Strategic Advances in Efficient Chitin Extraction from Black Soldier Fly Puparia: Uncovering the Potential for Direct Chitosan Production

Abstract

As a biodegradable material, chitin possesses exceptional physical and chemical properties, making it valuable in various industrial sectors. Compared to chitin, its derivative, chitosan, offers even more versatile applications due to its higher solubility and reactivity. As the key precursor for chitosan production, chitin is typically harvested from shrimp and crab exoskeletons. However, the quest for alternative sources has intensified to reduce reliance on crustacean-derived chitin. Black soldier fly (BSF, Hermetia illucens), particularly its puparium, has emerged as a promising alternative, though it is typically considered waste. In this study, we investigated different combinations and sequences of extraction treatments for chitin from the BSF puparium. The results demonstrate that sequential treatments of demineralization (DM), deproteination (DP), and decolorization (DC) produce chitin with the lowest ash, lipid, and protein contents—approximately 1.01%, 1.99%, and 3.01%, respectively, yielding degrees of DM and DP at 94.77% and 92.24%, and a chitin purity of 93.98%. In contrast, combining DP and DC following the DM treatment facilitates partial chitosan transformation with a degree of deacetylation (DD) of 65.90%, offering a direct alternative for producing chitosan without an additional deacetylation process.

1. Introduction

Chitin is a naturally occurring polysaccharide comprising a linear homopolymer of N-acetylglucosamine (GlcNAc) units linked by β(1,4) glycosidic bonds [1]. It is predominantly found in the exoskeletons of arthropods such as shrimp, crabs, and spiders, as well as in the cell walls of fungi [2]. In parallel, chitosan, a derivative of chitin, retains similar structural features but differs due to the partial removal of acetyl groups through deacetylation [3,4]. This deacetylation process produces a copolymer with significantly improved solubility in acidic solutions and distinct functional properties compared to chitin [5,6].

As a result, chitosan exhibits enhanced versatility in various applications, including tissue engineering, bioactive encapsulation, food preservation, and packaging [7,8,9]. Given its significance, the sustainability of chitin as the template platform plays a crucial role in both the current advancements and future progress in the extraction and functionalization of chitosan. However, with the projected growth of the global chitin and chitosan market size from approximately USD 1.96 billion in 2024 to USD 4.07 billion by 2030, driven by a compound annual growth rate (CAGR) of 12.8% [10], exploring alternative sources for chitin production to alleviate environmental pressures is essential. In this context, chitin extraction from insects emerges as a compelling solution. Insects, representing approximately 95% of the animal kingdom’s species, can contain up to 60% chitin in their exoskeletons, comparable to or even higher than the chitin content found in marine sources (~40%) [11,12].

Among insect species, the black soldier fly (BSF, Hermetia illucens) stands out as an up-and-coming candidate. The BSF undergoes a complete metamorphosis consisting of five larval stages, followed by pre-pupal and pupal stages before the emergence of the adult fly [13]. Chitin can be extracted from all developmental stages; however, the puparia are especially advantageous due to their higher content (~23%), despite being typically considered waste [14,15]. Furthermore, the BSF is distinguished by its rapid growth cycle of just a few days and high efficiency in converting organic waste into biomass [16]. This rapid conversion process contributes significantly to waste reduction and sustainable resource management, making BSF a valuable resource for chitin extraction.

Biological or chemical processes are generally employed for chitin extraction, though chemical methods are most used due to their efficiency [17]. These chemical processes involve deproteinization and demineralization using strong bases (KOH or NaOH) and acids (HCl or H₂SO₄) [18]. Additionally, decolorization is often performed to purify the chitin further by removing residual pigments, using oxidizing agents such as H₂O₂ or NaClO [16]. While previous studies have explored chitin extraction from BSF [19,20,21], to the best of our knowledge, an investigation into the effects of different treatment combinations and sequences, particularly involving BSF puparium, has been limited.

In our pursuit of an efficient chitin extraction process to meet the growing demand for this biopolymer, we present an alternative approach for isolating chitin and chitosan from the BSF puparium by refining various combinations and sequences of deproteination (DP), demineralization (DM), and decolorization (DC) treatments. Despite the relatively high purity of chitin (~93.98%) obtained from the treatment sequences of DM, DP, and DC—resulting in the lowest ash, lipid, and protein contents (approximately 1.01%, 1.99%, and 3.01%, respectively), we found that the simultaneous combination of DP and DC following the DM treatment exhibits the highest degree of deacetylation (DD) at 65.90%. This result provides an alternative approach to directly produce chitosan with a higher DD by eliminating the conventional deacetylation process, paving the way for efficient and cost-effective large-scale production in the future.

2. Materials and Methods

2.1. Insect Material and Chemicals

Puparia of the black soldier fly (BSF) were collected from a local farmer in Cimahi, West Java Province, Indonesia, after being cleaned and having the remaining molten pupa removed. All reagents used in the experiments were of technical grade and purchased from a local supplier unless otherwise specified. Hydrochloric acid (HCl), sodium hydroxide (NaOH), ethanol (C₂H₅OH, 96% v/v), n-hexane (C₆H₁₄), hydrogen peroxide (H₂O₂, 50% w/w), copper (II) sulfate pentahydrate (CuSO₄·5H₂O, ≥98%, Sigma-Aldrich, St. Louis, MO, USA), potassium sulfate (K₂SO₄, ≥99%, Sigma-Aldrich, St. Louis, MO, USA), sulfuric acid (H₂SO₄, 95–98%, Sigma-Aldrich, St. Louis, MO, USA), boric acid (H₃BO₃, ≥99.5%, Sigma-Aldrich, St. Louis, MO, USA), bromocresol green (BCG, ACS, Sigma-Aldrich, St. Louis, MO, USA), methyl red (C.I. 13020, ACS, Sigma-Aldrich, St. Louis, MO, USA), acetic acid (CH₃COOH, ≥99.7%, Sigma-Aldrich, St. Louis, MO, USA), and sodium acetate (CH₃COONa, ≥99%, Sigma-Aldrich, St. Louis, MO, USA) were used as received, without any further purification. Distilled water was used to prepare all reagent solutions and treatment procedures.

2.2. Chitin Extraction from BSF Puparium

The puparium of BSF, hereinafter referred to as puparium-BSF, was pretreated by drying in a convection oven (UNB-400, Memmert, Schwabach, Germany) at 378 K for 24 h before being ground and passed through a 60-mesh sieve to achieve a uniform particle size. Chitin (CT) extraction involves deproteination (DP), demineralization (DM), and decolorization (DC), with each step performed in various sequences to optimize extraction outcomes, such as efficiency and material quality. For clarity, the detailed methods are further outlined in

Table 1. Variants of the extraction sequences and parameters used in this study. The variant numbers listed are referenced throughout the text.

Variant	Treatment Sequence	Demineralization	Deproteination	Decolorization
CT-1	DP → DM	1.0 mol·L−1 of HCl at Tamb for 1 h.	1.0 mol·L−1 of NaOH at 353 K for 1 h.	-
CT-2	DM → DP			-
CT-3	DP → DM → DC			5% H2O2 at 363 K for 1 h.
CT-4	DM → DP → DC
CT-5	DP → DC → DM
CT-6	(DP + DC) → DM		5% H2O2 and 1.0 mol·L−1 of NaOH at 353 K for 1 h.
CT-7	DM → (DP + DC)

.

The DP step was initiated by mixing a calculated amount of puparium-BSF powder with 1.0 mol·L⁻¹ of NaOH solution at a 1:25 solid-to-liquid (m/v) ratio in a 500 mL conical flask. The mixture was stirred with a magnetic bar under moderate agitation and heated to 353 K for 1 h using a hot plate (Yellow Line, Mag HS7). In the DM step, a 1.0 mol·L⁻¹ HCl solution was used as a demineralization agent, mixed with the puparium-BSF powder at a 1:15 ratio at an ambient temperature for 1 h. This was followed by rinsing with distilled water and hot ethanol at the same ratio at least three times. For the DC step, the puparium-BSF powder was treated with a 5% H₂O₂ solution at a 1:20 ratio and heated to 363 K for 1 h. The degree of deproteination (DDP) and demineralization (DDM) were determined by comparing the protein and ash content before and after treatment, following Equations (1) and (2), where Cp₀, Ca₀, Cp₁, and Ca₁ represent the initial protein and ash contents and the contents after treatment, respectively.

$$\mathrm{D}\mathrm{e}\mathrm{g}\mathrm{r}\mathrm{e}\mathrm{e} \mathrm{o}\mathrm{f} \mathrm{D}\mathrm{e}\mathrm{p}\mathrm{r}\mathrm{o}\mathrm{t}\mathrm{e}\mathrm{i}\mathrm{n}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n}\left(\mathrm{\%}\right)={\displaystyle \frac{{\mathrm{C}\mathrm{p}}_0-{\mathrm{C}\mathrm{p}}_1}{{\mathrm{C}\mathrm{p}}_0}}\times 100$$

(1)

$$\mathrm{D}\mathrm{e}\mathrm{g}\mathrm{r}\mathrm{e}\mathrm{e} \mathrm{o}\mathrm{f} \mathrm{D}\mathrm{e}\mathrm{m}\mathrm{i}\mathrm{n}\mathrm{e}\mathrm{r}\mathrm{a}\mathrm{l}\mathrm{i}\mathrm{z}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n}\left(\mathrm{\%}\right)={\displaystyle \frac{{\mathrm{C}\mathrm{a}}_0-{\mathrm{C}\mathrm{a}}_0}{{\mathrm{C}\mathrm{a}}_0}}\times 100$$

(2)

A modified simultaneous DP and DC step was also conducted for comparison. In this approach, a 5% H₂O₂ solution at a 1:20 ratio was added dropwise to a mixture of puparium-BSF powder and 1.0 mol·L⁻¹ of NaOH under moderate stirring. The hot suspension was further heated to 353 K for 1 h. Following each step (DP, DM, DC, and simultaneous DP-DC), the suspensions were subjected to filtration and ultrasonication-assisted redispersion cycles with distilled water multiple times until a neutral pH was achieved. Finally, the resultant solid was dried at 353 K for 24 h to yield chitin and stored in a desiccator to prevent moisture uptake and maintain sample quality before further analysis.

2.3. Material Characterization

The proximate analysis of chitin, including moisture, ash, crude lipid, and protein content, was conducted using standard analytical procedures with minor modifications. Moisture and ash content were measured according to ISO 638-1:2022 [22] and ISO 1762:2019 [23]. Samples were initially heated in a drying oven at 378 K until a constant weight was achieved and then subjected to incineration in a muffle furnace (Vulcan D-550, NeyTech, Mifflinburg, PA, USA) at 823 K for 2 h. Crude lipid content was measured using a Soxhlet extractor with n-hexane as the solvent, following AOAC 2003.05 [24]. The moisture, ash, and crude lipid content were quantified as weight differences before and after the treatment.

The nitrogen content was estimated using the Kjeldahl method, following ISO 1871:2009 [25]. Briefly, 1 g of the samples was digested with 12 mL of concentrated H₂SO₄ and a mixture of 0.8 g K₂SO₄ and 7 g CuSO₄·5H₂O as a catalyst under heat at 683 K for 90 min, using a digestor (Tecator Digestor 8, FOSS, Hillerød, Denmark). After cooling to room temperature, the greenish-transparent mixture was diluted with 60 mL of distilled water and neutralized with 80 mL of 40% NaOH. The ammonia (NH₃) gas produced was distilled into a 20 mL mixture of 4% H₃BO₃, 10 mL of bromocresol green, and 7 mL of methyl red solution using an automatic distillation unit (Kjeltec^TM 8100, FOSS, Hillerød, Denmark). The amount of NH₃ was then quantified by back-titration with a standard 0.1 mol·L⁻¹ H₂SO₄ solution to determine the nitrogen content. Finally, the protein content was estimated using the conversion factor, as described in Equation (3).

$$\mathrm{P}\mathrm{r}\mathrm{o}\mathrm{t}\mathrm{e}\mathrm{i}\mathrm{n} \mathrm{C}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{e}\mathrm{n}\mathrm{t}\left(\mathrm{\%}\right)=\left[{\displaystyle \frac{({\mathrm{V}}_1-{\mathrm{V}}_0)\times {\mathrm{C}}_{\mathrm{t}}\times 0.014}{\mathrm{m}}}\times 100\right]\times 5.6$$

(3)

where V₀ and V₁ represent the volumes of H₂SO₄ titrant (in mL) used for the blank and sample titrations, respectively, C_t is the concentration of H₂SO₄ (in mol·L⁻¹), and m is the mass of the test sample (in grams). Factor 5.6 is a nitrogen-to-protein conversion factor (Kp) applied to estimate the protein content of black soldier flies [26].

The chemical structures were analyzed using Fourier-transform infrared spectroscopy (FT-IR, Bruker, Alpha II). This technique is used to identify specific functional groups in chitin by examining the IR absorption frequencies present in the spectra. Samples were prepared as potassium bromide (KBr) pellets, with approximately 1 mg of the samples finely ground with 100 mg of KBr. The infrared spectra were recorded over a frequency range of 4000 cm⁻¹ to 500 cm⁻¹ with a resolution of 4 cm⁻¹ and 16 scans. The degree of deacetylation (DD) was calculated to evaluate the extent of acetyl group removal from chitin. The DD was determined using Equation (4).

$$\mathrm{D}\mathrm{e}\mathrm{g}\mathrm{r}\mathrm{e}\mathrm{e} \mathrm{o}\mathrm{f} \mathrm{D}\mathrm{e}\mathrm{a}\mathrm{c}\mathrm{e}\mathrm{t}\mathrm{y}\mathrm{l}\mathrm{a}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n}\left(\mathrm{\%}\right)=\left[1-\left({\displaystyle \frac{{\mathrm{A}}_{1655}}{{\mathrm{A}}_{3450}}}\times {\displaystyle \frac{1}{1.33}}\right)\right]\times 100$$

(4)

In this Equation, A₁₆₅₅ and A₃₄₅₀ represent the absorbance intensity at 1655 cm⁻¹ and 3450 cm⁻¹, which corresponds to the carbonyl (C=O) stretching of the acetyl groups in chitin and the N-H stretching in the amine groups of chitosan [13,27], respectively. Constant 1.33 represents the theoretical value for complete deacetylation.

Surface morphologies were assessed using a Field-Emission Scanning Electron Microscope (FE-SEM, SU3500, Hitachi, Tokyo, Japan) at an accelerating voltage of 5 kV. The crystal structures were characterized by powder X-ray diffraction spectroscopy (PXRD, MiniFlex 6th, Rigaku, Tokyo, Japan) with Cu Kα radiation (40 kV, 30 mA, λ = 1.5406 Å); the crystallization index (CrI) was also determined to evaluate the chitin structural properties and behavior. The diffractograms were recorded over a 2θ range of 3° to 90° with a step size of 0.02° and scan rate of 10°·min⁻¹. Thermal stability, composition, and decomposition behavior were observed using thermogravimetry differential thermal analysis (TG/DTA, STA7300, Hitachi, Tokyo, Japan). The measurements were conducted over a temperature range of 303 K to 903 K under nitrogen purging (N₂) with a flow rate of 100 mL·min⁻¹ and a ramping rate of 10 K·min⁻¹.

The solubility properties were determined by dissolving 0.1 g of samples in 10 mL of 0.3 mol·L⁻¹ CH₃COOH at an ambient temperature under constant mixing for 18 h. The remaining solid was filtered using Whatman filter paper no. 40 and dried at 353 K overnight. Finally, the solubility of samples (%) was calculated as weight differences before and after the treatment. Meanwhile, the viscosity-average molecular weight (M_V) was determined as described previously by Yuan et al. [19], with modification. A specific amount of chitosan with pre-determined solubility was dissolved in a 0.3 mol·L⁻¹ CH₃COOH/0.2 mol·L⁻¹ CH₃COONa buffer (pH 4.5) to achieve a concentration of approximately 0.01 to 0.05 g·dL⁻¹. Viscosity was measured using a capillary viscometer (513-20/200, Schott Geräte) at ambient temperature, by recording the flow time of the solvent/chitosan solution within a fixed volume. M_V (g·mol⁻¹) was estimated by using the Mark–Houwink–Sakurada equation, as follows:

$$\left[\mathsf{\eta }\right]=\mathrm{K}\times {\left[{\mathrm{M}}_{\mathrm{V}}\right]}^a$$

(5)

where [η] is the intrinsic viscosity of the sample (dL·g⁻¹), and K = 0.0076 and a = 0.76 are constants that depend on the solute, solvent, and temperature used.

3. Results

The chitin extraction process began with drying the raw puparium at 378 K for 24 h to remove moisture, as excess moisture can hinder subsequent steps. The dried puparium was then physically ground and sifted through a 60-mesh screen to produce fine particles (puparium-BSF powder). This grinding step was essential for achieving uniform particle size and increasing the surface area, thereby ensuring an effective extraction process, as is typical in such procedures. The puparium-BSF was subsequently subjected to a series of treatments, including deproteination (DP), demineralization (DM), and/or decolorization (DC), to produce chitin, as illustrated in Figure 1a. Digital image documentation of the resulting puparium-BSF and CT variants is also documented and shown in Figure 1b–i.

From the optical perspective, the digital images clearly show that the puparium-BSF powder (Figure 1b) exhibits a dark brown color, characteristic of melanin pigment typically found in insect exoskeletons [28]. While CT-1 and CT-2 display colors like puparium-BSF, the other CT variants (CT-3 to CT-7) appear significantly brighter after purification with additional decolorization, with CT-6 being the brightest. This suggests that more pigment was removed, indicating an improvement in purity [13]. However, further analysis is needed to validate these results. Thus, the chitin was then characterized through proximate analysis and various advanced physical characterization techniques, including scanning electron microscopy (SEM), powder X-ray diffraction (PXRD), Fourier-transform infrared spectroscopy (FTIR), and thermogravimetric analysis/differential thermal gravimetry (TG/DTG). These analyses will further evaluate the effects of each treatment variant on the extracted chitin, as discussed in the following section.

3.1. Physicochemical Analysis of Chitin

3.1.1. Proximate Composition

Table 2. Proximate analysis of puparium-BSF and CT variants in dry weight (%-dw).

Variant	Proximate Analysis (%-dw)				DDM (%)	DDP (%)	DD (%)
	Ash	Lipid	Protein	Chitin
Puparium-BSF	19.33 ± 0.15	17.19 ± 0.52	38.85 ± 0.31	24.62	-	-	-
CT-1	1.61 ± 0.02	3.84 ± 0.55	4.24 ± 0.05	90.31	91.68 ± 0.15	89.09 ± 0.03	10.17
CT-2	3.18 ± 0.26	6.64 ± 0.00	7.43 ± 0.11	82.75	83.56 ± 1.22	80.86 ± 0.43	25.20
CT-3	1.16 ± 0.08	2.33 ± 0.14	3.95 ± 0.09	92.60	94.23 ± 0.37	89.82 ± 0.16	40.13
CT-4	1.01 ± 0.09	1.99 ± 0.18	3.01 ± 0.15	93.98	94.77 ± 0.41	92.24 ± 0.31	18.09
CT-5	2.43 ± 0.15	3.28 ± 1.11	3.78 ± 0.09	90.50	87.42 ± 0.86	90.25 ± 0.16	46.18
CT-6	3.27 ± 0.19	1.89 ± 1.02	3.28 ± 0.08	91.57	83.09 ± 1.10	91.57 ± 0.15	65.90
CT-7	2.22 ± 0.06	2.63 ± 0.53	5.56 ± 0.09	89.59	88.50 ± 0.42	85.71 ± 0.14	22.17

presents the summarized results for the proximate analysis in dry weight for ash, lipid, and protein content for CT variants after the moisture content correction. The proximate analysis of the puparium-BSF was also conducted as a reference point to evaluate the effectiveness of deproteination (DP) and demineralization (DM), which is indicated by its ash content. The total chitin content was finally determined by subtracting the ash, lipid, and protein percentage from 100 on a dry weight basis, excluding the moisture content. The effectiveness of DM and DP was expressed through the degree of demineralization (DDM) and deproteination (DDP), calculated by comparing the ash and protein content to the initial value from the puparium-BSF (19.33% and 38.85%, respectively). For comparison, Tables S1 and S2 in the Supporting Information show the proximate analysis of each deproteination, demineralization, and decolorization treatment. The moisture content of the samples ranges from 0.68% to 5.56%, with the discrepancy likely arising from the storage conditions before the analysis. A similar trend is observed in the main samples, puparium-BSF and CT variants, which range from 3.06% to 9.68% (Table S3).

The individual treatments of deproteination, demineralization, and decolorization show distinct trends in the proximate analysis results. Specifically, after deproteination, the sample exhibits nearly identical ash content to that of puparium-BSF but with a noticeable decrease in lipid and protein, which drops to 4.37% and 24.62%, respectively, resulting in a DDP of 36.5%. In contrast, demineralization reduces the ash content to 3.9% resulting in a DDM of 79.83%, with no significant change in protein and lipid content. In the case of decolorization, both ash and lipid content decrease to 10.56% and 7.4%, with no considerable change in protein content, yielding DDM and DDP values of 45.39% and 25.13%, respectively. These single treatments result in relatively low chitin purity (<60%), as each treatment plays a distinct role in the purification process. Therefore, integrating all treatments is essential for achieving efficient purification and higher chitin purity. A detailed discussion of the effects of each treatment will be provided in the following section, with a focus on ash and protein content, as they reflect the scope of this study.

As expected, a significant decrease in minerals and proteins is observed across all CT variants, with the degree of demineralization (DDM) decreasing to about 83.09 ~ 94.77% and protein content (DDP) decreasing to 80.86 ~ 92.24%. Nonetheless, CT-4 shows the lowest ash content among the CT variants statistically, which decreased significantly to 1.01% from 19.33%—observed in puparium-BSF, following sequential demineralization, deproteination, and decolorization. Similarly, lipid and protein content demonstrated a comparable trend, with concentrations decreasing significantly from 17.19% and 38.85% to 1.99% and 3.01%, resulting in a total chitin content of 93.98%. This value is comparable to, or even higher than, those reported in recent studies on chitin purity extracted from BSF at various life stages, particularly observed from the puparium (Table S4). While lipid removal is commonly performed in some studies, we highlight that the combined treatments of deproteination, demineralization, and decolorization are essential, as treatment with only deproteination/demineralization or both combinations results in lower chitin purity (<80%). On the other hand, CT-2 exhibits an opposite trend compared to CT-4; despite undergoing similar treatment sequences of demineralization and deproteination, CT-2 shows a higher ash and protein content (3.18% and 7.43%) than CT-1 (1.61% and 4.24%).

While demineralization appears to be essential as the initial step, the absence of decolorization may hinder further mineral removal in the overall extraction process. These results are consistent with those reported in previous studies, which showed that chitin subjected to decolorization after purification has significantly lower residual minerals [29]. When deproteination is adopted as the initial step (CT-1, CT-5, and CT-7), no significant differences are observed in proximate analysis, despite the different sequences of DM and DC treatments, further confirming the importance of demineralization as the initial step. On the other hand, the modified simultaneous deproteination and decolorization (CT-6 and CT-7) do not exhibit a significant decrease in ash and protein content. However, CT-6 shows the highest ash content of 3.27% while demonstrating a significantly high degree of deacetylation (DD) of 65.90%, compared to other variants ranging from 10.17% to 46.18%. This indicates that most of the acetyl groups (−COCH₃) in the chitin have been removed, indicating partial conversion to chitosan. Nonetheless, this DD aligns with typical values for commercial chitosan, which usually exceed 55% [30]. A more detailed discussion on DD will be provided in a subsequent section, focusing on related physical characterizations.

We hypothesize that the higher DD results from the exothermic reaction between NaOH and H₂O₂ (2NaOH (aq) + 3H₂O₂ (aq) → Na₂O₂ (aq) + 2H₂O (l)). This reaction generates additional heat, leading to a temperature increase of approximately 10° raising it to around 363 K. The use of H₂O₂ as a decolorization agent likely disrupts the chitin polysaccharide structure while also breaking down melanin into smaller, more soluble compounds through the generation of reactive oxygen species, particularly the perhydroxyl anion (HO₂^-), which forms via deprotonation under alkaline conditions [31,32]. Since decolorization can lead to further degradation of chitin, and given that the deacetylation reaction typically occurs at elevated temperatures (between 363 K and 393 K) [29,33,34], this combination of treatments and the additional heat facilitates partial deacetylation despite the lower NaOH concentration usually adopted for the reaction (~50% w/v) [17], thus leading to the conversion of chitin to chitosan. Additionally, while the increase in temperature may influence the deacetylation process, this reaction also produces sodium peroxide (Na₂O₂) as an additional mineral.

This result aligns with observations from the deproteination and decolorization, excluding the demineralization treatment (Table S2). The simultaneous deproteination and decolorization resulted in a higher ash content (17.47%) compared to the sequential treatment (15.56%), which may explain the increase in ash content observed in CT-6. We are particularly interested in the detailed reaction and its effect on chitin extraction, especially on chitosan, with findings to be reported in due course. On the other hand, the lower DD observed in CT-7, despite the similar simultaneous deproteination and decolorization, is close in magnitude to that of CT-2 and CT-4 (~20%), both of which underwent demineralization as the initial step. The presence of protein layers may act as an adhering layer, preventing hydrolysis, depolymerization, and mineral dissolution, thereby protecting the chitin from degradation, including deacetylation [35]. In addition, the simultaneous reaction of demineralization and decolorization was not performed due to safety concerns, as the interaction between HCl and H₂O₂ produces toxic chlorine (Cl₂) gas (H₂O₂ (aq) + 2HCl (aq) → Cl₂ (g) + 2H₂O (l)).

Given these results, it is clear that the purification process plays a vital role in tuning chitin purity (in terms of residual minerals, lipids, and proteins), as the extent of purity influences its suitability for various applications. The purity of chitin is particularly relevant in industries such as biomedicine and pharmaceuticals [36]. While the moderate purity typically used for non-medical applications may suffice for general purposes [37], achieving higher purity levels would likely involve refining key purification processes, such as demineralization, deproteination, decolorization, and others. Moreover, factors like the choice of reagents, their concentrations, and reaction times also significantly influence chitin purity, regardless of the methods used.

3.1.2. Solubility, Intrinsic Viscosity, and Viscosity-Average Molecular Weight

Table S5 (Supporting Information) presents the solubility properties in an aqueous acetic acid solution. Most of the chitin samples exhibit low or nearly negligible solubility (<8%), consistent with its inherent insolubility in most solvents [5]. However, CT-3, CT-5, and CT-6 show relatively higher solubility values of 25.30%, 27.80%, and 41.05%, respectively, with CT-6 exhibiting the highest. These results align with their considerable DD, as chitosan typically demonstrates greater solubility in weak organic acids [38]. Furthermore, the estimated viscosity-average molecular weight (M_V) reveals that CT-3 and CT-4 exhibit an increasing trend in line with their intrinsic viscosities (29.01 dL·g⁻¹ and 32.05 dL·g⁻¹, respectively). Despite this trend, both samples show similar molecular weight magnitudes of ~50 kDa (51.62 kDa and 58.85 kDa), indicating comparable molecular weight characteristics.

In contrast, CT-6 displays an opposite trend, showing a significantly lower M_V of 42.39 kDa with an intrinsic viscosity of 24.98 dL·g⁻¹. Similar findings were reported in the study by Triunfo et al. [13], in which the purification process with additional decolorization was shown to decrease M_V due to the scission of polysaccharide chains or depolymerization induced by strong deacetylation conditions. For CT-6 specifically, the exothermic reaction between NaOH and H₂O₂ during the combined deproteination and decolorization treatments likely exacerbated the degradation, resulting in a reduced M_V. Yet, these M_V values are typical for chitosan derived from insects, generally ranging from 26 kDa to 450 kDa [13,39].

This variability in molecular weight (MW) of chitosan is significant, as it also influences its diverse applications. Generally, low-molecular-weight (LMW) chitosan, characterized by low viscosity, is used in animal husbandry, pharmaceuticals, and certain medical applications [40]. On the other hand, medium- (MMW) and high-molecular-weight (HMW) chitosan, though more constrained by their high viscosity, are utilized in agriculture, veterinary care, cosmetics, and water treatment [41]. However, beyond molecular weight, intrinsic factors such as particle size, degree of deacetylation, and surface characteristics also play a crucial role in shaping its functional properties, including antibacterial and antioxidant activity, biodegradability, and/or solubility [42]. This complexity further allows chitosan to be tailored for specific uses, regardless of its molecular weight.

3.2. Characterization

3.2.1. Surface Morphology

The surface morphology of the puparium-BSF and the CT variants was further analyzed using SEM (Figure 2). Figure 2a shows an SEM image of the puparium-BSF at 250× magnification, revealing a cuticle with a rough and uneven structure characterized by small aggregates of CaCO₃ scattered across the surface, typical of untreated puparium-BSF [43]. In contrast, CT variants (Figure 2b–h) reveal enhanced clarity and a solid structure characterized by well-defined repeating units arranged in a hexagonal tile configuration, with each tile featuring a depression in its center resembling a honeycomb pattern, as also reported in other studies [14,34,43]. However, upon closer examination at a magnification of 5000×, the CT samples reveal noticeable surface differences. Specifically, CT-1 and CT-2 (Figure 2a,b) exhibit a ragged surface, while the other CT variants, which underwent additional decolorization treatment, show a relatively smooth surface.

The smooth surface may have originated from the decolorization treatment, as a similar finding was reported in the study by Ifuku et al. [29], in which shrimp-derived chitin nanofibers (CNFs) exhibited an exfoliated and thinner structure after decolorization compared to the surface before treatment. This suggests a degradation of the chitin surface layer due to decolorization, as also mentioned previously. However, while the other CT variants display a relatively similar surface morphology, CT-5 and CT-6 exhibit a fibrous-like structure commonly found in chitosan [34]. This result further supports the higher DD observed in this sample, with the highest in CT-6. Despite the differences in surface morphology between chitin and chitosan, it is noteworthy that the micromorphology of chitin can vary among studies, as it strongly depends on combinations of several factors, including insect species, growth/maturity stages, and gender [12]. Nevertheless, this homogeneous surface morphology indicates the effective removal of proteins and minerals [44], which aligns with the proximate analysis results.

3.2.2. Crystalline Structure

The crystal structure of the puparium-BSF and CT variants was characterized using PXRD, as illustrated in Figure 3a. In agreement with the literature [43], the diffraction pattern of puparium-BSF reveals the typical features of a polycrystalline structure consisting of a mixture of chitin and calcium carbonate (CaCO₃) phases. Diffraction peaks are perceived at 2θ values of approximately 9.5°, 19.5°, 23.3°, and 40.7°, which can be assigned to the (020), (110), (130), and (124) crystal planes, respectively, corresponding to the orthorhombic structure of α-chitin (JCPDS #35-1974). Additionally, peaks at 29.7°, 36.2°, 39.7°, 43.5°, 47.8°, 48.9°, 57.8°, and 61.3° are assigned to the (200), (020), (211), (022), ($\overline{3}$12), ($\overline{2}$22), (130), and ($\overline{1}$32) crystal planes, corresponding to the monoclinic structure of CaCO₃ (JCPDS #70-0095). In contrast, all CT variants exhibit relatively similar reflection; however, there are prominent existing α-chitin peaks along with new peaks appearing at approximately 26.5°, 28.2°, 28.8°, and 39.2°, corresponding to (140), (023), (132), and (171) planes. The absence of the CaCO₃ reflections suggests that the chitins are essentially pure within the limitations of the PXRD analysis and are free from other mineral or protein-related organic phases. For reference, no peaks corresponding to chitosan or significant shifts are observed according to the JCPDS 39-1894 database, except for a decrease in intensity in CT-3, CT-5, and CT-6, despite their higher DD.

In addition, the crystallinity index (CrI) of the samples was calculated (Table S5), revealing values of 17.8% for puparium-BSF. In contrast, CT variants exhibit significantly high CrI values of 35%, 40%, 45%, and 34% for α-chitin, observed from CT-1, CT-2, CT-4, and CT-7. Meanwhile, relatively lower values were observed in CT-3 (18%), CT-5 (20%), and CT-6 (23.9%), particularly in those with higher DD. These lower values mostly correspond to a decrease in the acetyl group, which leads to weak intermolecular hydrogen bonding between hydroxyl and acetyl groups in an antiparallel arrangement, which also explains the decreased peak intensity [45]. Nonetheless, the CT variant that underwent demineralization as initial treatment (CT-2, CT-4, and CT-7) tends to have higher CrI, which can be attributed to a protein adhering layer, as mentioned earlier. This layer retains mineral content, resulting in chitin with higher ash content and a consequential higher CrI [35]. When deproteination is the initial step, the mineral layer is exposed, leading to lower mineral content and a lower CrI in general [36,46].

However, despite the CrI values, which reflect the characteristics of disordered amorphous material due to the lower crystalline properties, these values are relatively consistent with the results reported for BSF-derived chitin, ranging from 24.9% to 96.4%, with specific values of 33.1% to 68.4% from the early growth stages [17,47]. However, the CrI may vary based on the source and the extraction process, such as the acid concentration and the inclusion of decolorization, as these factors could account for variations in the CrI of the polymer [13,35]. In general, CrI indicates the degree of crystallinity, which further influences the physical properties of chitin, including mechanical strength, thermal stability, and chemical resistance. These diverse properties lead to different potential applications depending on the CrI value [27,34]. More amorphous chitin is generally used in wastewater treatment due to its adsorbent properties [48], while highly crystalline chitin is commonly used in cosmetic and biomedical fields [49].

3.2.3. Chemical Functional Group

Specific functional groups in puparium-BSF and CT variants are evaluated with FTIR. The FTIR spectra of all samples are illustrated in Figure 3b and demonstrate that the puparium-BSF and CT variants show a relatively similar IR reflection of the characteristic functional groups associated with chitin’s structure in their specific wavelength, indicating that the chitin was successfully isolated. As observed, broad bands recorded at 3488, 3447, 3448, 3481, 3445, and 3481 cm⁻¹ in the spectra of CT variants indicate hydroxyl (OH) stretching vibrations. Additionally, weaker bands at 3280 cm⁻¹ observed in puparium-BSF and those at 3264, 3269, and 3266 cm⁻¹ for CT-1, CT-4, and CT-7 signify N-H asymmetric stretching [50]. On the other hand, bands at 3106, 3113, 3108, 3110, and 3108 cm⁻¹ for all CT variants in sequence, except CT-2 and CT-6, indicate N-H symmetric stretching associated with secondary amides, which is typical for chitin. Aliphatic C-H symmetrical stretching was observed in the range of 2851 cm⁻¹ to 2936 cm⁻¹, demonstrating the preserved aliphatic structure despite the different treatment sequences [27,51].

The sharp band at 1737 cm⁻¹ observed for CT-6 was attributed to the C=O stretching vibration of the ester group [52]. The primary amide band, associated with C=O stretching of secondary amides, was detected at 1660 cm⁻¹ in puparium-BSF and 1662, 1648, 1659, 1658, 1658, 1632, and 1660 cm⁻¹ in the CT variants, specifically confirming the α-crystalline structure of chitin [27]. Bands corresponding to Amide II, which include N-H bending and C-N stretching, were found at around 1544 for puparium-BSF and 1553, 1555, 1562, 1552, 1559, 1544, and 1558 cm⁻¹ for CT variants. The presence of Amide III, indicated by the C-N stretch [13], was identified in the range of 1314 cm⁻¹ to 1321 cm⁻¹, while the peaks in the region between 1026 cm⁻¹ and 1259 cm⁻¹ were attributed to asymmetric oxygen bridging and C-O stretching, signifying the glycosidic bonds in the chitin structure [51].

Additionally, bending vibrations of C-H groups in aromatic compounds were also observed at around 701 cm⁻¹ to 897 cm⁻¹ [16]. While all the samples exhibit a profile similar to typical chitin spectra, a significant difference is observed in CT-6. Compared to the other CT variant spectra, this chitin shows a decreasing trend in the peak at 1544 cm⁻¹, which could signify effective deacetylation. Generally, a signal in this frequency range indicates successful deproteination linked to the presence of peptide bonds; therefore, its absence suggests a low protein content in the CT [27,34]. These results further confirm the higher degree of deacetylation especially observed in CT-6.

3.2.4. Thermal Behavior

Thermal analysis (TG/DTG) was performed to assess the thermal stability of the sample. Thermal properties of chitin are essential, as they not only determine the effectiveness of extraction treatments but also indicate its resistance to irreversible changes in chemical/physical structure when exposed to elevated temperature, which further influences its potential applications [53]. Figure 4a–i shows the TG/DTG thermogram obtained from puparium-BSF and the CT variants, revealing a comparable thermodegradation event with three main decomposition steps. The initial thermal event occurred sharply between 299 K and 369 K, resulting in a weight loss of approximately 6 ~ 13% across all samples, attributed to adsorbed water evaporation. However, since the samples have relatively low moisture content, this could not be appreciated well in the thermogram. Following this, the second event associated with the decomposition temperature (T_D) of the samples occurred in the range of 376 K to 695 K.

This subsequent event was linked to the thermal dehydration, depolymerization, and decomposition of saccharide chains, involving the random breaking of C−O−C skeletal bonds, which leads to vaporization and the release of volatile compounds. Finally, the pyranose ring degrades at 673 K with about 6 ~ 11% weight loss contribution, making the last thermal event [53,54,55]. For reference, the sequential losses observed earlier at 467 K and 584 K for puparium-BSF, 587 K, 450 K, and 585 K for CT-2, CT-6, and CT-7 could be attributed to the degradation of protein, lipid, and pigment chains [12,56]. A similar phenomenon was also observed in other CT variants, though it overlaps with the chitin T_D peak due to minor lipid and/or protein chains in the samples. The total weight loss for puparium-BSF (Figure 4a) was around 65%, while the CT variants (Figure 4a–h) exhibited variable losses of 78% (CT-1), 70.7% (CT-2), 85.1% (CT-3), 78% (CT-4), 87.4% (CT-5), 84.5% (CT-6), and 82% (CT-7). The significantly higher total weight loss observed from the CT variants is attributed to their higher purity, as they contain a more significant proportion of chitin compared to puparium-BSF, which retains a substantial number of proteins, lipids, pigments, and non-degradable inorganic substances (e.g., minerals and salts).

In addition, while the TG/DTG analysis reveals that the optimum T_D of puparium-BSF occurs at 646 K, the T_D for the CT variants ranges between 640 K and 669 K. The observed shifting of the optimum T_D to a higher temperature region in the CT variants (Figure 4i), along with the purity factor, is also influenced by the CrI. Since the CT variants exhibit a higher CrI, this contributes to a more crystalline and stable structure, resulting in a higher optimum T_D [17]. While other CT variants showed relatively similar T_D values around 660 K, CT-3, CT-5, and CT-6 exhibit a slightly lower trend, with the lowest observed from CT-6 (T_D of 652 K). This phenomenon is mainly linked to the lower CrI, which is also correlated with its lower acetyl unit content, as the T_D tends to decrease with decreasing acetyl units [55]. This is further explained by its higher DD of 65.9%.

4. Conclusions

This study demonstrates an optimal method for extracting high-purity chitin and emphasizes the importance of treatment combinations and sequences. A series of chitin (CT) variants was successfully extracted from the BSF puparium through various treatment sequences. The proximate analysis, morphology, crystal structure, chemical functional groups, and thermal properties of the CT variants were thoroughly examined to elucidate the effects of each treatment sequence and combination. High-purity chitin was obtained from CT-4 through the treatment sequences of demineralization (DM), deproteination (DP), and decolorization (DC), achieving the lowest ash, lipid, and protein contents—approximately 1.01%, 1.99%, and 3.01%, respectively—with degrees of DM and DP of 94.77% and 92.24%, yielding a chitin purity of 93.98%. Conversely, CT-6, with the simultaneous combination of DP and DC following the DM process, exhibits the highest degree of deacetylation (DD) at 65.90%, suggesting partial deacetylation to chitosan. Furthermore, this study shows an alternative approach to directly produce chitosan with a higher DD by eliminating the conventional deacetylation process. This condition provides excellent prospects for making high-tech products at lower costs.