1. Introduction
Biodeterioration is one of the main causes of natural building material degradation [
1]. Protecting these indoor and outdoor materials from biological contaminants is a challenging task. Under appropriate conditions that support microbial growth and proliferation such as high relative humidity and high temperature, fungi and bacteria are able to significantly contaminate natural materials. This leads to different types of damage on these materials, ranging from aesthetic to mechanical and chemical deterioration [
2,
3,
4,
5]. Even more serious is the impact of biological contamination on health. In fact, the microbial production of harmful substances such as allergens, mycotoxins, etc. is often associated with direct and indirect health effects [
6].
Stone and wood are among the most widespread natural building materials. For this reason, their protection against microbial damage is a topic of great interest.
The bacteria
Pseudomonas vancouverensis and
Flavobacterium sp. were isolated from a sandstone slab of the Maximilian’s Fountain in Bratislava (Slovakia). Members of these genera are associated with the biodeterioration of sandstone [
7].
The fungal strain
Purpureocillium lilacinum was also isolated from the same sample of the Maximilian’s Fountain. This fungus is frequently encountered in soils and is also used as a bio-pesticide [
8].
P. lilacinum is known to release nematocidal metabolites such as serine proteases, acetic acid, small-chain fatty acids, and leucinostatins, which plays a vital role as key toxicity factor in controlling nematode populations in soil [
9].
P. lilacinum cutaneous infection can be highly mortal, due to the fungus’s ability to sporulate in tissues [
10].
Fungi from the
Pleurotus genus are wood-decay fungi responsible for the decomposition of wood [
10].
P. eryngi demonstrated nematicidal activity against the larvae and adults of different nematodes species, in addition to proteases and chitinases production [
11].
Essential oils (EOs) are natural green candidates for the protection and disinfection of many materials, including stone and whitewood, without using aggressive and toxic substances [
12,
13]. EOs are composed of complex mixtures of substances with antimicrobial activity. Due to this multicomponent composition, they can be used as effective antibiofilm, antifungal, insecticidal, and antibacterial agents [
14,
15,
16].
Unfortunately, EOs are characterized by low water solubility, high volatility, and light, heat, and oxygen sensitivity. In fact, different chemical reactions, such as dehydrogenation or oxidation, triggered either enzymatically or chemically, can cause the degradation of EOs constituents [
17]. Nanoencapsulation is a well-established approach for preserving EOs [
18]. Several articles have been dedicated to the encapsulation of plant-derived metabolites such as EOs [
19,
20,
21,
22,
23]. Choi et al. [
24] found that the encapsulation of eugenol into poly(ε-caprolactone) nanocapsules could improve its stability against light degradation. Another study showed increased heat resistance of jasmine EOs after encapsulation in arabic gum and gelatin nanocapsules [
25]. Jummes et al. [
26] developed
Cymbopogon martini EOs-loaded poly(ε-caprolactone) nanocapsules with a small particle size (282 nm) and high encapsulation efficiency (99.54%). Similarly, rosemary EO was efficiently entrapped in poly(ε-caprolactone) nanoparticles with an average size of 220 nm, a zeta potential equal to −19.9 mV, and an encapsulation efficiency of 99% [
27]. To protect EOs from degradation and increase their physicochemical stability, a nanotechnology approach can be effective.
In previous works, we prepared nanocapsules based on poly(ε-caprolactone), a biodegradable and biocompatible polymer. NCs were loaded with thyme and oregano EOs, which contain a high amount of bioactive phenols such as thymol and carvacrol. We showed that the prepared nanostructured systems have a higher biological activity than EOs as they are [
2,
28].
In the current study, we evaluated the antibiofilm activity of Or-NC and Th-NC against two bacterial and two fungal strains which contaminate widely used building natural materials such as sandstone and whitewood. Moreover, under laboratory conditions, samples of sandstone and whitewood were used to prove the protective and disinfecting effect of Or-NC and Th-NC against the selected strains. A study of reflectivity was also performed to verify the color change of the tested sample materials.
This is an innovative approach for the application of nanoencapsulated EOs as protective and disinfecting agents for building natural materials, as there are very few publications dealing with similar research [
2].
3. Materials and Methods
3.1. Nanocapsules and Essential Oils
Poly(ε-caprolactone) (PCL) nanocapsules loaded with oregano and thyme EOs were prepared according to the procedure described by Granata et al. [
28] using the interfacial deposition of the preformed polymer method. Briefly, an acetone solution (80 mL) (Sigma-Aldrich/Merck, Darmstadt, Germany) containing EO (1.0 g), PCL (320 mg), and sorbitan monostearate (112 mg) (Sigma-Aldrich/Merck) was dropped under stirring at 25 °C into a polysorbate 80 aqueous solution (275 mL in 160 mL) (Sigma-Aldrich/Merck). The removal of the organic solvent provided an EO-NC suspension. The NCs without essential oils were prepared in the same conditions. The physicochemical parameters of the nanocapsules such as particle size, polydispersity, and zeta potential, were determined by light scattering measurements. The encapsulation efficiency and loading capacity values were obtained by UV–Vis spectroscopy. The exact composition of the commercial thyme and oregano EOs (provided by Flora s.r.l., Lorenzana, Italy, and Esperia S.p.A, Milan, Italy) were determined by GC-FID and GC-MS analysis. The values allowing the characterization of the nanocapsules and EOs were already reported [
10] and are briefly shown in
Table 5 and
Table 6.
3.2. Microbial Strains and Growth Conditions
The environmental bacterial strains Pseudomonas vancouverensis and Flavobacterium sp. and the fungal strain Purpureocillium lilacinum were isolated from a sandstone slab (Maximilian’s Fountain in Bratislava, Slovakia) and belong to the collection of the Institute of Molecular Biology Slovak Academy of Sciences (IMB SAS). Pleurotus eryngii CCBAS471, was obtained from the Culture collection of Basidiomycetes (CCBAS), Institute of Microbiology, Academy of Sciences of the Czech Republic. The bacterial strains were kept frozen in stock cultures at −80 °C, and the fungal cultures were stored at 4 °C and subcultured once a month. The bacteria were grown at 26 °C in nutrient agar (NBA) (HiMedia, Mumbai, India) for 12–18 h. The fungal strains were grown at 26 °C on Malt Extract Agar (MEA) (HiMedia) for 5 days.
3.3. Minimum Inhibitory Concentration (MIC)
Microtiter plate assays were performed according to Poaty et al. (2015) [
43] with a modification to determine the MIC (minimum inhibitory concentrations) of Th-NCs and Or-NCs, inhibiting bacterial and fungal growth [
10]. The MIC for the fungal strains was determined in our previous work [
32]. We based our testing of anti-biofilm activity on BIC concentrations determined by Kapustova et al. 2021 [
10].
3.4. Microtiter Biofilm Assay
Biofilm formation was processed as described by Harriot et al. [
44], with minor modifications. The bacterial cultures (
P. vancouverensis,
Flavobacterium sp.) were grown overnight in nutrient broth and then were washed twice in sterile phosphate-buffered saline (PBS) (Sigma-Aldrich/Merck) by centrifugation at 3000×
g for 5 min at room temperature. The obtained bacterial cell suspensions were diluted to a concentration of 2 × 10
5 CFU/mL. The isolates of fungal spores of
P. lilacinum and
P. eryngii were harvested from a 5-day-aged pure culture in MEA by adding 5 mL of saline solution (0.85%) to the plate. The spore suspensions were resuspended by vortexing before quantification. The final pellets of fungal cells were dissolved in approximately 20 mL of Malt Extract Broth (MEB; HiMedia) and counted under a microscope; the final density of the fungal cell suspensions was 2 × 10
5 spores/mL.
The microtiter biofilm assay took place in 96-well microtiter plates. The EOs-NCs testing proceeded as follows: 100 µL of bacterial or fungal cell suspensions, 90 µL of nutrient broth (NB; HiMedia) or MEB, and 10 µL of EOs-NCs in various concentrations were added to the plate wells. The original concentrations of EOs in NCs was 5.7 mg/mL for the thyme EO and 5.8 mg/mL for the oregano EO. These concentrations were adjusted with respect to the MIC and sub-MIC values for bacterial and fungal cell suspensions from 0.5 to 0.125 mg/mL and from 0.125 to 0.03 mg/mL, respectively. The final volume in each well was 200 µL.
Each bacteria plate was statically incubated for 24 h at 26 °C. After incubation, the supernatant was removed, and each well was rinsed 2 times with 90 µL of sterile PBS. Subsequently, 10 µL of 3-(4,5-dimethyl-thiazoyl)-2,5-diphenyltetrazolium bromide (MTT) (5 mg/mL) (Sigma-Aldrich/Merck) was added to the plate, which was incubated again for 2 h at 26 °C. After incubation, 100 µL of detergent (mixture of 95% isopropyl alcohol and 2M HCl) (CentralChem, Bratislava, Slovakia) was added and mixed, and the biofilm was characterized by optical density measurements. The measurements took place at 540 nm using an 800™ TS Absorbance Reader (BioTek, Winooski, VT, USA). With the measured absorbance values, we determined the MIC and sub-MIC.
Each fungal plate was incubated statically for 48 h at 26 °C. After biofilm formation, the MEB medium was carefully removed, and the fungal biofilm was quantified with 0.1% crystal violet solution (CV) as follows: 110 μL of 0.1% CV was pipetted into each well of dried biofilm and left for 5 min at room temperature. Then, the CV solution was withdrawn, and the wells washed twice with 100 μL of PBS and dried for 15 min at room temperature. Next, we added 200 μL of 95% ethanol into each well and left it for 15 min to release CV from the cells. After this time, 150 μL of inoculum was transferred to a clean 96-well microtiter plate, and we determined the MIC and sub-MIC values by optical density measurements at 570 nm by an 800™ TS Absorbance Reader (BioTek) [
45].
The control samples of biofilm formation from bacterial and fungal cell suspensions without EOs-NCs consisted of 100 µL of cells and 100 µL of medium per one well.
The data are presented as means of 3 experiments ± one standard deviation (SD). The differences between the groups were tested for statistical significance using the Student’s t-test (* p < 0.05; ** p < 0.01; *** p < 0.001). Because the antimicrobial activity datasets were normally distributed, the independent samples t-test was performed to test for significant differences between groups.
3.5. Antimicrobial Activity on the Sandstone and Whitewood Samples
Sandstone and whitewood were cut into rectangular units with a size of approximately 30 × 20 × 10 mm (L × W × D) for sandstone and of 40 × 40 × 10 mm (L × W × D) for whitewood. Stone and whitewood were sterilized by autoclave (121 °C for 20 min) and kept under a laminar flow for about 1 h. Two methods were tested and compared: disinfection (for disinfecting contaminated surfaces) and protection (for safeguarding surfaces against microbial colonization).
Disinfection procedure: 100 μL of bacterial/fungal suspension (1 × 106 CFU/mL) was spread on the surface of each sample. After absorption/evaporation of the bacterial/fungal suspension, 200 μL of Th-NCs and Or-NCs (with a concentration of EOs of 0.5 mg/mL for the bacterial surfaces and of 0.125 mg/mL for the fungal surfaces) was added to the sandstone and whitewood surfaces.
Protection procedure: 200 μL of Th-NCs and Or-NCs (with a concentration of EOs of 0.5 mg/mL for the bacterial surfaces and 0.125 mg/mL for the fungal surfaces) was added to the sandstone and whitewood surfaces. After drying the surfaces, 100 μL of bacterial/fungal suspension (1 × 106 CFU/mL) was added to the sandstone and whitewood surfaces.
The two bacterial strains (
P. vancouverensis and
Flavobacterium sp.) and the fungus
P. lilacinum were inoculated on the sandstone samples, while
P. eryngii was applied on the whitewood surface. After applying the microbial suspensions and EOs-NCs to the material surfaces, the tested materials were dried at room temperature. The treated surfaces were printed on MEA or NBA in order to transfer the microorganisms from sandstone and whitewood to agar medium via direct contact for 15 min. The conditions were standardized by applying 0.02 kg/cm
2 of constant pressure. This ensured the absence of air bubbles and perfect adhesion of the agar to the sandstone and whitewood surfaces [
2]. The agar plates were incubated at 26 °C for 48 h (bacteria) and at 26 °C for 96 h (fungi), and colony counts were carried out. The test was performed three times, and the average colony count of duplicate printed plates was used to calculate the CFU/mL.
3.6. Detection of Microbial Activity on the Substrates’ Surface
The printed biofilms on Petri dishes (mm2) were evaluated by using ImageJ program. A digital optical microscope from Keyence (Osaka, Japan) with a long working distance zoom objective was used to display the grown microbial culture on the wood and sandstone surfaces.
3.7. Measurement of Optical Properties
The reflectivity measurements were used to determine the optical properties of whitewood and sandstone with empty NCs and EOs-NCs before and after exposure to the microorganisms. The measurements were performed using a SolidSpec-3700 UV–VIS-NIR spectrophotometer from Shimadzu (Kyoto, Japan). The diffuse reflectivity measurements were recorded in the range from 200 to 900 nm using an integrating sphere equipped with three detectors: a photomultiplier, InGaAs, and PbS. In addition, the baseline correction prior to the reflectivity measurement was performed using a standard white plate composed of BaSO4.