1. Introduction
Filamentous fungi are microorganisms of greatest concern for food safety as lower eukaryotic heterotrophic organisms that rely on external carbon sources [1]. These microorganisms are responsible for several plant diseases and cause the spoilage of fruits, vegetables, and grains, resulting in huge economic losses [2]. After spore germination, the fungi directly penetrate the plant epidermis through natural openings or wounds, as well as through specialized structures such as appressoria. They either attach to the fruit surface for penetration or use enzymes to degrade the plant cell walls [3]. Two of the recurrent filamentous fungi responsible for fruit spoilage are Botrytis cinerea and Rhizopus sp., which cause significant losses, particularly in strawberry, tomato, and grape cultivars [4]. There are few alternatives for controlling fungi in food due to either chemical residues or challenges in application. Therefore, it is essential to develop innovative solutions to reduce fungal contamination in fresh produce, substrates, and food products [4,5].
The natural antimicrobials currently receiving attention are the essential oils (EOs) [6,7]. EOs are derived from natural plant-based raw materials through various extraction processes, including mechanical (citrus fruits, orange, lemon, bergamot, lime, tangerine, and grapefruit), physical (aromatic plants, basil, and thyme), and chemical methods (fragile components of flowers, not separable by heat), as well as dry distillation (woods, barks, roots, or gums). Steam distillation is the most common extraction method, and it has been employed for obtaining several EOs, such as herbal leaves, mint, oregano, and tea tree [8]. EOs contain a complex mixture of natural polar and non-polar substances, volatile compounds, terpenes, including monoterpenes and sesquiterpenes, aromatic compounds (such as methoxy derivatives, aldehydes, alcohols, and phenols), and terpenoids [4,9].
The plant Baccharis dracunculifolia (Figure 1) belongs to the Asteraceae family and the Baccharis genus, which includes approximately 500 species. It is widely distributed in South America, particularly in the southern region of Brazil, and in countries such as Argentina, Uruguay, Paraguay, and Bolivia [10,11]. This plant is culturally regarded as a source of natural compounds and is traditionally used in the treatment of various diseases [6,11]. Moreover, it has been demonstrated that its secondary metabolites possess antimicrobial, antioxidant, antiparasitic, anti-inflammatory, and antiviral activities [11,12,13,14,15]. The natural sesquiterpene alcohol nerolidol (3,7,11-trimethyl-1,6,10-dodecatrien-3-ol) is the main compound found in the EO of B. dracunculifolia and is permitted by the US Food and Drug Administration (FDA) as a safe flavoring agent in foods [12,16]. Nerolidol has a wide range of pharmacological and biological activities attributed to it, such as antioxidant, antibacterial, antibiofilm, insecticide, and antiparasitic properties, being of great interest in the field of agriculture and medicine [16].
There has been a growing demand from consumers for foods without synthetic additives and preservatives [17]. The food industry has significant interest in EOs and their constituents, as the use of these bioactive compounds as natural antimicrobial agents has been described in various studies [18,19,20]. However, the interaction of pure EO with the food matrix can cause undesirable organoleptic effects when added in quantities sufficient to provide an antimicrobial effect [21]. In this context, nanoencapsulation appears among the alternatives for delivering natural antimicrobials [22]. Therefore, applying EOs in the form of nanoparticles can mask the sensory attributes of the EOs and their components, enhancing apparent solubility and stability while reducing interaction with the food matrix. This approach has the potential to increase biological activity, bioaccessibility, and bioavailability [22,23].
Among the polymers used in nanotechnology, the triblock copolymer Pluronic® F-127 appears as an interesting material in developing nanoparticles with antibacterial activity against foodborne bacteria [24,25], being effective in encapsulating geraniol to control enteric bacterial pathogens on spinach surfaces [26]. It is an inexpensive polymer with high biocompatibility and low toxicity, approved by the FDA for use in various pharmaceutical applications, including oral products [24,27]. However, its application for the delivery of natural compounds with activity against food spoilage fungi has been poorly explored [28,29]. Therefore, the aim of this study was to evaluate the inhibition of B. cinerea and Rhizopus sp. using Pluronic® F-127 nanoparticles loaded with EO from B. dracunculifolia and its major compound, nerolidol (NE).
2. Materials and Methods
2.1. Chemicals
The B. dracunculifolia EO was obtained from Harmonia Natural (Canelinha, Brazil). Nerolidol (98%, mixture of cis and trans), Pluronic® F-127 (powder, BioReagent, suitable for cell culture), 2,2-diphenyl-1-picrylhydrazyl (DPPH) and 2,2′-azino-bis-(3-ethylbenzothiazoline)-6-sulfonic acid (ABTS) radicals, and 6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid (Trolox) were purchased from Sigma Aldrich (St. Louis, MO, USA). Tetrahydrofuran (THF) and Potato-Dextrose agar (PDA) were obtained from Merck (Darmstadt, Germany). Polysorbate 80 (Tween 80®) was acquired from Synth (Diadema, Brazil). Capric/caprylic triglyceride was provided by Delaware (Porto Alegre, Brazil). Ultrapure water was obtained using a Milli-Q purifier (Millipore, Burlington, MA, USA). All other reagents were of analytical grade, and ultrapure water was used to prepare all solutions.
2.2. Microorganisms
The filamentous fungi Botrytis cinerea and Rhizopus sp. were isolated from grape and tomato, respectively. The strains were obtained from the culture collection of the Laboratory of Toxicology (ICTA-UFRGS, Porto Alegre, Brazil). Fungal identification was performed as described elsewhere [30] based on macroscopic and microscopic morphological criteria and molecular analyses of the internal transcribed spacer (ITS) region and part of beta-tubulin gene. PDA plates were inoculated and incubated at 25 °C for 7 days or 15 days for mycelium and spore analyses, respectively [31]. The isolates were maintained on PDA agar plates under refrigeration (7 °C) and subcultured in a fresh plate prior to each experiment.
2.3. Characterization of the EO
The manufacturer provided the chemical characterization and quantification of the EO compounds. The analysis was performed through gas chromatography coupled with mass spectrometry using Agilent MSD5977B equipment (Agilent, Santa Clara, CA, USA). Compounds were separated using a DB-5MS capillary column (30 m × 0.25 mm × 0.25 µm). The injector and detector temperatures were set at 280 °C and 260 °C, respectively. The oven temperature gradient was as follows: an initial temperature of 60 °C (2 min) with a rate of 4 °C/min to 200 °C, and then a rate of 6 °C/min to 260 °C (10 min). The ionization source temperature was 280 °C, and the acquisition mode was scan. A 1.0 μL aliquot was injected using the split mode (split ratio, 1:20). Compound identification was based on the comparison of mass spectra of peaks with those in the NIST17.L library (NIST Chemistry WebBook), with the similarity degree of each identification presented in the results table. The relative percentage area of each peak was calculated based on the sum of the areas of all peaks eluted from the column and originating from the analyzed sample, including peaks identified as “unidentified compounds” due to similarity values below reliable identification thresholds.
2.4. Antifungal Activity of EO and NE
2.4.1. Agar Contact Method
Antifungal activity was assessed using the direct contact method on agar [31] with modifications. The EO or NE was added to PDA (containing 0.05% Tween 80) at concentrations of 0, 1, 5, 10, 20, and 25 mg/mL. Then, 15 mL of the PDA solution was poured into sterile Petri dishes (90 mm in diameter). A mycelial disc (3 mm in diameter) was removed from 7-day-old cultures on PDA plates using a punch and placed in the center of each Petri dish. The Petri dishes were incubated for 7 days at 25 ± 2 °C, with measurements of mycelial growth taken at 72 h, 5 days, and 7 days in two perpendicular directions (diameter in mm). Comparison of the obtained dimensions with those of the controls allowed for the calculation of the percentage of inhibition (IP %) at the end of the incubation period, according to Equation (1):
where C: average colony diameter (mm) in the control; T: average colony diameter (mm) in the treatment.
2.4.2. Exposure to Volatiles
The antifungal activity of the volatiles from the EO and NE was also evaluated as described elsewhere [32]. Volumes of 5, 10, 20, and 25 µL were applied to filter paper discs (16 mm in diameter), which were then fixed to the center of the inner lid of Petri dishes containing solidified PDA. The plates were sealed with Parafilm to ensure a controlled environment. The volatiles were released from the impregnated paper discs into the dish during an incubation period of 7 days at 25 ± 2 °C. The pathogen inoculation, incubation, and growth measurement procedures were the same as those described for the direct contact method.
2.5. Preparation of Nanoparticles
The organic phase of the nanoparticle (NP) solution consisted of Pluronic diluted 1:1 (w/w) in EO or NE, followed by the addition of this solution to tetrahydrofuran (THF) at a ratio of 16% (w/w). The mixture was emulsified using a probe-type ultrasonic device (Unique OF S500, Unique, Americana, Brazil) operating at frequency 50 kHz, power 250 W, for two cycles of 5 min each with 2-min intervals [26]. The aqueous phase of the solution consisted of sterile ultrapure water. The organic phase was then added to the aqueous phase by the nanoprecipitation method at a 1:10 (w/w) ratio, followed by sonication for three additional cycles of 5 min with 2-min intervals to ensure complete dilution (Figure 2). The NP solution was subsequently placed under a hood for 20 h with moderate agitation to remove the THF. The NPs were collected, filtered through 0.22 μm membranes (Millipore, Billerica, MA, USA), and stored in sterilized glass containers at 25 °C before further use. In addition to the NPs containing EO (NP-EO) or NE (NP-NE), control formulations were prepared by replacing the EO/NE with caprylic/capric acid triglycerides (NP-B).
2.6. Characterization of the Nanoparticles
2.6.1. Particle Size and Zeta Potential
To determine the size (average diameter), polydispersity index (PDI), and zeta potential (ζ), the NP suspensions were diluted in ultrapure water (1:10, v/v). The size was measured using Dynamic Light Scattering (DLS) with a goniometer (BI200SM, λ = 632.8 nm, 75 mW He-Ne laser, and BI 9000 correlator; Brookhaven, Nashua, NH, USA) at a fixed angle (θ = 90°). Data processing was performed using the NNLS. The cumulant method was used for PDI determination. The zeta potential was measured by electrophoretic mobility using a ZetaPALS Potential Analyzer (Brookhaven Instruments, Nashua, NH, USA) at 25 °C.
2.6.2. Encapsulation Efficiency
Encapsulation efficiency (EE%) was determined in triplicate using the centrifugation filtration technique [33]. To determine the EE, it was necessary to identify the wavelength of maximum absorbance (λmax) for the EO and NE and to construct the standard curve. The λmax values were obtained by diluting the EO in methanol, followed by scanning with a UV/VIS spectrophotometer, where the maximum absorbance was detected at λ = 293 nm and λ = 262 nm for EO and NE, respectively. The standard curve was generated by diluting the EO or NE to concentrations of 0.0015, 0.003, 0.006, 0.012, and 0.025 mg/mL in methanol. The correlation coefficient (r2) was 0.9919 with the equation y = 2.9797x + 0.0011 for EO, and r2 was 0.9966 with the equation y = 3.78084x − 0.0054 for NE.
For the determination of EE, 500 μL of the NP suspension was added to centrifuge filters (Ultracel®-3k, Merck Millipore Ltd., Tullagreen, Ireland), followed by centrifugation at 10,000× g for 30 min at 4 °C. After centrifugation, the supernatant was diluted in methanol, and absorbance was measured using UV/VIS spectrophotometry at 293 nm and 262 nm for EO and NE, respectively, and quantified using the standard curve. The EE was calculated using the following Equation (2):
where NP is the total amount of EO or NE used in the preparation of the NPs (mg/mL), and S is the total amount of EO or NE in the supernatant (mg/mL).
2.6.3. Determination of pH
The pH of the NP suspensions was measured using a benchtop pH meter (Model HI 2221; Hanna Instruments, Smithfield, RI, USA), calibrated with standard buffer solutions of pH 4, 7, and 10. The NPs were dispersed in ultrapure water to ensure sample homogeneity. The pH meter probe was immersed in the NP solution, and readings were taken after the pH value stabilized to minimize interference or fluctuations.
2.6.4. Scanning Electron Microscopy
The shape of the NPs obtained was analyzed using scanning electron microscopy (Zeiss EVO MA10 SEM, Oberkochen, Germany). The colloidal suspension (10 µL) was deposited in stubs, followed by 24 h air-drying. After total solvent evaporation, samples were coated with a 5 nm Au/Pd layer by sputtering. The microscopy conditions were set as follows: a working distance (WD) of 8.5 mm, magnification of 20,000×, an ion probe current (I Probe) of 20 pA, and examination at an accelerating voltage of 10 kV.
2.7. Antifungal Activity of Nanoparticles
2.7.1. NP Contact Method
The antifungal activity of the NPs was determined using the direct contact method on in vitro agar. This method was similar to that described in the “Agar Contact Method” Section 2.4.1, with the free compounds being replaced by the NP-EO and NP-NE.
2.7.2. Spore Germination Assay
To evaluate the effect of NPs on spore germination, the method described by Zhao et al. [34] with modifications was followed. Spores of B. cinerea or Rhizopus sp. were suspended in 1 mL of Tween 80 (0.05%) using a Drigalski loop and collected in a sterile microtube. The spore concentration was 1 × 105 CFU/mL, quantified using a Neubauer chamber. The spores were incubated in Potato-Dextrose broth containing NP-EO or NP-NE at concentrations that showed the best antifungal activity against the mycelium (10, 20, and 25 mg/mL) and incubated for 72 h at 25 °C. At least 100 spores were counted to calculate the germination rate.
2.8. Storage Stability Analysis
To evaluate the storage stability of NP-EO or NP-NE under temperature stress conditions, the NPs were prepared as described in the “NP Preparation” section. The prepared NPs were then stored at different temperatures (4 °C, 25 °C, and 37 °C) at pH 7.0. Samples were collected on days 0, 3, 5, 7, 10, 13, and 15 for size analysis using a Nanotrac NPA150 (Microtrac Inc., York, PA, USA) DLS-based device.
2.9. Hemolysis Assay
Hemolytic activity was evaluated essentially as described previously [33]. The NPs were mixed at a 1:1 (v/v) ratio in heparinized sheep blood (4%, v/v in PBS, pH 7.4). The aliquots were incubated (60 min at 37 °C) and then centrifuged at 3000× g for 10 min. One mL of the supernatant was taken for reading on a spectrophotometer at 540 nm. Triton X-100 (0.1%, v/v) and PBS were used as positive and negative controls, respectively [35]. Hemolytic activity was determined with Equation (3):
where AS: sample reading, AN: negative control, AP: positive control.
2.10. Statistical Analysis
Experimental data were expressed in the form of mean ± standard deviation. The analysis software used was GraphPad Prism (Version 6.0, GraphPad Software Inc., Boston, MA, USA). For comparisons between two groups, the t-test was applied. For comparisons between three or more groups, one-way analysis of variance (ANOVA) was performed, followed by Tukey’s multiple comparisons post hoc test to identify significant differences. In all analyses, p-values lower than 0.05 were considered statistically significant.
4. Discussion
The EO of B. dracunculifolia has nerolidol as its major component, which is consistent with previous reports [15,33]. The presence of β-caryophyllene and β-pinene is also notable, given their association with antifungal activities [36]. This suggests that the antifungal activity observed could be partially attributed to the synergistic action of these major components, especially NE, which has shown potent antifungal effects against filamentous fungi, such as various Aspergillus and Fusarium species [36,37].
The results of antifungal activity indicated significant inhibition of B. cinerea and Rhizopus sp. by both EO and NE. EO was particularly effective against B. cinerea, achieving over 50% inhibition at intermediate concentrations and over 95% at the highest concentration. NE also showed good activity, although less effective, particularly at higher concentrations. These results highlight the pronounced antifungal activity of EO compared to NE, similar to that described for other EOs [29,38,39]. The results of volatile exposure further supported the superior antifungal activity of EO over NE, showing a concentration-dependent response for both fungi. This may be due to the synergistic effects of the volatile components in EO, as terpenes and monoterpenes can work together to enhance fungal inhibition [40]. Previous studies have demonstrated the synergistic effects of compounds such as linalool and caryophyllene in EO from Michelia alba [41]. Thus, the combined effects of EO components are greater than the sum of individual effects, as seen with the compound NE. Additionally, volatiles showed significantly lower inhibitory activity than the direct contact method. This difference arises because free compounds in the solid medium ensure a more consistent concentration, while the varying volatilities of individual EO compounds cause them to diffuse at different rates in a non-saturated environment, resulting in higher emission rates that can reduce efficacy [42].
Both EO and NE were effectively incorporated in Pluronic NPs. The use of ultrasound was important for achieving nanometer-sized particles during NP preparation. The ultrasonic energy broke larger droplets, promoting the formation of more stable and smaller NPs [43]. Although the nature of the encapsulated compounds may influence particle size due to specific interactions with the polymer matrix, both EO and NE caused no significant effects on this parameter, as NP-EO and NP-NE exhibited similar particle diameters to that observed for the control (NP-B). Comparable particle sizes ranging from 100 to 200 nm have been reported for different NP formulations encapsulating EOs [25,33].
Furthermore, PDI values suggest that the NPs prepared in this study showed high uniformity. Polydispersity is an indicative of the homogeneity of particle size distribution, and PDI values around 0.2–0.3 have been associated with narrow size distribution [22].
The zeta potential results from the arrangement of the materials used in the formulation, and a relatively high value is important to maintain the physicochemical stability of the colloidal suspension since large repulsive forces tend to hinder aggregation of adjacent NPs [44]. The incorporation of EO or NE improved the zeta potential of the bare NPs, as NP-B had a zeta potential close to zero, indicating low colloidal stability. NP-NE exhibited a more negative zeta potential as compared to NP-EO, suggesting higher colloidal stability for NP-NE due to greater electrostatic repulsion. EO droplets present negative zeta potential values [45], and therefore a negative zeta potential is generally expected for NPs with an oily core. However, despite the higher negative charge of NP-NE, which could enhance penetration into fungal cells, NP-EO showed better antifungal activity. This discrepancy may be attributed to the specific chemical composition of the EO, which contains active compounds with strong antimicrobial properties, as reported in previous studies [40,41]. While zeta potential influences colloidal stability, other factors, such as the bioactive properties of the encapsulated compounds, play a crucial role in determining antifungal efficacy.
The EE varied considerably, with NP-EO achieving about 80%, significantly higher than NP-NE at 51%. The complex composition of EO, rich in monoterpenes, appears to facilitate better cohesion and packing of nonpolar chains within the NP vesicles, enhancing stability and retention. Monoterpenes located in the polar regions of membranes influence surface curvature and promote higher encapsulation efficiency [46]. NE, being a single compound, may have limited interactions with Pluronic® F-127, resulting in higher leakage during encapsulation and lower EE. The EE of NE is consistent with those observed for other individual EO components, such as timol and carvacrol, showing 46.3% and 50.9%, respectively [47].
The NPs (NP-EO and NP-NE) demonstrated improvements in antifungal efficacy as compared to non-encapsulated compounds, indicating potential for food preservation against fungal contaminants [48]. Encapsulation allows for controlled release of active compounds, resulting in prolonged effects, which is crucial for food storage and shelf life [22,49]. NP-EO, for instance, showed up to 87% inhibition of B. cinerea, comparable to or exceeding non-encapsulated EO. Moreover, NP-EO effectiveness at relatively low concentrations, such as 1 mg/mL, indicates its potential for the development of active packaging and preservatives [48]. This suggests that encapsulation not only preserved but may have enhanced the antifungal efficacy of EO, offering a robust solution for food protection against pathogenic fungi [22,49,50]. NP-NE also showed good efficacy, with up to 78% inhibition. This difference may be due to the higher volatility of NE, which is partially mitigated by encapsulation, allowing for sustained and protected release of the active compound [37,51].
There is limited literature on the inhibition of Rhizopus species by B. dracunculifolia EO and NE. However, the results of this study indicate that for Rhizopus sp., NPs were less effective as compared to B. cinerea. NP-EO achieved a maximum inhibition of 32%, while NP-NE reached 24%, suggesting that Rhizopus sp. is more resistant to treatments. This point is relevant for the preservation of foods where this fungus might be present, such as strawberries, peaches, tomatoes, and sweet potatoes [19,52,53]. In this case, further optimization of the NP formulation would be necessary to increase the efficiency against this fungus.
The effectiveness of NP-EO and NP-NE in inhibiting spore germination reflects their ability to halt fungal spread and infection, particularly in fresh foods such as tomatoes and strawberries [19,39]. The NPs showed pronounced inhibition of spore germination, especially at higher concentrations, suggesting their potential effectiveness in preventing fungal contamination from the early stages of sporulation. These findings reinforce the potential of NPs as active additives in food preservation, offering an innovative approach for extending shelf life and ensuring food quality [54,55].
Overall, the NPs demonstrated promising effectiveness, with the potential to enhance the delivery of bioactive compounds such as EO and NE. The ability of NPs to protect active compounds from degradation and volatilization may explain their increased or comparable efficacy to non-encapsulated compounds [48,50]. NP-EO, in particular, performed better than NP-NE, especially against B. cinerea, likely due to its chemical composition and higher EE, resulting in greater retention of bioactive compounds and enhanced antifungal efficacy in both contact and spore germination assays [43].
The stability analysis indicated that NP-EO and NP-NE formulations maintained good stability at 25 °C, with no significant changes in hydrodynamic sizes over 15 days. This demonstrates that at room temperature, the NPs can maintain their physicochemical characteristics, which is favorable for storage and transportation under normal conditions. This makes them suitable for application in active food packaging that does not require refrigeration [56,57]. However, storage at 4 °C led to the formation of a secondary population of particles, suggesting some instability at lower temperatures. This effect is potentially due to decreased hydrophobicity of the poly(oxpropylene) at lower temperatures, which might have caused the loss of oil emulsification and micelle disintegration [22]. At elevated temperatures (37 °C), NP-EO showed a significant decrease in size, possibly indicating degradation processes from prolonged exposure, facilitating drug release [58]. In contrast, NP-NE showed a constant size during incubation, suggesting greater resistance to thermal degradation [26]. These data are important for evaluating the stability of formulations under various conditions, which is crucial for ensuring their long-term efficacy.
The hemolytic activity results indicate the NPs have a safe potential for use in food applications and packaging. All NP formulations showed values below 3%, indicating minimal damage to erythrocytes. This safety profile is crucial, as it suggests that NPs containing EO and NE do not increase cytotoxicity, which is an important factor for their potential future use in food preservation, active packaging, and antimicrobial films where consumer safety is a priority [57,59]. However, it is essential to conduct further research on the potential side effects of nanoparticle migration from packaging to food, as well as to carry out preclinical and clinical studies to validate the safety of these nanoparticles.
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