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Antimicrobial activity of highly stable silver nanoparticles embedded in agar–agar matrix as a thin film
Carbohydrate Research 345 (2010) 2220–2227
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Antimicrobial activity of highly stable silver nanoparticles embedded in agar–agar matrix as a thin film S. Ghosh a, R. Kaushik a, K. Nagalakshmi b, S. L. Hoti b, G. A. Menezes c, B. N. Harish c, H. N. Vasan a,* a
Solid State and Structural Chemistry Unit, Indian Institute of Science, Bangalore 560 012, India Vector Control Research Centre, Medical Complex, Indra agar, Puducherry 605 006, India c Department of Microbiology, Jawaharlal Institute of Postgraduate Medical Education and Research (JIPMER), Puducherry 605 006, India b
a r t i c l e
i n f o
Article history: Received 1 May 2010 Received in revised form 27 July 2010 Accepted 4 August 2010 Available online 9 August 2010 Keywords: Silver nanoparticles Surface plasmon resonance Agar–agar Thin film Antimicrobial activity
a b s t r a c t Highly stable silver nanoparticles (Ag NPs) in agar–agar (Ag/agar) as inorganic–organic hybrid were obtained as free-standing film by in situ reduction of silver nitrate by ethanol. The antimicrobial activity of Ag/agar film on Escherichia coli (E. coli), Staphylococcus aureus (S. aureus), and Candida albicans (C. albicans) was evaluated in a nutrient broth and also in saline solution. In particular, films were repeatedly tested for antimicrobial activity after recycling. UV–vis absorption and TEM studies were carried out on films at different stages and morphological studies on microbes were carried out by SEM. Results showed spherical Ag NPs of size 15–25 nm, having sharp surface plasmon resonance (SPR) band. The antimicrobial activity of Ag/agar film was found to be in the order, C. albicans > E. coli > S. aureus, and antimicrobial activity against C. albicans was almost maintained even after the third cycle. Whereas, in case of E. coli and S. aureus there was a sharp decline in antimicrobial activity after the second cycle. Agglomeration of Ag NPs in Ag/agar film on exposure to microbes was observed by TEM studies. Cytotoxic experiments carried out on HeLa cells showed a threshold Ag NPs concentration of 60 lg/mL, much higher than the minimum inhibition concentration of Ag NPs (25.8 lg/mL) for E. coli. The mechanical strength of the film determined by nanoindentation technique showed almost retention of the strength even after repeated cycle. Ó 2010 Elsevier Ltd. All rights reserved.
1. Introduction By manipulating materials at the atomic level, nanotechnology offers to achieve unique properties for various desired applications in biology.1 Most of the nature’s creation occurs at the nanoscale regime, thus fusion between nanotechnology and biology can mimic nature and bring about a revolution in the field of health and medicine, for example, in drug delivery,2 cancer therapy,3 and medical diagnostic kits.4 The growth of new resistant strains of bacteria causing multi drug resistance (MDR) due to repeated use of bactericides is a serious issue in public health. Therefore, there is a pressing need for alternative bactericides particularly inorganic-based materials to prevent bacteria from developing MDR.5 The inorganic antibacterial agents have advantage over the organic antimicrobial agents in terms of their stability, toxicity, preparation methods, and so on.6 Depending upon the membrane structure, bacteria are of two types; Gram positive and Gram negative. The structural difference lies in the organization of a key component of the membrane, peptidoglycan. Gram-negative bacteria exhibit only a thin layer of pep* Corresponding author. Tel.: +91 80 22933310; fax: +91 80 23601310. E-mail address: [email protected] (H.N. Vasan). 0008-6215/$ - see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.carres.2010.08.001
tidoglycan (2–3 nm) between the cytoplasmic membrane and the outer membrane.7 In contrast, Gram-positive bacteria lack the outer membrane but have a peptidoglycan layer about 30 nm thick.8 Fungi come under the eukaryotic organisms like humans and their cell structures are well organized, comprising a series of single cells. The cell wall of fungus is composed of chitin, the same carbohydrate that gives strength to the exoskeletons of insects.9 The antimicrobial properties of metallic nanoparticles10–14 and metal-oxide powders15–19 are promising for biotechnological applications, such as food packaging,20 water purification,13 and as sterile coatings for biomedical devices.21 Among the various metals, for a long time silver and silver ions have been known to have strong inhibitory and bactericidal effect as well as broad spectrum of antimicrobial activity.22–24 The germicidal effect of silver ions on microorganisms is very well known but the bactericidal mechanism of both Ag+ and Ag0 is still not fully understood.23,25 Nanoparticles have an advantage over particles of micron size and above, because of their high surface to volume ratio, thus providing a large number of particles at the surface and also having large surface area. This facilitates better interaction between the particles and microbes. The antibacterial activity of Ag NPs is dependent not only on particle size26 but also on their shape.27 Hence, the synthesis of mono disperse Ag NPs having the same shape and also their stability is
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the key factor for carrying out the antibacterial activity.26,28 To achieve this, various capping agents,27 surfactants,29 and polymers30,31 are commonly used. Most of the antibacterial effects of Ag NPs have been studied by spreading homogeneously the Ag NPs into broth containing microbes.24,32 However, the Ag NPs once used cannot be recycled for repetitive use. One way to recycle the Ag NPs is to immobilize the Ag NPs in a matrix, while using it as an antimicrobial agent. The immobilization also gives stability to the NPs. Various methods are available for immobilizing Ag NPs; for example, Furno et al.10 have studied the impregnation of Ag NPs into silicone polymers using super critical carbon dioxide and evaluated their antibacterial activity. In other studies, Balogh et al.33 have studied the antibacterial activity of Ag-poly (amidoamine) (PAMAM) dendrimers nanocomposite. Jeon et al.34 have studied the antibacterial effect of Ag in Ag–SiO2 thin films, while Su et al.35 have studied the antibacterial activity of Ag nanoparticles in silicate clay as nanohybrids. Very recently, Dong et al.36 have studied the synergic antibacterial effect of impregnating Ag nanoparticles into chitosan films. However, no reports are available with repetitive use of such hybrids/composites for antibacterial studies. Here, we report a simple method of synthesis of highly stable Ag NPs (15–25 nm) in agar–agar matrix as an inorganic–organic hybrid in the form of a free-standing film. The film is found to be stable for more than 18 months without any change in dispersion or morphology of the Ag NPs. The antimicrobial studies of the film are being carried out against Candida albicans (a lower fungi), Escherichia coli (Gram-negative), and Staphylococcus aureus (Gram-positive) both in nutrient broth and also in saline water. The films are also tested for their antimicrobial property on repeated cycling and found to be encouraging with respect to C. albicans. Cytotoxic experiments against human HeLa cells showed that the threshold concentration of Ag NPs in Ag/agar is much higher than the MIC of Ag NPs for E. coli. The mechanical strength of the film determined shows almost retention of the strength even on repeated cycle. 2. Experimental 2.1. Materials Silver nitrate (99.9%), absolute alcohol (99.9%) (S.D Fine-Chem Pvt. Ltd, India), Polyvinyl pyrrolidone (PVP) (Mw 55,000, Aldrich Chemical Co. Inc.), and agar–agar (Bacterial grade type-I, Hi-Media Laboratories Ltd, India) were used as such without any further purification for the preparation of Ag NPs. For the purpose of antimicrobial assay of Ag/agar film, nutrient broth obtained from Hi-Media Laboratories Ltd, India, was used. Standard reference strains, E. coli MTCC 1302, S. aureus ATCC 25923 (supplied by M.S. Ramaiah Hospital, Bangalore), and C. albicans ATCC 90028 (supplied by Jawaharlal Institute of Postgraduate Medical Education and Research, Puducherry), were used. 2.2. Synthesis of silver nanoparticles in agar–agar matrix In situ Ag NPs of 15–25 nm were synthesized in an agar–agar matrix according to the procedure reported by us earlier.37 In a typical reaction, 1.0 g of chloride free agar–agar was dissolved in 100 mL of Millipore water under boiling conditions with stirring. To this clear solution, 60 mL of ethanol and 0.1 g of PVP were added after lowering the temperature to 50 °C with constant stirring. 1.0 mmol (0.170 g) of AgNO3 was dissolved in 20 mL Millipore water and added dropwise to the above solution and heated at reflux for 2 h at 90 °C in a rotary evaporator. The solution turned to golden yellow ensuring complete reduction of Ag+ ions to Ag0. The excess solvent and water were pumped out until the solution became viscous; 20 mL of this solution was poured into plastic
Figure 1. UV–vis spectra of Ag/agar film at three different concentrations: (a) 762 lg/cm2, (b) 487 lg/cm2, and (c) 141 lg/cm2. Inset shows TEM micrograph of Ag NPs at concentration 762 lg/cm2.
Petri dish (diameter 90 mm) and dried at room temperature until the mass could be peeled off as a transparent, free-standing film. The loading of Ag in the film may be varied by varying the initial concentration of AgNO3. To test the effectiveness of Ag/agar film as antimicrobial agent on repeated cycle in saline water, 200 lL of 0.2 mmol Ag/Agar of original solution was drop-casted over a glass slide to form a film of area 1.0 cm2, having 11.7 lg of Ag NPs (ICP analysis) and used after air drying. 2.3. UV–vis absorption study UV–vis absorption studies were carried out during the reduction of Ag+ to Ag NPs and also at various stages of treatment of Ag/Agar film with microbes. Typically a 1 cm 1 cm Ag/agar film was dissolved in 5 mL of Millipore water and scanned over the wavelength range 300–800 nm in Perkin–Elmer Lambda 35 UV– vis spectrophotometer using a quartz cuvette with Millipore water as background. 2.4. Estimation of Ag concentration The concentration of Ag NPs in Ag/agar films was estimated by ICP emission spectroscopy. Known area of Ag/agar free-standing film or on glass slide was dissolved in dilute HNO3 and made up to mark in a 10 mL volumetric flask and estimated for Ag using Perkin–Elmer optima 2100 Inductively Coupled Plasma-Optical Emission (ICP-OES).
Table 1 Viable cell counts of the microbes Set up
Initial concentration Of Ag NPs (lg/cm2)
E. coli
S. aureus
C. albicans
(i) Control (iii) PVP + ethanol/agar (iv) PVP/agar–agar (v) Ethanol/agar–agar (ii) Ag/agar–agar
0 0 0 0 116 141 258 478 522 762
TNTC TNTC TNTC TNTC TNTC 50 0 0 0 0
TNTC TNTC TNTC TNTC TNTC TNTC 174 93 15 0
TNTC TNTC TNTC TNTC 29 0 0 0 0 0
Note: The values are the average of three repeated experiments. TNTC denotes too numerous to count (when the number of colonies are P1000).
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Figure 2. Photographs showing: (a1–c1) the nutrient agar plates of positive control and (a2–c2) the corresponding MIC plates for S. aureus, E. coli, and C. albicans, respectively.
2.5. Antimicrobial study Free-standing Ag/agar films of thickness around 20 lm and 1 cm 1 cm in size, containing different concentrations of Ag NPs, were cut and washed several times with Millipore water to remove any unreduced Ag+ ions and further dried before using for the antimicrobial studies. The desired bacteria and fungi were cultured in nutrient broth at 37 °C for overnight in a mechanical shaker. The cultured microbial concentration was further reduced to 108 CFU/mL and 106 CFU/mL for bacteria and fungi, respectively, by serial dilution technique. The test samples were prepared by taking 9.9 mL of nutrient broth/saline water with 0.1 mL of inoculums. To ensure that the antimicrobial studies reflect solely by the Ag NPs embedded in film, the following five samples were prepared in separate test tubes: (i) only the test sample (positive control), and test sample containing, (ii) Ag/agar film, (iii) agar–agar film containing 0.1 g PVP and ethanol, (iv) agar–agar film containing only 0.1 g PVP, and (v) agar–agar film containing only ethanol, and all the setups were incubated at 37 °C for 4 h in a shaker.
plate technique along with positive-control plate. Then the plates were incubated at 37 °C overnight and the colonies were counted to determine the MIC. 2.7. Evaluation of zone of Inhibition (ZOI) The bacterial lawn was prepared on sterile Muller–Hinton agar plates by using a sterile cotton swab. Then 0.5 cm diameter Ag/agar film of different concentrations were placed at the center of the lawn carefully without touching other parts and incubated at
2.6. Determination of minimum inhibition concentration (MIC) The MIC, the minimum concentration at which a material exhibits antimicrobial activity, was determined by taking eleven sterile test tubes separately with 9.9 mL of nutrient broth, and inoculated with the corresponding microorganism and diluted up to 108 CFU/mL for bacteria and 106 CFU/mL for fungi. To this 1 cm 1 cm Ag/agar film containing different concentrations of Ag NPs (116–762 lg/cm2) were taken and placed in individual test tubes. After 4 h of incubation at 37 °C, 0.1 mL each of this solution was taken and plated in sterile nutrient agar plates for bacteria and sterile potato dextrose agar (PDA) plates for fungi by using spread-
Figure 3. UV–vis spectra of silver film before and after treatment with microbes: (a) spr band of Ag/agar before contact with bacteria and fungi (b–d) the corresponding spectra after 1st cycle (e–g) after 2nd cycle with E. coli, S. aureus, and C. albicans, respectively.
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37 °C for 24 h and the zone around the film was measured on the subsequent day using a centimeter scale. 2.8. Transmission electron microscopic (TEM) study The size and morphology of silver nanoparticles before and after treating with microbes were analyzed by TEM using TECNAI F 30 transmission electron microscope. Care was taken to prepare and mount the samples on Cu grid each time to avoid any possible agglomeration of particles while taking films in water. All samples were prepared by similar condition, by placing a drop of well-sonicated 1 cm 1 cm Ag/agar film dissolved in 10 mL of Millipore water on carbon-coated copper grid and subsequently dried in air before transferring it to the electron microscope, which was operated at an accelerated voltage of 120 kV. 2.9. Scanning electron microscopic (SEM) study The morphological studies of microbes on treatment with Ag NPs were carried out using FE-SEM QUANTA field emission scanning electron microscope (SEM). Briefly, 10 lL of Ag/agar solution was spread over a glass cover slip and 1 mm diameter fresh colony of microbe was emulsified on it and incubated for 24 h at 37 °C. This is then coated with a very thin layer of gold by sputtering technique. The coated sample was then subjected to SEM studies. 2.10. Cytotoxicity assay In vitro cytotoxicity assay for the Ag nanoparticles in agar (Ag/ agar) was carried out by exposing human cervical cancer cell line
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(HeLa cells) in microtiter plates to various dilutions of Ag/agar aqueous solution as described in the literature.38 Various concentrations of Ag/agar aqueous solution, starting from 6 lg/mL to 240 lg/mL, were pipetted into 96-well microtiter plates and then 0.1 mL freshly trypsinized HeLa cells cultured as monolayer in Dulbecco’s Modified Eagle Medium (DMEM) (containing 10% FCS and antibiotic solution) to active growth phase (18,000 cell per well) were added and incubated for 20 h at 35 °C in 5% CO2 atmosphere. Wells containing equal number of cells, without Ag/agar, were left as controls. After incubation, the detached cells, medium, and Ag/agar were removed by vigorous shaking. The remaining live cells were fixed in 2% solution of formalin in 0.067 M phosphate-buffered saline (PBS) (pH 7.2) for 1 min. The plates were stained with 0.13% crystal violet in 5% ethanol–2% formalin-PBS for 20 min. Excess stain was removed through rinsing with water, and the plates were air dried. For quantification, the stain was eluted from the wells with four successive 50 lL samples of 50% ethanol and diluted in 0.9 mL of PBS. Absorbance of the pooled washings was recorded at 595 nm in a spectrophotometer. Percentage of cells surviving the treatment was computed for various doses of Ag/agar by considering the optical density (OD595) for control as hundred percent. 2.11. Nanoindentation Nanoindentation experiments were carried out on pure agar– agar film and after each cycle the film (10 mm 10 mm area) of 20 lm thickness coated on a glass slide, was dipped in a saline solution containing E. coli to determine the mechanical properties. A Berkovich tip (a three-sided pyramidal diamond tip) of diameter
Figure 4. TEM micrographs of silver nps: (a) before treatment with microbes (b1–b3) after 1st treatment with E. coli, S. aureus, and C. albicans at their corresponding MIC concentration. (Note: The magnifications are different; all the scale bars are equivalent to 20 nm.)
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100 nm was used for carrying out the nanoindentation experiments in Hysitron Triboindenter (Minneapolis, USA). Since the mechanical properties extracted from nanoindentation technique are sensitive to the tip geometry, the tip area function has to be calibrated accurately. This was done by using a standard quartz sample and by following a standard practice. Because the loads used in indenting agar–agar films are very small, the tip area function is calibrated in low depth ranges for precise determination of the modulus and hardness. Using this area function, nanoindentation experiments were performed on single-crystal Al to cross check the standard elastic modulus, E, and hardness, H, values as prescribed by the manufacturer (75.1 ± 5% GPa and 360 ± 10% MPa, respectively). The standard deviation is within 5% for both these values, validating the tip calibration process. A peak load of 400 lN was applied in all the cases, with loading and unloading rates of 40 lN/s. A pause time of 10 s was allowed at the peak load
to find out time-dependent plastic deformation, if any. Load-penetration depth curves of the Ag/Agar over glass slide after each cycle of use are shown in Figure 2S as Supplementary data. Ten indentations were made for each sample. 3. Results and discussion 3.1. Synthesis and characterization of Ag/agar film One of our group’s interests is to develop an affordable and effective inorganic-based material such as silver and zinc oxide as antimicrobial agents for the purification of water, which can also be recycled for repetitive use. As mentioned earlier many researchers have tried to develop various composites containing Ag NPs for the antimicrobial activity.10,31–33 However, in most of the cases, materials and process involved are either expensive or time con-
Figure 5. SEM micrographs of microbes: (a1–a3) before and (b1–b3) after treatment with corresponding MIC concentrations for E. coli, S. aureus, and C. albicans respectively.
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suming. Silver, being highly electropositive, is easy to reduce to zero valence from its Ag+ oxidation state. Various reducing agents such as borohydrides,39 aldehydes,40 sugars,41 and alcohols,42 including polyols,43 have been used to reduce Ag+ to Ag0. As we were interested in carrying out the in situ reduction of Ag+ in the presence of agar–agar in an aqueous medium containing a milder reducing agent, ethyl alcohol was chosen. Ethyl alcohol also has the advantage of mixing with water in all proportions, forming alcohol–water mixture. Agar–agar is biodegradable polysaccharide, and it gels easily and can be easily casted to form films. The Ag NPs are homogeneously distributed in an agar–agar matrix to form an organic–inorganic hybrid, and it can be obtained in various forms such as in solution, gel, and as a free-standing film. The thickness of the film can be varied by varying the initial volume of Ag/agar solution poured into a Petri dish. Different concentrations of Ag NPs can be loaded in situ into agar–agar by varying the initial concentration of AgNO3. The formation of Ag NPs is easily characterized by UV–vis spectra as shown in Figure 1. The sharp plasmon resonance (SPR) band around 410 nm indicates the presence of spherical Ag NPs and the band intensity increases as the concentration of Ag NPs increases. The TEM micrograph also shows more or less the spherical shape of Ag NPs with size in the range 15–25 nm (inset, Fig. 1). The SPR band of Ag NPs in UV–vis spectra and the photograph of the film are shown as an inset (Supplementary data, Figure S1), taken after storing the free-standing film for more than 18 months in a desiccator. The intensity and the FWHM of the peak remained almost the same after this period, indicating the long stability of Ag/agar film. 3.2. Antimicrobial activity Invariably the antimicrobial studies of Ag NPs are carried out in a homogeneous medium,24,32 where Ag NPs are evenly distributed in bacterial culture, however, the Ag NPs cannot be recycled. Whereas the Ag/agar film prepared by the procedure described above can be repeatedly cycled on washing and tested for its antimicrobial activity. The MIC determinations of Ag NPs in Ag/agar films are carried out in nutrient broth and also in saline water against C. albicans, E. coli, and S. aureus. Table 1 shows the viable cell counts of both bacteria and fungi in nutrient broth at dilution levels of 108 CFU/mL and 106 CFU/mL, respectively, after incubation at 37 °C for 24 h on different silver concentrations in agar– agar. It is quite evident from the results that with increasing silver concentration there is a decrease in microbial growth. The antimicrobial effect of silver is more effective and is in the order of C. albicans > E. coli > S. aureus. The corresponding MIC of Ag NPs where 100% reduction of microbial growth is seen are found to be 14.1 lg/mL, 25.8 lg/mL, and 76.2 lg/mL (in the film, this corresponds to 141 lg/cm2, 258 lg/cm2, and 762 lg/cm2, respectively) in nutrient broth solution. The photographs of nutrient agar plates of positive control and the corresponding MIC plates for E. coli, S. aureus, and C. albicans, respectively, are depicted in Figure 2. The number of microbial colonies grown on PVP + ethanol/agar film (setup iii), PVP/ agar film (setup iv), and ethanol/agar film (setup v) for E. coli, S. aureus, and C. albicans, respectively, is found to be more than 1000 (photographs not shown), thus indicating that other chemicals such as PVP and ethanol used initially while preparing the Ag NPs have no effect on the antimicrobial activity. After 4 h contact with microbial solution during incubation at 37 °C, each of the Ag/agar films is taken out and washed thoroughly with Millipore water several times for recycling and tested for antimicrobial activity. We presume that Ag NPs have diffused into the broth, due to the concentration gradient of Ag, thus attacking the cell walls, rather than the microbes coming in contact with Ag/agar. This is evident
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Figure 6. UV–vis spectra showing SPR bands of Ag NPs on glass plate for E. coli on repeated cycle. Inset table showing amount of silver present after each cycle.
from the UV–vis spectra (Fig. 3), where the SPR intensity of Ag NPs has decreased and broadened on subsequent cycle indicating the decrease in concentration and the possible shape change of Ag NPs in the matrix. However, it is not clear whether the antibacterial activity is due to Ag0 or Ag+ ions once Ag NPs have diffused into the broth solution. Before contact with microbe, spherical silver NPs (size = 15–25 nm) were almost evenly distributed throughout the agar–agar matrix, but after the first contact with microbe, the particles are found to be agglomerated with wide distribution in size and also exhibiting irregularity in shape as seen in TEM micrograph (Fig. 4). Thus the agglomeration and depletion of Ag NPs reduce the antimicrobial property of Ag/agar film on cycling. The bacterial death after Ag attack was observed by SEM micrographs (Fig. 5). On close observation, one can infer that the nature of attack on microbes studied is different, even though it is the cell walls that are destroyed. In case of E. coli the original rod shape has swollen into bigger size, with high agglomeration of the cells. However, in S. aureus, the spherical-shaped cell has broken down to smaller size with wide distribution. In C. albicans, but for the agglomeration
Figure 7. Antimicrobial activity versus number of cycles of Ag/agar: S. aureus (circle), E. coli (square), and C. albicans (triangle).
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Figure 8. Zone of inhibition (ZOI): (a) zone radius (cm) vs different concentrations of Ag NPs in Ag/agar (b) Optical images of ZOI for E. coli as concentration of Ag NPs increased in Ag/agar (anticlockwise).
the cell size and shape are not much affected. It is presumed that the death of microbial cells may be due to damage of the microbial enzymes or cell membrane. Considering the possible application of Ag/agar for the purification of water free from microbes, a separate sample preparation was carried out at a lower concentration of Ag. The film is casted onto a glass slide to ensure mechanical stability and tested for MIC of Ag NPs in saline water (0.85% NaCl), which represents closer to bacterial-contaminated water.18 The corresponding MIC determined for C. albicans, E. coli, and S. aureus are 0.914 lg/mL, 0.914 lg/mL, and 2.55 lg/mL (this corresponds to 9.14 lg/cm2, 9.14 lg/cm2, and 25.5 lg/cm2 of film, respectively). These values are correspondingly very low compared to the MIC in nutrient media. This is because the growth of microbes in nutrient media is many times higher (exponential rate) than in saline water. For antibacterial activity on recycling, the glass-coated films are washed in Milli-Q water and dried and dipped into saline water containing the corresponding microbes and tested for four cycles. UV–vis spectra of the slide were taken before and after each contact with microbes. Representative UV–vis spectra for E. coli are shown in Figure 6. The intensity of the SPR band decreased with broadening after each contact indicating loss of Ag NPs from the
film, which in turn decreases the antimicrobial activity, similar to as observed in nutrient broth. The inset of Figure 6 shows the Ag concentration in the film estimated from ICP analysis after each cycle. The decrease in Ag concentration on subsequent cycling further confirms the diffusion of Ag NPs into the saline solution. Arbitrarily, three times the concentration of MIC of S. aureus in saline solution was taken on a glass slide as before and cycled to determine the percentage survival of microbes and the results are shown in Figure 7. It is seen that the Ag/agar is more effective upto the third cycle in the case of C. albicans compared to E. coli and S. aureus, and there is a steep rise in the percent survival after the second cycle in the latter two. From the Kirby–Bauer test, it is found that as the concentration of Ag NPs increased, the zone radius also increased for all the microbes tested and they follow a linear plot. The zone radius for C. albicans is higher at a particular concentration of silver film (Fig. 8a) and the line is slightly steeper, compared to the other two microbes studied. A representative photograph of ZOI for E. coli in Figure 8b shows the increase in zone radius as Ag concentration is increased (anti clockwise). 3.3. Cytotoxicity experiments Upon conducting cytotoxicity experiments, the percentage survival of HeLa cells was plotted against the concentration of Ag NPs in Ag/agar and is shown in Figure 9. From the plot it can be seen that the Ag NPs exhibit a very low level of toxicity (72 lg/mL). Beyond this concentration of Ag NPs a drastic reduction of viability of the cells is seen. However, it may be noted that 9 lg/mL was the MIC found against the water-borne coliform bacterium, E. coli, and this concentration is almost seven times less than the threshold concentration of 60 lg/mL as seen in the plot.
Table 2 Mechanical properties of the films
Figure 9. Bar chart showing percentage survival of HeLa cells against concentrations of Ag NPs in Ag/agar.
Cycles
Elastic modulus (GPa)
Hardness (MPa)
Blank agar–agar 0th 1st 2nd 3rd 4th
1.229 ± 0.028 4.498 ± 0.117 4.139 ± 0.097 3.734 ± 0.162 3.804 ± 0.331 3.462 ± 0.057
36.6 ± 0.001 272.8 ± 0.009 268.2 ± 0.008 256.9 ± 0.133 241.8 ± 0.023 239.3 ± 0.008
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Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.carres.2010.08.001. References
Figure 10. Normalized hardness and elastic modulus of the film plotted as a function of number of cycles.
3.4. Mechanical properties The results of the elastic modulus (E) and hardness (H) of bare agar–agar and Ag/agar film after each cycle, calculated using Oliver–Pharr44 method, are listed in Table 2. These properties normalized with respect to Ag/agar film at zero cycle are plotted against the number of cycles (Fig. 10). The elasticity of Ag/agar film has increased almost four fold compared to the pure agar–agar film. Similarly the hardness has increased by seven times. There is only a decrease of hardness around 5% after second cycle and a decrease of around 10% on subsequent cycles, indicating good stability of the film. As a comparison, for example, Ag/agar films have mechanical strength more than fifty times compared to PVA-(polyvinyl alcohol)reinforced graphine films.45 4. Conclusions We have described a simple biocompatible, in situ synthesis of highly stable (more than 18 months) Ag nano particles in agar– agar matrix as free-standing film, using alcohol as reducing agent. Films are characterized by UV–vis absorption and TEM studies. The antimicrobial activity of the film is found to be in the order of C. albicans > E. coli > S. aureus. For the first time we have demonstrated the antimicrobial activity of the film on recycling in a saline solution. The activity reduces on cycling, mainly because of depletion of Ag NPs in the film on repeated use. Electron microscopic studies show the changes in morphology of microbes on the effect of Ag NPs and in turn the agglomeration of Ag NPs on exposure to microbes. Cytotoxicity experiments show a reasonably low toxic limit of Ag NPs. The Ag/agar film exhibits good mechanical stability. It is further worth studying to understand the exact mechanism of antimicrobial activity of Ag/agar film. Acknowledgments The authors thank the Nano Centre (IISc) for electron microscope facilities. The authors are indebted to Professor U. Ramamurty and Dr. S.R.N. Kiran Mangalampalli of the Department of Materials Engineering (IISc) for the timely help in the measurement of mechanical properties. H.N.V. thanks the generous internal funding from IISc, Bangalore.
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Carbohydrate Research journal homepage: www.elsevier.com/locate/carres
Antimicrobial activity of highly stable silver nanoparticles embedded in agar–agar matrix as a thin film S. Ghosh a, R. Kaushik a, K. Nagalakshmi b, S. L. Hoti b, G. A. Menezes c, B. N. Harish c, H. N. Vasan a,* a
Solid State and Structural Chemistry Unit, Indian Institute of Science, Bangalore 560 012, India Vector Control Research Centre, Medical Complex, Indra agar, Puducherry 605 006, India c Department of Microbiology, Jawaharlal Institute of Postgraduate Medical Education and Research (JIPMER), Puducherry 605 006, India b
a r t i c l e
i n f o
Article history: Received 1 May 2010 Received in revised form 27 July 2010 Accepted 4 August 2010 Available online 9 August 2010 Keywords: Silver nanoparticles Surface plasmon resonance Agar–agar Thin film Antimicrobial activity
a b s t r a c t Highly stable silver nanoparticles (Ag NPs) in agar–agar (Ag/agar) as inorganic–organic hybrid were obtained as free-standing film by in situ reduction of silver nitrate by ethanol. The antimicrobial activity of Ag/agar film on Escherichia coli (E. coli), Staphylococcus aureus (S. aureus), and Candida albicans (C. albicans) was evaluated in a nutrient broth and also in saline solution. In particular, films were repeatedly tested for antimicrobial activity after recycling. UV–vis absorption and TEM studies were carried out on films at different stages and morphological studies on microbes were carried out by SEM. Results showed spherical Ag NPs of size 15–25 nm, having sharp surface plasmon resonance (SPR) band. The antimicrobial activity of Ag/agar film was found to be in the order, C. albicans > E. coli > S. aureus, and antimicrobial activity against C. albicans was almost maintained even after the third cycle. Whereas, in case of E. coli and S. aureus there was a sharp decline in antimicrobial activity after the second cycle. Agglomeration of Ag NPs in Ag/agar film on exposure to microbes was observed by TEM studies. Cytotoxic experiments carried out on HeLa cells showed a threshold Ag NPs concentration of 60 lg/mL, much higher than the minimum inhibition concentration of Ag NPs (25.8 lg/mL) for E. coli. The mechanical strength of the film determined by nanoindentation technique showed almost retention of the strength even after repeated cycle. Ó 2010 Elsevier Ltd. All rights reserved.
1. Introduction By manipulating materials at the atomic level, nanotechnology offers to achieve unique properties for various desired applications in biology.1 Most of the nature’s creation occurs at the nanoscale regime, thus fusion between nanotechnology and biology can mimic nature and bring about a revolution in the field of health and medicine, for example, in drug delivery,2 cancer therapy,3 and medical diagnostic kits.4 The growth of new resistant strains of bacteria causing multi drug resistance (MDR) due to repeated use of bactericides is a serious issue in public health. Therefore, there is a pressing need for alternative bactericides particularly inorganic-based materials to prevent bacteria from developing MDR.5 The inorganic antibacterial agents have advantage over the organic antimicrobial agents in terms of their stability, toxicity, preparation methods, and so on.6 Depending upon the membrane structure, bacteria are of two types; Gram positive and Gram negative. The structural difference lies in the organization of a key component of the membrane, peptidoglycan. Gram-negative bacteria exhibit only a thin layer of pep* Corresponding author. Tel.: +91 80 22933310; fax: +91 80 23601310. E-mail address: [email protected] (H.N. Vasan). 0008-6215/$ - see front matter Ó 2010 Elsevier Ltd. All rights reserved. doi:10.1016/j.carres.2010.08.001
tidoglycan (2–3 nm) between the cytoplasmic membrane and the outer membrane.7 In contrast, Gram-positive bacteria lack the outer membrane but have a peptidoglycan layer about 30 nm thick.8 Fungi come under the eukaryotic organisms like humans and their cell structures are well organized, comprising a series of single cells. The cell wall of fungus is composed of chitin, the same carbohydrate that gives strength to the exoskeletons of insects.9 The antimicrobial properties of metallic nanoparticles10–14 and metal-oxide powders15–19 are promising for biotechnological applications, such as food packaging,20 water purification,13 and as sterile coatings for biomedical devices.21 Among the various metals, for a long time silver and silver ions have been known to have strong inhibitory and bactericidal effect as well as broad spectrum of antimicrobial activity.22–24 The germicidal effect of silver ions on microorganisms is very well known but the bactericidal mechanism of both Ag+ and Ag0 is still not fully understood.23,25 Nanoparticles have an advantage over particles of micron size and above, because of their high surface to volume ratio, thus providing a large number of particles at the surface and also having large surface area. This facilitates better interaction between the particles and microbes. The antibacterial activity of Ag NPs is dependent not only on particle size26 but also on their shape.27 Hence, the synthesis of mono disperse Ag NPs having the same shape and also their stability is
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the key factor for carrying out the antibacterial activity.26,28 To achieve this, various capping agents,27 surfactants,29 and polymers30,31 are commonly used. Most of the antibacterial effects of Ag NPs have been studied by spreading homogeneously the Ag NPs into broth containing microbes.24,32 However, the Ag NPs once used cannot be recycled for repetitive use. One way to recycle the Ag NPs is to immobilize the Ag NPs in a matrix, while using it as an antimicrobial agent. The immobilization also gives stability to the NPs. Various methods are available for immobilizing Ag NPs; for example, Furno et al.10 have studied the impregnation of Ag NPs into silicone polymers using super critical carbon dioxide and evaluated their antibacterial activity. In other studies, Balogh et al.33 have studied the antibacterial activity of Ag-poly (amidoamine) (PAMAM) dendrimers nanocomposite. Jeon et al.34 have studied the antibacterial effect of Ag in Ag–SiO2 thin films, while Su et al.35 have studied the antibacterial activity of Ag nanoparticles in silicate clay as nanohybrids. Very recently, Dong et al.36 have studied the synergic antibacterial effect of impregnating Ag nanoparticles into chitosan films. However, no reports are available with repetitive use of such hybrids/composites for antibacterial studies. Here, we report a simple method of synthesis of highly stable Ag NPs (15–25 nm) in agar–agar matrix as an inorganic–organic hybrid in the form of a free-standing film. The film is found to be stable for more than 18 months without any change in dispersion or morphology of the Ag NPs. The antimicrobial studies of the film are being carried out against Candida albicans (a lower fungi), Escherichia coli (Gram-negative), and Staphylococcus aureus (Gram-positive) both in nutrient broth and also in saline water. The films are also tested for their antimicrobial property on repeated cycling and found to be encouraging with respect to C. albicans. Cytotoxic experiments against human HeLa cells showed that the threshold concentration of Ag NPs in Ag/agar is much higher than the MIC of Ag NPs for E. coli. The mechanical strength of the film determined shows almost retention of the strength even on repeated cycle. 2. Experimental 2.1. Materials Silver nitrate (99.9%), absolute alcohol (99.9%) (S.D Fine-Chem Pvt. Ltd, India), Polyvinyl pyrrolidone (PVP) (Mw 55,000, Aldrich Chemical Co. Inc.), and agar–agar (Bacterial grade type-I, Hi-Media Laboratories Ltd, India) were used as such without any further purification for the preparation of Ag NPs. For the purpose of antimicrobial assay of Ag/agar film, nutrient broth obtained from Hi-Media Laboratories Ltd, India, was used. Standard reference strains, E. coli MTCC 1302, S. aureus ATCC 25923 (supplied by M.S. Ramaiah Hospital, Bangalore), and C. albicans ATCC 90028 (supplied by Jawaharlal Institute of Postgraduate Medical Education and Research, Puducherry), were used. 2.2. Synthesis of silver nanoparticles in agar–agar matrix In situ Ag NPs of 15–25 nm were synthesized in an agar–agar matrix according to the procedure reported by us earlier.37 In a typical reaction, 1.0 g of chloride free agar–agar was dissolved in 100 mL of Millipore water under boiling conditions with stirring. To this clear solution, 60 mL of ethanol and 0.1 g of PVP were added after lowering the temperature to 50 °C with constant stirring. 1.0 mmol (0.170 g) of AgNO3 was dissolved in 20 mL Millipore water and added dropwise to the above solution and heated at reflux for 2 h at 90 °C in a rotary evaporator. The solution turned to golden yellow ensuring complete reduction of Ag+ ions to Ag0. The excess solvent and water were pumped out until the solution became viscous; 20 mL of this solution was poured into plastic
Figure 1. UV–vis spectra of Ag/agar film at three different concentrations: (a) 762 lg/cm2, (b) 487 lg/cm2, and (c) 141 lg/cm2. Inset shows TEM micrograph of Ag NPs at concentration 762 lg/cm2.
Petri dish (diameter 90 mm) and dried at room temperature until the mass could be peeled off as a transparent, free-standing film. The loading of Ag in the film may be varied by varying the initial concentration of AgNO3. To test the effectiveness of Ag/agar film as antimicrobial agent on repeated cycle in saline water, 200 lL of 0.2 mmol Ag/Agar of original solution was drop-casted over a glass slide to form a film of area 1.0 cm2, having 11.7 lg of Ag NPs (ICP analysis) and used after air drying. 2.3. UV–vis absorption study UV–vis absorption studies were carried out during the reduction of Ag+ to Ag NPs and also at various stages of treatment of Ag/Agar film with microbes. Typically a 1 cm 1 cm Ag/agar film was dissolved in 5 mL of Millipore water and scanned over the wavelength range 300–800 nm in Perkin–Elmer Lambda 35 UV– vis spectrophotometer using a quartz cuvette with Millipore water as background. 2.4. Estimation of Ag concentration The concentration of Ag NPs in Ag/agar films was estimated by ICP emission spectroscopy. Known area of Ag/agar free-standing film or on glass slide was dissolved in dilute HNO3 and made up to mark in a 10 mL volumetric flask and estimated for Ag using Perkin–Elmer optima 2100 Inductively Coupled Plasma-Optical Emission (ICP-OES).
Table 1 Viable cell counts of the microbes Set up
Initial concentration Of Ag NPs (lg/cm2)
E. coli
S. aureus
C. albicans
(i) Control (iii) PVP + ethanol/agar (iv) PVP/agar–agar (v) Ethanol/agar–agar (ii) Ag/agar–agar
0 0 0 0 116 141 258 478 522 762
TNTC TNTC TNTC TNTC TNTC 50 0 0 0 0
TNTC TNTC TNTC TNTC TNTC TNTC 174 93 15 0
TNTC TNTC TNTC TNTC 29 0 0 0 0 0
Note: The values are the average of three repeated experiments. TNTC denotes too numerous to count (when the number of colonies are P1000).
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Figure 2. Photographs showing: (a1–c1) the nutrient agar plates of positive control and (a2–c2) the corresponding MIC plates for S. aureus, E. coli, and C. albicans, respectively.
2.5. Antimicrobial study Free-standing Ag/agar films of thickness around 20 lm and 1 cm 1 cm in size, containing different concentrations of Ag NPs, were cut and washed several times with Millipore water to remove any unreduced Ag+ ions and further dried before using for the antimicrobial studies. The desired bacteria and fungi were cultured in nutrient broth at 37 °C for overnight in a mechanical shaker. The cultured microbial concentration was further reduced to 108 CFU/mL and 106 CFU/mL for bacteria and fungi, respectively, by serial dilution technique. The test samples were prepared by taking 9.9 mL of nutrient broth/saline water with 0.1 mL of inoculums. To ensure that the antimicrobial studies reflect solely by the Ag NPs embedded in film, the following five samples were prepared in separate test tubes: (i) only the test sample (positive control), and test sample containing, (ii) Ag/agar film, (iii) agar–agar film containing 0.1 g PVP and ethanol, (iv) agar–agar film containing only 0.1 g PVP, and (v) agar–agar film containing only ethanol, and all the setups were incubated at 37 °C for 4 h in a shaker.
plate technique along with positive-control plate. Then the plates were incubated at 37 °C overnight and the colonies were counted to determine the MIC. 2.7. Evaluation of zone of Inhibition (ZOI) The bacterial lawn was prepared on sterile Muller–Hinton agar plates by using a sterile cotton swab. Then 0.5 cm diameter Ag/agar film of different concentrations were placed at the center of the lawn carefully without touching other parts and incubated at
2.6. Determination of minimum inhibition concentration (MIC) The MIC, the minimum concentration at which a material exhibits antimicrobial activity, was determined by taking eleven sterile test tubes separately with 9.9 mL of nutrient broth, and inoculated with the corresponding microorganism and diluted up to 108 CFU/mL for bacteria and 106 CFU/mL for fungi. To this 1 cm 1 cm Ag/agar film containing different concentrations of Ag NPs (116–762 lg/cm2) were taken and placed in individual test tubes. After 4 h of incubation at 37 °C, 0.1 mL each of this solution was taken and plated in sterile nutrient agar plates for bacteria and sterile potato dextrose agar (PDA) plates for fungi by using spread-
Figure 3. UV–vis spectra of silver film before and after treatment with microbes: (a) spr band of Ag/agar before contact with bacteria and fungi (b–d) the corresponding spectra after 1st cycle (e–g) after 2nd cycle with E. coli, S. aureus, and C. albicans, respectively.
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37 °C for 24 h and the zone around the film was measured on the subsequent day using a centimeter scale. 2.8. Transmission electron microscopic (TEM) study The size and morphology of silver nanoparticles before and after treating with microbes were analyzed by TEM using TECNAI F 30 transmission electron microscope. Care was taken to prepare and mount the samples on Cu grid each time to avoid any possible agglomeration of particles while taking films in water. All samples were prepared by similar condition, by placing a drop of well-sonicated 1 cm 1 cm Ag/agar film dissolved in 10 mL of Millipore water on carbon-coated copper grid and subsequently dried in air before transferring it to the electron microscope, which was operated at an accelerated voltage of 120 kV. 2.9. Scanning electron microscopic (SEM) study The morphological studies of microbes on treatment with Ag NPs were carried out using FE-SEM QUANTA field emission scanning electron microscope (SEM). Briefly, 10 lL of Ag/agar solution was spread over a glass cover slip and 1 mm diameter fresh colony of microbe was emulsified on it and incubated for 24 h at 37 °C. This is then coated with a very thin layer of gold by sputtering technique. The coated sample was then subjected to SEM studies. 2.10. Cytotoxicity assay In vitro cytotoxicity assay for the Ag nanoparticles in agar (Ag/ agar) was carried out by exposing human cervical cancer cell line
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(HeLa cells) in microtiter plates to various dilutions of Ag/agar aqueous solution as described in the literature.38 Various concentrations of Ag/agar aqueous solution, starting from 6 lg/mL to 240 lg/mL, were pipetted into 96-well microtiter plates and then 0.1 mL freshly trypsinized HeLa cells cultured as monolayer in Dulbecco’s Modified Eagle Medium (DMEM) (containing 10% FCS and antibiotic solution) to active growth phase (18,000 cell per well) were added and incubated for 20 h at 35 °C in 5% CO2 atmosphere. Wells containing equal number of cells, without Ag/agar, were left as controls. After incubation, the detached cells, medium, and Ag/agar were removed by vigorous shaking. The remaining live cells were fixed in 2% solution of formalin in 0.067 M phosphate-buffered saline (PBS) (pH 7.2) for 1 min. The plates were stained with 0.13% crystal violet in 5% ethanol–2% formalin-PBS for 20 min. Excess stain was removed through rinsing with water, and the plates were air dried. For quantification, the stain was eluted from the wells with four successive 50 lL samples of 50% ethanol and diluted in 0.9 mL of PBS. Absorbance of the pooled washings was recorded at 595 nm in a spectrophotometer. Percentage of cells surviving the treatment was computed for various doses of Ag/agar by considering the optical density (OD595) for control as hundred percent. 2.11. Nanoindentation Nanoindentation experiments were carried out on pure agar– agar film and after each cycle the film (10 mm 10 mm area) of 20 lm thickness coated on a glass slide, was dipped in a saline solution containing E. coli to determine the mechanical properties. A Berkovich tip (a three-sided pyramidal diamond tip) of diameter
Figure 4. TEM micrographs of silver nps: (a) before treatment with microbes (b1–b3) after 1st treatment with E. coli, S. aureus, and C. albicans at their corresponding MIC concentration. (Note: The magnifications are different; all the scale bars are equivalent to 20 nm.)
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100 nm was used for carrying out the nanoindentation experiments in Hysitron Triboindenter (Minneapolis, USA). Since the mechanical properties extracted from nanoindentation technique are sensitive to the tip geometry, the tip area function has to be calibrated accurately. This was done by using a standard quartz sample and by following a standard practice. Because the loads used in indenting agar–agar films are very small, the tip area function is calibrated in low depth ranges for precise determination of the modulus and hardness. Using this area function, nanoindentation experiments were performed on single-crystal Al to cross check the standard elastic modulus, E, and hardness, H, values as prescribed by the manufacturer (75.1 ± 5% GPa and 360 ± 10% MPa, respectively). The standard deviation is within 5% for both these values, validating the tip calibration process. A peak load of 400 lN was applied in all the cases, with loading and unloading rates of 40 lN/s. A pause time of 10 s was allowed at the peak load
to find out time-dependent plastic deformation, if any. Load-penetration depth curves of the Ag/Agar over glass slide after each cycle of use are shown in Figure 2S as Supplementary data. Ten indentations were made for each sample. 3. Results and discussion 3.1. Synthesis and characterization of Ag/agar film One of our group’s interests is to develop an affordable and effective inorganic-based material such as silver and zinc oxide as antimicrobial agents for the purification of water, which can also be recycled for repetitive use. As mentioned earlier many researchers have tried to develop various composites containing Ag NPs for the antimicrobial activity.10,31–33 However, in most of the cases, materials and process involved are either expensive or time con-
Figure 5. SEM micrographs of microbes: (a1–a3) before and (b1–b3) after treatment with corresponding MIC concentrations for E. coli, S. aureus, and C. albicans respectively.
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suming. Silver, being highly electropositive, is easy to reduce to zero valence from its Ag+ oxidation state. Various reducing agents such as borohydrides,39 aldehydes,40 sugars,41 and alcohols,42 including polyols,43 have been used to reduce Ag+ to Ag0. As we were interested in carrying out the in situ reduction of Ag+ in the presence of agar–agar in an aqueous medium containing a milder reducing agent, ethyl alcohol was chosen. Ethyl alcohol also has the advantage of mixing with water in all proportions, forming alcohol–water mixture. Agar–agar is biodegradable polysaccharide, and it gels easily and can be easily casted to form films. The Ag NPs are homogeneously distributed in an agar–agar matrix to form an organic–inorganic hybrid, and it can be obtained in various forms such as in solution, gel, and as a free-standing film. The thickness of the film can be varied by varying the initial volume of Ag/agar solution poured into a Petri dish. Different concentrations of Ag NPs can be loaded in situ into agar–agar by varying the initial concentration of AgNO3. The formation of Ag NPs is easily characterized by UV–vis spectra as shown in Figure 1. The sharp plasmon resonance (SPR) band around 410 nm indicates the presence of spherical Ag NPs and the band intensity increases as the concentration of Ag NPs increases. The TEM micrograph also shows more or less the spherical shape of Ag NPs with size in the range 15–25 nm (inset, Fig. 1). The SPR band of Ag NPs in UV–vis spectra and the photograph of the film are shown as an inset (Supplementary data, Figure S1), taken after storing the free-standing film for more than 18 months in a desiccator. The intensity and the FWHM of the peak remained almost the same after this period, indicating the long stability of Ag/agar film. 3.2. Antimicrobial activity Invariably the antimicrobial studies of Ag NPs are carried out in a homogeneous medium,24,32 where Ag NPs are evenly distributed in bacterial culture, however, the Ag NPs cannot be recycled. Whereas the Ag/agar film prepared by the procedure described above can be repeatedly cycled on washing and tested for its antimicrobial activity. The MIC determinations of Ag NPs in Ag/agar films are carried out in nutrient broth and also in saline water against C. albicans, E. coli, and S. aureus. Table 1 shows the viable cell counts of both bacteria and fungi in nutrient broth at dilution levels of 108 CFU/mL and 106 CFU/mL, respectively, after incubation at 37 °C for 24 h on different silver concentrations in agar– agar. It is quite evident from the results that with increasing silver concentration there is a decrease in microbial growth. The antimicrobial effect of silver is more effective and is in the order of C. albicans > E. coli > S. aureus. The corresponding MIC of Ag NPs where 100% reduction of microbial growth is seen are found to be 14.1 lg/mL, 25.8 lg/mL, and 76.2 lg/mL (in the film, this corresponds to 141 lg/cm2, 258 lg/cm2, and 762 lg/cm2, respectively) in nutrient broth solution. The photographs of nutrient agar plates of positive control and the corresponding MIC plates for E. coli, S. aureus, and C. albicans, respectively, are depicted in Figure 2. The number of microbial colonies grown on PVP + ethanol/agar film (setup iii), PVP/ agar film (setup iv), and ethanol/agar film (setup v) for E. coli, S. aureus, and C. albicans, respectively, is found to be more than 1000 (photographs not shown), thus indicating that other chemicals such as PVP and ethanol used initially while preparing the Ag NPs have no effect on the antimicrobial activity. After 4 h contact with microbial solution during incubation at 37 °C, each of the Ag/agar films is taken out and washed thoroughly with Millipore water several times for recycling and tested for antimicrobial activity. We presume that Ag NPs have diffused into the broth, due to the concentration gradient of Ag, thus attacking the cell walls, rather than the microbes coming in contact with Ag/agar. This is evident
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Figure 6. UV–vis spectra showing SPR bands of Ag NPs on glass plate for E. coli on repeated cycle. Inset table showing amount of silver present after each cycle.
from the UV–vis spectra (Fig. 3), where the SPR intensity of Ag NPs has decreased and broadened on subsequent cycle indicating the decrease in concentration and the possible shape change of Ag NPs in the matrix. However, it is not clear whether the antibacterial activity is due to Ag0 or Ag+ ions once Ag NPs have diffused into the broth solution. Before contact with microbe, spherical silver NPs (size = 15–25 nm) were almost evenly distributed throughout the agar–agar matrix, but after the first contact with microbe, the particles are found to be agglomerated with wide distribution in size and also exhibiting irregularity in shape as seen in TEM micrograph (Fig. 4). Thus the agglomeration and depletion of Ag NPs reduce the antimicrobial property of Ag/agar film on cycling. The bacterial death after Ag attack was observed by SEM micrographs (Fig. 5). On close observation, one can infer that the nature of attack on microbes studied is different, even though it is the cell walls that are destroyed. In case of E. coli the original rod shape has swollen into bigger size, with high agglomeration of the cells. However, in S. aureus, the spherical-shaped cell has broken down to smaller size with wide distribution. In C. albicans, but for the agglomeration
Figure 7. Antimicrobial activity versus number of cycles of Ag/agar: S. aureus (circle), E. coli (square), and C. albicans (triangle).
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Figure 8. Zone of inhibition (ZOI): (a) zone radius (cm) vs different concentrations of Ag NPs in Ag/agar (b) Optical images of ZOI for E. coli as concentration of Ag NPs increased in Ag/agar (anticlockwise).
the cell size and shape are not much affected. It is presumed that the death of microbial cells may be due to damage of the microbial enzymes or cell membrane. Considering the possible application of Ag/agar for the purification of water free from microbes, a separate sample preparation was carried out at a lower concentration of Ag. The film is casted onto a glass slide to ensure mechanical stability and tested for MIC of Ag NPs in saline water (0.85% NaCl), which represents closer to bacterial-contaminated water.18 The corresponding MIC determined for C. albicans, E. coli, and S. aureus are 0.914 lg/mL, 0.914 lg/mL, and 2.55 lg/mL (this corresponds to 9.14 lg/cm2, 9.14 lg/cm2, and 25.5 lg/cm2 of film, respectively). These values are correspondingly very low compared to the MIC in nutrient media. This is because the growth of microbes in nutrient media is many times higher (exponential rate) than in saline water. For antibacterial activity on recycling, the glass-coated films are washed in Milli-Q water and dried and dipped into saline water containing the corresponding microbes and tested for four cycles. UV–vis spectra of the slide were taken before and after each contact with microbes. Representative UV–vis spectra for E. coli are shown in Figure 6. The intensity of the SPR band decreased with broadening after each contact indicating loss of Ag NPs from the
film, which in turn decreases the antimicrobial activity, similar to as observed in nutrient broth. The inset of Figure 6 shows the Ag concentration in the film estimated from ICP analysis after each cycle. The decrease in Ag concentration on subsequent cycling further confirms the diffusion of Ag NPs into the saline solution. Arbitrarily, three times the concentration of MIC of S. aureus in saline solution was taken on a glass slide as before and cycled to determine the percentage survival of microbes and the results are shown in Figure 7. It is seen that the Ag/agar is more effective upto the third cycle in the case of C. albicans compared to E. coli and S. aureus, and there is a steep rise in the percent survival after the second cycle in the latter two. From the Kirby–Bauer test, it is found that as the concentration of Ag NPs increased, the zone radius also increased for all the microbes tested and they follow a linear plot. The zone radius for C. albicans is higher at a particular concentration of silver film (Fig. 8a) and the line is slightly steeper, compared to the other two microbes studied. A representative photograph of ZOI for E. coli in Figure 8b shows the increase in zone radius as Ag concentration is increased (anti clockwise). 3.3. Cytotoxicity experiments Upon conducting cytotoxicity experiments, the percentage survival of HeLa cells was plotted against the concentration of Ag NPs in Ag/agar and is shown in Figure 9. From the plot it can be seen that the Ag NPs exhibit a very low level of toxicity (72 lg/mL). Beyond this concentration of Ag NPs a drastic reduction of viability of the cells is seen. However, it may be noted that 9 lg/mL was the MIC found against the water-borne coliform bacterium, E. coli, and this concentration is almost seven times less than the threshold concentration of 60 lg/mL as seen in the plot.
Table 2 Mechanical properties of the films
Figure 9. Bar chart showing percentage survival of HeLa cells against concentrations of Ag NPs in Ag/agar.
Cycles
Elastic modulus (GPa)
Hardness (MPa)
Blank agar–agar 0th 1st 2nd 3rd 4th
1.229 ± 0.028 4.498 ± 0.117 4.139 ± 0.097 3.734 ± 0.162 3.804 ± 0.331 3.462 ± 0.057
36.6 ± 0.001 272.8 ± 0.009 268.2 ± 0.008 256.9 ± 0.133 241.8 ± 0.023 239.3 ± 0.008
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Supplementary data Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.carres.2010.08.001. References
Figure 10. Normalized hardness and elastic modulus of the film plotted as a function of number of cycles.
3.4. Mechanical properties The results of the elastic modulus (E) and hardness (H) of bare agar–agar and Ag/agar film after each cycle, calculated using Oliver–Pharr44 method, are listed in Table 2. These properties normalized with respect to Ag/agar film at zero cycle are plotted against the number of cycles (Fig. 10). The elasticity of Ag/agar film has increased almost four fold compared to the pure agar–agar film. Similarly the hardness has increased by seven times. There is only a decrease of hardness around 5% after second cycle and a decrease of around 10% on subsequent cycles, indicating good stability of the film. As a comparison, for example, Ag/agar films have mechanical strength more than fifty times compared to PVA-(polyvinyl alcohol)reinforced graphine films.45 4. Conclusions We have described a simple biocompatible, in situ synthesis of highly stable (more than 18 months) Ag nano particles in agar– agar matrix as free-standing film, using alcohol as reducing agent. Films are characterized by UV–vis absorption and TEM studies. The antimicrobial activity of the film is found to be in the order of C. albicans > E. coli > S. aureus. For the first time we have demonstrated the antimicrobial activity of the film on recycling in a saline solution. The activity reduces on cycling, mainly because of depletion of Ag NPs in the film on repeated use. Electron microscopic studies show the changes in morphology of microbes on the effect of Ag NPs and in turn the agglomeration of Ag NPs on exposure to microbes. Cytotoxicity experiments show a reasonably low toxic limit of Ag NPs. The Ag/agar film exhibits good mechanical stability. It is further worth studying to understand the exact mechanism of antimicrobial activity of Ag/agar film. Acknowledgments The authors thank the Nano Centre (IISc) for electron microscope facilities. The authors are indebted to Professor U. Ramamurty and Dr. S.R.N. Kiran Mangalampalli of the Department of Materials Engineering (IISc) for the timely help in the measurement of mechanical properties. H.N.V. thanks the generous internal funding from IISc, Bangalore.
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