Skip to main content
NCBI home page
ACS Omega logoLink to ACS Omega
. 2025 Apr 21;10(16):17015–17023. doi: 10.1021/acsomega.5c01987

Obtention and Characterization of Chitosan from Exuviae of Tenebrio molitor and Sphenarium purpurascens

M Selene Marín-Morales †, Celeste C Ibarra-Herrera †,*, María J Rivas-Arreola ‡,*
PMCID: PMC12044503  PMID: 40321518

Abstract

graphic file with name ao5c01987_0006.jpg

Chitosan is a versatile biopolymer with applications in various industries due to its biocompatibility and biodegradability. While crustacean shells are the primary source of chitosan, the extraction process can be environmentally taxing. This study focuses on evaluating chitosan from Tenebrio molitor and Sphenarium purpurascens, two insect species that offer a more sustainable alternative and require fewer resources for cultivation and produce large amounts of chitin-rich biomass. The acid–alkali method was applied using three experimental conditions (M1, M2, and M3) that involved modification in the demineralization, deproteinization, and deacetylation steps. The chitosan samples were characterized by determining the degree of deacetylation, solubility, molecular weight, antimicrobial activity, and swelling capacity and furthermore by measuring deproteinization and impurity content. Also, Fourier-transform infrared spectroscopy (FTIR) analysis was also performed on the chitosan samples. Samples M3 from the exuvia of T. molitor (DD 55.62% ± 0.79 and solubility 24.13% ± 2.07) and M1 from S. purpurascens (DD 61.86% ± 4.98 and solubility 27.31% ± 1.87) presented the best performance. The molecular weight was calculated between 75 kDa and 118 kDa. On the other hand, data obtained for swelling tests suggested that the film obtained from sample M1 (T. molitor, 5.07% ± 0.11) proved to be more resistant to degradation in an aqueous environment, suggesting that this chitosan could be used for designing a film with high resistance. By exploring these insect sources, this research aims to contribute to the development of chitosan production practices, with potential applications in water treatment, biomedicine, and food packaging, thereby expanding the availability and uses of this valuable biopolymer.

1. Introduction

Biopolymers have gained attention due to their biodegradability and eco-friendly properties, making them suitable for various applications.1 For instance, they are used in biodegradable packaging, healthcare2 (e.g., sutures and drug delivery systems), agriculture as mulch films,3 and even automotive and construction industries as biobased materials.4 This reduces reliance on traditional plastics and fossil fuels, addressing environmental concern. After cellulose, chitin is the second most abundant biopolymer in nature and is found in crustaceans, fungi, and insects. Chitin is a linear polymer chain made up of repeating units of N-acetylglucosamine linked by hydrogen bonds between polymer chains and the acetyl groups attached to the monomers conferring to chitin a highly stable and resilient structure.5 Chitosan is the biopolymer obtained when chitin is partially or completely deacetylated, and it is formed by units of glucosamine and N-acetylglucosamine linked by β-(1,4) glycosidic bonds. Chitosan is mainly extracted from crustacean shells, accounting for about 90–92% of total commercial production. Globally, approximately 70,000 tons of chitosan are produced each year.6 This biopolymer has gained significant importance due to the versatility of its applications, for example, chitosan nanoparticles are explored for their potential in drug delivery, in biosensors, and as carriers of genetic material in gene therapy.7 In food industry, chitosan is used for food preservation, for packaging, and as an additive to enhance the shelf life of products.8,9 Also, chitosan is effective in wastewater treatment,10 where it aids in the removal of heavy metals and pollutants due to its adsorption capabilities.11 Insects are currently being reported as a potential source of chitin for chitosan production offering a sustainable alternative to crustaceans, reducing environmental impact and waste using farming byproducts.12Tenebrio molitor, commonly known as mealworm, stands out as one of the most promising edible insect species globally. During specific larval stages in its life cycle, a substantial quantity of exuviae is produced.13 According to reported data, the primary composition of its exuviae comprises 10–25% of chitin.14−16 Previous studies have demonstrated the production of chitin and chitosan from various sources of T. molitor, including exuviae, larvae, and adult beetles highlighting their potential as sustainable alternatives to commercial chitosan and demonstrating the feasibility of utilizing insect-derived materials for industrial applications.14,16 On the other hand, Sphenarium purpurascens which is an edible grasshopper has a chitin content between 8 and 15% dry base;17,18 this compositional profile suggests that the grasshopper could serve as a sustainable source for chitosan production. The extraction of chitin from crustaceans is carried out through chemical processes that include demineralization with hydrochloric acid (HCl), deproteinization with sodium hydroxide (NaOH), and in some cases, decolorization.19 The advantages of these methods include their effectiveness and speed, while the disadvantages encompass the use of aggressive chemicals that can be pollutants and the high cost of waste treatment,20 and these methods are widely applied in insects to obtain chitin and chitosan;21 however, certain methods for extracting chitin and chitosan can be excessively aggressive, leading to undesirable modifications, including the depolymerization of chitosan.22 Employing less aggressive processes during chitin extraction is essential for preserving the polymer’s integrity and producing biopolymers with desirable properties as solubility, degree of deacetylation, the capability to form films or gels, and so on.

In this study, the acid–alkali method was used for the extraction of chitin and production of chitosan from the exuviae of T. molitor and S. purpurascens. Three different milder experimental conditions were evaluated to analyze their effectiveness in chitin for chitosan production. The physicochemical properties, such as solubility, molecular weight, and degree of deacetylation, were evaluated; also, biological properties, such as antimicrobial activity, were also assessed to determine their impact on chitosan’s performance. Fourier-transform infrared spectroscopy was used for the characterization of functional groups. Additionally, mechanical properties as swelling degree were measured, and the production of chitosan films from these sources was evaluated for their potential applications in packaging, biomedical, and environmental solutions.

2. Materials and Methods

The exuvia of T. molitor was obtained from Zuustento Company located in Tequisquiapan, Queretaro, México, and S. purpurascens was obtained from a maize field located in Coronango, Puebla, México. The exuviae of T. molitor were ground (Nutribullet NBR-0601) and then sieved through an Elvec 100 mesh to remove straw residues, bran, and other contaminants. Additionally, the grasshoppers were freeze-dried (Labconco Freezone 4.5, Kansas City, MO, USA), then ground and sieved through a 100-mesh. All reagents used in the proximal analyses were of analytical grade. As standards, a low molecular weight chitosan and a medium molecular weight chitosan (Sigma-Aldrich, Cas number 9012-76-4) were used. Spectrograms were obtained using Origin (Pro) Version 2024, OriginLab Corporation, Northampton, MA, USA.

2.1. Determination of Proximal Composition

For proximate analysis, moisture (AOAC 926.12), content of ashes (AOAC 942.05), percentage of protein (AOAC 920.87), crude fat (AOAC 920.39), and dietary fiber (AOAC 985.29) were the parameters determined with AOAC methodologies. Total carbohydrates were determined by the sulfuric acid spectrophotometric method23 using spectrophotometer HACH (mod DR6000 CO, USA).

2.2. Extraction of Chitin and Chitosan Obtention

For the extraction of chitin, the following steps described by Reyes24 were followed (M1): first demineralization was carried out mixing 20 g of samples with 5% of HCl in a 1:6 w/v ratio, at 40 °C during 2 h in agitation using a stirring hot plate (Cole-Parmer, 03407-31, IL, US), and then the samples were filtered through mesh cloth and washed with distilled water until acid residues were eliminated and subsequently dried overnight at 60 °C in drying oven Memmert IFP-500 (Bellevue WA, USA). The next step was deproteinization which was carried out by mixing 3% of NaOH at a 1:6 w/v ratio and heating at 60 °C for 4 h under agitation. The samples were washed until alkaline residues were removed and dried overnight at 60 °C. Finally, deacetylation was carried out mixing the samples with 50% of NaOH in a 1:6:2 w/v ratio (sample, NaOH, water), with agitation at 80 °C for 2 h, washing until the residues of NaOH are eliminated and drying overnight at 50 °C. The second experimental condition (M2) was like described above, but the demineralization step was omitted. The third experimental condition (M3) was carried out according to Song et al.;14 this process consisted of two steps: deproteinization using 5% of NaOH in a 1:6 w/v ratio for 3.5 h at 70 °C in agitation, and deacetylation using 50% of NaOH at 121 °C (15 psi) (Viresa, ALL-50X-120 V, MX) for 30 min.

Then, to calculate the yield of chitin relative to the raw material, eq 1 was used. To determine the yield of chitosan relative to the raw material, eq 2 was employed.

2.2. 1
2.2. 2

2.3. Physicochemical Characterization

2.3.1. Demineralization

To evaluate the percentage of demineralization, ash content was measured according to AOAC 942.05 using a furnace of the brand Felissa (mod FE-360, MX). The result was calculated using eq 3

2.3.1. 3

where ash content1 corresponds to the value obtained in proximal analyses and ash content2 is obtained after the demineralization process.

2.3.2. Deproteinization

Deproteinization was determined by the Kjeldahl method using Labconco distiller mod 56000 (Kansas City, MO, USA). All experiments were carried out in triplicate. The DP (%) was calculated using eq 4

2.3.2. 4

where protein content1 corresponds to the value obtained in proximal analyses and protein content2 is obtained after the deproteinization process.

2.3.3. Degree of Deacetylation

The determination of the degree of deacetylation (DD) was performed by the potentiometric method using a potentiometer (Hanna Instruments HI 2550, RU, EUR). Briefly, 0.1 g of chitosan was dissolved in 25 mL of 0.1 M HCl. For the titration, a standard solution of 0.1 M NaOH was used. All experiments were carried out in triplicate. The DD was calculated using eq 5(24)

2.3.3. 5

where 16.1 is the protein-related factor, y is the major inflection point expressed in volume units, x corresponds to the minor inflection point expressed in volume units, w is the mass in grams of the sample, and f is the molar concentration of NaOH.

2.3.4. Solubility

The solubility of chitosan was tested using a method adapted from Luo et al.25 First, 0.1 g of chitosan was mixed with 10 mL of 1% of acetic acid, then agitated at 30 °C for 1 h, and centrifuged (Eppendorf 5703, Germany) at 3220g for 15 min, and finally, the sediment was dried at 60 °C overnight. The solubility was calculated using eq 6.25

2.3.4. 6

where M0 is the initial weight of the tube and M1 and M2 are the initial weight of the tube plus the sample. All experiments were carried out in triplicate.

2.3.5. Determination of Molecular Weight by Viscosity

The solutions of chitosan were prepared in concentrations of 0.08, 0.12, 0.16, 0.2, and 0.24 g mL–1 in a solution of acetic acid (0.20 M) and sodium acetate (0.15 M). A capillary viscosimeter in a water bath at 25 °C was used. The intrinsic viscosity was calculated using Mark–Houwink–Sakurada eq 7.5

2.3.5. 7

where [η] is the intrinsic viscosity and K and a are the empirical constants; for the chitosan solution system, the values of the constants are K = 7.4 × 10–4 cm3 g–1 and a = 0.86, as reported by Rinaudo 2006.5

2.3.6. Fourier-Transformed Infrared Spectroscopy

The spectral analysis was conducted on multiple samples: 3 samples of chitosan from T. molitor, 3 samples of chitosan from S. purpurascens, and two standards of chitosan Sigma-Aldrich. Each sample was analyzed three times, and the spectra report represents the average of the three acquisitions made at different points on the same sample. The FTIR technique was performed using a Shimadzu IRTracer-100 spectrophotometer (Kyoto, Japan) with a frequency range of 4000–600 cm–1. The analysis was conducted using the Attenuated Total Reflectance (ATR) technique.

2.3.7. Film Ability and Swelling Degree

For these tests, 0.5 g of chitosan was dissolved in 50 mL of a 1% (v/v) acetic acid solution according to Triunfo et al.26 From the solution before prepared, 25 g of this mixture was weighed and placed into a Petri dish with a 100 mm diameter. The Petri dishes are left uncovered at room temperature for a period of 2 days. Subsequently, pieces of the film of 2 × 2 cm were weighed and placed in distilled water. After 10 min, 30 min, and 60 min and 24 and 48 h, the weight of the films was measured.27 The swelling rate was calculated using the following eq 8

2.3.7. 8

where Wi is the initial weight (g) of the film and Wf is the final weight (g) after each period.

2.3.8. Determination of Antimicrobial Activity

The methodology employed followed the approach proposed by Hongpattarakere and Riyaphan,28 with modifications, and included the use of a multiple dilution test. The strains used were Staphylococcus aureus ATCC 25923, Escherichia coli ATCC 25922, Salmonella spp., Pseudomonas aeruginosa ATCC 77853, and Enterobacter aerogenes ATCC 13048. Bacterial standardization was performed by inoculating one colony into trypticase soy broth (TSB) and incubating at 35 °C for 24 h. Using the dilution method, bacterial counts were determined, obtaining cultures of approximately 1 × 108 CFU/mL. This process was repeated for inoculum standardization and testing. A 1% chitosan solution in 0.5% acetic acid (pH adjusted to 4.8) was prepared, and 5 mL were added to tubes containing TSB with bacteria. After incubation at 35 °C for 24 and 48 h, viable cells were quantified using the dilution method.

2.4. Statistical Analysis

The characterization data of chitosan underwent statistical analysis through one-way analysis of variance (ANOVA) and Tukey’s test, using Minitab 19 software from Minitab Inc. (State College, PA, USA). A p-value threshold of less than 0.05 was established to determine statistical significance. Each sample was analyzed in triplicate, with the standard deviation calculated, and a normality test was conducted to assess the distribution of the data.

3. Results and Discussion

3.1. Chitin Extraction and Chitosan Production

Edible insects generally exhibit a low ashes content (see Table 1), which for the exuviae of T. molitor was 5.4% ± 0.01 and S. purpurascens was 4.4% ± 0.3, indicating a reduced presence of mineral impurities. The yields of chitin and chitosan raw material are shown in Table 2. In this study, the yields of chitin obtained from the three experimental conditions employed for the exuviae of T. molitor ranged from 11% to 12.4% and among these treatments did not present statistical differences (p < 0.05). Notably, M1 had a demineralization step, which did not present any change in yield, likely due to the inherently low mineral content on insect exuviae. For S. purpurascens, the yields were M1: 16.85% ± 0.86, M2: 18.41% ± 0.59, and M3: 13.09% ± 1.12. While no statistical differences were observed between M1 and M2, M3 was significantly different from both M1 and M2 (p < 0.05). The significantly lower yield observed in M3 compared to M1 and M2 can be attributed to the more aggressive deproteinization conditions which may have led to partial chitin degradation. Compared with other works, Purkayastha and Sarkar29 reported chitin yields ranging from 9% to 23% from the pupal exuviae and imago, respectively, of Hermetia illucens using harsher conditions (1 M HCl solution at 100 °C for 30 min and 1 M NaOH solution at 80 °C for 24 h). Similarly, Kim et al.30 reported chitin yields from Gryllus bimaculatus of 11.84% using 50% NaOH at 95 °C for 3 h. Also, Kleden et al.31 reported yields of the imago and exuviae from Locusta migratoria of 8.79 and 12.05%, respectively, boiling it with 4% NaOH during 1 h, then 1% HCl solution for 24 h, and finally, 3 N NaOH solution, and heating at 90 °C. The yields reported of chitin extraction from the insects are in the range of 8–20% and depend on the method of extraction, raw material, and insect’s developmental stage.43

Table 1. Proximal Composition from the Exuviae of Tenebrio molitor and Sphenarium purpurascensa.

component T. molitor exuviae S. purpurascens
moisture 6.4b ± 0.28 72.7a ± 0.91
ash 5.4a ± 0.01 4.4a ± 0.30
crude fat 3.7b ± 0.1 11.4a ± 1.6
protein 26.3b ± 0.4 57.6a ± 3.1
carbohydrates 49.2a ± 0.6 26.6b ± 0.83
dietary fiber 22.8a ± 1.4 11.1b ± 0.87
Open in a new tab
a

Content is expressed in g/100 g of dry matter. All values represent the mean ± SD by triplicate. a,b Different letters between columns indicate a statistically significant difference based on Tukey’s test p values = 0.05.

Table 2. Yields of Chitin Extraction, Chitosan Obtained, and Physicochemical Characterizationa.

parameter physicochemical characterization
  exuviae of T. molitor
 S. purpurascens
  M1 M2 M3 M1 M2 M3
yield of chitin/raw material (dw) 12.04a ± 0.51 12.43a ± 0.92 11.32a ± 0.88 16.85a ± 0.86 18.41a ± 0.59 13.09b ± 1.12
yield of chitosan/raw material (dw) 6.77b ± 0.38 8.27a ± 0.60 8.87a ± 0.35 6.76b ± 0.33 8.84a ± 0.14 8.70a ± 0.23
DM (%) 63.89a ± 0.31 50.55b ± 0.14 46.85c ± 0.06 86.81a ± 0.04 63.18b ± 0.04 25.45c ± 0.12
DP (%) 69.49a ± 0.23 69.45a ± 0.17 74.97b ± 0.20 58.52a ± 0.95 59.46a ± 0.13 60.53a ± 0.44
DD (%) 42.62b ± 0.38 25.25c ± 1.2 55.62a ± 0.79 61.86a ± 4.98 44.87b ± 1.38 43.77b ± 1.52
solubility (%) 12.74b ± 0.82 9.81c ± 0.81 24.13a ± 2.07 27.31a ± 1.87 13.64b ± 1.37 27.11a ± 1.21
molecular weight (kDa) 71b ± 0.05 97b ± 1.17 116a ± 2.53 118a ± 3.2 94ab ± 1.18 89b ± 2.56
Open in a new tab
a

All values are means ± SD by triplicate. a,b,c Different letters among columns of each insect indicate a statistically significant difference based on Tukey’s test p value = 0.05. Dw: dry weight. DD: degree of deacetylation expressed in (%), DM: demineralization reported in (%), and DP: deproteinization reported in (%).

Chitosan yields using T. molitor were 6.77% ± 0.38 (M1), 8.27% ± 0.60 (M2), and 8.87% ± 0.35 (M3) and S. purpurascens were 6.76% ± 0.33 (M1), 8.84% ± 0.14 (M2), and 8.70% ± 0.23 (M3), where M2 and M3 outperform. These results show that tougher deacetylation conditions (M2 and M3) significantly improve chitosan production compared to M1. In comparison, similar yields were reported for G. bimaculatus (8.40%)30 and Musca domestica (5.87%)21 but under harsher conditions (50% NaOH solution at 95 and 105 °C for 3 and 5 h). Also, the yield of chitosan from H. illucens pupal exuviae reached 8% (2 M NaOH for 4 h at 100 °C).26 These results fall within the range typically observed for chitosan from crustacean sources (7–15%).14,19 For example, Parthiban et al.34 reported higher chitosan yields of 15.40% ± 0.16 for white shrimp (Fenneropenaeus indicus), 13.25% ± 0.09 for mud crab (Scylla serrata), and 12.56% ± 0.14 for squilla (Squilla leptosquilla) but used a longer method (3% HCl at room temperature for 16 h, 4% NaOH at room temperature for 20 h, and 50% NaOH at 65 °C for 20 h). According to the observation by Younes and Rinaudo,35 chitosan yield can vary significantly depending on the source of chitin and the deacetylation method.

Demineralization (DM) is a key step in chitosan extraction, as it reduces the mineral content in the raw material. In this study, DM percentages varied significantly depending on the method used. For T. molitor, the values were 63.89% ± 0.31 (M1), 50.55% ± 0.14 (M2), and 46.85% ± 0.06 (M3), while S. purpurascens showed 86.81% ± 0.04 (M1), 63.18% ± 0.04 (M2), and 25.45% ± 0.12 (M3), all with statistically significant differences (p < 0.05). Notably, M1 included a DM step using 5% HCl at 40 °C for 2 h, while M2 and M3 underwent only deproteinization and deacetylation. Compared to earlier studies, Song et al.14 reported a 32.56% DM from T. molitor larval exuviae using 2 N HCl at 25 °C, while Kim et al.21 achieved 38.58% in M. domestica pupal shells under the same conditions. The insects analyzed in this study had low ash content (T. molitor: 5.4% ± 0.01, S. purpurascens: 4.4% ± 0.3), therefore, extensive demineralization may not always be necessary for insect-derived chitin or chitosan. Deproteinization (DP) involves the use of chemical methods to break down and remove proteins while preserving the chitin structure. Methods include alkaline treatment, where chitin is subjected to alkaline hydrolysis to solubilize proteins. The samples of exuviae from T. molitor showed ranges from 60 to 75%. Regarding samples of S. purpurascens, the results obtained ranged between 58 and 60%. The lower DP in S. purpurascens may be attributed to its higher protein content (57.6% ± 3.1), which could influence the efficiency of the deproteinization process compared to T. molitor exuviae (26.3% ± 0.4) under these milder conditions. Insects of the order Orthoptera as G. bimaculatus and Schistocerca gregaria have shown DP between 45 and 60% under 1.25 N NaOH at 95 °C for 3 h.21,30 Higher DPs are reported by several authors that worked with different insects under more severe conditions. For example, 94% and 97% are reported for H. illucens larvae and beetles26 (2 M NaOH for 2 h at 80 °C) and 86% for M. domestica(21) (1.25 N NaOH at 95 °C for 3 h). Certain conditions, such as the concentration of the alkali, the reaction time, and the raw material, are factors that influence the DP.36 The removal of proteins and minerals improves the characterization of chitosan by eliminating interferences from these compounds.36 Additionally, it can enhance its biocompatibility, biodegradability, antimicrobial activity, and other properties.

The degree of deacetylation (DD) is a fundamental parameter for determining the quality of chitosan. The DD depends on the source of chitin and the method used.5,25 Regarding the degree of deacetylation, the results for T. molitor were 42.62% ± 0.38 (M1), 25.25% ± 1.2 (M2), and 55.62% ± 0.79 (M3), while for S. purpurascens, they were 61.86% ± 4.98 (M1), 44.87% ± 1.38 (M2), and 43.77% ± 1.52 (M3). For T. molitor, M3 achieved the best result; it could be an effect of the conditions used in M3 that lead to a higher degree of deacetylation compared to the conditions in M1 and M2. The increased temperature and pressure in M3 could accelerate the deacetylation reaction, potentially resulting in a greater conversion of chitin to chitosan,14 whereas for S. purpurascens, the M1 resulted in the highest deacetylation that could be achieved by removing the mineral component, due to which the matrix becomes exposed, which enhances deacetylation.36 While some authors reported higher degrees of DD under more severe conditions, for example, 83–94% from exoskeletons of G. bimaculatus(30) and larval exuviae and exoskeletons of H. illucens(26) using harsher conditions, others reported 47%14 from the exuviae of T. molitor using similar conditions.

3.2. Physicochemical Properties of Chitosan Obtained

In addition, in this work, the percentage of solubility obtained for chitosan from the exuviae of T. molitor was M1: 12.74% ± 0.82, M2: 9.81% ± 0.81, and M3: 24.13% ± 2.07 and S. purpurascens was M1 27.31% ± 1.87, M2 13.64% ± 1.37, and M3 27.11% ± 1.21. The samples M3 from T. molitor exuviae exhibits higher solubility compared to M1 and M2 (p < 0.05), meaning that the conditions of samples M3 favored production of chitosan with higher solubility in comparison with M1 and M2. This result is advantageous for antimicrobial solutions, drug delivery systems, or coagulation of contaminants, where chitosan needs to dissolve effectively. In the case of lower solubility, such as that observed in M2 (9.81%) is beneficial for applications requiring structural stability, such as membranes or filters.37 Although the samples from S. purpurascens M1 and M3 exhibit similar solubility, a higher yield of chitosan was obtained in M3. The differences could be attributed to variations in deacetylation conditions, such as NaOH concentration, reaction time, or temperature, which affected the structure of the produced chitosan.38 Besides, the samples of grasshopper S. purpurascens generally presented higher solubility percentages compared to the exuvia from T. molitor; the chitin from the grasshopper is more susceptible to the deacetylation process due to its molecular arrangement or less compact crystalline structure.39 Additionally, the higher protein and fat content in S. purpurascens may contribute to a less rigid structure, while the carbohydrate-rich composition of T. molitor exuviae could lead to a more compact and crystalline arrangement, reducing reagent penetration and process efficiency.13 Other authors have reported higher results of solubility, for example, 94% for chitosan from grasshoppers (demineralization: 1 M HCl at 30 °C and 200 rpm for 2 h, deproteinization: 1 M NaOH at 90 °C for 2 h, and deacetylation: 60 wt % NaOH at 100 °C for 8 h at 200 rpm).25 Solubility of chitosan derived from insects is largely influenced by the deacetylation method employed and the structure of chitin.40

The molecular weight was calculated by the Mark–Houwink–Sakurada equation, and the results are found in Table 2. Chitosan with low molecular weight oscillates between 10 kDa and 50 kDa, with medium molecular weight 50 kDa and 150 kDa, and with high molecular weight >150 kDa.5 The exuviae samples from T. molitor exhibited molecular weights of 71 ± 0.05 kDa (M1), 97 ± 1.17 kDa (M2), and 116 ± 2.53 kDa (M3), corresponding to medium molecular weights. M1 exhibited the lowest molecular weight in comparison with M2 and M3, possibly due to the demineralization process being more aggressive or pronounced, which could have led to the depolymerization of the chitosan chain.22 In contrast, M3 had the highest molecular weight, suggesting that the processing conditions for M3 allowed a better preservation of the chitosan structure. In the same way, the molecular weight for samples from S. purpurascens was reported for M1 118 ± 3.2 kDa, M2 94 ± 1.18 kDa, and M3 89 ± 2.56 kDa being chitosans of medium molecular weight furthermore to present statistical differences (p < 0.05). M2 and M3 are comparable and lower than M1, despite undergoing a demineralization process; M1 apparently managed to preserve the polymer chain of chitosan. Authors have reported chitosan with variable molecular weights obtained from different insect species and conditions. For instance, a low molecular weight of chitosan (16 kDa) was obtained from Periplaneta americana that was subjected to more severe conditions.41 Psarianos et al.42 obtained chitosan from Acheta domesticus with a molecular weight of 103.4 ± 2.4 kDa and employed 1 M HCl during 2 h at 98 °C, 1 M NaOH (1:50 w/v) for 24 h at 80 °C, and 50% NaOH, 3 h at 130 °C. In contrast, high molecular weight chitosans were obtained from other cricket species, such as Gryllodes sigillatus with 524 kDa43 applying less aggressive conditions. The variations observed in molecular weight suggest that the extraction process or the deacetylation treatment had a direct impact on the integrity of the chitosan. More aggressive or prolonged processes may be fragmenting the chitosan chains, resulting in lower molecular weights. In summary, the degree of deacetylation (DD), molecular weight (MW), and solubility are highly influenced by the extraction process, as well as by the raw material. Depending on this, it suggests different applications, for instance, in applications requiring enhanced bioavailability, such as antimicrobial coatings28 or water treatment as coagulants.10 Hence, M3 from T. molitor and S. purpurascens presented the highest yield in chitosan production.

3.3. Fourier-Transformed Infrared Spectroscopy

Chitosan standards of low (STD 1) and medium (STD 2) molecular weight were analyzed to compare their spectrum with the data from this study. The spectrum displayed characteristic bands at 3300–3200 cm–1 (O–H and N–H groups), 1650 cm–1 (amide I), and 1060 cm–1 (carbohydrates) (Figure 1). Figure 2 presents the FTIR spectra of chitosan from T. molitor exuviae, highlighting characteristic functional groups. Signals at 3000–2800 cm–1 correspond to C–H stretching, while bands at 1650 cm–1 and 1550 cm–1 represent amide I (C=O stretching) and amide II (N–H bending, C–N stretching). The fingerprint region (1500–1200 cm–1) confirms amine and amide groups, and signals at 1200–900 cm–1 indicate C–O–C and C–O–H bonds in the glucosamine ring. Notably, 2910 cm–1 and 2848 cm–1 confirm the polysaccharide nature of chitosan, while 1030–1020 cm–1 and 990–890 cm–1 suggest polymer ring vibrations. Unique signals at 1469 cm–1 and 1656 cm–1 indicate methyl/methylene groups and hydrogen bonding, respectively. Figure 3 displays the spectra of chitosan from S. purpurascens, showing consistent signals at 3262–3270 cm–1 (N–H and O–H bonds), amide II at 1546–1553 cm–1, and C–O vibrations at 1000–1065 cm–1. Amide I at 1614–1622 cm–1 was present in all samples, with slight variations likely due to impurity removal. A signal at 1371–1376 cm–1 suggests partial deacetylation. In general, all the samples presented the characteristic functional groups of chitosan. These findings confirm structural similarities across samples despite extraction method variations and raw materials, with FTIR characterization verifying preserved chitosan functional groups.

Figure 1.

Figure 1

Open in a new tab

Spectrograms of the chitosan STD 1: low molecular weight standard and STD 2: medium molecular weight as a function of the wavelength (cm–1). The main absorption bands are observed around 3300 cm–1 (O–H and N–H bonds), 1650 cm–1 (C=O of amide I), and 1080 cm–1 (C–O of hydroxyl groups).

Figure 2.

Figure 2

Open in a new tab

Spectrograms of chitosan samples from Tenebrio molitor exuviae, labeled based on extraction methods (A: method 1, B: method 2, and C: method 3). Key functional groups include C–H stretching (3000–2800 cm–1), acetyl (−C=O–CH3), amide I, II, and III (1650–1580 cm–1), N–H flexion, and polysaccharide bands (1000–800 cm–1). Spectrograms were created using OriginPro 2024 (OriginLab, USA).

Figure 3.

Figure 3

Open in a new tab

Spectrograms of Sphenarium purpurascens chitosan samples, labeled by extraction methods (A: method 1, B: method 2, and C: method 3), show key functional groups: C–H stretching (3000–2800 cm–1), acetyl (−C=O–CH3), amide I, II, and III (1650–1580 cm–1), N–H flexion, and polysaccharide bands (1000–800 cm–1). Spectrograms were created using OriginPro 2024 (OriginLab, USA).

3.4. Swelling Degree

The results of the swelling test are shown in Table 3 for exuviae of T. molitor and S. purpurascens. The SD obtained for the chitosan samples from exuvia of T. molitor indicates significant variability in water retention capacity, which is directly related to the structural properties and integrity of the chitosan. M1 (1.5%) presented SD 5.07% ± 0.11; this relatively low value suggests that sample M1 has a denser or more compact structure, which limits its ability to absorb water. On the other hand, M1 (1%) had 7.77% ± 0.76 and it exhibits a SD greater than that of M1, indicating a higher capacity for water absorption. This may reflect a more porous or less cross-linked structure, allowing water to penetrate and be retained within the chitosan matrix.44 This behavior showed a balance between structural stability and swelling capacity. Finally, M2 (1%) presented a SD of 13.3% ± 1.2; this value suggests that it has an open or porous structure, allowing for significant water absorption. Mechanical properties observed in M1 suggest that it had a denser and less porous structure, which is preferable for applications requiring stability in aqueous environments, such as the development of filtration membranes. For the samples of S. purpurascens were obtained the following results: M1 (1.5%) and M1 (1%) have similar swelling values, 3.86% ± 0.4 and 3.71% ± 1.9, respectively, and showed consistency in the structure of the chitosan derived from these films. These values indicate that the films have a moderate capacity for water absorption, likely due to a balance between porosity and density.9 In contrast, M2 (1%) showed the highest SD 4.75% ± 0.06, which could show greater porosity or a less cross-linked structure, allowing for slightly higher water absorption. Lastly, M3 (1%) had the lowest SD of 2.75% ± 0.32, showing a more compact structure that limits the film’s ability to absorb water. This may be associated with a higher degree of cross-linking or lower porosity compared to the other samples. Chitosan films from H. illucens(26) have shown uniformity, whereas films from T. molitor exuviae and S. purpurascens were heterogeneous, and, despite this, their lower swelling capacity and slower degradability make them suitable for filter design. A low swelling degree may indicate that the material has greater hydrophobicity, which can be beneficial for applications that require higher resistance to water and moisture.45

Table 3. Swelling Degree (SD) Result and Loss Weight Percentage (LW)a.

  exuviae T. molitor
 S. purpurascens
  M1 (1.5%) M1 (1%) M2 (1%) M3 (1%) M1 (1.5%) M1 (1%) M2 (1%) M3 (1%)
% SD 5.07a ± 0.11 7.77a ± 0.76 13.3b ± 1.2 * 3.86a ± 0.4 3.71a ± 1.9 4.75a ± 0.06 2.75a ± 0.32
% LW 2.46a ± 0.43 4.88a ± 0.41 * * 4.61b ± 1.45 4.79b ± 0.97 12.3a ± 2.35 15.7a ± 28
Open in a new tab
a

All values are means ± SD by triplicate. a,b Different letters among columns indicate a statistically significant difference based on Tukey’s test.

3.5. Antimicrobial Activity Test

A more significant reduction in microbial cells was observed, notably affecting the reduction in Gram-negative bacteria E. coli, E. aerogenes, P. aeruginosa, and S. spp. Chitosan exhibited notable antimicrobial activity after 48 h against the evaluated microorganisms, as can be observed in the results of Figure 4 from exuviae of T. molitor and Figure 5 for S. purpurascens. However, this study identified a significant exception with S. aureus, where the antimicrobial activity was less effective at 24 h. While both types of chitosan exhibited antimicrobial efficacy, the chitosan from S. purpurascens demonstrated a more consistent and higher performance in inhibiting S. aureus, especially at 48 h, compared to the chitosan from T. molitor. This behavior could be attributed to the medium molecular weight of chitosan. According to the literature, a lower molecular weight chitosan may penetrate the cell structure more effectively,46 thus increasing the antimicrobial activity. Then, this reduced efficacy could be attributed, also, to the cell membrane due to the robust structure formed by the peptidoglycan layer in S. aureus that is denser compared to Gram-negative bacteria, which could hinder the effective interaction of chitosan with the cell membrane.47 Studies on various microorganisms show that lower molecular weight chitosan is more effective in inhibiting microbial growth and multiplication having bactericide and bacteriostatic effects.46 These results underscore that while medium molecular weight chitosan performs well against Gram-negative pathogens, factors such as degree of deacetylation which enhance cellular interaction are critical for maximizing antimicrobial activity. Overall, optimizing these parameters is essential for tailoring chitosan for specific antimicrobial applications.

Figure 4.

Figure 4

Open in a new tab

Results of percentages of the inhibition test of chitosan from exuviae of Tenebrio molitor at 24 h and 48 h. All values are means ± SD by triplicate. a,b,c,d,e Different letters among bars indicate a statistically significant difference based on Tukey’s test.

Figure 5.

Figure 5

Open in a new tab

Results obtained of the antimicrobial test in chitosan from Sphenarium purpurascens. All values are means ± SD by triplicate. a,b,c,d Different letters among columns indicate a statistically significant difference based on Tukey’s test.

4. Conclusions

Chitosan obtained under milder processing from edible insects such as T. molitor and S. purpurascens represents a promising alternative to crustacean-derived chitosan. The extraction methods played a critical role in determining the final quality of the product, as evidenced by our results, which demonstrated that the molecular weight, solubility, swelling degree, and degree of deacetylation of insect-derived chitosan had the quality to form films which could have industrial and biomedical applications. The findings provide a foundation for future studies to explore the utilization of insect waste as a valuable resource.

Author Contributions

The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript.

This research was supported by the Secretariat of Science, Humanities, Technology and Innovation (SECIHTI) through the scholarship 769152; the authors acknowledge to SECIHTI, the Department of Bioengineering from Tecnologico de Monterrey, Campus Puebla, and the Institute of Design and Technological Innovation from IBERO Puebla.

The authors declare no competing financial interest.

References

  1. Encyclopedia of Polymer Science and Technology; John Wiley & Sons, Inc., 2002. [Google Scholar]
  2. Udayakumar G. P.; Muthusamy S.; Selvaganesh B.; Sivarajasekar N.; Rambabu K.; Banat F.; Sivamani S.; Sivakumar N.; Hosseini-Bandegharaei A.; Show P. L. Biopolymers and Composites: Properties, Characterization and Their Applications in Food, Medical and Pharmaceutical Industries. J. Environ. Chem. Eng. 2021, 9 (4), 105322. 10.1016/j.jece.2021.105322. [DOI] [Google Scholar]
  3. Kashyap P. L.; Xiang X.; Heiden P. Chitosan Nanoparticle Based Delivery Systems for Sustainable Agriculture. Int. J. Biol. Macromol. 2015, 77, 36–51. 10.1016/j.ijbiomac.2015.02.039. [DOI] [PubMed] [Google Scholar]
  4. Rahman M. Z.; Rahman M.; Mahbub T.; Ashiquzzaman M.; Sagadevan S.; Hoque M. E. Advanced Biopolymers for Automobile and Aviation Engineering Applications. J. Polym. Res. 2023, 30 (3), 106. 10.1007/s10965-023-03440-z. [DOI] [Google Scholar]
  5. Rinaudo M. Chitin and Chitosan: Properties and Applications. Prog. Polym. Sci. 2006, 31 (7), 603–632. 10.1016/j.progpolymsci.2006.06.001. [DOI] [Google Scholar]
  6. Expert Market Research . Global Chitosan Market Report and Forecast 2023–2028, 2023. https://www.expertmarketresearch.com/reports/chitosan-market.
  7. Dhillon G. S.; Kaur S.; Brar S. K. Facile Fabrication and Characterization of Chitosan-Based Zinc Oxide Nanoparticles and Evaluation of Their Antimicrobial and Antibiofilm Activity. Int. Nano Lett. 2014, 4 (2), 107. 10.1007/s40089-014-0107-6. [DOI] [Google Scholar]
  8. Vallejo-Domínguez D.; Rubio-Rosas E.; Aguila-Almanza E.; Hernández-Cocoletzi H.; Ramos-Cassellis M. E.; Luna-Guevara M. L.; Rambabu K.; Manickam S.; Siti Halimatul Munawaroh H.; Loke Show P. Ultrasound in the Deproteinization Process for Chitin and Chitosan Production. Ultrason. Sonochem. 2021, 72, 105417. 10.1016/j.ultsonch.2020.105417. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Wang L.; Dong Y.; Men H.; Tong J.; Zhou J. Preparation and Characterization of Active Films Based on Chitosan Incorporated Tea Polyphenols. Food Hydrocolloids 2013, 32 (1), 35–41. 10.1016/j.foodhyd.2012.11.034. [DOI] [Google Scholar]
  10. Zubair M.; Arshad M.; Ullah A.. Chitosan-Based Materials for Water and Wastewater Treatment. In Handbook of Chitin and Chitosan; Elsevier, 2020; pp 773–809. [Google Scholar]
  11. Abdullah H. A.; Jaeel A. J. Turbidity, Color and Chemical Oxygen Demand Removals from Synthetic Textile Wastewater Using Chitosan as a Coagulant. IOP Conf. Ser.: Mater. Sci. Eng. 2019, 584 (1), 012016. 10.1088/1757-899X/584/1/012016. [DOI] [Google Scholar]
  12. Mishyna M.; Glumac M. So Different, yet so Alike Pancrustacea: Health Benefits of Insects and Shrimps. J. Funct.Foods 2021, 76, 104316. 10.1016/j.jff.2020.104316. [DOI] [Google Scholar]
  13. Jantzen da Silva Lucas A.; Quadro Oreste E.; Leão Gouveia Costa H.; Martín López H.; Dias Medeiros Saad C.; Prentice C. Extraction Physicochemical Characterization, and Morphological Properties of Chitin and Chitosan from Cuticles of Edible Insects. Food Chem. 2021, 343, 128550. 10.1016/j.foodchem.2020.128550. [DOI] [PubMed] [Google Scholar]
  14. Song Y.-S.; Kim M.-W.; Moon C.; Seo D.-J.; Han Y. S.; Jo Y. H.; Noh M. Y.; Park Y.-K.; Kim S.-A.; Kim Y. W.; Jung W.-J. Extraction of Chitin and Chitosan from Larval Exuvium and Whole Body of Edible Mealworm, Tenebrio Molitor. Entomol. Res. 2018, 48 (3), 227–233. 10.1111/1748-5967.12304. [DOI] [Google Scholar]
  15. Meyer-Rochow V. B.; Gahukar R. T.; Ghosh S.; Jung C. Chemical Composition, Nutrient Quality and Acceptability of Edible Insects Are Affected by Species, Developmental Stage, Gender, Diet, and Processing Method. Foods 2021, 10 (5), 1036. 10.3390/foods10051036. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Mwita C. S.; Muhammad R.; Nettey-Oppong E. E.; Enkhbayar D.; Ali A.; Ahn J.; Kim S.-W.; Seok Y.-S.; Choi S. H. Chitosan Extracted from the Biomass of Tenebrio Molitor Larvae as a Sustainable Packaging Film. Materials 2024, 17 (15), 3670. 10.3390/ma17153670. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Ibarra-Herrera C. C.; Acosta-Estrada B.; Chuck-Hernández C.; Serrano-Sandoval S. N.; Guardado-Félix D.; Pérez-Carrillo E. Nutritional Content of Edible Grasshopper (Sphenarium Purpurascens) Fed on Alfalfa (Medicago Sativa) and Maize (Zea Mays). CyTA--J. Food 2020, 18 (1), 257–263. 10.1080/19476337.2020.1746833. [DOI] [Google Scholar]
  18. Marín-Morales M. S.; Ibarra-Herrera C. C.; Luna-Vital D. A.; Monribot-Villanueva J. L.; Guerrero-Analco J. A. Biological Activity of Extracts and Hydrolysates from Early- and Adult-Stage Edible Grasshopper Sphenarium Purpurascens. Front. Nutr. 2022, 9, 1028543. 10.3389/fnut.2022.1028543. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Pakizeh M.; Moradi A.; Ghassemi T. Chemical Extraction and Modification of Chitin and Chitosan from Shrimp Shells. Eur. Polym. J. 2021, 159, 110709. 10.1016/j.eurpolymj.2021.110709. [DOI] [Google Scholar]
  20. Mohan K.; Ganesan A. R.; Ezhilarasi P. N.; Kondamareddy K. K.; Rajan D. K.; Sathishkumar P.; Rajarajeswaran J.; Conterno L. Green and Eco-Friendly Approaches for the Extraction of Chitin and Chitosan: A Review. Carbohydr. Polym. 2022, 287, 119349. 10.1016/j.carbpol.2022.119349. [DOI] [PubMed] [Google Scholar]
  21. Kim M.-W.; Han Y. S.; Jo Y. H.; Choi M. H.; Kang S. H.; Kim S.-A.; Jung W.-J. Extraction of Chitin and Chitosan from Housefly, Musca Domestica, Pupa Shells. Entomol. Res. 2016, 46 (5), 324–328. 10.1111/1748-5967.12175. [DOI] [Google Scholar]
  22. No H. K.; Nah J. W.; Meyers S. P. Effect of Time/Temperature Treatment Parameters on Depolymerization of Chitosan. J. Appl. Polym. Sci. 2003, 87 (12), 1890–1894. 10.1002/app.11628. [DOI] [Google Scholar]
  23. DuBois M.; Gilles K. A.; Hamilton J. K.; Rebers P. A.; Smith F. Colorimetric Method for Determination of Sugars and Related Substances. Anal. Chem. 1956, 28 (3), 350–356. 10.1021/ac60111a017. [DOI] [Google Scholar]
  24. Reyes V. Producción de quitosano a partir de desechos de camarón generados del procesamiento industrial. Res. Dev. Food Sci. Technol. 2019, 4, 897–901. [Google Scholar]
  25. Luo Q.; Wang Y.; Han Q.; Ji L.; Zhang H.; Fei Z.; Wang Y. Comparison of the Physicochemical, Rheological, and Morphologic Properties of Chitosan from Four Insects. Carbohydr. Polym. 2019, 209, 266–275. 10.1016/j.carbpol.2019.01.030. [DOI] [PubMed] [Google Scholar]
  26. Triunfo M.; Tafi E.; Guarnieri A.; Salvia R.; Scieuzo C.; Hahn T.; Zibek S.; Gagliardini A.; Panariello L.; Coltelli M. B.; De Bonis A.; Falabella P. Characterization of Chitin and Chitosan Derived from Hermetia Illucens, a Further Step in a Circular Economy Process. Sci. Rep. 2022, 12 (1), 6613. 10.1038/s41598-022-10423-5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Lavorgna M.; Piscitelli F.; Mangiacapra P.; Buonocore G. G. Study of the Combined Effect of Both Clay and Glycerol Plasticizer on the Properties of Chitosan Films. Carbohydr. Polym. 2010, 82 (2), 291–298. 10.1016/j.carbpol.2010.04.054. [DOI] [Google Scholar]
  28. Hongpattarakere T.; Riyaphan O. Effect of Deacetylation Conditions on Antimicrobial Activity of Chitosans Prepared from Carapace of Black Tiger Shrimp (Penaeus Monodon). Songklanakarin J. Sci. Technol. 2008, 30, 1–9. [Google Scholar]
  29. Purkayastha D.; Sarkar S. Physicochemical Structure Analysis of Chitin Extracted from Pupa Exuviae and Dead Imago of Wild Black Soldier Fly (Hermetia Illucens). J. Polym. Environ. 2020, 28 (2), 445–457. 10.1007/s10924-019-01620-x. [DOI] [Google Scholar]
  30. Kim M.-W.; Song Y.-S.; Han Y. S.; Jo Y. H.; Choi M. H.; Park Y.-K.; Kang S. H.; Kim S.-A.; Choi C.; Jung W.-J. Production of Chitin and Chitosan from the Exoskeleton of Adult Two-Spotted Field Crickets (Gryllus Bimaculatus). Entomol. Res. 2017, 47 (5), 279–285. 10.1111/1748-5967.12239. [DOI] [Google Scholar]
  31. Kleden Y.; Mukkun L.; Ndapamuri M.; Pramatana F. The Migratory Locust (Locusta Migratoria) as a Potential Source of Protein and Biopolymer Compounds in the Future. Biodiversitas 2023, 24, 5979–5987. 10.13057/biodiv/d241117. [DOI] [Google Scholar]
  32. Parthiban F.; Balasundari S.; Gopalakannan A.; Rathnakumar K.; Felix S. Comparison of the Quality of Chitin and Chitosan from Shrimp, Crab and Squilla Waste. Curr. World Environ. 2017, 12 (3), 670–677. 10.12944/CWE.12.3.18. [DOI] [Google Scholar]
  33. Younes I.; Rinaudo M. Chitin and Chitosan Preparation from Marine Sources. Structure, Properties and Applications. Mar. Drugs 2015, 13 (3), 1133–1174. 10.3390/md13031133. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Rasweefali M. K.; Sabu S.; Muhammed Azad K. S.; Raseel Rahman M. K.; Sunooj K. V.; Sasidharan A.; Anoop K. K. Influence of Deproteinization and Demineralization Process Sequences on the Physicochemical and Structural Characteristics of Chitin Isolated from Deep-Sea Mud Shrimp (Solenocera Hextii). Adv. Biomarker Sci. Technol. 2022, 4, 12–27. 10.1016/j.abst.2022.03.001. [DOI] [Google Scholar]
  35. Chitin and Chitosan: Properties and Applications, 1st ed.; Broek L. A. M., Boeriu C. G., Eds.; Wiley, 2019. [Google Scholar]
  36. Braspaiboon S.; Osiriphun S.; Peepathum P.; Jirarattanarangsri W. Comparison of the Effectiveness of Alkaline and Enzymatic Extraction and the Solubility of Proteins Extracted from Carbohydrate-Digested Rice. Heliyon 2020, 6 (11), e05403 10.1016/j.heliyon.2020.e05403. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Kaya M.; Erdogan S.; Mol A.; Baran T. Comparison of Chitin Structures Isolated from Seven Orthoptera Species. Int. J. Biol. Macromol. 2015, 72, 797–805. 10.1016/j.ijbiomac.2014.09.034. [DOI] [PubMed] [Google Scholar]
  38. Hahn T.; Roth A.; Ji R.; Schmitt E.; Zibek S. Chitosan Production with Larval Exoskeletons Derived from the Insect Protein Production. J. Biotechnol. 2020, 310, 62–67. 10.1016/j.jbiotec.2019.12.015. [DOI] [PubMed] [Google Scholar]
  39. Chen S.; Wei X.; Sui Z.; Guo M.; Geng J.; Xiao J.; Huang D. Preparation of Antioxidant and Antibacterial Chitosan Film from Periplaneta Americana. Insects 2021, 12 (1), 53. 10.3390/insects12010053. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Psarianos M.; Dimopoulos G.; Ojha S.; Cavini A. C. M.; Bußler S.; Taoukis P.; Schlüter O. K. Effect of Pulsed Electric Fields on Cricket (Acheta Domesticus) Flour: Extraction Yield (Protein, Fat and Chitin) and Techno-Functional Properties. Innov. Food Sci. Emerg. Technol. 2022, 76, 102908. 10.1016/j.ifset.2021.102908. [DOI] [Google Scholar]
  41. Malm M.; Liceaga A. M. Physicochemical Properties of Chitosan from Two Commonly Reared Edible Cricket Species, and Its Application as a Hypolipidemic and Antimicrobial Agent. Polysaccharides 2021, 2 (2), 339–353. 10.3390/polysaccharides2020022. [DOI] [Google Scholar]
  42. Leceta I.; Guerrero P.; de la Caba K. Functional Properties of Chitosan-Based Films. Carbohydr. Polym. 2013, 93 (1), 339–346. 10.1016/j.carbpol.2012.04.031. [DOI] [PubMed] [Google Scholar]
  43. Ortiz-García T.; Rapado-Paneque M.; Peniche-Covas C. Hidrogeles superabsorbentes basados en poliacrilamida para aplicación agrícola: estudio de hinchamiento. Rev. Cubana Quím. 2021, 33 (2), 46–68. [Google Scholar]
  44. Tsai G.-J.; Zhang S.-L.; Shieh P.-L. Antimicrobial Activity of a Low-Molecular-Weight Chitosan Obtained from Cellulase Digestion of Chitosan. J. Food Prot. 2004, 67 (2), 396–398. 10.4315/0362-028X-67.2.396. [DOI] [PubMed] [Google Scholar]
  45. Helander I. M.; Nurmiaho-Lassila E. L.; Ahvenainen R.; Rhoades J.; Roller S. Chitosan Disrupts the Barrier Properties of the Outer Membrane of Gram-Negative Bacteria. Int. J. Food Microbiol. 2001, 71 (2–3), 235–244. 10.1016/S0168-1605(01)00609-2. [DOI] [PubMed] [Google Scholar]

Articles from ACS Omega are provided here courtesy of American Chemical Society

RESOURCES