2.4.1. Growth media and strain.

During this study, C. auris CBS10913T strain (Clade II) was used for the antifungal studies [16]. Prior to the experiment, the C. auris strain was resurrected in YPD broth that contained 1% (w/v) yeast extract, 2% (w/v) dextrose, and 2% (w/v) peptone, along with 2% (w/v) agar for the plates. Every strain of Candida was kept at −80 °C in 30% (v/v) glycerol stocks.

2.4.2. Broth dilution assay.

The Clinical and Laboratory Standards Institute’s (CLSI) method M27-A3 guidelines were adhered [17] for determination of MIC with slight modifications. On a 96 well plate, 100 ul of YPD media was added to each well which was followed by addition of Ag-MO and Ag-Zn-MO and then diluted serially. 100 ul of C. auris cell suspension was added to each well and then finally OD₆₀₀ was measured after 48 hrs of incubation at 30° C.

2.4.3. Spot assay and Static/Cidal assay.

Drop dilution assay was used for accessing spot assay [18]. Yeast extract peptone dextrose (YPD) agar plates containing nanocomposites Ag-MO and Ag-Zn-MO at their MIC₉₀ concentrations, i.e., 125 μg/mL and 250 μg/mL and control plates (without compounds) were prepared. In spot assay, 5 μl of Candida cell culture (0.1 OD₆₀₀) was applied to these plates in presence as well as absence of compounds, using fivefold serial dilutions. The plates were incubated at 30°C for 48 hours, and growth differences were recorded.

For static/cidal assay, C. auris ((0.1 OD₆₀₀) cells were initially grown overnight at 30°C in the presence of Ag-MO and Ag-Zn-MO, each at its minimum inhibitory concentration (MIC). The next day, 100 μl of these cultures were transferred to fresh YPD medium (lacking Ag-MO and Ag-Zn-MO) and incubated again overnight at 30°C. Finally, the following day, the growth of the C. auris cultures was assessed by measuring their optical density at 600 nm (OD₆₀₀) using a spectrophotometer [19].

2.4.4. Checkerboard titration method for drug synergism of Ag-MO and Ag-Zn-MO with known antifungal drugs (FLU, CAS, AMP B).

Checkerboard assays were conducted to evaluate the potential for synergistic, antagonistic, or indifferent interactions between both the nanocomposites and fluconazole (FLU), caspofungin (CAS) and amphotericin B (AMP B). Prior to testing, stocks and serial dilutions were made in accordance with the National Committee for Clinical Laboratory Standards (NCCLS) [17]. Except for the first and last two columns, 100 μL of YPD (Yeast Extract Peptone Dextrose) media was added to each well of a 96-well microdilution plate. Next, 200 μL of the compounds and medium were added to the first column. The final two columns were used as positive controls, which contained Candida cells, and negative controls, which contained only medium. While the second antifungal medication was serially diluted horizontally, the first ingredient in combination was diluted serially and vertically. 100 μL of Candida cells (0.1 OD₆₀₀) were added to all the wells, and the wells were then incubated for 48 hours at 30°C. Each combination of two medications was organized according to the checkerboard design, with the maximum concentrations of each drug in opposite corners. The lowest antibiotic concentration that, as seen with the naked eye, totally stopped the organism’s development [19].

2.4.5. Effect on Biofilm formation.

C. auris biofilms were grown in 96-well tissue culture plates. To start, 100 µl of a 1.0 x 10⁶ cells/ml C. auris suspension in RPMI 1640 medium was added to each well. The plates were then left undisturbed at 37°C. After 90 minutes, allowing cells to adhere, the liquid was removed and any unattached cells were washed away. Fresh medium was then added, and the plates were incubated at 37°C for an additional 24–48 hours to allow mature biofilms to form [18].

Crystal Violet staining of biofilm.

Briefly, C.auris cells were diluted with YPD media and then 200 µl cell suspensions were added to flat-bottomed 96-well microplates. Both the nanocomposites were simultaneously added to the wells of microplates and incubated fat 37^oC for 24 hrs. After incubation, the planktonic cells were removed from the wells, and the wells were washed with PBS. A 2.3% (w/v) crystal violet solution was then added and allowed to stain the biofilms for 5 minutes at room temperature. The stain was subsequently removed, and the wells were washed twice with PBS. Finally, the stained biofilms were observed under microscope [19].

2.4.6. Cell adherence assay.

Adherence assays were carried out in accordance with prior study [18]. After being grown on YPD media for the entire night at 37^o C, the yeast cells were resuspended in 2 milliliters of sterile PBS and centrifuged twice before being resuspended in spider media. Epithelial cells were obtained from the cheek mucous membrane of a co-author voluntarily by gently scraping sterile cotton swabs which were then gently agitated and centrifuged at 3000 rpm for three minutes each to wash with PBS. In order to conduct adherence tests, 1 milliliter of each suspension was combined in a test tube. This was followed by two hours of gentle stirring at 37^oC in the presence of Ag-MO and Ag-Zn-MO separately. Following incubation, each test tube received two drops of 0.4% trypan blue solution, and the mixture was gently mixed. Finally, 10 ul stained suspension was examined under a microscope [19].

2.4.7. Cell wall staining and analysis.

Exponential-phase C. auris cells (2.5 × 10⁶ cells) were obtained from overnight cultures of control, Ag-MO, and Ag-Zn-MO- treated samples. The cells were then fixed for 30 minutes using 4% paraformaldehyde in phosphate-buffered saline. Fixed cells were washed and stained with ConA-fluorescein conjugate (50 μg/mL) for mannan, AB (100 μg/mL) for β-1, 3-glucan, and CFW (5 μg/mL) for chitin. For half an hour, the staining was done at room temperature. Finally, the fluorescence-stained cells were seen at 100 X magnification using a Nikon fluorescent microscope. NIS Elements program was used to take pictures [18].

2.4.8. Propidium Iodide Uptake.

Propidium Iodide (PI) is a fluorescent dye that glows when it binds to nucleic acids. It cannot easily pass through the membranes of living cells. PI is used to tell the difference between both healthy and injured Candida cells. Candida cells (0.1 OD₆₀₀) were inoculated with Ag-MO and Ag-Zn-MO and were allowed to grow by gentle shaking for 3 hours. After that, the cells were cleaned, centrifuged, and stained for 15 minutes with PI (1 μg/mL). The cells were then examined using a fluorescent microscope [18].

2.4.9. Estimation of Ergosterol.

0.1 OD₆₀₀ C. auris cells were inoculated in 50 ml of YPD in the presence of Ag-MO and AgZn-MO. The cells were treated with a solution of potassium hydroxide (KOH) dissolved in alcohol. The strong alkaline conditions of KOH help to disrupt cell membranes and release the sterols and %age of the wet weight of the cells was calculated to estimate ergosterol content by following equation:

% Ergosterol + % 24(28)-DHE= [A281.5/290). F]/ pellet weight; 24(28)-DHE= [A230/518). F]/ pellet weight and % Ergosterol = [% ergosterol + % 24(28) DHE]—% 24(28) DHE, where F is the factor for dilution in petroleum ether and 290 and 518 are the E values (in percent per centimeter) determined for crystalline ergosterol and 24(28)-DHE, respectively [18,19].

2.4.10. Vacuole acidification & morphology study.

Untreated cells (used as control); Ag-MO & Ag-Zn-MO were overnight cultured in YPD media. Following a 4-hour suspension in fresh YPD, cells were washed in 1x PBS and FM4–64 (a fluorescent lipophilic, styryl dye) at 40μM was introduced, and further incubation was done at 30°C for 15 minutes. Washing and resuspension in newly prepared YPD was repeated after incubating at 30°C for 60 minutes. Lastly, pictures were taken at a magnification of 100x using a fluorescent microscope [18].

2.4.11. R6G extracellular efflux assay.

A previously established technique was used to determine the efflux of R6G [18]. In short, a culture of yeast that had grown overnight was pelleted, cleaned, and resuspended in PBS without glucose, resulting in about 1 x 10⁶ cells. After being de-energized in 2-DOG and 2, 4 DNP for 45 minutes, the cells were cleaned and reconstituted in PBS devoid of glucose. After adding R6G to a final concentration of 10 μM, it was incubated at 30°C for 40 minutes. To evaluate efflux, the equilibrated cells were cleaned and resuspended in PBS devoid of glucose. One milliliter of samples was taken out at the designated intervals and centrifuged for 2 minutes at 9000 x g. After collecting supernatant, the optical density (OD) at 527 nm was determined. The cells reconstituted in PBS devoid of glucose were supplemented with 2% glucose to determine energy-dependent efflux. Every experiment had glucose-free negative controls and positive control had only untreated Candida cells with glucose and R6G [19,20]. The time intervals when readings were taken were 10 minutes apart. For competition assays, Ag-MO (100 μg/mL) and Ag-Zn-MO (200 μg/mL) were added respectively to the de energized cells 30 min before the addition of R6G at various concentrations (10 μM, 20 μM, 30 μM, 40 μM) and allowed to equilibrate to follow glucose for energy-dependent efflux as above. Various concentrations of R6G were added in order to determine the rate of the reactions to get the slope for competitive or non-competitive plots.

2.4.13. Caenorhabditis elegans studies.

Caenorhabditis elegans (C. elegans) were infected with C. auris and monitored for survival. To evaluate the antimycotic effects of both the nanocomposites, C. elegans were co-cultured in presence or absence of Ag-MO and Ag-Zn-MO with C. auris at their sub-MIC concentrations which are 100 μg/mL and 200 μg/mL respectively. Intestinal persistence was visualized using fluorescence microscopy. Survival studies of C. elegans infected with C. auris were conducted using the Kaplan-Meier method. Approximately 50 young adult female C. elegans worms in the L4 developmental stage were transferred from a culture of E. coli OP50 bacteria to BHI growth medium supplemented with Ag-MO (100 μg/mL) and Ag-Zn-MO (200 μg/mL). C. auris was introduced to worms and their survival was tracked daily for 7 days. Images were obtained on the seventh day of infection, and the infected C. elegans were cultured at 25 °C for seven days, scoring as live or dead every day. To visualize C. auris in the worm intestine, fluorescence microscopy was used. Images were captured using a stereo microscope and a fluorescence microscope [18].

2.4.14. Macrophage killing & infection.

THP-1 cells, a type of human white blood cell, were grown in a nutrient-rich medium. The cells were treated with 15 nM Phorbol 12-myristate 13-acetate (PMA), a pharmacological stimulant, for 48 hours before being rinsed thrice. Afterward, these macrophage-like cells were infected with C. auris. This fungus was prepared by centrifuging a culture, resuspending it in the same growth medium, and breaking up any clumps with a syringe. The infection was carried out at a high fungal-to-cell ratio (5:1) (pathogen: host) for 4 hours. Following the infection, the cells were treated with an antibiotic, amikacin (200 µg/ml). Following a PBS wash, the cells were resuspended in RPMI with 10% FBS added, and the remaining intracellular fungi were allowed to grow for 24 hours. After lysing the cells with 0.5% Triton-X, the cell lysate was serially diluted in triplicate and plated on agar plates to count the number of surviving fungi. The number of fungal colonies that grew on the plates was counted to assess the effectiveness of the treatment [21].

2.4.15. Hemolytic activity assay.

A modified version of a previously established approach was used to determine the hemolytic activity of Ag-MO and Ag-Zn-MO [18]. The absorbance of hemoglobin at 540 nm was used to quantify its discharge into the supernatant. PBS was used as a negative control, while 1% Triton X-100 was utilized as a positive control. The percentage of hemolysis was calculated based on the absorbance of the sample, the blank, and the Triton X-100-treated sample.

2.4.16. Statistical analysis.

All experiments were conducted in triplicate (n = 3). Student’s t-test was used to examine the data, and P < 0.05 was deemed statistically significant. The mean ± standard deviation is used to express the data.

3.1.1. X-ray diffraction.

X-ray Diffraction (XRD) is very useful in determining the crystal structure, atomic arrangement and crystal size. Fig 1a shows the X-ray diffraction pattern of AgNPs derived from MO leaves. The nanoparticle showed a sharp high intensity peak indexed to (111), (200), (220) and (311) planes corresponding to face centered cubic lattice of AgNPs. The absence of broad peaks shows the existence of pure form of AgNPs without Moringa plant extract residues. Fig 1 b shows the X-ray diffraction pattern of ZnO NPs derived from MO leaves. The formation of hexagonal wurtzite structure in pure phase has been confirmed with the distinct planes corresponding to (100), (002), (101), (102), (110), (103), (200), (112) and (201), respectively [22]. The presence of two additional amorphous peaks at lower 2 theta range below 30° could be attributed to the residues of MO. The peaks confirm the deposition of plant extracts on the surface of nanoparticles and such composites clearly show the role of antioxidants, flavonoids and associated organic contents in the formation of ZnO nanoparticles [22]. Fig 1 c shows the X-ray diffraction pattern of Ag-ZnO NPs derived from MO leaves. The XRD pattern exhibits peaks corresponding to both face-centered cubic structure of AgNPs and hexagonal shaped wurzite ZnO. AgNPs peaks corresponding to (111), (200), (220) and (311) planes were dominant, while less intense peaks of wurtzite structure of ZnO were visible at lower 2 theta values. Such large crystallinity of AgNPs indicates the deposition of metallic phase of Ag on the surface of ZnO nanoparticles.

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Fig 1. XRD pattern of Ag-MO and Ag-Zn-MO.

a) Peaks (111), (200), (220) and (311) planes correspond to face centered cubic lattice of AgNPs. b) XRD pattern of ZnO NPs. (100), (002), (101), (102), (110), (103), (200), (112) and (201) planes show hexagonal wurtzite structure in pure phase. c) XRD pattern of Ag-Zn-MO. Pattern exhibits peaks corresponding to both face-centered cubic structure of AgNPs and hexagonal shaped wurzite ZnONPs. Such crystallinity of AgNPs indicates the deposition of metallic phase of Ag on the surface of ZnO nanoparticles.

https://doi.org/10.1371/journal.pone.0336309.g001

3.1.2. Zeta potential measurement.

Zeta potential measurement shows the surface charge measurements. Fig 2 shows the zetapotential of Ag, ZnO and Ag-ZnO NPs derived from Moringa Oleifera leaves. The technique is useful to study the zeta size and colloidal state stability of synthesized Ag, ZnO and Ag-ZnO NPs. AgNPs showed the highest negative zeta potential of −14.6 mV, while ZnO nanoparticles showed the lower negative zeta potential of −3.0 mV. The composite of Ag-ZnO showed an intermediate negative zeta potential of −10.5 mV. The present study shows that AgNPs and Ag-ZnO NPs possess dispersion stability than ZnO nanoparticles. This is in accordance with a prior study where the presence of capping agent on NPs surface was responsible for repulsive force between NPs that hindered their aggregation [23,24].

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Fig 2. Zetapotential of Ag-MO and Ag-Zn-MO.

a) Ag-MO showed the highest negative zeta potential of −14.6 mV, while ZnO nanoparticles showed the lower negative zeta potential of −3.0 mV while Ag-Zn-MO showed the intermediate zeta potential of −10.5 mV. b) thermogravimetric analysis of Ag, ZnO and Ag-ZnO NPs derived from Moringa oleifera leaves. Ag-MO showed three distinct weight losses between 100−280 °C, 300−500 °C and 500−900 °C with total weight loss of about 50%. In Ag-Zn-MO, the weight loss was reduced to 30% with a distinct decomposition trend. ZnO alone showed a broad decomposition weight loss starting from 100−440 °C and second weight loss from 450−900 °C with total weight loss of about 53%. c) FTIR spectra of Ag-MO, ZnO and Ag-Zn-MO. The spectra of Ag-MO and Ag-Zn-MO showed a distinct broad peak at about 3245 cm^-1 corresponding to the hydroxyl groups polyphenols and amino compounds. The peak at 1606 cm-1 is ascribed due to the carbonyl stretching functional group of amides. d) DLS data and size distribution graph of AgNPs derived from Moringa oleifera leaf extract.

https://doi.org/10.1371/journal.pone.0336309.g002

3.1.3. Thermogravimetric analysis.

Thermogravimetric Analysis (TGA) is used in determining physical phenomena like thermal decomposition and phase transition based on measuring the mass of substance after heat treatment on elevated temperatures. Fig. 2 b shows the thermogravimetric analysis of Ag, ZnO and Ag-ZnO NPs derived from MO leaves. AgNPs showed three distinct weight loss between 100–280 °C, 300–500 °C and 500–900 °C with total weight loss of about 50%. ZnO NPs showed a broad decomposition weight loss starting from 100–440 °C and second weight loss from 450–900 °C with total weight loss of about 53%. In the case of Ag-ZnO NPs, the weight loss was reduced to 30% with a distinct decomposition trend observed. For instance, a loss of less weight was observed between 100–280 °C followed by a gradual weight loss up to 900 °C. For the three samples, the observed weight at about 100 °C was ascribed due to moisture adsorbed on the surface of Ag, ZnO and Ag-ZnO nanoparticles. The significant weight loss observed up to 700 °C is ascribed due to the disintegration of phytocomponents of plant extracts [25], in our case it is related to the MO.

3.1.4. FTIR analysis.

FTIR is used to determine the surface chemistry of synthesized nanocomposite. The functional groups of biosynthesized Ag-MO and Ag-Zn-MO nanocomposites were determined by FTIR analysis. Fig 2 c shows the FTIR spectra of Ag, ZnO and Ag-ZnO NPs derived from MO leaves. The spectra of Ag and Ag- ZnO showed a distinct broad peak at about 3245 cm^-1 corresponding to the hydroxyl groups polyphenols and amino compounds [25]. The peak at 1606 cm-1 is ascribed due to the carbonyl stretching functional group of amides. In the case of ZnO, a characteristic stretching peak of Zn-O was observed at about 670 cm^-1 [26]. The presence of such band at about 665 cm^-1 in Ag-ZnO shows the composite formation between Ag and ZnO NPs. [27,25] (Fig 2 c).

Dynamic light scattering analysis (DLS) was also done to check the size of the nanoparticles. The results indicate that the average hydrodynamic particle size is approximately 769.5 nm, suggesting possible nanoparticle aggregation in the aqueous medium (Fig 2d). This observation is consistent with the biological synthesis route, which often leads to surface-bound phytochemicals contributing to increased particle size and agglomeration.

3.1.5. SEM analysis.

SEM analysis was also performed to determine the morphology of Ag-MO and Ag-Zn-MO nanocomposite. Fig 3 (a-c) shows the SEM image of Ag, ZnO and Ag-ZnO NPs using MO leaf extract. In our study, the chemical precursor was found to play a key role in the formation of different shaped Ag, ZnO and Ag-ZnO NPs. For instance, metal precursor sources such as silver nitrate, zinc acetate and mixed metal precursors tend to form variable morphological features of nanoparticles (Fig 3 (a-c) Ag NPs using silver nitrate precursor indicating formation of mixed large, aggregated nanoparticles covered with small, layered particles (Fig 3a). In the case of green ZnO NPs, the formation of chucks occurs with variable sizes (Fig 3b). Bimetallic Ag and low density ZnO nanocomposite formation tends to form aggregation of small size Ag nanoparticles on microforest flower ZnO structure [27]. In this case, the mixed precursor showed better formation of large morphology of Ag-ZnO indicating an effective bioreduction by MO (Fig 3 c). Fig 3(d-f) shows the EDX spectrum of Ag, Zn and Ag-Zn NPs. The formation of distinct Ag and Zn peaks indicates the successful reduction of AgNO₃ to Ag NPs and Zn acetate to Zn NPs, respectively (Fig 3d and e). Fig 3f shows the synergistic presence of both Ag and Zn NPs indicating cohabitation of both types of nanoparticles formation in a single pot facilitated by MO leaf plant reduction [27,28].

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Fig 3. SEM images and EDX spectra.

a) Mixed large, aggregated nanoparticles covered with small, layered particles of AgNO3-MO. b) Formation of variable chucks in ZnO-MO nanocomposite. c) Aggregation of small size Ag nanoparticles on microforest flower ZnO structure. d) Distinct peak shows AgNO3 reduced to Ag NPs. e) Peak shows reduction of ZnO to Zn NPs. f) Cohabitation of Ag and Zn after reduction.

https://doi.org/10.1371/journal.pone.0336309.g003

Additionally, XPS was also carried out to observe the presence of different metallic peaks that has been presented as a supplementary file. The study of elemental composition and oxidation states of Ag-MO and Ag-Zn-MO were analyzed which revealed the presence of carbon and oxygen that might have come from residual of Moringa extract used in the preparation of nanocomposites. These results confirm the formation of metallic AgNO₃ without any presence of AgO in Ag-MO and the formation of ZnOH and other zinc oxide derivates in Ag-Zn-MO [29–32] (Supplementary file, S1 and S2 File).