Harnessing Wild Jackfruit Extract for Chitosan Production by Aspergillus versicolor AD07: Application in Antibacterial Biodegradable Sheets

Abstract

A fungal strain with comparably high chitosan yield was isolated from the Shivaganga hills and identified as Aspergillus versicolor AD07 through molecular characterization. Later, the strain was cultivated on Sabouraud Dextrose Broth (SDB) and wild jackfruit-based media to evaluate its potential for chitosan production. Among the various media formulations, the highest chitosan yield (178.40 ± 1.76 mg/L) was obtained from the jackfruit extract medium with added peptone and dextrose. The extracted chitosan was characterized through FTIR, XRD (reported a crystallinity index of 55%), TGA/DTG, and DSC analysis, confirming the presence of key functional groups and high thermal resistance. The extracted chitosan was fabricated into a sheet incorporated with 1% lemongrass oil; the sheet exhibited strong antibacterial activity against Escherichia coli (30 mm) and Bacillus megaterium (48 mm). The biodegradation studies reported a weight loss of 38.93 ± 0.51% after 50 days of soil burial. Further, the chitosan film was tested as a packaging material for paneer, demonstrating better preservation by maintaining nutritional quality and reducing microbial load over a 14-day storage period. These findings highlight the potential of A. versicolor AD07-derived chitosan, cultivated on a waste substrate medium, as a sustainable biopolymer for food packaging applications.

1. Introduction

The rising accumulation of agro-industrial waste and its mismanagement have become a global concern. The improper disposal of these wastes contributes to environmental threats such as greenhouse gas emissions, water and soil contamination [1]. In recent times, new sustainable approaches have been designed to convert the waste biomass into high-value bioproducts, aligning with the “waste-to-wealth” and circular bioeconomy paradigms [2]. Among the various approaches that are under development, the conversion of fruit and food processing waste into functional biomaterials is gaining considerable attention owing to the rich nutrient content that supports microbial growth and metabolite synthesis [3]. According to the recent reports by the Food and Agriculture Organization (FAO), nearly 1.3 billion tonnes of food waste are being generated every year, and the agro-wastes contribute to half of it [4].

The wild jackfruit (Artocarpus hirsutus), commonly known as “Anjili chakka”, is a tropical tree species native to the Western Ghats of India, in the regions of Kerala and Karnataka. It is primarily cultivated for its highly valued timber used in construction and furniture making [5]. When compared to its cultivated relative, Artocarpus heterophyllus (common jackfruit), the wild jackfruit is comparatively underutilized. As a result, a large portion of the fruit biomass remains unutilized, contributing to notable organic waste accumulation [6]. The wild jackfruit shared morphological similarities with cultivated jackfruits, consisting of a spiky rind, fibrous perianth, central core, edible bulbs, and seeds. The underutilization of wild jackfruits represents a missed opportunity for economic valorization. The fruit pulp is rich in carbohydrates, fibers, and minerals, rendering it a suitable substrate for microbial growth and bioconversion processes. Hence, transforming this biomass into substrates for microbial growth offers a low-cost alternative to synthetic media, additionally promoting sustainable bioprocessing.

Chitosan is a naturally occurring biopolymer, composed of partially deacetylated derivatives of chitin [7]. It is mainly known for its biocompatibility, antimicrobial properties, biodegradability, and non-toxicity. Generally, chitosan is extracted from the exoskeletons of crustaceans such as shrimp and crabs [8]. These traditional chitosan productions are facing issues such as seasonal availability, allergenic potential, and the overexploitation of marine resources. In this aspect, fungal-derived chitosan has emerged as a promising alternative to address the limitations associated with crustacean-derived chitosan. The filamentous fungal genera such as Aspergillus, Rhizopus, and Mucor are capable of producing high quantities of chitin, which can be converted into chitosan through chemical treatments [9]. Fungal chitosan possesses some superior properties when compared with crustacean chitosan. The fungal-derived chitosan is comparatively purer, vegan, and is better suited for biomedical and food-grade applications, as it avoids the impurities and heavy metals often associated with marine sources [10].

The biomass generation and the chitosan yield highly depend on the composition of the culture medium. The incorporation of fruit waste-based media for biomass production can significantly enhance the quality of the chitosan, additionally lowering its production cost [11]. Several studies have reported the successful enhancement of fungal polysaccharide production using agro-residues such as sugarcane bagasse, banana peel, and orange pulp. Conversely, studies related to the usage of wild jackfruit-based medium for fungal chitosan biosynthesis remain modest.

Paneer (Indian cottage cheese) is a widely consumed dairy product in India, and it has a very short shelf life because of its high moisture content, neutral pH, and rich nutrient profile. These conditions make it an ideal medium for microbial growth. In India, Paneer spoilage is a major concern, especially in the localities where transportation and cold storage facilities are limited [12]. Paneer has a shorter shelf life than meat, but lasts longer than milk and most fresh vegetables under similar storage conditions. Although the exact national statistics related to paneer spoilage are scarce, the broader data from the Food Safety and Standards Authority of India (FSSAI) suggests that nearly 20–30% of dairy products are lost or wasted in India due to poor handling, storage, and distribution practices. Paneer production in India accounts for 5% of the total dairy production [13]. These production statistics could translate to a total paneer spoilage or wastage of 3000–4000 tonnes every day. Paneer has a shelf life of 2 or 3 days at room temperature, and in cold storage it can last for about 5–6 days [14]. Spoilage due to microbial activity is the primary cause for paneer degradation, and the common spoilage organisms include Bacillus spp., Escherichia coli, Staphylococcus aureus, Pseudomonas Spp., Lactobacillus spp., Candida spp., and fungal stains from genera such as Aspergillus and Penicillium [15,16]. The activity of these organisms causes undesirable changes in the appearance, odor, and flavor of paneer; at times, these organisms render compounds unsafe for consumption. Hence, the development of effective preservation methods and processing is significant in extending the shelf life of paneer, thereby ensuring consumer safety [14].

To explore this uninvestigated aspect, the present study focused on the isolation and identification of a novel fungal strain capable of yielding high quantities of chitosan using the wild jackfruit-based medium. Later, a sheet was synthesized from the extracted chitosan with the incorporation of 1% lemongrass essential oil. Particularly, lemongrass oil is rich in citral and other terpenoids, which are known to have a broad-spectrum antimicrobial activity. The biodegradability and the antimicrobial aspects of the sheet were subsequently tested. This investigation also aims to utilize this synthesized biodegradable sheet to wrap Indian cottage cheese (paneer) to evaluate its effectiveness in extending shelf life.

2. Materials and Methods

2.1. Isolation of Fungal Strain for Chitosan Production

The soil samples were procured from Shivagange Hills (13.172017913025°, 77.2251228520333°), Karnataka, India (Figure 1). This site is renowned for its distinct geological landscape and cultural importance, features rugged, rocky terrain, and a semi-arid climate. Such an environment supports a diverse and abundant microbial population, making the site highly suitable for isolating fungal strains with promising industrial and biotechnological potential [17]. All the media components and chemicals (analytical grade) used in this study was procured from M/s Hi-Media, Mumbai, India.The samples were maintained at 4 °C for transport and storage. Later, 1g portion of the soil sample was mixed in 100 mL of autoclaved distilled water, followed by serial dilution up to 10⁻⁶ [18]. The dilutions 10⁻⁴, 10⁻⁵, and 10⁻⁶ were spread plated onto sterile Sabouraud dextrose agar (SDA) plates (dextrose—4% w/w, peptone—1% w/w, Agar—1.8% w/w) supplemented with ampicillin (40 mg/L) to inhibit bacterial growth and promote fungal proliferation. These plates were incubated at room temperature for 14 days [19]. After the incubation period of the plates, 10 visibly distinct fungal colonies were observed and sequentially labeled (AD01–AD10). These isolates were separately grown on fresh SDA plates and were stored for further study. Later, these isolates were inoculated in Sabouraud dextrose broth (SDB) (dextrose—40 g/L, peptone—10 g/L) and incubated at room temperature for 7 days for fungal biomass production. Chitosan biopolymer was extracted from each fungal biomass, and their yields were compared; one of the highest-yielding isolates (AD07) was selected for this study. A microscopic examination of the AD06 fungal isolate was performed using the lactophenol cotton blue (LPCB) staining technique [20]. The LPCB acts as a clearing agent that could remove the debris and enhance the visualization of fungal stains. For ensuring a uniform inoculum size for the subsequent studies, fungal inoculum disks of AD07 were prepared by placing Whatman filter paper disks (5 mm diameter) on SDA plates inoculated with AD07 [21].

2.2. Biochemical and Molecular Characterization

Fungal DNA was extracted from AD07 fungal biomass cultured on SDA. Isolated DNA was subjected to 1% agarose gel electrophoresis. Polymerase chain reaction (PCR) was performed for the high-molecular-weight DNA band, which was used to amplify the ITS region. Further, the PCR amplicons were subjected to agarose gel electrophoresis to reveal the amplicon band of 600 bp. The purified PCR amplicon was subjected to forward and reverse sequencing using ITS1 and ITS4 primers with the BDT v3.1 Cycle Sequencing Kit on the ABI 3730xl Genetic Analyzer. An aligner software was used to align the obtained forward and reverse sequence data to generate a consensus sequence. The obtained consensus sequence underwent a BLAST (version 2.14.0) search on the NCBI GenBank database to obtain ten sequences with the highest identity were selected [22]. These sequences were aligned with the ClustalW software, and later the distance matrix analysis and the phylogenetic tree construction were performed based on nucleotide homology and phylogenetic analysis using MEGA 10 [23].

2.3. Production of Fungal Biomass

The fungal biomass was grown using both commercial medium and waste fruit-derived medium (Wild Jackfruit), and the chitosan yield from both sources was compared.

The commercially available SDB powder was used to produce the commercial medium and was inoculated using the fungal inoculation disk. This medium was kept at room temperature for 14 days to achieve maximum fungal growth.

Ripe, fallen wild jackfruits were gathered from Kottayam, Kerala, India, for utilizing them in the preparation of fruit waste-derived medium for the production of fungal biomass. The utilization of fruit waste-based media for fungal growth was designed to develop a sustainable and cost-effective method for producing value-added chitosan. The fruits were washed and the seeds were removed, and the remaining pulp was dried at 50 °C in a hot air oven and was powdered. A proximate analysis of the powdered fruit pulp was conducted to determine the sugar and protein content, which would have an effect on fungal biomass production [24].

Later, 50 g of the wild jackfruit powder was blended in 100 mL of distilled water for 1 h, and the resulting liquid extract was filtered. This extract was maintained as the base component to prepare four different media combinations by supplementing with peptone and dextrose:

(i)

JE + P: Wild jackfruit extract (100 mL) + peptone (10 g/L)

(ii)

JE + D: Wild jackfruit extract (100 mL) + dextrose (40 g/L)

(iii)

JE + P + D: Wild jackfruit extract (100 mL) + dextrose (40 g/L) + peptone (10 g/L)

(iv)

JE: Wild jackfruit extract (100 mL)

The medium components were thoroughly mixed and were autoclaved at 121 °C for 15 min. The sterile media were later inoculated with a fungal inoculum disk and were incubated under static conditions at room temperature for 14 days [25].

2.4. Extraction of Chitosan from the Cultivated Fungal Biomass

The fungal biomass was harvested after 14 days of incubation and was washed with distilled water to remove the residual media components. For the chitosan extraction, the fungal biomass was subjected to alkali treatment in 1M NaOH (1:40 g (biomass)/mL (NaOH)) at 121 °C and 15 psi for 15 min in an autoclave. Alkali treatment will help in the removal of alkali-soluble proteins and other impurities. The mixture was then centrifuged at 10,000 rpm for 15 min to separate out the fungal biomass from the alkali solution. Fungal biomass was subjected to repeated washes to neutralize the pH. Alkali-treated fungal biomass was added to 2% (v/v) acetic acid treatment (1:30 g(fungal biomass)/mL) for 8 h at 95 °C in a water bath. This treatment facilitates the dissolution of the fungal chitosan in the acetic acid solution. The solution was filtered, and the pH was adjusted to 11 using 1 M NaOH to induce fungal chitosan precipitation. The precipitated chitosan was separated out by centrifuging the solution at 10,000 rpm for 15 min. The recovered chitosan was neutralized by repeated washing with distilled water [26].

2.5. Characterization of Extracted Fungal Chitosan

The extracted AD07 fungal chitosan was subjected to characterization techniques such as Fourier transform infrared spectroscopy (FTIR), X-ray diffraction (XRD), thermogravimetric analysis (TGA), differential thermal analysis (DTA), and differential scanning calorimetry (DSC) to assess the functional groups and physicochemical properties.

FTIR was employed to understand the functional groups present in the AD07 fungal chitosan. The infrared absorption bands were obtained by scanning the extracted chitosan within the IR frequency range of 4000–400 cm⁻¹ at a resolution of 4 cm⁻¹ [27] using a Shimadzu IR Spirit-L FTIR spectrophotometer at the Common Instrumentation Lab, Department of Life Sciences, CHRIST (Deemed to be University).

The XRD analysis was performed to determine the crystal phase of the AD07 fungal chitosan. The samples were scanned across the 2θ values ranging from 5° to 90° to obtain the XRD spectrum [28]. The XRD analysis was performed using the MiniFlex 600 XRD at CHRIST (Deemed to be University), Bangalore. The crystallinity index of the fungal chitosan was calculated by dividing the area of the crystalline peaks in the XRD spectrum by the total peak area.

The TGA and DTA thermograms were obtained by heating the sample from 40 °C to 800 °C at a rate of 20 °C/min. The thermogram reflects the thermal stability and degradation behaviour of the sample [29]. The TGA and DTA were performed using the PerkinElmer STA 6000 at the Research & Development Cell, St. Joseph’s University, Bengaluru, Karnataka.

The DSC thermogram of the fungal chitosan was obtained by heating the samples from room temperature to 800 °C at a rate of 10 °C/min. The DSC analysis reveals the thermal phase transitions, including the glass transition and thermal degradation ranges of the biopolymer [30]. DSC thermogram was acquired from the Sophisticated Test and Instrumentation Centre (STIC), Cochin University of Science & Technology, Kochi.

2.6. Synthesis of Chitosan Sheet Incorporating Lemongrass Oil

The chitosan extracted from the AD07 fungal isolate was added to 1% (v/v) acetic acid solution (100 mL) with 1 mL of lemon grass oil, and the mixture was kept on a magnetic stirrer at 500 rpm for 3h at 60 °C to dissolve the chitosan. The lemongrass oil for the study was extracted from Cymbopogon flexuosus (Cochin grass). It contains the key compound called citral, which was proven to have a broad range of antibacterial effects on both the Gram-positive (Staphylococcus aureus and Bacillus sp.) and Gram-negative (Escherichia coli, Klebsiella pneumoniae, and Pseudomonas aeruginosa) bacteria [31]. Subsequently, the chitosan-acetic acid solution was cast into a casting tray and was kept at 55 °C for 6 h to facilitate chitosan sheet formation [32]. The mechanical properties, including tensile strength, Young’s modulus, elongation at break, and elongation at peak load of the chitosan sheet, was procured by testing using a Universal Testing Machine (UTM) at the Department of Physics, CHRIST University, Bangalore.

Antimicrobial Activity of the Chitosan Sheet

The casted sheets with different concentrations of lemongrass oil were cut into squares of 1 cm × 1 cm dimension and were placed onto sterile Müller–Hinton (beef extract—2 g/L, acid hydrolysate of casein—17.5 g/L starch—1.5 g/L, and agar—17.0 g/L) Petri dishes plated with E. coli (Gram-negative) and Bacillus megaterium (Gram-positive). The plating of the microbes was carried out using 100 µL of the inoculum, with CFU adjusted to 1 × 10⁴ CFU/mL. The plates were incubated for 24 h at 37 °C to facilitate bacterial growth [33].

2.7. Biodegradation Studies Using the Fabricated Chitosan Sheet

The sheets, which were cut into 1 cm × 1 cm dimensions, were weighed and enclosed in non-biodegradable plastic mesh. These setups were buried at a depth of 1 cm in 1 kg of garden soil in triplicate to study the degradation behaviour of the chitosan sheet. The moisture content was maintained at 60% throughout the study [34]. The weight of the fungal chitosan sheets was recorded every 10 days, and the degradation percentage of the fungal chitosan sheet was calculated using the formula:

Degradation percentage (%) (w/w%)= ([Initial weight − Final weight]/Initial weight) × 100

(1)

2.8. Shelf Life Extension Study Coating the Paneer with Chitosan Sheet

The shelf life extension properties of the chitosan sheet were tested on paneer. Fresh paneer was procured from a dairy outlet in K. R. Market, Bangalore. The fresh paneer was cut into cubes of 1 cm × 1 cm × 1 cm for easy wrapping of the chitosan sheet to study their shelf life (Figure 2). Typically, paneer has a refrigerated shelf life of up to 30 days when stored in aseptic packaging. However, once the packaging is opened, its shelf life reduces to approximately 5 to 6 days under refrigeration. Hence, the chitosan-packed paneer was subjected to refrigeration for 14 days along with an unpacked control. On the 14th day, the paneer samples were analyzed for the microbial count, moisture content, total protein content, total lipid content, and total carbohydrate content. The analysis was carried out according to the test methods specified by the Association of Official Analytical Chemistry (AOAC Method) [35].

3. Results and Discussions

3.1. Isolation of Fungal Strain for Chitosan Production

The ten fungal isolates (AD01–AD10), which were isolated from the Shivagange soil samples, were cultivated for biomass production in SDB. The biomasses obtained were subjected to chitosan extraction. Among the isolates, AD07 was selected for further studies due to its high chitosan yield. Macroscopically, the fungal colony showed a slow-growing, flat appearance with a velvety, powdery texture (Figure 3a). The colony initially appeared white during the early stages of growth and gradually transitioned to a yellow-green hue as it matured. The reverse side of the agar plate presented a pale yellow hue. Microscopically, AD07 displayed hyaline (transparent) septate and branched hyphae (Figure 3b). The conidiospores appeared short, smooth-walled, and unbranched. The tip of conidiophores terminated in a spherical body which appeared to be covered with biseriate phialides—consisting of both metulae and phialides—arranged over the entire surface, forming either a radiating or loosely columnar structure. Upon the preliminary microscopic and macroscopic examination, the fungal strain AD07 can be related to a member of the Aspergillus genus [36].

3.2. Biochemical and Molecular Characterization

The ITS sequence of the AD07 fungal strain was acquired through PCR amplification and sequencing (Figure 4a). The sequence was then subjected to BLAST analysis, and the results revealed that the AD07 fungal isolate exhibited a high degree of resemblance to Aspergillus versicolor. A phylogenetic tree was created using the top ten sequences with the highest identity scores (Figure 4b). Later, the sequence was submitted to the GenBank database, and the isolate used in this study was confirmed as Aspergillus versicolor AD07, with the assigned accession number PP182246.1. In the study conducted by Manaswini et al., (2025), A. versicolor was isolated from the roots of P. amboinicus collected from Bengaluru, Karnataka [37]. A. versicolor was obtained during the characterization of the fungal microflora from Mattavara Forest in Chikkamagaluru, Karnataka [38]. Mukunda et al., (2013), obtained A. versicolor while screening for the industrially important fungi from the Western Ghats forests of Agumbe and Koppa, Karnataka [39]. These studies conclude the extensive distribution of Aspergillus versicolor in the regions adjacent to the sampling site of the present study.

3.3. Extraction of Chitosan from the Cultivated Fungal Biomass

A substantial presence of carbohydrates and proteins can promote robust fungal biomass growth. Hence, the proximate chemical composition of the wild jackfruit was found out (

Table 1. Proximate chemical composition of Jackfruit pulp.

Biochemical Composition	Average
Moisture content (w/w%)	82.61 ± 0.030%
Ash content (w/w%)	2.11 ± 0.007%
Carbohydrate (w/w%)	17.07 ± 0.017%
Protein (w/w%)	11.40 ± 0.027%

Data represents the mean of triplicate ± standard deviation.

). After the 14th day of static incubation of the wild jackfruit extract (JE)-based media inoculated with Aspergillus versicolor AD07, the fungal biomass was harvested and subjected to chitosan extraction (Figure 5). The chitosan yielded from the AD07 biomass cultivated on SDB was recorded as 75.88 ± 2.99 mg/L. The chitosan yield obtained from fungal biomass grown on commercial media was comparatively lower than that from all setups where the fungi were cultivated using JE-based media (Figure 6).

The highest chitosan yield, recorded at 178.40 ± 1.76 mg/L, was achieved from fungal biomass cultivated in the JE + P + D medium. This was followed by the JE + P setup, which produced a chitosan yield of 163.43 ± 1.56 mg/L. The biomass grown on the jackfruit extract alone produced a chitosan yield of 107.91 ± 3.96 mg/L. The fungal biomass from the JE + D setup yielded 78.23 ± 3.07 mg/L of chitosan.

In a similar study using Aspergillus niger BBRC20004, Rasmussen et al., (2007), used soybean residue as the substrate and obtained a fungal chitosan yield of 17.03 g/kg [40]. More recently, Davis et al. (2024) cultivated Aspergillus niger DEL01 on pineapple peel waste-based medium and obtained a chitosan yield of 139 ± 0.25 mg/L [41]. The chitosan yield obtained in the present study was higher compared to those reported in the aforementioned studies.

The higher yield observed in JE compared to the commercial SDB indicates that JE alone contains sufficient nutritional components to support fungal biomass growth and chitin production. The closeness in yield from the biomass obtained using JE + P + D and JE + P indicates that the addition of peptone and dextrose can provide extra nutrients, potentially enhancing the fungal chitosan yield in the resulting biomass. The slight reduction in chitosan yield from the biomass obtained from the JE + D setup when compared with other setups can be due to carbon–nitrogen imbalance. Fungi tend to produce lipids or spore formation rather than chitin synthesis at a high C/N ratio. These findings align with the studies of Rasmussen et al., (2007), in which a 50% reduction in fungal growth under high-sugar conditions and a 40% decline when nitrogen was in surplus was reported [40]. The proximate analysis of wild jackfruit showed a higher carbohydrate content relative to its total protein concentration, which may account for the comparatively lower chitosan yield observed in the JE + D setup.

Among the studies utilizing fruit waste-based media to cultivate fungal biomass, this research is one of the first to specifically employ Aspergillus versicolor for chitosan extraction. This is also one of the first studies to utilize wild jackfruit extract-based media for cultivating fungal biomass. In a related work, Rasmussen et al., (2007), cultivated Aspergillus niger BBRC20004 on soybean residue, resulting in a chitosan yield of 17.03 g/kg after extraction [40]. Habibi et al., (2021), cultivated Aspergillus terreus on apple waste extract and reported a chitosan to substrate yield of 49.32 mg/g [11]. Similarly, Davis et al. (2024) extracted fungal chitosan from Aspergillus niger DEL01 biomass cultivated on pineapple peel waste-based media, reporting a yield of 139 ± 0.25 mg/L [41]. In the study by Elgohary et al., (2024), 7.8 g/L of chitosan yield was obtained from Aspergillus flavus grown on potato dextrose broth (PDB) [42].

3.4. Characterization of Extracted Fungal Chitosan

The FTIR spectra (Figure 7) revealed the characteristic peaks observed in chitosan samples. The broad peak at 3524.36 cm⁻¹ can be due to the presence of water [43]. The peak at 3447.25 cm⁻¹ indicates the O-H stretching vibrations related to the hydroxyl groups present in the chitosan sample. The peak at 2930.30 cm⁻¹ is attributed to the methyl (CH₃) groups’ C-H stretching vibrations [44]. The C-H bending vibrations of the chitosan are associated with the peak at 1472.29 cm⁻¹. The peak at 1567.97 cm⁻¹ can be associated with the N–H bending vibrations of the amide II band [45]. The C-O stretching, which is observed in the glucosamine rings, can be attributed to the peak observed at 968.20 cm⁻¹. The 592.63 cm⁻¹ peak corresponds to the N-H bending vibrations [25]. The sharp peak at 502.66 cm⁻¹ may be attributed to the N–H wagging/bending vibrations of the amine salt deformations [46]. The absence of the peaks related to the impurities highlights the considerable purity of the extracted fungal chitosan.

The XRD spectrum of the fungal chitosan is illustrated in Figure 8. The prominent peak observed at 2θ = 22.02° represents the (110) crystalline phase of chitosan [47]. The broad peak observed at 2θ = 13.44° corresponds to the amorphous (020) phase of chitosan. The peaks can be correlated to JCPDS card No. 00-039-1894, which represents chitosan [48]. The additional peaks observed in the XRD spectrum of the fungal chitosan at 2θ = 34.92° (002) and 2θ = 60.42° signify the crystallographic reflections of chitosan molecules, which are related to the crystallinity index of the chitosan (CI%) [49]. The CI% of the fungal chitosan was found to be 55.49%, which indicates a relatively high degree of structural order. The higher crystallinity suggests the moderate rigidity and flexibility of the material, which enhances the tensile strength of the sheet produced with them. Sheets prepared from materials with a considerably high CI% are strong enough for packaging, yet flexible enough for biomedical use [50]. Additionally, higher crystalline substances are less soluble in water, which makes them suitable for moisture-sensitive packaging. Chemically, moderate CI% represents a good number of reactive sites for chemical modifications, which enables them to form stable composites [51]. Upon comparison with similar studies, the chitosan exhibited a comparable or slightly higher CI%. When chitosan extracted from Ugandan mushrooms was subjected to XRD, Ssekatawa et al., (2021), obtained a CI% of 48.4  ±  0.44% [52]. Kaya et al., (2015), reported a CI% of 52% from the chitosan extracted from Fomitopsis pinicola [53]. In a similar study, Davis et al. (2024) obtained a CI of 51.61% for chitosan extracted from Aspergillus niger [41].

The TGA spectrum of the AD07 fungal chitosan was illustrated in Figure 9a. The initial weight loss of 11.927% (w/w) observed between 30 °C and 221.33 °C is linked to the evaporation of water present in the sample. The significant weight loss of 52.814% (w/w) between 231.33 °C and 541.11 °C can be attributed to the fast pyrolysis phase of the biopolymer. Fast pyrolysis can be aligned with the breakdown of acetylated chitin structures within the chitosan matrix. This phase also represents the thermal degradation of its polymeric components [54]. Upon inspecting the DTA spectrum (Figure 9b), three distinct thermal events were observed at 288.51 °C, 358.22 °C, and 450.6 °C within the same temperature range of the fast pyrolysis [55]. These peaks can be indicative of the progressive depolymerization of the chitosan structure.

The TGA curve further enters a slower pyrolysis phase between 541.1 °C and 800 °C, which accounts for a weight loss of 14.568% (w/w). The slower pyrolysis can be related to the degradation of pyranose ring structures [56]. A residual mass of 20.64% (w/w) was observed upon the completion of the thermal analysis up to 800 °C [57]. Reasonably low residual weight implies the high purity of the extracted fungal chitosan. Overall, the thermal analysis validates the excellent thermal stability of the extracted fungal chitosan, making it suitable for industrial processes that deal with high temperatures.

The DSC thermogram of the extracted fungal chitosan is depicted in Figure 10. The first endothermic peak at 41.99 °C can be connected with the evaporation of the moisture content from the chitosan, as chitosan’s hydrophilic nature causes it to absorb atmospheric water [58]. The prominent exothermic transition, which peaked at 423.93 °C, refers to the principal degradation stages associated with the disintegration of the glycosidic linkage within the chitosan backbone. This exothermic peak highlights the higher thermal resilience of the extracted chitosan biopolymer. The peak at 526.65 °C could be with the oxidative decomposition or the combustion of carbonaceous residues [59]. The overall thermal behaviour closely resembles that of commercial chitosan as reported by Soon et al. (2018), further supporting the purity and quality of the AD06 fungal chitosan [60].

3.5. Synthesis of Chitosan Sheet Incorporating Lemongrass Oil

The synthesized chitosan sheet resembled conventional plastic films (Figure 11). The transparent chitosan sheet had a thickness of 0.43 mm and had a light yellow tinge. The stress–strain curve for the fungal chitosan sheet is depicted in Figure 12. The tensile strength of the chitosan sheet was found to be 34.97 MPa, and the Young’s modulus value of the sheet was calculated to be 162 MPa. The sheet exhibited the elongation at Peak of 6.05% and the elongation at break of 7.03%. These values are promising, considering the sheet was made without any other additives; the mechanical properties can be improved by the addition of other polymerizing agents such as PVA. The overall appearance and the characteristics of the fungal chitosan film produced in this study closely matched those reported by Zhang and Jiang (2020) [61]. In the study conducted by Kanthiya et al., (2022), a sheet of poly(lactic acid) and chitosan was produced. This sheet exhibited a tensile strength of 30 MPa [62]. Prashanth et al., (2021), have reported a maximum tensile strength of 22.5 MPa for 3% chitosan with 2% coir fibres [63]. When compared to these previously reported results, the tensile strength observed in the current study was found to be comparatively higher.

Antimicrobial Activity of the Chitosan Sheet

The sheets showed zones of inhibition for both E. coli as well as Bacillus megaterium with diameters of 30 mm and 48 mm, respectively (Figure 13). This reveals the property of the film to release the compound to the surrounding environment and to arrest the growth of both Gram-positive and Gram-negative bacteria. The antimicrobial properties of the chitosan in various studies are discussed in detail by Ke et al., (2021) [64]. Studies have already reported that 1% (v/v) of lemongrass showed antimicrobial activity against the Gram-positive as well as Gram-negative organisms. Perdana et al., (2021), demonstrated that the minimal inhibitory concentration of lemon grass oil against E. coli and Bacillus cereus was 1.56 µL/mL and 0.39 µL/mL [65]. The antimicrobial activity of lemongrass oil was demonstrated against various organisms, including Bacillus subtilis, Bacillus cereus, and E. coli in the study conducted by Shendurse et al., (2021) [66].

3.6. Biodegradation Studies Using the Fabricated Chitosan Sheet

Figure 14 depicts the results of the biodegradation of the chitosan sheets after 50 days of soil burial. The biodegradation of biopolymers follows three stages [67]. The initial degradation begins with biodeterioration, within which the enzymes disintegrate the polymer into simpler molecules [68]. This biodeterioration happens through the interference of both the physical and enzymatic activities. The depolymerization happens in the second phase, in which the polymer breaks into monomers, resulting in a reduction in polymer weight [69]. The final phase is the mineralization, which involves the complete oxidation of the polymer residues [70].

The degradation percentage after the 10th day of soil burial was observed to be 2.38 ± 0.61% (w/w), which increased steadily till the 30th day of soil burial. The biodegradation percentage recorded after the 20th and the 30th day was 5.43 ± 0.53% (w/w) and 10.39 ± 0.73% (w/w), respectively. The biodegradation percentage of the fungal chitosan sheet exhibited a steep increase from the 30th to the 50th day, reaching 19.38 ± 0.93% (w/w) after 40 days and 38.93 ± 0.51% (w/w) after 50 days of soil burial. This initial steady increase, followed by the steep jump in the biodegradation percentages, can likely be due to the slow initial biodegradation phase, followed by rapid depolymerization and a notable reduction in polymer weight.

The study conducted by Santhosh and Umesh (2024) reported the biodegradation percentage of plastic sheets to be 12.08 ± 0.15% (w/w). In comparison, the transparent biopolymer sheet in the current study exhibited more than three times the biodegradation rate of the plastic sheet. This suggests that the fungal chitosan sheet can be used as a biodegradable alternative to plastic sheets. In a study conducted by Pavoni et al., (2021), the enhanced structural breakdown of chitosan films was observed when modified with acetic and lactic acids [71]. A complete degradation of the starch-based chitosan nanocomposite made from brown rice starch was observed after 20 days of soil burial by Hasan et al., (2020) [72].

Upon comparisons with the degradation rates of packaging plastic sheets, the results of the current study suggest its potential to replace synthetic plastics as a packaging material. The high thermal performance and comparatively good mechanical properties further support this conclusion.

3.7. Shelf Life Extension Study Coating the Paneer with Chitosan Sheet

The current study investigated the shelf life extension of the fungal-derived chitosan sheet for its utilization as a bioactive packaging material. The visual inspection and microbial analysis over the 14-day period clearly demonstrated that the chitosan-based wrapping provided a significant protective effect against microbial spoilage (Figure 15).

The paneer samples, which were wrapped with the fungal chitosan sheet, maintained their appearance and texture, remarkably superior to the unwrapped control sample. The unwrapped paneer samples showed visible signs of spoilage, such as discoloration and surface degradation. The difference was supported by the microbial load measurements, as the control exhibited a high bacterial count of 1.59 × 10⁷ CFU/mL after 14 days, while the wrapped paneer samples had a lower count of 6.96 × 10⁵ CFU/mL. This substantial reduction in microbial load can be attributed to the antimicrobial properties of both chitosan and the incorporated lemongrass oil. There was no visible evidence of any biofouling during the course of the shelf life extension study, thereby indicating the films to be stable for packaging applications.

The proximate analysis presented in

Table 2. Changes in nutritional parameters of paneer during storage (day 0 vs. day 14—wrapped vs. unwrapped).

Parameters	Day 0	Day 14
		Unwrapped	Wrapped
Protein (w/w%)	23.37	21.87	21.12
Fat (w/w%)	19.61	18.11	18.27
Carbohydrate (w/w%)	11.23	8.5	11.13

Data represents the mean of triplicate ± standard deviation.

revealed that the wrapping did not significantly alter the nutrient contents of the paneer; further, it preserved the total carbohydrate content of the paneer. On day 0, the protein content was 23.37% (w/w), while on day 14, it measured 21.87% (w/w) in the unwrapped sample and 21.12% (w/w) in the wrapped sample. The total fat content was 19.61% (w/w) on day 0, which reduced to 18.11% (w/w) for the unwrapped and 18.27% (w/w) for the paneer sample wrapped with the chitosan sheet. The total carbohydrate content on the initial day was 11.23% (w/w), which was reduced to 8.5% (w/w) for the unwrapped paneer sample, which could be attributed to the microbial fermentation of the lactose sugar into lactic acid. However, the paneer sample wrapped with the chitosan sheet preserved the total carbohydrate content, which was reported to be 11.13% (w/w). This can be related to the antimicrobial results of the chitosan, which were discussed above.

Suresh et al., (2015), tested the shelf life of eggs coated with chitosan. The study revealed that a three-layer coating of chitosan preserved the internal quality and prevented weight loss [73]. In another study conducted by Kanatt et al., (2013), ready-to-cook meat products (chicken balls, chicken seekh kababs, and mutton seekh kababs) were coated with chitosan (2 g/100mL) to test the shelf life extension capability. The study showed that the chitosan-coated samples exhibited minimal changes in organoleptic properties, resulting in a significant extension of shelf life [74].

The results outlined in the study confirm that wrapping paneer with a chitosan–lemongrass oil sheet effectively slows down microbial growth and spoilage, thereby extending its shelf life. The bioactive packaging developed in this study offers a sustainable and health-conscious packaging solution, which could replace synthetic preservatives or plastic films, making it highly relevant for improving food safety and reducing waste in dairy supply chains.

4. Conclusions

The current study explored the cost-effective production of chitosan biopolymer via waste valorization. The fungal strain Aspergillus versicolor AD07 isolated from soil samples collected from Shivaganga hills, Karnataka, was cultivated on the wild jackfruit-derived media to enable sustainable chitosan synthesis. The highest chitosan yield of 178.40 ± 1.76 mg/L was achieved from fungal biomass cultivated in the jackfruit extract media supplemented with peptone and dextrose. This yield significantly surpassed the yield from the fungal biomass cultured on synthetic media (75.88 ± 2.99 mg/L). The chitosan extracted from the AD07 fungal biomass was subjected to characterization techniques (FTIR, XRD, DSC, and TGA). The FTIR confirmed the characteristic functional groups present in chitosan, and the XRD spectrum exhibited the characteristic peaks present in chitosan. The crystallinity index (CI) was calculated to be 55.49%. Thermal analysis (TGA, DTA, and DSC) demonstrated the high thermal stability of the extracted fungal chitosan. A chitosan sheet was fabricated by incorporating lemongrass oil; the sheet resembled synthetic plastic in appearance and exhibited a biodegradability rate of 38.93 ± 0.51%% (w/w) after 50 days of soil burial. The biodegradable chitosan sheets were later used for packing paneer cubes to study the impact on shelf life. The results revealed excellent shelf life extension and minimal microbial growth for the paneer samples wrapped with the chitosan sheet. These findings suggest that fungal-derived chitosan film can serve as a thermally stable, biodegradable alternative to synthetic packaging plastic films.