Extraction and Characterisation of Chitosan-Based Biomaterials for Prostate Cancer Tissue Regeneration

 Corresponding author: Abiodun Adedoyin ADELEYE. Copyright © 2025 Author(s) retain the copyright of this article. This article is published under the terms of the Creative Commons Attribution License 4.0. Extraction and Characterisation of Chitosan-Based Biomaterials for Prostate Cancer Tissue Regeneration Arinze Daniel OZIEME 1, 2, Abiodun Adedoyin ADELEYE 1, * and Olusegun Olufemi AJIDE 1, 3 1 Department of Biomedical Engineering, University of Ibadan, Nigeria. 2 Department of Chemistry, Covenant University, Ota, Nigeria. ³ Department of Mechanical Engineering, University of Ibadan, Nigeria. Global Journal of Engineering and Technology Advances, 2025, 25(02), 046-062 Publication history: Received on 25 September 2025; revised on 08 November 2025; accepted on 12 November 2025 Article DOI: https://doi.org/10.30574/gjeta.2025.25.2.0328 Abstract In this work, extraction and characterisation of chitosan derived from crab shells was studied as a potential biomaterial for prostate cancer tissue regeneration. Chitosan was obtained through sequential demineralization, deproteinization, and deacetylation, yielding a light-yellow powder with a 23.6±1.2% recovery. Fourier Transform Infrared (FTIR) spectroscopy confirmed successful deacetylation through characteristic amine and hydroxyl absorption bands and the disappearance of acetyl-related peaks. Scanning Electron Microscopy (SEM) micrographs revealed a porous, irregular microstructure (5–20 µm) favourable for cell attachment and nutrient diffusion, while Energy Dispersive Spectroscopy (EDS) verified high elemental purity (C, O, N). X-Ray Diffraction (XRD) analysis indicated semi-crystallinity (CrI = 68.4%), and Thermogravimetric Analysis (TGA) demonstrated good thermal stability with a two-stage degradation profile. Physicochemical assessments revealed a swelling index of 268 ±12 % and a density of 0.42 ± 0.03 g cm⁻³, indicating high hydrophilicity and low compactness suitable for biomedical scaffolds. The Loss on Ignition (LOI) of 97.3% confirmed effective demineralization and high organic purity. The pH of a 1% chitosan solution remained stable (5.8 ± 0.2), demonstrating chemical consistency, while enzymatic degradation with lysozyme confirmed biodegradability under physiological conditions. The foregoing findings imply that crab shell-derived chitosan possesses the structural, morphological, and compositional attributes required for biomedical applications. Its hydrophilic, biocompatible, and thermally stable nature positions it as a sustainable candidate for integration with polymers such as Poly(Lactic-co-Glycolic Acid) (PLGA) in future scaffold fabrication for prostate tissue regeneration. Keywords: Crab shell-derived chitosan; Tissue regeneration; Biodegradability; Hydrophilicity; Demineralization 1. Introduction Prostate cancer is one of the most reported diagnosed malignancies among men and a major contributor to global cancer mortality. According to epidemiological analyses as reported by Sung et al. (2021), there was more than 1.4 million new prostate cancer worldwide in 2020, accounting for approximately 14 % of all cancers in men. The burden is particularly high in sub-Saharan Africa, where late presentation, limited diagnostic facilities, and genetic predisposition contribute to poor survival rates (Ikuerowo et al., 2019; Adeloye et al., 2021). Conventional treatment strategies—including radical prostatectomy, androgen deprivation therapy, chemotherapy, and radiotherapy—are effective for tumour control but frequently cause collateral tissue injury, urinary incontinence, erectile dysfunction, and systemic toxicity (Attard and de Bono, 2016; Resnick et al., 2013). These adverse outcomes highlight the urgent need for regenerative interventions capable of restoring functional tissue following ablative therapy. Global Journal of Engineering and Technology Advances, 2025, 25(02), 046-062 47 Tissue engineering integrates biomaterials, cells, and biochemical signals to reconstruct damaged organs. A key requirement for successful regeneration is the development of a scaffold that mimics the extracellular matrix, promotes cell adhesion and proliferation, and supports angiogenesis (Langer and Vacanti, 2016; O’Brien, 2011). Biopolymers— both natural and synthetic—have been extensively investigated for this role. Natural polymers such as collagen, alginate, and chitosan provide intrinsic biocompatibility and bioactivity, whereas synthetic polymers including PLGA, PCL, and PEG offer mechanical strength, tunable degradation, and processing versatility (Jayakumar et al., 2020; Chen et al., 2021). The combination of these two classes in hybrid composites frequently yields materials with synergistic physicochemical and biological performance (Dash et al., 2011; Li et al., 2020). Chitosan is a partially deacetylated derivative of chitin obtained from crustacean shells. Structurally composed of β- (1→4)-linked D-glucosamine and N-acetyl-D-glucosamine units, it exhibits cationic behaviour in acidic media, facilitating electrostatic interactions with negatively charged cell membranes and biomolecules (Rinaudo, 2006; Younes and Rinaudo, 2015). Its remarkable biocompatibility, biodegradability, hemostatic, and antimicrobial properties have established it as one of the most promising candidates for biomedical and pharmaceutical applications (Khor and Lim, 2003; Mi et al., 2002). Moreover, chitosan’s functional amine and hydroxyl groups allow facile chemical modification and blending with other polymers to tailor mechanical strength, degradation rate, and hydrophilicity (Singla and Chawla, 2001; Kumar et al., 2022). For prostate tissue regeneration, chitosan-based scaffolds can be engineered to provide an optimal microenvironment supporting epithelial and stromal cell growth, while reducing inflammatory responses. Studies have shown that chitosan composites with PLGA or PCL can deliver bioactive molecules, modulate cell behaviour, and enhance neovascularisation in soft-tissue constructs (Chen et al., 2021; Jayakumar et al., 2020). The ability of chitosan to form porous, hydrated matrices further facilitates nutrient transport and waste removal, essential for three-dimensional tissue culture (Kumar et al., 2020). Nigeria’s coastal zones produce large volumes of crab and shrimp waste, which are typically discarded as refuse. Transforming these by-products into biomedical-grade chitosan not only supports environmental sustainability, but also provides an indigenous source of raw material for the biomedical industry (Olatunji et al., 2022). Extraction processes typically involve deproteinisation, demineralisation, decolourisation, and deacetylation. The physicochemical properties of the resulting chitosan—including degree of deacetylation (DD), crystallinity, and molecular weight— govern its solubility, mechanical strength, and bioactivity (Brugnerotto et al., 2001; Farrag et al., 2016). Hence, accurate characterisation using analytical techniques such as FTIR, XRD and SEM-EDS is essential for confirming quality and ensuring reproducibility (Jayakumar et al., 2020). The focus of the present study is on the extraction of chitosan from locally sourced crab shells collected along the Eko Atlantic Seashore, Lagos State, Nigeria, and its subsequent characterisation. The objective was to evaluate the suitability of the synthesised chitosan for potential use in prostate tissue engineering when combined with PLGA. By comparing experimentally obtained physicochemical data—including FTIR, XRD, SEM-EDS, swelling index, and density—with standards. Attempt was made to establish a reference framework for chitosan–PLGA composite development in regenerative medicine. 2. Materials and Methods 2.1. Materials Crab shells (Callinectes sapidus) were sourced from local seafood vendors in Lagos, Nigeria. Analytical-grade reagents such as hydrochloric acid (HCl), sodium hydroxide (NaOH), acetic acid, and ethanol were procured from Sigma-Aldrich (USA). All reagents were utilised as received without any further purification, and deionized water served as the solvent for all experimental procedures. 2.2. Preparation of Crab Shell Powder The harvested crab shells were carefully rinsed with running water to eliminate sand, flesh remnants, and other contaminants. After cleaning, the shells were boiled in water for 30 minutes to detach any remaining organic matter, then oven-dried at 60 °C for 24 hours. The dried samples were subsequently milled using a mechanical grinder and sieved to achieve a fine crab shell powder with a particle size of ≤ 500 µm. The resulting powder was preserved in airtight containers until further use. Global Journal of Engineering and Technology Advances, 2025, 25(02), 046-062 48 2.3. Extraction of Chitosan from Crab Shells The extraction process consisted of three main stages which are demineralization, deproteinization, and deacetylation, conducted using a modified method adapted from previously reported studies. 2.3.1. Demineralization A 50 g portion of crab shell powder was subjected to treatment with 1 M HCl at a solid to liquid ratio of 1:10 (w/v) and stirred at room temperature for 3 hours to eliminate calcium carbonate and other inorganic salts. The resulting mixture was then filtered and rinsed several times with deionized water until a neutral pH was achieved. The obtained demineralized residue was dried at 60°C overnight and weighed to evaluate the mass reduction resulting from mineral removal. 2.3.2. Deproteinization The demineralized crab shells were subjected to treatment with 1 M NaOH solution at 80 °C for 2 hours under continuous stirring at a solid to liquid ratio of 1:10 (w/v) to eliminate proteins and lipids. The resulting mixture was filtered, and the residue was rinsed repeatedly with deionized water until a neutral pH was achieved. The residue was then dried at 60 °C for 24 hours, yielding chitin as the final product from this stage. 2.3.3. Deacetylation The obtained chitin was transformed into chitosan by treatment with 50% (w/v) NaOH solution at 100°C for 3 hours under reflux conditions. Following the reaction, the material was filtered and rinsed repeatedly with deionized water until the filtrate reached a neutral pH. The resulting chitosan was then oven dried at 60 °C for 24 hours and stored in a desiccator for further analysis. 2.4. Fourier Transform Infrared Spectroscopy (FTIR) The FTIR analysis was carried out using a PerkinElmer Spectrum 100 spectrometer within the wave number range of 4000 to 400 cm⁻¹ at a resolution of 4 cm⁻¹. The dried chitosan sample was blended with spectroscopic grade KBr, compressed into a pellet, and scanned. The resulting absorption bands were analysed to identify functional groups and evaluate the degree of deacetylation. 𝐷𝐷 (%)= [ 1 − (𝐀𝟏𝟔𝟓𝟓 𝐀𝟑𝟒𝟓𝟎) ∕ 1.33 ] 𝑋 100 1 2.5. Scanning Electron Microscopy (SEM) The surface morphology of the produced chitosan was analysed using a Zeiss EVO LS10 scanning electron microscope. Prior to imaging, the samples were sputter coated with an approximately 10 nm thick layer of gold to improve conductivity. Imaging was conducted at an accelerating voltage of 15 kV, and the obtained micrographs were evaluated for surface roughness, pore structure, and particle size distribution. 2.6. X-Ray Diffraction (XRD) X-ray diffraction (XRD) patterns were obtained using a Bruker D8 Advance diffractometer equipped with Cu-Kα radiation (λ = 1.5406 Å) operating at 40 kV and 40 mA. The diffraction data were recorded over a 2θ range of 5°–60°. The crystallinity index (CrI) of chitosan was determined using the Segal method as shown in Figure 1 below: Cr𝐼 (%)= 𝑰₁₁₀ − 𝑰ₐₘ 𝑰₁₁₀ 𝑋 100 2 where I₁₁₀ is the intensity of the crystalline peak (around 20°) and Iₐₘ is the intensity of the amorphous region (around 12°). 2.7. Thermogravimetric Analysis (TGA) Thermal stability was studied using a Mettler Toledo TGA analyzer. Approximately 10 mg of chitosan sample was heated from 25 °C to 800 °C at 10 °C/min under a nitrogen flow rate of 50 mL/min. The percentage weight loss was plotted against temperature to identify thermal degradation stages. Global Journal of Engineering and Technology Advances, 2025, 25(02), 046-062 49 2.8. Hydrophilicity (Contact Angle Measurement) The hydrophilicity of the extracted chitosan samples was evaluated using the static water contact angle method. Dried chitosan films were prepared by solvent casting from 1% (w/v) chitosan solution in 1% acetic acid and air-dried at room temperature. A contact angle goniometer (or manual method using a calibrated digital microscope) was used to measure the angle formed between a distilled water droplet (5 µL) and the chitosan surface at room temperature. The contact angle (θ) was recorded within 5 seconds of droplet deposition at five different locations on each film, and the mean value was calculated. A lower contact angle (<90°) indicates higher hydrophilicity, implying better wettability and potential for cell adhesion, while a higher contact angle (>90°) suggests hydrophobic behaviour. The hydrophilicity of the samples was also correlated with the swelling index and surface roughness observed in SEM analysis to establish structure–property relationships. 2.9. Swelling Index Swelling behaviour was analysed by immersing dried chitosan samples (Wd) in phosphate-buffered saline (PBS, pH 7.4) at 37 °C for 24 h. After removal, excess surface liquid was gently blotted, and swollen samples were weighed (Wt). The swelling index (SI %) was calculated using Equation (3): 𝑆𝐼 (%)= 𝑊𝑡− 𝑊0 𝑊0 𝑋 100 3 where W0 is the dry weight and Wt the swollen weight at time t. This test evaluated the water absorption capacity, which reflects the hydrophilicity and potential biological fluid interaction of the extracted chitosan. 2.10. Density Measurement The density (ρ) of the extracted chitosan was determined using a pycnometer (displacement method), as shown in Equation (4): 𝜌= 𝑚 𝑉 4 where m is the sample mass (g) and V is the volume (cm³). The measurement was conducted in triplicate, and the mean values were reported to evaluate structural compactness and material consistency. 2.11. Loss on Ignition (LOI) Loss on ignition was determined to evaluate the organic matter content and purity of the chitosan. Approximately 2 g of dried chitosan was weighed (W₁) and heated in a muffle furnace at 750 °C for 3 hours. After cooling in a desiccator, the residue was weighed again (W₂). LOI was calculated as using equation 5 below: LOI (%)=𝑊1−𝑊2 𝑊1 𝑋100 5 Where: where W₁ is the initial weight and W₂ the post-ignition weight. A high LOI indicates significant organic purity and successful removal of mineral residues. 2.12. Enzymatic Degradation of Chitosan Powder Chitosan powder (initial dry weight, Wo) was subjected to enzymatic degradation using lysozyme to simulate physiological breakdown conditions. Approximately 100 mg of the sample was suspended in phosphate-buffered saline (PBS, pH 7.4) containing 1.5 mg/mL lysozyme and incubated at 37 °C under gentle shaking. At predetermined time intervals (1, 7, 14, 21, and 30 days), aliquots were withdrawn to assess degradation. After incubation, the chitosan powders were carefully filtered and rinsed with distilled water to eliminate residual enzyme and buffer salts. The samples were subsequently dried in an oven at 50°C until a constant weight (Wt) was Global Journal of Engineering and Technology Advances, 2025, 25(02), 046-062 50 achieved. The percentage weight loss, which indicates the extent of enzymatic degradation, was calculated according to Equation (6): Degradation (%)=Wo−𝑊𝑡 Wo 𝑋 100 6 where Wo is the initial dry weight and Wt is the final dry weight after degradation. This method provides an accurate estimation of the biodegradability of chitosan under enzyme-rich physiological conditions. Lysozyme, an enzyme naturally found in human saliva and tears, effectively hydrolyses the β-(1→4)- glycosidic linkages between N-acetylglucosamine units in chitosan. The rate and extent of degradation are influenced by several parameters, including the degree of deacetylation, molecular weight, crystallinity, and porosity of the material. High deacetylation levels and amorphous structures typically enhance lysozyme access and degradation kinetics. 2.13. Statistical Analysis All experiments were performed in triplicate (n = 3), and the results were expressed as mean ± standard deviation (SD). Statistical analysis was carried out using one way analysis of variance (ANOVA), with a significance level set at p < 0.05. 3. Results 3.1. Visual and Physical Observations The extraction process yielded a light-yellow to off-white chitosan powder with a smooth texture and mild odour, indicating successful deproteinization and deacetylation. The yield of chitosan obtained was approximately 23.6±1.2% of the original crab shell mass, consistent with reports by Ogunlana et al. (2022) and Rahman et al. (2020), who reported yields in the range of 20–30% depending on shell source and processing conditions. The reduction in colour intensity after alkali treatment confirmed the effective removal of pigments and organic residues. The extracted chitosan was insoluble in water and organic solvents but dissolved completely in 1% acetic acid solution, producing a clear and viscous gel — a key indicator of high-quality chitosan suitable for biomedical applications such as wound dressing, tissue regeneration, and drug delivery. 3.2. Fourier Transform Infrared Spectroscopy (FTIR) The FTIR spectrum of the extracted chitosan displayed characteristic absorption bands corresponding to amine and hydroxyl functional groups as shown in Table 1 and depicted in Figure 1. The characteristic absorption bands confirmed the structural transformation of chitin to chitosan. A broad absorption band observed around 3420–3450 cm⁻¹ corresponds to the O–H and N–H stretching vibrations, indicating the presence of hydroxyl and amino groups, respectively. This broadness reflects strong intermolecular hydrogen bonding within the chitosan matrix. The band near 2920–2870 cm⁻¹ is attributed to C–H stretching vibrations of aliphatic –CH₂ and –CH₃ groups in the glucosamine backbone. A distinct band at 1650–1630 cm⁻¹ represents the amide I (C=O stretching) of the remaining N-acetyl groups, confirming that partial deacetylation occurred during processing. The amide II band around 1590–1560 cm⁻¹ is due to N–H bending, indicating the presence of free amino groups that result from successful deacetylation of chitin. A medium-intensity band between 1420–1380 cm⁻¹ corresponds to CH₂ bending and C–H deformation, characteristic of polysaccharide structures. The strong and broad peaks in the region 1150–1020 cm⁻¹ are assigned to C–O–C stretching vibrations, which confirm the polysaccharide nature and glycosidic linkages of chitosan. Furthermore, a distinct peak near 895–720 cm⁻¹ corresponds to β-1,4-glycosidic linkages, validating the polymeric backbone of Dglucosamine units typical of chitosan. Global Journal of Engineering and Technology Advances, 2025, 25(02), 046-062 51 Table 1 FTIR Interpretation of Crab Shell–Derived Chitosan Wave number (cm⁻¹) Functional Groups Interpretation 3420–3450 (broad, weak) O–H and N–H stretching vibrations Indicates hydroxyl and amino groups — characteristic of chitosan’s polysaccharide backbone and deacetylated structure. 2920–2870 C–H stretching (–CH₂, – CH₃) Associated with aliphatic groups in the glucosamine unit. 1650–1630 Amide I (C=O stretching of –NHCOCH₃) Confirms partial deacetylation; represents remaining acetyl groups. 1590–1560 Amide II (N–H bending) Suggests presence of primary amine groups from deacetylated chitosan. 1420–1380 CH₂ bending / C–H deformation Typical of polysaccharide structures. 1150–1020 C–O–C stretching (glycosidic linkage) Strong band representing the saccharide structure of chitosan. 897–720 β-1,4-glycosidic linkage vibration Confirms polymer backbone connectivity between D-glucosamine units. These observed functional group vibrations align with previously reported spectra of pure chitosan (Dash et al., 2011; Kumar, 2000), confirming the effective removal of acetyl groups and the successful conversion of crab shell chitin to chitosan. The presence of free amino groups and hydroxyl functionalities enhances hydrophilicity and provides reactive sites for further modification or composite formation, supporting the potential use of this chitosan in biomedical scaffold applications, including prostate tissue regeneration. The absence of a strong peak near 1730 cm⁻¹ (characteristic of acetyl groups in chitin) confirmed the effective removal of acetyl moieties and successful deacetylation to chitosan. These observations align with previous findings reported by Adekoya et al. (2021) and Shanmugam et al. (2020), suggesting a high degree of deacetylation suitable for biomedical material formulation. Figure 1 FTIR spectra of Crab Shell–Derived Chitosan Global Journal of Engineering and Technology Advances, 2025, 25(02), 046-062 52 3.3. Scanning Electron Microscopy (SEM) The Scanning Electron Microscopy (SEM) micrograph of the synthesized chitosan from crab shells reveals a rough, porous, and flake-like surface morphology characteristic of partially deacetylated chitin structures. The surface exhibits irregular cavities, cracks, and granular nodules distributed across the matrix, indicating efficient removal of calcium carbonate and protein residues during the demineralization and deproteinization stages. At a magnification of 500× (Figure 2), the image shows micropores ranging between 1–10 µm in diameter. Such porosity is beneficial for biomedical applications, particularly in tissue engineering scaffolds, as it promotes cell adhesion, nutrient diffusion, and metabolic exchange. The interconnected pore channels observed suggest a high surface area that may enhance drug loading or biomolecule immobilization when used as a composite material. The presence of spherical agglomerates on the surface may be attributed to partial aggregation of polymer chains during drying, which is commonly observed in chitosan derived from natural sources (Dash et al., 2011). Additionally, the cracks and fissures visible across the surface could result from moisture loss and internal stress developed during solvent evaporation, indicating a brittle microstructure typical of purified chitosan films. These morphological features confirm the successful conversion of chitin to chitosan and the formation of a biocompatible porous structure, suitable for further blending with polymers such as PLGA or hydroxyapatite in the fabrication of biomedical scaffolds for prostate tissue regeneration. Figure 2 Morphological Image of the Synthesised Chitosan 3.4. Elemental Composition of the Extracted Chitosan The EDS spectrum showed dominant peaks corresponding to carbon (C), oxygen (O), and nitrogen (N), confirming the organic composition of chitosan. Minor traces of calcium (Ca) were also detected, indicating minimal residual mineral content post-demineralization. Elemental quantification revealed C (45.8%), O (38.3%), N (15.1%), and Ca (0.8%), consistent with high-purity chitosan as shown in Table 1. The C: N ratio (≈3:1) corroborated successful deacetylation and removal of inorganic matter. These morphological and compositional features indicate that the extracted chitosan possesses a favorable microarchitecture for scaffold fabrication and potential prostate tissue regeneration applications. Global Journal of Engineering and Technology Advances, 2025, 25(02), 046-062 53 Table 2 EDS Elemental Composition of Crab Shell–Derived Chitosan Element Symbol Weight (%) Atomic (%) Carbon C 45.8 48.2 Oxygen O 38.3 40.5 Nitrogen N 15.1 10.9 Calcium Ca 0.8 0.4 3.5. X-Ray Diffraction (XRD) The X-ray diffraction (XRD) spectrum (Figure 3) of the crab shell–derived chitosan displays several characteristic peaks between 2θ = 10° and 60°, indicating a semi-crystalline structure. Prominent diffraction peaks were observed at approximately 2θ = 10.1°, 19.8°, 26.5°, and 32.8°, corresponding to SiO (Q), Silica-Aluminate (SA), Ferrite (Fe), and Calcite (C) phases, respectively as shown in Table 3. The intense peaks at 2θ ≈ 10° and 20° are typical of partially deacetylated chitosan, confirming the presence of both amorphous and crystalline domains. These peaks represent the regular arrangement of polysaccharide chains through intermolecular hydrogen bonding between the amino and hydroxyl groups. The broadness of the peaks indicates that the chitosan retains an amorphous polymeric structure, which enhances its solubility and reactivity for biomedical modification. Figure 3 X-ray diffraction (XRD) spectrum of the synthesised chitosan Table 3 XRD Peak Positions and Corresponding Phases of Crab Shell–Derived Chitosan 2θ (°) Phase Identified Compound/Structure Relative Intensity (%) Remarks 10.1 Q Silicon Oxide (SiO) 25 Typical amorphous chitosan peak 14.8 SA Silica-Aluminate 18 Minor impurity phase 19.8 C Calcite (CaCO₃) 100 Major crystalline peak of chitosan 22.4 Q Silicon Oxide (SiO) 38 Semi-crystalline ordering 26.5 SA Silica-Aluminate 30 Residual inorganic trace 29.6 Fe Ferrite (Fe₂O₃) 22 Trace metal oxide content Global Journal of Engineering and Technology Advances, 2025, 25(02), 046-062 54 32.8 C Calcite (CaCO₃) 45 Minor residual mineral peak 38.1 Q Silicon Oxide (SiO) 20 Secondary broad chitosan halo 47.9 Fe Ferrite (Fe₂O₃) 17 Low intensity metal oxide trace 50.3 SA Silica-Aluminate 15 Minor amorphous peak The minor diffraction signals associated with calcite (CaCO₃) suggest trace amounts of residual mineral content from the crab shell matrix, consistent with EDS data that showed less than 1% calcium content. The appearance of silicaaluminate (SA) and ferrite (Fe) peaks may arise from environmental impurities or trace inorganic residues inherent in the natural crab shell source. Furthermore, the XRD pattern confirms successful deacetylation and purification of the extracted chitosan, with a predominant amorphous phase ideal for scaffold fabrication and biopolymer blending. The amorphous structure is advantageous in biomedical applications, as it allows enhanced swelling, flexibility, and interaction with biological tissues, all of which are essential for prostate tissue regeneration scaffolds. Also, the crystallinity index (CrI) was calculated to be 68.4%, indicating partial ordering due to hydrogen bonding among polymer chains. This moderate crystallinity is desirable for biomedical materials as it offers a balance between mechanical stability and flexibility. In comparison, highly crystalline chitosan tends to be brittle and less soluble, whereas low crystallinity may reduce structural integrity. The obtained results are consistent with the findings of Chen et al. (2019), who reported CrI values between 60% and 75% for chitosan prepared from crab and shrimp shells. Table 4 Crystallinity and Structural Properties of Extracted Chitosan Parameter Observed Value Literature Range Remarks Crystallinity Index (CrI, %) 68.4 60–75 Semi-crystalline Major Diffraction Peaks (°2θ) 10°, 20° 9–21 Consistent with chitosan structure Structural Nature Semi-crystalline - Suitable for biomedical use 3.6. Thermogravimetric Analysis (TGA) The TGA thermogram showed two major weight-loss stages. The first stage (below 120 °C) corresponded to the loss of adsorbed water (~8.3% weight loss). The second major stage occurred between 250 °C and 380 °C, corresponding to the degradation of chitosan polymer chains (48.7% weight loss). The residual char at 800 °C was approximately 21%, indicating good thermal stability. This decomposition profile is consistent with chitosan derived from other marine sources (Rahman et al., 2021; Patel et al., 2020). The relatively high decomposition temperature suggests potential for moderate-temperature processing; such as scaffold fabrication or film casting. Table 5 Thermal Decomposition Characteristics of Chitosan Degradation Stage Temperature Range (°C) Weight Loss (%) Interpretation I 30–120 8.3 Evaporation of bound water II 250–380 48.7 Decomposition of polymer backbone III 380–800 21.0 (residue) Formation of carbonaceous residue 3.7. Loss on Ignition (LOI) The LOI value for the extracted chitosan was 97.3%, indicating high organic matter content and minimal inorganic residue. This result corroborates EDS findings, confirming effective demineralization during extraction. 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