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Colloids and Surfaces B: Biointerfaces 84 (2011) 338–345

Contents lists available at ScienceDirect

Colloids and Surfaces B: Biointerfaces

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iomimetic preparation of polymer-supported free radical scavenging,ytocompatible and antimicrobial “green” silver nanoparticles using aqueousxtract of Citrus sinensis peel

ocktotpal Konwarha, Biswajit Gogoia, Ruby Philipb, M.A. Laskarb, Niranjan Karaka,∗

Advanced Polymer and Nanomaterial Laboratory, Department of Chemical Sciences, Tezpur University, Napaam-784028, Sonitpur District, Tezpur, Assam, IndiaDepartment of Biotechnology, St. Anthony’s College, Shillong-793001, Meghalaya, India

r t i c l e i n f o

rticle history:eceived 21 October 2010eceived in revised form 14 January 2011ccepted 16 January 2011vailable online 25 January 2011

eywords:

a b s t r a c t

In the pursuit of making the nanoscale-research greener, the utilization of the reductive potency of acommon byproduct of food processing industry i.e. orange peel is reported here to prepare biopolymer-templated “green” silver nanoparticles. Aqueous extract of orange peel at basic pH was exploited toprepare starch supported nanoparticles under ambient conditions. The compositional abundance ofpectins, flavonoids, ascorbic acid, sugars, carotenoids and myriad other flavones may be envisaged forthe effective reductive potential of orange peel to generate silver nanoparticles. The nanoparticles were

itrus sinensis peelilver nanoparticlesnti-microbialree radical scavengingHP-1 cytocompatibilitynti-lipid peroxidation

distributed within a narrow size spectrum of (3–12 nm) with characteristic Bragg’s reflection planes offcc structure, and surface plasmon resonance peak at 404 nm. Anti-lipid peroxidation assay using goatliver homogenate and DPPH scavenging test established the anti-oxidant potency of the silver nanopar-ticles. Their synergy with rifampicin against Bacillus subtilis MTCC 736 and cytocompatibility with thehuman leukemic monocytic cell line, THP-1 were also investigated. Thus, the present work deals with thepreparation of starch assisted anti-microbial, cytocompatible and free radical scavenging “green” silvernanoparticles.

. Introduction

The accord of the medical world and the material sciences continually strengthened by the ever widening spectrum ofpplications of silver nanoparticles (NPs). However, many of theonventional methods [1] for their generation rely on the use ofrganic solvents and toxic reducing agents like sodium borohydridend N, N-dimethylformamide. For cleaner preparation of theseoble metal nanoparticles, various biomimetic approaches [2,3]re being explored. Green technology, in the niche of preparativerotocols of nanomaterial stands for switching to environmen-ally benign starting materials, water as the reaction medium, aiodegradable polymer as the stabilizing template with minimalastage in terms of energy and raw materials [4]. In this pursuit,

he conjugation of amyloglucosidase onto magnetically recyclable

ilver nanoparticles, prepared using Mesua ferrea L. leaf extract [5]as been recently reported.

In this work, we wish to report the utilization of orange peel,common waste material of the food processing industry for the

∗ Corresponding author. Tel.: +91 3712 267009; fax: +91 3712 267006.E-mail address: karakniranjan@yahoo.com (N. Karak).

927-7765/$ – see front matter © 2011 Elsevier B.V. All rights reserved.oi:10.1016/j.colsurfb.2011.01.024

© 2011 Elsevier B.V. All rights reserved.

preparation of starch supported silver nanoparticles. The compo-sitional abundance of pectins, flavonoids, ascorbic acid, sugars,carotenoids, limonene and various other flavones [6,7] in the epi-carp of different citrus species has been reported. Different in vitroassays in brain homogenates have been used to establish the excel-lent antioxidant potency of the various polar fractions of orangepeel [7]. These components may be envisaged for the effectivereductive potential of orange peel to generate silver nanoparticlesby a greener route.

Polymer encapsulation or entrapment is an excellent meanto tune particle size distribution and to improve their stabil-ity, dispersibility and compatibility with organic systems [8].These are critical factors for silver nanoparticles to complementtheir biomedical applications based on plasmonic attributes, anti-microbial potency and target-specific localization. In this context,starch, a biodegradable polymer can be used as matrix for thestabilization of silver nanoparticles, as it forms right-handedhelical conformation in aqueous solution and displays complex-

ation of metal ions onto the molecular matrix with tremendousmorphology-directing action [2]. This green capping potency ofstarch has already been confirmed [2] for the preparation of sil-ver nanoparticles, with the requisite of microwave-assistance orhydrothermal energy. But, in the present study, starch is used as

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he templating matrix and the reductive potential of orange peel istilized for the preparation of silver nanoparticles at ambient tem-erature, eliminating the additional energy input, unlike the above2].

Free radicals, leading to oxidative stress can cause biologicalavoc. Jain et al. [9] have reported that antimicrobial gel formu-

ation containing silver nanoparticles elicit oxidative stress withubsequent trigerring of the cellular anti-oxidant system. On theontrary, a crucial role of novel Ag+-loaded zirconium phosphateanoparticles in diabetic wound healing has been documented10]. Thus, investigating the free radical scavenging potency of thereen silver nanoparticles forms an interesting proposition. Fur-hermore, tunable antibacterial coatings with silver nanoparticles,enerated through in situ route, which support mammalian cellrowth, have been developed [11]. Establishing the biocompati-ility of the reported “green” silver nanoparticles is important inhe context of their biomedical utility.

Therefore, the authors wish to report the reduction potency ofrange peel for the preparation of starch supported silver nanopar-icles in addition to the utilization of its anti-oxidant potency. Theanoparticles are characterized using various techniques and their

ree radical scavenging potency is also evaluated. Furthermore, thenti-bacterial potential of silver nanoparticles using Bacillus sub-ilis as the test organism and their cytocompatibility with humaneukemic monocytic cell line (THP-1) are also evaluated.

. Materials and methods

.1. Materials

Silver nitrate (Qualigens®), soluble starch (Qualigens®) andotassium hydroxide (Rankem) were used as received. RPMI 1640Roswell Park Memorial Institute 1640) medium (HiMedia), 90%BS (fetal bovine serum) (Sigma), MTT [3-(4,5-dimethyl thiazol--yl)-2,5-diphenyltetrazolium bromide] (HiMedia), dimethyl-ulphoxide (DMSO) (Merck), 0.25% trypsin (HiMedia), glycineuffer (pH 10.5) comprising of 0.1 M glycine (HiMedia), 0.1 Modium chloride (Rankem), peptone (HiMedia), beef extract (HiMe-ia), agar (Fisher Scientific), DPPH (1′-1′ diphenyl picryl-hydrazyle)HiMedia), trichloro acetic acid (TCA) (HiMedia), thiobarbituric acidTBA) (Himedia), iron sulphate (Qualigens) and 96 well Microtestlates (Corning).

B. subtilis MTCC 736 was used to evaluate the antimicrobialotency of the prepared nanoparticles. The human leukemic mono-ytic cell line (THP-1) was obtained from the National Center for Cellciences (NCCS), Pune, India and used for the cytotoxicity analysis.

.2. Preparation and characterization of the nanoparticles

About 0.2 g of orange peel was washed with de-ionized water,round using a domestic blender followed by stirring for about0 min in 50 mL of water at 50 ◦C. The aqueous extract was fil-ered through a muslin cloth at ambient temperature. 200 �l ofM AgNO3 was taken in 7.5 mL of 5% (w/v) soluble starch solutionith 6 mL of the extract. The pH was adjusted to basic condition

y adding adequate amount of aqueous 2 M KOH solution. Theolour of the solution gradually turned black indicating the forma-ion of silver nanoparticles. The matrix-supported particles wereubjected to centrifugal decantation followed by washing with Mil-ipore water and subsequently with acetone.

UV–visible spectra of the samples were recorded in HitachiU-2001, Tokyo, Japan) UV spectrophotometer. FTIR spectra forhe samples were taken from Nicolet (Impact 410, Madison, WI)TIR spectrometer by using KBr pellets. An X-ray diffractometer,

Miniflex’ (Rigaku Corporation, Japan), was used for the analysis

: Biointerfaces 84 (2011) 338–345 339

of film sample at room temperature (approx. 23 ◦C). The scanningrate of 5.0◦ min−1 over the range of 2� = 10–90◦ was used. Sizeand distribution of nanoparticles were studied using JEOL, JEM-CXII transmission electron microscope (TEM) at operating voltageof 100 kV.

2.3. Anti-lipid peroxidation assay

Goat liver, used for anti-lipid peroxidation (ALP) assay, wascollected from slaughter house immediately after slay and theexperiment was conducted within 1 h after the collection. Mam-malian liver was rinsed with phosphate buffered saline (PBS) toget rid of the residual blood, and 0.9 mL of 1.5 g L−1 KCl solutionwas added to 100 mg liver tissue to make 100 g L−1 homogenates ofliver, then stored at −20 ◦C. The anti-lipid peroxidation of the goatliver homogenate was measured according to the method used byMandal and Chaterjee [12] after slight modification. 2.8 mL of 10%goat liver homogenate, 0.1 mL of 50 mM FeSO4 and various volumesof the polymer supported green silver nanoparticles were mixedtogether with a micro-stirrer. The mixture was then incubated for30 min at 37 ◦C. 1 mL of this mixture was taken with 2 mL of 10%TCA-0.67% TBA in acetic acid (50%) to stop the reaction. This was fol-lowed by boiling for 1 h at 100 ◦C and subsequent centrifugation at10,000 rpm for 5 min. Absorbance of the supernatant was recordedat 535 nm against the reagent blank. Identical experiments wereperformed to determine the absorbance of the control (without thenanoparticles and FeSO4) and induced (without the nanoparticles)peroxidation. Vitamin C was used as the standard in this study. ALPpercentage was calculated using the following formula

%ALP = absorbance of Fe2+ induced proxidation − absorbance of sample

absorbance of Fe2+ induced peroxidation − absorbance of control× 100

(1)

2.4. DPPH scavenging assay

Antioxidant activity was also measured by using the modifiedDPPH method as reported previously [13]. To examine the concen-tration effect of the silver nanoparticles, 20 �l, 50 �l, 100 �l, 150 �land 200 �l of the polymer supported nanoparticles were mixedwith 2 mL of 100 �M DPPH solution. The samples were vortexedand allowed to scavenge DPPH in dark for 30 min. The absorbanceof the supernatants after centrifugation at 9200 × g for 2 min wasmeasured at 517 nm in UV–vis spectrophotometer. In all the cases,measurements were done in triplicates. The scavenging percentagewas calculated using the formula:

DPPH scavenging = (AC − As) × 100Ac

(2)

where AC and AS are absorption of blank DPPH and DPPH subjectedto interact with the sample at 517 nm, respectively.

2.5. Anti-bacterial assay

Nutrient agar (1.5%) was prepared by dissolving 5 g of peptone,5 g of beef extract, 8 g of NaCl and 15 g of agar in 1 L of water, fol-lowed by autoclaving under 15 lbs pressure at 121 ◦C for 20 min. Themedia was poured onto sterilized petriplates and allowed to solid-ify. In similar lines, the soft agar was prepared (0.7%) and whenit had acquired a temperature of about 35 ◦C, it was inoculatedwith the bacterial culture in the ratio of 1:10 (v/v). About 25 �l

of the inoculum was poured onto the agar plates. After the agarhad solidified, wells were punched as per the requirement. The sil-ver nanoparticles at various concentrations (Table 1) were vortexedwith desired concentration of rifampicin for 15 min. These combi-nations were injected into the wells with rifampicin (1 mg mL−1)

340 R. Konwarh et al. / Colloids and Surfaces B

Table 1Mean zone of inhibition (mm) ± standard deviation of various combinations ofantibiotic and silver nanoparticles against Bacillus subtilis MTCC 736.

Plate no./well number

Antibiotic(�g)

Silvernanoparticles(�g)

Mean zone of inhibition(mm) ± standard deviation

(a)/(1) 20 0 20 ± 0.00(a)/(2) 0 20 14 ± 0.5(a)/(3) 12 8 16 ± 1.0(a)/(4) 10 10 17 ± 0.5(a)/(5) 8 12 17 ± 0.5(a)/(6) 4 16 16 ± 1.0(a)/(7) 16 4 17 ± 0.5(b)/(1) 20 0 20 ± 0.0(b)/(2) 1 20 17 ± 0.5

aww

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peel extract. No characteristic silver’s SPR peak was observed when

Ft

(b)/(3) 2 20 17 ± 0.5(b)/(4) 3 20 17 ± 0.5(b)/(5) 4 20 17 ± 0.5(b)/(6) 5 20 17 ± 0.5

s the positive control. After 24 h of incubation at 37 ◦C, the platesere checked to measure the zone of inhibition. All experimentsere done in trplicates.

.6. MTT assay

MTT assay was used for the cytotoxicity evaluation of theanoparticles. THP-1 cell line was maintained in RPMI-1640 [14].or this test, briefly, 1 × 105 cells mL−1 at the exponential growthhase were placed in a flat-bottomed 96-well polystyrene-coatedlate and incubated for 24 h in a CO2 incubator at 37 ◦C in 5% CO2.o study the concentration effect of the nanoparticles, 14.1 �M,6.4 �M and 98.7 �M of the starch supported nanoparticles were

ntroduced into the medium. Each concentration was added tohe plate in triplicates. After termination of the incubation period,00 �l of MTT solution (5 mg mL−1) was added to the wells. Thelates were wrapped with aluminium foil and incubated for 4 h

n the CO2 incubator. This was followed by dissolution of formazonrystals formed in the respective wells in 100 �l of DMSO and 25 �lf glycine buffer (to adjust the pH). The plates were read in the ELISAlate reader (Eppendorf Biophotometer Plus Hamburg, Germany)t 540 nm. Wells with complete culture medium, nanoparticles,

nd MTT, but without cells were used as the blank for each case. Allxperiments were performed twice in triplicates, and the averagef all the experiments has been shown as a cell-viability percentagen comparison with the control experiment, while silver untreatedontrols were considered as 100% viable.

ig. 1. UV–vis spectra of (a) Orange peel extract and (b) starch templated “green” silver nahe progressive time course evolution of silver nanoparticles.

: Biointerfaces 84 (2011) 338–345

3. Results and discussion

3.1. Preparation and characterization of the “green” silvernanoparticles

In the present study, the first objective was to use the aqueousextract of orange peel to prepare polymer assisted “green” silvernanoparticles. The UV–vis spectra supported the generation of thesilver nanoparticles through the devised green-route. No absorp-tion peak was observed in UV–vis spectrum of Ag+ solution beforereduction. This is attributed to Ag+ ion’s d10 configuration. How-ever, addition of the reducing agent adjusted to basic pH, led togradual generation of silver nanoparticles as indicated by the pro-gressive increase (from 2 min to 45 min) in the absorbance intensityat around 400–410 nm in the UV–vis spectra. This is due to surfaceplasmon resonance (SPR) of silver nanoparticles (Fig. 1). The band isattributed to the collective oscillation of electron gas in the particleswith a periodic change in the electronic density at the surface. Var-ious parameters like particle size, shape, dielectric constant of themedium and surface adsorbed species determine the position andshape of the plasmon absorption [15]. The plasmonic coupling ofmetal nanoparticles with light augment a number of useful opticalphenomena that finds application in ultra-sensitive biomoleculardetection and lab-on-a-chip sensors [16].

It has been reported that the time required for completion ofthe reaction during biosynthesis of metal nanoparticles (i.e., com-plete reduction of the metal ions) using bacteria and fungi rangedfrom 24 to 124 h [3]. Gold nanoparticles, though of considerable sizediversity (20–40 nm), have been prepared using Geranium leaves in60 min, while a basic solution had reduced the time from 72 h to 1 hfor the preparation of protein stabilized spherical silver nanopar-ticles by Coriolus versicolor [3]. Here the rapid generation of thenanoparticles (in about 45 min) owing to the excellent reducingpotential of the active components of orange extract and their poly-meric stabilization within a narrow size spectrum (discussed in thesections for TEM and XRD analyses) has obvious advantages overthe previously reported systems.

At this juncture, the readers might be intrigued about the agentactually responsible for the reduction i.e. starch versus the orange

starch alone was used at ambient conditions without the input ofexternal thermal energy. This additional energy is required for thebreakdown of starch to generate active –OH groups through releaseof glucose units for the reduction of the metal ions. However, the

noparticles, showing the surface plasmon resonance at 404 nm. The inset (c) shows

faces B: Biointerfaces 84 (2011) 338–345 341

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onovalent silver ion is primarily reduced by –OH groups of thective components of the peel extract, which is oxidized to cor-esponding carbonyl group. This was authenticated from the IRpectra (image given as supplementary figure).

As mentioned earlier, orange peel represents a complex store-ouse of myriad of biomolecules like ascorbic acid, vitamin A,avonoid fractions, including hesperidin, neohesperidin, diosminnd various other polymethoxylated flavones like nobiletin andangeritin (rarely found in other plants) (Fig. 2). It is very muchikely that the reduction process of silver ion is interplay of numberf active components of the orange peel. For simplicity, let us con-ider the role of ascorbic acid as an effective reductant, illustratedn Fig. 3.

Ag+ + C6H8O6 � 2Ag0 + C6H6O6 + 2H+ (3)

he silver ions in the starch matrix are predominantly stabilized bylectrostatic interaction, while the narrow size and good dispersionf the silver nanoparticles in the polymeric support is primarilyttributed to steric stabilization.

Furthermore, pH played an important role (image not shown)n dictating the reducing potency of the orange extract. It is perti-ent to mention that under acidic conditions the active componentsf the peel extract are not effectively utilized for the reductionrocess. This may be due to their structural stabilization and asso-iation predominantly through H-bonding. The reduction potencyf ascorbic acid (the biomolecule under consideration as effectiveeducing agent) also decreases under acidic condition for anothereason. The nitrate group (of the silver salt) at low pH, is a strongerxidant than Ag+ [17] and can oxidize ascorbic acid as shown under-eath.

O3− + 4H3O+ + 3e− � NO ↑ +6H2O E0 = 0.96 V (4)

g+ + 1e− � Ag0 E0 = 0.799 V (5)

O3− + H2O + 2e− � NO2

− + 2OH− E0 = 0 V (6)

owever, these components are free to take part in the reductionnder alkaline conditions due to dissociation of H-bonding. It was

nteresting to note that when no base was added to the system,he silver ion was not reduced by the extract at all, implying thathere is an energy barrier for this reduction process. Chou et al.18] have proposed a two-step reaction mechanism to illustratehe role of alkaline ion in the synthesis of nanosized silver col-oids using either formaldehyde or dextrose as the reducing agent.he alkaline ion reacted initially (the first stage) with silver ion toorm Ag2O. Its conversion to silver is instantaneous. In the nexttage, the decrease in the silver ion concentration was gradual andontinuous. Our observed results could also be interpreted in simi-ar lines. The UV–vis absorbance spectrum could be visualized justfter 2 min of addition of all the reacting components of the system.owever, the subsequent increase in the intensity was gradual andontinuous.

The intensity of the peak at 404 nm continuously increased withhe increase of pH up to 11. This was indicative of the progressiveeneration of the nanoparticles. Further increase in the basicity ofhe medium resulted in a red shift of the �max peak position indica-ive of larger particles. Very high alkaline medium may weaken thessociation (H-bonding) even for the starch matrix. This possiblyiminishes the templating potential of the biopolymer to supportmall sized nanoparticles and may be bracketed together with thebserved red-shift for the nanoparticles formed at pH above 11.uthors would like to mention that the samples prepared at pH

1 have been used for various analyses described underneath. Theommon problem of agglomeration of nanoparticles upon storageas not observed in the present study due to their stabilization

y the polymeric chains. This was indicated by the retention ofhe peak sharpness and position at 404 nm with almost no red

Fig. 2. Representative biomolecules present in orange peel.

342 R. Konwarh et al. / Colloids and Surfaces B: Biointerfaces 84 (2011) 338–345

ing as

st

t

Fig. 3. Probable mechanism of reduction of silver ion in starch matrix us

hift (analyzed after 3 months of preparation) on storage at roomemperature.

The X-ray diffractrogram (Fig. 4) of the sample match the litera-ure values of silver nanoparticles. All the distinct peaks at 2� values

Fig. 4. XRD pattern of the starch templated “green” silver nanoparticles.

corbic acid as representative effective reductant present in orange peel.

of about 38◦, 44.5◦, 64.5◦, 77.5◦ and 82◦ representing the 1 1 1, 2 0 0,2 2 0, 3 1 1 and 2 2 2 Braggs reflections of fcc structure of silver, con-firms the formation of Ag nanoparticles [19]. These indexed peakswere compared with the JCPDS data file (4-0787) indicating thepresence of metallic silver in the fcc lattice. The average crystallitesize was determined by Scherrer’s formula:

D = K˛

ˇ cos �(7)

where D is the crystalline domain size, K (the Scherrer con-stant) = 0.9 in this case, ˛ = 15.74 nm is the wavelength of the X-rays,ˇ is the peak angular width and � is the diffraction angle. Thecrystalline average domain size for the polymer assisted silvernanoparticles was found to be 5.29 nm. TEM imaging (Fig. 5) showswell-dispersed spherical particles of 3–12 nm size. It was also inter-esting to note that the highest fraction of particles had a diameterof 6 nm.

3.2. Action at the bio-nano interface

The free radical scavenging, antimicrobial potency and the cyto-

compatibility of the silver nanoparticles were evaluated in thepresent study. Fig. 6 highlights the linear correspondence of thepercent anti-lipid peroxidation and the increasing volume of thesilver nanoparticles. This raises the prospects of incorporating theprepared nanoparticles for development of topical antimicrobial

R. Konwarh et al. / Colloids and Surfaces B: Biointerfaces 84 (2011) 338–345 343

Fig. 5. (a) TEM micrograph and (b) histogram showing the particle size distribution.

Fr

frsli

D

Fto

ig. 6. Anti-lipid peroxidation assay using goat liver tissue homogenate. Resultsepresent mean percent of anti-lipid peroxidation ± S.D. of triplicate determination.

ormulation with added facet of free radical scavenging. The freeadical scavenging attribute of the green silver nanoparticles is

hown in Fig. 7. The percent scavenging of DPPH increases almostinearly with the increase in the concentration of the nanoparticlesn the test samples.

Numerous anti-oxidants of the orange peel act synergistically.uring the preparation of the nanoparticles, these biomolecules

ig. 7. DPPH scavenging by (a) 20 �l, (b) 50 �l, (c) 100 �l, (d) 150 �l and (e) 200 �l ofhe starch templated silver nanoparticles. Results represent mean scavenging ± S.D.f triplicate determination.

Results represent mean size ± S.D. of observations under three different fields.

are pooled into the system. These may get adsorbed onto the activesurface of the noble metal nanoparticles reported here. With a highsurface area to volume ratio and an ambient electrostatic field withanti-oxidant bio-moities on the surface, these silver nanoparticlesdevelop a high tendency to interact with and reduce DPPH likespecies. Interestingly, Paul et al. [13] have recently documentedanti-oxidant activity for iron oxide particles as a surface dependentproperty. The nanochemistry involved in this free radical scav-enging attribute of nanoparticles needs further investigation butnevertheless, this facet of the noble metal nanoparticles can haveprofound impact in the domain of nanomedicine.

Nano-silver targets a broad spectrum of gram-negative andgram-positive bacteria including antibiotic-resistant strains [20].Different mechanisms have been put forward to explain the anti-microbial action of Ag nanoparticles. These include alteration of thecell membrane permeability primarily by immediate dissipation ofthe proton motive force and interaction with sulphur and phospho-rus containing macromolecules like DNA and thereby affecting thereplication machinery. In yet another work, it has been suggestedthat silver nanoparticles may modulate the phosphotyrosine pro-file of putative bacterial peptides. This could affect cellular signalingand therefore inhibit the growth of bacteria [21]. Surface to volumeratio and the crystallographic structure of the nanoparticles dictatetheir anti-mirobial potency. It has been demonstrated that silverreactivity is favored by {1 1 1} facets [22]. Generally, spherical silvernanoparticles (generally with cubooctahedral, multiple-twinneddecahedral, or quasi-sperical morphology) have {1 0 0} facets alongwith a small percentage of {1 1 1} facets [23]. Spherical silvernanoparticles with {1 1 1} facets (which is also a characteristicfeature of the nanoparticles reported here) attach directly to thebacterial surface of the cell membrane and are located inside bac-teria [24]. Shahverdi et al. [25] have reported that the antibacterialactivities of penicillin G, amoxicillin, erythromycin, clindamycin,and vancomycin were increased in the presence of Ag-NPs againstStaphylococcus aureus and Escherichia coli. The antimicrobial actionof the prepared nanoparticles was tested against Bacillus subtilis.The antimicrobial potency of the silver nanoparticles was alsoinvestigated in presence of the antibiotic-rifampicin (Rif) in thewells of the agar plates. Rifampicin inhibits DNA-dependent RNApolymerase in bacterial cells, especially those that are gram stainpositive, by binding its beta-subunit, thus preventing transcriptionand subsequent translation.

The mean zones of inhibition observed in the anti-bacterial test

have been represented in the Table 1. Fig. 8 depicts representativeculture plates showing antibacterial activity. In the present studythe antibiotic (20 �g) i.e. the positive control produced a meanzone of inhibition of 20 mm (Fig. 8a1). The smaller zone of inhi-bition (14 mm) produced by the polymer templated nanoparticles

344 R. Konwarh et al. / Colloids and Surfaces B: Biointerfaces 84 (2011) 338–345

F tibiotic, rifampicin (Rif) and silver nanoparticles (a) both at different concentrations, (b)w e antibiotic at the peripheral wells. The central well contains Rif as positive control.

(ontdgfstpscTpatosaf

cpe2tacbtintcvnicTmpoug

ig. 8. Representative culture plate showing antibacterial test: Combination of anith constant concentration of the nanoparticles and increasing concentration of th

Fig. 8a2) may be attributed to their lesser diffusion than the antibi-tic in the agar plates. Next, the synergy of the antibiotic with theanoparticles was investigated. The concentration of the nanopar-icle was progessively increased while that of the antibiotic wasecreased in the wells. It is very much likely that the antibioticets adsorbed onto the active surface of the nanoparticles. Thisurther reduces the mobility of the starch templated nanoparticlesince diffusion is indirectly proportional to the mass. Even thoughhe coupled system was expected to have a higher antimicrobialotency, the latter never reached that of the positive control. Irre-pective of the increment in the rifampicin concentration, all theombinations produced a zone of inhibition of 17 mm (Fig. 8b).hese observations suggest that a combination of antibiotic and theolymer templated nanoparticle may not merely show synergisticntimicrobial potency. Also, the probable interaction of the starchemplate and the agar medium might further restrict the diffusionf the particles. The interaction of the partners and the overall diffu-ion bears a relation with their respective concentrations, availablective surface area and nanoscale surface chemistry, which meritsurther investigation.

In our report, the cytotoxicity of the nanoparticles on THP-1ells was examined in terms of the former’s effect on the latter’sroliferation by the MTT assay. Untreated cells as well as cellsxposed to different concentrations of the silver nanoparticles for4 h were subjected to the MTT assay for cell viability determina-ion. MTT assay, first described by Mosmann [26], is based on thebility of mitochondrial dehydrogenase enzyme from viable cells toleave the tetrazolium rings of the pale yellow MTT and form darklue formazan crystals, largely impermeable to cell membranes,hus resulting in its accumulation within healthy cells. Solubil-sation of the cells results in the liberation of the crystals. Theumber of surviving cells is directly proportional to the level ofhe formazan product formed that can be quantified photometri-ally. After 24 h post treatment, the THP-1 cells showed excellentiability (almost 100%) even up to 98.7 �M concentration of theanoparticles (Fig. 9). These results indicate that the phytochem-

cals present in the aqueous orange extract provide a non-toxicoating on the nanoparticles. It is however, critical to mention thatHP-1 cells have Fc and C3b receptors and lack surface and cytoplas-

ic immunoglobulins. The response of a single cell-line towards a

articular nanomaterial is not representative of all the plausibleutcome of the events at the bio-nano material interface. The safetility of the reported silver NPs demands in vivo assessments andreater risk analysis.

Fig. 9. MTT assay for cytocompatibility of the silver nanoparticles with THP-1 cellline. Results represent mean cell percent viability ± S.D. of triplicate determination.

4. Conclusions

The reported system highlights a common by-product of thefood sector i.e. orange peel that normally finds applications in thecosmetic industry can be useful even in the domain of nanotech-nology. The biopolymer templated silver nanoparticles synthesisedin the ambient conditions shows considerable potential for appli-cations in a number of niches. These nanoparticles with free radicalscavenging, biocompatibility and antimicrobial potency may beexploited for various biomedical applications including develop-ment of catheters, topical antimicrobial gel formulations and soon.

Acknowledgements

Mr. Rocktotpal Konwarh sincerely acknowledges the receiptof his Senior Research Fellowship from the Department of

Biotechnology, New Delhi. RSIC, NEHU, Shillong is thankfullyacknowledged for the TEM imaging. Mr. Gautam Das is thankfullyacknowledged for his suggestions offered during the manuscriptpreparation.

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ppendix A. Supplementary data

Supplementary data associated with this article can be found, inhe online version, at doi:10.1016/j.colsurfb.2011.01.024.

eferences

[1] X. Chen, H.J. Schluesener, Toxicol. Lett. 176 (2008) 1–12.[2] V.K. Sharma, R.A. Yngard, Y. Lin, Adv. Colloid Interface Sci. 145 (2009) 83–96.[3] H. Korbekandi, S. Iravani, S. Abbasi, Crit. Rev. Biotechnol. 29 (4) (2009) 279–306.[4] K.F. Schmidt, “Green nanotechnology: It’s easier than you think” Presentation

given at Green Nanotechnology Event hosted by the Projects on EmergingNanotechnologies at the Woodrow Wilson International Center for Scholar,(2007).

[5] R. Konwarh, D. Kalita, C.L. Mahanta, M. Mandal, N. Karak, Appl. Microbiol.Biotechnol. 87 (6) (2010) 1983–1992.

[6] S. Li, T. Lambros, Z. Wang, R. Goodnow, C.-T. Ho, J. Chromatogr. B 846 (2007)291–297.

[7] R. Guimarães, L. Barros, J.C.M. Barreira, M.J. Sousa, A.M. Carvalho, I.C.F.R. Fer-reira, Food Chem. Toxicol. 48 (2010) 99–106.

[8] R. Konwarh, N. Karak, S.K. Rai, A.K. Mukherjee, Nanotechnology 20 (2009)225107 (10 pp).

[9] J. Jain, S. Arora, J.M. Rajwade, P. Omray, S. Khandelwal, K.M. Paknikar, Mol.Pharm. 6 (5) (2009) 1388–1401.

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[

[

: Biointerfaces 84 (2011) 338–345 345

10] G. Mohammad, V.K. Mishra, H.P. Pandey, Dig. J. Nanomater. Bios. 3 (4) (2008)159–162.

11] K. Vasilev, V. Sah, K. Anselme, C. Ndi, M. Mateescu, B. Dollmann, P. Martinek, H.Ys, L. Ploux, H.J. Griesser, Nano Lett. 10 (2010) 202–207.

12] T.K. Mandal, S.N. Chatterjee, Radiat. Res. 83 (2) (1980) 290–302.13] S. Paul, J.P. Saikia, S.K. Samdarshi, B.K. Konwar, J. Magn. Magn. Mater. 321 (2009)

3621.14] N. Mahajan, V. Dhawan, G. Sharma, S. Jain, D. Kaul, J. Hum. Hypertens. 22 (2008)

141–143.15] J. Zhao, A.O. Pinchuk, J.M. Mcmahon, S. Li, L.K. Ausman, A.L. Atkinson, G.C.

Schatz, Acc. Chem. Res. 41 (12) (2008) 1710–1720.16] J. Zhao, X.Y. Zhang, C.R. Yonzon, A.J. Haes, R.P. Van Duyne, Nanomedicine 1

(2006) 219–228.17] L. Suber, W.R. Plunkett, Nanoscale 2 (2010) 128–133.18] K.-S. Chou, Y.-C. Lu, H.-H. Lee, Mater. Chem. Phys. 94 (2005) 429–433.19] O.L.A. Monti, J.T. Fourkas, D.J. Nesbitt, J. Phys. Chem. B 108 (2004) 1604.20] S.L. Percival, P.G. Bowler, J. Dolman, Int. Wound J. 4 (2007) 186–191.21] S. Shrivastava, T. Bera, A. Roy, G. Singh, P. Ramachandrarao, D. Dash, Nanotech-

nology 18 (2007) 225103 (9pp).22] D.W. Hatchett, S.J. Henry, Phys. Chem. 100 (1996) 9854–9859.

23] B. Wiley, Y. Sun, B. Mayers, Y. Xia, Chem. Eur. J. 11 (2005) 454–463.24] J.R. Morones, J.L. Elechiguerra, A. Camacho, K. Holt, J.B. Kouri, J.T. Ramirez, M.J.

Yacaman, Nanotechnology 16 (2005) 2346–2353.25] A.R. Shahverdi, A. Fakhimi, H.R. Shahverdi, S. Minaian, Nanomed-Nanotechnol.

(2) (2007) 168–171.26] T. Mosmann, J. Immunol. Methods 65 (1–2) (1983) 55–63.