Biological synthesis and characterisation of silver nanoparticles using Pseudomonas geniculata H10 for pharmaceutical activity

Abstract

In the present study, silver nanoparticles (SNPs) were synthesised for the first time using Pseudomonas geniculata H10 as reducing and stabilising agents. The synthesis of SNPs was the maximum when the culture supernatant was treated with 2.5 mM AgNO₃ at pH 7 and 40°C for 10 h. The SNPs were characterised by field emission scanning electron microscopy‐energy‐dispersive spectroscopy, transmission electron microscopy, dynamic light scattering, X‐ray diffraction and UV–vis spectroscopy. Fourier transform infrared spectroscopy indicated the presence of proteins, suggesting they may have been responsible for the reduction and acted as capping agents. The SNPs displayed 1,1‐diphenyl‐2‐picrylhydrazyl (IC₅₀  = 28.301 μg/ml) and 2,2′‐azinobis‐3‐ethylbenzothiazoline‐6‐sulphonate (IC₅₀  = 27.076 μg/ml) radical scavenging activities. The SNPs exhibited a broad antimicrobial spectrum against several human pathogenic Gram‐positive and Gram‐negative bacteria and Candida albicans. The antimicrobial action of SNPs was due to cell deformation resulting in cytoplasmic leakage and subsequent lysis. The authors’ results indicate P. geniculata H10 could be used to produce antimicrobial SNPs in a facile, non‐toxic, cost‐effective manner, and that these SNPs can be used as effective growth inhibitors in various microorganisms, making them applicable to various biomedical and environmental systems. As far as the authors are aware, this study is the first to describe the potential biomedical applications of SNPs synthesised using P. geniculata.

1 Introduction

The synthesis of metal nanoparticles (NPs) is an interesting topic in nanobiotechnology. These NPs have unique chemical, physical, biological, magnetic and electrical properties due to their large surface area‐to‐volume ratios. Furthermore, these properties make them suitable for various industrial applications, such as in the photonics, electronics and energy generation fields, for cell imaging, drug delivery and biomedical targeting, and as catalysts and biosensors [1]. Among metal NPs, silver nanoparticles (SNPs) have a wide range of applications from electronics to medicine, and are used as optical receptors, intercalating materials and chemical catalysts for molecular labelling [2]. Furthermore, SNPs are finding novel biomedical applications as larvicidal, anticoagulant and thrombolytic agents [3]. In particular, they are effective antimicrobial agents that can be used to treat various infectious diseases without inducing antibiotic resistance [4]. In addition, SNPs, as an antibacterial agent, are effective in a variety of equipment, including metal, fabric, polymers and glass. One gram of SNPs is known to impart antimicrobial characteristics to hundreds of square metres of substrate material [5].

Generally, SNPs are synthesised using physico‐chemical methods [6], but these are energy demanding and expensive. The obvious disadvantages of chemical methods are that they require toxic reducing agents that pose a potential risk to human health and the environment and that the SNPs produced are often present in non‐polar organic solutions that preclude biomedical applications [2, 7, 8]. Therefore, there is an ever‐growing need to develop clean and environmentally safe methods for synthesising SNPs. On the other hand, biological methods using microorganisms [7, 9, 10, 11] or plants [8, 12, 13, 14] require normal temperature and pressure, and thus, could potentially be used to synthesise biocompatible, inexpensive, eco‐friendly and size controlled SNPs [15].

Among the biological sources used for the synthesis of SNPs, microorganisms have been extensively studied and shown to provide one of the best ways of producing SNPs due to their diverse natures and adaptive abilities. In particular, bacteria‐mediated synthesis of SNPs offers advantages over plant and fungi‐based methods, because reaction times are shorter, bacteria are easily cultured and handled, and because their genetic manipulations are relatively straightforward [10]. Microorganism‐based methods of NP production can be classified into intracellular and extracellular synthesis according to the place where NPs are formed. Extracellular biosynthesis is preferred because it reduces the complexity of downstream processing as no additional process is requires to release NPs from biomass [16]. In addition, the extracellular approach results in products containing high levels of secreted biomolecules/proteins that stabilise NPs and improve yield [17]. For this reason, researchers have focused on the use of cultures containing whole cells for SNPs synthesis [15, 17, 18]. However, Ag ions are toxic to cells at concentrations of >1 mM [19], and thus, cell‐free culture supernatants are used for the bulk synthesis of SNPs. The previous studies have showed that culture supernatants of some bacterial strains, such as Morganella sp., Bacillus safensis LAU 13, Streptomyces sp., Pseudomonas spp. and Pseudomonas aeruginosa, could induce the synthesis of SNPs [9, 10, 15, 20, 21]. Nevertheless, the number of bacterial strains, evaluated so far for their ability to synthesise SNPs, is limited and needs to be extended to include bacteria from various habitats in order to understand the mechanisms of synthesis and rapid scale‐up processing. In other words, although there have been many studies on synthesis of SNPs using microorganisms, the isolation and characterisation of new strains for eco‐friendly synthesis are a continuing challenge for mass production and application of SNPs.

In a previous study, we isolated a novel Pseudomonas geniculata H10 from human hair and characterised feather degradation by this strain [22]. Interestingly, this strain produces various bioactive compounds, such as indoleacetic acid and siderophores and a variety of hydrolytic enzymes, including proteases, lipases, cellulases and chitinases, and this led us to investigate the SNPs biosynthesis by P. geniculata H10. In the present study, we optimised reaction parameters for SNPs synthesis by the culture supernatant of P. geniculata H10 and investigated the antimicrobial and antioxidant activities of the SNPs produced. Additionally, this study was extended to characterise the SNPs produced.

2 Materials and methods

2.1 Materials

All chemicals were of analytical grade and purchased from Sigma‐Aldrich (USA) unless otherwise stated. Culture media were purchased from BD (USA).

2.2 Bacterial strain and culture condition

P. geniculata H10 was isolated from human hair in our laboratory, as we previously described [22]. Cells were freshly inoculated into nutrient broth (peptone 1%, beef extract 1%, NaCl 0.5% and pH 6) and incubated at 200 rpm for 24 h at 30°C. The culture was centrifuged at 10 000 × g for 20 min, and the supernatant was used to synthesise SNPs. Nitrate reductase activity in culture was measured using the method described by Harley [23].

2.3 Extracellular synthesis of SNPs

AgNO₃ solution (50 ml, 1 mM) was mixed with 50 ml of culture supernatant in a 250 ml Erlenmeyer flask and incubated at 30°C for 24 h in the dark. The formation of SNPs in the solution was monitored by observing colour changes. As a control, culture supernatant alone and 1 mM AgNO₃ were run simultaneously under the same conditions.

2.4 Optimisation of reaction parameters for SNPs biosynthesis

The effects of different parameters on SNP synthesis were optimised by varying parameters in that order [temperature (30–60°C), pH (5–10) and AgNO₃ concentration (0.5–2.5 mM)]. The pH was adjusted using 0.1 N HCl and 0.1 N NaOH. The influence of reaction time was also investigated by incubating reaction solutions under the optimised condition for up to 12 h. Samples were taken at regular times and the absorbances were measured spectrophotometrically. In addition, the stabilities of as‐synthesised SNPs were examined by exposing them to the ambient conditions for 2 months [24].

2.5 Characterisation of SNPs

SNPs were isolated by centrifuging the reaction mixture at 12 000 × g for 30 min, washed with sterile deionised water three times to remove excess silver ions and other components, and freeze dried for further characterisation.

The UV–visible (UV–vis) absorption spectrum of SNPs was obtained using a UV–vis spectrophotometer (Pharmacia Biotech, Ultrospec 4000, UK) from 300 to 600 nm, and the Fourier transform infrared spectroscopy (FTIR) spectrum of SNPs was obtained from 4000 to 400 cm⁻¹ using a Jasco FTIR 6300 (Japan). The sample for FTIR was prepared by crushing isolated SNPs with KBr pellets. X‐ray diffraction (XRD) analysis of SNPs was carried out using a X‐ray diffractometer (PANalytical, Empyrean series 2, Netherlands) with reflection geometry using CuKα radiation (λ  = 0.154 nm) at 40 kV and 30 mA. Scanning was performed from 2θ  = 20° to 80° at a scan speed of 10°/min. SNP morphology was analysed by field emission scanning electron microscopy (FESEM) (Carl Zeiss, SUPRA 40VP, Germany) at an accelerating voltage of 15 kV. Prior to FESEM, SNPs were spread and sputter coated with platinum. Elemental analysis was conducted by energy‐dispersive spectroscopy (EDS) combined with FESEM. Morphological characteristic of SNP was also investigated using a Tecnai 12 (FEI, Netherlands) transmission electron microscopy (TEM) operated at 120 kV. Size distribution, hydrodynamic diameter and zeta potential were determined by dynamic light scattering (DLS) using a Zeta‐sizer (Malvern Instruments, ZS nano, UK) at room temperature, after dispersing freeze‐dried SNPs in water.

2.6 Determination of antioxidant activity

SNP antioxidant activity was determined using both 1,1‐diphenyl‐2‐picrylhydrazyl (DPPH) and 2,2′‐azino‐bis(3‐ethylbenzthiazoline‐6‐sulphonic acid (ABTS) assay as previously described [25, 26] with some modification. For the DPPH assay, 1 ml of different concentrations (0–100 μg/ml) of SNPs were added to 2.5 ml of 0.3 mM DPPH in ethanol, and incubated at 30°C for 30 min in the dark. Absorbance was then measured at 517 nm. For the ABTS assay, first, the ABTS radical cation was produced by reacting 7 mM ABTS stock solution with 2.45 mM potassium persulphate and allowing the mixture to stand in the dark for 12 h. Separately, the absorbance of ABTS radical solution at 734 nm was reduced to 0.70 ± 0.02 by dilution with distilled water, and then 100 μl of different concentrations (0–100 μg/ml) of SNPs were added to 1 ml of the diluted ABTS radical solution. Absorbance was measured at 734 nm after incubation for 6 min.

Ascorbic acid was used as the positive control. Radical scavenging activity was calculated using the following equation:

$$\begin{array}{rl}\mathrm{Radical}\mathrm{scavenging}\mathrm{activity}(\text{\%})& =\left[1-(A_{\mathrm{sample}}-A_{\mathrm{sample}\mathrm{blank}})\right.\\ & \left.\times /(A_{\mathrm{control}}-A_{\mathrm{control}\mathrm{blank}})\right]\times 100\end{array}$$

IC₅₀ values were calculated from plots of inhibitory or scavenging activity versus SNP concentration.

2.7 Determination of antimicrobial activity

The antimicrobial activity was evaluated using the agar well‐diffusion method [27]. The human pathogenic microorganisms tested were: the Gram‐negative bacteria (Escherichia coli KCCM40880, Klebisiella pneumoniae ATCC13883, Proteus vulgaris ATCC13315, Salmonella choleraesuis ATCC6994, Salmonella typhimurium KCCM 40253, Serratia marcescens KCCM21204 and Vibrio cholerae KCTC2715), the Gram‐positive bacteria (Listeria monocytogenes KCCM40307 and Staphylococcus aureu s ATCC6538) and a yeast (Candida albicans IFO 1385). Test strains were incubated in nutrient broth at 200 rpm for 24 h at 37°C. Culture broths were then spread on Muller–Hinton agar plates. Wells were cut in plates using cork borer (5 mm diameter) and 100 μl (50 μg/ml) of SNP solution was dispensed into each well. Plates were incubated at 37°C for 24 h and inhibition zones were measured. Minimum inhibitory concentration (MIC) of SNP was estimated using the method of Gallucci et al. [12]. FESEM was used to examine the morphological changes in microbial cells after SNP treatment.

2.8 Statistical analysis

All experiments were carried out in triplicate and results were presented as means ± standard deviations. Statistical analysis was performed using SPSS ver. 18.0 (SPSS Inc., Chicago, IL, USA). P values <0.05 were considered statistically significant.

3 Results and discussion

3.1 Visual observations and UV–vis spectrophotometric study of SNPs

SNPs absorb radiation in the visible region (ca. 380–450 nm) due to the excitation of surface plasmon vibrations which is responsible for the brown colour of SNPs in different media [28]. For this reason, SNP synthesis can be confirmed by visual observation and by measuring its surface plasmon resonance (SPR) band using a UV–vis spectrophotometer. The extracellular biosynthesis of SNPs was first investigated using the culture supernatant of P. geniculata H10 in the absence of any reducing agent. Fig. 1 (inset) shows solutions containing culture supernatant and 1 mM AgNO₃ before and after the reaction for 24 h. The colour of the solution turned from pale yellow before to dark‐brown after reaction, indicating the formation of SNPs. No change was observed for controls (culture supernatant and 1 mM AgNO₃) under the same conditions.

Fig. 1.

UV–vis spectrum and colour change of culture supernatant of P. geniculata H10

(a) Before reaction and (b) After reaction

SNP synthesis was confirmed by UV–vis spectral analysis. As shown in Fig. 1, an intense SPR peak was observed at 420 nm. This extracellular reduction of Ag ions offers considerable advantages over intracellular reduction, as intracellular reduction would have required additional processing steps (e.g. ultrasound or detergent treatment) to separate NPs from cells. Furthermore, it has been reported spherical SNPs produce absorption bands at ∼420 nm in the UV–vis spectrum [29], and this was confirmed by our SEM and TEM results (below).

3.2 Optimisation of reaction parameters for SNPs biosynthesis

The synthesis of SNPs is influenced by microorganism type and reaction parameters [30]. Many reports have been published on the extracellular synthesis of SNPs by microorganisms [9, 15, 16, 17, 18], but few studies have examined the optimisation of reaction parameters such as temperature, pH and precursor concentration [31, 32]. To maximise yield, temperature, pH, precursor concentration and time were consecutively optimised step by step.

The influence of temperature on SNP absorption intensity was investigated in the range 30–60°C. As shown in Fig. 2 a, maximum absorption intensity was observed during the reaction at 40°C, followed by at 45°C and then at 50°C. SNP absorption intensities declined at reaction temperatures below 30°C or above 60°C. However, these results did not agree with the results of Deepak and Kalishwaralal [31], who reported reaction kinetics were linearly related to the reaction temperature. Our data indicate that the SNP synthesis was probably the result of the effect of temperature on the biomolecules responsible for the reduction of Ag. To study the effect of time on SNP synthesis at the optimal temperature (40°C), the UV–vis absorption spectra of samples taken at regular times were examined. As shown in Fig. 2 d, an SPR band appeared after ∼4 h of reaction at 420 nm, and this increased in intensity up to 24 h.

Fig. 2.

Synthesis of SNPs by culture supernatant of P. geniculata H10. Error bars (±SDs) are shown when larger than the symbol

(a) Effect of temperature, (b) Effect of pH, (c) Effect of AgNO₃ concentration, and (d)–(f) Spectra show extracellular synthesis of SNPs in a time‐dependent manner under each optimised condition

The influence of pH on SNP synthesis was studied in the range of pH 5–10. As shown in Fig. 2 b, the SNPs were efficiently synthesised over a wide range of pH values (7–10). At pH 6, absorption intensity was much reduced, and at pH 5, Ag ions were precipitated in the form of AgCl (white precipitate) and the formation of SNPs was inhibited. Sintubin et al. [32] reported increased SNPs synthesis at alkaline conditions using lactic acid bacteria due to more competition between H⁺ ions and Ag⁺ ions for negatively charged OH⁻. On the other hand, the acidic solution contains H⁺ ions which compete with Ag⁺ ions for negatively charged binding sites and this decreases the rate of SNPs synthesis [14]. In the present study, although SNPs were synthesised at pH 7–10, broadening of their absorption spectrum at pH 8–10 indicated an increase in the NPs polydiversity. In addition, another peak at ∼510 nm was observed at these pH levels (data not shown), which might have been due to the result of the coupling of NPs. Actually, it is known that a peak at ∼510 nm is observed from dipole–dipole couplings between SNPs [33]. Based on these observations, pH 7 was selected as optimum. At this pH 7, absorption intensity at 420 nm steadily increased with time from 2 to 12 h, indicating continuous SNP synthesis (Fig. 2 e).

The concentration of Ag ions is a critical parameter of SNP synthesis and nucleation. To determine the influence of AgNO₃ concentration on SNPs synthesis, different concentrations of AgNO₃ ranging from 0.5 to 3 mM were added to culture supernatant. Absorption intensity was increased on increasing AgNO₃ concentration to 2.5 mM, but further increases sharply decreased SNP synthesis (Fig. 2 c). Fig. 2 f shows the absorption intensities in supernatant treated with 2.5 mM AgNO₃. As was observed for temperature and pH, the SPR band occurred at ca. 420 nm and the absorption intensity increases up to 12 h, with no considerable shift in peak wavelength. Since the silver ion is toxic to bacterial cells, the concentration of silver ions used to synthesise SNPs should be less than the threshold level (1 mM) [19]. Thus, the synthesis of SNPs at higher concentrations of AgNO₃ in cell‐free culture supernatant favours the mass production of SNPs.

Based on the above, the optimal condition 40°C, pH 7 and 2.5 mM AgNO₃ was established for SNP synthesis. To study the kinetics of SNP synthesis under the optimal conditions, we plotted the absorption intensity at 420 nm against the reaction time. As shown in Fig. 3, the synthesis of SNPs took place within 2 h and the absorption intensity of SNPs increased with time until 10 h. Consequently, the reaction time was reduced by 14 h by the optimisation (24 h). Hence, our optimisation study proved to be an important step in the development of green processes for SNPs synthesis. The stability of synthesised SNPs was monitored regularly for 2 months after the reaction completion, and found to be stable with no evidence for aggregation (data not shown). The stability of the SNPs could be due to the capping agent which probably may be biomolecules secreted by strain H10. In the previous studies on the synthesis of SNPs using bacteria, microalgae or fungi, time required for complete reaction ranged from 24 h to 7 days (24 h for Bacillus sp. GP‐23 [16] and Morganella sp. [10]; 72 h for Aspergillus flavus [7] and Anabena doliolum [11]; 7 days for Streptomyces sp. VITSJK10 [15]. Accordingly, the time required in the present study was considerably shorter. On the other hand, B. safensis LAU13 synthesised SNPs from 8 min and the reaction was complete at 15 min [7], and Desulfovibrio desulfuricans reduced palladium ions to palladium NPs within minutes, although an exogenous reducing agent was added to achieve this rate of formation [34].

Fig. 3.

Kinetic profile of SNPs synthesis under optimised conditions. The inset picture illustrates the test tubes containing SNPs photographed at different working times (2, 4, 6, 8 and 10 h). Error bars (±SDs) are shown when larger than the symbol

3.3 Characterisation of SNPs

FESEM and TEM were used to determine SNP shapes and sizes. As shown in Fig. 4 a, FESEM revealed the synthesis of spherical SNPs with different sizes along with some nanoclusters. Biomolecules bound to the surface of NPs tend to aggregate the NPs partially during sample drying for SEM observation, and as a result, the nanoclusters may have been observed [35]. Similar results were observed for SNPs produced using Bacillus licheniformis [19]. TEM showed that the SNPs formed were spherically shaped and polydispersed. TEM also revealed the size of SNPs between 6.51 and 56.25 nm. It was noticed that the centres of the SNPs were denser than the edges, suggesting that biomolecules such as proteins capped the SNPs [36].

Fig. 4.

SNPs synthesised by culture supernatant of P. geniculata H10

(a) FESEM image, (b) EDS spectrum, (c) TEM images at lower magnifications and (d) TEM images at higher magnifications

EDS was used to confirm SNP synthesis, and showed an intense absorption peak (48.58% by weight) at 3 keV, which is characteristic of metallic silver nanocrystals (Fig. 4 b). This finding indicated SNPs were successfully synthesised in culture supernatants. In addition, peaks corresponding to C, O, N, S, Cl and K were also observed, which were probably caused by the biomolecules present in culture supernatant.

XRD was used to determine the crystalline structure of the SNPs produced (Fig. 5 a), and exhibited four intense peaks at 2θ values of 38.1°, 46.3°, 64.4° and 77.4°, which corresponded to (111), (200), (220) and (311) lattice planes of face‐centred cubic silver, respectively, and agreed with those of the standard spectrum of silver (JCPDS 04‐0783).

Fig. 5.

SNPs synthesised by culture supernatant of P. geniculata H10

(a) XRD pattern, (b) Particle‐size distribution, (c) Zeta potential and (d) FTIR spectrum

Particle size, size distribution and zeta potential are important properties of SNPs because they govern other properties, such as solubility at saturation, dissolution velocity, physical stability and even biological performance [37]. First, hydrodynamic diameter and size distribution were determined by DLS in water. As shown in Fig. 5 b, the hydrodynamic diameter of SNPs produced was ∼42.18 nm (range 20–100 nm). SNP also had a relatively high polydispersity index of 0.244, indicating that the reaction suspension contains some aggregated particles. This result was consistent with UV–vis spectral data and TEM observation. It is notable that the size of the SNPs measured by DLS is larger than that measured by TEM. This difference is due to the fact that the particle size thus obtained is substantially increased by the hydrated capping agents and surface water. That is, biomolecules, such as proteins, surrounding the NPs form a highly hydrated layer, and consequently, larger hydrodynamic diameters were obtained [38, 39]. Zeta potentials provide a means of characterising NP surface charges and stabilities in aqueous suspensions, and usually NPs with zeta potential >±30 mV are considered to be stabilised [34]. The zeta potential curve of SNPs exhibited a single peak at −21.7 mV (Fig. 5 c), indicating SNPs were well dispersed in the colloidal dispersion due to electrostatic repulsion between the adjacent particles and were moderately stable [40]. These findings suggest that the surface capping molecules possessed negatively charged groups.

3.4 Identification of functional groups using FTIR

Several authors have suggested that biological macromolecules secreted by microorganisms play important roles as reducing and capping agents during the NP synthesis reaction [7, 16, 29, 31, 35]. Therefore, FTIR was used to identify the functional groups of macromolecules responsible for the bioreduction and capping/stabilisation of SNPs synthesised using the culture supernatant of P. geniculata H10. Fig. 5 d shows the FTIR spectrum of SNPs. The peaks observed at 3745 and 3448 cm⁻¹ were assigned to the stretching vibrations of secondary and primary amines, respectively. The peaks at 2930–2960 cm⁻¹ were assigned to C–H stretching vibration of protein methylene groups; the peaks at 2848 and 1380 cm⁻¹ to C–H and C–N stretching vibrations of aromatic amines; and the peaks at 2373 and 2321 cm⁻¹ to the nitrile group. The intense peak at 1630 cm⁻¹ was attributed to C=O stretch of the amide I band of a peptide linkage; the peak at 1450 cm⁻¹ to symmetric stretch of the –COO⁻ groups of amino acid residues with free carboxylate groups; the peaks at 1265 and 1096–1060 cm⁻¹ to C–N stretch of aliphatic amines, which are commonly found in proteins; the peak at 800 cm⁻¹ to the amide V band arising from out‐of‐plane N–H bending of peptide linkages [41]; and the peak at 737 cm⁻¹ to metal bound carboxylic (M‐COO⁻) groups. Most of peaks observed in the FTIR spectrum of SNPs are commonly observed for proteins [42, 43], and thus, suggested the presence of proteins in P. geniculata H10 culture supernatant. Furthermore, the result was consistent with the UV–vis spectrum of culture supernatant, which showed an absorption peak at 280 nm (data not shown), corresponding to the aromatic amino acids of proteins, and with EDS results for C, N and O (Fig. 4 b). It was reported that enzymes like nitrate reductase [5], proteins [16, 41] or polysaccharides [44] control SNPs biosynthesis in microorganisms. In this study, the activity of nitrate reductase was not detected in the culture supernatant (data not shown). Therefore, the results indicate that proteins present in supernatant could possibly perform synthesis and stabilisation of SNPs, as it has been demonstrated that protein–NP interactions can occur due to free amino groups, cysteine residues and the carboxylate groups of proteins [31, 45].

3.5 Antioxidant activity of SNPs

Excess free radicals can cause oxidative stress in the human body. These radicals not only cause lipid peroxidation but also oxidise biomolecules and increase the risks of disease, such as atherosclerosis, cancer, emphysema, cirrhosis and arthritis [46]. Since some plant‐mediated SNPs are known for free radical scavengers, we evaluated the free radical scavenging activity of our P. geniculata H10 synthesised SNPs using a DPPH radical scavenging assay. DPPH radical scavenging activity was found to be dependent on SNP concentration (Fig. 6 a). At 5–100 μg/ml, the scavenging activity of SNPs was 6.8 ± 0.8–70.5 ± 1.2%, which was lower than that of the ascorbic acid standard (23.6 ± 0.3–95.1 ± 0.5%). The IC₅₀ values of SNPs and ascorbic acid were 28.301 and 9.114 μg/ml, respectively. The total antioxidant activity of SNPs was using the ABTS assay. SNPs concentration dependently scavenged the ABTS radical, and completely scavenged at 100 μg/ml, whereas ascorbic acid did so at 50 μg/ml (Fig. 6 b). The IC₅₀ value of the SNPs was found to be 27.076 μg/ml, whereas that of ascorbic acid was 21.417 μg/ml. It has been reported that the antioxidant properties of plant‐mediated SNPs are retained due to the effects of plant extract derived capping agents on SNP surfaces [47]. The antioxidant capacity of our SNPs could also be explained to functional groups adhered to them which were produced from the strain H10. These results suggest that P. geniculata H10 synthesised SNPs could potentially be used to scavenge reactive oxygen species in the biomedical and food processing fields.

Fig. 6.

Radical scavenging activities of SNPs synthesised by culture supernatant of P. geniculata H10. Error bars (±SDs) are shown when larger than the symbol. Radical scavenging activities of ascorbic acid (filled circle) and SNPs (open circle)

(a) DPPH and (b) ABTS

3.6 Antimicrobial activity and action mode of SNPs

Metallic SNPs have been reported to have antimicrobial properties [14, 48]. We investigated the antimicrobial activities of SNPs against various pathogenic organisms using the agar well‐diffusion method. SNPs were found to show broad spectrum antimicrobial activity against Gram‐negative, Gram‐positive bacteria, and C. albicans (Fig. 7). Observed inhibitory zone diameters were: E. coli (14.8 ± 1.2 mm), K. pneumoniae (13.6 ± 1.0 mm), P. vulgaris (12.3 ± 1.0 mm), S. choleraesuis (12.1 ± 0.8 mm), S. typhimurium (8.2 ± 0.6 mm), S. marcescens (8.3 ± 0.9 mm), V. cholerae (12.5 ± 0.9 mm), L. monocytogenes (12.3 ± 0.9 mm) and S. aureus (12.1 ± 0.0 mm) at a test concentration of 50 μg/ml, respectively. Interestingly, the highest antimicrobial activity was observed against C. albicans (16.8 ± 0.3 mm), which is a representative pathogenic yeast that causes recurrent mucosal and potentially life‐threatening contagious infections [11]. Furthermore, the number of drugs available for treating C. albicans is limited [49]. Thus, we suggest that the synergistic effect on SNPs and low‐dose amphotericin B (a commonly used anticandidal) be examined for the treatment of C. albicans infections. The cell growth inhibitory effect of SNPs on pathogenic microorganisms was measured by MIC. The MIC of SNPs against E. coil, K. pneumoniae, S. choleraesuis and C. albicans was 1.56 μg/ml; 3.12 μg/ml against P. vulgaris, S. marcescens and V. cholerae. In the case of S. typhimurium, S. aureus and L. monocytogenes, the MIC was obtained at a higher concentration of 6.25 μg/ml. Thus, the SNPs synthesised were found to be a more potent antimicrobial agent in terms of concentration.

Fig. 7.

Antimicrobial activity of synthesised SNPs against pathogenic microorganisms by agar well diffusion assay

(a) E. coli, (b) K. pneumoniae, (c) P. vulgaris, (d) S. choleraesuis, (e) S. typhimurium, (f) S. marcescens, (g) V. cholerae, (h) L. monocytogenes, (i) S. aureus and (j) C. albicans

FESEM analysis was used to investigate the mode of action of SNPs on pathogenic microorganisms. For this purpose, we examined the surface morphologies and shapes of SNP treated and untreated cells. As shown in Figs. 8 a –c, cells of untreated Gram‐negative E. coli exhibited a typical rod‐shape with a well‐developed cell structure and an intact, smooth surface morphology with no apparent extracellular debris. In contrast, E. coli treated with SNPs exhibited significant changes in shape and surface morphology (blebbing, shrinkage and cell distortion), and a tubular shape with the end being destroyed, indicating both outer and inner membranes had been damaged. Furthermore, some cell debris was observed, which was probably caused by release of intracellular constituents following cell membrane damage. Untreated Gram‐positive S. aureus was typically spherical (Figs. 8 d –f), and cell surfaces were intact without evidence of damage. However, SNP treated cells had an irregular shape, and cell surfaces were rougher. There was also congealing of the cell wall. Resultantly, cells of S. aureus were broken down with local rupture in the cell membranes. On the other hand, untreated yeast C. albicans was typically oval shaped with smooth surface (Fig. 8 g) and exhibited a normal budding profile. Whereas C. albicans treated with SNPs showed extensive structural changes including bending, dent and rupture or the collapse of the cell membrane (Figs. 8 c and d). In addition, we observed some intracellular content, which means serious damage to the candida walls took place resulting in the yeast loosing cellular content. Furthermore, it was observed that bud growth was inhibited in SNP‐treated cells, suggesting SNPs inhibit the normal budding process, probably through the destruction of membrane integrity.

Fig. 8.

FESEM image of native and SNPs treated‐cells. White arrows show significant morphological changes including surface roughness, blebbing, distortion and bending. Black arrows show partial disruption of cells, leakage of intracellular content and bursting of cell

(a) Native E. coli, (b, c) Treated E. coli, (d) Native S. aureus, (e, f) Treated S. aureus, (g) Native C. albicans and (h, i) Treated C. albicans

The mechanism responsible for the antimicrobial effect of SNPs is not fully understood. Several authors have proposed SNPs attach to the cell wall and damage membrane integrities, increase membrane permeability and adversely affect respiration functions [50]. It is also possible that the release of Ag ions contributes to the antimicrobial effect of SNPs [48]. Our results suggest that SNPs can interact with microbial cells, damage cell walls and cause cytoplasm leakage, and thus, cell death. On the other hand, it has been reported that Gram‐negative bacteria are more sensitive to SNPs than Gram‐positive bacteria, and suggested that this is due to their thinner walls (7–8 versus 20–80 nm) [51]. However, our results did not show this tendency (Figs. 7 and 8), which suggests antimicrobial mechanisms are species dependent and its mechanism is a matter for further investigation.

4 Conclusion

To develop simple, cost‐effective and eco‐friendly method for synthesis of SNPs, this study focused on extracellular synthesis of SNPs using the culture supernatant of P. geniculata H10 without the use of any chemical reducing agents. The present study showed P. geniculata H10 could be used for the extracellular synthesis of mostly roughly spherical SNPs with a broad antimicrobial spectrum against human pathogenic microorganisms under mild conditions, which offers the advantages large‐scale operation, straightforward processing, and environmental and human safety over chemical synthesis. FTIR analysis demonstrated the presence of proteins in the culture supernatant of P. geniculata H10 as responsible for the reducing silver ions instead of using toxic reducing agents. The synthesised SNPs could scavenge free radicals such as DPPH and ABTS. This study suggests SNPs synthesised by P. geniculata have potential uses in a variety of biomedical and pharmaceutical applications. However, in the present study, we did not confirm the toxicity of SNPs to higher cells, and therefore, we are currently investigating the effects of SNPs on the growth of fibroblasts. As far as we are aware, this study is the first to describe the potential biomedical applications of SNPs biologically synthesised using P. geniculata.