Antifungal Activity of Nanochitosan in Colletotrichum musae and Colletotrichum chrysophillum

Abstract

The search for developing materials of natural origin has become imperative due to the resistance shown by phytopathogenic microorganisms to traditional antimicrobial agents. Natural polymers such as chitosan offer a new alternative to fungal infections because, in most cases, these polymers are biocompatible, nontoxic, and natural. This study aimed to synthesize nanochitosan using ultrasonication and evaluate its antifungal activity on Colletotrichum chrysophillum and Colletotrichum musae. Nanochitosan of 302.4 ± 92.3 nm and a zeta potential of +35.9 ± 2.3 Mv, amorphous in shape, and a rough surface, was obtained. Nanochitosan reduced the radial growth 21%, for C. chrysophillum while C. musae showed a maximum inhibition of 26% at a concentration of 1.5 mg mL⁻¹ of nanochitosan. C. musae was the species most affected, with a 38% increase in hyphal diameter to 12 h. Also, nanochitosan affected the integrity of the fungi cell walls, plasma membrane, and generated low oxidative stress level. Our findings indicate that nanochitosan induces notable changes in the intracellular structures of the tested phytopathogens. Nevertheless, additional investigations are required to clarify the mechanisms underlying adaptability or resistance in fungal strains that exhibit reduced sensitivity to this biopolymer.

1. Introduction

The widespread application of fertilizers and pesticides in the agri-food industry has resulted in contamination, microbial resistance, and adverse effects on human health. Today, nanotechnology emerges as a promising alternative for creating new environmentally friendly antimicrobials. This approach promotes sustainable practices, enhances efficiency, and contributes to food security [1]. Chitin is a polysaccharide found in nature, and the deacetylation of at least 60% of its acetyl groups is known as chitosan [2]. Extensive research has been conducted on chitosan’s antifungal properties against various fungi, including Aspergillus, Botrytis, Penicillium, Lasiodiplodia, and Colletotrichum [3,4,5,6]. Colletotrichum is a fungal genus responsible for developing anthracnose, one of the most significant diseases that threaten fruit crops such as papaya, avocado, strawberry, banana, and mango [7,8]. Earlier studies have shown that chitosan causes growth inhibition in C. asianum, C. fructicola, C. tropicalis, C. karstii, C. lupini, C. nymphaeae, C. acutatum, C. musae, and C. siamense [9,10,11,12]. The antifungal efficacy of chitosan can be improved at the nanoscale level, attributable to the amplified contact surface area, which yields a heightened area–volume ratio when compared to micrometric particles [13]. In the nanotechnology field, a nanoparticle is 1–100 nm in size regardless of its morphology [14,15]. In the context of polymeric nanomaterials, the size may extend up to 1000 nanometers [16,17]. Chitosan nanoparticles, or nanochitosan, have been reported with sizes between 40–600 nm, according to the original features and the synthesis method at the nanoscale [18].

Ultrasonication is one of the methods studied for the synthesis of nanochitosan. This method uses high-frequency sonic waves (>20 kHz), and its main mechanism is based on acoustic cavitation. It creates bubble formation, growth, and implosion, resulting in shear forces capable of breaking the chemical bonds of the chitosan polymer [19].

Smaller particle diameters and reduced polydispersity index increase nanochitosan biological activity [20]. Ultrasonication makes these properties easily possible due to the ease of controlling parameters such as frequency, ultrasonic power, sonication times, and temperature [19]. Therefore, the ultrasound method provides an environmentally friendly and simple solution, avoiding the use of additional substances, mostly toxic, requiring a subsequent stage of washing or purification of the suspended particles [20].

Few studies evaluate only chitosan nanometric particles in fungi [3,6,21]. This study aimed to synthesize chitosan nanometric particles by ultrasonication and evaluate their antifungal activity in C. chrysophillum and C. musae, both species of the Colletotrichum gloeosporioides complex.

2. Materials and Methods

2.1. Materials

Chitosan particles were made from chitosan (CS) (Sigma-Aldrich, lot #STBF3282V, Saint Louis, MO, USA), with a molecular weight of 32.0 kDa and a degree of deacetylation of 78% [22], with glacial acetic acid (J.T. Baker, Phillipsburg, NJ, USA), ultrapure water, and NaOH (1 and 5 M). Fluorescent dyes acquired from Sigma-Aldrich (St. Louis, MO, USA) were fluorescent brightener (calcofluor white), propidium iodide (>94% purity), and 2,7-dichlorodihydrofluorescein diacetate (>94% purity).

2.2. Microorganisms

Colletotrichum musae (H5-5) and Colletotrichum chrysophillum (H1-2) were previously isolated and identified [22]. The fungi were activated in 125 mL flasks with 50 mL of potato dextrose agar culture medium (PDA, Bioxon^®, Franklin Lakes, NJ, USA) and incubated at 25 °C with a 12 h photoperiod for seven days. Subsequently, mechanical stress was generated with a soft scrape on the mycelium using a microbiological loop, and they were incubated for an additional three days to favor sporulation.

2.3. Synthesis and Characterization of Chitosan Particles

A 100 mL solution of ultrapure water acidified with acetic acid (pH 3.5 and 0.03 M acetic acid) was mixed with 0.2 g of chitosan using magnetic stirring at 500 rpm (SH Digital HotPlate Stirrer FAGA) for 10 min. Subsequently, the pH was adjusted to 5.4 ± 0.2 (NaOH, 1 and 5 M). The chitosan solution was ultrasonicated at a frequency of 42 kHz and ultrasonic power of 70 W for 60 min (Branson 1510 Ultrasonic Cleaner) [20] and autoclaved (121 °C, 15 lb, 15 min). Finally, the chitosan particles solution was stored at 4 °C (7 days maximum). From this section onwards, the solution of ultrasonicated chitosan particles will be referred to as nanochitosan.

2.3.1. Particle Size, Polydispersity Index, and Zeta Potential (ζ)

The particle size, dispersion, and zeta potential of the nanochitosan were calculated by dynamic light scattering (DLS) (Möbiuz equipment, Wyatt Technology Corp., Santa Barbara, CA, USA) at 25 °C. The chitosan solution (20 μL) was deposited in the chamber of equipment, adapted with a vertically polarized laser. The detection angle was kept at 90° with respect to the incident light beam, and the wavelength was 488 nm (2 W). Subsequently, the averages of three repetitions were estimated for 60 s at room temperature [23].

2.3.2. Fourier Transform Infrared Analysis

The spectra of the samples were obtained by Fourier transform infrared (FT-IR) spectroscopy using a Nicolet Protegé 460 spectrophotometer (Instrument Corp., Madison, WI, USA) to identify the characteristic functional groups of chitosan. Measurements were made in 2 mg mL⁻¹ nanochitosan solutions at 25 °C, taking an average of 64 scans in a spectral range of 4000–600 cm⁻¹. Additionally, the degree of deacetylation (DD) was calculated based on the model proposed by Brugnerotto et al. [23] using the spectra at 1320 cm⁻¹ (amine III) and 1420 cm⁻¹ (-CH₂).

2.3.3. Morphology of Chitosan Particles

The nanochitosan particle morphology and size were determined using a transmission electron microscope at 120 kV (FEI Tecnai Spirit, Hillsboro, OR, USA). A 1:100 dilution with nanochitosan at 2 mg mL⁻¹ was made and sonicated (Bradson M1800 50–60 HZ) for 5 min to disperse. Subsequently, 20 microliths of the solution were placed on a microporous copper TEM (200 mesh) and dried for 48 h. Micrographs were acquired using a Veleta Olympus Soft Imaging camera.

2.4. Radial Growth

Nanochitosan (0.1, 0.5, 0.75, 1, and 1.25 mg mL⁻¹) was mixed with sterile PDA culture medium at 45 °C, deposited in 15 mL of the mixture in Petri dishes (60 mm diameter), and cooled until solid. Subsequently, a hole (6 mm diameter) was made in the center of each culture medium using a sterile disposable tip for a micropipette (1 mL capacity). The hole was inoculated with 100 µL of each mycelium suspension of C. chrysophillum and C. musae in Tween 80 solutions (0.05%). Mycelium suspensions were obtained from a 10-day-old culture in PDA medium at 25 °C and 12 h photoperiods. A PDA control without nanochitosan was used. All Petri dishes were incubated at 25 °C, photoperiod of 12 h, and relative humidity (HR ≈ 100). The radius of the colonies was measured manually every 12 h until the mycelium covered 95% of the surface of the culture medium in the PDA control [24]. Four measurements of the radius were carried out from each treatment. In addition, the percentage of mycelial inhibition at 96 h was calculated online according to Quintana-Obregón et al. [25], and the radial extension velocity (Vm) was estimated using the Gompertz model.

2.5. Morphometric Analysis of Hyphaes

The diameters of hyphae from at least 60 measured samples were obtained (optical microscope, Olympus CX31, Tokyo, Japan) with a camera (Infinity 1; Lumenera Corp., Ottawa, ON, CA) at 400× magnification using Image-Pro Plus version 6.3 software (2008 Media Cybernetics, Inc., Bethesda, MD, USA). The suspension of fungal mycelium was deposited in a culture medium with and without nanochitosan (1 mg mL⁻¹) in Petri dishes and incubated at 25 °C with photoperiods of 12 h on a humidified surface, as described previously. The terminal hyphal width was measured at 0, 12, 16, 20, and 24 h [26].

2.6. Fluorescence Microscopy

Staining and observation under an inverted microscope (Model DMi8, Leica Microsystems, Wetzlar, HE, Germany) and fluorescence filters (excitation 546/10 RHOD filter and emission 585/40, excitation filter 350/50 DAPI and emission 460/40, and FITC excitation filter 480/40 and FITC filter 527/30) allowed researchers to assess the damage that nanochitosan (1 mg mL⁻¹) caused to the membrane. Digital images were obtained with a cooled monochrome DFC 450C camera (Leica Microsystems, Wetzlar, HE, Germany) and fluorescence overlay software (LAS AF version 3.1.0, Leica Microsystems, CMS GmbH, Mannheim, LfDI, Germany).

Assays were performed in a 12-well microplate. A volume of 500 µL of inoculum mixed with Czapek medium and Tween 80 (0.05%) was deposited in each well. Each treatment was assayed two times. Subsequently, 500 μL of nanochitosan solution (1 mg mL⁻¹) was added. The microplate was incubated for 12 h at 28 ± 2 °C. After incubation, the supernatant was extracted, and fluorescent dyes were added [26]. The dyes used were propidium iodide, calcofluor, and 2′,7′-dichlorofluorescein diacetate.

2.6.1. Plasma Membrane Integrity

The propidium iodide (PI) staining method (P4170, Sigma, USA) was used to determine the changes in the integrity of the plasma membrane. PI is a universal marker used to assess plasma membrane permeability. Generally, PI can penetrate apoptotic or damaged cells through defective membranes, emitting red fluorescence because of intercalation with some DNA bases. Ten microliters of the PI solution at 3 µM (w v⁻¹) were deposited in the fungal cells and incubated for 30 min in refrigerated and dark conditions [27].

2.6.2. Cell Wall Integrity

The presence of chitosan as a disturbing agent of the fungal cell wall and septa was determined using Fluorescent Brightener 28 (calcofluor white, Sigma-Aldrich, St. Louis, MO, USA), a dye commonly used to identify polysaccharides of the fungal cytoskeleton, emitting blue fluorescence when bound with β (1,4) glucans such as chitin, chitosan, and cellulose. The samples with the previously incubated cells in each well were washed with PBS, and the supernatant was subsequently removed. Next, 10 µL of the 10 µL mL⁻¹ (w v ⁻¹) calcofluor white solution was added and incubated for 30 min at 4 °C in the dark [21].

2.6.3. Oxidative Stress

To determine whether nanochitosan produces intracellular reactive oxygen species (ROS), 2′,7′dichlorofluorescein diacetate (2′,7′-dichlorodihydrofluorescein diacetate, or 2,7-DCFH2-DA, D6883, Sigma, USA) was used, a dye capable of diffusing through the plasma membrane of cells and being hydrolyzed by intracellular esterase enzymes, forming a nonfluorescent molecule. When there is increased intracellular ROS production, 2,7-DCFH2-DA deesterifies to 2′,7′-dichlorofluorescein due to oxidation, emitting intense green fluorescence. To each well with the samples, 10 µL of the dye solution at 10 µM (w/v) was added. The samples were incubated for 30 min at 4 °C in the dark [21].

2.7. Statistical Analysis

A randomized unifactorial block design was used to test five concentrations of nanochitosan. Radial growth analysis (n = 4) and morphometric analysis (n = 60) were performed. The design was determined using the one-way analysis of variance (ANOVA) at a significance level of p = 0.05. The differences between concentrations and fungus species were estimated using the Tukey multiple comparisons test (Tukey’s post hoc test) at a 95% confidence interval procedure in the Infostat Version 2020 software. All data were presented as mean values with their standard deviation indicated (mean ± SD). Differences were accepted as significant when p ≤ 0.05.

3. Results and Discussion

3.1. Characterization of Chitosan Particles

3.1.1. Particle Size, Polydispersity Index, and Zeta Potential (ζ)

Figure 1 shows the particle size distribution of nanochitosan after ultrasonication treatment for 60 min. The nanochitosan exhibited a bimodal distribution; the predominant population of nanochitosan (95%) had an average diameter of 302.4 ± 92.3 nm, while the other had less intensity (≈15 nm) (Figure 1). The nanometric scale achieved by using ultrasound may be due to acoustic cavitation. This phenomenon creates bubbles or cavities that grow and implode, generating strong forces on the chitosan molecules. As a result, the 1,4-glucosidic bonds break, and the clumps produced by the Brownian motion of chitosan particles dissociate throughout the solution. Furthermore, the sonic waves increase the temperature, favoring chitosan’s depolymerization; this process causes chitosan to breakdown into smaller particles [28,29].

Previous studies have successfully used the ultrasound method to synthesize chitosan nanoparticles. Diameters of 538 nm have been reported using a frequency of 20 kHz and an irradiation power of 750 W for 1.4 min [29]. In contrast, other studies have achieved particle sizes below 30 nm, using a frequency of 24 kHz and a power of 360 W [30]. In their research, the smaller size achieved compared to that reported by Salehi et al. [29] may be attributed to the chitosan used, derived from a 300–400 nm chitin obtained by exposure to ultrasound for 2 h. This suggests that a longer exposure time to ultrasonic radiation increases cavitation energy, which facilitates particle size reduction.

Esmaeilzadeh-Gharedaghi et al. [20] successfully obtained chitosan nanoparticles with a diameter of 300 nm using a sonication frequency of 20 kHz. Their study reported an inverse relationship between sonication time and particle size, identifying a critical time at which the minimum particle size is achieved. It was noted that beyond this crucial point, no further size reduction occurred. This minimum size was attributed to the presence of bonds requiring greater cavitation intensity to break. Similarly, Khoerunnisa et al. [31] achieved chitosan nanoparticles with a size of 426 nm after 5 min of sonic irradiation. Increasing the sonication time to 60 and 120 min reduced the particle diameters to 220 nm and 109 nm, respectively, with unimodal size distribution. These results, obtained using a frequency of 20 kHz and a power of 750 W, demonstrated the significant impact of irradiation time on particle size reduction.

Compared with our study, sonic irradiation with a higher frequency (42 kHz) and lower power (70 W) was used for 60 min. Although the prolonged exposure time facilitated size reduction within the 300 nm range, a narrow size distribution, as previously reported by Khoerunnisa et al. [31], was not achieved. This may primarily be attributed to the combination of low power and high frequency used, which generates less intense cavitation compared to lower frequencies (<20 kHz). Under these conditions, cavitation bubbles are smaller and collapse more easily, reducing the time to act on chitosan chains [32,33].

The zeta potential is a measure that defines the electrostatic potential at the imaginary hydrodynamic shear surface (at the particle surface), facilitating interactions between particles or with other surfaces. It is frequently used to theoretically deduce the surface charge of nanoparticles and analyze their behavior in suspension, biological interactions, and potential toxicity [34].

The zeta potential obtained in our study was +35.9 ± 2.3 mV. It has been reported that zeta potential values greater than 30 mV provide greater electrostatic repulsion between polymer particles in aqueous media, avoiding aggregation [34,35,36].

The antifungal activity of chitosan nanoparticles can also be interpreted through the zeta potential value. Some studies have indicated that particles with high zeta potential values and positive charge promote interaction with the charges on the surface of microorganism membranes, affecting their growth [3,21,26]. Regarding the effects of nanochitosan observed in our study on fungal species, the most significant impact was observed on the membranes and cell walls of the fungi, consistent with findings reported in previous research.

3.1.2. Fourier Transform Infrared Analysis

FT-IR spectroscopy was used to analyze the degree of effect of ultrasound treatment on the bonds in the molecular structure of chitosan. The infrared absorbance spectra of the chitosan (lot #STBF3282V) used in this study were reported by Valenzuela-Ortiz et al. [22]. According to the results, the characteristic absorption bands coincide with those of the previously reported chitosan (Figure 2). However, after ultrasonic wave treatment, a decrease in the intensity of the absorption bands of the residual N-acetyl group (–CH and –NH) was observed, as well as a slight signal in the region of 1039–1159 cm⁻¹ corresponding to the vibrations of the symmetrical stretching of the primary amine (–CN), secondary amine (–CO), and the alcoholic group (–COH). A slight absorption peak at 1451 cm⁻¹ is associated with CH₂ stretching. The peaks corresponding to the NH₂ group of the polymer’s amide II were observed at 1613 cm⁻¹. A slight peak at 1649 cm⁻¹ was also observed due to the C=O stretching vibration of the amide I bonds [30]. In addition, a wide band at 3293–3356 cm⁻¹ was found, indicating the presence of –OH stretching. The reduction in the absorption intensities of certain characteristic groups of chitosan after ultrasound treatment is related to the time of exposure to the ultrasound, suggesting degradation of the polymer. Vallejo-Domínguez et al. [37] used ultrasound for the deproteinization of chitin to chitosan, finding a significant reduction in the intensity of the peaks of some characteristic groups of chitosan, especially at longer sonication times. Meanwhile, Suryani et al. [38] reported that the structure of chitosan remains stable during the ultrasound depolymerization process. However, there was also a decrease in the intensity of the peaks related to the acetyl group compared to chitosan without treatment.

The DD of the nanoparticles was 82.1 ± 0.09%. Previously, Valenzuela-Ortiz et al. [22] reported a DD of 78% for the chitosan (lot STBF3282V) used in this study to elaborate nanochitosan; the increase in the degree of deacetylation after nanoparticle formation may be related to the depolymerization induced during the ultrasonication–autoclaving process [37]. Baxter et al. [39] reported a slight 5–10% increase in DD after sonication. Similarly, a DD% of up to 100% was reported in sonicated chitosan samples [39]. The variability could be attributed to the intensity of the emission of sonic waves and the treatment times, in addition to the characteristics of chitosan, as previously discussed in Section 3.1.1.

3.1.3. Morphology of Chitosan Particles

The size and shape of the chitosan nanoparticles are part of the characteristics that confer the biopolymer’s antimicrobial effect [40]. Transmission electron microscopy (TEM) analysis showed amorphous nanochitosan particles with sizes ranging between 30 and 200 nm. Figure 3 displays variations in the size of the population. The operation of the TEM equipment requires dry nanoparticle samples, resulting in the appearance of rough surfaces and a smaller particle size than that obtained by DLS, where the sample is used in suspension, which can lead to particle swelling and larger size.

The difference in sizes obtained during sonication can be affected by the force variation in the intermolecular bonds of chitosan, easily breaking the weaker bonds or interactions and resisting those of higher energy [20]. The results confirmed that the physical ultrasonication technique and autoclaving effectively reduced the particle size.

3.2. Radial Growth

The data showed that nanochitosan caused a slight decrease in the development of mycelial during the first hours of growth of C. chrysophillum (Figure 4a). The same pattern was shown in C. musae; nanochitosan caused a reduction in radial growth, concentrations of 1 and 1.5 mg mL⁻¹ showing the most significant impact (Figure 4b).

At 96 h, nanochitosan showed an inhibition (p ≤ 0.05) on the radius of the fungal sample vs. control (

Table 1. Effect of nanochitosan on radial growth of C. chrysophillum and C. musae to 96 h (25 °C and 12 h photoperiod).

Nanochitosan (mg mL−1)	Inhibition Percentage of Radial Growth *
	C. chrysophillum	C. musae
0.1	9.15 ± 1.58 Aa	0 ± 0.00 Ab
0.5	16.46 ± 1.58 Bb	6.19 ± 2.43 Ba
0.75	17.07 ± 1.73 Bb	12.83 ± 3.42 Ca
1	20.12 ± 0.7 Ca	24.19 ± 1.99 Db
1.5	21.34 ± 0.7 Ca	25.66 ± 4.24 Da

* Values correspond to the mean ± standard deviation. The uppercase superscript indicates statistically different groups in the columns (concentration effect of nanochitosan for each species). Lowercase letters indicate statistically different groups between the species according to Tukey’s test (p ≤ 0.05).

). However, none presented more than 26% inhibition.

Table 2. Radial growth velocity (Vm) of C. chrysophillum and C. musae (25 °C, photoperiod 12 h).

Treatment	C. chrysophillum	C. musae
	* Vm (h−1)
Control PDA	0.30 ± 0.01 Bb	0.24 ± 0.003 Ba
NCS0.1	0.28 ± 0.01 Bb	0.21 ± 0.001 Aa
NCS0.5	0.26 ± 0.004 Ab	0.20 ± 0.01 Aa
NCS0.75	0.27 ± 0.01 Aa	0.28 ± 0.04 Ba
NCS1	0.27 ± 0.003 Aa	0.21 ± 0.06 Aa
NCS1.5	0.26 ± 0.0008 Aa	0.28 ± 0.09 ABa

PDA: potato dextrose agar, NCS: nanochitosan. * Values correspond to the mean ± standard deviation. The uppercase superscript indicates statistically different groups in the columns (the concentration effect of nanochitosan for each species is reported as mg mL⁻¹). The lowercase letters indicate statistically different groups between the species according to Tukey’s test (p ≤ 0.05).

shows the radial growth velocity (Vm), and significant differences (p < 0.05) concerning the control were found in the two species.

Numerous studies carried out with chitosan-based nanoparticles have obtained antifungal effects above 70% for fungi, for example, Aspergillus parasiticus [3], Fusarium oxysporum [41], and Sclerotinia sclerotiorum [42], evaluated during the mycelial growth stage. In contrast, Colletotrichum species exhibit inhibitions below 30% [42,43,44]. The fungicidal or fungistatic effect of chitosan relies on characteristics such as the nature of the microorganism and growth stage, the concentration, DD, molecular weight, and particle size [45,46]. It has been previously reported that Colletotrichum species exhibit low sensitivity to chitosan [22]. The variation in the sensitivity of each species to nanochitosan may be related to their ability to produce fewer fluid membranes and enzymes, such as chitin deacetylase or chitosanase, which assist in degrading chitosan. It has been demonstrated that chitosan can modulate the expression of chitosan-related genes and accelerate the development of certain fungi. Some studies have shown that the variation in sensitivity to chitosan among various fungi may be related to the lipid components of the plasma membrane, mainly fatty acids, reporting that fungal membranes rich in saturated fatty acids are less fluid and have less sensitivity. In contrast, the membranes of fungi with a low content of saturated fatty acids show greater fluidity and, therefore, greater sensitivity to chitosan [47]. In addition to the stress response of each fungus to tolerate the main effects of chitosan, the modification of genes related to the integrity of the cell membrane and wall, the variation in metabolism, and the production of reactive oxygen species that mainly affect the plasmatic membrane and its permeabilization are some of the factors responsible for the antifungal effect of chitosan [48].

3.3. Morphometric Analysis

The morphometric analysis of the species was evaluated with nanochitosan (1 mg mL⁻¹). Figure 5 shows that nanochitosan affected the size of the hyphae of C. chrysophillum and C. musae (25 °C). The nanochitosan showed structural damage for the two species, increasing the (p < 0.05) diameter of the hyphae compared to the control during the range of hours evaluated. The greatest affectation observed was for C. musae (12 h), obtaining a value of 4.75 ± 0.46 μm, an increase of 38% concerning the control.

Galvez-Iriqui et al. [21] found irregularities in the hyphae morphometry of A. niger exposed to chitosan—pyrrole-2-carboxylic acid biocomposite, and, moreover, to an increase in the diameter of the hyphae. They suggest that the increase in the hyphae diameter may be due to the loss of turgor due to the passage of liquids into the cell, caused by an osmotic imbalance attributed to the fungus–chitosan interaction. The development of hyphae involves a process of nutrient delivery through transport vesicles, which, under stress conditions, can lose coordination with the structural function [49].

At 12–20 h of incubation, the shape of the mycelium of C. chrysophillum and C. musae changed, showing bulges along the hyphae (Figure 6a,b, respectively).

It should be emphasized that hyphae growth relies on a nutrient supply process facilitated by transport vesicles. However, unfavorable conditions can cause these vesicles to lose synchronicity with the structural role, causing morphological changes, protuberances, and distortion in the terminal hyphae growth [11,49]. It is important to note that these conjectures need further research for confirmation. Chitosan adheres to fungal plasma membranes by interacting with extracellular components.

The relationship between the positive charge of chitosan’s free amino acid groups and the negatively charged components of cell membranes causes structural and metabolic damage. The ergosterol content, glycosphingolipid acids, and phosphatidylglycerol of phospholipids are the main characteristic components of the cell membrane associated with chitosan binding [50].

3.4. Fluorescence Microscopy Analysis

3.4.1. Plasma Membrane Integrity

Figure 7 shows that the fluorescence emanating from the hyphae of C. chrysophillum and C. musae was more significant when exposed to nanochitosan. The impact of 1 mg mL⁻¹ nanochitosan on the plasma membrane of the Colletotrichum species was determined by monitoring PI dye uptake. The permeability of PI within the cell indicates an unstable, lost, or destroyed membrane structure [51], consistent with previous studies on Colletotrichum [43]. The fluorescence intensity reflects the changes in membrane permeability of the species when exposed to nanochitosan. The increase in PI intensity in isolates exposed to chitosan can be attributed to factors such as leakage of essential intracellular ions and alterations in the morphology of the mycelium, which have been discussed in detail in an earlier section of morphometric analysis. Furthermore, according to Meng et al. [51], chitosan stimulates the expression of enzymes that break down the main components of the membrane, such as phospholipids, ergosterols, and sterols, implying that chitosan affects the fluidity and integrity of the membrane.

3.4.2. Cell Wall Integrity

In the nanochitosan (1 mg mL⁻¹), the cell walls of C. chrysophillum and C. musae were distorted, granulated, and had a lower fluorescence intensity (Figure 8). The fungal cell wall contains glucans and chitin that maintain its integrity and structure [51]. Calcofluor white allows us to see if the chitin–calcofluor or chitin–glucan interaction changes the way these polysaccharides are spread out. Likewise, chitosan has been shown to exert fungal intracellular activity and affect the expression of proteins that maintain cell wall integrity [52]. Previous studies on Fusarium oxysporum and Aspergillus parasiticus observed morphological and microstructural changes caused by chitosan and caused by alterations in the content of hydrolysis proteins or chitin and glucan synthesis, which were evidenced by the intensity of calcofluor staining [26,51]. Therefore, it can be hypothesized that the morphological and growth alterations in C. chrysophillum and C. musae exposed to nanochitosan are manifested by alterations in the expression of cell wall proteins. However, further research is needed at the gene expression level to detect the specific action of chitosan on the cell surface of fungi such as Colletotrichum.

3.4.3. Oxidative Stress

Fluorescent staining (2′,7′dichlorofluorescein diacetate) was used to observe the effect of chitosan particles on the production of oxidative stress in fungal hyphae. The formation of reactive oxygen species (ROS) induced by electron transfer forms highly oxidizing radicals, such as superoxide, hydrogen peroxide, and the hydroxyl radical [53]. Our results show that exposure of the fungus C. chrysophillum to nanochitosan decreased fluorescence compared to the control group. Naturally, the production of intracellular reactive oxygen species (ROS) is a byproduct of certain metabolic pathways, and antioxidant molecules regulate this process [54]. This suggests that treatment with nanochitosan reduced ROS levels in the cells of C. chrysophillum.

However, in the case of C. musae, there was no significant difference in ROS production. From this, it can be concluded that a concentration of 1 mg mL⁻¹ of nanochitosan does not cause oxidative stress in C. musae.

When the balance between the production and natural elimination of intracellular ROS is disrupted, they can react with macromolecules such as DNA, proteins, lipids, and carbohydrates, causing irreversible damage that can ultimately lead to cell death [54]. This effect has been documented in Aspergillus parasiticus, where chitosan nanoparticles increased the generation of free radicals in a concentration-dependent manner [55].

An inverse correlation was noted between the production of ROS and fungal cell viability, with higher ROS production due to an increased concentration of nanochitosan (0.6 mg mL⁻¹) leading to reduced spore viability [55]. The same effect was observed when chitosan was applied to inhibit the growth of Sphaeropsis sapinea, a fungus commonly found in wood. In this study, the formation of superoxide and peroxide radicals was observed as the chitosan concentration in the medium (1 mg mL⁻¹) increased, resulting in a decrease in fungal growth rate and severe alterations to the cellular structure [56].

Similar results were obtained for Neurospora crassa, which suggests that chitosan causes an intracellular burst of ROS by oxidizing free fatty acids from the membranes of fungi sensitive to chitosan, generating permeabilization of the plasma [57]. According to their hypothesis, the results obtained from the PI staining assay regarding the alteration of cell membranes, sensitivity, and damage caused by nanochitosan on the membranes of C. chrysophillum and C. musae are not associated with the induction of intracellular ROS.

4. Conclusions

The study discovered that nanochitosan, synthesized through sonication, damaged Colletotrichum species’ cell walls and membranes. However, it did not produce significant oxidative stress. This suggests that nanochitosan affects the cell structure as a sensitivity parameter. Future research efforts should identify the exact defense mechanism of fungi and determine the degree of their sensitivity to nanochitosan.