Broad‐spectrum inhibitory effect of green synthesised silver nanoparticles from Withania somnifera (L.) on microbial growth, biofilm and respiration: a putative mechanistic approach

Abstract

Multi‐drug resistance in pathogenic bacteria has created immense clinical problem globally. To address these, there is need to develop new therapeutic strategies to combat bacterial infections. Silver nanoparticles (AgNPs) might prove to be next generation nano‐antibiotics. However, improved efficacy and broad‐spectrum activity is still needed to be evaluated and understood. The authors have synthesised AgNPs from Withania somnifera (WS) by green process and characterised. The effect of WS‐AgNPs on growth kinetics, biofilm inhibition as well as eradication of preformed biofilms on both gram‐positive and gram‐negative pathogenic bacteria was evaluated. The authors have demonstrated the inhibitory effect on bacterial respiration and disruption of membrane permeability and integrity. It was found that WS‐AgNPs inhibited growth of pathogenic bacteria even at 16 µg/ml. At sub‐minimum inhibitory concentration concentration, there was approximately 50% inhibition in biofilm formation which was further validated by light and electron microscopy. WS‐AgNPs also eradicated the performed biofilms by varying levels at elevated concentration. The bacterial respiration was also significantly inhibited. Interaction of WS‐AgNPs with test pathogen caused the disruption of cell membrane leading to leakage of cellular content. The production of intracellular reactive oxygen species reveals that WS‐AgNPs exerted oxidative stress inside bacterial cell causing microbial growth inhibition and disrupting cellular functions.

1 Introduction

There are different ways of producing metal nanoparticles that can be broadly categorised in three groups, i.e. physical, chemical and biological methods [1, 2]. The chemical method of nanoparticle synthesis is relatively expensive and requires the use of aggressive and toxic chemicals as capping and/or reducing agents including sodium borohydride, hydrazine and dimethyl formamide [3]. These chemicals are potentially exerting hazardous effect on health, environmental and biological systems. The gap between need and application of nanoparticles and its negative effect on environment is creating a vacuum for the researchers to focus on development of new techniques that can substantiate both these issues. For green synthesis of nanoparticles, biomolecules (such as carbohydrates, proteins etc.), various types of whole cells systems (fungi, bacteria and algae) and plant parts (leaves, roots, bark, flowers, fruits, roots etc.) have been successfully tested [2]. There is increasing demand of silver nanoparticles (AgNPs) owing to their application in chemical stability, conductivity and antimicrobial activity [4, 5]. Due to its high antibacterial potential, silver ions were used for many centuries for the treatment of wound infections and as an eye drop against Neisseria gonorrhoeae infection [6]. Although AgNPs have been extensively tested for antimicrobial potential, but the exact underlying mechanism of interference is still poorly understood. Greener route of nanoparticle synthesis is especially for antimicrobial potential have many advantages. Now, medicinal plants are targeted for nanoparticle because there is synergistic effect between nanoparticles and phytocompounds [7].

Withania somnifera (L.) (WS) (family: Solanaceae), commonly known as winter cherry or Ashwagandha is used in Ayurvedic and Unani medicine due to its high medicinal value. The non‐toxic nature of this plant has attained high therapeutic potential and the major bioactive compounds detected so far are mainly different types of withanolide [8]. Different parts of this plant, mainly roots are used in traditional medicine in the form of syrup, ointment, infusions, decoction, powder and so on [9]. The plant is now cultivated on large scale to meet the high demand in pharmaceutical industry [10]. WS has been reported to possess antioxidant, aphrodisiac, adaptogen, liver tonic, antimicrobial and anti‐inflammatory activities [11]. Previously, WS leaf powder was used for the synthesis of AgNPs that only evaluated for a brief antibacterial activity [12]. Also, AgNPs synthesised form WS were incorporated into cream to assess its antibacterial potential [9]. To the best of our knowledge, there is no detailed broad‐spectrum and mechanistic report of antimicrobial potential of AgNPs synthesised from WS. Therefore, we have studied the broad‐spectrum effect of AgNP synthesised from WS on bacterial growth, biofilm, respiration and reactive oxygen species (ROS) production on both gram‐positive and gram‐negative pathogenic bacteria to obtain a mechanistic view on its mode of action and potential application.

2 Material and method

2.1 Collection of plant sample and preparation of aqueous extract

WS (root) was obtained as a gift from Himalaya Drug Company, Dehradun, India. The material was authenticated by Himalaya Drug Company as well as Department of Botany, Aligarh Muslim University, Aligarh. Voucher specimen (WS/R‐AGM/HDCO/01‐2017) was deposited in Department of Agricultural Microbiology, Aligarh Muslim University, Aligarh, India. To make 5% extract, 5 g of freshly prepared dry powder was suspended in 100 ml of double distilled water and heated at 100°C for 1 h. The suspension was centrifuged at 10,000 rpm for 10 min at 25°C and then filtered by using a Whatman filter paper no.1. The filtrate obtained was stored at −20°C for further use.

2.2 Synthesis of WS‐AgNPs by aqueous plant extract

To optimise the green synthesis, varying amount of WS aqueous extract (0.25, 0.5, 1.0 and 2.0 ml) was added separately to 20 ml of 1 mM AgNO₃ solution at room temperature. The reduction of AgNO₃ to Ag⁺ was confirmed by the change in colour of suspension from colourless to dark brown. The combination of 0.5 ml extract and 20 ml silver nitrate solution was found to be optimum which was further used in this study. The synthesised AgNPs were harvested by centrifuging at 10,000 rpm for 25 min. The pellet obtained was washed twice with double distilled water and dried overnight in oven at 60°C.

2.3 Characterisation of WS‐AgNPs

2.3.1 UV–Vis spectral analysis

The synthesis of nanoparticles due to reduction of silver nitrate was preliminary monitored by recording the changes in absorption spectra of reaction mixture until maxima was obtained. The absorbance spectra of WS‐AgNPs suspension was recorded in 300–600 nm range using Cintra 10e spectrophotometer (GBC Scientific Equipment Ltd.) at 1 h interval and double distilled water was used as blank for baseline correction.

2.3.2 X‐ray diffraction (XRD) and particle size analysis

The XRD pattern of WS‐AgNPs was obtained on X‐ray diffractometer (MiniFlex^TM II XRD system, Rigaku Corporation, Tokyo, Japan) using CuK_α radiation (λ  = 1.54060 Å) with nickel monochromator in 2θ range of from 20° to 80°. The average crystal size of WS‐AgNPs was calculated using Debye‐Scherrer's equation

$$D=\frac{K\lambda }{\beta \cos \theta }$$ (1)

where D is average crystal size of nanoparticle, K is constant of Debye‐Scherrer's equation, λ is wavelength of X‐ray source used (1.54060 Å) and β is full width at half maximum of the diffraction peak.

Average particle size was also obtained using DynaPro‐TC‐04 dynamic light scattering (DLS) instrument attached to ZetaSizer‐HT (Malvern, UK). WS‐AgNPs were suspended in double distilled water and sonicated for 30 min at 40 W prior to DLS analysis.

2.3.3 Fourier transform infrared (FTIR) spectroscopy

This is an infrared spectroscopic technique that provides information regarding vibrational and rotational modes of motion of a molecule, commonly used for characterisation of nanoparticles. The air‐dried powder of WS‐AgNPs was mixed with spectroscopic grade KBr (in ratio of 1:100) and the spectra were recorded from 4000 to 400 cm⁻¹. FTIR analysis were performed on Perkin Elmer FTIR spectrometer Spectrum Two (Perkin Elmer Life and Analytical Sciences, CT, USA) in the diffuse reflectance mode at a resolution of 1 cm⁻¹ [13].

2.3.4 Scanning electron microscopy (SEM) and transmission electron microscopy (TEM) analysis

Finely powdered WS‐AgNPs were subjected to scanning electron microscope (JSM 6510LV scanning electron microscope, JEOL, Tokyo, Japan) to obtain their surface morphology and elemental composition. Elemental composition of WS‐AgNPs was obtained using Oxford Instruments INCAx‐sight EDAX spectrometer equipped to SEM. Images were captured at acceleration voltage of 20 kV.

Transmission electron micrographs of WS‐AGPs were obtained using transmission electron microscope (JOEL‐2100, Tokyo, Japan). For TEM analysis, slide of nanoparticles was prepared by placing 10 μl of aqueous suspension of WS‐AgNPs on a TEM grid. The SEM and TEM images were recorded at different magnifications at USIF, AMU, Aligarh, India.

2.4 Bacterial strains and growth condition

WS‐AgNPs were tested for their antibacterial and antibiofilm activity against both gram‐positive and gram‐negative bacteria. Staphylococcus aureus (MTCC 3160), Streptococcus mutans (MTCC 497), Pseudomonas aeruginosa (PAO1) and Salmonella typhimurium (MTCC 98) were used in this study. S. mutans was cultured in brain‐heart infusion medium and other used bacteria were grown in Luria‐Bertani medium.

2.5 Antibacterial activity of WS‐AgNPs

Sensitivity of WS‐AgNPs was preliminarily investigated using agar well diffusion assay [14]. About 100 µl of overnight grown cultures (10⁷ CFU/ml) were spread over petri plates to obtain bacterial lawns. Wells of 8 mm were punctured onto the plates in which 100 µl of different concentration of nanoparticles were loaded and incubated overnight at 37°C.

2.6 Determination of minimum inhibitory concentration (MIC)

MIC values of WS‐AgNPs against each bacterium were determined on 96‐well microtitre plate using TTC (2,3,5‐triphenyl tetrazolium chloride) as adopted by Klančnik et al. [15]. Two‐fold dilution of WS‐AgNPs of varying concentrations (08–256 µg/ml) were made and 10 μl of overnight grown culture was inoculated. The group without nanoparticles was used as control. The MIC is the lowest concentration at which there is no visible growth after 24 h on the basis of metabolic activity [16]. The presence of metabolically active bacterial cells was determined after adding 10 μl/well of TTC (2 mg/ml) followed by incubation for 30 min. The minimum concentration at which no colour was observed was taken as MIC.

2.7 Growth curve analysis

The effect of WS‐AgNPs was tested on the growth kinetics of each bacterium and was assessed by optical density measurements as described earlier [17]. Bacteria were grown in 50 ml flasks in the presence of WS‐AgNPs and group without nanoparticles were taken as control. Aliquots of 1 ml were drawn at 2 h interval and optical density was measured at 620 nm over 24 h.

2.8 Biofilm inhibition assay

Biofilm inhibition potential of WS‐AgNPs against tested bacteria was quantitatively estimated using microtitre plate assay as described by Kalishwaralal et al. [18] with slight modifications. Briefly, 10 µl of overnight grown cultures were seeded in 96‐well sterile plates containing nutrient media and WS‐AgNPs were added to make final concentrations of (1/4) × MIC, (1/2) × MIC and 1 × MIC of respective bacterial cultures followed incubation for 24 h at 37°C. Wells without nanoparticles were taken as control. The media was decanted and planktonic bacterial cells were removed by washing thrice with phosphate buffer saline (PBS) pH 7.4. Biofilms formed by sessile cells in plate were stained with crystal violet (0.1% w/v) and excessive stain was removed by gentle washing. Biofilms formed were finally dissolved in 70% ethanol and absorbance was recorded at 595 nm.

2.9 Biofilm inhibition on glass coverslip

Inhibition of biofilm formation on surface of sterile glass coverslips was determined using 24‐well polystyrene plate containing medium. To each well, test organisms were seeded in the presence of varying concentrations of WS‐AgNPs [(1/4) × MIC, (1/2) × MIC and 1 × MIC]. Wells without WS‐AgNPs were taken as control. The bacteria were allowed to grow for 24 h at 37°C. After incubation, media was discarded and coverslips were washed thrice with autoclaved PBS. Coverslips were then stained with 0.1% crystal violet and then incubated for 10 min. Excess amount of dye was washed in PBS and coverslips were visualised under light microscope (Olympus BX60, Model BX60F5, Olympus Optical Co. Ltd. Japan) equipped with colour VGA camera (Sony, Model no. SSC‐DC‐58AP, Japan). The images presented are at 40× magnification.

2.10 Scanning electron microscopy

The ability of WS‐AgNPs to inhibit biofilms was analysed by SEM as described earlier [19]. Briefly, test bacteria were cultured in 24‐well microtitre plates for 24 h in the presence of varying concentrations [(1/4) × MIC, (1/2) × MIC] of WS‐AgNPs containing sterile glass cover slips. Test group without WS‐AgNPs was taken as control. Unbound bacterial cells were then washed in 1× sterile PBS and subsequently fixed with 2.5% glutaraldehyde. The biofilm and attached cells were dehydrated and gently washed with a gradient of ethanol (30, 50, 70, 80, 95 and 100%) for 10 min and left at room temperature to completely dry it. The topography of biofilms was visualised using SEM as mentioned above.

2.11 Biofilm disruption assay

For assay of disruption of preformed biofilm, method adopted by Gaidhani et al. [20] was used with slight modifications. Briefly, bacterial cultures were grown in 96‐well microtitre plates for 24 h at 37°C. The unbound planktonic cells were removed by washing and fresh medium was added to each well. WS‐AgNPs were added to wells to make desired final concentration (25 × MIC, 50 × MIC and 100 × MIC) and incubated overnight. The wells were then washed thrice with PBS and stained with 0.1% crystal violet. Excess amount of crystal violet was removed by gentle washing. The biofilm in wells was then dissolved in 70% ethanol and optical density (595 nm) was measured. Wells without nanoparticles were taken as control.

2.12 Effect of WS‐AgNPs on membrane integrity of bacteria

Disruption of bacterial membrane leads to leakage of cytoplasmic constituents. The relative amount of intracellular content (that may be DNA or RNA) released was assayed by recording the absorbance of samples at 260 nm as reported earlier [21]. Briefly, overnight grown bacterial cultures were aliquoted into a 96‐well microtitre plate and cells were treated with varying concentration (2 × MIC, 4 × MIC and 8 × MIC and 16 × MIC) of WS‐AgNPs. After 2 h of incubation, cell suspension was filtered with 0.2 µm syringe filters to remove out bacterial cells. The supernatant was diluted accordingly and absorbance was recorded 260 nm. Treatment group with no WS‐AgNPs was taken as control. The data presented is the ratio of OD_260 nm of treated group to control group.

2.13 Respiration inhibition assay

Inhibition in cellular respiration by WS‐AgNPs was evaluated by dehydrogenase assay, as described earlier [22]. Briefly, bacterial cells were harvested from log phase of their growth by centrifugation. Cells were washed twice with PBS to remove media and then re‐suspended in buffer to obtain OD_600 nm value of 0.5. About 100 µl of the cell suspension was transferred to 96‐well microtitre plates. To each well, WS‐AgNPs were added to obtained desired concentrations of (1/2) × MIC, 1 × MIC and 2 × MIC. Group without WS‐AgNPs was taken as control. Subsequently, TTC (0.1% w/v) was added to each well. The colour of cell suspension changed from colourless solution to pink coloured formazan which was measured by recording absorbance at 450 nm.

2.14 Production of intracellular ROS

The relative amount of intracellular ROS produced in bacterial cells was evaluated using 2,7‐dichlorofluorescein diacetate (DCFH‐DA) which is oxidation‐sensitive fluorescent probe [23]. DCFH‐DA enter inside cell by passive diffusion through the cell membrane where it is deacetylated by esterases to form 2,7‐dichlorofluorescein (DCFH). DCFH reacts with ROS to form dichlorofluorescein (DCF) which gets trapped inside cells making them fluorescent. Briefly, bacterial cells were harvested from their log phase to obtain metabolically active cells by centrifugation and washing with PBS. Cells were then re‐suspended in sterile media and the DCFH‐DA was added. The bacteria were incubated for 30 min in shaking incubator at 37°C. Bacterial cells were then centrifuged and washed twice to remove free DCFH‐DA from medium. These cells were then aliquoted in 96‐well microtitre plate and treatment of varying concentration [(1/2) × MIC, 1 × MIC and 2 × MIC] of WS‐AgNPs were given. Group without WS‐AgNPs was taken as control. The fluorescence intensity of cell suspension was recorded at λ _ex of 488 nm and λ _em of 535 nm using spectrofluorophotometer (RFPC5301, Shimadzu, Japan).

2.15 Statistical analysis

All experiments were performed in triplicate and the data obtained were presented as average values and the level of significance was analysed using Student's t‐ test in Microsoft excel 2016.

3 Results and discussion

3.1 Green synthesis of AgNPs using aqueous root extract of WS

Till date, various biological agents such as medicinal plants, microbes and natural products have been extensively used for synthesis of nanoparticles [24, 25]. In this investigation, aqueous extract of WS was used for synthesis of AgNPs. For optimisation of synthesis, various combinations of WS extract (0.25, 0.5, 1.0 and 2.0 ml) was added separately to 20 ml of 1 mM AgNO₃ solution. Combination of 0.5 ml extract and 20 ml AgNO₃ was found to be optimum, therefore, this combination was used further. Initially, silver nitrate solution was colourless which changed to brown and finally to black in 6 h as shown in Fig. 1 a (inset). The change in colour indicated the reduction of AgNO₃ to Ag⁺ demonstrating the reducing potential of WS extract for biosynthesis of AgNP [26].

Fig. 1.

Characterisation of WS‐AgNPs

(a) UV–Vis spectra of formation of AgNPs using WS‐AgNPs at different time intervals and change in colour of silver nitrate solution (inset), (b) XRD pattern of WS‐NPs, (c) Hydrodynamic size of WS‐AgNPs, (d) FTIR spectrum of WS‐AgNPs and aqueous extract of W. somnifera

3.2 Characterisation of WS‐AgNPs

3.2.1 UV–Vis spectral studies

The UV–Vis absorption spectra of WS‐AgNPs synthesis are shown in Fig. 1 a, there was absorption band at 440 nm that increased progressively with time. The absorption maximum at 440 nm is characteristic feature of spherical AgNP which is attributed to the surface plasmon band of AgNPs [5]. The broad absorption peak is due to polydispersed size of the particles [27, 28]. AgNPs exhibit brown colour in water due the oscillation modes arising from electromagnetic field in the visible range and also due to the collective oscillations of conduction electrons [29]. The absorption spectra of WS‐AgNPs were found to be slightly asymmetrical indicating the presence of different sized particles with some anisotropy in the shape of the AgNPs [30]. After synthesis, various phytoconstituents might cap the AgNPs that act as stabilising agent.

3.2.2 XRD and particle size analysis

The green synthesised WS‐AgNPs were characterised by monitoring its XRD pattern. The nano‐crystalline nature of WS‐AgNPs is clearly evident by the peak broadening as shown in Fig. 1 b. The diffractions peak index suggests that the WS‐AgNPs are crystalline in nature (JCPDS File No. 03‐0921) [31]. The average particle size was found to be 52.19 nm. The stability of the nanoparticle is evident from the DLS data (58.8 nm) obtained after 1 year of its synthesis. The size range and shape were validated further using TEM.

3.2.3 FTIR spectroscopy

FTIR analysis has been performed to assess the role of various phytoconstituents of WS, responsible for the synthesis as well as capping and stabilisation of WS‐AgNPs. Fig. 1 d shows the FTIR spectra of WS‐AgNPs and aqueous extract of WS. A broad peak at about 3440 cm⁻¹ of both WS‐AgNPs and aqueous extract is attributed to vibrations of hydroxyl (–OH) group [32]. The absorption band at 1642 cm⁻¹ is associated to N–H bend of amines revealing that proteins are one of the major capping agents in synthesised nanosilver [33]. This amide peak got shifted to lower wavenumbers which indicates the changes in secondary conformation of proteins [34]. A prominent peak around 1022 cm⁻¹ was also recorded which is ascribed to the C–Br stretching [35]. Previously, it has been reported by Niraimathi et al. [36] that the proteins present in the plant extract act as a capping agent. The possible mechanism of absorption of plant material mainly flavanones or terpenoids is through carbonyl groups interactions in absence of other strong ligating agents at sufficiently high concentrations [37]. The presence of reducing sugars terpenoids in plant extract plays a role in reduction of metal ions (Ag⁺) and formation of AgNPs [38]. All these phytoconstituents present in aqueous extract of WS would have played a significant role in synthesis as well as stabilisation of WS‐AgNPs.

3.2.4 SEM and TEM analysis

Scanning electron micrographs were used to examine the morphology of WS‐AgNPs. Fig. 2 b shows the electron micrograph of WS‐AgNPs which shows that most WS‐AgNPs were spheroidal with rough surfaces. Energy dispersive X‐ray (EDX) spectrum of WS‐AgNPs depicts the presence of carbon, oxygen, chlorine and silver atoms as major components, consisting of 28.28, 25.22, 7.96 and 38.55%, respectively. The size of nanoparticles and their distribution depends on the relative rate of nucleation and growth processes as well as extent of agglomeration [39].

Fig. 2.

Scanning and transmission electron micrographs

(a) Transmission electron micrographs of WS‐AgNPs at 100,000× and 200 kV, (b) Panel (i) shows the SEM images of WS‐AgNPs; panel (ii) represents histogram of weight% of major elements in WS‐AgNPs; panel (iii) represents the EDX spectrum of WS‐AgNPs

Fig. 2 a is the representation of transmission electron micrograph of WS‐AgNPs. It shows a good illustration that how nanoparticles formed by extract of WS are in the size range of 40–60 nm. Furthermore, most of the particles revealed by TEM appeared spheroidal along with few anisotropies. Nanoparticles were found to form small aggregates that might be due to improper capping. The variation in size and shape of nanoparticles formed by different green synthesis approaches had also been reported earlier [40, 41].

3.3 Antibacterial activity of AgNPs

Preliminary antibacterial activity of WS‐AgNPs was investigated against S. aureus, P. aeruginosa, S. mutans and S. typhimurium using agar well diffusion method. The zones of inhibition around each well by WS‐AgNPs, silver nitrate solution and aqueous plants extract are represented in Table 1. Among tested organisms, S. mutans was found to be sensitive with inhibition zone of 13 and 15 mm for WS‐AgNPs at 100 and 150 µg, respectively. WS‐AgNPs exhibited largest zone of inhibition of 15 and 16 mm at 100 and 150 µg against P. aeruginosa. It is interesting to note that inhibition zone of WS‐AgNPs was found to be slightly greater for approximately all tested bacteria compared to silver nitrate solution. The relatively enhanced antimicrobial activity of WS‐AgNPs as compared AgNO₃ is due to their large surface area, which provides better contact with microorganisms and the synergistic effect when particles combine with other natural compounds [7, 42]. Cardozo et al. [43] showed that AgNPs and phenazine‐1‐carboxamide together increased the antibacterial effect against methicillin‐resistant S. aureus strains by 32‐fold, resulting in morphological alterations of the bacterial cell wall. The mechanism of action involves the attack on the respiratory chain, cell division and bacterial membrane that leads to cell death. The nanoparticles also release Ag⁺ in bacterial cells, enhancing the bactericidal activity [44].

Table 1.

Zone of inhibition of WS‐AgNPs, AgNO₃ solution and aqueous extracts of W. Somnifera by agar well diffusion assay against bacteria

Bacteria	Zone of inhibition, mm
	Doxycycline	AgNO3	WS‐AgNPs		W. Somnifera extract
	100 µg	100 µg	100 µg	150 µg	100 µg
S. aureus (MTCC 3160)	34.66 ± 2.08	13.66 ± 1.52	14.33 ± 3.21	17.33 ± 2.08	—
S. mutans (MTCC 497)	35.33 ± 3.05	16.33 ± 2.08	20.66 ± 2.51	22.66 ± 1.52	—
P. aeruginosa (PAO1)	20.33 ± 2.08	15.33 ± 1.52	16.66 ± 1.52	20.33 ± 1.52	—
S. typhimurium (MTCC 98)	36.00 ± 2.64	16.00 ± 1.73	16.66 ± 1.15	17.66 ± 2.08	—

The data presented is mean ± SD of three replicates. No significant difference was obtained between AgNO₃ and WS‐AgNPs (100 µg) and WS‐AgNPs (100 µg) and WS‐AgNPs (150 µg) groups of treatment.

3.4 Determination of MIC of WS‐AgNPs

MIC of WS‐AgNPs against selected bacteria was determined to find out its effective concentration under in vitro conditions. Among tested bacterial strains, P. aeruginosa exhibited maximum MIC value of 64 µg/ml whereas S. mutans was found to be most sensitive with MIC of 16 µg/ml. WS‐AgNPs was found to be inhibitory to S. typhimurium and S. aureus at 32 µg/ml. Assessment of inhibitory effect on biofilm formation was done at MIC and sub‐MIC (i.e. concentrations below the MICs).

3.5 Effect of WS‐AgNPs on growth kinetics of bacteria

The dose‐dependent effect of WS‐AgNPs was studied on the growth kinetics of test bacteria at their respective (1/2) × MIC, 1 × MIC and 2 × MIC. The group without treatment was taken as control. It is evident from Fig. 3 b that at (1/2) × MIC there was only 05.51% (the data was insignificant with respect to control) growth inhibition of S. mutans after 24 h and WS‐AgNPs exhibited 68.97 and 91.79% inhibition at 1 × MIC and 2 × MIC. Similarly, there was 64.59 and 95.35% inhibition in growth of P. aeruginosa at 1 × MIC and 2 × MIC of WS‐AgNPs, respectively (Fig. 3 c). For S. aureus, at 1 × MIC and 2 × MIC, there was 58.90 and 81.08% growth inhibition (Fig. 3 a). In a similar fashion, the reduction in viability of S. typhimurium was observed as 69.39% at inhibitory concentration (1 × MIC) and twice amount (2 × MIC) inhibited the growth by more than 90% as depicted in Fig. 3 d. It has been found earlier that AgNPs synthesised from Allium cepa extract exhibited an articulate effect on the growth curve of S. typhimurium [45]. AgNPs have shown toxic effect on S. aureus that increased the lag phase of their growth. It was found that interaction of AgNPs with DNA resulted in loss of their replicating ability and breakdown of cell wall [46]. AgNPs (0.1%) in combination with tissue conditioner exhibited bactericidal effect against S. aureus and S. mutans. One percent of Ag⁺ is bactericidal with complete loss in viable of tested cells [47]. Strong antibacterial activity of AgNPs is reported due to their increased surface area which provides maximum contact with the environment [45]. Another mechanism for bactericidal activity of AgNPs is by interaction with sulphur containing protein that leads to H₂ O₂ production ultimately causing cell death [48].

Fig. 3.

Growth curve of tested bacteria in absence and presence of varying concentration [(1/2) × MIC, 1 × MIC and 2 × MIC] of WS‐AgNPs

(a) S. aureus, (b) S. mutans, (c) P. aeruginosa, (d) S. typhimurium

* represents significance at p  ≤ 0.05 with respect to control; # represents significance at p  ≤ 0.01 with respect to control

3.6 Inhibition of biofilm formation by WS‐AgNPs

The biofilm inhibition of tested bacteria at different concentrations of WS‐AgNPs is shown in Fig. 4 a and MIC was taken as positive control. Among tested bacterial strains, the maximum inhibition in S. mutans was recorded. At MIC, there was 90.56% reduction in biofilm as expected due to growth inhibition effect. However, sub‐MICs [(1/4) × MIC and (1/2) × MIC] also significantly reduced the biofilm formation by 26.96 and 54.47%, respectively. WS‐AgNPs successfully inhibited the biofilm formation of S. aureus by 88.32% at MIC value. At (1/2) × MIC, there was more than 40% inhibition in biofilm formation against S. aureus. Biofilm of P. aeruginosa was also inhibited by 71.84, 48.56 and 18.50% at 1 × MIC, (1/2) × MIC and (1/4) × MIC. Similarly, treatment of (1/4) × MIC and (1/2) × MIC of WS‐AgNPs resulted in 20.61 and 50.72% decrease in biofilm formation of S. typhimurium. Earlier, Kalishwaralal et al. [18] found that AgNPs synthesised from B. licheniformis impeded biofilm formation of P. aeruginosa and the probable mechanism they proposed that the diffusion of AgNPs through water channels (pores) resulted in its anti‐microbial function. AgNPs synthesised from Eucalyptus globulus exhibited a dose‐dependent inhibitory effect on biofilm formation (>80%) of S. aureus and P. aeruginosa at 30 μg/ml [37]. The tested microorganisms are potent biofilm formers that make them causative agents of several infections including superficial skin suppurations and septicemias [49]. Moreover, other nanoparticles such as zinc oxide and copper oxide nanoparticles have also shown to inhibit biofilm formation of S. mutans on teeth model [50]. While with best of our knowledge, AgNPs has not been previously reported against biofilms of S. mutans. This is the first report on in vitro biofilm inhibition of S. mutans.

Fig. 4.

Effect of MIC and sub‐MICs of WS‐AgNPs on inhibition of biofilm formation

(a) Effect of MIC and sub‐MICs [(1/2) × MIC and (1/4) × MIC] of WS‐AgNPs on inhibition of biofilm formation on S. aureus, S. mutans, P. aeruginosa and S. typhimurium, (b) Effect of WS‐AgNPs (25 × MIC, 50 × MIC and 100 × MIC) on disruption of preformed biofilm of S. aureus, S. mutans, P. aeruginosa and S. typhimurium

* represents significance at p  ≤ 0.05 with respect to control; # represents significance at p  ≤ 0.01 with respect to control

3.7 Biofilm inhibition on glass coverslip

To validate biofilm inhibition data, the architecture biofilms grown in absence and presence of WS‐AgNPs [(1/4) × MIC, (1/2) × MIC and 1 × MIC] was analysed by light microscopy. The microscopic image of biofilms of tested bacteria with and without WS‐AgNPs is shown in Fig. 5. It is quite clear from the microscopic images that biofilm formation was significantly impaired by supplementation of nanosilver. The untreated samples showed quite thick cluster of biofilms in all tested microbes. WS‐AgNPs had a dose‐dependent effect on the biofilm formation and at highest concentration, there was almost no visible biofilm.

Fig. 5.

Inhibition of biofilm of test bacteria at varying concentration of WS‐AgNPs under light microscope

(a) Untreated control, (b) (1/4) × MIC of WS‐Ag‐NPs, (c) (1/2) × MIC of WS‐Ag‐NPs, (d) 1 × MIC of WS‐Ag‐NPs

Panel I – S. aureus, panel II – S. mutans, panel III – P. aeruginosa and panel IV – S. typhimurium

3.8 Scanning electron microscopy

The biofilm of S. aureus, P. aeruginosa and S. mutans grown on glass slide in 24‐well titre plate for 24 h in absence and presence of (1/2) MIC value of WS‐AgNPs is shown in Fig. 6. It is evident from electron micrograph that biofilm formation by untreated S. aureus showed extensive biofilm formation (Fig. 6 a). S. aureus cultured without WS‐AgNPs exhibited normal cellular morphology and smooth cell surfaces. When the same bacterial strain was grown in presence of (1/2) × MIC, the colonisation and biofilm formed by bacteria were fairly inhibited as depicted in Fig. 6 b. Similarly, untreated cells of P. aeruginosa were found in dense biofilm as clumped and clustered mass of cell (Fig. 6 a). The rod‐shaped cell of P. aeruginosa can be easily seen in polymeric matrix with normal smooth cell morphology. The aggregation of P. aeruginosa cells was remarkably reduced in presence of WS‐AgNPs (Fig. 6 b) and the presence of bound nanoparticles onto their surfaces are clearly visible that resulted in damage to their membrane. The polymeric matrix encapsulating bacterial cells was absent. The control cells of S. mutans were present in chains and patches with smooth cell surfaces (Fig. 6 a). Treatment with sub‐MIC concentration of WS‐AgNPs resulted in reduced clumping and was found in much shorter chains as well as rough cell surfaces that has been damaged by the AgNPs (Fig. 6 b). Scanning electron microscopic images clearly demonstrated that glass surfaces treated with WS‐AgNPs significantly reduced the colonisation and biofilm formation compared to their respective untreated bacterial controls.

Fig. 6.

Inhibition of biofilm of S. aureus, S. mutans and P. aeruginosa by WS‐AgNPs under scanning electron microscope

(a) Untreated control, (b) In presence of (1/2) × MIC of WS‐Ag‐NPs

SEM provided a visual evidence that also somewhat led us to understand the putative mechanism of biofilm inhibition by nanoparticle. The results showed the reduced clumping and aggregation of cells with increased roughness of the cell surface suggesting that it has been caused by the nanosilver. It has been shown by Kostenko et al. that nanocrystalline nature of AgNPs produced a significant decrease in number of viable cells within tested biofilms of P. aeruginosa and S. aureus. Even the smaller amounts of nanosilver (2.5–6.5 µg/ml) was enough to kill 90% of the biofilm cells and they suggested that application of silver in dressings could significantly improve wound healing by enhancing the susceptibility of bacterial cells within biofilms [51]. Moreover, confocal microscopic study using double staining (propidium iodide and Con A‐FITC) revealed that treatment of AgNPs resulted in significant decrease in live bacterial cells. The three‐dimensional structure of biofilm was also found to be disrupted with negligible amount of glycocalyx matrix [19]. Biofilm matrix contains water channels for nutrient transportation through which AgNPs may diffuse and impart anti‐microbial function. Nanoparticles also have tendency to attach and penetrate bacterial membranes, accumulating inside bacterial cells, and ultimately killing of bacteria [52]. Our findings on inhibition of biofilm of four bacterial pathogens by WS‐AgNPs indicated a broad‐spectrum activity and also validate the findings of above works on S. aureus. Inhibition of biofilm may also be attributed to interference in the bacterial quorum sensing [53].

3.9 Eradication of preformed biofilms

Bacteria under biofilm state may tolerate antibiotic and other toxic substances to several hundred times more than its planktonic mode of growth [54]. Therefore, eradication of preformed biofilms by chemical agents is difficult. WS‐AgNPs were tested for their ability to eradicate the preformed biofilms. The results showed that at 100 × MIC, the amount of residual biofilm was reduced to 54.30% in P. aeruginosa. The treatment at 25 × MIC and 50 × MIC also disrupted the preformed biofilm by 18.85 and 28.34% as compared to untreated P. aeruginosa cells. However, at lower concentrations (3.12 × MIC, 6.25 × MIC and 12.5 × MIC), no significant biofilm eradication was observed (data not shown). For S. aureus, WS‐AgNPs successfully reduced the 24 h old formed biofilm to 43.03% at 100 × MIC. The tested lowest concentration, i.e. 25 × MIC also diminished preformed biofilm by more than 15% against S. aureus. The residual biofilm was found to be 70.27, 57.53 and 53.30% at 25 × MIC, 50 × MIC and 100 × MIC, respectively, in S. typhimurium. The preformed biofilm of S. mutans was also successfully eradicated by 11.78, 27.21 and 51.06% by the addition of 25 × MIC, 50 × MIC and 100 × MIC of WS‐AgNPs, respectively. The above presented data (Fig. 4 b) clearly demonstrate the potency of WS‐AgNPs to eradicate mature biofilm of both gram‐positive and gram‐negative bacteria. Biofilms are embedded in exopolymeric substances, mainly composed of polypeptides, polysaccharides and nucleic acids that blocks the entry of metal ions, antibiotics and other antibacterial agents [55]. Our findings revealed that WS‐AgNPs successfully penetrated the biofilm and achieved more than 50% eradication of preformed biofilm of tested pathogenic bacteria. Previously, it has been found that nanosilver synthesised form A. calcoaceticus inhibited and disrupted the biofilm of 22 tested pathogenic bacteria. The diffusion tendency of AgNPs in biofilm matrix has been shown earlier by Peulen and Wilkinson [56] that might be the possible mechanism of biofilm eradication. The present study further highlights the role of WS‐AgNPs in eradicating biofilm of both gram‐positive and gram‐negative bacteria.

3.10 Effect of WS‐AgNPs on bacterial membrane permeability and integrity

The release of intracellular contents from bacteria is useful indicator for determination of their membrane integrity [21]. The relative amount of bacterial intracellular content released can be determined by recording optical density at 260 nm. The ratio of OD_260 nm of WS‐AgNPs treated sample to OD_260 nm untreated sample is shown in Fig. 7 a. It is evident from the data that WS‐AgNPs produced dose‐dependent effect on all tested bacteria. Exposure of 2 × MIC, 4 × MIC, 8 × MIC and 16 × MIC of WS‐AgNPs to P. aeruginosa resulted in 1.21, 1.81, 2.01 and 2.90‐fold increase in OD_260 nm, respectively, compared to untreated bacterial cells. Similarly, in presence of 2 × MIC of nanosilver, there was 25.9% increase in OD_260 nm of S. aureus compared to their respective untreated control cells. When concentration was increased to 16 × MIC, relative OD_260 nm value increased to 2.10 times. Amount of intracellular content released from S. mutans was also remarkable that increased to 2.25‐fold of control at 16 × MIC. Similarly, there was 1.47, 2.65, 3.03 and 3.29‐fold increase in OD_260 nm value for S. typhimurium at 2 × MIC, 4 × MIC, 8 × MIC and 16 × MIC, respectively. A mechanistic study revealed that AgNPs caused the destruction of cell wall S. aureus. Previously it has been reported that nanosilver induced the structural variation in peptide portion of peptidoglycan that caused the release of muramic acid. Interaction of AgNPs to bacterial wall created pits on both sides of the peptide and glycan ports. AgNPs attached to β‐1,4 bonds N‐acetylglucosamine and N‐acetylmuramic acid that destroyed the bond of glycan strands [57]. Copper nanoparticles are also documented to exhibit similar activity on both S. aureus and P. aeruginosa [58].

Fig. 7.

Effect of WS‐AgNPs on bacterial membrane permeability and integrity

(a) Effect of WS‐AgNPs on bacterial membrane permeability and integrity of S. aureus, S. mutans, P. aeruginosa and S. typhimurium. The data is presented in terms of ratio of absorbance at 260 nm of treated bacteria in presence of 2 × MIC, 4 × MIC, 8 × MIC and 16 × MIC of WS‐AgNPs compared to their respective untreated controls, (b) WS‐AgNPs induced ROS generation in treated S. aureus, S. mutans, P. aeruginosa and S. typhimurium in absence and presence [(1/2) × MIC, 1 × MIC and 2 × MIC] of WS‐AgNPs

* represents significance at p  ≤ 0.05 with respect to control; # represents significance at p  ≤ 0.01 with respect to control

3.11 Effect of WS‐AgNPs on bacterial respiration

One of known mechanism for interruption of bacterial growth is by inhibiting its respiration. Fig. 8 shows concentration‐dependent respiration inhibition by WS‐AgNPs. After 2 h of growth, there was more than five‐fold decrease in respiratory activity of P. aeruginosa at 2 × MIC as compared to control. Similarly, S. aureus exhibited more than three times lesser respiration on comparing untreated control. In S. typhimurium, exposure of (1/2) × MIC, 1 × MIC and 2 × MIC of WS‐AgNPs resulted in decrease in its respiration by 20.18, 53.21 and 89.81% after 120 min of incubation. Among all tested microorganisms, the respiration of S. mutans was least inhibited that only by ∼60% at 2 × MIC. It has been found that silver ions gave significant inhibition (∼80%) of respiration [59]. In a comparative study, it was found that nanosilver more effectively (86%) inhibited bacterial respiration as compared to silver ions (42%) and silver chloride colloids (46%) at 1 mg/l [60]. Not only AgNPs, but respiration inhibition by copper oxide micro‐spheres were also found in P. aeruginosa and S. aureus at varying concentrations [61].

Fig. 8.

WS‐AgNPs induced ROS generation in treated S. aureus, S. mutans, P. aeruginosa and S. typhimurium in absence and presence [(1/2) × MIC, 1 × MIC and 2 × MIC] of WS‐AgNPs.* represents significance at p ≤ 0.05 with respect to control; # represents significance at p ≤ 0.01 with respect to control

3.12 Effect of WS‐AgNPs on intracellular ROS production

The relative amount of ROS generated intracellularly by WS‐AgNPs has been evaluated using the dye, DCFH‐DA which is fluorescent ROS indicator [62]. Fig. 7 b shows ROS generation in absence and presence of WS‐AgNPs. Under non‐stress conditions, ROS produced are counter balanced by ROS scavenging enzymes present in bacteria cells. However, the exposure of WS‐AgNPs had a remarkable effect on intracellular ROS. In the presence of (1/2) × MIC, 1 × MIC and 2 × MIC of WS‐AgNPs, P aeruginosa exhibited 19.98, 51.65 and 112.62% increase in ROS level as compared to untreated control cells. Likewise, at sub‐MIC concentration ((1/2) × MIC), cells of S. aureus showed 02.27% (insignificant as compared to control) that elevated to 27.73 and 54.72% at 1 × MIC and 2 × MIC, respectively. ROS level of S. typhimurium also enhanced by 39.78 and 52.01% in presence of 1 × MIC and 2 × MIC of WS‐AgNPs as compared to untreated cells. Similarly, amount of ROS in S. mutans also increased to 09.67, 23.32 and 52.01 by the supplementation of (1/2) × MIC, 1 × MIC and 2 × MIC of WS‐AgNPs to the cells. The differences in ROS production can be linked to the difference in magnitude of change in interfacial potential of different bacteria, both the gram‐positive and gram‐negative types of bacteria [63]. The neutralisation of surface potential in bacteria has been documented for induction of ROS production that found to be responsible for DNA, protein and lipid damage and finally causing the loss in viability of cells [64, 65].

4 Conclusion

The green synthesised WS‐AgNPs in size range 40–60 nm exhibited broad‐spectrum antibacterial activity against gram‐positive and gram‐negative test bacterial pathogens through interference on multiple cellular functions mainly inhibition of growth, cell membrane damage, ROS generation and biofilm inhibition. Further studies on in vivo efficacy in suitable infection model and safety issue have to be investigated to exploit the combatting potential in problematic bacterial infection.