Influence of chitosan addition on the mechanical and antibacterial properties of carrot cellulose nanofibre film
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Films of carrot cellulose nanofibrils (CCNFs) with the addition of low-viscosity chitosan (CHIT) were prepared by the vacuum filtration. The chitosan content in the films varied from 9 to 33% (dry wt. basis). The surface morphology of the films was investigated by scanning electron microscopy, and it was found that chitosan was dispersed in the CCNF matrix. The interaction between CCNFs and CHIT was evaluated in terms of Fourier transform infrared spectroscopy (FTIR). The obtained results suggested physical interactions rather than hydrogen bonding between CCNFs and CHIT. This finding also supports the results of the water wettability experiment. The addition of chitosan to the nanocellulose matrix causes an increase in the water contact angle, i.e., the surface of the composites becomes more hydrophobic. This increase is probably connected to an interaction between nanocellulose and chitosan forming a denser structure. Analyses of thermal properties showed that the composites are stable under high temperature, and the degradation occurred above 300 °C. It was found that the addition of CHIT to CCNF matrices caused a decrease in the Young’s modulus—the higher that the concentration of chitosan in the composite was, the lower the Young’s modulus (decreased from 14.71 GPa for CCNFs to 8.76 GPa for CCNF/CHIT_5). Additionally, the tensile strength of composites, i.e., the maximum force that causes a fracture decreased after the addition of chitosan (decreased from 145.83 MPa for CCNFs to 129.43 MPa for CCNF/CHIT_5). The results indicated the highest inhibitory effect of the investigated composites against E. coli and S. epidermidis. Whereas M. luteus was inhibited only by the higher concentration of chitosan in the tested composites, inhibition was not found against C. krissii and all tested filamentous fungi.
KeywordsNanocellulose Chitosan Antibacterial properties Mechanical properties Thermal properties
Cellulose and chitin are two of the most abundant polymers on Earth. Both are biodegradable and biocompatible and are derived from natural renewable sources (Chen 2017; Dufresne 2012). Chitin, a linear polysaccharide of β-1,4-N-acetylglucosamine, is a component of the fungal cell wall and exoskeletons of insects or crustaceans (Chen 2017). Due to its high crystallinity and strong intermolecular bonds, chitin has low reactivity and is water insoluble. Therefore, to overcome that limitation, after deacetylation, chitosan is obtained. Chitosan is easily dissolved at low pH, and its composites with a wide range of polymers were analysed (Deng et al. 2017). Subsequently, due to their biocompatibility, biodegradability, antibacterial activity, and non-toxicity, their applications were studied in agriculture as a fertilizer, in the food industry as a stabilizer and thickener and in the field of biomedical engineering (de Alvarenga 2011; de Mesquita et al. 2010). The main disadvantage of chitosan composites is their poor mechanical properties (Abdul Khalil et al. 2016). On the other hand, positively charged chitosan is a natural antimicrobial agent against an extensive variety of microorganisms having potential as an antimicrobial agent to improve food safety and quality (Kim et al. 2011). Chitosan’s antimicrobial activity is influenced by the type of chitosan, its degree of deacetylation, its degree of polymerization, the tested host, the age of the microbial cell, the source of the medium, the chemical composition of the medium, water activity inside the medium, its concentration and pH of the medium (Dutta et al. 2009; Kong et al. 2010).
Cellulose is a linear polymer of β-1,4-glucose organized into fibrils, the main constituent of the plant cell wall (Szymańska-Chargot et al. 2011). Cellulose individual fibrils are well known for their unique properties such high mechanical strength, making them comparable to such materials as the Kevlar fibre or steel wire (Brinchia et al. 2013; Moon et al. 2011; Wang et al. 2015). Moreover, there is growing research interest in the use of cellulose nanomaterials such as nanofibrillated cellulose and cellulose nanocrystals as eco-friendly fillers and reinforcement for existing composites (Moon et al. 2011; Ämmälä et al. 2013). So far nanocellulose is produced from both forest and agriculture resources (Alemdar and Sain 2008; Chen et al. 2011; Liu et al. 2012, Rambabu et al. 2016; Rahmi and Julinawati 2017). Recently, the possibility of the use of fruit and vegetable pomaces as the source of cellulose was presented (Szymańska-Chargot et al. 2017). The cellulose was extracted under mild acid–alkali conditions, and cellulose nanofibres (CNF) were produced by high-intensity ultrasound (Szymańska-Chargot et al. 2017, 2018, 2019). CNF or, in general, nanocellulose films are often called nanopapers due to the analogous production methods with cellulosic-based paper (Klemm et al. 2011; Stark 2016). The CNF films themselves are highly flexible due to the entangled network of ultrathin nanofibrils (Wang et al. 2015). Moreover, CNF-only films have good optical transparency, and the addition of CNF to polymers does not affect the transparency (Abdul Khalil et al. 2016). Once the film is produced, the CNFs cannot be easily spread again in water, although, generally, nanocellulose films are not fully water-resistant. The main advantage of CNF films are small pores as a results of both hydrogen bonding between nanofibrils and high crystallinity, giving materials with good barrier properties useful for packaging materials (Klemm et al. 2011; Stark 2016). However, the material source, morphology, and chemical composition affect the barrier properties of CNF films (Lavoine et al. 2012). Cellulose nanostructures are usually used as a reinforcing agent for several polymers, but they can also be used as matrices for various materials, including films for food-packaging materials (Azeredoa et al. 2017). Rahmi and Julinawati (2017) showed that hydrogen bonds forming between cellulose and chitosan helps to obtain good adhesion between both materials even without using surface modificators or coupling agents (Rahmi and Julinawati 2017). Most of the publication reporting results of chitosan reinforced by cellulose focused on commercial cellulose which unfortunately results in high-cost material (Abo-Elseoud et al. 2018; Hassan et al. 2011, 2016; Trana et al. 2013).
Thus, although the properties and applications of chitosan–cellulose blends and nanocellulose-reinforced chitosan biocomposites have been reported, there is still a lack of publication reporting of the use of cellulose isolated from fruit or vegetable pomace as the composite with chitosan (Abdul Khalil et al. 2016). Previously, it has been shown that films obtained from nanocellulose originating from apple and carrot pomaces present very good mechanical properties (Szymańska-Chargot et al. 2019). The utilization of cellulose isolated from pomaces may result in a low-cost material. Here, cellulose was isolated from carrot pomace, and nanocellulose was obtained by the high-intensity ultrasonication method (HIUS) (Szymańska-Chargot et al. 2017). Chitosan, although it has interesting antimicrobial properties, is known to have poor mechanical properties. Thus, reinforcing agents are still being searched (Elsabee and Abdou 2013). Nanocellulose, with its biocompatibility, biodegradability and good optical transparency, is the potential reinforcing agent for chitosan films (Fernandes et al. 2010). Thus far, nanocellulose, both in the form of nanofibres or nanowhiskers, was used as the reinforcement of chitosan films (Fernandes et al. 2010; Falamarzpour et al. 2017; Dehnad et al. 2014) and most of the studies are focused on the characterization of their mechanical and antibacterial properties (Li et al. 2009; Rahmi and Julinawati 2017; Niu et al. 2018; Toivonen et al. 2015; Wu et al. 2014). Moreover, the addition of nanocellulose could increase the flexibility and thermal stability of chitosan-nanocellulose films compared with native chitosan films (Fernandes et al. 2010). This characteristic of chitosan–CNF films makes them useful for various applications such as electronic devices, medical and antibacterial packaging. However, reports are lacking concerning nanocellulose as the matrix and chitosan as an antimicrobial agent (Hänninen et al. 2018). Here, for the first time, composites based on nanocellulose prepared from carrot cellulose with different concentrations of chitosan from shrimp shells were prepared. The nanocellulose was the base matrix for which chitosan played only a role as an antimicrobial agent.
Materials and methods
Never-dried cellulose was isolated from carrot pomace prepared in a de-pulping machine with a double-screw shredder (Twin Gear Juice Extractor; Green Star Elite GSE-5000, Anaheim, CA, USA) as described by the method of Szymańska-Chargot et al. (2017). Carrot was purchased in a local grocery shop. Low-viscosity chitosan from shrimp shells was purchased from Sigma Aldrich Poland.
Preparation of nanofibrillated cellulose from carrot pomace
The carrot cellulose nanofibrils (CCNFs) were prepared using high-intensity ultrasonication (Szymańska-Chargot et al. 2019). The ultrasonic homogenizer Vibra Cell VCX 130 (SONICS & MATERIALS Inc.) with a net power output equal to 130 watts and a frequency of 20 kHz was used. The processor was equipped with a 6-mm-diameter probe with a maximum oscillation amplitude equal to 114 μm. The sonication system contained a temperature probe, and, to avoid heating of the sample, an ice-bath was used. The operating amplitude of the ultrasonic homogenizer was maintained to 90% of the nominal amplitude.
Each time, 250 g of the 0.2% water suspension of carrot cellulose was prepared. First, the Ultra-Turrax (T10 basic ULTRA TURRAX, IKA) was used for 10 min to initial disperse the obtained suspensions. Next, the dispersed samples were introduced to ultrasound treatment for 30 min. Thereafter, the sample was divided into two portions and diluted to obtain the 0.1 wt% dispersion. Each portion was introduced to ultrasound treatment for 30 min. Thereafter, the 0.1 wt% CCNFs dispersion was obtained. The procedure of CCNFs preparation together with the photo of CCNFs dispersion and AFM height image of the CCNFs is presented in Supplementary Material Fig. 1.
Weight proportions of CHIT and CCNFs in each composite
Content of CHIT in composite (%)
FTIR spectra were collected in the Nicolet 6700 Fourier Transform infrared spectrometer (FTIR, ThermoScientific, Waltham, MA, USA). The Smart iTR ATR sampling accessory was used. The composite films were directly placed on the ATR crystal and measured over the range of 4000–650 cm−1. For each material, 3 samples under the same conditions were examined. For each sample, 200 scans were averaged with a spectral resolution of 4 cm−1. Next, for a given material, the final average spectrum was calculated. These spectra were normalized to 1.0 at 1017 cm−1 (COH stretching vibration). The differential spectra between the CCNF/CHIT spectra and CCNFs spectrum were calculated to determine and highlight differences between composites. All the spectra manipulation was carried out using The Origin Pro 8.5 (OriginLab Corporation, USA).
Differential scanning calorimetry (DSC)
DSC analysis was performed using a TA Instruments DSC 250 system (Waters, DE, USA) and 5–10 mg of the film’s samples sealed in aluminium pans and a 20–400 °C heating increase under a nitrogen flux of 50 mL min−1; the heating rate was 10 °C min−1. Analyses were preceded by scans in the heat–cool–heat mode with a maximum temperature at 200 °C to eliminate water from the sample but not cause its decomposition. The data were analysed using Trios v.4.2.1 (TA Instruments, Waters, DE, USA) software. Enthalpy was determined as the area under the peak of the curve in the range 240–380 °C.
Mechanical properties of composites
All the composites under the study were tested for tensile strength, Young’s modulus and elongation at break. Samples of composites were prepared as rectangular strips, with a length of 40 mm and a width of ~ 2 mm. Precise measurements of the sample width were carried out using the Olympus SZX16 (Olympus Corporation, Japan) microscope with an SDF PLAPO 0.5 XPF lens, equipped with a DFK 51BU02.H digital camera (The Imaging Source Europe GmbH, Bremen, Germany). Image resolution was equal to 8.26 µm per pixel. The width of each sample was calculated as the mean value from three measurements. The thickness of each sample was measured using a digital micrometre BAKER IP54 (Baker Gauges India Private Limited, India) with a measurement accuracy equal to 0.001 mm. The composite film strips were subjected to uniaxial tensile testing using a miniature tensile stage (Deben Microtest, Suffolk, UK). The initial gap between grips was equal to 10 mm. Mechanical experiments were carried out up to sample rupture with a deformation speed of 0.2 mm min−1. Tensile force and elongation of the sample were recorded and converted into stress and strain, respectively. Stress was determined as the ratio of the tensile force to the sample cross-sectional area. The strain was defined as the ratio of the sample elongation to its initial length. The Young’s modulus was determined as the slope of the longest linear part of the stress–strain curve. Mechanical tests were repeated ten times for each composite film.
The contact angles of 5-μL water droplets on the film’s surface attached to microscopic slides by double-sided tape (T: 23.5 ± 0.1 °C; RH: 31.6%) were measured. The Rame Hart 200Std goniometer was used. Measurements were performed for 10 min with 1-s time intervals. A longer measurement did not give any reliable results, and it was decided to be completed after 10 min. Each point on the plot corresponds to the mean value of the left and right contact angle at a given point in time. Additionally, photos of droplets were taken immediately after droplet setting, after 10 and after 20 min (Supplementary Material Fig. 2).
Scanning electron microscopy (SEM)
The morphology of the CCNFs and CCNF/CHIT composite surface was examined by scanning electron microscopy (SEM; Hitachi SU3500) at 1.5 kV under high-vacuum conditions. The small cut of each sample was applied on the aluminium stage covered by carbon tape. Next, the samples were coated with an ultrathin gold layer (Au) using an ion-sputtering machine (Cressington Sputter Coater 108 Auto).
Composite antibacterial properties
To analyse the inhibition of CCNFs and CHIT composites, four bacterial species (Staphylococcus epidermidis, Escherichia coli, Bacillus cereus, and Micrococcus luteus), one yeast species (Candida krissii), and four filamentous fungal species (Botrytis cinerea, Neosartorya fischeri, Petriella setifera and Fusarium oxysporum) were selected from the microorganism collection of the Laboratory of Molecular and Environmental Microbiology, Institute of Agrophysics, Polish Academy of Sciences (Lublin, Poland). The information about the microorganisms and conditions of pre-culturing were described in Supplementary Material SM Table 1. Micrococcus luteus, yeast and all filamentous fungi were incubated at 28 °C, whereas the remaining bacterial strains were incubated at 37 °C. All microorganisms were shaken at 180 rpm for 24 h and 72 h for bacteria and fungi, respectively.
Antimicrobial assay through the inhibition zone test
The antimicrobial properties were determined by measurements of the inhibition zone of bacteria and fungi using the modified Kirby–Bauer Disk diffusion susceptibility test protocol (Hudzicki 2009). For each medium (Supplementary materials Table 1), the pH was corrected to 6 using HCl. The Petri dishes were divided into four equal parts. Each quarter was tested with CCNFs and CCNF/CHIT_1–5 films. Each composite cut with a diameter of 5 mm was prepared using a paper puncher. To confirm the antimicrobial activity of chitosan, the spreading on each medium of 300 µL 0.25% (w/v) of chitosan dissolved in broth medium or adding 0.25% (w/v) of chitosan to the medium before sterilization was used. Each treatment was prepared in three replicates. Each Petri dish was inoculated with 300 µL and 500 µL of the bacterial and fungal cell suspensions, respectively. The initial number of bacteria and fungi was approximately of 107–108 CFU mL−1, with an OD of 1 at 610 nm and 750 nm for bacteria and fungi, respectively. The OD was measured using the Infinite® M200PRO spectrophotometer (Tecan, Switzerland). Inoculated Petri dishes were incubated at 28 °C for Micrococcus luteus, yeast and all filamentous fungi and 37 °C for other bacteria. The diameter of the inhibition zone (in mm) was measured every 24 h for 4 days. For the fungi, during the experiment, the inhibition of chitosan applied alone was not observed; therefore, a higher concentration of chitosan at 1% (w/v) was tested. The procedure was the same as described above, but measurements of the inhibition zone were performed every 24 h for 7 days.
Inhibition test using the optical density (OD)
The inhibitory effects of different concentrations of CCNFs and chitosan on microorganism growth were estimated by the optical density (OD) measurements of each culture. For antimicrobial analysis, aliquots of 200 µL of each bacterial or fungal suspension (at 1 OD) were added to each well together with the film comprising different concentrations of CCNFs and CHIT or CHIT alone. Furthermore, to avoid overestimation of the OD, during plate reading, each composite (CCNF, CCNF/CHIT) and chitosan alone were loaded into separate wells without the inoculum as a negative control. The results were standardized by subtracting the OD of the composite (CCNF/CHIT) or chitosan alone from the OD of the inoculum with the different composites. The study was performed in three replicates. The optical density was read at 610 nm and 750 nm for bacteria and fungi, respectively, using the Infinite® M200PRO spectrophotometer (Tecan, Switzerland) every 24 h for 7 days.
To illustrate the significant differences, one- and two-way ANOVA were performed between the optical density and microorganisms, time and treatment. Subsequently, significant differences were calculated by post hoc analysis using Tukey’s test. Moreover, Pearson’s correlation between the concentration of chitosan, nanofibrils, and water availability of each film was assessed. All the statistical analyses were performed using STATISTICA 13.1 software (StatSoft, Inc., USA).
Structure and morphology of composites
The CCNFs used in this study was produced in the form of nanofibrils. Previously, it was shown that the average diameter of CCNFs was 3.31 nm, and the length did exceed 1 µm. The crystallinity degree, as evaluated by XRD, was 80% (Szymańska-Chargot et al. 2019).
In the chitosan films spectrum (Fig. 2a) region between 3500 and 3250 cm−1 can be assigned to the stretching vibrations of intra- and intermolecular O–H and –CH2OH, but also stretching vibrations of –NH2 (3358 cm−1) and –NH secondary amide vibrations (3288 cm−1) can be visible. Additionally, region 2960–2870 cm−1 is typical for symmetric and asymmetric vibrations of C–H (Khan et al. 2012). The peaks at 1647 cm−1, 1583 cm−1 and 1557 cm−1 are due to the amide I, primary amino group and amide II, respectively (Li et al. 2011; Khan et al. 2012).
In the case of CCNF/CHIT composites, some differences can be observed on the FTIR spectra (Fig. 2b). The main differences are related to the presence of bands originated from chitosan amine and amide group vibration: 1647, 1583 and 1557 cm−1. For pure chitosan, the intensity of 1647 and 1586 cm−1 bands were the same. In the case of CCNF/CHIT, the primary amine band at 1583 cm−1 diminished, and the band related to amide II increased. On the other hand, CCNF/CHIT_5 had higher intensity for the 1583 cm−1 than that for the 1647 cm−1 band. Moreover, in the case of CCNF/CHIT composites, the band connected to the carboxyl ester disappeared. Previously, the band at 3343 cm−1 was related to the hydrogen bonding between nanocellulose and chitosan (Khan et al. 2010). However, here, it is less likely because this band is also present in the pure CCNFs spectrum (reflecting the intramolecular hydrogen bonds in cellulose; please see ref.: Szymańska-Chargot et al. 2019), and, besides that, no visible shift of this band occurred.
Data from DSC curves corresponding to the material decomposition process
Mechanical properties of CCNF/CHIT composites
Young’s modulus (GPa)
Yield strength (MPa)
Yield strain (%)
Tensile strength (MPa)
Maximum strain (%)
14.71 ± 1.27
92.90 ± 15.05
0.83 ± 0.09
145.84 ± 37.66
1.35 ± 0.35
10.82 ± 2.00
78.65 ± 12.18
0.96 ± 0.12
141.37 ± 22.80
2.68 ± 0.80
10.89 ± 1.02
78.58 ± 10.23
0.95 ± 0.12
139.53 ± 20.27
2.08 ± 0.55
9.68 ± 0.88
79.65 ± 11.60
0.98 ± 0.17
138.50 ± 19.01
2.35 ± 0.58
8.82 ± 0.79
70.54 ± 6.36
0.97 ± 0.10
129.28 ± 15.75
2.76 ± 0.79
8.76 ± 1.10
76.37 ± 6.04
1.09 ± 0.09
129.43 ± 15.06
2.68 ± 0.68
Antimicrobial assay through the inhibition zone test
Percent of the inhibition zone with the standard deviation for each tested microorganism (n = 3)
1.95 ± 0.00
1.46 ± 0.72
1.95 ± 0.00
1.95 ± 0.00
1.95 ± 0.00
1.95 ± 0.00
1.75 ± 2.49
0.87 ± 1.40
1.26 ± 1.63
1.80 ± 2.60
1.65 ± 2.95
4.03 ± 5.64
0.00 ± 0.00
0.00 ± 0.00
0.00 ± 0.00
0.00 ± 0.00
0.00 ± 0.00
0.61 ± 1.17
35.52 ± 22.53
37.89 ± 35.63
38.63 ± 29.94
16.02 ± 8.27
29.64 ± 30.29
34.01 ± 32.10
1.26 ± 1.63
0.97 ± 1.02
1.13 ± 1.58
1.60 ± 1.23
2.48 ± 1.60
3.16 ± 2.13
1.30 ± 1.54
1.83 ± 3.40
1.46 ± 0.88
10.68 ± 15.71
3.18 ± 3.73
8.34 ± 14.19
0.81 ± 1.00
1.14 ± 1.00
1.30 ± 0.96
0.49 ± 0.88
0.49 ± 0.88
0.49 ± 0.88
0.81 ± 1.00
1.14 ± 1.00
1.14 ± 1.00
2.68 ± 2.68
0.49 ± 0.88
1.36 ± 1.90
11.63 ± 18.45
7.11 ± 6.79
2.70 ± 1.32
7.10 ± 6.81
10.48 ± 14.74
3.59 ± 2.50
Antimicrobial assay through the optical density (OD) measurement
Significant differences between the average optical density (OD) of all microorganisms for all tested composites (including nanocellulose and chitosan alone)
The composites based on nanocellulose prepared from carrot cellulose with different concentrations of chitosan from shrimp shells were prepared. The optical transparency of films was very good only for the film with the highest content of chitosan (CCNF/CHIT_5) starts to become opaque. The deterioration of the transparency, in this case, could be the result of the aggregation of CCNFs (Toivonen et al. 2015). Additionally, SEM micrographs showed no difference between composites and CCNF film surfaces, indicating the proof of obtaining a homogenous mixture of chitosan and nanocellulose. Toivonen et al. (2015) showed that nanocellulose cross-linked with chitosan formed tough films with good optical properties. The interaction between CCNFs and CHIT was evaluated in terms of FTIR spectra. Even though the mechanism of interaction between nanocellulose and chitosan is still not fully understood the most probable are hydrogen or ionic interactions, covalent imine linkage, and also hydrophobic interactions (Toivonen et al. 2015). The most pronounced changes in the FTIR spectra should be related to hydrogen bonding represented by bonds at approximately 3600–3000 cm−1 (Khan et al. 2012). However, no differences in the spectra region mentioned above were observed for the CCNF/CHIT composites and compared with the pure CCNFs and CHIT spectra. This lack of differences allows the assumption that there was likely no hydrogen bonding between nanocellulose and chitosan. On the other hand, the FTIR spectra of the CCNF/CHIT composites showed that only bands related to amide II and amide I were slightly changed. Previously, the redshift of amide bands i.e., towards lower wavenumbers was related to the interaction between nanocellulose and chitosan (Hu et al. 2015). Moreover, the disappearance of the ester band at 1740 cm−1, characteristic for oxidized CCNFs, could suggest the interaction between the oxidized surface of cellulose and amide groups of chitosan. Toivonen et al. (2015) suggested that crosslinking between cellulose nanofibres and chitosan is based on a physical interaction between the reducing ends of cellulose and primary amines of chitosan, promoting the dehydration of chitosan. This finding has been supported by the water wettability experiment. Nanocellulose is known to have poor water resistance and low water contact angle. Tangpasuthadol et al. (2003) showed that water contact angle of chitosan films is around 89 ± 6° which is much higher than value obtained in this study for CCNF films. The addition of chitosan to the nanocellulose matrix causes an increase in the water contact angle i.e., the surface of the composites become more hydrophobic. This increase could be connected to the interaction between nanocellulose and chitosan resulting in a denser structure. The degradation temperatures ca. 300 °C are similar to those obtained by Jia et al. (2017) using the TGA method. High degradation temperatures assure that the composites are not affected by the temperature (Ostadhossein et al. 2015). Additionally, none of the peak temperatures matches the temperatures of decomposition of pure CCNFs or CHIT, implying interactions between these polymers in composites (Jia et al. 2017). Moreover, the addition of chitosan to the nanocellulose matrix caused an increase in the degradation process enthalpy.
The addition of chitosan to CCNF matrices caused a decrease in Young’s moduli the higher concentration of chitosan in the composite leads to a lower Young’s modulus. The yield strength describes the maximum force that can be applied for reversible deformation. The addition of chitosan caused a decrease in the yield strength of the composites. Additionally, the tensile strength of composites i.e., the maximum force that causes a fracture was decreased after the addition of chitosan. Only the strain had the tendency to increase after chitosan addition. These results slightly differed from those that could be found in the literature. Toivonen et al. (2015) analysed the cellulose nanofibres–chitosan composites, and they found that chitosan improved the mechanical properties of the composites compared with the pure nanocellulose film. On the other hand, most of the papers reported changes in the mechanical properties of composites where the nanocellulose is additive to the chitosan matrix (Wu et al. 2014; Fernandes et al. 2010). Generally, nanocellulose was an additive of the chitosan matrix improved the mechanical properties of composites. Wu et al. (2014) found that cellulose nanofibrils incorporated in chitosan films produced by the solution casting method caused a significant increase in the tensile strength and Young’s modulus. However, Li et al. (2009) reported that, by increasing the cellulose nanowhisker concentration from 0 to 20 wt%, the tensile strength of the films raised from 85 to 120 MPa for dry composites and from 9.9 to 17.3 MPa for wet chitosan composites. It was also observed that the incorporation of cellulose nanowhiskers enhanced water resistance and the thermal stability of chitosan films.
The main purpose of using chitosan as an additive to films was to exploit its antimicrobial properties. The microorganisms chosen for antimicrobial testing were either human pathogens (Staphylococcus epidermidis, Escherichia coli, Bacillus cereus, Micrococcus luteus, Candida krissii) or plant pathogens (Botrytis cinerea, Neosartorya fischeri, Petriella setifera and Fusarium oxysporum). The greatest inhibition was observed for Escherichia coli and Staphylococcus epidermidis. Micrococcus luteus was inhibited at the highest chitosan concentration in film. Moreover, for Bacillus cereus and E. coli, modification of their growth mode was observed. Benhabiles et al. (2012) discovered that chitosan can lead to a change in the gene expression in Staphylococcus aureus SG511. Consequently, the modification in the growing modes seen in Supplementary materials Fig. 5 could be the effect of chitosan exposure. The disk diffusion analysis revealed that the combination of cellulose nanofibrils and chitosan has a strong effect on Escherichia coli and Staphylococcus epidermidis, confirming the effect of chitosan on both Gram-positive and Gram-negative bacteria (Benhabiles et al. 2012; Dutta et al. 2009; Hosseinnejad and Jafari 2016; Leceta et al. 2013; Poonguzhali et al. 2017). The fungal cultures were resistant to the chitosan presence in films. However, disk diffusion analysis showed that, after 1 day of incubation, an inhibition zone around the film was observed (Supplementary materials Fig. 4 and 6). Subsequently, this inhibition zone decreased with the experimental duration, and finally, the mycelium completely covered the film. This behaviour of the fungal culture can be connected to the fluidity of the cell membrane because the increase in the antimicrobial activity of chitosan is connected to the increase in the amount of unsaturated fatty acids in the cell membrane (Verlee et al. 2017). This outcome could be the reason for the fungal resistance to the presence of chitosan.
Additionally, chitosan dispersed in incubation medium was used as the control experiment for the antimicrobial properties of chitosan alone. The degradation of the chitosan inside the Petri dish was observed only for Botrytis cinerea, Neosartorya fischeri, and Petriella setifera (Supplementary materials Fig. 4). Chitosan degradation can be obtained through chitinases (E.C. 18.104.22.168), chitosanases (E.C. 22.214.171.124) (Zhang and Neau 2001), and exo-glucosaminidase (E.C. 126.96.36.199) (Nidheesh et al. 2015). Chitonase enzyme is produced from bacteria (e.g., Myxobacter, Sporocytophaga, Arthrobacter, Bacillus, and Streptomyces) and fungi (Rhizopus, Aspergillus, Penicillium, Chaetomium, and Basidiomycetes that are very rich in glucanase) (Gooday 1990). Thus, it could be deduced that B. cinerea, N. fischeri, and P. setifera likely produced some of these enzymes and chitosan is simply a source of carbon used by fungi, explaining their resistance to the CCNF/CHIT composites.
The optical density (OD) was used to evaluate the antimicrobial activity of the CCNF/CHIT composites. The OD value obtained for all tested microorganisms in the presence of CCNF/CHIT composites was lower than that of the control samples (Figs. 5, 6). Additionally, both concentrations of chitosan (1% and 0.25%) had OD values lower than that of the control sample (Figs. 5, 6), indicating that the composites had bacteriostatic and fungistatic activities on the analysed microorganisms; however, they did not kill analysed microorganisms completely. Particularly, during the incubation time, we saw a significant decrease in the OD value of the analysed film after 72 h. Chitosan, in contact with water, most likely exerted antimicrobial activity against all the analysed microorganisms, reducing the growth density (measured as optical density). Furthermore, only P. setifera presented a higher OD; this result could be connected to the presence of β-glucosidase. Zhang and Neau (2001) studied the β-glucosidase activity on chitosan and found that the higher the degree of deacetylation (DD) of chitosan is, the slower is its degradation. In that study, the preparation of β-glucosidase also contained chitinase. P. setifera studied here could degrade chitosan through the utilization of both enzymes. Moreover, the analysis of OD values indicates that chitosan alone has less efficiency than that of the CCNF/CHIT composite; the combination of chitosan with cellulose nanofibrils could improve the antimicrobial activity of chitosan on bacteria and fungi. Additionally, studies on the bacterio- or fungistatic properties of chitosan, which is soluble in an acidic environment, are usually conducted at low pH (Badawy and Rabea 2009). In the present study, although chitosan was dissolved in acidic solution, the CCNF/CHIT composites were neutralized (Schillinger and Lücke 1989). In this way, we analysed the real effect of chitosan-cellulose nanofibril composites on bacteria and fungi without the influence of low pH.
Films of carrot cellulose nanofibrils (CCNFs) with the addition of low-viscosity chitosan (CHIT) were prepared by the vacuum filtration method. The chitosan content in the films was varied from 9 (CCNF/CHIT_1) to 33% (CCNF/CHIT_5) of dry wt. The optical transparency of films was very good except for those with the highest content of chitosan. The deterioration of the transparency, in this case, could be the result of the aggregation of CCNFs. SEM showed that chitosan was dispersed homogeneously in the CCNF matrix. FTIR spectroscopy suggested that there was interaction between the oxidized surface of the CCNFs and the amide groups of CHIT. This finding is also supported in the water wettability experiment. The addition of chitosan to the nanocellulose matrix causes an increase in composite hydrophobicity. This increase is probably connected with the interaction between nanocellulose and chitosan to form a denser structure. Analyses of the thermal properties showed that composites are stable at high temperature, and the degradation occurred above 300 °C. It was found that the addition of chitosan to CCNF matrices caused a decrease in Young’s moduli, indicating that chitosan acts as a plasticizer and enhances the ductility of nanocellulose itself. Additionally, the tensile strength of composites decreased after the addition of chitosan. The analysed antimicrobial properties indicated the highest inhibitory effect of cellulose nanofibrils–chitosan composite films against E. coli and S. epidermidis. It should also be highlighted that this is the first work analysing the antimicrobial properties of nanocellulose–chitosan composites without the influence of acetic acid present in composites to dissolve chitosan. Therefore, we can confirm that the composites of carrot nanocellulose with the addition of chitosan has antimicrobial activity, especially fungistatic and bacteriostatic activity.
The work was partially funded by The National Centre for Research and Development–Poland (Grant No. LIDER/300/L-6/14/NCBR/2015).
MSC conceived and designed the experiment, wrote the manuscript, and obtained and analysed the FTIR and Raman spectra. MSC and MC designed and performed the experiments involving cellulose isolation and nanocellulose preparation. MC performed and analysed the DSC curves and contact angle and participated in the results and discussions. GP and MF designed, performed and discussed the results of microbiological tests. PMP performed the mechanical tests and was involved in the analysis of their results. KJC obtained SEM micrographs. AZ helped in manuscript preparation.
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Conflict of interest
The authors declared that they have no conflict of interest.
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