Bacterial tolerance to silver nanoparticles (SNPs): Aeromonas punctata isolated from sewage environment

Journal of Basic Microbiology 2011, 51, 183 – 190

183

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Research Paper

Bacterial tolerance to silver nanoparticles (SNPs):
Aeromonas punctata isolated from sewage environment

S. Sudheer Khan, E. Bharath Kumar, Amitava Mukherjee and N. Chandrasekaran

Nanobio-Medicine Laboratory, School of Bio-Sciences and Technology, VIT University, Vellore, India

Use of silver nanoparticles (SNPs) is increasing in a large number of consumer products. Thus,
the possible build-up of the nanoparticles in the environment is becoming a major concern.
Aeromonas punctata isolated from sewage showed tolerance to 200 μg/ml SNPs. The growth
kinetics data for A. punctata treated with nanoparticles were similar to those in the absence of
nanoparticles. There was a reduction in the amount of exopolysaccharides (EPS) in bacterial
culture supernatant after nanoparticle-supernatant interaction. EPS capping of the nanopartic-
les was confirmed by UV-visible, XRD and comparative FTIR analysis. The EPS-capped SNPs
showed less toxicity to Escherichia coli, Staphylococcus aureus and Micrococcus luteus compared to
the uncapped ones. The study suggests capping of nanoparticles by bacterially produced EPS as
a probable physiological defense mechanism.

Keywords: SNPs / Aeromonas punctata / EPS / Encapsulation / Bacterial tolerance / Toxicity reduction

Received: February 19, 2010; accepted: July 08, 2010

DOI 10.1002/jobm.201000067

Introduction

Silver nanoparticles (SNPs) are potential candidates of
strong antimicrobial activity and are used in significant
amounts in consumer products, in the food industry
for storage, packaging, and processing [1], in textiles [2],
in medical applications for wound care products and
therapeutic devices [3], and in diagnostics and drug
delivery [4, 5].
But increasing concentrations of SNPs with varied
physical and surface properties could pose a threat to
human and environmental health [6]. Impellitteri et al.
2009 [7] revealed that the SNPs impregnated in clothes
and washing systems can easily leak into wastewater
during washing, thus potentially disrupting helpful
bacteria used in wastewater treatment facilities or en-
dangering aquatic organisms in lakes and streams.
In vitro and in vivo toxicity studies in mammalian
species proved that SNPs have the capability to enter
cells and cause cellular damage [8]. They have the po-
tential to cause chromosomal aberrations and DNA

Correspondence: Prof. Dr. N. Chandrasekaran, Nano Bio-Medicine
Laboratory, School of Bio-Sciences and Technology, VIT University,
Vellore-632014, India
E-mail: nchandrasekaran@vit.ac.in; nchandra40@hotmail.com
Phone: +91 416 2202624

damage and are capable of inducing proliferation arrest
in cell lines [9].
Indiscriminate use of nano-sized silver materials in
commercial products leads to their release into the
environment and ultimately harms microbial commu-
nities in the ecosystem [10]. The release of SNPs enter-
ing sewage treatment plants was estimated to be 270 t/
year [11]. Due to this, sensitive bacteria get disturbed,
besides harming some beneficial forms as well. Ulti-
mately, the released SNPs destabilize the functioning of
ecosystems [11]. On the other hand, some resistant
bacteria exist due to their adaptive nature [12]. The
hypothesis of the paper was to study the principle un-
derlying bacterial tolerance to SNPs and its possible
mechanism.

Materials and methods

Materials
All chemicals and media were obtained from Himedia
Laboratories Ltd., Mumbai, India. The bacterium was
isolated from sewage (Vellore, India). Manufactured
SNPs were obtained from Sigma Aldrich, USA. The
nanoparticles were dispersed using an ultrasonic pro-
cessor at a frequency of 132 kHz (Crest, USA).

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Characterization of SNPs
The SNPs were characterized using UV-visible spectro-
scopy and a high-resolution transmission electron mi-
croscope (TEM; Tecnai G-20) after dispersion in LB me-
dium. The samples were prepared by placing a drop of
homogeneous suspension on a copper grid with a lacey
carbon film and air-drying. The mean particle size was
analyzed from the digitized images with Image Tool
software. The morphological features of the manufac-
tured SNPs were characterized by scanning electron
microscopy (SEM; FEI Sirion, Eindhoven, The Nether-
lands). The surface area was measured using a Smart
Sorb 93 single-point BET surface area analyzer (Smart
Instruments Co. Pvt. Ltd., Mumbai, India). The received
particles were also subjected to X-ray diffraction (XRD)
analysis using a JEOL-JDX 8030 diffractometer. The
target was Cu Kα (

= 1.54 Å). The generator was oper-

ated at 45 kV and with a 30 mA current. The scanning
range (2

) was selected from 10 to 100°.

Isolation and identification of microorganisms
The sample collected from the sewage environment
(1 ml) was grown on LB broth supplemented with
100 µg/ml SNPs and incubated at 30 °C for 24 h with
agitation (200 rpm). The bacterial culture isolated from
the above medium was identified using the Sherlock
microbial identification system (MIS) following the
method of Wintzingerode et al. [13] and 16S rRNA analy-
sis. The 16S rRNA gene sequencing analysis was done by
using the primers 27F (5′-AGAGTTTGATCCTGGCTCAG-
3′) and 1492R (5′-GGTTACCTTGTTACGACTT-3′) [14].
Nearly complete 16S rRNA nucleotides were aligned
and bacterial identities were deduced by BLAST search
to ascertain their closest relatives. A phylogenetic tree
was constructed by using ClustalW. pls check

Disc diffusion and agar well diffusion method
Bacterial sensitivity to SNPs was tested by the disc dif-
fusion method (25, 50 and 100 µg per disc) [15] and the
well diffusion method (10, 25, 50, 100 and 200 μg/well)
[16]. The average diameter of the inhibition zone sur-
rounding the disc/well was measured. Six replicates
were employed for all concentrations. The bacteria
Escherichia coli (ATCC 13534 and ATCC 25922), Staphylo-
coccus aureus (ATCC 25923) and Micrococcus luteus (clinical
isolate) were used as positive control. The agar well
diffusion method was carried out with silver nitrate
solution at concentrations of 10, 25, 50, 100, 250 and
500 µg/ml.

Dilution plate count method
The different concentrations of SNPs (10, 25, 50, 100,
and 200 µg) were applied uniformly on the surface of

LB agar plates and the colony-forming units (CFU) were
determined at each concentration. Nanoparticle-free
plates incubated under the same conditions were used
as control. The counts on six plates corresponding to a
particular concentration were averaged.

Silver ion concentration measurement
SNPs dispersed in LB broth at 100 and 200 µg/ml were
centrifuged at 15,000 × g for 30 min. The clear super-
natant was carefully collected and filtered through a
0.1 µm sterile filter. The ion concentrations were meas-
ured by an AAS after acidification by 1% nitric acid
[17].

Growth kinetics study
To examine the bacterial growth profile in the presence
of SNPs at increasing concentrations (10–200 µg/ml), LB
broth was sonicated for 30 min after adding the SNPs to
prevent the aggregation of nanoparticles. Subsequently,
the flasks were inoculated with 1 loop full of freshly
prepared bacterial culture and incubated in an orbital
shaker at 200 rpm and room temperature (30 °C). Nano-
particle-free media were used as control. The bacterial
growth was measured every 2 h by using a colorimeter
(Elico CL 157) at 600 nm. A positive control (flask con-
taining nanoparticles and nutrient media, devoid of
inoculum) and a negative control (flask containing
inoculum and nutrient media, devoid of nanoparticles)
were included in this experiment. The absorbance value
for the positive control was subtracted from the ex-
perimental values (flasks containing nutrient media,
inoculum and nanoparticles). The growth profiles of
positive control strains were recorded at a concentra-
tion of 100 µg/ml SNPs. The growth studies of all the
bacteria were performed with corresponding amounts
of silver ions that were released during the time of
dispersion of SNPs in the medium.

FTIR analysis
A loop full of culture of A. punctata was inoculated into
50 ml of LB broth and grown for 48 h at room tempera-
ture, under shaking at 180 rpm. The exopolysaccha-
rides (EPS) were extracted [18] and quantified [19]. The
purified EPS [18] were subjected to FTIR analysis
(Perkin-Elmer spectrometer – one instrument in diffuse
reflectance mode at a resolution of 4 cm

–1

in KBr pel-

lets).
At the same time, another part of the A. punctata
culture supernatant (10 ml) was collected and inter-
acted with 100 µg/ml SNPs for 4 h, in a rotary shaker at
200 rpm to maintain a proper interaction with nano-
particles. After the interaction, the mixture was centri-

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Bacterial tolerance to silver nanoparticles

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fuged and the nanoparticle pellets were separated, ly-
ophilized and subjected to FTIR analysis. The super-
natant was collected, and the EPS left in the culture
supernatant after interaction with SNPs were quanti-
fied. The EPS extraction and FTIR analysis were also
done with positive control strains.

UV-visible spectral analysis and XRD analysis
The purified EPS were redissolved in deionized (DI)
water and interacted with different concentrations of
SNPs (10–100 µg/ml), in a rotary shaker at 200 rpm to
maintain proper interaction with the nanoparticles.
The absorbance of SNPs was measured after 4 h of in-
teraction, by using a UV-visible spectrophotometer
(Shimadzu UV – 1700, Japan) at 200–700 nm. After the
interaction, the SNPs were separated, lyophilized and
analyzed by XRD (PANalytical XPert Pro, Eindhoven,
The Netherlands).

Toxicity test for EPS-coated SNPs

To examine the effect of EPS-coated nanoparticles, the
coated and uncoated SNPs were interacted with positive
control strains at a concentration of 100 µg/ml as per
the method described earlier.

Results and discussion

Characterization of SNPs
UV-visible spectroscopy is one of the techniques used
for the structural characterization of SNPs. UV-visible
absorption spectra for the manufactured SNPs showed
an absorption band at 425 nm (Fig. 1). The surface
area of the manufactured SNPs was determined to be

Figure 1. UV-visible absorption spectra of dispersed SNPs in LB
broth (a) before dilution, (b) after 10 times dilution.

0.26 m

/g. The TEM images showed that the SNPs were

spherical in shape and polydispersed with diameters in
the range of 5 to 40 nm. The SEM images showed SNPs
whose size was below 40 nm. The XRD pattern of the
dispersed SNPs was characterized by a major peak at
38.115 and minor peaks at 44.299, 64.443, 77.397,
81.541 and 98.241. This confirms the stability of the
nanoparticles in the LB medium.

Microbial identification
MIS studies and 16S rRNA analysis showed that our
bacterial isolate from sewage was A. punctata (data not
shown). Fig. 2 represents the phylogenetic tree with
boot strap values. The BLAST search shows 99% similar-
ity to the A. punctata strain. The 16S rRNA sequence was
submitted to Genbank (Accession number GQ401237)
and the organism was named A. punctata VITSCA01.

Antimicrobial tests
The antimicrobial properties of SNPs against the isolate
A. punctata were investigated by disc diffusion test, agar
well diffusion method, dilution plate count method and
growth kinetics studies. The disc impregnated with
SNPs in the A. punctata culture plate gave no zone of
inhibition. There was also no zone of inhibition ob-
served in the agar well diffusion method (Fig. 3). In the
disc diffusion test, E. coli 13534, E. coli 25922 and
M. luteus (clinical isolate) showed a small inhibition at
50 µg/disc. However, S. aureus 25923 gave an inhibition
zone at 100 µg/disc. In the agar well diffusion method,
the zone of inhibition was observed in all control or-
ganisms at 100 and 200 µg (Fig. 3). Three different agar
media (Muller Hinton agar, Nutrient agar and LB agar)
were used in both experiments. There was no differ-
ence in the activity of SNPs observed in all the media
tested. The study of Ruparelia et al. [15] has shown that
a disc impregnated with spherical SNPs showed a large
inhibition zone for E. coli, S. aureus and Bacillus subtilis.
The study with spherical SNPs in the size range from 10
to 100 nm diameter exhibited excellent antibacterial
activity against the bacteria S. aureus, B. cereus, E. coli and
Pseudomonas aeruginosa, with the agar well diffusion
method, and clearing zones around the holes with bac-
teria growth were observed [16].
A.

punctata showed tolerance to silver nitrate at 100,

250, and 500 µg/ml concentration. A narrow zone at
500 µg/ml that measured approximately 1 mm in di-
ameter was observed (not shown). This indicates that
A. punctata was resistant to 100 and 250 µg/ml silver and
showed the lowest resistance at 500 µg/ml. The studies
proved the mechanism of resistance to be plasmid me-
diated [20–22].

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Figure 2. Phylogenetic tree based on 16S rRNA gene sequence comparison showing the position of the silver nanoparticle-tolerant
bacterial strain isolated from sewage.

Figure 3. Representative images of agar plates containing SNP-
impregnated discs and wells. (A) A. punctata, (B) E. coli (ATCC
13534), (C) E. coli (ATCC 25922), and (D) S. aureus (ATCC
25923); (a) 25, (b) 50, (c) 100, and (d) 200

μg SNPs.

  When nanoparticles were present on the surface of
the nutrient agar plates, they could more completely
inhibit bacterial growth as compared to liquid broth.
The growth of A. punctata in agar plates supplemented
with different concentrations of SNPs was not found to
be different compared to the control plate. The ob-
served bacterial counts (mean ± standard error) in the
agar plates were 84.6 ± 0.8, 84.8 ± 0.9, 83.3 ± 0.8, 84.8 ±
0.6, 84.3 ± 0.9, and 83.5 ± 0.8 for control, 10, 25, 50,
100, and 200 µg SNPs, respectively (not shown).
  Fig. 4 shows the bacterial growth curves for diffe-
rent concentrations of SNPs. In comparison with the
control, no growth inhibition was observed at all the
concentrations of SNPs tested. The present study with
positive control strains showed a great reduction in
the growth of the bacteria (Fig. 5), similar to the
studies of Sondi and Salopek-Sondi [23]. Fig. 6 shows
the interaction of A. punctata with SNPs. Most of the
studies reported bacterial growth inhibition by nano-
particles at several optimal concentrations [15, 23–25].

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Figure 4. Representative batch growth profile of A. punctata in LB
broth dosed with 10, 25, 50, 100 and 200

μg/ml SNPs and control

broth (without nanoparticles) at 30 °C.

Overuse of nanoparticles in consumer products and
washing of these into sewage systems might induce
resistance to environmental strains [26].
On dispersion, a substantial amount of silver ions
was quickly released from SNPs into the LB medium. At
100 µg/ml SNPs, 1.4 ± 0.002 µg/ml silver ions were re-
leased, and 200 µg/ml of SNPs released 2.9 ± 0.01 µg/ml
silver ions into the LB medium. To evaluate the toxic
effect of silver ions released during the dispersion of
SNPs, the LB medium was centrifuged at 15,000 × g for
30 min after dispersion. The SNPs settle down and the
silver ions remain in the medium. The growth kinetics
studies of A. punctata did not show any growth reduction

Figure 5. Toxicity studies of EPS-coated and uncoated SNPs with
E. coli 13534. The same pattern was observed with three other
control species.

Figure 6. SEM image of A. punctata interacted with SNPs. The pro-
duction of EPS can be observed in the figure.

in the above medium. A. punctata may not be affected
by these silver ions due to its above-mentioned resistant
nature. The growth kinetics studies with E. coli (ATCC
13534, ATCC 25922), S. aureus (ATCC 25923) and M. lu-
teus (clinical isolate) showed only a negligible growth
reduction. This suggests that there might be some
other mechanism behind the action of SNPs. The re-
lease of silver ions from SNPs may vary depending on
the medium used for dispersion.
Morones et al. [25] reported that the bactericidal
mechanisms of SNPs and silver ions are distinctly dif-
ferent. For treatment with silver nitrate, a low-mole-
cular-weight central region was formed within the cells
as a defense mechanism, whereas for treatment with
nanoparticles no such phenomenon was observed, al-
though the nanoparticles were found to penetrate the
cell wall [23].

Role of EPS in defense mechanism
The results of EPS quantification showed that the
amount of EPS present in the supernatant after interac-
tion with SNPs was lower (82.3 ± 0.9 µg/ml) compared
to the amount of EPS extracted before interaction
(113.7 ± 0.6 µg/ml). A reduction in the amount of EPS
was observed in the supernatant after SNPs-supernatant
interaction. This result suggests that the EPS might
have coated the SNPs during interaction.
The FTIR spectral profile obtained in the 1000–
1200 cm

–1

region mainly reflected the absorption of

sugars present in the EPS. The studies showed that FTIR

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spectroscopy affords a rapid and easy means for indicat-
ing the nature of major components of A. punctata EPS
(Fig. 7a). The three peaks near 1000–1200 cm

–1

indicate

that the polysaccharide was α-pyranose. After interac-
tion of the SNPs with A. punctata culture supernatant,
the FTIR spectra matched all the peak values of the
extracted EPS (Fig. 7b). The coating with EPS during the

interaction of SNPs with the bacterial culture super-
natant was confirmed by evaluating the FTIR peak
value in both these cases.
Hydroxyl, carboxyl, carbonyl and amine groups were
found in the FTIR spectra of EPS. The FTIR spectra of
EPS secreted by E. coli (ATCC 13534, ATCC 25922) and
S. aureus were entirely different from that of the test

Figure 7. (a) FTIR spectra of EPS extracted from A. punctata culture supernatant. (b) FTIR spectra of lyophilized SNPs after 4 h of inter-
action with A. punctata culture supernatant.

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strain. Production of EPS was not found in the case of
M. luteus.
Noble metal SNPs exhibit unique and tunable optical
properties, on account of their surface plasmon reso-
nance (SPR) which depends on shape, size and size dis-
tribution of the nanoparticles. When SNPs were inter-
acted with EPS extracted from the culture supernatant
of A. punctata, UV-visible spectrophotometric analysis
gave a peak at 394 nm for the SNPs. On the other hand,
the

max

value for the uninteracted SNP dispersion was

noted at 425 nm. A decrease in absorbance at 425 nm
was observed when it was interacted with EPS. After
interaction with EPS, a peak shift was observed (blue
shift). This might be due to the repelling action of EPS-
capped SNPs. The breaking up of the agglomerates/flocs
would result in a blue shift. The blue shift (towards
lower wavelength) in the

max

value after interaction

with EPS demonstrated that the shape of the nanopar-
ticles did not suffer distinct modifications. The blue
shift indicates that the decrease in size might be due to
the coating with EPS that prevents the further agglom-
eration of SNPs.
  EPS coating was confirmed by XRD analysis. The XRD
study corroborated the adsorption of EPS on the nano-
particle surface; the coverage was so strong that no
characteristic band of silver could be revealed through
XRD, whereas the manufactured SNPs gave six peaks in
XRD analysis. Ravindran et al. [27] reported that, in XRD
analysis, SNPs that were interacted with BSA did not
give the characteristic peak of silver, due to the coating
of SNPs with BSA.
  The toxicity test of EPS-coated and uncoated SNPs
(100 µg/ml) with four different strains, E. coli 13534,
E. coli 25922, S. aureus 25923 and M. luteus (clinical iso-
late), showed a great reduction in the growth rate of
these bacteria when treated with uncoated nanoparti-
cles compared to the control growth profile (without
nanoparticles). But not much reduction was observed
when the culture medium was supplemented with
coated SNPs, indicating lower toxicity (Fig. 5).
  Nanosilver may compromise to control harmful bac-
teria. Besides that it may affect the beneficial bacteria
present in the soil and sewage treatment plants. This
study confirmed that the release of nanoparticles
into the wastewater in a wastewater treatment plant
would not affect all the beneficial microbes involved
in the sewage treatment process. A. punctata might
protect the survival of the beneficial microbes by pro-
viding an EPS capping to the nanoparticles. On the
other hand, the EPS-capped SNPs can be used for com-
mercial application including drug delivery and bio-
sensing.

Conclusions
A. punctata isolated from sewage showed tolerance to
SNPs at the tested concentrations of up to 200 μg/ml.
The findings from this study suggest that, when grown
in the presence of increasing concentrations of SNPs,
environmental isolates may acquire tolerance/resist-
ance. One of the important implications of our findings
is that future predictions on the environmental toxicity
of nanoparticles, especially for microbes, need to take
into account the possible adaptation of the environ-
mental strains to high concentrations of nanoparticles.
This physiological defense mechanism developed might
be due to the presence of other toxic compounds in the
sewage system. Although the toxicity effects in situ will
definitely differ from those observed in a controlled
laboratory environment, we strongly feel that, in the
absence of prior reports on acquired nanoparticle toler-
ance in environmental isolates, our study gains impor-
tance. Finally, we stress here the need to take up future
in situ studies involving different nanoparticles and
further microbial strains for a clearer understanding of
the environmental toxicity effects of nanoparticles.

Acknowledgement

The authors thank the Management of VIT University
for the provision of funding to carry out this research.

References

[1] Das, M., Saxena, N., Dwivedi, P.D., 2009. Emerging trends

of nanoparticles application in food technology: Safety

paradigms. Nanotoxicology, 3, 10–18.

[2] Perelshtein, I., Applerot, G., Perkas, N., Guibert, G., Mik-

hailov, S., Gedanken, A., 2008. Sonochemical coating of

silver nanoparticles on textile fabrics (nylon, polyester

and cotton) and their antibacterial activity. Nanotechno-

logy, 19, 245705.

[3] Furno, F., Morley, K.S., Wong, B., Sharp, B.L., Arnold, P.L.

et al., 2004. Silver nanoparticles and polymeric medical

devices: A new approach to prevention of infection? J. An-

timicrob. Chemother., 54, 1019–1024.

[4] Roe, D., Karandikar, B., Savage, N.B., Gibbins, B., Roullet,

J.B., 2008. Antimicrobial surface functionalization of

plastic catheters by silver nanoparticles. J. Antimicrob.

Chemother., 61, 869–876.

[5] Li, P., Li, J., Wu, C.,Wu, Q., Li, J., 2005. Synergistic anti-

bacterial effects of β-lactam antibiotic combined with sil-

ver nanoparticles. Nanotechnology, 16, 1912–1917.

[6] Panyala, N.R., Pena-Mendez, E.M., Havel, J., 2008. Silver or

silver nanoparticles: A hazardous threat to the environ-

ment and human health? J. Appl. Biomed., 6, 117–129.

[7] Impellitteri, C.A., Tolaymat, T.M., Scheckel, K.G., 2009.

The speciation of silver nanoparticles in antimicrobial

190 S.

Sudheer

Khan

et al.

Journal of Basic Microbiology 2011, 51, 183 – 190

www.jbm-journal.com

fabric before and after exposure to a hypochlorite/deter-

gent solution. J. Environ. Qual., 38, 1528–1530.

[8] Hussain, S.M., Schlager, J.J., 2009. Safety evaluation of

silver nanoparticles: Inhalation model for chronic expo-

sure. Toxicol. Sci., 108, 223–224.

[9] AshaRani, P.V., Mun, G.L.K., Hande, M.P., Valiyaveettil, S.,

2009. Cytotoxicity and genotoxicity of silver nanoparticles

in human cells. ACS Nano, 3, 279–290.

[10] Senjen, R., 2007. Nanosilver – a threat to soil, water and

human health? Friends of the Earth, Australia.

[11] Blaser, S., Scheringer, M., MacLeod, M., Hungerbuhler, K.,

2008. Exposure of modeling of nano silver in the envi-

ronment. nanoECO Conference, March 3; Monte Verita.

[12] Chopra, I., 2007. The increasing use of silver-based pro-

ducts as antimicrobial agents: A useful development or a

cause for concern? J. Antimicrob. Chemoth., 59, 587–590.

[13] Wintzingerode, F., Rainey, F.A., Kroppenstedt, R.M., Sta-

ckebrandt, E., 1997. Identification of environmental

strains of Bacillus mycoides by fatty acid analysis and spe-

cies-specific 16S rDNA oligonucleotide probe. FEMS Mic-

robiol. Ecol., 24, 201–209.

[14] Yim, W.J., PoonGuzhali, S., Madhaiyan, M., Palaniappan,

P., Siddikee, M.A., Tongmim, S., 2009. Characterization of

plant growth promoting diazotrophic bacteria isolated

from field grown Chinese cabbage under different fertili-

zation conditions. J. Microbiol., 47, 147–155.

[15] Ruparelia, J.P., Chatterjee, A., Duttagupta, S.P., Mukherji,

S., 2008. Strain specificity in antimicrobial activity of sil-

ver and copper nanoparticles. Acta Biomater., 4, 707–716.

[16] Maliszewska, I., Sadowski, Z., 2009. Synthesis and anti-

bacterial activity of silver nanoparticles. J. Phys: Confer-

ence Series, 146, 012024.

[17] Jiang, W., Mashayekhi, H., Xing, B., 2009. Bacterial toxici-

ty comparison between nano- and micro-scaled oxide par-

ticles. Environ. Pollut., 157, 1619–1625.

[18] Ashok Kumar, Sunish Kumar, R., Sakthivel, N., 2003.

Compositional difference of the EPS produced by the viru-

lent and virulence-deficient strains of Xanthomonas oryzae

pv. oryzae. Curr. Microbiol., 46, 251–255.

[19] Mitchel, D., Gilled, K.A., Hamilton, J.K., Roberts, P.A.,

Smith, F., 1956. Calorimetric method for determination of

sugars and related substances. Anal. Chem., 28, 350–356.

[20] Deshpande, L.M., Chopade, B.A., 1994. Plasmid mediated

silver resistance in Acinetobacter baumannii. BioMetals, 7,

49–56.

[21] Starodub, M.E., Trevors, J.T., 1990. Mobilization of Escheri-

chia coli R1 silver-resistance plasmid pJT1 by Tn5-Mob into

Escherichia coli C600. BioMetals, 3, 24–27.

[22] Kaur, P., Saxena, M., Vadehra, D.V., 1985. Plasmid medi-

ated resistance to silver ions in Escherichia coli. Indian J.

Med. Res., 82, 122–126.

[23] Sondi, I., Salopek-Sondi, B., 2004. Silver nanoparticles as

antimicrobial agent: A case study on E. coli as a model for

gram-negative bacteria. J. Colloid Interf. Sci., 275, 177–

182.

[24] Shrivastava, S., Bera, T., Roy, A., Singh, G., Ramachandra-

rao, P., Dash, D., 2007. Characterization of enhanced an-

tibacterial effects of novel silver nanoparticles. Nano-

technology, 18, 225103.

[25] Morones, J.R., Elechiguerra, J.L., Camacho, A., Holt, K.,

Kouri, J.B., Ramirez, J.T., 2005. The bactericidal effect of

silver nanoparticles. Nanotechnology, 16, 2346–2353.

[26] Melhus, A., 2007. Silver threatens the use of antibiotics

(unpublished manuscript).

[27] Ravindran A., Singh A., Raichur A. M., Chandrasekaran N.,

Mukherjee A., 2009. Studies on interaction of colloidal Ag

nanoparticles with bovine serum albumin (BSA). Colloid

Surface B, 76, 32–37.