Yield and moisture of chitosan
Chitosan yield from A. anntenus in this research is relatively lower where it was 18%. This was higher than that reported by Brzeski and Sauer48 (14% yield of chitosan from shrimp), which may be due to depolymerization of the chitosan polymer, loss of sample mass/weight from excessive removal of acetyl groups from the polymer during deacetylation, and loss of chitosan particles during washing.
It almost exactly matches William and Wid’s35 findings, who found that the average amount of chitosan produced from the wet weight of shrimp shell waste was 19.01%. Because the natural structure of chitin (a protein) was lost during the deproteination process, less chitosan was generated when extraction was started with deproteinization. The weight of the chitosan was ultimately reduced by the prior process, which led to increased hydrolysis and losses of solid material. Moisture content in this study the shrimp shell chitosan samples had a moisture content of 7.65 ± 0.13%. The produced chitosan’s moisture content was 7.77 ± 0.15%, compared to 7.50 ± 0.10% for commercial chitosan. This discovery is consistent with other studies’ findings. According to Li et al.49, commercial chitosan can include more than 10% moisture. According to Rege et al.50, the moisture content of chitosan powder ranged from 7 to 11% (w/w) and had nothing to do with its DD or MW. On the other hand, Mucha et al.51 found that DD had a favorable impact on moisture content. The ability to absorb water will decrease as the DD increases.
Degree of deacetylation of chitosan (DD%)
Numerous physiochemical and organic properties of chitosan, including crystallinity, hydrophilicity, degradation, and cell response, are determined by the degree of deacetylation (DD), which is usually recognized as an essential metric. According to the degree of d-glucosamine and N-acetyl d-glucosamine deacetylation, the biopolymer is classified as either chitin or chitosan. Various methods have been described for guaranteeing the degree of chitosan deacetylation52. The DD of the extracted chitosan from the current study was found to be 86%, which is very similar to Puvvada et al.53 DD of 89.79% and their conclusion that higher DD values were brought on by the presence of more protein, which affects the chemical, physical, and biological properties of chitosan, such as adsorption, covalent linking, and encapsulation. While Younes and Rinaudo54 have recorded that 70.9% and it’s the lowest value for characteristic of chitin; (DD) value depends on the raw material and the processes used for the deproteinization and the demineralization. Additionally, Muñoz et al.55 found that 73.6% of the chitin was deacetylated and noted that the process of DD is dependent on the type of chitin utilized as well as the time, temperature, and alkaline concentration.
Solubility of extracted chitosan
In contrast to the 99% reported for commercial chitosan by Hossain and Iqbal56, the solubility from extracted chitosan in this work was 98.15%. To achieve the requisite solubility, deacetylation is thought to need to be at least 85%. Chitosan solubility increased correspondingly as the degree of deacetylation increased; samples treated with 50 and 60% NaOH yielded solubility ranging from 96.01 to 97.2%, respectively. The degree of deacetylation affects both the solubility properties of chitosan. A high degree of deacetylation appears to have higher solvency, and a degree of deacetylation appears to destitute solubility57,58. Moreover, the partial removal of the acetyl group and protein gives lower solubility values of chitosan solubility in inorganic acid56, due to the highly protonated free amino group that attracts ionic compounds32.
FTIR spectrum of extracted and standard chitosan
The FTIR analysis was used to assess the functional groups present in both extracted and standard chitosan. In Fig. 1A (See them separately in Supplementary data file Figs. 1S–4S), the IR of extracted chitosan can be observed where a strong band in the region 2920.5, 2845.3 and 1463 cm−1 corresponds to OH, NH and NH2 stretching chemical bond, as well as the intramolecular hydrogen bonds. The weak bands presented 2660, 2663, 2509, 2016, and 1425 cm−1 due to C–H, H–C–H and C=O stretching were also detected. According to Kumari et al.37, the methyl group in NHCOCH3, the methylene group in CH2OH, and the methane group in the pyranose ring all exhibit stretching vibrations in the range of 2921–2879 cm−1. This was nearly range with our study peaks at 2920.5 and 2845.3 cm−1. The absorption bands at around 1076, 719.8 cm−1 were medium peaks N–H, C–O, and C–O–C groups, while the characteristic peaks assignment of the extracted chitosan at 3602 and 1799 cm−1 was attributed to the stretching vibration of OH chemical bond, also the peak was showed at 492.6 cm−1 may be due to Si–O–Si asymmetrical and symmetrical stretching. On the other hand, the FTIR spectrum analysis of standard chitosan was a broad strongest band at 3379 and 1959 cm−1 corresponding to the stretching of OH groups in Fig. 1B 59,60,61,62.
FTIR spectra of extracted chitosan (A) and standard chitosan (B).
The peak at 2880 cm−1 is related to N–H in NH2 association in primary amines due to different stretching vibration bands. The peak at 1378 cm–1 is due to C–C stretching. However, the observation in this spectrum study is identical to what was seen by Muñoz et al.55, even though the absorption bands at 1115 cm–1 correspond to the vibration of the C-N bond and confirm the joining of the monomers through α-glycosidic links.
Biofilms formation and characterization
As mentioned before, the biofilms were prepared according to the formulation presented in Table 1. The greatest casted biofilms with perfect strength were selected for further analyses. In general, Fig. 2 represents a side of different products from biofilms based on chitosan, which saved the highest harmony and homogeneity for films and showed flexibility. The most acceptable biofilms were the formulations of; Chitosan (90%)/Starch (5%)/Castor (5%) and Chitosan (70%)/Starch (10%)/Castor (20%) in comparison to extracted chitosan film. Besides that, the polypropylene (PP) film was taken into consideration. Indeed, there was no clear reason for such a result. Coupling chitosan with renewable and abundant agro-resources such as starch can be utilized to improve moisture resistance accelerate degradation, and amazingly reduce the manufacturing cost63. Generally, the properties of chitosan movies are influenced by a few components such as atomic mass and degree of deacetylation of chitosan, as well as the nearness or nonappearance of plasticizers64. However, one of the polymers that has been thoroughly investigated for application as a biodegradable plastic is biofilm starch-chitosan.
A side of the most acceptable biofilms synthesized from chitosan, starch, and castor.
FTIR of synthesized bioplastic films
Figure 3 shows several FTIR spectra of the synthetic bioplastic sheets. The absorption peaks at 3280 cm–1 and 2924.3 cm–1 in the spectra of the chitosan film alone are caused by the overlap of NH2 and OH65,66,67,68. The N–H band is responsible for the band absorption band at 1650.4 cm–1. The presence of the OH group is indicated by the absorption peak at 1417.69 cm–165. There are C-N group attributed to the amino group at an absorption peak of 1313.99 cm–169. The absorption peaks at 1153.4 and 1029.2 cm–1 indicate the existence of the C–O group65. Occurring at 2323.2 cm−1 and 2111.49 cm–1 are ascribed to the O–H stretching mode in chitosan indicating the existence of the acid group. The absorption band at 1793.79 cm−1 is due to the C=O group65.
Different FTIR spectra of; chitosan (70%)/starch (10%)/castor (20%) (a), polypropylene (as a petroleum synthetic plastic) (b), extracted chitosan (c), and castor chitosan (90%)/starch (5%)/castor (5%) (d).
Additionally, data shows the FTIR spectrum of chitosan (70%)/starch (10%)/castor (20%), which was characterized by the presence of amide I, amide II, and amide III bands near 1640.98, 1586.08, and 1421.6 cm–1, respectively. The presence of amide can be confirmed by the N–H stretching band near 3357.46 cm–1. On the other hand, carbohydrates can be determined by the presence of broad C–O bands near 1148.83 and 1071.05 cm–1. There are O–H stretching groups in the region of 2916.54 cm–1 indicating the existence of the acid group. Also, there are C–N group attributed to the amino group at an absorption peak of 1375.24 cm–169. The peak was shown at 655.7, 562.3 and 515.5 cm–1 may be due to Si–O–Si asymmetrical and symmetrical stretching70,71,72,73.
Moreover, the data explains the FTIR spectrum of chitosan (90%)/starch (5%)/castor (5%), which was characterized by the presence of amide I, amide II, and amide III bands near 1623.92, 1552.91, and 1423.85 cm–1, respectively. In addition, the N–H stretching band between 3432.36 and 3257.06 cm–1 confirms the presence of amide. On the other hand, the presence of large C–O bands between 1066.04 and 1014.47 cm–1, as well as the O–H group at 3257.06 cm–1, can be used to identify carbohydrates65. It is seen that the O–H group at 3101.30 cm–1 broadens. There are O–H stretching groups in the region of 2879.12 cm–1 indicating the existence of the acid group. The FTIR spectrum shows a broad change in the region of 3257.06 cm–1. This is due to the Maillard reaction within the castor oil where the ester bond underwent a reaction with the carbohydrate74. Also, there are C–N groups attributed to the amino group at absorption peaks of 1373.57, 1310.55, 1257.88, 1205.97, 1152.85, and 111.86 cm–169. The absorption bands at around 955.70, 899.39 and 746.33 cm–1 were medium peaks N–H, C–O, and C–O–C groups. The peak was shown at 695.38, 561.91 and 528.33 cm–1 may be due to Si–O–Si asymmetrical and symmetrical stretching. The peaks on 1623 and 1374 cm–1 are included in the unique spectrum of PP featuring CH2 deformation and symmetric CH3 deformation, while the peaks on 3429 and 3256 cm–1 are CH stretch spectra. The spectra on 559 and 527 cm–1, on the other hand, indicate an isotactic polypropylene band. Finally, the peaks 1555 and 1412.42 cm–1 spectrum suggest C–C bending, which is the backbone of PP.
Swelling capacity of produced biofilm films
Chitosan is hydrophobic, and the swelling behaviors of unplasticized films formed from extracted chitosans are soluble in glacial acetic acid 99%. However, Fig. 4 represents the films of extracted chitosan, chitosan (90%)/starch (5%)/castor (5%), chitosan (70%)/starch (10%)/castor (20%) and PP from 0 to 20 min in the water. Different synthesized biofilms exhibited a high degree of swelling ranging from 0.0 to 12%. It was observed that the degree of swelling of the chitosan (70%)/starch (10%)/castor (20%) had the highest percentage presented at 20 min and reached 12%, while the lowest percentage of the degree of swelling was in the PP had the same time stable consequently. At the same time, the degree of swelling of the chitosan (90%)/starch (5%)/castor (5%) was 2.5% after 20 min, while the absorption capacity of the extracted chitosan film was less than chitosan (90%)/starch (5%)/castor (5%) approximately 1% after 20 min.
Degree of swelling (water effect) of different bioplastic films synthesized.
According to Tan et al.75, the biofilm’s resistance property decreases with increasing chitosan content, which suggests a reduced ability of the biofilm to hold onto water. It results from a polymer chain in chitosan that has many –NH2 or hydrophobic amine groups, indicating the significant hydrophobicity of the polymer that makes up chitosan. A biopolymer called chitosan might have excellent qualities that make water safe for biofilm. It is brought on by the hydrophobicity (hatred of water) of starch-chitosan properties. Less water is retained in biodegradable plastic the more chitosan is used. It is exhibited at the highest level of water resistance, or at a 40% chitosan addition with the highest level of water assimilation of 0.38%76.
Tensile strength (TS)
Because of this, chitosan-derived materials have poor mechanical qualities, which makes it difficult for them to be used in more difficult applications. Expanding the use of plasticizers is one strategy used to solve this issue77. The maximal pull before breaking or ripping is used to determine the biofilm’s tensile strength. The goal of this measurement is to ascertain the amount of force needed to draw the film to its fullest extent across all of its surface areas. The results of the TS test for different biofilms are collected in Table 2. There are several significant criteria used to evaluate the mechanical properties of biofilms made from chitosan extracted from different microbial cell walls and/or crustaceans’ shells. Specifically, the different analyses can be summarized in TS mechanical properties of biofilm can be known from the reaction of ductile quality and elongation tests. Tensile Strength is the greatest constraint that might be stood up to by a biofilm until breakage78. The TS value increased in proportion to the increasing chitosan concentration. This is because more hydrogen bonds form in the biofilm at higher chitosan concentrations, making them stronger and more difficult to break. After all, more energy is required to do so79. Based on the data in Table 2, the TS of biofilm samples is influenced by the variation of the composition of chitosan, where in extracted chitosan film high tensile strength was found at 5.71% MPa. Based on the TS test findings, it was determined that adding starch and castor oil to chitosan films significantly increased the value of the TS biofilm.
The elongation at the break of the isolated chitosan film was 10.4%. The percentage change in the specimen’s initial length between the grips of a film to stretch (extend) reveals the flexibility and elongation capacity of the film, which is calculated at the point at which the film breaks during tensile testing. Young’s modulus measures how rigid the film is, and a lower value denotes a more flexible substance. Chitosan was added, and this resulted in a considerable decrease in the young’s modulus and the development of more flexible films. In the addition of chitosan, the Young’s modulus, which was 0.002 MPa, decreased. In addition, the elongation for the chitosan (90%)/starch (5%)/castor (5%) was breaking at 9.0%, and that shows the film capacity for flexibility, the rigidity of the film determined by Young’s modulus was 0.05 MPa. As well as the biofilm made of chitosan (70%)/starch (10%)/castor (20%) appears to have the maximum tensile strength at 5.51 MPa. The films’ ability to elongate and bend may be seen by looking at the elongation at break, which was 5.92%. The inclusion of chitosan decreased the rigidity of the film, as measured by Young’s modulus, which was 0.005 MPa and the breaking point was 0.0017 Mpa. Further, the data illustrates the TSA stress–strain relationship of PP. The behavior of all curves is essentially the same; they all start linearly, defining the slope, and then they all start to taper off after a given amount of strain, showing the stiffness of the material as it gets closer to the failure load. These plots are used to extract all significant tensile parameters, which are then provided in terms of the average, upper and lower bounds for the tensile modulus, strength, and maximum elongation. This resulted in a higher diameter of elongation which was 10.8%, lower Young’s modulus and TSA, and higher residual strain as well at 0.009 Mpa, and the breaking point was 0.0095 Mpa. The elongation of polypropylene decreases with the rise in the natural filler loading in the natural rubber composites, the PP was breaking at 11.2% (See them separately in Supplementary data file Fig. 5S, 6S, 7S, 8S).
The addition of up to 30% chitosan results in the maximum value of TS, which is 38.25 MPa. The important aspect that affects the mechanical behavior of biofilm material, according to Fathanah et al.80, is an affinity between its constituent parts. A certain atom or molecule may have a propensity to form bonds due to a phenomenon called affinity. The more molecules that bind, the stronger the affinity. The chemical bonding of a biofilm’s constituents has an impact on its strength. The number of molecules that are bound and the type of binding determine how strong a chemical is. It takes a lot of energy to break a strong chemical bond because strong chemical bonds are difficult to break.
Thermogravimetric analysis (TGA)
The curve of the extracted chitosan shows two stages of weight loss between 50 and 400 °C, with the first stage occurring between 50 and 100 °C due to water molecule loss and resulting in a weight loss of roughly 10.9%. A weight loss of about 41% resulted from the primary deterioration of the extracted chitosan, which started at 220 °C and was destroyed at 400 °C. The TGA of the chitosan (90%) / starch (5%) / castor (5%) showed three different stages of weight loss. The loss of adsorbed water may correspond to the initial stage of weight loss of 10.38%, which occurs at 60 °C, the thermal degradation causes the second disintegration step, which results in a weight loss of roughly 25.52% in the range of 250 °C. The third degradation was at approximately 300–400 °C because of thermal degradation with weight loss of about 25.8%. The TGA curve of chitosan (70%)/starch (10%)/castor (20%) exhibited that three stages 190 to 450 °C. The first stage results in a weight loss of 5.94% at 190 °C. The second decomposition stage occurs at 230–330 °C due to thermal degradation with a weight loss of about 16.6%. The third degradation was at approximately 350–500 °C because of thermal degradation with weight loss of about 56.1%. Furthermore, the TGA analysis for the PP film sample contains two stages in mass: the first stage indicates the thermal degradation of 3.09% occurs at 450 °C and the second stage is the thermal decomposition at 84.6% at 450 °C (See them separately in Supplementary data file Figs. 9S–12S).
TGA–DSC study
The DSC of extracted chitosan shows two broad endothermic peaks at 90.7 °C and the exothermic decomposition peaks at onset at 282.3–547.8 °C. The DSC of chitosan (90%)/starch (5%)/castor (5%) exhibited the broad endothermic peaks attributable to water loss at approximately 92.2 °C, the exothermic decomposition peaks at onset at 232.9–650 °C. As well as, the DSC thermogram of the films collected during the first heating cycle showed three signals endothermic peaks below 100 °C at 98.4 °C, while the exothermic peaks are broad variations 293, 379.6 and 645.5 °C. In addition, The TGA of the chitosan starch with castor 5% showed three different stages of weight loss. The loss of adsorbed water may correspond to the initial stage of weight loss of 10.38%, which occurs at 60 °C, the thermal degradation causes the second disintegration step, which results in a weight loss of roughly 25.52% in the range of 250 °C. The third degradation was at approximately 300–400 °C because of thermal degradation with weight loss of about 25.8%. Moreover, the TGA result of chitosan (70%)/starch (10%)/castor (20%) exhibited that shows that three stages 190 to 450 °C. The first stage results in a weight loss of 5.94% at 190 °C. The second decomposition stage occurs at 230–330 °C due to thermal degradation with a weight loss of about 16.6%. The third degradation was at approximately 350–500 °C because of thermal degradation with weight loss of about 56.1%. Furthermore, the TGA analysis for the PP sample contains two stages in mass: The first stage indicates the thermal degradation in 3.09% occurs at 450 °C and the second stage is the thermal decomposition at 84.6% at 450 °C (See them separately in Supplementary data file Figs. 13S–16S).
X-ray diffraction (XRD)
To design a particular application in the field of packaging industries crystallinity is one of the important ‘criteria for food packaging films. The X-ray diffraction measurements were carried out to analyze the nature of each film sample of extracted chitosan film, chitosan (90%)/starch (5%)/castor (5%), chitosan (70%)/starch (10%)/castor (20%) and PP as shown in Figs. 17S–20S, respectively. The X-ray patterns of chitosan film blends exhibited diffraction peaks of chitosan at around 9°, 20° and 64° of 2θ. When compared with Tripathi et al., X-ray patterns of chitosan. The diffraction peak of chitosan is at around 10° and 20° of 2θ. the highest intensity of about 800 counts at 2θ = 19°81. Moreover, the X-ray diffraction patterns of chitosan (90%)/starch (5%)/castor (5%) showed diffractive peaks in the region from 9°, 20° and 64°of 2θ with the highest intensity about 280 counts at 2θ = 20° than chitosan 220 counts. Furthermore, the XRD diffraction studies chitosan (70%)/starch (10%)/castor (20%), there are four peaks presented around 2θ = 9°, 2θ = 28° and sharp peaks at 2θ = 20°, 2θ = 43.5°. The highest intensity about 440 counts around 43.5° of 2θ. While the XRD patterns of PP were broad peaks at 2θ = 20°, 2θ = 23° and small peaks at 2θ = 9°, 2θ = 29°, 2θ = 36°, 2θ = 43.5°, and 2θ = 64°. Also, the highest intensity is about 1350 counts in 2θ = 23° (See them separately in Supplementary data file Figs. 17S–20S).
Surface area analysis
The Adsorption–desorption isotherm of Chitosan (70%)/Starch (10%)/Castor (20%)was studied as an example and the data were reported in the supplementary data (Figs. 21S–23S). The BET and BJH analysis showed that the film is mesopore with 3.1 nm mean pore diameter and 11.74 m2/g specific surface area.
Morphological analysis
In only a few straps, the chitosan film was uniform and continuous, with a high basic and no interface layer, as illustrated in various SEM micrographs of removed chitosan surfaces and manufactured bioplastic films in Fig. 5. This displays both the consistency and compatibility with which chitosan chains are depicted throughout the films. In general, hydrogen bonding between the functional bunches of the mixed component (–Gracious and –NH2 bunches in chitosan) was what caused the homogeneous mixtures of extracted chitosan-starch and castor oils, in both percentages; 5 and 20%. The SEM images at a magnification of 25 × 103 were obtained. When viewed on a computer, it was clear that they had accurately shown every pore on the pore surface and that, in addition to the PP scaffold membrane surface, the extracted chitosan and bioplastic films also had unevenly distributed pores. By SEM to watch the morphology of the break surface of the biofilms, and whether all components of biofilms have been blended homogeneously. A nonporous, smooth membranous phase with dome-shaped orifices, crystallite, and microfibrils was visible in the SEM images of the extracted chitosan film. Both chitosan (90%)/starch (5%)/castor (5%) and chitosan (70%)/starch (10%)/castor (20%) derivative electron micrographs showed that they had a porous, chain-like structure. Further, these images demonstrated the close spherical morphology of our synthesized biofilms, which may be utilized in biomedical applications37.
Scanning electron microscope showed (A, B) extracted chitosan film with different magnifications 25 × 103 and 250, respectively, (C, D) presented castor oil 5% mixed with starch-chitosan complex with 25 × 103, 250 dramatically, (E, F) also castor oil 20% mixed with starch-chitosan complex as 25 × 103 and 250 magnifications, finally (G, H) given as standard polypropylene bag at 25 × 103 and 250 consequently.
Antimicrobial activity of bioplastic films
There are several methods to evaluate the antimicrobial activity of the biofilms. This research used a disc diffusion test as an antimicrobial activity test for our bioplastic film but no clear result was recorded. It may be due to the weak contact between the bioplastic films and reference strain in the agar medium. Therefore, the antimicrobial activity was re-evaluated via suppression percentages of reference strains in liquid broth as mentioned in the “Materials and methods” section.
In particular, the data presented in Table 3 confirmed that there was no suppression detected after 2 h against any reference strain used (E. feacalis, K. pnumina, B. subtilis, and P. aeruginosa). It was also observed that the positive values of suppression % ranged from 14.8 to 96.8%. The lowest suppression was recorded for biofilm of chitosan (70%)/starch (10%)/castor (20%) against K. pnumina, while the highest one was recorded for the same formula against B. subtilis. From this result, the most susceptible reference strain was B. subtilis, while the most resistant reference strain was K. pnumina. Specifically, our bioplastic films have possessed effective antimicrobial activities. Thus, they are promising in such a manner with additional value.
Initially, chitosan has good properties to be moulded into plastic and has antimicrobial properties because it is easily biodegraded and biocompatible when combined with other materials and possesses biofunctional characteristics18. Consequently, chitosan has been utilized as a material for food packaging to preserve food quality82.
The development of chitosan-based solutions was the subject of a lot of research since the inherent qualities of chitosan might improve the packaging’s antimicrobial effectiveness. Due to chitosan’s ability to form films, several natural polysaccharides have been combined to create film packaging materials. Cha and Chinnan20 illustrated when they examined the antibacterial action of a chitosan-polysaccharides film against Listeria monocytogenes and showed that the growth of the bacteria was completely inhibited. The antibacterial activity of chitosan-HPMC films was significantly reduced when the amino groups of chitosan were cross-linked with citric acid, revealing the vital importance of the protonated amino groups of chitosan for antimicrobial action. Moreover, the biofilms prepared by Agustin and Padmawijaya83 showed fair antibacterial activity against S. aureus and E. coli. Recently, Chang et al.84 prepared chitosan-based plastic films, which had high antibacterial activity; especially with an inhibitory efficiency of over 95% against E. coli, P. fluorescens, and S. aureus.
Numerous studies have demonstrated that additional -NH3+ residues in chitosan make it easier for it to bind to bacteria, which causes structural instability in the bacteria85. However, as the amount of chitosan in the film rises, it may tighten the film’s structure, prevent amine groups from being exposed, or create electrostatic attraction between or between chitosan molecules, decreasing the antibacterial activity of the material84. Nevertheless, the addition of a plasticizer into chitosan biofilms may decrease the antimicrobial activity of biofilms because it fills the space between hydrogen bonds and will weaken the chitosan strength which can affect chitosan antimicrobial activity.
Biodegradation of bioplastic films
The biodegradability of the generated biofilm was evaluated to gauge its eco-friendliness. The outcomes shown in Table 4 supported the biodegradability of the synthetic biofilms we created, with weight loss during burial indicating genuine degradation. However, after the experiment, bioplastic films made with isolated chitosan became lighter and more translucent. In the case of the chitosan (90%)/starch (5%)/castor oil (5%) recipe, the biofilm’s color faded and it also became lighter and pore-filled. Various biofilm formulations become clear and cutting-friendly. Observably, PP as a synthetic plastic was used to compare all previous bioplastic films, and this has not changed at all. All of the examined bioplastic films lost weight over time, except PP, which essentially maintained its weight in all treatment media. On day 7, it was discovered that both films had lost more than 40% of their original weight. Chitosan-reinforced starch-based biofilm decomposed by 53.6%, while pure starch-based biofilm film deteriorated by 47.5%. No appreciable weight loss was observed for any biofilm the second and third weeks saw a 47.5% degradation of the biofilm75. As well as, day 28 marked the end of the weight loss of pure starch-based and chitosan-reinforced starch-based and 52.1 and 67.7%, respectively, of the total. Weight reduction was greater with the starch-only biofilm than with the chitosan-reinforced version.
Starch-based biofilm layers during burial may be related to higher starch matrix hydrophilicity86. The biofilm made of starch and chitosan exhibited a slower rate of deterioration, possibly because chitosan has antibacterial characteristics. In addition, the hydrogen bonding interaction between starch and chitosan, may lowered the starch matrix’s hydrophilicity, which slowed the biodegradation87.
The most important variables in biodegradation are more particularly the physiochemical characteristics of the biodegradable polymers, the ambient circumstances, and the microbial populations to which the biofilms are exposed. This process can take place in both natural and artificial settings, under aerobic and anaerobic circumstances latter of which has received the least attention in the literature. Compost, soil, and a few aquatic habitats are among the researched aerobic environments, whereas anaerobic environments include anaerobic digestion plants and a few aquatic habitats16. However, the breakdown process is influenced by both biotic (microbial activity) and abiotic (UV radiation, temperature, moisture, and pH) factors88. The thickness of the biodegradable material is another element that affects the rate of biodegradation; the thicker the product, the longer it will take to degrade89.
