Antimicrobial N-halamine modified chitosan films

Article abstract of DOI:10.1016/j.carbpol.2012.08.115

The inherent antimicrobial properties and biodegradability of chitosan make it an ideal candidate for antimicrobial materials. In this study, N-halamine precursor 3-glycidyl-5,5-dimethylhydantoin (GH) was synthesized and bonded onto chitosan by a ring ope

Full text of DOI:10.1016/j.carbpol.2012.08.115

Carbohydrate Polymers 92 (2013) 534–539
Contents lists available at SciVerse ScienceDirect
Carbohydrate Polymers
journal homepage: www.elsevier.com/locate/carbpol
Antimicrobial N-halamine modified chitosan films
Rong Lia, Pei Hua, Xuehong Rena,∗, S.D. Worleyb, T.S. Huangc
a Key Laboratory of Eco-textiles of Ministry of Education, College of Textiles and Clothing, Jiangnan University, Wuxi, Jiangsu 214122, China
b Department of Chemistry and Biochemistry, Auburn University, Auburn, AL 36849, USA
c Department of Poultry Science, Auburn University, Auburn, AL 36849, USA
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 12 May 2012
Received in revised form 30 August 2012
Accepted 31 August 2012
Available online 7 September 2012
The inherent antimicrobial properties and biodegradability of chitosan make it an ideal candidate for
antimicrobial materials. In this study, N-halamine precursor 3-glycidyl-5,5-dimethylhydantoin (GH) was
synthesized and bonded onto chitosan by a ring opening reaction between chitosan and GH. The chitosan
film modified with the N-halamine precursor could be rendered biocidal after exposure to a dilute house-
hold bleach solution. Syntheses routes, characterization data, and antimicrobial test results are presented.
The chlorinated films with 2.60 × 1018 atoms/cm2 of active chlorine were challenged with Staphylococcus
aureus (ATCC 6538) and Escherichia coli O157:H7 (ATCC 43895) and showed good efficacy against these
two bacterial species with log reductions of 7.4 and 7.5 within 10 and 5 min of contact time, respectively.
These films may serve as potential materials for food packaging and biomedical applications.
© 2012 Elsevier Ltd. All rights reserved.
Keywords:
Chitosan
N-halamine
Antibacterial
Biocidal
Films
1. Introduction
of these compounds might also affect the film properties and food
qualities.
Recently, there has been a growing interest in the devel-
opment of antimicrobial films in various applications including
biomedical surface coatings and food packaging. The antimicrobial
packaging films have shown great potential to control growth of
food borne pathogens, including Listeria monocytogenes, Escherichia
coli O157: H7 and Salmonella typhimurium (Cagri, Ustunol, &
Ryser, 2004). Extensive research work has been performed to
develop antimicrobial packaging to inactivate the pathogens and
spoilage microorganisms on the surface of food (Moura, Mattoso,
& Zucolotto, 2012; Tankhiwale & Bajpai, 2012). One of the sim-
plest and most feasible methods of producing antimicrobial films
is to load antimicrobial agents, such as nanoparticles (Au, Ag, Cu,
ZnO, TiO2) (Fayaz, Balaji, Girilal, Kalaichelvan, & Venkatesan, 2009;
Li, Xing, Jiang, Ding, & Li, 2009; Sikong, Kongreong, Kantachote, &
Sutthisripok, 2010), essential oils (Kuorwel, Cran, Sonneveld, Miltz,
& Bigger, 2011), bacteria originated antibacterial proteins (bacteri-
ocins) (Seydim & Sarikus, 2006), enzymes (Buonocore et al., 2003),
fruit extracts (Du et al., 2008), and chitosan (Aider, 2010; Tripathi,
Mehrotra, & Dutta, 2010) onto films. However, the release of these
additives from the films, especially water-soluble compounds and
metal nanoparticles, may raise issues regarding adverse environ-
mental problems and their safe use in food products. The addition
Chitosan, the linear and partly acetylated (1-4)-2-amino-
2-deoxy-␤-d-glucan (Muzzarelli et al., 2012), is a biopolymer
obtained from chitin. It is the second most abundant polysaccharide
in nature after cellulose, and has received great attention because
of its non-toxicity, biocompatibility, versatility, biodegradability,
and antimicrobial properties (Xu, Xin, Li, Huang, & Zhou, 2010).
Because chitosan has intrinsic antimicrobial properties and good
film-forming ability, it has been used in antimicrobial films and
coatings to inhibit the growth of not only Gram-positive and Gram-
negative bacteria, but also yeasts and molds (Chen, Yeh, & Chiang,
1996). However, the antimicrobial activity of chitosan is only mod-
erate which may limit its many applications. In order to improve
its antibacterial activity, numerous modifications of chitosan have
been reported such as N-carboxybutylation (Muzzarelli et al.,
1990; Muzzarelli, Ilari, & Petrarulo, 1994), O-carboxymethylation
(Chen & Park, 2003), quaternization (Sajomsang, Ruktanonchai,
Gonil, & Warin, 2010), sugar-modification (Sajomsang, Gonil, &
Tantayanon, 2009), N-alkylation (Yang, Chou, & Li, 2005), and chi-
tosan/nanoparticle complexes (Li, Deng, Deng, Liu, & Li, 2010).
For over three decades, N-halamine compounds have gained
growing attention as antimicrobial agents due to their efficacies
against a broad-spectrum of microorganisms, long-term stabili-
ties, non-toxicity to humans, and regenerabilities upon exposure
to aqueous free chlorine solutions (Worley et al., 2003; Kocer,
Cerkez, Worley, Broughton, & Huang, 2011). N-halamines refer to
compounds that contain amine, amide, and imide halamine bonds,
which have the capability of rapid and total inactivation of various
∗
Corresponding author. Tel.: +86 051085912007; fax: +86 051085912009.
E-mail address: xhren@jiangnan.edu.cn (X. Ren).
0144-8617/$ – see front matter © 2012 Elsevier Ltd. All rights reserved.
http://dx.doi.org/10.1016/j.carbpol.2012.08.115

R. Li et al. / Carbohydrate Polymers 92 (2013) 534–539
535
microorganisms without causing the microorganisms to develop
resistance to them. Furthermore, N-halamine biocides are capable
of killing microorganisms directly without the release of free chlo-
rine into the system (Barnes et al., 2006). These N-halamines could
be incorporated into or attached onto textiles, medical devices, and
other solid surfaces related to health care (Worley, Chen, Wang, &
Wu, 2005). The introduction of the N-halamines in textiles by cova-
lent bonding between the precursor moieties and host polymers
has been extensively explored in these laboratories and elsewhere
over the past decade (Liang et al., 2007; Luo & Sun, 2008). N-
halamine moieties have been applied onto cotton by using grafting
techniques or other coating methods to produce antimicrobial cel-
lulose (Sun & Sun, 2001), nylon (Lin, Cammarata, & Worley, 2001),
and polyester (PET) (Ren et al., 2008).
However, little research has been done on the modifications
of chitosan by covalently incorporating N-halamine moieties to
produce biocidal chitosan. Recently, Cao and Sun (2008) reported
that antimicrobial chitosan was prepared by exposure of chitosan
to dilute sodium hypochlorite. The N-halamine chitosan provided
total kill of 108–109 CFU/mL of E. coli (Gram-negative bacteria) and
Staphylococcus aureus (Gram-positive bacteria) in 10 and 60 min,
respectively. The physical state of chitosan is a crucial factor affect-
ing antimicrobial activity, and extensive works have been done
for chitosan in water solution. Little attention has been paid to
the investigation of inactivation of microorganisms by chitosan
in the solid state, such as for films and fibers. In this study, an
N-halamine precursor 3-glycidyl-5,5-dimethylhydantoin was syn-
thesized and attached to chitosan by a ring-opening reaction. The
newly synthesized N-halamine chitosan derivative was character-
ized by FT-IR, NMR, DTG, and TGA. The chitosan derivative was
dissolved in acetic acid solution and coated onto polyester trans-
parency slides and then rendered antimicrobial upon chlorination.
The chlorinated films were evaluated for biocidal efficacies against
both Gram-negative bacteria E. coli O157:H7 and Gram-positive
bacteria S. aureus.
Scheme 1. The schematic description of synthesis of chitosan–GH and
chitosan–GH–Cl.
heating rate of 5 ◦C min−1 under nitrogen atmosphere (flow rate of
40 mL min−1).
2.3. Synthesis of 3-glycidyl-5,5-dimethylhydantoin (GH)
3-Glycidyl-5,5-dimethylhydantoin (GH) was synthesized using
the method reported by Liang et al. (2007). In brief, an equimo-
lar mixture of NaOH and 5,5-dimethylhydantoin were added to
water and stirred for 5–10 min at ambient temperature. Then an
equimolar quantity of epichlorohydrin was added followed by stir-
ring for 10 h at ambient temperature. After the reaction water was
removed at reduced pressure, and acetone was added to the flask
to dissolve the product. Sodium chloride produced in the reaction
was removed by filtration, and the desired product was obtained
by the evaporation of the solvent acetone. The yield of the product
was 89.23%. 1H NMR (d-acetone): ı 1.41 (6H), 2.77–2.91 (2H), 3.20
(1H), 3.48–3.65 (2H).
2. Experimental
2.1. Materials
Chitosan with a molecular weight of 200 kDa and deacetylation
degree of 95% was purchased from Zhejiang Aoxing Biochemi-
cal Co., Ltd., China. 5,5-Dimethylhydantoin was purchased from
Hebei Yaguang Fine Chemical Co., Ltd.; epichlorohydrin, acetone,
household bleach (the active chlorine content was 5%), and acetic
acid (analytical grade) were purchased from Sinopharm Chemical
Reagent Co., Ltd, Shanghai. All reagents were used as received with-
out further purification. The PET film was purchased from Wenzhou
Kewen Teaching Equipment Co., Ltd. The bacteria employed were S.
aureus ATCC 6538 and E. coli O157:H7 ATCC 43895 (American Type
Culture Collection, Rockville, MD). The Trypticase soy agar used was
from Difco Laboratories, Detroit, MI.
2.4. Modification of chitosan with GH
Appropriate amounts of GH and chitosan (the molar ratios of
GH to chitosan were 0.8, 1.0, 1.2 and 1.5) were dissolved in 1% (v/v)
acetic acid solution. The mixture was stirred at 65 ◦C for 10 h. Water
was removed under vacuum, and excess acetone was added to the
flask. The precipitate was filtered and washed with abundant ace-
tone, then dried in a vacuum oven at 60 ◦C for 48 h. The yields of the
products were about 93%. The syntheses of GH and chitosan–GH are
shown in Scheme 1.
2.5. The degree of substitution of chitosan by GH
2.2. Instruments
The degree of substitution (DS) of chitosan by GH was measured
by UV–VIS spectroscopy; the absorbance of different concentra-
tions of GH were employed in the standard curve. DS is defined
as the molar ratio of bonded GH of glucosamine calculated from
the original mass of chitosan and its degree of deacetylation (DD).
The absorbance of chitosan–GH acetic acid solution was measured
first, and then the corresponding concentrations were found from
the standard curve (c2). The DS were determined by the following
equation:
The FT-IR spectra of chitosan, chitosan–GH and chitosan–GH–Cl
were recorded by a Nicolet Nexus 470 spectrometer in the opti-
cal range of 400–4000 cm−1 by averaging 32 scans at a resolution
of 4 cm−1. All samples were prepared as potassium bromide pel-
lets. The UV–VIS spectra of chitosan, GH, and chitosan–GH were
measured by a UNICO UV-2802S in the range of 190–900 nm.
The 1H NMR of GH and 13C NMR spectrum of chitosan–GH’s was
recorded on a Bruker AV-300 spectrometer. Thermogravimetric
analysis (TGA) was carried out using a TGA unit (NETZSCH STA 499
F3). About 5 mg of sample was heated from 20 ◦C to 600 ◦C at a
c2
DS =
× 100%
(1)
(C1 − c2 × M2/M1) × DD

536
R. Li et al. / Carbohydrate Polymers 92 (2013) 534–539
where c2 was the concentration of GH grafted on chitosan, mol/L;
C1 was the concentration of chitosan–GH, g/L; M2 was the molecu-
lar weight of GH; M1 was the monomer average molecular weight
of chitosan; and DD was the degree of deacetylation of chitosan,
respectively.
2.6. Preparation of chitosan–GH films
The synthesized chitosan–GH was dissolved in 2% acetic acid
solution at different concentrations (from 1% to 6%). Then 10 mL
of polymer solution were coated on transparency PET slides
(9 × 9 cm), followed by evaporation of the solvent in the fume
hood overnight. The polymer films were neutralized by immersing
in 1 mol/L aqueous NaOH solution at ambient temperature under
constant stirring for 1 h, followed by thoroughly washing with
deionized water. After air drying, the polymer films were stored
in a desiccator for further characterization and testing.
2.7. Chlorination procedure
Fig. 1. The effect of molar ratio of GH on active chlorine content in chitosan–GH–Cl.
The chitosan, chitosan–GH and chitosan–GH films were soaked
in a 5% solution of household bleach (pH adjusted to 7 using 1 N
H2SO4) with stirring at ambient temperature for 60 min. After
chlorination, the samples were washed thoroughly with deionized
water to remove free chlorine. The chlorinated films were dried at
45 ◦C for 1 h and stored in a desiccator for further testing.
incubated at 37 ◦C for 24 h. The bacterial colonies were recorded
and enumerated for biocidal efficacy.
3. Results and discussion
3.1. The effect of molar ratio of chitosan and GH on the chlorine
loadings
2.8. Analytical titration
The degree of substitution (DS) of chitosan by GH was measured
by UV–VIS spectroscopy; the absorbances of different concentra-
tion of GH were employed in constructing a standard curve. In
this study, four different DS of chitosan–GH were synthesized by
changing the molar ratio of GH to chitosan from 0.8 to 1.5. The DS
increased from 3.5% to 4.6% by the increasing of the molar ratio.
The effects of molar ratio of GH and chitosan on the active chlo-
rine content of the chitosan–GH samples are shown in Fig. 1. The
molar ratio of GH to chitosan ranged from 0.8 to 1.5 (DS was 3.5%,
4.1%, 4.6% and 4.5%, respectively). Fig. 1 shows that the increase of
molar ratio within the above range does not significantly affect the
oxidative chlorine content. The increase of the molar ratio of GH
could not further increase the active chlorine content which con-
firms that the amine groups in chitosan are favorable sites for the
reaction; this was in agreement with that of the DS measurement. A
molar ratio of 1:1 was employed in this study. However, the active
chlorine loading of chitosan after the introduction of GH was sig-
nificantly increased from 8.9% of chitosan to 12.9% of chitosan–GH
indicating that the amide nitrogen atoms on the hydantoin ring
were being chlorinated.
The iodometric/thiosulfate titration procedure was employed
to determine the oxidative chlorine loading of the modified
compounds and the coated films. For example, about 0.05 g of
chlorinated sample was suspended in a 50 mL water solution, into
which, 0.5 g of KI and 1 mL of 1% of starch water solution were
added. The solution was titrated with 0.005 N sodium thiosulfate.
The bonded oxidative chlorine on the chitosan–GH compounds was
calculated with the equation below:
N × V × 35.45
Cl+% =
× 100
(2)
W × 2
where N and V are the normality (equiv./L) and volume (L) of the
Na2S2O3 consumed in the titration, respectively, while W is the
weight of the chitosan and chitosan–GH–Cl samples in grams.
The chlorine loadings of the films (the amount of chlorine on the
surfaces) on the slides were calculated with the following equation:
N × V × 6.02 × 1023
Cl+(atoms/cm2) =
(3)
2 × S
where N and V are the normality (equiv./L) and volume (L) of the
titrant sodium thiosulfate, respectively, and S is the area of the film
in cm2.
3.2. Preparation of chlorinated chitosan–GH films with different
concentrations
2.9. Antimicrobial test
Different concentrations of the synthesized chitosan–GH, 1%, 2%,
4%, 6%, were dissolved in 2% acetic acid solution. The chitosan–GH
solution was coated on PET transparency slides. After neutral-
ization and drying, the films were chlorinated according to the
method mentioned in experimental section. The chlorine loadings
of chitosan–GH on these transparency slides are shown in Fig. 2.
The active chlorine contents increase from 2.15 × 1018 atoms/cm2
to 2.60 × 1018 atoms/cm2 within the range from 1% to 4%, while
further increase of concentration of chitosan–GH (increased to 6%)
does not significantly increase the active chlorine contents. Thus,
4% of chitosan–GH solution was used for coating the films in further
studies. Films with high chlorine loading of 2.60 × 1018 atoms/cm2
would be expected to be quite antibacterial.
Both chlorinated and unchlorinated modified chitosan films
were challenged with S. aureus (ATCC 6538) and E. coli O157:H7
(ATCC 43895) using a modified AATCC Test Method 100-1999. The
testing began with the addition of 25 L of the bacterial suspensions
buffered at pH 7 to the centers of 6.45 cm2 portions of slides in a
sterile Petri dish, and with second identical slides placed upon the
first ones held in place by a sterile weight. Different slides were
exposed to the bacteria with contact times of 1, 5, 10, and 30 min,
respectively; they were placed in tubes containing 5.0 mL of sterile
0.02 N sodium thiosulfate to remove all oxidative chlorine. Series
of the quenched solutions were diluted with pH 7, 100 mM phos-
phate buffer, and plated on Trypticase soy agar. The plates were

R. Li et al. / Carbohydrate Polymers 92 (2013) 534–539
537
Fig. 2. Effect of the concentration of chitosan–GH on active chlorine content.
Fig. 4. The 13C NMR spectra of chitosan–GH using D2O and CF3COOD as the solvents.
3.3. Characterization
carbonyl bands upon chlorination have been reported previously
in the literature (Ren, Kocer, Worley, Broughton, & Huang, 2009).
The 13C NMR spectrum of chitosan–GH in D2O and CF3COOD
solution is presented in Fig. 4. The signals ascribed to the car-
bons in the chitosan backbone at 100.7 ppm (C-1), 63.6 ppm (C-2),
73.2 ppm (C-3), 80.6 ppm (C-4), 78.0 ppm (C-5), and 59.2 ppm (C-
6) are shown in Fig. 4 (Jiang et al., 2011). The signals at 56.9 ppm,
68.1 ppm, 63.6 ppm, 179.2 ppm, and 22.9 ppm may be assigned to
3.3.1. Structural determination by 13C NMR, FT-IR and UV–VIS
The FT-IR spectra of chitosan, chitosan–GH and chitosan–GH–Cl
are recorded in Fig. 3. The spectrum of chitosan showed the char-
acteristic absorption bands at 1655 (amide I, C O stretching), 1597
(amide II, –NH2 bending) and 1381 cm−1 (–CH2 bending). Other
characteristic bands of the polysaccharide appeared at 1152 cm−1
(symmetric stretching of the C–O–C) and 1080 cm−1 (skeletal
vibration of the C–O stretching) (Sajomsang, Gonil, & Tantayanon,
2009). The strong band at around 3400 cm−1 was assigned to the
stretching vibration for O–H, the extension vibration of N–H, and
inter-molecular hydrogen bonds of the polysaccharide moieties.
All of the bands mentioned above were common in the spectra of
chitosan–GH samples due to the presence of the chitosan backbone.
Compared with the FT-IR spectrum of pure chitosan, the spectra of
chitosan–GH showed two strong absorption peaks at 1569 cm−1
(C O stretching) (Haro et al., 2005), which indicates that chitosan
was successfully modified with GH through covalent bonds. The
vibrational band of C O shifts from 1569 cm−1 to 1630 cm−1 after
chlorination. The shifts to higher wave number of the hydantoin
the carbons of the –CH2 (C-7, C-9), –CH (C-8), –CH (C-12), –C
O
(C-10, C-11), and –CH3 (C-13) (Lu, Wu, & Fu, 2007; Ren et al.,
2009) groups, respectively. These signals on the GH units detected
above further confirmed that GH was bonded to chitosan success-
fully.
The UV–VIS spectra of chitosan, GH, and chitosan–GH are shown
in Fig. 5. The UV absorption at 240 nm can be attributed to the
hydantoin ring structure (Chen & Sun, 2006), which is evident in the
spectra of GH and chitosan–GH. Chitosan does not show absorption
under UV analysis. The band at 240 nm of chitosan–GH provided
further evidence that the hydantoin ring structure was covalently
bonded onto chitosan.
Fig. 3. The FT-IR spectra of chitosan, chitosan–GH and chitosan–GH–Cl.
Fig. 5. UV spectra of chitosan, GH, and chitosan–GH.

538
R. Li et al. / Carbohydrate Polymers 92 (2013) 534–539
Fig. 6. TGA and DTG curves for chitosan, un-chlorinated chitosan–GH and chlorinated chitosan–GH.
Table 1
Antibacterial property against S. aureusa and E. coli O157:H7b.
Sample
Contact time (min)
BacterialreductionS. aureusa
Bacterial reduction
E. coli O157:H7b
%
Log
%
Log
Un-chlorinated chitosan–GH Film
Chlorinated chitosan–GH Film
30
1
5
10
30
39.34
99.90
99.93
100
100
0.22
2.98
3.14
7.39
7.39
3.97
99.62
100
100
100
0.02
1.52
7.48
7.48
7.48
a
The inoculum population was 2.43 × 107 cfu/sample.
The inoculum population was 3.73 × 107 cfu/sample.
b
3.3.2. Thermal analysis
Gram-negative E. coli O157:H7 are presented in Table 1. The films
were challenged with S. aureus and E. coli O157:H7 bacteria at
2.43 × 107 cfu/sample and 3.73 × 107 cfu/sample, respectively. The
unchlorinated chitosan–GH films produced only a small log reduc-
tion for both bacteria, 0.218 log reduction of S. aureus and 0.017 log
reduction of E. coli after 30 min of contact due to the adhesion
of the bacteria to the films and perhaps a small degree inactiva-
tion of the bacteria. The chlorinated chitosan–GH films containing
2.60 × 1018 atoms/cm2 of active chlorine completely inactivated
both S. aureus and E. coli O157:H7 with log reductions of 7.39 and
7.48 within 10 min and 5 min of contact, respectively. About 99.9%
of both S. aureus and E. coli O157:H7 could be killed by the chlori-
nated films within 1 min of contact.
The TGA thermograms of chitosan, chitosan–GH, and
chitosan–GH–Cl are shown in Fig. 6. The TGA of chitosan showed
two different stages of weight loss. The first stage weight loss
extended from 32 to 101 ◦C, and Tmax was located at 51 ◦C. This may
correspond to the loss of adsorbed and bound water (Kumar, Koh,
Kim, Gupta, & Dutta, 2012). The second stage of weight loss starts
at 217 ◦C and continues up to 350 ◦C (Tmax was located at 286 ◦C)
and is due to the degradation of chitosan biopolymer (Kumar
et al., 2012). Adelaida Ávila et al. also reported that the weight loss
between 40 ◦C and 200 ◦C may be attributed to the adsorbed and
bound water (Ávila, Bierbrauer, Pucci, López-González, & Strumia,
2012). Therefore, the weight loss of chitosan–GH extending from
28 to 80 ◦C (Tmax, 51 ◦C) and from 80 to 156 ◦C (Tmax was located
at 108 ◦C), and the weight loss of chitosan–GH–Cl extending from
28 to 101 ◦C (Tmax was located at 62 ◦C) and 101 to 118 ◦C (Tmax
,
4. Conclusions
108◦C) also may be attributed to the adsorbed and bound water.
The weight loss of chitosan–GH from decomposition from 200 to
360 ◦C (Tmax, 275 ◦C) is very close to that of chitosan, which indi-
cates that incorporation of GH in chitosan does not significantly
affect its thermal performance. The chlorinated chitosan–GH–Cl
N-halamine chitosan (chitosan–GH) was prepared by a ring-
opening reaction between chitosan and GH. The incorporation
of GH into chitosan was confirmed by FT-IR, 13C NMR, UV, and
TGA analyses. The active chlorine content of chitosan–GH–Cl and
chitosan–GH–Cl film could reach 12.9% and 2.60 × 1018 atoms/cm2,
respectively, upon exposure to dilute household bleach. The
N-halamine chitosan provided excellent antimicrobial efficacies
against both Gram-negative and Gram-positive bacteria by inac-
tivating 7 logs of S. aureus and E. coli O157:H7 within 5–10 min.
The addition of stable cyclic N-halamine with a hydantoin ring
into chitosan significantly increases the active chlorine loadings on
the films which enhances the antimicrobial activity and durability.
These encouraging results show that N-halamine-based chitosan
has potential for a wide range of biomedical applications including
showed that the weight loss temperature from 145 to 280 ◦C (Tmax
,
192 ◦C) is lower than that of the un-chlorinated chitosan. The
decomposition of the N–Cl bonds in the chlorinated chitosan–GH
may accelerate the thermal decomposition of chitosan–GH through
a free radical process.
3.4. Antibacterial efficacy
The biocidal efficacy data for the unchlorinated and chlo-
rinated chitosan–GH films against Gram-positive S. aureus and

R. Li et al. / Carbohydrate Polymers 92 (2013) 534–539
539
wound dressings, coatings for medical devices, and food packag-
ing.
Liang, J., Chen, Y., Ren, X., Wu, R., Barnes, K., Worley, S. D., et al. (2007). Fabric
treated with antimicrobial N-halamine epoxides. Industrial & Engineering Chem-
istry Research, 46, 6425–6429.
Lin, J., Cammarata, V., & Worley, S. D. (2001). Infrared characterization of biocidal
nylon. Polymer, 42, 7903–7906.
Lu, G. Q., Wu, D. C., & Fu, R. W. (2007). Studies on the synthesis and antibacterial
activities of polymeric quaternary ammonium salts from dimethylaminoethyl
methacrylate. Reactive & Functional Polymers, 67, 355–366.
Luo, J., & Sun, Y. (2008). Acyclic N-halamine coated Kevlar fabric materials: Prepa-
ration and biocidal functions. Industrial & Engineering Chemistry Research, 47,
5291–5297.
Muzzarelli, R. A. A., Boudrant, J., Meyer, D., Manno, N., DeMarchis, M., & Paoletti,
M. G. (2012). Current views on fungal chitin/chitosan, human chitinases, food
preservation, glucans, pectins and inulin: A tribute to Henri Braconnot, precursor
of the carbohydrate polymers science, on the chitin bicentennial. Carbohydrate
Polymers, 87, 995–1012.
Muzzarelli, R. A. A., Tarsi, R., Filippini, O., Giovanetti, E., Biagini, G., & Varaldo, P.
E. (1990). Antimicrobial properties of N-carboxybutyl chitosan. Antimicrobial
Agents and Chemotherapy, 34, 2019–2023.
Muzzarelli, R. A., Ilari, P., & Petrarulo, M. (1994). Solubility and structure of N-
carboxymethylchitosan. International Journal of Biological Macromolecules, 16,
177–180.
Ren, X., Kocer, H. B., Kou, L., Worley, S. D., Broughton, R. M., Tzou, Y. M., et al. (2008).
Antimicrobial polyester. Journal of Applied Polymer Science, 109, 2756–2761.
Ren, X., Kocer, H. B., Worley, S. D., Broughton, R. M., & Huang, T. S. (2009). Recharge-
able biocidal cellulose: Synthesis and application of 3-(2,3-dihydroxypropyl)-
5,5-dimethylimidazolidine-2,4-dione. Carbohydrate Polymers, 75, 683–687.
Sajomsang, W., Gonil, P., & Tantayanon, S. (2009). Antibacterial activity of quaternary
ammonium chitosan containing mono or disaccharide moieties: Preparation
and characterization. International Journal of Biological Macromolecules, 44,
419–427.
Sajomsang, W., Ruktanonchai, U. R., Gonil, P., & Warin, C. (2010). Quaternization of
N-(3-pyridylmethyl) chitosan derivatives: Effects of the degree of quaterniza-
tion, molecular weight and ratio of N-methylpyridinium and N,N,N-trimethyl
ammonium moieties on bactericidal activity. Carbohydrate Polymers, 82,
1143–1152.
References
Aider, M. (2010). Chitosan application for active bio-based films production and
potential in the food industry: Review. Lwt-Food Science and Technology, 43,
837–842.
Ávila, A., Bierbrauer, K., Pucci, G., López-González, M., & Strumia, M. (2012). Study
of optimization of the synthesis and properties of biocomposite films based on
grafted chitosan. Journal of Food Engineering, 109, 752–761.
Barnes, K., Liang, J., Wu, R., Worley, S. D., Lee, J., Broughton, R. M., et al. (2006).
Synthesis and antimicrobial applications of 5,5ꢀ-ethylenebis 5-methyl-3-(3-
triethoxysilylpropyl)hydantoin. Biomaterials, 27, 4825–4830.
Buonocore, G. G., Del Nobile, M. A., Panizza, A., Bove, S., Battaglia, G., & Nico-
lais, L. (2003). Modeling the lysozyme release kinetics from antimicrobial films
intended for food packaging applications. Journal of Food Science, 68, 1365–1370.
Cagri, A., Ustunol, Z., & Ryser, E. T. (2004). Antimicrobial edible films and coatings.
Journal of Food Protection, 67, 833–848.
Cao, Z., & Sun, Y. (2008). N-halamine-based chitosan: Preparation, characterization,
and antimicrobial function. Journal of Biomedical Materials Research Part A, 85,
99–107.
Chen, M. C., Yeh, G. H. C., & Chiang, B. H. (1996). Antimicrobial and physicochemi-
cal properties of methylcellulose and chitosan films containing a preservative.
Journal of Food Processing and Preservation, 20, 279–290.
Chen, X.-G., & Park, H.-J. (2003). Chemical characteristics of O-carboxymethyl
chitosans related to the preparation conditions. Carbohydrate Polymers, 53,
355–359.
Chen, Z., & Sun, Y. (2006). N-halamine-based antimicrobial additives for polymers:
Preparation, characterization, and antimicrobial activity. Industrial and Engineer-
ing Chemistry Research, 45, 2634–2640.
de Moura, M. R., Mattoso, L. H. C., & Zucolotto, V. (2012). Development of cellulose-
based bactericidal nanocomposites containing silver nanoparticles and their use
as active food packaging. Journal of Food Engineering, 109, 520–524.
Du, W. X., Olsen, C. W., Avena-Bustillos, R. J., McHugh, T. H., Levin, C. E., & Friedman,
M. (2008). Antibacterial activity against E. coli O157: H7, physical properties,
and storage stability of novel carvacrol-containing edible tomato films. Journal
of Food Science, 73, M378–M383.
Fayaz, A. M., Balaji, K., Girilal, M., Kalaichelvan, P. T., & Venkatesan, R. (2009).
Mycobased synthesis of silver nanoparticles and their incorporation into sodium
alginate films for vegetable and fruit preservation. Journal of Agricultural and
Food Chemistry, 57, 6246–6252.
Haro, M., Giner, B., Lafuente, C., Lopez, M. C., Royo, F. M., & Cea, P. (2005). Proton
sponge and fatty acid interactions at the air–water interface. Thermodynamic,
spectroscopic, and microscopic study. Langmuir, 21, 2796–2803.
Jiang, X., Cai, K., Zhang, J., Shen, Y., Wang, S., & Tian, X. (2011). Synthesis of a novel
water-soluble chitosan derivative for flocculated decolorization. Journal of Haz-
ardous Materials, 185, 1482–1488.
Kocer, H. B., Cerkez, I., Worley, S. D., Broughton, R. M., & Huang, T. S. (2011). Poly-
meric antimicrobial N-halamine epoxides. Acs Applied Materials & Interfaces, 3,
2845–2850.
Seydim, A. C., & Sarikus, G. (2006). Antimicrobial activity of whey protein based
edible films incorporated with oregano, rosemary and garlic essential oils. Food
Research International, 39, 639–644.
Sikong, L., Kongreong, B., Kantachote, D., & Sutthisripok, W. (2010). Inactivation of
salmonella typhi using Fe(3+) doped TiO(2)/3SnO(2) photocatalytic powders and
films. Journal of Nano Research, 12, 89–97.
Sun, Y. Y.,
& Sun, G. (2001). Novel regenerable N-halamine polymeric
biocides. I. Synthesis, characterization, and antibacterial activity of
hydantoin-containing polymers. Journal of Applied Polymer Science, 80,
2460–2467.
Tankhiwale, R., & Bajpai, S. K. (2012). Preparation, characterization and antibacterial
applications of ZnO-nanoparticles coated polyethylene films for food packaging.
Colloids and Surfaces B-Biointerfaces, 90, 16–20.
Tripathi, S., Mehrotra, G. K.,
& Dutta, P. K. (2010). Preparation and physico-
chemical evaluation of chitosan/poly(vinyl alcohol)/pectin ternary film for
food-packaging applications. Carbohydrate Polymers, 79, 711–716.
Worley, S. D., Chen, Y., Wang, J. W., & Wu, R. (2005). Novel N-halamine siloxane
monomers and polymers for preparing biocidal coatings. Surface Coatings Inter-
national Part B-Coatings Transactions, 88, 93–99.
Kumar, S., Koh, J., Kim, H., Gupta, M. K., & Dutta, P. K. (2012). A new chitosan–thymine
conjugate: Synthesis, characterization and biological activity. International Jour-
nal of Biological Macromolecules, 50, 493–502.
Worley, S. D., Li, F., Wu, R., Kim, J., Wei, C. I., Williams, J. F., et al. (2003). A novel
N-halamine monomer for preparing biocidal polyurethane coatings. Surface
Coatings International Part B-Coatings Transactions, 86, 273–277.
Xu, T., Xin, M., Li, M., Huang, H., & Zhou, S. (2010). Synthesis, characteristic and
antibacterial activity of N,N,N-trimethyl chitosan and its carboxymethyl deriva-
tives. Carbohydrate Polymers, 81, 931–936.
Yang, T.-C., Chou, C.-C., & Li, C.-F. (2005). Antibacterial activity of N-alkylated dis-
accharide chitosan derivatives. International Journal of Food Microbiology, 97,
237–245.
Kuorwel, K. K., Cran, M. J., Sonneveld, K., Miltz, J., & Bigger, S. W. (2011). Essential oils
and their principal constituents as antimicrobial agents for synthetic packaging
films. Journal of Food Science, 76, R164–R177.
Li, L.-H., Deng, J.-C., Deng, H.-R., Liu, Z.-L., & Li, X.-L. (2010). Preparation, charac-
terization and antimicrobial activities of chitosan/Ag/ZnO blend films. Chemical
Engineering Journal, 160, 378–382.
Li, X., Xing, Y., Jiang, Y., Ding, Y., & Li, W. (2009). Antimicrobial activities of ZnO
powder-coated PVC film to inactivate food pathogens. International Journal of
Food Science and Technology, 44, 2161–2168.