gel using CHCl₃-MeOH as the eluent and identified via spectroscopic and spectrometric methods (as in [70]).

Both the α-chitin standard derived from snow crab (Chionoecetes opilio) and the β-chitin standard isolated from sepia (Sepia officinalis) were purchased from INTIB GmbH (Freiberg, Germany).

The isolated skeletal fragments of I. basta demosponge were immersed in 20% acetic acid for 24 h at RT, then washed with distilled water until the pH reached 6.5. Subsequently, the samples were transferred to a 10% NaOH solution for 24 h at RT. This alternating acid-base treatment was repeated over five days to obtain colorless chitinous scaffolds. The schematic view of the procedure is presented in Figure 5.

Figure 5.

Schematic representation of the procedure of isolating chitinous scaffolds from I. basta demosponge skeleton together with aqueous bromotyrosines-containing extract.

2.12. Natural and Graphitized 3D Spongin Scaffolds for Tissue Engineering with Human MG-63 Osteoblastic Cells

2.12.1. Preparation of Natural and Carbonized Spongin Scaffolds for Tissue Engineering Experiments

3D spongin scaffolds were isolated from marine demosponges H. communis, S. tampa, and S. lamella using established purification protocols that ensure the preservation of the native fibrous architecture. For carbonized variants, native scaffolds were subjected to thermal treatment at 1200 °C under an inert atmosphere, yielding carbonized replicas of the original structures [32]. After processing, all scaffolds were cut, sterilized, and placed into sterile 48-well cell culture plates prior to biological testing.

2.12.2. Cell Culture and Seeding

MG-63 osteoblast-like cells were selected as a representative model for evaluating osteoblast–scaffold interactions on porous biomaterials. MG-63 cells were cultured under standard conditions (37 °C, 5% CO₂) in complete Dulbecco’s Modified Eagle Medium (DMEM, Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 10% fetal bovine serum (FBS). The medium was supplemented with penicillin (100 U mL⁻¹) and streptomycin (100 mg mL⁻¹). Sponges were cut to fit the 48-well plate, and cells were seeded onto the top of the spongin scaffolds at a density of 30,000 cells per well. Standard cell culture polystyrene (TCPS) surfaces served as reference substrates for monitoring indirect cytotoxic effects. MG-63 cells were selected due to their well-documented sensitivity to substrate stiffness, surface topography, and 3D scaffold architecture, making them suitable for comparative evaluation of species-dependent scaffold effects.

2.12.3. Statistical Analysis

All data are presented as mean ± standard deviation (SD). Statistical significance was assessed using GraphPad ™ software (version 8.x; GraphPad Software, San Diego, USA) by the unpaired Student’s t-test for the comparison of two datasets with normal distribution. Data with p < 0.05 were considered statistically significant.

2.13. Characterization Techniques

2.13.1. Digital Optical Microscopy

Samples were observed and analyzed using an advanced imaging system, the VHX-6000 digital optical microscope (Keyence, Osaka, Japan), equipped with a VH-Z20R zoom lens (magnification up to 200×), as well as the VHX-7000 digital optical microscope (Keyence, Japan), which features VHX-E20 (magnification up to 100×), VHX-E100 (magnification up to 500×), and VH-Z100T (magnification up to 1000×) zoom lenses.

2.13.2. Scanning Electron Microscopy (SEM)

Morphological analysis was conducted using a Quanta 250 FEG scanning electron microscope (FEI Ltd., Brno, Czech Republic), producing high-resolution images of the sample surfaces with established parameters: for spongin-chromium and control samples before carbonization-detector LFD, 10 kV at 70 Pa (low vacuum); and for the spongin-chromium sample after carbonization-detector ETD, 10 kV at 3.64 × 10⁻³ Pa or 4.76 × 10⁻³ Pa (high vacuum).

2.13.3. X-Ray Diffraction (XRD)

The X-ray diffraction data was collected using a Bruker D8 Advance diffractometer equipped with a Cu X-ray tube. The setup included a Goebel mirror that provided a parallel beam to illuminate the sponge sample placed on a glass plate to avoid sample holder reflections. The diffraction pattern was collected using a LynxEye silicon strip detector operating in 1D mode within the 2θ range from 5° to 120°, with a resolution of 0.04°.

2.13.4. Micro X-Ray Fluorescence (MXRF)

MXRF measurements were carried out using a modified M4 Tornado MXRF spectrometer (Bruker Nano GmbH, Berlin, Germany) equipped with a 30 W Rh-microfocus X-ray tube (50 kV, 600 μA), a polycapillary full lens in the excitation channel for X-ray focusing (spot size ≤ 20 μm for Mo Kα), and a 30 mm² silicon drift detector (SDD) [35]. The samples were fixed on a polymer slide with double-sided carbon tape, and element distribution images were received by lateral movement of the samples laterally with an xyz motorized stage. MXRF measurements were performed for a total sample area of 13.0 × 8.0 mm² (or 18.0 × 8.0 mm²) with a step size of 5 μm. A spot measure time of 10 ms was utilized, resulting in an overall measurement time of approx. 12 h. Additionally, regarding the presence of light elements in the spongin samples, a vacuum of 20 mbar was applied during measurements. Data analysis of the generated xy area maps was performed using the spectrometer software, which provided impulse count values and element distribution images for signals of Si-Kα (1.740 keV), S-Kα (2.309 keV), K-Kα (3.314 keV), Ca-Kα (3.692 keV), Fe-Kα (6.405 keV), Br-Kα (11.924 keV), and I-Lβ (4.221 keV).

2.13.5. Water Contact Angle (WCA)

Water contact angle (WCA) analyses were performed by the sessile drop technique at room temperature and atmospheric pressure with a Krüss DSA100 (Goniometer, Hamburg, Germany). Square samples of approximately 1 cm × 1 cm were prepared. Three independent measurements were taken for each sample using 5 µL water drops, and the obtained results were averaged.

2.13.6. Optical Microscopy

Cell attachment and morphology were evaluated using optical microscopy after 1 and 7 days of culture. This analysis provided a qualitative assessment of indirect cytotoxicity, based on the cells’ ability to adhere to the TCPS surface in the presence of the scaffolds.

2.13.7. Live/Dead Assay

Cell viability within the 3D scaffolds was evaluated after 3 and 7 days using fluorescein diacetate (FDA) and propidium iodide (PI) (F7378, P4170, Sigma-Aldrich) according to the manufacturer’s protocol and adopted from previously described 3D cell culture and imaging workflows [71]. 3D scaffolds were incubated with the staining solution at room temperature for 5 min in the dark. After removal of the staining solution and washing with PBS, the samples were maintained in medium without serum for analysis. Confocal laser scanning microscopy was used to visualize viable (green) and non-viable (red) cells within the scaffold volume. Autofluorescence from spongin fibres was taken into account during image interpretation. Image 3D visualization was performed using Imaris software (version 11.0, Oxford Instruments, Abingdon, Oxfordshire, United Kingdom).

2.13.8. Immunocytochemical Staining

For carbonized spongin scaffolds, immunocytochemical staining was performed after 7 days of culture. Cell nuclei were labeled with DAPI, the cytoskeleton with phalloidin (F-actin), and proliferating cells with Ki67. Sponges were fixed in 4% paraformaldehyde overnight at 4 °C. The next day, samples were washed three times for 30 min each with 1X PBS on a rotary shaker. Sections were incubated for 4 h in blocking buffer (1X PBS, 0.5% Triton X-100, 1% BSA). For F-actin cytoskeleton visualization, Oregon Green 488 Phalloidin (O7466, Thermo Fisher Scientific, Waltham, MA, USA) was used. Primary antibodies were applied to detect antigens: human anti-Ki67 conjugated with Phycoerythrin (1:100, 2352515, Sony, Tokyo, Japan). After PBS washing, the samples were incubated with DAPI (4′,6-diamidino-2-phenylindole) (Thermo Fisher Scientific, Waltham, MA, USA) solution for 1 h at RT (21 °C). Samples were imaged using an Olympus FV1200 confocal microscope with a 10× dry objective. All measurements were repeated at least three times. Image 3D visualization was performed using Imaris software (version 11.0, Oxford Instruments).

2.13.9. Cell Viability and Metabolic Activity Assessment

Cell metabolic activity was measured using the Alamar Blue assay following the manufacturer’s protocol (Cat.786–921, G-Biosciences, St. Louis, MO, USA) and interpreted as metabolic activity in 3D scaffolds, as previously described [72,73,74]. MG-63 cells were cultured in the presence of natural and carbonized spongin scaffolds derived from S. lamella and H. communis. Tissue culture polystyrene served as the control. Prior to the Almar Blue assay, the DMEM culture medium was removed, and wells containing control cells and cells seeded on spongin samples were rinsed twice with PBS. Alamar Blue solution in phenol red-free DMEM at (1:9 v/v) ratio was added to each well, followed by incubation at 37 °C for 2 h. During incubation, resazurin was reduced to resorufin by metabolically active cells. Plates were checked in fluorescence reader with excitation wavelength at 530–560 nm and emission wavelength at 590 nm. Measurements were performed after 3 and 7 days of culture. Fluorescence intensity was expressed as relative metabolic activity. For each time point, values were normalized to the tissue culture polystyrene (TCPS) control (set to 100%). In this study, the Alamar Blue assay was used as an indicator of cellular metabolic activity and adaptation within in 3D scaffolds and was interpreted in conjugation with Live/Dead staining and imaging, rather than as a standalone ISO 10993 [75] cytotoxicity test. Similar approaches for assessing cell metabolic activity in 3D porous scaffolds have been widely applied in previous studies using different biomaterial systems [72,73,74].

3. Results and Discussion

3.1. On Structural Architectonics of Spongin-Based Skeletons of Bath Sponges

For people and scientists not directly involved in spongology (the study of sponges as living organisms), the mention of “marine, or sea sponge” immediately evokes the image of a well-known biomaterial mainly used for cosmetic purposes. These marine bath sponge products, which can be found in various shapes and sizes commercially (Figure 6), are actually decellularized 3D skeletons of the sponges themselves, initially cleaned mechanically and chemically.

Figure 6.

A typical variety of American commercial sponges (A–D) (for overview see [76]) collected, cleaned, and bleached to the proper condition for sale. These photos were taken in November 2025 during our tour of the company “Tarpon Sponge Factory” in Tarpon Springs, FL, USA.

However, despite the general morphology of the sponge’s body being similar to the architecture of its skeleton, the external appearance of this organism in its natural habitat is very different from the physical form that is called a commercial sponge. For example, the surface layer of the widely known commercial sponge Spongia officinalis, known as ectosome (for details see [77]), is black, as can be seen in underwater photos or photos of freshly caught sponges (Figure 7A,B). This ectosome represents some kind of dark pigmented dermal crust (Figure 7D) made of debris and foreign spicules. Directly beneath this cortical layer is the sponge’s skeleton, represented by a dense 3D network of spongin fibres of various thicknesses arranged in different levels of hierarchical reticulation [77]. According to the classical work by Ernest Maylard from 1891, “this framework is composed of a substance called keratode, and is itself in the natural state, when living, coated with a soft gelatinous material, called “sarcode”" [78]. In contrast to the dark-colored surface layer, the interior of the bath sponge body ranges from white through cream to pale brown or from pale to bright yellow [79].

Figure 7.

Both photographs of S. officinalis bath sponge, originally found on the sea bottom (A) and recently collected (B), display the sponge’s natural black color. (Images courtesy of Donat Petricioli and Ivana Petrov Rančić). This coloration is caused by the dermal ectosome layer that covers the spongin-based fibrous skeleton, as shown in the schematic view (D). Such a skeleton can be extracted and chemically bleached to produce a commercial-quality product, as shown in (C) with the example of H. communis bath sponge.

A brief but informative overview of the main steps in the technology for manufacturing spongin scaffolds from bath sponges can also be found in Maylard’s work. He wrote, “on removing the sponges from the water, the gelatinous matter, which is of a brownish yellow color, begins to run freely from it. It is necessary, however to hasten this process in order to prevent the evil effects of putrefaction. For this purpose the sponge gatherers dig round, shallow holes in the sand along the shore which are filled with water. In these, the living sponges are placed and then trampled upon until the canals are entirely free of the gelatinous protoplasm, or “milk of the sponge”, and nothing remains but the keratose skeleton, known as the sponge of commerce. After this, sponges are submitted to further processes of cleansing, which may have some slight antiseptising effect. They are subjected, to baths of dilute hydrochloric acid and permanganate of potash.” [78]. It is therefore important to understand that the 3D biomaterial of bath sponge origin (Figure 1, Figure 6 and Figure 7C), which researchers from various scientific fields study, results from a chemical process called bleaching, well documented in literature and patents [80,81]. Reagents such as hydrogen peroxide, sodium bisulfite, and potassium permanganate have traditionally been used for bleaching bath sponges [82].

It is precisely the purified skeletons of bath sponges that enable us to examine this kind of multi-porous biological structure in detail, using various microscopy techniques, with digital and scanning electron microscopy (SEM) being the most revealing for studying the structural features of these ancient bioarchitectures (Figure 8).

Figure 8.

SEM images of selected fragments (A–D) of spongin-based scaffolds isolated from the H. communis commercial sponge under different magnifications. For better understanding, see also Figure 9.

It becomes well visible that the spongin-based skeleton of bath sponges belonging to the Spongiidae family (149 valid species, distributed among 6 genera) [77,79] contains diverse types of spongin fibres (primary, secondary, and tertiary), which can vary in thickness and density (Figure 8) as well as be located in different parts of the sponge body [77]. The texture of the entire bath sponge is resilient and compressible due to the hierarchical organization of interconnected spongin fibres. According to a classic definition, for example, for the S. officinalis keratosan sponge, “the skeletal network is composed of a reduced number of cored, primary fibers and highly developed, uncored secondary fibers, which gives this sponge its flexibility and water-retentive properties. The anastomosing of secondary fibers produces the skeletal reticulum, creating a meshwork of fibers. This mesh is formed by the intersection and joining of fibers, with each intersection always having three fibers branching from it” [79]. The structural principles of the spongin microfibres within the sponge skeleton become clearer when analyzing the SEM image of selected fragments, where the spongin-based meshes are arranged in layers, and the connecting “bridges” are shown in different colors (Figure 9).

Figure 9.

SEM imagery of selected fragments of H. communis bath sponge skeleton shows spongin-based meshes interconnected and arranged in layers (A). Additionally, larger-diameter fibres are connected at various levels by “microbridges” formed by the processes of the main fiber (B).

Such Lissajous element-like [83] 3D microstructural organization of the spongin scaffold allows it to be compressed, stretched, and torsionally stretched, as well as to absorb liquids through capillary action. According to the classical work by von Ledenfeld published in 1889, the shape of the spongin meshwork determines its compressibility, while its hardness is due to the regularity of the polygonal meshes and the rigidity depends on the thickness of spongin fibres [84]. Polygonal meshes in the skeletal architecture of an individual layer of the spongin scaffold can also be conveniently observed using digital microscopy. For example, in the skeletal monolayer of H. communis bath sponge scaffold carbonized at 2000 °C, trigonal, square, pentagonal, hexagonal, and heptagonal meshes become clearly visible (Figure 10).

Figure 10.

Digital microscopy image of polygonal meshes observable in structural monolayer of the scaffold fragment discovered on the surface of H. communis demosponge skeleton after carbonization at 2000 °C.

Statistically reliable information on the mechanical properties of 3D spongin scaffolds is a key factor in exploring their biomimetic potential for both tissue engineering and various technological applications. Up today such characteristics as absorption capacity, firmness, strength, elasticity, including compression modulus, compressive strength, tensile strength, elastic limit, elastic strain, modulus of elasticity, modulus of resilience have been obtained only for selected species of bath sponges skeletons (for details see [31,85,86,87]). It is suggested that a commercial sponge if applicable for industrial cleaning may require a minimum water retention efficiency of 20%, minimum elastic limit (strength) of 100 kPa, minimum firmness of 30 kPa and minimum modulus of elasticity (rigidity) of 700 kPa [86]. There is no doubt that the mechanical properties as well as the structural organization of keratosan spongin-containing skeletons, are species-specific. Therefore, the relevant research on the selection of spongin scaffolds with a specific architecture and porosity appears to be an insurmountable obstacle to optimizing these natural formations for effective use. Cultivating bath sponges under strictly controlled marine ranching conditions to produce skeletons that are more or less structurally homogeneous [86] appears to be an attractive, albeit still risky, approach.

3.2. Mineral Phases and Natural Strengthening of Spongin Microfibres

Marine keratosan demosponges represent one of the most outstanding natural examples of structural optimization of their spongin-based fibrous skeletons through both environmentally driven biomineralization and the collection of minerals in the form of microparticles. These features give the sponge high resistance to environmental stresses and help preserve the organism over the long term when the sponge body is exposed to underwater conditions currents.

Among keratosan demosponges, H. communis serves as a model for studying mineral incorporation in the skeletal spongin network. This species lives in shallow coastal areas with sandy or rocky substrates and acts as an effective suspension filter, pumping hundreds of liters of seawater daily through its aquiferous system [88,89,90]. Through natural filtration, mineral microparticles, colloids, microorganisms, and detritus are captured and gradually incorporated into the growing fibres of spongin [88,91,92]. This incorporation follows the growth lines of spongin microfibres, indicating that mineral phases are embedded during biomineralization process rather than sticking after death (Figure 11).

Figure 11.

Digital microscopic images of the mineral phases within the spongin fibres of H. communis demosponge: (A) natural spongin matrix; (B,C) mineral incorporations (arrows) following the growth lines of spongin fibres from the bottom to the top of the sponge body. (D) mineral residue after partial dissolution of spongin in 10% NaOH. White-colored mineral particles of similar size remain agglomerated into fibrous structures, suggesting the presence of spongin, which binds the particles together. These particles protect residual spongin from the chemical action of the alkali. Rust-colored microparticles (arrows) represent iron-based minerals.

Spongin microfibres of H. communis typically range from 5 to 100 μm in diameter, depending on the anatomical region and specimen maturity [31,93]. Thus, the mineral particles incorporated within these fibres vary in size and are distributed along the main axis of the fibres, often forming linear or clustered arrangements (Figure 11). Microscopy images reveal that the inclusions are not merely surface-adhered but are deeply integrated into the interior of the microfibres (Figure 12), forming continuous organic–inorganic structural hybrids. It can be suggested that such microstructural organization enhances the stiffness and compressive resistance of the spongin framework, converting it into a naturally reinforced biocomposite optimized for hydrodynamic environments.

Figure 12.

Digital microscopic images of the mineral phase within the spongin fibres of H. communis demosponge: (A) mineral phase embedded in the non-stained spongin matrix, appearing as light brown fibres; (B–D) spongin stained with Coomassie Blue, which indicates the presence of proteins, clearly showing the integration of mineral microparticles into the spongin fibres.

Micro X-ray fluorescence (MXRF) analyses performed on selected H. communis samples confirmed the heterogeneous mineralogical nature of the inclusions (For details see Supplementary Materials). The main detected elements include Si, S, K, Ca, Fe, Br, and I. The H. communis samples showed higher spatial concentrations of Si, K, and Fe aligned with the spongin fibres (Figure 13), while S, Ca, Br and I were distributed more uniformly, corresponding to the organic matrix [26,31] (Figure S1).

Figure 13.

MXRF analysis of the H. communis spongin mineral residue: (A) video image of the sample with the partially dissolved organic matrix; (B) element mapping of Si, K, and Fe with lines indicating typical spongin fibres microarchitecture.

The presence of various SiO₂ mineral phases in the spongin scaffold from H. communis demosponge is confirmed using X-ray diffraction (XRD), (see Figure 14). At least four mineral variants of SiO₂ are identified in the qualitative phase analysis: α-Quartz (ICSD 16331 [94]), Coesite (ICSD 172295 [95]), Cristobalite (ICSD 47219 [96]), Tridymite (ICSD 40896 [97]). The exact phase composition depends on the examined volume fraction of the sponge sample.

Figure 14.

XRD analysis of the H. communis spongin mineral residue showing presence of α-Quartz, Coesite, Cristobalite and Tridymite.

These findings align well with earlier studies on the mineralization of spongin-based skeletons, which reported the formation of natural iron-spongin biocomposites resulting from the sponge’s filtration of detrital microparticles [35,98,99]. It was demonstrated that ferruginous mineralization can lead to in situ formation of iron oxyhydroxides, goethite, and hematite-like phases within the spongin framework, thereby acting as a natural reinforcement mechanism [35,98,99]. Furthermore, microanalytical data indicate the presence of trace halogens (Br, I), suggesting long-term ionic exchange between spongin and the surrounding seawater. The presence of these halogens is consistent with the chemical structure of brominated collagen-like proteins typical for dictyoceratid demosponges [26,100]. The mineral phases embedded within the spongin microfibres serve both structural and mechanical function. The inorganic particles act as distributed load-bearing reinforcements, increasing the local stiffness of the spongin network and reducing deformation. These reinforcement mechanisms operate similar to those in engineered fiber-reinforced composites but are achieved here through passive environmental processes. The combination of flexible organic fibres and rigid mineral inclusions creates a hierarchical composite capable of resisting mechanical fatigue, bioerosion, and sediment abrasion—key factors in the dynamic benthic environment of H. communis habitats.

Importantly, the detection of several SiO₂ polymorphs within the spongin scaffold suggests that H. communis captures siliceous microparticles originating from geologically diverse sediment sources rather than undergoes in vivo phase transformations. Thermodynamically stable quartz, metastable cristobalite and tridymite, and even high-pressure coesite are all naturally present in coastal sediments [94,95,96,97]. Their coexistence within the bath sponge skeleton likely results from episodic sediment transport, storm-driven resuspension, or long-distance movement of fine particulate matter.

From a mechanical standpoint, the presence of SiO₂ grains with different crystallographic structures and hardness may introduce subtle microscale variations in stiffness within the organic scaffold. This contributes to the overall mechanical robustness of the sponge. This naturally formed, mineral-graded architecture is similar to engineered composites with spatially distributed hard phases, offering increased resistance to abrasion and hydrodynamic stress [32].

The ability of spongin to capture microscopic mineral particles from the environment should also be considered when studying paleontological sponge specimens [24]. It is entirely possible that the mineral remains of fossilized skeletons of keratosan sponges originate from the process described above. In these studies, staining spongin residues with a specific protein dye like Coomassie blue can be very effective for identifying the exceptionally well-preserved organic material phase.

The incorporation of mineral particles also has ecological implications. As the sponge continuously filters water, it accumulates mineral grains that reflect the geochemical signature of the surrounding seabed [88,91,101]. Therefore, chemical analysis of the spongin mineral fraction can provide valuable insights into local sediment composition, water chemistry, and particulate fluxes. This property makes H. communis and related demosponges potential bioindicators of benthic mineral dynamics and marine pollution.

Understanding the mechanism of natural mineral strengthening in spongin is highly valuable for modern biomimetics and tissue engineering. The hybrid spongin–mineral composite demonstrates how environmental particles can be transformed into functional reinforcement structures through self-organization. Insights from such systems inspire the design of self-mineralizing collagen-based scaffolds with gradient mechanical properties, suitable for biomedical applications (e.g., bone and cartilage regeneration) and sustainable material engineering [93,102]. Controlled mineral impregnation of spongin-like biomaterials could replicate the adaptive stiffness and durability found in marine sponge skeletons.

3.3. Liquid Absorption Capacity of Spongin Scaffolds

The widespread use of bath sponges by humans since ancient times has been primarily driven by their ability to absorb water and other liquids, as well as by their capacity to squeeze these liquids out, which determines their many possible uses. Because of their high water absorption capacity, skeletons of marine sponges were initially used as natural containers for drinking water. According to Creswell, “every Roman soldier had a sponge which he used in lieu of a cup for drinking purposes on long marches” [103].

The phenomenon relies on a combination of the sponge’s internal and surface structure, porosity, surface tension, wettability, and capillary action. For example, as Von Lendenfeld states, “the more numerous the fibres per unit of volume, the greater the capillary action and the more water the sponge can hold” [84]. The shape of the pores also plays a key role; small pores hold water tightly but fill more slowly and release less easily, whereas larger pores allow for faster intake and release (see [104] for details). The influence of pressure gradients should also be considered when water moves within a spongin scaffold through surface pores into an interconnected, channel-based sophisticated fibrous network (Figure 7C).

Cui and co-workers reported that the commercial sponge typically exhibits a water absorption capacity of about 0.23 mL cm⁻³ [105]. Water absorption and water retention efficiency were also measured for various bath sponges in the study by Louden [85]. Nowadays, the capillary effect of natural sponge inner architecture serves as the basis for bioinspiration in developing high-performance artificial sponges [106]. Our preliminary experiments with wetting a 50 × 1 × 1 cm flat S. lamella spongin scaffold, cut from water-wetted material, demonstrated its remarkable ability to alter the rigidity of the skeleton to the point that knotting became possible (Figure 15).

Figure 15.

A cord-like fragment cut from a water-soaked S. lamella spongin scaffold (A) becomes so elastic that it can be twisted into knots (B,C).

We varied the behavior of the S. lamella spongin scaffolds during immersion in different liquids. In most cases, the spongin sample sinks after absorbing the solution; however, in simulated seawater, red wine, beer, and glycerin, the spongin rises to the surface of the liquid, as shown in Figure 16. A unique property of the spongin scaffold is its ability to fully retain absorbed liquid within its structure, as demonstrated in Figure 17. The only exception to liquid retention was observed with pork blood.

Figure 16.

Photographs of spongin scaffolds from S. lamella bath sponge immersed in various liquids used to measure the liquid absorption capacity test.

Figure 17.

Photographs of spongin scaffolds from S. lamella bath sponge taken 5 min after removing them from the subsequent liquids used for measuring liquid absorption capacity test.

The volumetric results of the liquid absorption test for various liquids with different properties are presented in Table 2. All data collected during this test are also available in the Supplementary Materials (Table S1). This test demonstrates how large volumes of liquids can be absorbed by the tiny, lightweight skeletons of the marine sponge under study. The amount of liquid absorbed ranges from 30 to 45 mL per 1 g of S. lamella spongin sample (see Table 2). Therefore, we found that the spongin scaffold can increase its weight by as much as 50 times (see Table S1 in Supplementary Materials). Comparing water absorbency to the results from Louden [85], spongin from S. lamella demosponge has higher water absorbency than that from Rhopaloeides odorabile and Coscinoderma species, and it reaches a similar level to H. lachne, also known as “wool sponge,” which had the highest absorbency among all materials tested by Louden.

Table 2.

Liquid volume absorbed per 1 g of S. lamella spongin scaffold. Data are presented as an average value ± standard deviation, calculated from 3 samples repetitions.

No.	Liquid	Density * [g/mL]	Absorbed Liquid Volume per 1 g of Spongin Scaffold
1	tap water	0.99	31.3 ± 5.8 mL
2	distilled water	0.99	33.5 ± 7.7 mL
3	simulated seawater	1.16	32.7 ± 6.7 mL
4	dental water	1.00	29.6 ± 4.1 mL
5	sugar syrup	1.24	40.9 ± 3.2 mL
6	pork blood	1.04	41.6 ± 7.1 mL
7	physiological saline (0.9% NaCl)	1.00	45.0 ± 6.9 mL
8	human plasma	1.01	38.1 ± 7.3 mL
9	serum (FBS)	1.01	34.2 ± 4.8 mL
10	cells medium (DMEM)	1.00	39.5 ± 7.2 mL
11	vinegar, 10%	1.01	36.4 ± 3.9 mL
12	red wine	1.01	31.6 ± 1.8 mL
13	beer	1.00	35.0 ± 1.8 mL
14	glycerin	1.20	30.5 ± 1.2 mL
15	hyaluronic acid, 1%	1.01	36.5 ± 2.2 mL

* density measured under the experimental conditions.

The highest absorption level by spongin scaffolds from S. lamella bath sponge is observed for physiological saline, while the lowest is seen with dental water. Significant absorption is also observed with biological media such as defibrinated delipidated human plasma, cell medium (DMEM), as well as with sugar syrup and pork blood. This indicates that marine spongin scaffolds have high potential for biomedical applications, including tissue engineering. The errors observed in the results for three repetitions, shown in Table 2, are related to the structural features of this spongin scaffold, which is naturally never completely homogeneous.

The relationship between spongin matrices and blood, both historically rooted and still under modern research, highlights their ongoing connection. It is well recognized that hemostasis is the first step in wound healing [107]. Inspired by the blood absorption behavior of natural bath sponge skeletons, a wide variety of hemostatic sponge-like scaffolds composed of different natural and synthetic polymers, as well as silicon- and metal-based materials, have been developed over the last decade, including 3D printing approaches (see overview in [108,109,110,111,112]).

Unfortunately, modern studies on the hemostatic properties of spongin matrices are almost nonexistent. Therefore, the work by Wang and colleagues titled “Rapid Hemostatic Biomaterial from a Natural Bath Sponge Skeleton” [37], deserves special attention. In this study, the hemostatic effect of spongin from S. officinalis bath sponge and the derived spongin materials called SX, SFM, and SR was investigated on rats and rabbits. It was shown that SX, SFM, and SR were significantly more effective than type I collagen, used as the positive control, in shortening the whole blood clotting time in vitro and promoting hemostasis. Spongin derivatives have been recommended as potential hemostatic agents for uncontrolled bleeding and hemorrhagic diseases [37].

3.4. Surface Wettability of Spongin Microfibres and Capillary Effects

Spongin specimens have been obtained from bath sponges such as S. lamella, S. tampa, and H. communis. The water contact angle (WCA) analysis performed on selected spongin surfaces revealed high values exceeding 120° (Figure 18, Table 3). Such elevated contact angles typically indicate a potentially hydrophobic surface; however, in the case of the examined samples, this behavior does not result from the intrinsic chemical properties of the biomaterial but rather from morphological and capillary effects characteristic of highly porous, fibrous structures [113,114].

Figure 18.

Positioning a water droplet during contact angle measurement of spongin isolated from such sponges as (A) S. lamella, (B) S. tampa, (C) H. communis using a goniometer. (D) Image of the water droplet on the surface of a 3D spongin scaffold of H. communis demosponge origin.

Table 3.

Results of water contact angle measurements on spongins.

Sample	Water Contact Angle [°]	SD [°]
Spongin from S. lamella	122.3	0.3
Spongin from S. tampa	122.5	0.5
Spongin from H. communis	132.9	1.3

The surface of this type of material does not form a uniform, continuous contact with the liquid. During measurement, water only interacts with the outermost parts of the fibres, while a significant portion of the droplet remains suspended above the surface, supported by its microstructure [115]. As a result, the initial contact angle is high despite the lack of true chemical hydrophobicity of the material. This behavior can be explained by the limited and delayed penetration of the liquid into the interior of the 3D structure. In the early stage of contact with water, the fibrous network of the spongin surface does not saturate immediately, as the flow pathways are discontinuous and the fibres slow down the absorption process.

Similar phenomena have been described in the literature for wool fabrics with micro- and nanostructural features [116,117]. In such systems, a high contact angle results from complex topographical and capillary effects, and the measured liquid-surface interaction is only apparent, which may significantly differ from the true value. Furthermore, fabrics contain a capillary network both between and within yarns, making their surface far from perfectly flat, smooth, and uniform [116,117].

The high contact angles observed for spongin in this study result from its heterogeneous, rough microstructure rather than from any intrinsic chemical hydrophobicity. After prolonged contact with water, the examined spongin scaffolds exhibit hydrophilic behavior typical of capillary materials, characterized by liquid absorption and penetration into the material’s fibrous skeletal network.

Various natural sponge-like 3D structures have already been fabricated from materials such as different synthetic polymers, including melamine, polyurethane (PUR), and poly(dimethylsiloxane) (PDMS), as well as carbon-based materials (see overview by [118]). However, creating these sponge scaffolds analogs involves extra costs and specialized technological methods, along with environmental concerns linked to hazardous manufacturing processes. Chemical modification of spongin matrices, a renewable natural raw material, seems to be a promising approach alternative.

Coating the spongin with molding silicone and PUR (blue-colored sample, Figure 19) caused significant changes in the surface morphology. Compared to the natural samples, which display a highly developed, rough fibrous structure, the surfaces of the treated spongins became noticeably smoother and more uniform after impregnation. The images (Figure 19) reveal that the fibres were covered with a thin, flexible layer of silicone or polyurethane, which partly filled the pores and connected adjacent parts of the 3D structure. As a result, the exterior surfaces of the spongin samples coated with silicone or PUR show fewer protruding fibres than the unmodified ones. A similar effect was seen inside the spongin matrix as well. Areas fully immersed in silicone or PUR during impregnation also exhibit significant smoothing. The treated fibres in these regions are partly bonded together by a thin layer of silicone or PUR. The contact angles of the modified spongin samples range from 118–129° (Table 4), due to the chemical hydrophobicity of the coating. Additionally, the observations suggest that impregnation partially seals the spongin structure, reducing its absorbency but not completely eliminating it. Both molding silicone and PUR effectively modify the topography and wetting properties of spongin, giving its surfaces a more uniform and durable hydrophobic character (Figure 19).

Figure 19.

(A) Spongin from H. communis sponge coated with PUR. (A′) Water droplet during contact angle measurement on the marked outer surface. (B) Interior of spongin coated with PUR. (B′) Water droplet during contact angle measurement in the interior region. (C) Spongin coated with molding silicone. (C′) Water droplet during contact angle measurement on the marked outer surface. (D) Interior of spongin scaffold coated with molding silicone. (D′) Water droplet during contact angle measurement in the marked interior region.

Table 4.

Results of water contact angle for H. communis spongin/silicone and spongin/PUR surfaces.

Sample	Water Contact Angle [°]	SD [°]
Spongin coated with PUR	118.1	4.1
Interior region of spongin coated with PUR	126.9	3.1
Spongin coated with molding silicone	128.8	2.4
Interior region of spongin coated with molding silicone	125.1	3.8

Although natural spongins initially show high water contact angles, this is mainly due to their mesoporous, fibrous structure, which ultimately leads to liquid absorption. Chemical modification, such as coating spongins with silicone or PUR, creates a more even surface and provides long-lasting hydrophobic qualities. This approach offers a promising way to stabilize wettability and expand the potential uses of spongin-based materials scaffolds.

In practice, this means that the measured initial contact angle is high not because the material is chemically hydrophobic, but because the water droplet contacts only the tips of the fibers, while a significant part of the droplet remains “suspended” above the surface due to the material’s microstructure. Importantly, this angle is not constant over time. The study also presents surface modification of the spongin with silicones. In this case, a high contact angle is also observed; however, its origin is different, as it results from the chemical hydrophobicity of the silicone coating. Consequently, no water absorption is observed because the silicone surface effectively prevents liquid penetration into the interior of the material.

From the perspective of practical applications, this behavior is significant: unmodified spongin, despite the high initial contact angle, retains the ability to absorb liquids, whereas after silicone modification, they become hydrophobic materials that limit water ingress, which may be advantageous in applications requiring moisture resistance.

3.5. Advances in Metallization of Spongin Scaffolds

Research on the interaction of metal ions and the metals themselves in various nanophase forms with spongin matrices appears to be in demand for the following reasons. First, metal ions such as mercury [119] or copper [33], which are often found in highly toxic wastewater, actively interact with the surface of spongin, forming complexes and nanostructured composites (Figure 20) with catalytic properties [33,120].

Figure 20.

Top immersion of a portion of a spongin scaffold isolated from H. communis bath sponge in a toxic wastewater solution of copper chloride in ammonia results in the formation of a rigid spongin-atacamite composite (A). The resulting nanostructured 3D material (B) remains stable after ultrasonic treatment and can be used as a highly effective catalyst on an industrial scale (for details see [33]).

Recent research on the metallization process of spongin has resulted in the creation of unique 3D “iron-spongin” composites containing lepidocrocite (γ-FeO(OH)). These composites have been produced using two methods, biomimetics and extreme biomimetics, and exhibit strong capacities for detecting dopamine and tryptophan [35,98,121]. Furthermore, by melting steel at 1450–1600 °C on initially carbonized H. communis spongin scaffolds at 1200 °C, new magnetic 3D spongin-based materials were recently produced [122]. The development of magnetic sponges using natural spongin scaffolds appears to be a very important direction and a clear alternative to the many synthetic analogues that have already found application in controlled drug delivery [123]., force sensing, energy harvesting [124], remediation [125], and tissue engineering [126].

In the following subsections, we present the results of our studies on coating spongin scaffolds with titanium nanoparticles using the ion–plasma deposition method, chromium plating, and extreme biomimetic interaction with iron ions.

3.5.1. Creation of Spongin–Titanium 3D Composites Using Ion–Plasma (Vacuum Arc) Deposition Method (VAD)

The aerospace industry, chemical processing, and medicine are the main consumers of titanium in the form of porous structures. The so-called “titanium sponge” represents a form of porous agglomerated powder particulate [127]. It should be immediately noted that, in the classical sense, "titanium sponge" differs significantly from the spongin-based skeletons of commercial sponges. However, titanium foams, for example, due to their mechanical robustness, corrosion resistance, biocompatibility, and bioactivity, have already found diverse orthopedic and dental applications [128,129,130,131]. Also, 3D-printed, highly porous titanium implants have recently been introduced into knee and hip arthroplasty [132]. The porosity of titanium structures for biomedical purposes is an important factor. For example, as reported by Müller and co-workers, human osteoblasts can grow well through the interconnected porosity of the titanium foam [133].

Given the above facts, the idea of coating a ready to use 3Dspongin scaffolds covered with titanium nanoparticles in such a way that the skeletal architecture itself would be preserved seemed intriguing, albeit ambitious at the same time. The question remained open whether spongin as a biomaterial would withstand the specific conditions under which titanium nanoparticle deposition would occur. A wide range of metallic coating deposition methods has been developed to date, among which Physical Vapor Deposition (PVD) techniques occupy a prominent position [134]. The most commonly used PVD technique is magnetron sputtering, which, despite its advantages, demonstrates relatively low deposition rates—particularly when applied to materials such as Ti, Mo, or Si—and is therefore mainly employed for producing thin films. From both technological and design perspectives, a promising alternative is the ion–plasma (vacuum arc) deposition method (VAD) (see for overview [135]). This technique enables the formation of dense, uniform coatings from virtually any metal or alloy with strong adhesion to the substrate, which is attributed to the high ionization degree of the cathode erosion products.

Titanium was selected as the cathode material because of its high chemical reactivity, which allows it to wet non-metallic and ceramic surfaces, including carbonized ones. The cathode, made of this coating material, erodes through a vacuum arc, enabling high deposition rates on various types of surfaces [136]. This specific method and the associated equipment were used for metallizing H. communis bath sponge 3D spongin scaffolds, with more detailed descriptions provided in [137] (see also Section 2). Consequently, the experiments (Figure 21 and Figure 22) demonstrate, for the first time, the feasibility of metallizing spongin with titanium using a very harsh ion–plasma deposition method. The spongin biomaterial, in an argon atmosphere, withstood an arc current of 100 amperes and an arc voltage of 70 volts while being sprayed with titanium nanoparticles for 5 and 10 min, without significant changes to the 3D architecture of the scaffold. However, a mass loss (Δm) ranging from 0.01 to 0.1 g was observed, depending on the sample size.

Figure 21.

Digital microscopy images reveal a distinct difference in the surface of spongin scaffolds before (A) and after metallization with titanium nanoparticles (B) under ion–plasma deposition conditions. A similar pattern is observed when examining the surface of these spongin fibres using SEM at higher magnification. The characteristic microstructural organization of the natural spongin fiber surface (C) is drastically changed when coated with titanium. The nanostructured spherical architecture of the metallized spongin surface becomes clearly visible (D).

Figure 22.

SEM image of a titanium-coated spongin fiber of H. communis demosponge origin at a site of mechanical rupture (A). Elemental mapping (B) clearly shows the distribution of titanium within the surface layer of spongin, a structural biocomposite material.

As a result of metallization, the surface of the spongin fibres was coated with a uniform titanium film. Interestingly, the thickness of this titanium-coated layer (Figure 21D) reaches 500 nm (Figure 22), corresponding to the so-called cuticular layer of spongin fibres [138,139]. Unlike the collagen core of spongin fibres [26], the nature and origin of this layer are still unknown, although it is believed to be where sulfur-containing proteins are concentrated [35].

It is logical to assume that now, after creating a 3D spongin–titanium composite material, it will be necessary to study its physical and mechanical properties in detail, considering the prospects for practical application in biomedicine, similar to the already well-tested porous titanium implants. Titanium can also serve as an interlayer for applying a functional coating layer of various metals, including Ni, Ag, and Au [140,141]. Consequently, metallized spongin scaffolds may also find applications in various fields of engineering and technology.

3.5.2. Chromium Tanning of Spongin Scaffolds

The measurement of the sample mass before and after the chromium tanning process demonstrates that a reaction has occurred between spongin from H. communis demosponge containing lepidocrocite and chromium ions. The increase in the mass of the chromium-tanned sample after ultrasound application, compared to the initial mass, is 69.4%. Using an extreme biomimetic approach to chromium tanning of a bath sponge skeleton and subsequent carbonization of the resulting composite led to the creation of a new material with a complex structure. Digital and scanning electron microscopy (SEM) analyses showed that exposure to high concentrations of chromium solution does not compromise the 3D architecture of the spongin scaffold.

Images from the camera (Figure 23A) and digital microscopy (Figure 23C) show a noticeable greenish tint of the fibres after the chrome tanning process, compared to the untreated control sample (Figure 23B). Additionally, many dark regions are visible, indicating the incorporation of chromium compounds into the spongin matrix (Figure 23C). SEM images further confirm the formation of an extra layer of spongin fibres on the chrome-tanned sample compared to the control (Figure 23G vs. Figure 23F). These findings are in agreement with previous studies on chromium-tanned leather materials [142,143]. After carbonization, the spongin fibres display microspherical and microcrystalline formations on their surface, along with distinct needle-like nanostructures (Figure 23D,E,H,I). These structures show the scaffold’s ability to maintain its structure and support the formation of chromium-containing micro- and nanostructures under high temperatures under high-temperature conditions.

Figure 23.

(A) Image from the camera of H. communis spongin containing lepidocrocite after chromium tanning in 50% Cr(OH)SO₄ at 40 °C for 48 h after ultrasound treatment (UST). Digital microscopy images of H. communis spongin containing lepidocrocite: (B) control sample; (C) sample after chromium tanning in 50% Cr(OH)SO₄ at 40 °C for 48 h after UST—the highlighted areas show the accumulation of chromium compounds covering the spongin fibres; (D,E) sample after chromium tanning in 50% Cr(OH)SO₄ at 40 °C for 48 h after UST and after carbonization at 1450 °C for 90 min and after UST. SEM images of H. communis spongin containing lepidocrocite: (F) control sample; (G) sample after chromium tanning in 50% Cr(OH)SO₄ at 40 °C for 48 h after UST; (H,I) sample after chromium tanning in 50% Cr(OH)SO₄ at 40 °C for 48 h after UST and after carbonization at 1450 °C for 90 min and after UST.

The analysis of the chromium tanning process and the needle-like structures observed suggests the possible presence of Cr₂O₃ and chromium carbides after the carbonization process. Under specific furnace cooling conditions, these could have formed needle-like crystals in the analyzed sample [144,145,146,147]. At high temperatures, the residual chromium (III) oxide remaining in the material after chromium tanning acts as a precursor for carbide formation during carbonization. We propose that as the organic matrix decomposes, carbon becomes available and participates in reducing Cr₂O₃. The overall mechanism begins with an initial CO/CO₂ transport stage, followed by a reaction between the reduced chromium species—mainly Cr₃C₂—and the remaining oxide phase [148]. The composition of the resulting carbides depends on both the available carbon and the efficiency of carbon transfer to the oxide surface. According to Berger and colleagues [148], when temperatures exceeds approximately 1600 °C, Cr₃C₂ becomes the dominant and ultimately the only stable phase. If the carbon supply is insufficient, additional carbide phases such as Cr₇C₃ or Cr₂₃C₆ may also form. These carbides tend to form a protective layer around unreacted Cr₂O₃, affecting further reduction kinetics [148].

XRD analyses (Figure 24) showed that after carbonization of the chromium-tanned sample, the crystalline form of Cr₂O₃ is formed. As a result of its partial reduction, crystalline forms of chromium carbides Cr₃C₂ and Cr₇C₃ are also formed. The process of reducing Cr₂O₃ to chromium carbides is well documented in the literature [148]. Peaks indicating the presence of Cr₂O₃ are observed at 2θ values: 24.5°, 33.6°, 36.1°, 54.9°, 63.5°, and 65.2°, respectively [149,150]. Peaks at 42.8° and 43.7° indicate the presence of Cr₇C₃ [151], and peaks at 39.0° and 40.3° confirm the presence of Cr₃C₂ [151,152].

Figure 24.

XRD analyses of H. communis spongin containing lepidocrocite after carbonization at 1450 °C: (A) control sample and (B) sample after chromium tanning (before carbonization) in 50% Cr(OH)SO₄ at 40 °C for 48 h.

3.6. Coating of Spongin Scaffolds with Iron Oxides

Iron oxides and oxyhydroxides have been extensively studied for their catalytic, adsorptive, magnetic, and sensing properties and are widely used in various technological applications [35,98,153,154,155]. However, the performance of these materials is strongly dependent on deposit morphology and matrix support characteristics [156]. Consequently, the creation of iron oxide-based 3D bioinorganic templates, including poriferan chitin and spongin, remains a trend in modern bioinspired materials science.

Up to today, several biomimetic and extreme biomimetic approaches for creating lepidocrocite (γ-FeOOH) and goethite (α-FeOOH)-based 3D spongin scaffolds have been reported [35,98]. Additionally, selective deposition of both the lepidocrocite and the goethite mineral phases on spongin fibres using electro-assisted deposition has been proposed [153].

In addition to the previous methods, a new two-step process was created to produce iron oxide–spongin composites. First, spongin scaffolds from H. communis demosponge were soaked at room temperature in artificial seawater enriched with Fe ions for seven days to structure the scaffolds, following a biomimetic biomineralization approach. After a week, the partially mineralized scaffolds were placed inside a sealed 3L glass chamber, with a separate chamber containing 35% HCl solution (Figure 25B). The scaffolds were not directly submerged in the acid but were instead exposed to the HCl vapor for 12 h.

Figure 25.

Preparation and characterization of iron oxide−spongin composites via the vapor-assisted biomimetic method. (A) H. communis spongin scaffold after 7 days of incubation in artificial seawater enriched with Fe ions. (B) Exposure of the pre−mineralized scaffold to HCl vapor in a closed glass chamber. (C) Iron oxide−spongin composite after 2 h ultrasonication, showing a homogeneous reddish-brown coating. (D) Digital microscopy image of pristine spongin with its typical smooth fibrous 3D network. (E) Digital microscopy of the composite after mineralization, HCl vapor treatment, and ultrasonication, revealing a well-preserved structure with a uniform iron oxide layer indicated by the rusty color. (F) SEM image of created composite after ultrasonication, confirming continuous and densely packed inorganic layers strongly adhered to spongin fibres. (G) Higher−magnification SEM image of the composite surface, showing tightly bound iron oxide layers on spongin fibres. (H) SEM image of pure H. communis spongin, displaying clean, deposit-free fibres for comparison. (I) FTIR spectra of pure H. communis spongin and the iron oxide–spongin composite obtained after HCl vapor-assisted treatment, confirming chemical modifications related to lepidocrocite mineral deposition.

Microscopic examinations revealed distinct differences between pristine spongin and the composites produced (Figure 25D,E). After biomimetic mineralization and subsequent HCl vapor treatment, the spongin fibres were uniformly coated with a brownish iron oxide layer, creating a rougher surface texture without damaging the scaffold’s overall structure or microporosity. Importantly, even after two hours of ultrasonic treatment at room temperature, the composite remained intact, demonstrating its overall robustness. Additionally, SEM observations (Figure 25F–H) confirmed the successful mineralization and stability of the iron oxide coating on the spongin surface fibres.

FTIR spectroscopy confirmed the successful mineralization of spongin and the formation of iron oxyhydroxide phases through steam-assisted HCl treatment (Figure 25I). For example, the bands around 1150 and 1021 cm⁻¹ are generally associated with Fe–O–H bending vibrations, while the bands around ~740 and ~570 cm⁻¹ likely represent Fe–O stretching modes and are frequently reported for γ-FeOOH (lepidocrocite) [157,158]. In contrast, the IR spectrum of pure spongin mainly consists of absorptions indicative of the protein material: amide I (~1630–1650 cm⁻¹), amide II (~1540 cm⁻¹), amide III (~1240 cm⁻¹) bands, and the broad O–H / N–H stretching around ~3300 cm⁻¹. These pattern matches modern FTIR descriptions of collagen-based matrices and reflect spongin’s collagenous nature [26,159,160].

Overall, these findings demonstrate the versatility of spongin as a natural 3D scaffold for creating iron oxide composites. The methods described vary in terms of time, controllability, and phase selectivity, but all highlight the potential of organic scaffolds and iron oxides to generate durable, multifunctional materials suitable for catalysis, sensing, and environmental applications.

3.7. Carbonization of 3D Spongin Scaffolds

Biopolymer spongin obtained from industrially cultivated marine sponges has already been recognized as a promising and proven candidate for carbonization up to 1200 °C in the case of H. communis marine demosponge [32] (Figure 26). Its unique 3D hierarchical architecture, combined with thermal stability up to 360 °C, makes spongin an exceptional, sustainable natural scaffold for developing functional carbon materials via carbonization. Furthermore, the possibility of optimizing carbonization parameters, followed by surface activation or modification, allows for the creation of advanced spongin-derived carbon composites at the nanoscale, suitable for diverse technological, catalytic, and environmental applications.

Figure 26.

Comparative photographs of H. communis spongin scaffold before (A) and after (B) carbonization at 1200 °C demonstrate the exceptional preservation of this type of matrix at the macro level, which still retains its unique macro- and microporosity, including a branched and interconnected channel system. The carbonized scaffold consists of turbostratic graphite and remains mechanically strong enough to be cut into pieces or slices of the desired shape and size using a metal saw.

Digital light microscopy images shown in Figure 27 demonstrate that the spongin scaffolds from various sponge sources retain their strength, 3D structure, microporosity, and uniformity even after being exposed to carbonization temperatures up to 2000 °C.

Figure 27.

Digital microscopy images of typical 3D architecture of spongin scaffolds isolated from bath sponge species such as H. communis (A), S. lamella (C), and S. tampa (E), before (A,C,E) and after carbonization at 2000 °C (H. communis), 1200 °C (S. lamella), and 800 °C (S. tampa), respectively (B,D,F).

The data presented in Supplementary Materials Table S2 show the results of analysis of selected regions of the carbonized spongin structure shown in Figure 28B. The measurements clearly demonstrate that the polygonal pores—like meshes within the material- are highly variable in size and shape, including both larger and smaller meshes, as indicated by the wide range of measured areas and diameters. Such heterogeneity is typical for spongin architecture, and importantly, it remains well preserved even after the carbonization process. The analyzed regions have an average area of 11,373.14 µm², with maximum values reaching 23,762.56 µm², confirming substantial variability in pore size. The total measured pore area is 159,223.97 µm², which accounts for 75.29% of the analyzed region.

Figure 28.

Digital microscopy image of the H. communis spongin fragment carbonized at 2000 °C (A), and the image selected for calculations of the area, pore size distribution, and pore diameter (B).

Recent studies have confirmed the potential of carbonized spongin scaffolds as effective biological templates for creating advanced functional composites. For example, Szatkowski and co-workers [161] produced a 3D MnO₂/carbonized spongin composite that preserved the sponge’s fibrous structure and demonstrated stable electrochemical performance over multiple charge/discharge cycles. Petrenko and co-workers developed a carbonized spongin composite after carbonization at 1200 °C, resulting in a turbostratic graphite-like structure with high thermal and mechanical stability [32]. Copper electroplating of this composite led to a hybrid material with excellent catalytic performance for reducing p-nitrophenol in both freshwater and marine environments. More recently, Leśniewski and co-workers demonstrated that molten steel can be integrated with carbonized spongin scaffolds processed at 1200 °C to produce composites coated with magnetic iron oxides. These constructs exhibited electrocatalytic activity toward the hydrogen evolution reaction (HER) [122]. Application of mechanically stable carbonized spongin scaffolds in tissue engineering and biomedicine remains to be the next challenging task in near future.

3.8. 3D Spongin-Based Scaffolds and the Shape-Memory Phenomenon

Digital microscopy observation of the at 100 °C compressed spongin revealed a uniform, compact, and densely packed structure (Figure 29), indicating effective material densification. Despite significant compression, the fibres remained intact and preserved their structural integrity—no breakage or deformation was observed. This demonstrates the high elasticity of the biomaterial and its resistance to permanent deformation even after heating up to 100 °C. Such behavior reflects the continuity of the fibrous microstructure of spongin, which is essential for the functional performance of porous materials. It is to note, that spongin remains to be thermostable up to 360 °C as reported previously [162]. The pressed specimen showed no deformation over time, retaining a stable, cohesive form, confirming its mechanical durability in the dry state.

Figure 29.

Schematic illustration of the compression and rehydration cycle of spongin from bath sponge S. tubilifera, demonstrating its shape-memory behavior and reversible structural recovery after 2 months. Digital microscopy images show spongin after the pressing process (A) and after shape recovery and drying (B). Translation for "Uwaga gorąca powierzchnia!" is "Caution hot surface!"

After two months, the sample was immersed in distilled water for two hours. During this time, a gradual recovery of the original shape was observed (Figure 29), indicating the presence of a shape-memory phenomenon and the ability of this unique biomaterial to undergo reversible deformation in an aqueous environment.

Microscopic imaging after soaking in water (Figure 29B) revealed intact fibres and restored inter-fiber spaces of spongin, confirming complete structural regeneration. This behavior is characteristic of smart materials such as shape-memory polymers (SMPs) [163,164,165].

The presented experiment provides the first evidence of shape-memory behavior in native 3D spongin scaffolds after heating up to 100 °C. This finding is particularly significant, as it demonstrates spongin’s capacity to undergo reversible deformation and full structural recovery after compression—a phenomenon previously mainly associated with synthetic shape-memory polymers [166]. Equally interesting results were obtained when pressing under the same conditions selected rusty-colored bath sponges, in which the spongin fibres were originally biomineralized with various iron oxides, including lepidocrocite and goethite (see [35,98]). As shown in Figure 30, even in biomineral-containing spongins, shape memory was observed without any noticeable interference.

Figure 30.

Placing a drop of water on the far left corner of a strip of rusty S. officinalis bath sponge pressed two months ago (A) causes it to start swelling within 15 min (B). After 45 min, this process continues to progress (C,D), even though the spongin fibres remain biomineralized with iron oxides.

The above results served as an incentive to conduct a series of additional experiments on pressing spongin matrices using S. lamella and H. communis bath sponge scaffolds pre-modified with silver nitrate.

S. lamella spongin initially treated with silver nitrate, after pressing at both 120 °C (Figure 31) and at room temperature (Figure 32), exhibits tightly packed fibres that resist deformation and breakage. After inserting the pressed samples into distilled water for 19 h (Figure 31C,D and Figure 32B,C), the fibres return to their original shape, and importantly, they remain coated with silver, just as they were before pressing.

Figure 31.

(A,C) Camera images of S. lamella spongin treated with a 0.1 M silver nitrate solution (reduction method with fructose) after pressing (150 kN) at 120 °C: (A) before immersing in distilled water and (C) after immersing in distilled water. (B,D) Digital microscope images of the sample: (B) before immersing in distilled water and (D) after immersing in distilled water.

Figure 32.

(A,C) Camera images of S. lamella spongin treated with silver nitrate (1% solution, autoreduction of AgNO₃) after pressing into a tablet form (150 mbar): (A) before and (C) after immersing in distilled water. (B,D) Digital microscope images of the same sample in tablet form: (D) before immersing and (B) after immersing in distilled water.

This experiment demonstrates that the spongin-silver composite is very elastic and resistant to compression, similar to untreated spongin (see Figure 29), and exhibits shape memory phenomenon. After pressing, metallic silver is clearly visible on the material’s surface as tightly packed microstructures (Figure 31B and Figure 32C). This fact could have a positive and decisive impact on the future applications of such materials in medicine, biomaterials engineering, filters, or catalysis.

Although we have documented for the first time the memory shape phenomenon in both native and modified spongin as 3D scaffolds, the mechanism behind this phenomenon using this unique biomaterial still needs to be clarified. Nonetheless, the results already obtained open new opportunities for applying spongin scaffolds in various fields including:

• •Medicine (as active shape-memory-dressings, which contact with corresponding biological fluids, ensuring better conformity and absorption [167,168,169]
• •Biomaterials engineering (as scaffolds for cell culture or temporary implants, benefiting from natural biocompatibility and elasticity [167,170].
• •Filtration and sorption technologies (as 3D filtering systems) [171].

This discovery not only enhances the current understanding of spongin’s structure-function features but also paves the way for developing new bio-based technologies that combine high functionality with ecological sustainability. An additional experiment was performed to produce structures capable of maintaining the container’s shape without reverting to their original form. The preparation involved impregnating the spongin scaffolds with a molding silicone mixture, then placing the material in a vessel and compressing it to the desired mold height at 100 °C for 60 s. The resulting materials, shown in Figure 33, preserved the imposed geometry and showed no recovery after compression, indicating permanent structural fixation.

Figure 33.

A composite of spongin and molding silicone can be used as a source for creating materials of a given shape and size after transformation in a hydraulic press.

The ability to mechanically process carbonized spongin scaffolds into defined shapes and dimensions further enhances their practical applicability. Such processability, combined with preserved cytocompatibility, may support future patient-specific scaffold design following appropriate biological validation.

3.9. Poriferan 3D Skeletal Scaffolds and Tissue Engineering

Representatives of both the Verongiida and Dictioceratida orders of keratosan demosponges serve as an exceptional source of naturally pre-organized 3D skeletal architectures, attracting increasing interest in structural biomimetics and tissue engineering [172,173,174,175]. Their porous skeletons, primarily composed of fibrous chitin (Figure 34) or spongin, display hierarchical porosity, mechanical resilience, and long-term stability in aqueous environments with varying salinity levels and temperature ranges between −1.9 °C and 30 °C. Their open porous networks facilitate fluid transport, capillary effects, and mechanical flexibility, while the biological nature of branched, multilayered interconnected fibres allows surface interactions with cells and biomolecules. Unlike synthetic scaffolds, sponge-derived structures are formed through biological self-assembly, inherently incorporating multiscale architectural features that are challenging to replicate with conventional fabrication methods.

Figure 34.

Digital microscopy image of the typical 3D chitinous scaffold isolated from Aplysina sp. marine verongiid sponge. Despite their external structural similarity to spongin fibres, these chitinous fibres represent microtubules with nanoporous walls.

Both chitin- and spongin-based scaffolds derived from keratosan demosponges have been investigated as renewable biomaterials for various bioinspired applications, including tissue engineering. Successful applications of chitin scaffolds from sponges are shown in Figure 35.

At the same time, publications related to the use of spongin in tissue engineering are not very numerous to date. One previous study conducted on Spongia marine bath sponges showed that spongin in the form of 3D scaffolds promotes the growth of human osteoprogenitor cells and also confirms the formation of bone matrix [172].

In addition, a study conducted by Lin et al. evaluated the potential of Callyspongiidae marine sponge as a bioscaffold for bone engineering. Mouse osteoblasts cultured on such 3D scaffolds showed strong adhesion capacity, confirmed by F-actin staining. Alkaline phosphatase (ALP) analysis, von Kossa staining, and PCR demonstrated continued osteoblastic differentiation and the formation of corresponding mineralized nodules for up to 21 days. Osteocalcin and osteopontin expression also confirmed osteogenic activity [102]. Additionally, spongin from Hymeniacidon sinapium marine demosponge effectively stimulates bone mineralization in MG-63 osteoblastic cells [176]. It was reported that this spongin increases ALP activity, collagen synthesis, and osteocalcin secretion. Furthermore, it was shown that spongin also has an anti-inflammatory effect by inhibiting the production of inflammatory mediators such as TNF-α, IL-1β, and PGE₂ in RAW264.7 macrophages [176].

Figure 35.

An overview of application fields for 3D chitinous scaffolds isolated from diverse verongiid demosponges in tissue engineering [64,177,178,179,180,181,182,183,184].

For the first time, we present below the results of our work on isolating flat chitinous 3D matrices with a unique square architecture from I. basta keratosan sponge [184] using the distilled water extraction method. We also demonstrated the use of selected 3D spongin scaffolds for cultivating human MG-63 osteoblastic cells, one of the most popular cell lines in osteogenesis studies [185].

3.10. Gentle Method for Isolation of 3D Flat Chitinous Scaffolds from Ianthella basta Demosponge

Obtaining water extracts from marine sponges as renewable sources of various bioactive compounds and biological materials (such as chitin or spongin) is part of a green chemistry strategy. Unfortunately, this approach is rarely used due to the traditional use of organic solvents such as methanol. Aqueous extracts from sponges have been described in only a few studies [70,186,187,188,189].

For the first time, in this study, the I. basta sponge skeleton was decellularized using distilled water (Figure 36) and purified through alternating acid and alkali treatments to remove calcium carbonates, lipids, proteins, and pigments while maintaining the native flat 3D structure of the sponge skeleton. After removing all associated bromotyrosines, a colorless, purified chitin matrix was obtained, consisting of regularly bridged, pipe-like, translucent fibres with diameters up to 200 µm, which form characteristic chamber-like structural units (see [39,44,189,190]).

Figure 36.

Dried skeleton of I. basta marine demosponge before (A) and after immersion in distilled water (B). Decellularized flat skeletal scaffold (D) has been cut into fragments (C) for further isolation of pure chitin matrices (E).

Scanning electron microscopy (SEM) micrographs (Figure 37A,B) offer a closer look at the morphology of chitin fibres. They show a characteristic wrinkled, deeply fissured surface, a feature commonly observed in chitinous scaffolds of verongiids in previous studies [43,44,191,192].

Figure 37.

SEM micrographs of I. basta chitinous scaffold at different magnifications (A,B). (C,D) FTIR spectra of the obtained chitin scaffold compared with α-chitin and β-chitin standards in the ranges 4000–400 cm⁻¹ (C), and 1800–400 cm⁻¹ (D).

Infrared spectroscopy is a well-established method for characterizing polysaccharides, including chitin [186]. Our ATR-FTIR study (Figure 37C,D) revealed characteristic spectral features of chitin isolated from the I. basta sponge skeleton, compared to α- and β-chitin standards. All spectra showed amide bands corresponding to CONH group vibrations. The amide I band was split into two components at ca. 1630 and 1650 cm⁻¹, indicating hydrogen bonding effects [189,190,192,193]. The amide II band appeared as a narrow peak around 1550 cm⁻¹ in α-chitin and I. basta chitin, but was broader and shifted to 1539 cm⁻¹ in β-chitin. Amide III vibrations were observed between 1308 and 1203 cm⁻¹, while the band at 1375 cm⁻¹ corresponded to methyl group rocking. Strong C–O–C and C–O stretching bands were detected at 1154, 1112, and 1064 cm⁻¹ in all three samples. Additionally, β-chitin showed a band at 1032 cm⁻¹, whereas α-chitin and I. basta chitin displayed band splitting at 1023 and 1008 cm⁻¹, and at 1025 and 1008 cm⁻¹, respectively. The band of ß-glycosidic bond occurs at 897 and 898 cm⁻¹ in α-chitin and I. basta chitin, but in β-chitin, it shifts to 904 cm⁻¹. The FTIR results confirm that chitin isolated from the I. basta demosponge skeleton closely resembles α-chitin rather than β-chitin, as previously reported by Brunner and colleagues [189,190].

The chitin matrix from the I. basta demosponge skeleton represents a promising, highly biocompatible biomaterial for stem cell-based tissue-engineering applications [178,192,193], and it also has potential as a sustainable, innovative biomaterial for controlled drug release and wound dressing applications [43]. However, in this work, for the first time, the skeleton of the I. basta sponge was immersed in distilled water to produce an aqueous extract, which was then thoroughly analyzed for the isolation of biologically active compounds, including bromotyrosines.

For the first time, analytical investigations of the water extract of I. basta sponge skeleton help to identify the following bioactive bromotyrosines: bastadin-6 [193,194,195], bastadin-19 [195,196], 2-(3,5-dibromo-4-methoxyphenyl)acetic acid [196,197,198] and (Z)-3-(4-methoxy-3,5-dimethylphenyl)-2-(sulfooxy)acrylic acid [196,197].

3.11. Natural and Graphitized 3D Spongin Scaffolds for Tissue Engineering with Human MG-63 Osteoblastic Cells

3.11.1. Cytocompatibility of Natural Spongin Scaffolds Assessed by Optical Microscopy

The initial cytocompatibility of natural spongin scaffolds derived from S. lamella, S. tampa, and H. communis bath sponges was assessed using optical microscopy after one day after 1 day of MG-63 human osteoblast-like cell culture (Figure 38). In both cases, cells adhered and spread on the tissue culture polystyrene surface surrounding the sponge fragments. No signs of acute cytotoxicity, such as cell detachment, rounding, or decreased cell density, were observed near either scaffold type.

Figure 38.

Optical microscopy images of MG-63 cells cultured in the presence of natural spongin scaffolds derived from S. tampa, S. lamella and H. communis bath sponges after 1 day and 7 days. Scale bars: 1 mm and 200 µm.

The presence of well-adhered and morphologically normal cells indicates that neither S. lamella nor H. communis spongin scaffolds release toxic components affecting early cell attachment. At this time point, no qualitative differences in cellular response between the two sponge species were detected, suggesting comparable short-term cytocompatibility of both natural spongin materials.

3.11.2. Cell Viability and Infiltration Within Natural Spongin Scaffolds Evaluated by Live/Dead Staining

Cell viability and spatial distribution within natural spongin scaffolds were further evaluated using Live/Dead (FDA/PI) staining and confocal microscopy after 3 and 7 days of culture (Figure 39). Live cells (green fluorescence) were observed near and within the spongin structures for both sponge species, confirming that cells could migrate into the three-dimensional scaffold architecture.

Figure 39.

MG-63 cells cultured on natural spongin scaffolds derived from such bath sponges as S. tampa, S. lamella and H. communis after 3 and 7 days. Live/Dead (FDA/PI) confocal microscopy images. Live cells are shown in green. Scale bars 100 µm.

After 7 days, a higher number of viable cells was observed, indicating ongoing cell survival and proliferation. No accumulation of dead cells was detected, further confirming no evidence of acute cytotoxic responses under the applied experimental conditions. These results demonstrate that natural spongin scaffolds not only support cell viability but also enable gradual cellular infiltration into their porous structure. Live/Dead staining was used here as a qualitative assessment of cell survival and spatial distribution rather than as a quantitative viability assay.

3.11.3. Metabolic Activity of Cells Cultured with Natural Spongin Scaffolds

The metabolic activity of MG-63 cells cultured with natural spongin scaffolds was measured using the Alamar Blue assay after 3 and 7 days (Figure 40). At day 3, all natural spongin scaffolds supported cellular metabolic activity, indicating preserved cellular metabolic activity during early adaptation to the 3D scaffold environment. Although metabolic activity values were lower than those of the control, statistical analysis showed no significant differences for most sponge-derived conditions after adjusting for multiple comparisons. This suggests that early metabolic activity remains intact in the presence of natural spongin materials. By day 7, metabolic activity increased for all sponge-derived scaffolds, indicating sustained cell viability and adaptation to the scaffold environment. H. communis showed the most significant increase in metabolic activity between day 3 and day 7, suggesting progressive cell adaptation rather than improved early cell infiltration. These findings confirm that natural spongin scaffolds from both sponge species are cytocompatible and support cellular metabolic activity in a species-dependent manner, with H. communis inducing a stronger, time-dependent increase in metabolic activity. Notably, the significant rise in metabolic activity for H. communis at day 7 did not directly correlate with homogeneous three-dimensional cell infiltration, suggesting that metabolic measurements alone might overestimate scaffold performance in highly heterogeneous architectures. Importantly, reduced metabolic activity at early time points should not be interpreted as cytotoxicity, but rather as a characteristic feature of osteoblast adaptation to highly porous 3D architectures, where cell attachment, migration, and cytoskeletal reorganization precede full metabolic upregulation.

Figure 40.

Alamar Blue assay showing metabolic activity of MG-63 cells cultured with natural spongin scaffolds derived from such bath sponges as S. tampa, S. lamella, and H. communis, and after 3 and 7 days. Tissue culture polystyrene served as the control. Data are presented as mean ± SD, ** p < 0.01, unpaired two-tailed Student’s t-test.

3.11.4. Cytocompatibility of Carbonized Spongin Scaffolds Assessed by Optical Microscopy and Live/Dead Staining

As previously reported, carbonization of spongin scaffolds at 1200 °C produces 3D porous matrices made of turbostratic graphite, which have found various technological applications [32]. However, to the best of our knowledge, systematic biological studies evaluating these graphitic scaffolds for tissue engineering applications have not been reported.

The cytocompatibility of carbonized spongin scaffolds (1200 °C) derived from S. lamella and H. communis bath sponges was evaluated using optical microscopy and Live/Dead staining after 1 and 7 days of culture (Figure 41). Optical microscopy images revealed that cells remained adherent and appeared morphologically normal on the tissue culture polystyrene surface surrounding the carbonized scaffolds at both time points. No morphological signs consistent with acute cytotoxic responses were observed, indicating that high-temperature carbonization does not introduce toxic effects.

Figure 41.

Optical microscopy images of MG-63 cells cultured in the presence of carbonized (1200 °C) spongin scaffolds derived from S. lamella and H. communis bath sponges after 1 and 7 days. Scale bars: 1 mm and 200 µm.

Live/Dead confocal microscopy further confirmed high cell viability in the presence of carbonized scaffolds, with predominantly live cells detected and only sporadic dead cells observed. These findings demonstrate that carbonization maintains the cytocompatibility of spongin scaffolds from both sponge species. Figure 42 shows cells cultured on natural spongin and carbonized scaffolds for 3 and 7 days. Natural scaffolds supported viable cell attachment and infiltration, however, qualitative differences in cell density and spatial distribution were observed in comparison to carbonized scaffolds. In particular, S. lamella spongin scaffolds after 7 days in culture with osteoblasts displayed a high density of living cells spread throughout the fibrous structure, a more uniform cell distribution, and increased cell migration compared to H. communis spongin scaffolds.

Figure 42.

MG-63 cells cultured on control and carbonized (1200 °C) spongin scaffolds derived from S. lamella and H. communis bath sponges after 1 and 7 days. Live/Dead (FDA/PI) confocal microscopy images. Live cells are shown in green. Scale bars 100 µm.

3.11.5. Immunocytochemical Evaluation of Cell Morphology and Proliferation on Carbonized Spongin Scaffolds

Confocal microscopy images showing MG-63 cells cultured for 7 days on natural and carbonized (1200 °C) spongin scaffolds derived from S. lamella and H. communis. Cell nuclei were stained with DAPI (blue), the actin cytoskeleton was visualized using phalloidin (F-actin, green), and proliferating cells were identified by Ki67 immunostaining (red). Carbonized S. lamella scaffolds maintained favorable cell–material interactions, characterized by elongated cell morphology, aligned actin filaments, and numerous Ki67-positive nuclei, indicating active proliferation. Importantly, these differences were not attributable to cytotoxic effects but rather reflected architecture-dependent modulation of cell behavior. In contrast, carbonized H. communis scaffolds showed reduced cell density, limited cytoskeletal spreading, and fewer proliferating cells, despite overall cell viability shown in Figure 43. These observations confirm that carbonization preserves cytocompatibility but amplifies species-dependent differences, with S. lamella scaffolds providing a more supportive microenvironment for cell adhesion, migration, and proliferation after thermal treatment. These qualitative differences directly reflect the underlying fibre organization and pore interconnectivity of the respective spongin architectures, which become more pronounced after carbonization.

Figure 43.

Immunocytochemical staining of MG-63 cells cultured on selected carbonized spongin scaffolds from S. lamella and H. communis bath sponges origin after 7 days, showing nuclei (DAPI, blue), cytoskeleton (F-actin, green), and proliferating cells (Ki67, red). Scale bars 100 µm.

3.11.6. Effect of Carbonization on Cellular Metabolic Activity

The influence of carbonization on cellular metabolic activity was quantitatively evaluated using the Alamar Blue assay (Figure 44). Statistical significance between selected groups was evaluated using an unpaired two-tailed Student’s t-test (p < 0.05). Pairwise comparisons were performed within each time point to compare natural vs. carbonized scaffolds of the same species and to compare carbonized S. lamella vs. carbonized H. communis.

Figure 44.

Alamar Blue assay comparing metabolic activity of cells cultured on selected natural and carbonized spongin scaffolds under study after 3 and 7 days. Data are presented as mean ± SD, ** p < 0.01, *** p < 0.001, **** p < 0.0001, unpaired two-tailed Student’s t-test.

On day 3, carbonized S. lamella scaffolds showed significantly higher metabolic activity compared to carbonized H. communis scaffolds. No significant differences were observed between carbonized S. lamella and its natural counterpart at this early time point. In contrast, carbonized H. communis exhibited a marked decrease in metabolic activity compared to its natural counterpart and to carbonized S. lamella at day 3. By day 7, carbonized S. lamella scaffolds displayed a significant increase in metabolic activity relative to its natural counterpart and carbonized H. communis scaffolds. Interestingly, the metabolic activity of carbonized S. lamella scaffolds approached that of natural H. communis scaffolds, indicating improved cell–scaffold interactions after carbonization. Conversely, carbonized H. communis scaffolds remained significantly lower compared to carbonized S. lamella and its natural counterpart, despite a time-dependent rise in metabolic activity. Notably, despite reduced early metabolic activity observed for some carbonized scaffolds, Live/Dead staining and immunocytochemical analyses confirmed preserved cell viability, indicating that carbonization did not induce cytotoxic effects but modulated the kinetics of osteoblast metabolic adaptation.

Together, these results (Figure 44) demonstrated that carbonization maintains cytocompatibility but triggers a strong, species-dependent biological response, favoring homogenously structured S. lamella over H. communis in terms of cellular metabolic performance.

Thus, the present study offers a direct comparative evaluation of osteoblast response to natural and carbonized spongin scaffolds derived from S. lamella and H. communis bath sponges, emphasizing the crucial role of species-specific skeletal architecture in influencing cell–material interactions. An important aspect is understanding that the sophisticated polygonal mesh geometry, along with the diversity in mesh opening sizes, remains a key element in providing spatial organization for appropriate nutritional conditions and, consequently, for cell growth within such matrices. Both cell migration and mass transport depend on the initially established microporous architecture of the sponge skeleton, which was optimized over evolution for the sponge’s own cells as a living organism. Therefore, cells foreign to the sponge, including mammalian osteoblasts, face a choice: either reject the proposed scaffold or partially or entirely adapt to it, subsequently actively using it for growth and proliferation.

Both natural spongin scaffolds supported osteoblast viability and metabolic activity, confirming their cytocompatibility. However, clear differences in cellular behavior were observed between sponge species. Natural H. communis scaffolds promoted a pronounced, time-dependent increase in metabolic activity, suggesting progressive cellular adaptation. In contrast, S. lamella scaffolds exhibited a more stable metabolic profile, accompanied by improved cell distribution and infiltration within the scaffold structure. This indicates that higher metabolic activity alone does not necessarily correlate with effective three-dimensional growth colonization.

These differences can be attributed to the intrinsic architectural features of the spongin skeletons. S. lamella skeleton is characterized by a more homogeneous fibre network and interconnected pore structure, which likely facilitates osteoblast migration and spatial organization throughout the scaffold. In contrast, the more heterogeneous architecture of H. communis skeleton may favor surface-associated proliferation rather than deep infiltration.

It could theoretically be expected that carbonized spongin scaffolds would have a beneficial effect on the growth and development of the osteoblastic cells being studied. It has previously been reported that, for example, graphite-containing 3D hydrogels showed high biocompatibility and enhanced L929 fibroblast cell proliferation [198, 199]. Additionally, a tunable scaffold of microtubular graphite has been successfully used for 3D growth of rat embryo fibroblasts [199, 200]. Graphite oxide nanoparticles with a diameter greater than 20 nm have been reported as biocompatible with mouse embryonic stem cells and applicable in various tissue engineering systems [200,201]. Scaffolds of PLA/Graphite nanoplatelet composites created via 3D printing were suitable for ligament tissue engineering [201,202]. Graphite derived from Terminalia arjuna—fruit has been reported as a versatile material for bone tissue engineering applications [202,203]. In principle, all previously studied graphite materials with a graphene layer on their surface are promising for tissue engineering (for overview, see [203,204,205].

Carbonization at 1200 °C maintained the overall 3D structure of both scaffolds but notably enhanced species-specific biological responses. Carbonized S. lamella scaffolds showed improved osteoblast adhesion, cytoskeletal organization, and proliferation, as indicated by well-formed actin networks and numerous Ki67-positive cells. Importantly, carbonization did not hinder early metabolic activity on the S. lamella scaffold, suggesting that its architecture effectively accommodates changes in surface chemistry and stiffness caused by thermal treatment.

The superior biological performance of carbonized S. lamella scaffolds can be mechanistically attributed to the combination of its homogeneous fibre distribution, interconnected pore architecture, and favorable mechanical stability after carbonization. High-temperature treatment likely increases scaffold stiffness and surface roughness while preserving pore continuity, thereby improving focal adhesion formation and cytoskeletal tension in osteoblasts. In contrast, the more heterogeneous architecture of H. communis skeletal scaffolds may undergo local collapse or excessive densification during carbonization, impairing early cell anchorage despite preserved viability. Notably, carbonization transforms the highly elastic native spongin into a mechanically rigid scaffold that can be cut or machined into defined shapes, which is difficult to achieve with the native material due to its pronounced elasticity [32].

Conversely, carbonized H. communis scaffolds showed a marked reduction in early metabolic activity and limited osteoblast proliferation, despite maintaining cytocompatibility. This suggests that carbonization exacerbates architectural constraints inherent to this species, negatively impacting early cell–scaffold interactions. The observed recovery at later time points indicates delayed adaptation rather than an optimal scaffold performance.

Collectively, these findings demonstrate that the biological outcome of carbonization is not universal but strongly depends on the original architecture of spongin-based scaffolds. The S. lamella matrix acts as a scaffold for osteoblast growth in both natural and carbonized forms, highlighting the importance of selecting appropriate biological templates prior to thermal or physicochemical modifications. It cannot be ruled out that the technically unlimited possibilities of cutting fragments of the required shape, porosity, and size from spongin scaffolds will, in the future, enable the use of the patient’s own cells for tissue and organ production, which could then be re-transplanted without rejection [204,205].

4. Conclusions

Marine keratosan demosponges represent an exceptional source of naturally pre-organized and evolutionarily approved 3D skeletal architectures in the form of scaffolds, which have garnered increasing interest in the fields of structural and functional biomimetics as well as tissue engineering. Unlike synthetic 3D scaffolds, sponge-derived structures are created through biological self-assembly, inherently incorporating multi-scale architectural features that are difficult and costly to reproduce with conventional fabrication techniques including 3D printing.

In this experimental review, spongin-based scaffolds derived from keratosan demosponges are examined as renewable biomaterials for various bioinspired applications. Their open porous networks facilitate fluid transport, capillary effects, and mechanical flexibility, while the keratin-collagenous nature of spongin allows surface interactions with diverse biomolecules and cells. It was demonstrated that the highly sophisticated, multilayered, and polygonal mesh-based structural architecture of spongin skeletons in bath sponges, which includes Lissajous-like motifs, enables them to be compressed and torsionally stretched while still absorbing liquids through capillary action. 3D spongin scaffolds effectively absorb 15 different liquids, including human plasma, pork blood, serum (FBS), and DMEM medium used for cell cultivation, highlighting their biomimetic potential for biomedical applications. Importantly, these scaffolds can be further modified via physical or chemical treatments, such as metallization and carbonization, allowing the creation of functional biomaterials while preserving their unique species-dependent hierarchical 3D architecture. For the first time, such advances in metallization of spongin scaffolds—including the creation of spongin–titanium 3D composites using ion–plasma (vacuum arc) deposition, chromium tanning followed by carbonization resulting in crystalline Cr₂O₃ and chromium carbides on the surface of spongin, as well as mineralization with iron oxyhydroxide phases through steam-assisted HCl treatment—have also been reported in this study.

In addition, this work presented remarkable results that illustrate the compression and rehydration cycle of the spongin scaffold. It demonstrates its fascinating shape-memory behavior due to reversible structural recovery, even after two months of exposure to high mechanical pressure at temperatures above 100 °C. Research aiming to understand the principles of the shape-memory phenomenon in spongin matrices is now increasingly in demand.

Despite the significant biomimetic and structural importance of marine bath sponge-derived scaffolds, surprisingly little is known about how species-specific skeletal architectures influence functional osteoblast responses under biologically relevant conditions. In particular, systematic comparisons of osteoblast growth, migration, and metabolic activity on spongin scaffolds from different natural sources—and how these responses are affected by high-temperature carbonization—are still limited. Since carbonization significantly changes surface chemistry and mechanical properties while maintaining native 3D structure, it serves as a powerful yet architecture-dependent modification strategy.

This study demonstrates that both natural and carbonized spongin scaffolds derived from S. lamella and H. communis are cytocompatible and support mammalian cell survival. Optical microscopy, Live/Dead staining, immunocytochemical analysis, and metabolic assays consistently indicated no evidence of acute cytotoxic responses under the applied experimental conditions for all tested materials.

Quantitative and qualitative analyses revealed pronounced species-dependent differences in cellular response. Natural H. communis scaffolds promoted a strong, time-dependent increase in metabolic activity, whereas S. lamella scaffolds displayed a more stable and homogeneous cellular response, which was linked to better cell migration and spatial distribution within the scaffold. These results emphasize the role of the natural skeletal architecture in regulating cell–scaffold interactions.

Carbonization at 1200 °C preserved the 3D structure and cytocompatibility of spongin scaffolds, while also significantly enhancing species-specific biological effects. Although the carbonized S. lamella scaffold displayed enhanced cell migration, proliferation, and metabolic activity, the carbonized H. communis scaffold exhibited decreased early metabolic performance and limited recovery at later time points. These findings suggest that the biological effects of carbonization are primarily determined by the original sponge architecture rather than the thermal process itself.

Overall, this work identifies the carbonized S. lamella skeleton as the most biologically favorable scaffold among the tested biomaterials and emphasizes the crucial role of naturally evolved bath sponge skeletons as renewable templates for bioinspired 3D scaffolds. The presented comparative approach offers a framework for selecting and engineering sponge-derived materials for future applications, especially in structural biomimetics and tissue engineering.

The presented comparative strategy shows that naturally evolved sponge architectures should be considered not merely as passive templates but as active design parameters that dictate the biological results of subsequent physicochemical transformations.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/biomimetics11020124/s1, Table S1. The results of the liquid absorption capacity test performed for spongin scaffolds from demosponge S. lamella; Table S2. Measurement results for analyzed regions (fields) of the carbonized spongin structure presented in Figure 28B, obtained 50 using original software of Keyence VHX-6000 microscopy system; Figure S1. MXRF analysis of the H. communis spongin sample: (A), (B), (C), (D) element mapping of 55 the I, S, Ca, Br showing uniform distribution in spongin matrix and Si aligned with the fibres; (E), 56 (F) element mapping of K and Fe with lines repeating typical spongin fibres microarchitecture.

Author Contributions

Methodology, J.L., A.S. (Anna Szczurek), A.V., R.E.P., I.S. (Igor Smirnov) and C.V.; software, M.K.; validation, M.K.; formal analysis, M.K., A.K., B.L., L.M., H.S., K.H. and C.V.; investigation, J.L., A.S. (Anna Szczurek), A.V., D.P., M.F., R.E.P., I.S. (Igor Smirnov), S.P., I.S. (Ihor Sieliverstov), M.K., A.K., B.L., I.D., L.M., H.S., A.S. (Armin Springer), K.H., A.F., K.N. and M.P.; resources, H.E. and Z.S.; data curation, J.L.; D.P., M.F., I.S. (Igor Smirnov), I.S. (Ihor Sieliverstov) and D.T.; writing—original draft preparation, H.E., A.S. (Anna Szczurek), I.D. and D.T.; writing—review and editing, J.L.; visualization, Z.S., A.S. (Armin Springer), A.F. and M.P.; supervision, H.E., J.L., R.E.P. and S.P.; project administration, H.E.; funding acquisition, H.E. and A.V. All authors have read and agreed to the published version of the manuscript.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

The original contributions presented in this study are included in the article/supplementary material. Further inquiries can be directed to the corresponding authors.

Conflicts of Interest

The authors declare no conflicts of interest.

Funding Statement

This research was funded by the National Science Centre, Poland, under the project MAESTRO 12 No. 2020/38/A/ST5/00151.