Agar-lignin-silver

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Preparation and characterization of
agar/lignin/silver nanoparticles composite
films with ultraviolet lig....
Article in Food Hydrocolloids · October 2017
DOI: 10.1016/j.foodhyd.2017.05.002
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Food Hydrocolloids 71 (2017) 76e84
Contents lists available at ScienceDirect
Food Hydrocolloids
journal homepage: www.elsevier.com/locate/foodhyd
Preparation and characterization of agar/lignin/silver nanoparticles
composite films with ultraviolet light barrier and antibacterial
properties
Shiv Shankar a , Jong-Whan Rhim b, *
a Department of Food Engineering and Bionanocomposite Research Institute, Mokpo National University, 61 Dorimri, Chungkyemyon, Muangun, 534-729
Jeonnam, Republic of Korea
b Center for Humanities and Sciences, and Department of Food and Nutrition, Kyung Hee University, 26 Kyungheedae-ro, Dongdaemun-gu, Seoul 120-701,
Republic of Korea
article
info
abstract
Article history:
Received 3 April 2017
Received in revised form
2 May 2017
Accepted 2 May 2017
Available online 3 May 2017
Keywords:
Agar
Lignin
Silver nanoparticles
Composite film
Antibacterial activity
Lignin was used to prepare silver nanoparticles (AgNPs), and they were incorporated into agar-based
films. The composite films were characterized using UVevisible spectroscopy, FE-SEM, FTIR, XRD, and
TGA. The color, mechanical, water vapor barrier, and antibacterial properties of the composite films were
also evaluated. The composite films exhibited characteristic light absorption peaks at 250e300 nm and
430 nm due to the lignin and AgNPs, respectively. The interaction of lignin and AgNPs with agar
biopolymer was analyzed using FTIR. XRD results showed the characteristic peaks of crystalline AgNPs.
Incorporation of lignin and AgNPs increased the mechanical, UV-light barrier, and water vapor barrier
properties of the composite films. The loading of 1 wt% of AgNPs was the optimum concentration to
improve the mechanical and water vapor barrier properties of the films. The films containing AgNPs
exhibited antibacterial activity against food-borne pathogenic bacteria, Escherichia coli and Listeria
monocytogenes.
© 2017 Elsevier Ltd. All rights reserved.
1. Introduction
There have been serious concerns about the contamination of
food by micro-organisms which can reduce the shelf-life of food as
well as increase the risks of various food-borne infections causing
serious illness to human beings (Devlieghere, Vermeiren, &
Debevere, 2004). Traditionally, various physical and chemical
preservation methods have been used in the food industry to
reduce the spoilage of food, maintain food quality, and extend the
shelf-life of food products. The most common method to control
the growth of micro-organisms on the surface of the food is to
apply antimicrobial agents or to dip the food materials into an
antimicrobial solution (Sung et al., 2013). However, concerns on the
diffusion of such antimicrobial agents to the inside of foodstuffs
have limited the use of this technology. Also, recently increased
consumer demands for the minimally processed and ready-to-eat
fresh foods have inspired researchers to develop new
* Corresponding author.
E-mail address: jwrhim@khu.ac.kr (J.-W. Rhim).
technologies for securing food safety and maintaining food quality
(Bhat, 2013). Packaging materials with antimicrobial function have
been recognized as one of the most promising active packaging
systems for extending shelf-life of food, maintaining food safety
and quality, and improving storage stability by destroying or
inhibiting the spoilage and pathogenic microorganisms (Falguera,
Quintero, Jimenez, Mu~noz, & Ibarz, 2011; Han, 2000). Antimicrobial
packaging is often achieved by incorporation of antimicrobial
agents into the packaging system. Some organic and inorganic
materials have been used as antimicrobial agents for the antimicrobial
packaging application (Kanmani & Rhim, 2014; Shankar,
Teng, Li, & Rhim, 2015; Shankar, Teng, & Rhim, 2014). However, a
low thermal stability of organic antimicrobial materials have
limited their broad use in food packaging applications. In contrast,
the high thermostability of inorganic antimicrobial agents such as
metallic nanoparticles has opened the way for their use in the food
packaging industries (Llorens, Lloret, Picouet, Trbojevich, &
Fernandez, 2012).
Among the metallic nanoparticles, silver nanoparticles (AgNPs)
have been most widely used for the preparation of nanocomposite
in the food packaging and biomedical applications due to their high
http://dx.doi.org/10.1016/j.foodhyd.2017.05.002
0268-005X/© 2017 Elsevier Ltd. All rights reserved.

S. Shankar, J.-W. Rhim / Food Hydrocolloids 71 (2017) 76e84 77
surface area, unique optical, magnetic, electric, and catalytic
properties with high thermal stability and broad-spectrum of
antimicrobial activity (Duncan, 2011; Rhim, Park, & Ha, 2013). The
antimicrobial effect of AgNPs is known to be greatly influenced by
the size, shape, and method of preparation of the AgNPs (Lischer
et al., 2011). The AgNPs synthesized by green methods are recognized
as safe for the use in the food packaging applications. For the
green synthesis of AgNPs, various biopolymers and biomolecules
such as gelatin (Kanmani & Rhim, 2014), banana powder
(Orsuwana, Shankar, Wang, Sothornvit, & Rhim, 2016), chitosan
(Huang & Yang, 2004), vitamins (Shankar & Rhim, 2016), aminoacids
(Shankar & Rhim, 2015), and plant extracts (Shankar,
Chorachoo, Jaiswal, & Voravuthikunchai, 2014) have been used.
As one of such biopolymeric materials with high potential for the
preparation of AgNPs, lignin is interesting since it can be used not
only as reducing and stabilizing agent for the AgNPs, but also as a
filler for the preparation of nanocomposite films.
Lignin is a complex phenolic compound which is abundantly
present in the cell wall of plants in association with cellulose and
hemicellulose (Tuomela, Vikman, Hatakka, & Itavaara, 2000).
Although lignin is chemically heterogeneous and structurally
complex material, it possesses multiple functional groups such as
reductive aliphatic hydroxyls, phenolic hydroxyls, and thiols,
which can serve as reducing and stabilizing agents for the synthesis
of silver nanoparticles. Thiols have shown high affinity to
silver nanoparticles, hence facilitating its adsorption and capping
onto particle surface (Laibinis et al., 1991). The polar sulfonate
groups in lignin can also help to disperse the formed nanoparticles
in aqueous solutions. Also, lignin possesses a strong UV
light absorption property (Monties, 1991; Shankar, Reddy, &
Rhim, 2015).
Recently, biopolymers from various natural resources have been
used as eco-friendly packaging materials to substitute the nonbiodegradable,
petroleum-based plastic based packaging materials
(Gimenez, Lopez de Lacey, Perez-Santín, Lopez-Caballero, &
Montero, 2013). Among biopolymers, agar has been widely used for
the preparation of biodegradable packaging films due to its good
film forming property with abundance, renewability, and biocompatibility
(Shankar & Rhim, 2016; Shankar, Reddy et al., 2015;
Shankar, Teng et al., 2014). Agar is a hydrophilic polysaccharide
extracted from the Gelidiaceae and Gracilariaceae families of seaweeds
and mainly composed of alternating repeating units of D-
galactose and 3, 6-anhydro-ß-galactopyranose (Tako, Higa,
Medoruma, & Nakasone, 1999).
Therefore, the present study was aimed to develop agar-based
biodegradable food packaging films incorporated with lignin and
lignin capped silver nanoparticles. The films were characterized
using various analytical techniques. The UV-light barrier, mechanical,
thermal, water vapor barrier and antibacterial properties of the
films were also evaluated.
2. Materials and methods
2.1. Materials
The food grade agar was obtained from Fine Agar Co., Ltd.
(Damyang, Jeonnam, Korea). Glycerol and alkali-soluble low sulfonate
lignin were procured from Sigma-Aldrich Co. (St. Louis, MO,
USA). The lignin was dried at 80 C for 6 h before use. Silver nitrate
(AgNO 3 ), brain heart infusion broth (BHI), and tryptic soy broth
(TSB) were obtained from Duksan Pure Chemicals Co., Ltd. (Ansan,
Gyeonggi-do, Korea). Escherichia coli O157: H7 ATCC 43895 and
Listeria monocytogenes ATCC 15313 were procured from the Korean
Collection for Type Cultures (KCTC, Seoul, Korea).
2.2. Synthesis of agar/lignin/AgNPs composite films
Silver nanoparticles were synthesized using lignin as reducing
and capping agents. 0.12 g of lignin (3 wt% based on agar) was
dispersed in 150 mL of distilled water and stirred for 20 min at
90 C using a magnetic stirrer. Different amounts of AgNO 3 (0.5, 1.0,
1.5, and 2 wt% based on agar) were added dropwise into the lignin
solution and continued heating the mixture for 20 min. 1.2 g of
glycerol (30 wt% based on agar) was added into the mixture as a
plasticizer with vigorous stirring for 10 min followed by addition of
4 g of agar and heated continuously with stirring at 90 C for 20 min
using a hot plate. The fully solubilized film forming solution was
cast evenly onto a leveled Teflon film-coated glass plate
(24 cm 30 cm) and allowed to dry at room temperature for about
48 h. The dried films were peeled off from the plate and conditioned
in a humidity chamber set at 25 C and 50% RH for 48 h
before further analysis. Additionally, neat agar and agar with 3 wt%
of lignin films were prepared by the same procedure except adding
AgNO 3 .
2.3. Characterization of agar/lignin/AgNPs composite films
2.3.1. Surface morphology and optical properties
The surface morphology of the neat agar, agar/lignin, and agar/
lignin/AgNPs composite films was observed using a field emission
scanning electron microscope (FE-SEM, S-4800, Hitachi Co., Ltd.,
Matsuda, Japan). The film samples were attached on the specimen
holder, sputter coated with platinum, and the image was analyzed
at an accelerating voltage of 3 kV.
The optical properties of the composite films were determined
by measuring the absorption of light between 200 and 700 nm
using UVevisible spectrophotometer (Mecasys Optizen POP Series
UV/Vis, Seoul, Korea).
2.3.2. FTIR and XRD
Fourier transform infrared (FTIR) spectra of the composite films
were measured using an attenuated total Reflectance-Fourier
Transform infrared (ATR-FTIR) spectrophotometer (TENSOR 37
spectrophotometer with OPUS 6.0 software, Billerica, MA, USA).
The FTIR spectra were recorded as 32 scans per samples at a resolution
of 4 cm 1 .
X-ray diffraction (XRD) pattern of the films was analyzed using
an X-ray diffractometer (PANalytical X’ pert pro MRD diffractometer,
Amsterdam, Netherlands). The film samples (2.5 cm 2.5 cm)
were placed on a glass slide, and the XRD spectra were recorded
using Cu Ka radiation (wavelength of 0.1541 nm) and a nickel
monochromator filtering wave at 40 kV and 30 mA. The diffraction
pattern was recorded at 2q ¼ 30e80 with a scanning speed of 0.4 /
min at room temperature.
2.3.3. Thermogravimetric analysis
Thermal stability of the composite films was determined using a
thermogravimetric analyzer (Hi-Res TGA 2950, TA Instrument, New
Castle, DE, USA). The film samples were taken in a standard
aluminum pan and heated from 30 to 600 C at the rate of 10 C/
min under a nitrogen flow of 50 mL/min. A derivative form of TGA
(DTG) was obtained and the maximum decomposition temperature
(T max ) of the composite films was obtained from DTG curve.
2.3.4. Surface color and light transmittance
The Hunter color values (L, a, and b) of the composite films were
measured using a Chroma meter (Minolta, CR-200, Tokyo, Japan). A
white color standard plate (L ¼ 97.75, a ¼ - 0.49, and b ¼ 1.96) was
used as a background for the color measurements. The total color
difference (DE) was calculated as follows:

78
S. Shankar, J.-W. Rhim / Food Hydrocolloids 71 (2017) 76e84
qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi
DE ¼ ðDLÞ 2 þðDaÞ 2 þðDbÞ 2
where DL, Da, and Db are differences between each color value of
the standard color plate and film specimen. Five measurements
were taken for each film, and the average values are reported.
The UV-barrier property and transparency of the films was
determined by measuring percent transmittance of light at 280 and
660 nm, respectively.
2.3.5. Thickness and mechanical properties of the composite films
The films were cut into 2.54 cm 15 cm strips using a precision
double blade cutter (model LB.02/A, Metrotec, S.A., San Sebastian,
Spain) and the thickness of films was measured using a digital
micrometer (Digimatic Micrometer, QuantuMike IP 65, Mitutoyo,
Japan) with an accuracy of 0.001 mm. Five random locations around
each film sample were measured, and the average thickness was
used for the calculation of tensile strength.
The mechanical properties of the composite films in terms of
tensile strength (TS), elongation at break (EAB), and elastic modulus
(EM) were determined using Instron Universal Testing Machine
(Model 5565, Instron Engineering Corporation, Canton, MA, USA) in
a tensile mode with an initial grip separation and a cross-head
speed set at 50 mm and 50 mm/min, respectively (Shankar &
Rhim, 2016). The TS was calculated in MPa by dividing the
maximum load (N) by the initial cross-sectional area (m 2 ) of the
film sample. The EAB was calculated in % by dividing the extension
at the rupture of the film by the initial length of the film (50 mm)
and multiplied by 100. The EM was determined in GPa from the
slope of a linear portion of the stress-strain curve.
2.3.6. Water vapor permeability and water contact angle of
composite films
The water vapor permeability (WVP) of films was determined by
the standard method of ASTM E96-95 with modification
(Gennadios, Weller, & Goodings, 1994). The WVP measuring cup
was filled with 18 mL of distilled water and placed film sample
(7.5 cm 7.5 cm) on the top of the cup and sealed tightly to prevent
the leakage of water vapor. The assembled WVP cup was weighed
and subsequently placed in a controlled environmental chamber
set at 25 C and 50% RH. Weight change of the cup was determined
every 1 h for 8 h. The water vapor transmission rate (WVTR; g/m 2 s)
of the film was calculated by using the slope of the steady-state
(linear) portion of weight loss versus time plot. Then, the WVP
(g.m/m 2 .s.Pa) of the film was calculated as follows:
ðWVTR LÞ
WVP ¼
Dp
where L was the mean thickness of the film (m) and Dp was water
vapor partial pressure difference (Pa) across the film, which was
calculated by the method of Gennadios et al. (1994).
Surface hydrophobicity of the film was evaluated by measuring
the water contact angle (WCA) of the film using a water contact
angle analyzer (model Phoenix 150, Surface Electro-Optics Co., Ltd.,
Kunpo, Korea). The film was cut into 3 cm 10 cm strips and placed
on the horizontal movable stage (black Teflon coated steel,
7cm 11 cm) fitted with the WCA analyzer. About 10 mL of water
was dropped on the surface of the film using a microsyringe, and
the contact angle on both sides of the water droplet was measured.
Five measurements were taken for each sample, and average values
were presented.
2.3.7. Water solubility, and swelling ratio of composite films
The water solubility (WS) of the film samples was determined as
the percentage of dissolved dry matter after immersion in water.
Three randomly selected specimens of each type of film
(3 cm 3 cm) were first dried at 60 C for 24 h to determine the
initial dry matter (W 1 ). Each film was immersed into 30 mL of
distilled water in a 50 mL beaker with a gentle stirring for 24 h. The
film samples were removed after 24 h and dried in a drying oven at
105 C for 24 h to determine the undissolved final dry weight (W 2 ).
The WS of the sample was calculated as follows:
WS ¼ W 1 W 2
W 1
100
The swelling ratio (SR) of the films was determined gravimetrically.
Pre-weighed specimens (2.5 cm 5 cm) were immersed in
distilled water for 1 h, and then the film samples were removed
from the water and weighed after removing the surface water with
blotting paper. The percent SR of the films was calculated as
follows:
SRð%Þ ¼ W 2 W 1
W 1
100
where W 1 and W 2 are the weight of the film samples before and
after soaking in the water, respectively.
2.3.8. Antibacterial activity
The antibacterial activity of composite films was examined
against food-borne pathogenic Gram-positive bacteria,
L. monocytogenes and Gram-negative bacteria, E. coli (Shankar &
Rhim, 2016). The test bacteria were aseptically inoculated in the
TSB and BHI broth, respectively, and subsequently incubated at
37 C for 16 h. The inoculum was diluted, and 200 mL of diluted
inoculum (10 8 e10 9 CFU/mL) was aseptically transferred to 50 mL of
TSB and BHI broth containing 100 mg film samples and incubated at
37 C for 15 h with mild shaking. The samples were taken out at 3, 6,
9, 12, and 15 h of incubation, and the viable cell count was determined
by diluting and plating the samples on agar plates. Broth
without film was used as a positive control. Antimicrobial tests
were performed in triplicate with individually prepared films.
2.3.9. Statistical analysis
Measurement of each property of the films was performed in
triplicate with individually prepared film samples as the replicated
experimental units, and the results were presented as mean
values ± standard deviations (SD). One-way analysis of variance
(ANOVA) was performed, and the significance of each mean property
value was determined (p < 0.05) with Duncan’s multiple range
tests using the SPSS statistical analysis computer program for
Windows (SPSS Inc., Chicago, IL, USA).
3. Results and discussion
3.1. Preparation of AgNPs
The lignin solution was yellow, but it changed to dark brown
after the addition of silver nitrate solution. The change in color of
the solution indicated the formation of AgNPs, which was formed
through the reduction of Ag ions by the lignin. The solution
exhibited the maximum absorption peak around 420 nm (data not
shown), which is also evidence for the formation of AgNPs (Shankar
& Rhim, 2015). The color intensity of the solution was dependent on
the amount of silver nitrate used. Lignin possesses multiple functional
groups such as aliphatic hydroxyl, phenolic hydroxyls, and

S. Shankar, J.-W. Rhim / Food Hydrocolloids 71 (2017) 76e84 79
thiols, which has potential as reducing and capping agents during
silver nanoparticle synthesis (Hu & Hsieh, 2016). A thiol group of
lignin has a strong affinity to silver nanoparticles. Hence it facilitated
to adsorb AgNPs and to stabilize AgNPs (Laibinis et al., 1991).
The polar sulfonate groups of lignin can also help to disperse the
formed nanoparticles in aqueous solutions.
3.2. Characterization of agar/lignin/AgNPs composite films
3.2.1. Morphology and optical properties
All the films prepared were free standing, intact, and smoothsurfaced
without any apparent defects. The microstructure of the
composite films at the surface and cross-section was observed using
FE-SEM as shown in Fig. 1. As expected, the surface of the neat
agar and agar/lignin composite films was smooth and compact.
However, the surface of composite films incorporated with AgNPs
was slightly rough compared with the control films. As shown in
both the surface and the cross-sectional views, the AgNPs were
uniformly distributed through the agar/lignin/AgNPs composite
films. The size of AgNPs increased with increase in the concentration
of the precursor of AgNPs, AgNO 3 . This result indicated that the
size of the AgNPs was greatly influenced by the concentration of
silver precursor.
Light absorption characteristics of the films were evaluated by
measuring light absorption spectra in the wavelength of
200e700 nm as shown in Fig. 2a. The neat agar film did not show
any light absorption peak. On the other hand, lignin incorporated
films exhibited broad maximum peaks in the range of 250e300 nm,
which was attributed to the chromophore groups in the lignin
(Chaochanchaikul, Jayaraman, Rosarpitak, & Sombatsompop,
2012). In addition to the peak of lignin, the agar/lignin/AgNPs
composite films exhibited the light absorption peak at around
430 nm, which was due to the surface Plasmon resonance of AgNPs
in the composite films (Shankar, Chorachoo et al., 2014; Shankar,
Jaiswal, Selvakannan, Ham, & Rhim, 2016).
3.2.2. FTIR and XRD
The FTIR spectra of the composite films were shown in Fig. 2b.
The neat agar film showed broad bands at 3330 cm 1 and 2930,
which were due to the stretching of hydroxyl (OeH) group and
stretching of CeH that was associated with the ring of methane
hydrogen atoms, respectively (Tako et al., 1999). The peak at
1640 cm 1 was attributed to the stretching vibration of the peptide
bond formed by an amine (NH) and acetone groups in agar (Kumar,
Negi, Choudhary, & Bhardwaj, 2014). The peak at 1372 cm 1 was
attributed to the ester sulfate group of agar (Volery, Besson, &
Schaffer-lequart, 2004). The bands at 1040 and 930 cm 1 were
associated with the CO stretching group of 3,6-anhydrogalactose
(Shankar, Teng et al., 2014). The peak around 890 cm 1 was due
to the CeH residual carbon of b-galactose. The lignin exhibit
characteristic peak around 1515 cm 1 , however, there was no such
peak observed when lignin was composite with agar (Oun & Rhim,
2016). It was presumably because the peak of lignin was masked by
the peak of agar between 1500 and 1650 cm 1 . All the FTIR spectra
of composite films were similar to those of agar film, suggesting
that there were no chemical bonds formed among agar, lignin, and
AgNPs.
The crystalline nature of lignin and AgNPs in the composite
films was tested using the X-ray diffraction (XRD) analysis. The
results of XRD of the agar, agar/lignin, and agar/lignin/AgNP 1.0 films
were shown in Fig. 2c. The XRD patterns of agar/AgNPs composite
films exhibited distinctive diffraction peaks at 2q values around 32,
48, and 68 corresponding to the (111), (200), and (220) lattice
planes of AgNPs in the composite films, which were attributed to
the formation of crystalline AgNPs (Martínez-Casta~non, Ni~no-
Martinez, Martínez-Gutierrez, Martínez-Mendoza, & Ruiz, 2008).
However, the neat agar and agar/lignin composite film did not
show any diffraction peak in the 2q range of 30e80 .
3.2.3. Thermogravimetric analysis
The TGA and DTG thermograms of the composite films were
shown in Fig. 3. TGA curves showed the weight loss pattern of the
films and DTG curves clearly showed the maximum decomposition
temperature at each step of thermal decomposition. All the films
showed multi-step thermal degradation. The first step of thermal
degradation occurred at 80e120 C, which was attributed to the
evaporation of moisture in the composite films (Shankar, Teng et al.,
2014). The subsequent thermal degradation patterns were varied
depending on the type of films. The main steps of thermal degradation
were observed between 210 and 325 C which was due to
the decomposition of glycerol (a plasticizer) and polymers (agar
and lignin) (El-Hefian, Nasef, & Yahaya, 2012; Martucci &
Ruseckaite, 2010). Agar/lignin/AgNPs composite films exhibited
higher degradation temperature compared with the neat agar and
agar/lignin composite films. The residual char content at 600 Cof
the neat agar film was 10.5%, and it increased to 24.8, 25.6, and
24.3% in the agar/lignin, agar/lignin/AgNP 1.0 , and agar/lignin/
AgNP 2.0 composite films, respectively. The increased thermostability
of the composite films was mainly due to the thermostable
AgNPs.
Fig. 1. FE-SEM micrographs of surface and cross-section of the composite films.
3.2.4. Apparent color and light transmittance
The apparent color and light transmittance of the films were
presented in Table 1. The neat agar film was transparent without
any color tint, but it became yellow after addition of lignin. The

80
S. Shankar, J.-W. Rhim / Food Hydrocolloids 71 (2017) 76e84
(a)
(b)
(c)
Fig. 2. (a) UVevisible, (b) FTIR, and (c) XRD spectra of the composite films.
surface color of agar/lignin/AgNPs composite films exhibited dark
brown. The intensity of color from yellow to brown increased with
the increase in the concentration of silver nitrate. The Hunter L-
value (lightness) decreased significantly (p < 0.05) with increasing
AgNPs content. However, Hunter-a and b and total color difference
(DE) values of the composite films were higher than those of the
neat agar film. This result indicates that the lightness of the film
decreased with increased redness and yellowness after blending
with lignin and AgNPs. Similar results of decreased L values and
increased a and b values were found when lignin (Shankar, Reddy
et al., 2015) and AgNPs (Shankar & Rhim, 2015) were incorporated
into the agar films.
The light transmission characteristics of the composite films
were measured using a UV/Vis spectrophotometer at 280 and
660 nm (Table 1). The T 660 (a measure of transparency) of the neat
agar film were 88.8 ± 0.5% indicating the high transparency of the
film. The T 660 decreased slightly to 85.5 ± 1.3 after incorporation of
lignin and decreased significantly after composite formation with
AgNPs. On the other hand, the light transmittance at 280 nm (T 280, a
measure of UV-barrier) decreased significantly (p < 0.05) after
incorporation of lignin and AgNPs. The nanocomposite film incorporated
with lignin and AgNPs prevented UV-light passage almost
completely. Since lignin is composed of a complex aromatic polymer
of aromatic alcohols, it absorbs light in the ultraviolet region
due to the presence of aromatic rings in its structure (Monties,
1991). This result indicated that the composite film had a UVscreening
property with a slight sacrifice of the transparency. The
composite film has a high potential for the UV-screening food
packaging film with the see-through property.
3.2.5. Mechanical properties
The mechanical properties of the films determined in term of
the tensile strength (TS), elongation at break (EAB), and elastic
modulus (EM) and the results were presented in Table 2. The
thickness of the biopolymer films increased after incorporation of
lignin and AgNPs. The increase in the thickness of the composite
films was mainly due to the increased solid content by the lignin
and AgNPs (Shankar, Teng et al., 2014). The mechanical properties
were greatly influenced by the addition of lignin and AgNPs. The TS
of neat agar film was 40.3 ± 4.0 MPa, which was increased to
44.1 ± 3.6 MPa after blending with lignin and it increased further
after blending with AgNPs. The TS of agar/lignin/AgNPs composite
films was also influenced by the concentration of AgNO 3 . The TS of
the composite films increased up to 1 wt% of AgNO 3 , then
decreased when a higher concentration of AgNO 3 was used. Agar/
lignin/AgNP 1.0 exhibited the highest TS (49.7 ± 3.3 MPa) among the
composite films. The increase in the TS of the composite films
might be due to a high compatibility between the nanofiller and
agar biopolymer. As shown in the SEM images (Fig. 1), the lignin
and silver nanoparticles were dispersed well in the agar
biopolymer, indicating they were highly compatible. Since the
lignin served as a reducing as well as capping agent for the silver
nanoparticles, the interaction between AgNPs stabilized by the
lignin and agar polymer matrix was presumably increased to form
strong composite films. The effectiveness of the reinforcement and
dispersion of nanofillers in the polymer matrix functions controlling
role for the effective stress transfer at the interface of the
matrix and the filler and ultimately increase the tensile strength of
the polymeric biocomposite materials (Ahmed, Arfat, Castro-

S. Shankar, J.-W. Rhim / Food Hydrocolloids 71 (2017) 76e84 81
(a)
Table 2
Tensile properties of agar, agar/lignin, and agar/lignin/AgNPs composite films. 1)
Film Thickness (mm) TS (MPa) EAB (%) EM (GPa)
Agar 45.1 ± 2.3 a 40.3 ± 4.0 a 19.4 ± 4.2 a 1.39 ± 0.23 a
Agar/lignin 55.9 ± 2.6 a 44.1 ± 3.6 b 16.1 ± 4.3 a 1.48 ± 0.13 ab
Agar/lignin/AgNP 0.5 54.9 ± 7.4 b 45.5 ± 4.6 bc 16.2 ± 5.1 a 1.49 ± 0.15 ab
Agar/lignin/AgNP 1.0 54.3 ± 2.9 a 49.7 ± 3.3 d 16.7 ± 4.7 a 1.60 ± 0.08 c
Agar/lignin/AgNP 1.5 57.2 ± 1.9 a 47.0 ± 3.5 c 16.2 ± 3.4 a 1.52 ± 0.23 ab
Agar/lignin/AgNP 2.0 53.0 ± 1.4 a 45.6 ± 2.2 bc 16.7 ± 2.9 a 1.50 ± 0.13 ab
1) Each value is the mean of three replicates with the standard deviation. Any two
means in the same column followed by the same letter are not significantly
(p > 0.05) different by Duncan’s multiple range test.
the EAB of composite films was slightly decreased after incorporation
of lignin and AgNPs, though the decrease was not significantly
different (p > 0.05). The results showed that the strength and
stiffness of the composite films increased without sacrificing the
flexibility of the films.
(b)
Fig. 3. (a) TGA and (b) DTG of the composite films.
Aguirre, & Auras, 2016; Díez-Pascual & Díez-Vicente, 2014).
Shankar, Reddy et al. (2015) also reported that the TS of agar film
increased with increase in the concentration of lignin up to 3 wt%.
Chen et al. (2009) also reported that the addition of lignin into
chitosan increased the tensile strength of the film. The effect of
lignin and AgNPs on the EM of the film was similar to the TS. The
EM of agar film increased after lignin incorporation, and it
increased further with the incorporation of AgNPs up to 1 wt%, then
started to decrease with a higher concentration of AgNPs. The
decreased mechanical strength of the composite films above 1 wt %
of AgNPs might be due to aggregation of AgNPs. On the contrary,
3.2.6. Water vapor permeability (WVP) and water contact angle
(WCA)
The WVP of composite films were presented in Table 3. The WVP
of the neat agar film was 1.99 ± 0.26 10 9 g m/m 2 .Pa.s, which was
decreased to 1.85 ± 0.13 10 9 g m/m 2 .Pa.s after incorporation of
lignin. The WVP of the film decreased further when AgNPs were
incorporated, however, the degree of decrease was dependent on
the content of AgNPs. The WVP of the composite films decreased
down to 1.56 ± 0.11 10 9 g m/m 2 .Pa.s when 1 wt% of AgNPs was
incorporated, then increased with further increase in the AgNPs.
The decreased WVP of the nanocomposite films was mainly due to
the increased tortuous path for water vapor diffusion by the
impermeable nanoparticles distributed in the polymer matrix (Yu,
Yang, Liu, & Ma, 2009). Also, the compatibility between lignin and
agar might help to form a strong intermolecular interaction and to
reduce water vapor penetration through the composite films. The
increase in the WVP of the composite films incorporated with more
than 1 wt% was probably due to the aggregation of the nanoparticles
at a higher concentration as observed in the SEM result.
This result also indicates 1 wt% of AgNPs is optimum to increase
water vapor barrier property of the composite film. It has
frequently been observed that the incorporation of nanofillers
decreased the WVP of the composite films within a certain level of
concentration, but the effect was reversed when the concentration
of nanofiller was higher than the optimum level (Shankar & Rhim,
2016; Shankar, Reddy et al., 2015).
The hydrophobicity and wettability of the films are usually
evaluated by the water contact angle (WCA). The WCA of the test
films were also shown in Table 3. The WCA of the neat agar film was
54.3 ± 1.3 , it increased to 55.6 ± 0.7 and 59.4 ± 0.6 for the agar/
lignin and agar/lignin/AgNP 1.0 composite films, respectively. The
hydrophobicity of the composite film increased due to the hydrophobicity
of the AgNPs. It was interesting to note that the WCA of
the agar/lignin/AgNPs composite films also increased with the
Table 1
Apparent color and light transmittance of agar, agar/lignin and agar/lignin/AgNPs biocomposite films. 1)
Film L a b DE T 660 (%) T 280 (%)
Agar 94.81 ± 0.29 d 0.99 ± 0.02 a 3.67 ± 0.06 a 1.05 ± 0.09 a 88.8 ± 0.5 b 52.9 ± 1.7 b
Agar/lignin 84.30 ± 1.74 c 0.98 ± 0.41 a 23.57 ± 2.08 b 23.08 ± 2.62 b 85.5 ± 1.3 b 0.1 ± 0.0 a
Agar/lignin/AgNP 0.5 56.19 ± 5.16 b 5.51 ± 1.58 b 29.07 ± 3.18 d 47.05 ± 2.43 c 60.6 ± 7.7 a 0.1 ± 0.0 a
Agar/lignin/AgNP 1.0 49.11 ± 2.13 a 17.81 ± 0.31 c 28.73 ± 1.75 cd 55.38 ± 0.97 d 65.8 ± 7.0 a 0.1 ± 0.0 a
Agar/lignin/AgNP 1.5 46.05 ± 5.55 a 17.86 ± 0.31 c 26.11 ± 4.35 bcd 57.02 ± 2.81 de 66.6 ± 5.3 a 0.1 ± 0.0 a
Agar/lignin/AgNP 2.0 43.77 ± 1.39 a 24.43 ± 3.35 d 24.08 ± 1.49 bc 60.42 ± 2.05 e 68.2 ± 0.9 a 0.1 ± 0.0 a
1) Each value is the mean of three replicates with the standard deviation. Any two means in the same column followed by the same letter are not significantly (p > 0.05)
different by Duncan’s multiple range test.

82
S. Shankar, J.-W. Rhim / Food Hydrocolloids 71 (2017) 76e84
Table 3
Water vapor permeability (WVP), water contact angle (WCA), water solubility (WS), and swelling ratio (SR) of agar, agar/lignin, and agar/lignin/AgNPs composite films. 1)
Film WVP (10 9 g m/m 2 Pa s) WCA (degree) WS (%) SR (%)
Agar 1.99 ± 0.26 d 54.3 ± 1.3 a 27.3 ± 0.8 bc 2785.6 ± 343.2 c
Agar/lignin 1.85 ± 0.13 c 55.6 ± 0.7 ab 31.2 ± 0.6 d 2587.4 ± 483.8 bc
Agar/lignin/AgNP 0.5 1.71 ± 0.09 bc 58.3 ± 0.8 c 28.4 ± 0.9 c 2268.7 ± 453.9 bc
Agar/lignin/AgNP 1.0 1.56 ± 0.11 a 59.4 ± 0.6 d 25.5 ± 0.8 ab 1873.8 ± 559.4 ab
Agar/lignin/AgNP 1.5 1.62 ± 0.09 a 56.7 ± 1.2 b 24.0 ± 0.5 a 1428.1 ± 289.6 a
Agar/lignin/AgNP 2.0 1.67 ± 0.07 ab 54.2 ± 0.8 a 23.6 ± 1.1 a 1398.7 ± 195.4 a
1) Each value is the mean of three replicates with the standard deviation. Any two means in the same column followed by the same letter are not significantly (p > 0.05)
different by Duncan’s multiple range test.
concentration of AgNPs up to 1 wt%, then decreased. The decrease
in WCA at higher concentration of AgNPs might be due to the
increased surface roughness of the films (Orsuwana et al., 2016).
3.2.7. Water solubility and swelling ratio
The results of water solubility (WS) and swelling ratio (SR) of the
composite films were also shown in Table 3. The agar/lignin composite
film showed the highest WS among the tested films. The WS
of the composite films increased after incorporation of lignin, but it
decreased when AgNPs were incorporated. The WS of the composite
films decreased linearly with the increasing concentration of
AgNPs. The higher WS of agar/lignin composite films compared to
the neat agar films might be due to the higher water solubility of
lignin than agar. On the other hand, the decrease in WS of AgNPs
incorporated films might be due to the presence of water-insoluble
AgNPs in the composite films.
The SR of the composite films was also greatly affected by the
incorporation of the fillers. The neat agar film exhibited the highest
SR (2785.6 ± 343.2%), which was reduced to 2587.4 ± 483.8% in
agar/lignin composite films. The SR of AgNPs incorporated films
was decreased linearly with the increase in the concentration of
AgNPs.
3.2.8. Antibacterial activity
The antibacterial activity of agar, agar/lignin, and agar/lignin/
AgNPs composite films was tested against food-bore pathogenic
Gram-positive (L. monocytogenes) and Gram-negative (E. coli) bacteria
(Fig. 4). The neat agar and agar/lignin films did not show any
antibacterial activity against both Gram-positive and Gramnegative
bacteria. Similar to the control, they showed exponential
growth of both bacteria from an initial concentration of
10 6 e10 7 CFU/mL to 10 10 CFU/mL within 12e15 h of incubation. On
the contrary, the AgNPs incorporated composite films exhibited
distinctive antimicrobial activity against both Gram-positive and
Gram-negative bacteria, and their activity was dependent on the
concentration of AgNPs. Here, we used two composite films, agar/
lignin/AgNP 1.0 and agar/lignin/AgNP 2.0 to test the antibacterial activity
of the composite films. The one was chosen as the optimum
concentration of AgNPs based on the result of the mechanical and
water vapor barrier properties. The other was chosen to test the
effect of the concentration of AgNPs. Both composite films exhibited
stronger antimicrobial activity against E. coli than
L. monocytogenes. The composite films with 1 wt% AgNPs reduced
the bacterial population of L. monocytogenes from 10 6 CFU/mL to
10 2 CFU/mL after 15 h. However, it destroyed E. coli completely after
9 h of incubation. The antimicrobial activity increased when the
concentration of AgNPs increased, i.e., the composite films with
2 wt% AgNPs destroyed L. monocytogenes and E. coli completely in
12 h and 6 h, respectively. It is well known that the AgNPs have
stronger antimicrobial activity against Gram-negative than Grampositive
bacteria (Shankar & Rhim, 2015), which agrees well with
the present results. The difference in the antimicrobial activity of
Fig. 4. Antibacterial activity of composite films.
AgNPs between Gram-positive and Gram-negative bacteria might
be due to the difference in the cell wall structure. Gram-positive
bacteria have thick peptidoglycan layer and complex cell wall
structure that makes more difficult for AgNPs to penetrate into the
cell (Priyadarshini, Gopinath, Priyadharsshini, Ali, & Velusamy,
2013). On the contrary, Gram-negative bacteria have thin peptidoglycan
layer coated with the negatively charged outer membrane,
which makes the entry of AgNPs easier. Several probable
mechanisms have been reported on the antimicrobial activity of

S. Shankar, J.-W. Rhim / Food Hydrocolloids 71 (2017) 76e84 83
AgNPs, though a precise mechanism has not been established yet.
The most widely known mechanism is the interaction between
positively charged silver ions and negatively charged phosphorus
or sulfur-containing biomolecules such as proteins and nucleic
acids, which causes structural changes in the bacterial cell walls
leading to disruption of metabolic processes and cell death (Butkus,
Edling, & Labare, 2003). The other antibacterial mechanism of silver
nanoparticles is related to membrane damage due to free radicals
derived from the surface of the AgNPs, increasing the membrane
permeability and cell death (Kim et al., 2007). Above all, the antibacterial
action of the composite films would be initiated by the
contact of AgNPs, exposed to the surface of the film or free AgNPs or
Ag ions released from the film, with the bacterial cells. As seen from
the SR test results, it is evident that the water molecules diffused
into the film will swell the film and facilitate the release of AgNPs
into the liquid medium to be oxidized into silver ions.
The composite films with strong antimicrobial activity have a
high potential for the application in an antimicrobial active food
packaging. However, there is a safety issue on AgNPs with possible
migration from packaging film to food materials or the environment.
Presently, very little information is available to assess the
migration of nanoparticles from food packaging materials to the
packaged food and possible hazard on human beings and environment
(Rhim et al., 2013). However, it has been reported that the
impact of AgNPs synthesized by the green methods is much lower
than that synthesized by chemical or physical methods (Shankar,
Prasad, Selvakannan, Jaiswal, & Laxman, 2015). Therefore, the
nanocomposite films developed in the present study have the potential
for their applications in food packaging to prolong the shelflife
of packed food.
4. Conclusion
Lignin and AgNPs were used to prepare antimicrobial agarbased
films with improved film properties. The lignin was used as
reducing agent for the synthesis of AgNPs and a reinforcing filler for
the preparation of the composite films. The incorporation of lignin
increased the tensile strength, water vapor barrier, and UV-light
barrier properties. The inclusion of AgNPs further increased the
tensile strength and water vapor barrier properties of the composite
films and provided strong antibacterial property. The combined
use of lignin and AgNPs enhanced the functional properties
of the composite films. The incorporation of AgNPs up to 1 wt%
increased the mechanical and water vapor barrier properties of the
composite films. The composite films exhibited potent antibacterial
activity against E. coli and L. monocytogenes. These results suggest
that the agar/lignin/AgNPs nanocomposite films with UV-light
barrier property, antimicrobial activity, and improved film properties
have a high potential for their application in active food
packaging to secure the food safety and to prolong the shelf-life of
the packaged foods.
Acknowledgements
This research was supported by the Agricultural Research
Council (ARC 710003) program of the Ministry of Agriculture, Food
and Rural Affairs, Korea, and Korea Research Fellowship Program
through the National Research Foundation of Korea (NRF) funded
by the Ministry of Science, ICT and Future Planning
(2016H1D3A1903910).
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