Fresh flowers of Hibiscus rosa-sinensis were collected from the Horticulture Nursery, AMU, Aligarh, India, and washed several times with deionized water. A known volume of distilled water (100 ml) was mixed with 10 g (wet weight) of thoroughly washed HRS flowers. The mixture was boiled till the original volume of aqueous suspension was reduced to 50 ml and filtered using Whatman No. 1 filter paper. The resulting HRS flower extract was used in the synthesis of as-fabricated AgNPs. Similarly, HRS buds and leaf extracts were prepared and employed in the fabrication of AgNPs.

Increasing volumes (3–5 ml) of HRSF extract were added to 5 ml aqueous solution [10^–3 M] of silver nitrate (AgNO₃). The final volume of each reaction mixture was made up to 10 ml by adding a sufficient amount of distilled water and was kept on a stirrer at 100 rpm overnight at room temperature (Figure 1). Synthesis of AgNPs was also done by employing buds and leaf extracts of the HRSP. The as-synthesized AgNPs were characterized by various spectrophotometry-, transmission electron microscopy (TEM)-, and dynamic light scattering (DLS)-based analysis. The characterization and antibacterial activity data of buds and leaf extracts based on synthesized AgNPs are shown in the Supplementary Table S1.

We used the following bacterial strains in the study:

A UV-Vis spectroscopic analysis of the as-synthesized AgNPs was performed to ascertain the biogenic reduction of precursor AgNO₃. The double beam spectrophotometer (Jasco model V-750), operated at a resolution of 1 nm, was used to execute the analysis (Bhosale et al., 2014). The UV-Vis spectrum was recorded over the 200–800 nm range.

Dynamic light scattering (DLS) measurement was employed to determine the hydrodynamic size distribution of the particles. DLS measurements were analyzed by using Malvern Zetasizer Nano-ZS90 (ZEN3590, UK) to determine the average size and size dispersal of the as-synthesized AgNPs. The lyophilized powder was resuspended in PBS, and the solution obtained after passing through a 0.22-μm filter (Millipore) was subjected to various size determining measurements. The scattered light intensity was detected at 90° of the incident beam, and data analysis was performed in the default mode. The measured size was presented as the average value of 20 runs, with triplicate measurements within each run.

The as-synthesized AgNPs were characterized by TEM. We also studied the effect of increasing volumes of the HRSF extract on the shape and size of the as-formed AgNPs with the help of the JEOL Transmission Electron Microscope at an accelerating voltage of 200 kV. A copper grid mesh, covered with the carbon-stabilized film, was employed to analyze the as-synthesized AgNPs. The uranyl acetate (2% w/v) solution was used for negative staining of the sample.

For the determination of zeta potential of as-synthesized AgNPs, Malvern Zetasizer Nano-ZS90 (ZEN3590, UK) was used. Reconstitution of the samples was carried out in 5 mM phosphate buffer, pH 7.4, and the electrophoretic mobility was measured in the electrophoresis cell. The samples were run in triplicate to determine the zeta potential.

FTIR spectra of as-synthesized AgNPs was analyzed on an FTIR spectrophotometer (PerkinElmer, UK) in the diffuse reflectance mode operating at a resolution of 4 cm⁻¹ (Iravani et al., 2014). The spectra of AgNPs and HRSF extract were taken between 4,000 cm⁻¹ and 400 cm⁻¹. As-synthesized AgNPs were co-prepared again along with KBr crystals as a beam splitter.

The bactericidal activity of the as-synthesized AgNPs was assessed by employing the agar well diffusion method. Various bacterial strains such as Listeria monocytogenes, Streptococcus mutans, and Escherichia coli were used in the study. The bacterial strain S. mutans MTCC-497 (Institute of Microbial Technology, Chandigarh, India) was cultured in the brain heart infusion (BHI) broth (HiMedia Labs, Mumbai, India) at 37°C. We also used S. mutans serotype c strain (ATCC 700610/UA159) to assess the antibacterial potential of as-synthesized AgNPs. Next, we determined the antibacterial potential of as-synthesized AgNPs against standard E. coli (ATCC-25922). The antibacterial potential of as-synthesized AgNPs was also established against L. monocytogenes ATCC 19115. The clinical isolate of S. mutans, used in the study, was a kind gift from Prof. Shahid of the Department of Microbiology, JNMC, AMU, Aligarh.

The nutrient broth was used for subculturing of the pure culture of S. mutans, E. coli, and L. monocytogenes. They were grown overnight to attain the colony-forming unit (CFU) count of ∼10⁶ units per ml. An aliquot of 100 μl, from overnight grown culture, was spread uniformly on nutrient agar plates by using a sterile plastic spreader.

The agar plates, inoculated with S. mutans, were cocultured with the AgNP-Ni-Ti orthodontic wire. The uncoated and plain AgNO₃-coated Ni-Ti orthodontic wires were used as the control. The plates were incubated at 37°C, and the zone of inhibition was observed after the stipulated time period of 24 h.

In the next set of experiments, the agar plates were first exposed with E. coli and L. monocytogenes followed by punching of the wells using a punching machine. Subsequently, an increasing amount of as-synthesized AgNPs (1 mg/ml stock solution) or standard antibiotic, vancomycin (control), was dispensed into various wells in a given agar plate. The plates were incubated at 37°C, and the zone of inhibition was determined by measuring the clear region around each well after the stipulated time period of 24 h.

The anti-biofilm potential of as-synthesized AgNPs was established by employing the XTT assay, which is performed to analyze the presence of viable cells. The principle of the assay involves the cleavage of the tetrazolium salt XTT (2,3-bis(2-methoxy-4-nitro-5-sulfophenyl)-5-{phenylamino}carbonyl-2H-tetrazolium hydroxide) in the presence of an electron-coupling reagent, which ensues in the production of a soluble formazan salt. The XTT assay was performed following published protocols with minor changes as standardized in our lab. In brief, 100 µl of bacterial cell culture (1 × 10⁸ cells/ml) was dispensed in each well of 96-well culture plates. The bacteria were allowed to form adherent mature biofilm in the presence of an increasing concentration of the as-synthesized AgNPs for 48 h. Just before the commencement of the assay, a solution of menadione (0.4 mM), which acts as an artificial electron transporter to reduce tetrazolium salt to formazan, was prepared and filtered.

The plate was incubated for the stipulated time period, followed by washing with PBS (200 µl). Next, the XTT reagent [2,3-bis(2-methoxy-4-nitro-5-sulfophenyl)-5-{phenylamino}carbonyl-2H-tetrazolium hydroxide (XTT) at a final concentration of 250 μg/ml] was dispensed to each test well of the plate followed by the addition of menadione solution (2 μl/well). The plate was further incubated for the next 4 h at 37°C in the dark. Next, the solution was transferred into a fresh plate to assess the colorimetric change, and the absorbance was taken at 490 nm using an iMark microplate absorbance reader (BIORAD, United States). The XTT experiments were performed in triplicate to validate the data.

We studied the effect of as-synthesized AgNPs on the adherence of S. mutans on the smooth glass surface. We also assessed the effect of the presence of sucrose in the surrounding medium during AgNP-mediated inhibition of S. mutans adherence. The bacteria were cultured at 37°C for 24 h at an angle of 30° in a glass tube containing 10 ml of BHI with or without 5% (w/v) sucrose. The cultured tubes were also exposed to the sub-MIC concentrations of the AgNPs. The culture medium consisted of BHI (sucrose dependent) and without sucrose (sucrose independent). After stipulated incubation, the glass tubes were slightly rotated to decant the planktonic cells. Subsequently, the adhered cells were removed by adding 0.5 M of sodium hydroxide with stirring. The cells were thoroughly washed and suspended in saline. The adherence was determined spectrophotometrically at 600 nm. The percentage adherence was determined by using the following formula: p e r c e n t a g e   a d h e r e n c e   = ( O .D .   o f   a d h e r e d   c e l l s / O .D .   o f   t o t a l   c e l l s ) × 100.

The effect of AgNPs on the cell surface hydrophobicity of S. mutans was determined by measuring its adhesion with hydrocarbon. The cells were grown in the BHI medium supplemented with 5% sucrose along with the increasing concentration of AgNPs. The cells were washed twice and suspended in sterile saline (0.85%) to adjust optical density (O.D.) at around 0.3 at 600 nm. A given volume of the cell suspension (3 ml) was mixed with 0.25 ml toluene. The tubes were stirred for 2 min and allowed to equilibrate at room temperature for 10 min. The cell suspension was kept aside to allow phase separation. We next determined the O.D. of the aqueous phase spectrophotometrically at 600 nm.

Overnight cultured bacterial cells (treated and control) were suspended in 1 ml LB medium to obtain optical density (at a wavelength of 600 nm) at around 1.0 ± 0.01. Furthermore, toluene (1 ml) was added to the suspension (both treated and control) and vortexed. The test tubes were incubated for 30 min to let the biphasic solution settle. Next, the optical density of the aqueous phase was measured, and the hydrophobicity index was calculated by using the following equation: H I % = [ ( A i − A f ) / A i ] × 100 ,

where A_i and A_f denote the initial and final optical densities of the aqueous phase. The adherence of the bacterial cells to the organic solvent was evaluated to ascertain the hydrophobicity (Farkas et al., 2015).

Sterilized Ni-Ti orthodontic wires were placed into three separate Eppendorf tubes containing fresh sterilized nutrient broth. Next, one of the two sets of orthodontic wires was co-incubated with as-synthesized AgNPs (50 μg/ml) and the other with aqueous AgNO₃ solution (50 μg/ml). The wire, incubated in media only, served as the control. The standardized suspension (1 × 10⁸ cells/ml) of S. mutans was added to each Eppendorf tube (harboring orthodontic wire) and was incubated for 24 h at 37°C. Once the incubation period was over, wires were taken out and placed into fresh Eppendorf tubes containing PBS and subjected to sonication to remove adhered bacteria from wires. A known volume (100 µl) of the resultant bacterial suspension (diluted 100 times), collected post sonication, was spread on NB agar plates and incubated for 24 h at 37°C, and CFU/ml was calculated for each group.

S. mutans was cultured in the NB medium at 37°C overnight. An aliquot of log-phase bacterial culture (having 10⁶ CFU/ml) was suspended in broth (500 µl) and dispensed on the sterile coverslip placed in wells of a multi-well polystyrene plate. The setup was left for 24 h at 37°C for the growth of biofilm. Mature biofilm was washed and incubated with AgNPs (50 μg/ml) suspension and AgNO₃ solution (50 μg/ml) for the duration of 3 h. An untreated mature biofilm was considered as a control. Coverslips were then fixed in 4% paraformaldehyde (PFA) followed by washing, and cells were then stained with the fluorescein isothiocyanate (FITC) dye. Stained coverslips were then washed with sterile PBS, and the anti-biofilm activity was observed under the Zeiss Axiocam Imager MRM M2 fluorescence microscope. Both the treated and control samples were observed under a fluorescence microscope (×40 magnification).

The phenol/H₂SO₄ method (Decker et al., 2011) was used to quantify the glucan in the cultured sample. S. mutans was incubated in media containing 1% sucrose both in the absence and presence of as-synthesized AgNPs for 20 h, after which the bacteria were recovered. Extracellular water-soluble glucan was isolated by employing ethanol precipitation of the supernatants from the recovered bacterial cultures. The remaining cell pellet was resuspended in 1 M NaOH and centrifuged to remove bacterial cells. Water-insoluble glucan was recovered by ethanol precipitation of the supernatant.

Ni-Ti orthodontic wires were incubated with as-synthesized AgNPs (50 μg/ml) and aqueous AgNO₃ (50 μg/ml) for 24 h with mild and continuous shaking for homogenous adherence. Topographical changes on coated Ni-Ti orthodontic wires as compared to uncoated one (control) were assessed by scanning electron microscopy (JSM67500F, a JEOL model, Japan).

To facilitate the future plausible biomedical application, we assessed the cytotoxicity of the as-synthesized AgNPs against erythrocytes (RBCs) and PBMCs. It was done by performing an in vitro hemolysis assay of RBCs and killing of PBMCs employing the MTT assay as described later.

Data were analyzed by one-way and two-way ANOVA to assess the differences among groups. Statistical calculations were performed with the help of GraphPad Prism version 8.0. Significance was indicated as *** for p < 0.001; ** for p < 0.01, and * for p < 0.05.

The potential of HRSF extract to synthesize nanoparticles was established by evaluating its ability to convert silver salt to as-synthesized silver nanoparticles. The aqueous silver ions were exposed to the HRSF extract. The color of the reaction mixture changed from light pinkish-red to dark brown (Supplementary Figure S1). The color transformation corresponds to the characteristic surface plasmon resonance (SPR) effect of different size silver nanoparticles. The SPR analysis establishes HRSF-mediated synthesis of AgNPs.

The optical properties of the as-synthesized AgNPs were determined by employing surface plasmon absorption spectroscopy. The SPR properties usually depend on the shape of the NPs population. At the outset, a UV-Vis absorption spectroscopic technique was used to analyze the synthesis of the nanoparticles. Figure 2A shows the UV-Vis absorption spectra recorded for as-synthesized silver nanoparticles harvested after 24 h of the reaction (upon treatment with 3 ml of HRSF extract). The results indicate that the reaction solution displayed an absorption maximum at about 460 nm that can be attributed to the SPR of the as-synthesized silver nanoparticles (Lim, 2014). It was observed that with the increase in time, the silver nanoparticle synthesis rate also increased (Supplementary Figure S2). Although the wavelength maximum was stably positioned at 460 nm, the intensity steadily increased with time and eventually got saturated after 24 h.

We also studied the kinetics of silver nanoparticle synthesis upon incubation of AgNO₃ with increasing content of HRSF extract. The synthesis of AgNPs augmented upon increasing the concentration of plant extract in the reaction medium. The increase in HRSF content (3 vs. 5 ml HRSF extract) resulted in an upsurge in absorbance at 460 nm (Figure 2A).

The green synthesis of silver nanoparticles was further confirmed by representative TEM images of silver nanoparticles generated employing increasing amounts of flower extract for their synthesis. The shape of the particles was found to be dependent on the concentration of flower extract. Higher abundance of HRSF extract favored the occurrence of a largely homogenous population of small-sized nanoparticles. At lower concentrations of flower extract various triangular, hexagonal, and ovoid shaped particles in the size range 10–40 nm (Figure 2B) were seen coexisting. However, as the concentration of HRSF extract was increased, the average diameter of the silver nanoparticles was found to decrease as evidenced by the occurrence of spherical particles (15–25 nm) at higher concentrations of HRSF extract (Figure 2C). Also, as the flower extract concentration increased, the total number of particles in a given volume increased simultaneously as seen in the TEM micrograph (Figure 2C). Initially, the increase in the HRSF extract causes the nanoparticles to be formed in the range of 30–50 nm. This is more evident from the fact that a higher concentration of flower extract led to the synthesis of the large number of isomorphic spherical nanoparticles of small dimensions (20–30 nm), which is accompanied by a decrease in the number of large-sized compounds with the size range of 30–50 nm.

For the determination of the particle size of as-synthesized AgNPs in an aqueous buffer, the dynamic light scattering (DLS) analysis was carried out. To have better information pertaining to the size distribution of as-synthesized AgNPs, DLS is performed in conjunction with the TEM analysis. Analysis of the HRSF extract-mediated synthesized AgNPs’ dynamic light scattering (DLS) pattern showed the average size of as-synthesized AgNPs to be around 30–80 nm with a polydispersity index (PDI) of 0.28 (Figure 2D). The TEM analysis revealed the morphology of the as-synthesized AgNPs to be spherical/ovoid and having a size range of around 30–50 nm. The discrepancy in the size as determined by DLS versus TEM analyses is usually explained on the basis of the hydrodynamic diameter of AgNPs, which correspond to the size of the particles suspended in the liquid medium (Figure 2 B&C).

Zeta potential is a physical property which relates to the net surface charge of the nanoparticles. Depending upon the magnitude and nature of the charge, the particles may repel each other (because of the Coulomb explosion force) eventually preventing agglomeration of the as-synthesized nanoparticles. The stability of NPs has been correlated well with the zeta potential ranging from +30 mV to −30 mV. The as-synthesized AgNPs were found to have zeta potential of around −25 mV [Figure 2E].

FTIR spectroscopy was performed to characterize the putative biomolecules (organic and inorganic functional groups) responsible for the reduction of Ag⁺ ions and the stabilization of reduced silver nanoparticles to prevent agglomeration and their capping in the aqueous medium. The FTIR spectra of HRSF extract and the as-synthesized AgNPs are shown in Figure 3. Both the spectra showed bands at 3,446, 2,927, 2,856, 2074, 1,638, and 708, cm⁻¹. The strong peak at 3,446 cm⁻¹ corresponds to the O–H stretching of alcoholic and phenolic groups. The absorption bands in the region of 3,000–2,700 cm⁻¹ are characteristic of the C–H group in alkanes. So, the peaks at 2,927 and 2,856 cm⁻¹ with medium to weak intensity suggest aliphatic and aromatic C–H stretching. The peak at 2074 cm⁻¹ accounts for C=N stretching of R–N=C=S. The sharp band at 1,638 cm⁻¹ can be assigned to the amide I bond of proteins in HRSF extract as the band in the region of 1,680–1,500 cm⁻¹ has been shared between the amide bond of protein (sourced from flower extract) and also due to stretching vibrations in the secondary structure of protein resulting in the formation of two new peaks at 1,642 and 1,634 cm⁻¹ (as clearly revealed in the zoomed-in image) in silver nanoparticles. The lower frequency bands at 708 cm⁻¹ can be assigned to C–Cl stretching with C–H bending vibrations out-of-plane. In the IR spectrum of the AgNPs, new bands are observed at 1,338 and 1,314 cm⁻¹ (showed as the zoomed-in image), corresponding to the C–O stretching.

The antimicrobial activity of AgNPs was assessed against all three strains of S. mutans (MTCC-497, ATCC-700610, and the clinical isolate) using the agar well diffusion assay (data pertaining ATCC-700610 and clinical isolate is presented as Supplementary Figure S3 in the supplementary section). The as-synthesized AgNPs were found to be effective against all the strains under study, but the data shown in the whole study corresponds to a resistant strain of S. mutans, MTCC-497. The zone of inhibition (in mm) around each well containing AgNPs was assessed (Figure 4). We also determined the antibacterial activity of the as-synthesized AgNPs against E. coli and L. monocytogenes. The AgNPs exhibited significant antibacterial activity against all the tested organisms. Ni-Ti orthodontic wires were coated with as-synthesized AgNPs or AgNO₃ solution. The impregnated wires were placed on the agar plate that was previously inoculated with resistant isolates of S. mutans. The wire impregnated with AgNPs (50 µg/ml) showed significant S. mutans inhibition (clear zone of inhibition) as compared to AgNO₃ (50 µg/ml) solution-coated wire (Figure 5).

The anti-adherence potential of as-synthesized AgNPs was deduced by incubating Ni-Ti orthodontic wires with AgNPs and their further incubation with S. mutans. It was found that there was maximum bacterial load in terms of (CFU/ml), as shown in the case of uncoated wires (control), while wires coated with aqueous AgNO₃ solution and as-synthesized AgNPs shown less bacterial load than the control. The least bacterial load was found in the case of wires impregnated with AgNPs as compared to both uncoated and AgNO₃ coated wires (Figures 6A,B).

The as-synthesized AgNPs inhibited adherence of S. mutans to the surface of glass tubes. The as-synthesized AgNPs inhibited both sucrose-independent and sucrose-dependent adherence of the treated S. mutans bacteria. The inhibition was more pronounced in sucrose-dependent adherence as compared to the sucrose-free medium. The as-synthesized AgNPs inhibited the S. mutans biofilm formation in a dose-dependent manner (Figure 7A). The AgNPs at a concentration of around 25 μg/ml inhibited around 50% of the formed biofilm.

The as-synthesized AgNPs were found to inhibit cell surface hydrophobicity of S. mutans in a concentration-dependent manner, as shown in Figure 7B. The as-synthesized AgNPs reduced the hydrophobicity to more than 50% at a concentration of ∼100 μg/ml. On the other hand, the precursor AgNO₃ solution failed to reduce the hydrophobicity.

We also determined the anti-biofilm potential of as-synthesized AgNPs against less susceptible isolates of L. monocytogenes and E. coli. The increasing amount of AgNPs (ranging from 1.56 to 200 μg/ml) was dispensed in each well of the microtiter plate. Subsequently, the plate was inoculated with a bacterial strain (for 24 h) specified in the Method section. It can be inferred from the results that the biologically synthesized AgNP-based formulation was successful in inhibiting the drug-resistant bacterial biofilm (less susceptible against gentamicin), as compared to the negative control (Figure 7E). The adherence potential of bacterial cells can be attributed to the high hydrophobicity associated with the bacterial cell surface (Hu et al., 2019). The percentage of hydrophobicity index for E. coli (less susceptible isolate) decreased from ∼58% to ∼34% on treatment with as-synthesized AgNPs. Similarly, the hydrophobicity index for less susceptible L. monocytogenes isolate decreased from ∼64 % to ∼43% (Figure 7F).

We determined the potential of the as-synthesized AgNPs to inhibit glucan synthesis in the treated S. mutans. The AgNPs efficiently inhibited glucan synthesis in a concentration-dependent manner. The AgNPs inhibited around 50% glucan synthesis at around 100 µg/ml concentration (Figure 7C).

To ascertain the anti-biofilm potential of the as-synthesized AgNPs, we assessed S. mutans (drug-resistant MTCC-497)-mediated synthesis of biofilm on the glass surface in the presence of AgNO₃ solution or AgNPs. The fluorescence microscopy revealed the formation of mature biofilm when S. mutans was cultured in absence of AgNPs. The synthesis of profuse biofilm could be attributed to the unhindered proliferation of untreated cells. On the other hand, the treatment with AgNPs resulted in the inhibition of biofilm formation on the glass surface (Figure 8).

Surface morphology or the topographic conditions of Ni-Ti orthodontic wires was studied by the SEM analysis (Figure 9). As revealed by SEM micrographs, the surface of AgNP-coated Ni-Ti orthodontic wires showed less irregular topography in comparison to both aqueous AgNO₃ solution coated and uncoated wires. The uncoated Ni-Ti orthodontic wires were found to possess a significant number of well-distributed irregular cavities and a large number of pores on their surface. On the other hand, fewer topographic irregularities were observed on Ni-Ti wires that were coated with AgNO₃ solution.

In order to investigate biocompatibility, we carried out toxicity assays on RBCs (hemolytic assay) and PBMCs (MTT assay). The obtained results revealed that even at a very high dose (256 μg ml) of AgNP exposure, only around 30% of RBCs were hemolyzed, as shown in Figure 10A; similarly, for PBMCs, it showed around 77% cell viability after exposure to 128 μg/ml (Figure 10B). However, exposure of cells to increasing doses of AgNPs resulted in decreased cell viability. The findings indicate that AgNPs are a stable and non-toxic entity with significant antibacterial action that does not cause toxicity in the host cells.