Biobased Nanocomposites from Layer-by-Layer Assembly of

Biomacromolecules 2010, 11, 473–480

473

Biobased Nanocomposites from Layer-by-Layer Assembly of Cellulose Nanowhiskers with Chitosan Joa˜o P. de Mesquita, Claudio L. Donnici, and Fabiano V. Pereira* Departamento de Quı´mica, Universidade Federal de Minas Gerais. Av. Antoˆnio Carlos, 6627, Pampulha, Belo Horizonte, MG. CEP 31270-901 Received October 21, 2009; Revised Manuscript Received December 15, 2009

A new biodegradable nanocomposite was obtained from layer-by-layer (LBL) technique using highly deacetylated chitosan and eucalyptus wood cellulose nanowhiskers (CNWs). Hydrogen bonds and electrostatic interactions between the negatively charged sulfate groups on the whisker surface and the ammonium groups of chitosan were the driving forces for the growth of the multilayered films. The film growth was followed by UV-vis spectroscopy through the maximum value of the absorption band at 194 nm and showed the deposition of 14.7 mg · m-2 of chitosan polymer in each cycle. Scanning electron microscopy showed high density and homogeneous distribution of CNWs adsorbed on each chitosan layer. Cross-section characterization of the assembled films indicates an average of ∼7 nm of thickness per bilayer. The results presented in this work indicate that the methodology used can be extended to different biopolymers for the design of new biobased nanocomposites in a wide range of applications such as biomedical and food packaging.

Introduction Biobased nanocomposites are a relatively new class of materials that have attracted significant attention during the past decade, mainly due to ecological and climatic factors.1 They are produced by incorporating fillers that have at least one dimension at the nanometer scale into a natural polymer matrix.1,2 Nanoparticles of inorganic3–5 or natural organic fillers6,7 have been used in different biopolymers to prepare different bionanocomposites to improve the mechanical properties of the material when compared with the pristine matrix polymers. Cellulose, the most abundant organic compound on earth, is a classical example of a natural source to produce organic nanoreinforcement elements called cellulose nanowhiskers (CNWs). Compared with inorganic fillers, the main advantages of using CNWs to formulate biobased nanocomposites include their renewable nature, low cost, low density, high specific mechanical properties, and nonabrasive nature, which allows easy processability.8 Besides these characteristics, CNWs could find economical interest because they can be prepared from a large variety of natural sources including agricultural residue such as sugar cane bagasse, rice hulls, and maize straw. In this context, CNWs appear as an alternative way to prepare new ecologically friendly biobased nanocomposites.7,9 They are highly crystalline rod-shaped nanoparticles that can be obtained from different cellulose sources through a hydrochloric or sulfuric acid hydrolysis.8 The controlled acid hydrolysis destroys the amorphous regions of the cellulose fibers first, leaving the crystalline segments intact and leading to the formation of highpurity single crystals. The use of sulfuric acid leads to a more stable dispersion than the one using hydrochloric acid due to the grafting of sulfate groups on the surface of the whiskers that stabilize the aqueous whisker suspensions by electrostatic repulsion. The length and lateral dimension of the CNWs depend mainly on the source of cellulose and acid hydrolysis conditions. * To whom correspondence [email protected].

should

be

addressed.

E-mail:

For the cotton whiskers, for instance, TEM and DLS10 techniques show an average dimension of about 250 nm in length and 15 nm in diameter and about 1200 nm in length and 15 nm in diameter for tunicate whiskers. The mechanical properties of the CNWs depend on the processing conditions and the modulus of these materials can reach values as high as 145 GPa for tunicate nanowhiskers,11–13 whereas acid-hydrolyzed nanowhiskers are thought to have a modulus around 100 GPa.14 All these important characteristics make CNWs an attractive bionanoreinforcement to the preparation of novel biodegradable nanocomposites and they generate opportunities to the development of composites for a large number of applications. However, there are still significant scientific and technological challenges to make CNWs an alternative to prepare novel bionanocomposites at the industrial level. One of the major challenges in this area is the compatibilization of the nanoreinforcements with the polymer to achieve acceptable dispersion levels of the filler within the polymeric matrix. Considering hydrophobic polymers, several attempts have been developed to achieve dispersion of the nanowhiskers in these materials including the use of surfactants15–17 or by chemical surface modification of the whiskers.11,18–21 Regarding water-soluble polymers, they represent a preferred dispersion medium due to the good stability of aqueous CNWs dispersion. Several authors described improvement of mechanical properties and good dispersion level of CNWs in hydrophilic polymers such as starch,22,23 pullulan,24 and chitosan.2,25 Unfortunately, given the properties of the CNWs,26 the results described in the literature for mechanical reinforcement for both, hydrophobic or hydrophilic polymers are far from the expected values. Chitosan, one of the strongest natural polymers, is a natural hydrophilic cationic polysaccharide derived from deacetylation of chitin, the second most abundant natural polymer on earth. Currently, chitosan is produced large-scale in several countries, and due to the ease of obtaining the polymer in different physical forms, many industrial applications have emerged.27 Important properties of chitosan are biocompatibility, biodegradability, and antibacterial properties, making this polymer suitable for

10.1021/bm9011985  2010 American Chemical Society Published on Web 01/07/2010

474

Biomacromolecules, Vol. 11, No. 2, 2010

biomedical applications such as drug delivery, tissue engineering, wound healing, and for use in antimicrobial strategies.28 Besides the biomedical applications, chitosan films have also been used for food packaging, mainly due to its nontoxicity and biodegradability.29 However, inherent water sensitivity and relatively low mechanical properties (in comparison to petroleumbased polymers), restrict the use of chitosan films for a wider range of applications, especially in moist environments. In this way, several authors have described the preparation of different bionanocomposites using chitosan as the matrix and different nanofillers for several applications, including those that need improvement on the mechanical properties and others such as for separation of substances using the barrier properties of the nanocomposite30,31 and signal amplification in sensors.32,33 Darder et al.32 promoted the intercalation of the chitosan in Namontmorilonite, through adsorption in mono- or bilayers of chitosan chains, providing compact and robust three-dimensional nanocomposites with interesting functional properties. These nanocomposites were used in the development of bulk-modified electrodes exhibiting numerous advantages such as easy surface renewal, ruggedness, and long-time stability. Liu et al.30 prepared complex membranes with nanocomposites of chitosan/ sulfonic acid functionalized silica nanoparticles, which were applied to pervaporation dehydration of ethanol-water solutions. Some authors described the preparation of chitosan/carbon nanotube nanocomposites.34–37 Wang et al.34 prepared chitosan/ multiwalled carbon nanotubes (MWNTs) nanocomposites by a simple solution evaporation method. The authors observed that the MWNTs were homogeneously dispersed throughout the chitosan matrix. When compared with neat chitosan, the mechanical properties, including the tensile modulus and strength of the nanocomposites are greatly improved for 93 and 99%, respectively, with incorporation of only 0.8 wt % of MWNTs into the chitosan matrix. Using CNWs as the nanofiller and chitosan as the matrix, Li et al.2 described the preparation of films by solvent evaporation with a good dispersion of the CNWs and the formation of strong filler-matrix interaction.2 The as-prepared nanocomposite films displayed a significant increase of the tensile strength, excellent thermal stability, and water resistance with the incorporation of cellulose whiskers. The authors attributed the results mainly to the strong interactions between the sulfate groups on whiskers surface and the ammonium groups of chitosan.2 In the layer-by-layer (LBL) technique, each layer of material is alternately deposited using components that have strong interactions between them.38,39 Because of the nature of this approach, which combines properties of different materials, the technique can be used to prepare nanocomposites with a high level of dispersion and at the same time with a high filler load. Another advantage is that the nanocomposite can be prepared with various nanostructures, such as carbon nanotubes, nanoclays, or metal nanoparticles, giving unique mechanical, optical, or electronic properties.40–45 Recently, Podsiadlo et al.39 described the preparation of high performance composite films using the LBL assembly of the montmorillonite (MTM)/ poly(vinyl alcohol) (PVA) nanocomposite. The authors concluded that a high level of ordering of the nanoscale building blocks, strong interactions between the polymer and the filler, combined with glutaraldehyde cross-link agent resulted in a high efficient load transfer between the polymer phase and the nanofiller. In another paper, Podsiadlo et al.26 reported the preparation of CNWs multilayer composites combined with a polycation, poly-(dimethyldiallylamonium chloride) (PDDA), using LBL technique. In this study the authors conclude that

de Mesquita et al.

the multilayer films presented high uniformity and dense packing of CNWs. Cranston et al. described the preparation of orientated self-assembled films using a strong magnetic field46 or spin coating technique.47 Jean B. el al.48 prepared thin films composed of alternating layers of orientated rigid cellulose nanorods and flexible polycation chains. In this work, the authors achieved alignment of the whiskers using anisotropic suspensions of the nanocrystals. More recently, Jean et al.49 prepared a green composite based on cellulose nanocrystals/xyloglucan multilayers using the nonelectrostatic, cellulose-hemicellulose interaction and gave a detailed characterization of the thin films using neutron reflectivity experiments and AFM measurements. Using chitosan and clay nanoplatelets, Podsiadlo et al.50 prepared a hybrid nanocomposite using the same technique. In this paper the authors conclude that chitosan lacks the flexibility necessary for strong adhesion and efficient load transfer between the organic matrix and the MTM platelets. The purpose of the present work is to develop a new biodegradable and biocompatible thin film combining anionic rod-like cellulose nanocrystals with chitosan, a cationic polysaccharide, using the LBL assembly technique. The hydrogen bond and electrostatic interaction between the chitosan polymer and CNWs are the driving forces for the intercalation and good interfacial adhesion between these biopolymers. The as-prepared multilayered composite film represents a strategy to maximize the interaction and the load transfer between the biopolymers and the nanofibers without having nanofillers agglomeration in the polymer matrix. In this way, this work represents a first step to preparing a high performance multilayered composite film fully obtained from biodegradable and renewable resources presenting possible biomedical applications.

Experimental Section Materials. Eucalyptus wood pulp was gently supplied by Laborato´rio de Celulose e Papel (Minas Gerais, Brazil). Highly deacetylated chitosan polymer was supplied by Phytomare Food Supplements. The degree of chitosan deacetylation (92%) was determined by potentiometric titration, Fourier transform infrared spectrometry (FTIR), and 1H NMR. Sulfuric acid, sodium hydroxide, and hydrogen peroxide were purchased from Aldrich. Acetic acid 99%, hydrochloric acid, and sodium chloride were purchased from Synth. All reagents were used without further purification. Cellulose Nanowhisker Preparation. Sulfuric acid hydrolysis of eucalyptus wood pulp was performed as described in the literature with minor modifications.51 Briefly, the wood pulp was ground until a fine particulate was obtained using a Willey mill. Then, 1.0 g of cellulose was added to 9.0 mL of 64 wt % sulfuric acid under strong mechanical stirring. Hydrolysis was performed at 45 °C for 25 min. After hydrolysis, the dispersion was diluted 10-fold, and the suspensions were then washed using two repeated centrifuge cyclings. The last washing was carried out through dialysis with deionized water until the dispersion reached pH ∼ 5. Afterward, the dispersions were sonicated (Unique Sonicator, 40 kHz) for 7 min in an ice bath to avoid overheating and finally filtered using a Whatman filter paper No. 41 (20 µm pore size filter). The final concentration of the CNWs dispersions were about 1 wt %. Layer-by-Layer Assembly. The chitosan/CNWs multilayers were prepared on substrates with a negative excess of charge, such as glass or quartz slides. The glass slides were cleaned using piranha solution (H2SO4:H2O2, 3:1 volume ratio) at room temperature for a minimum of 1 h. The quartz surfaces used to UV measurements were treated with NH4OH/H2O2/H2O (1:1:5 v/v) and HCl/H2O2/H2O (1:1:6 v/v) for 10 min at 80 °C. Each treatment was followed by intensive rinsing with Milli-Q water. The multilayer films were manually prepared immediately after substrate cleaning using the LBL technique. The

Nanocomposites from Cellulose Nanowhiskers


slides were sequentially immersed into the solutions of chitosan and CNWs, following the procedure of (1) dipping of the substrate for 10 min in a 2% w/v chitosan solution (5% acetic acid), (2) rinsing with Milli-Q water for a fixed time in three different beakers to rinse the excess of the material, (3) dipping for 10 min in a 1% w/v CNWs aqueous dispersion, and (4) rinsing with Milli-Q water with the same procedure described in (2). The entire cycle was then repeated until the desired quantities of bilayers were deposited. Instrumental Analysis. Potentiometric titration curves were performed at 25 °C using a Metrhomn 670 automatic titrator. In the titration of chitosan, 25.0 mg of the polymer was dispersed in 20.0 mL of aq HCl solution (0.0114 mol L-1) directly into the electrochemical cell and titrated with a CO2-free NaOH solution (0.0792 mol · L-1). The titration of CNWs was performed by the dilution of an appropriate volume of 1% w/v solution with CO2-free NaOH solution and titrated with HCl solution (0.0114 mol L-1). A nonlinear method for fitting acid-base potentiometric titration data was applied to determine the amount of acid groups on the surface of the CNWs.52–55 The nonlinear regression program used in this work makes use of a general equation that describes the mixture titration of a strong acid with nitrogenated weak acids.

{

f(Vi, [H+]i) ) (Vi - VHAo)Cb + [H+]i -

Kw [H+]

N

∑ (V j)1

HAn

- VHAn-1)Cb

475

Figure 1. TEM image of the eucalyptus CNWs. Scale bar is 200 nm.

}

i

(V0 + Vi) KHAn

KHAn + [H+]1

(1)

In this equation it is necessary to provide the pH values and volumes of titrant added (Vi), the initial volume present in the titration cell (V0), the value of Kw in its ionic strength, the number of surface functional groups, and initial estimates of KHAn and VHAn, which will be the adjustable parameters in the calculation. The adjustment of ionization constants and volumes of equivalence (KHAn and VHAn) were performed by a mathematical method known as Levenberg-Marquardt.56 Zeta potential (ζ-potential) of the CNWs and chitosan were measured using a Zetasizer Nano-ZS (Malvern Instruments, U.K.). The reported values are an average of 10 measurements. Fourier transform infrared spectroscopy (FTIR) of chitosan, CNWs, and the LBL films was recorded with a Nicolet 380 FT-IR spectrometer (Nicolet, MN). LBL films of chitosan/CNWs were prepared on a ZnSe window following the same procedure as with the glass slides. Films of pristine chitosan and CNWs were obtained using a solvent casting method on the ZnSe window. UV-vis spectroscopy was performed using a SHIMADZU UV 2550. Measurements of chitosan solutions were performed with 1.0 cm quartz cells, and measurements of films were made from films deposited on quartz slides. The chitosan/CNW multilayers were deposited onto the quartz substrate, and the spectrum was obtained after the deposition of each two bilayers. The amount of the material deposited in each cycle was calculated using the same procedure described by Benedetti et al.57 Transmission electron microscopy (TEM) images of CNW whiskers were taken using a FEI Tecnai G2-Spirit with 120 kV acceleration voltage. The CNW suspensions were deposited from an aqueous dispersion on a carbon-Formvar-coated copper (300-mesh) electron microscopic grid. The samples were subsequently stained with a 2% uranyl acetate solution to enhance the microscopic resolution. Scanning electron microscopy (SEM) images were taken using a Quanta 200 FEG-FEI. The SEM samples were prepared on a glass slide to study the surface and the cross section of the film. Samples were coated with a 2 nm layer of gold using a BAL-TEC MC5 010 automated sputter coater.

Results and Discussion Figure 1 shows an electron micrograph (TEM) of the eucalyptus wood CNWs. Observations of the CNWs obtained

Figure 2. CNW dispersion at 4% (w/w) concentration, observed between crossed polarizers.

from diluted suspensions (near 0.01% m/v) show individual nanocrystals and some aggregates. The appearance of laterally aggregated elementary crystallites in TEM images is expected due to the high specific area and strong hydrogen bonds established between the whiskers. These aggregates may exist even in suspension but, when the dispersing medium is removed, as in the case of the TEM sample preparation, bundles of whiskers can be even more numerous than individualized rods.58 From several TEM images, the mean values of the length (L) and diameter (D) of the isolated CNWs were determined to be 145 ( 25 nm and 6 ( 1.5 nm, respectively, giving an aspect ratio (L/D) of around 24. Figure 2 shows typical flow birefringence found in the CNW suspensions at 4% (w/w) concentration, observed between two crossed polarizers. The appearance of flow birefringence, which results from induced alignment of the rodlike particles under flow, indicates the presence of isolated cellulose whiskers in the dispersion.59 Generally, the LBL method is based on the alternating sequential adsorption of polyelectrolytes of opposite charge. In this way, surface charge density of both polyelectrolytes is an important parameter for multilayer buildup in polyelectrolyte systems. This parameter can determine some characteristics of the nanocomposite, such as cohesive strength between the polymers and surface roughness of the film. Furthermore, for

476


de Mesquita et al.

Figure 3. Potentiometric titration curves obtained for (a) CNWs and (b) chitosan. Table 1. Characterization of Surface Charge Density for Both Polyelectrolytes Obtained from Potentiometric Titration and Zeta Potential Measurements chitosan -1

total anionic sites/mmol · kg total cationic sites/mmol · kg-1 ζ-potential/mV charge densitya/e · nm-2

CNWs 47

5000 +60 b

-32 0.068

a Charge density values were obtained considering CNWs as cylinders 145 nm in length and 6 nm in diameter. b Not determined.

CNWs, the amount of sulfate anionic groups provides electrostatic stabilization in aqueous dispersion. It is known that insufficient hydrolysis of cellulose may result in lower surface charge, larger particles, and therefore less surface area.51 Besides the occurrence of electrostatic interaction, it is important to mention at this point that the formation of hydrogen bonds between chitosan and CNWs plays an important role in the growth of this film. The formation of hydrogen bonds in this system will be discussed later in the text. Potentiometric titration and zeta potential were used to characterize quantitatively the surface charge density of both polyelectrolytes. Figure 3 shows the potentiometric titration curves obtained for chitosan and CNWs. The origin of the cationic sites of chitosan are the protonated NH2 groups (pH = 3), whereas for the CNWs the density of the negative charge is due to the presence of sulfate ester groups (-O-SO3-) introduced during the acid hydrolysis by the esterification of hydroxyl or carboxylic groups.60 Table 1 shows the amount of cationic and anionic groups for the chitosan and CNWs, respectively. With the average particle size of the CNWs and the amount of anionic groups, the charge density value was estimated as 0.068 e · nm-2 (Table 1). The nanowhiskers obtained in this work are weakly charged in comparison to the usually reported values.61 However, some authors have measured similar values for the amount of anionic groups present on the CNWs surface.60,62 Moreover, this amount was large enough to generate stable water suspensions, as shown in Figure 2. In addition, the ζ-potential of CNWs in water (Table 1) also indicates a stable suspension, and this value is similar to that found by Hasani et al. for CNWs obtained from cotton.61 As expected, the ζ-potential of chitosan solution is greater than the CNWs due to the large amounts of -NH3+ functional groups present in this polymer. Before the growth of chitosan/CNWs films, the optimum adsorption time was estimated to be five minutes. This estima-

tion was based on UV-vis measurements using different dipping times. From 5 min of dipping we found almost equal amount of deposition in each cycle (∼14.7 mg · m-2 of chitosan) to 10 or 15 min. However, just to ensure the equilibrium in each cycle, we choose 10 min for the deposition time. The growth of the chitosan/CNWs LBL assembly was followed by the absorbance measurements at 194 nm, typical for chitosan polymer (Figure S1). Chitosan is composed of two far UV chromophoric groups, N-acetylglucosamine and glucosamine. The two monomer units contribute in a simple, additive way to the total absorbance of the material at a particular wavelength (190-220 nm). However, the absorptivity of the acetyl-glucosamine monomer is greater.63,64 The UV-vis absorbance spectrum was obtained at each two bilayers deposited on the quartz substrate and showed a linear increase of the absorbance at 194 nm (Figure 4a). This result indicates the deposition of the same amount of material in each cycle. A molar absorption coefficient of 0.61 m2 · g-1 (Figure S2) was obtained from measurements of different concentrations of chitosan solutions, and this value was used to calculate the mass of the polymer deposited on each layer. Remembering that the bilayers are deposited on both sides of a quartz slide, we calculated the amount of chitosan adsorbed in each bilayer, and the result is shown in Figure 4. From the slope of this plot, it was estimated that the mass of chitosan per layer was 14.7 mg · m-2. Moreover, the amount of chitosan polymer is related to the slope of the plot in Figure 4. The measured absorbance of the adsorbed chitosan increases rapidly in this system, indicating rapid LBL deposition and high loading of the polymer in each cycle. A high loading of chitosan on each layer plays a role in the quantity of CNWs adsorbed and the formation of a dense coverage of cellulose nanowhiskers. The dense packing of nanofibers which can be seen on the multilayered film surface will be discussed later (Figure 7). The amount of surface charge density of the CNWs is quite small if compared to the ammonium groups on chitosan (Table 1). The large difference in surface charge density between CNWs and positively charged polyelectrolytes was already considered by Jean et al.48 The authors stated that the polymer adsorption occurs after the adsorption of two layers of the nanowhiskers and subsequent multilayer growth occurs even when charge reversal is not reached. In this case, the author stated that this may be possible because the outer surface of



477

Figure 4. (a) Absorbance at 194 nm as a function of the number of bilayers of chitosan/CNWs; (b) amount of chitosan deposited in each cycle.

the nanocrystals most probably remains negatively charged after adsorption on the polymer surface. It may be the case in the system studied in this work because charge reversal is not possible with only one CNW layer. Additionally, the formation of hydrogen bonds between cellulose nanowhiskers and chitosan is highly expected, given the chemical structure of these polysaccharides. To evaluate and identify the interactions between the chitosan polymer and the CNWs in the nanocomposites, we prepared the LBL films using a ZnSe window as substrate. For pure chitosan and CNW films, the samples were obtained by solvent evaporation on the same substrate. Figure 5 shows the FTIR spectra of cellulose nanowhiskers, the chitosan, and the nanocomposite film having 30 bilayers. The main bands that appear in the spectrum of chitosan are located around 1643 and 1558 cm-1. The first one corresponds to the CdO stretching of acetyl groups and the second one arises from N-H bending vibrations (amide and amine groups). The bands at 1070 and 1030 cm-1 are due to the C-O stretching vibrations, whereas the broadband at 3200-3450 is assigned to the O-H and N-H stretching. The peaks at 1375 and 1450 cm-1 are attributed to the -CH3 groups of N-acetylglucosamine residue and -CH2 groups, respectively.65,66 In pure CNWs film, the band at 3340 cm-1 is attributed to the O-H stretching vibration. The bands at 2893 and 1431 cm-1 are characteristic of C-H stretching and bending of -CH2 groups, respectively, whereas the peaks at 1160 and 1070 cm-1 are attributed to the saccharide structure.2 The sulfate groups were not detected because the amount of these groups was very small. The spectrum of the LBL nanocomposite shows characteristic bands of both materials. The saccharide and the C-H stretching bands could be observed in the spectrum of the nanocomposite film as well as the bands related to CdO stretching and N-H bending. The broadband around 3200-3450 cm-1 in the nanocomposite moved slightly and became broader compared to the spectrum obtained for pure CNWs film. Zhang et al.2 stated that these effects are due to the interaction between the negatively charged sulfate groups on the whisker surface and the ammonium groups of chitosan. Besides these interactions, the formation of hydrogen bonds between cellulose nanowhiskers and chitosan is highly expected given the chemical structure of these polysaccharides. Moreover, the number of sulfate groups on the whiskers surface is low compared to the number of ammonium groups on chitosan, leading to a low number of electrostatic interactions. In this way, we conclude

Figure 5. FTIR of the CNWs (I), chitosan/CNWs nanocomposite film with 30 bilayers (II), and pure chitosan (III). (a) FTIR spectrum from 500 to 4000 cm-1. (b) Close-up of the spectrum showing the displacement of the N-H bending in the nanocomposite from 1558 to 1530 cm-1.

that this displacement is also due to the hydrogen bonds between the OH groups of the cellulose nanowhiskers and the ammonium groups of chitosan. Moreover, compared to pure chitosan film,

478


Figure 6. Thickness characterization of a 20-bilayer film at 40.000 (a) and 80.0000× (b) magnifications. The white arrow in (b) indicates the span of the thickness.

the band corresponds to N-H bending of the nanocomposite moved to lower wavenumbers (1558 f 1530 cm-1), Figure 5b, suggesting also the formation of hydrogen bonds between chitosan and CNWs. Another evidence for the hydrogen bond formation is the remarkable change in the bands attributed to

de Mesquita et al.

N-H and O-H stretching from the pure chitosan as starting material, which shows a very broadband, from 3400 to 3000 cm-1 due to well-known intermolecular interactions among chitosan units. The nanocomposite chitosan/CNWs shows a narrower band (in comparison to pure chitosan) in the same region, more similar to the pure cellulose nanowhisker, due to the formation of these intermolecular interactions between chitosan and the nanowhiskers. Thickness characterization of the assembled films was made using scanning electron microscopy (SEM). The glass slides with LBL films were cut using a diamond blade and the SEM images were taken on the opposite side of the cut just to ensure a flat (i.e., not folded) sample. The obtained images, in fact, were not exactly a cross-section because the sample was inclined 10° with respect to the horizontal plane. Figure 6a,b shows the thickness characterization of a 20-bilayer film grown on a glass substrate. The images were taken from different regions of the film, and Figure 6b is a close-up of the lateral view of the film. SEM characterization revealed uniform thickness of 140 nm for the 20-bilayers film, indicating an average of ∼7 nm of thickness per bilayer (Figure 6b). We can also observe a dense packing of the layers and a planar orientation of the CNWs. Some nonplanar CNWs that can be seen at the edge are due to the shearing force resulting from cutting the film during sample preparation generating comb-hair morphology. Characterization of the assembly grown on a glass slide using atomic force microscopy (AFM) reveals a uniform and relatively smooth surface because the root-mean-squared (rms) roughness values are lower than 11 nm (Figure S3). Figure 7 shows typical SEM images of the surface of the thin film with 20 bilayers grown on glass substrate. The images clearly show a dense packing and uniform nanofibers coverage on the multilayered film surface. A dense packing of the CNWs has been previously described in other systems.26,48,49 The surface of the film also shows a morphology made by laterally aggregated nanowhiskers generating a morphology that looks like a bundle of spaghetti-like fibers. The formation of this type of morphology is possible because the nature of the cellulose whisker nanofibers. Recently, Rochas et al.58 studied three different sources of CNWs and found that these structures can be described as twisted ribbons presenting some lateral aggregation even in solution. The characteristics of the CNWs

Figure 7. Top-down view of a 20-bilayer chitosan/CNWs film on glass substrate at 50.000 (a) and 100.000× (b) magnifications.


described in the literature together with the morphology observed in Figure 6 can play a role in the growth of the LBL films if we compare cellulose nanorods with rigid clay platelets. The well-known lack of flexibility of chitosan polymer was described in the literature50 as a disadvantage to the development of LBL films with clay nanosheets. High rigidity of the chitosan chains does not allow them to acquire a conformation necessary for maximizing the interfacial adhesion with the clay platelets. We believe that using cellulose nanowhiskers as nanofiller, even as rigid rod-like crystalline objects, the presence of twisted ribbons presenting different numbers of lateral aggregations between them may maximize the nanofiller-polymer interactions. Moreover, the formation of a porous network structure of cellulose whiskers that can clearly be seen in Figure 7 increases the contact area and the interfacial adhesion between the rigid polymer and the nanofillers. It is important to remember that the surface of the multilayered film can be thought as representative of each CNWs layer intercalated with the chitosan layers. As a result, we believe that due to the well-known lack of flexibility of chitosan biopolymer, the CNWs could be a better choice than clay platelets as nanofiller for the preparation of multilayered nanocomposite films. Additionally, some parameters could be changed for the preparation of these thin films, such as pH values and ionic strength of chitosan medium, in order to have a more flexible chain, maximizing the number of polymer/nanofiller interactions. The effects of these parameters in the film growing are under investigation and will be studied in future work.

Conclusion Layer-by-layer assembly technique was successfully used to prepare a new biodegradable and biocompatible nanocomposite based on alternating layers of two renewable biobased materials: chitosan polymer and cellulose nanowhiskers. The length and diameter of the CNWs were characterized as being 145 ( 25 and 6 ( 1.5 nm, respectively. Potentiometric titration gave 47 mmol · kg-1 of sulfates groups on the whiskers surface, which were sufficient to generate stable suspensions with ζ-potential ) -32 mV. Chitosan and CNWs presented interaction due to the formation of hydrogen bonds and also electrostatic interaction which occurred between the sulfate groups on the whisker surface and the ammonium groups of chitosan. The film surface was characterized as a relatively smooth surface having the roughness values lower than 11 nm. Moreover, the use of the LBL technique could maximize the interaction between both components and allowed the incorporation of high amounts of the nanofillers, which presented a dense and homogeneous distribution of the cellulose nanowhiskers in each layer. The average thickness of a single bilayer of the film was found to be 7.0 nm and the amount of chitosan deposited in each cycle was 14.7 mg · m-2. Each layer of CNWs also showed a structure formed by laterally aggregated nanowhiskers originating a porous morphology that looks like a bundle of spaghetti-like fibers. Mechanical, thermal, and barrier properties of the freestanding films are under investigation and will be studied in the next work. Finally, given the initial results presented here, the preparation of multilayered LBL nanocomposites using CNWs as nanofillers and a biodegradable polymer as matrix can potentially be used for the development of high performance materials fully based on renewable materials. The methodology can be extended using different biopolymers for the design of new biobased nano-


479

composites in a wide range of applications such as biomedical and food packaging. Acknowledgment. Authors thank CNPq, Capes (NanobiotecEDT No. 04/2008), FAPEMIG (PRONEX EDT 479/07, PPM III 0207/09) for financial support. J.P.d.M. thanks CNPq for the scholarship. Centro de Microscopia-UFMG is also gratefully acknowledged for TEM and SEM images. We also thank Maximiliano D. Martins for AFM images. Supporting Information Available. UV-vis absorbance measurements and tapping mode AFM height images of 20bilayer chitosan/CNWs film on a glass substrate. This material is available free of charge via the Internet at http://pubs.acs.org.

References and Notes (1) Petersson, L.; Oksman, K. Compos. Sci. Technol. 2006, 66, 2187– 2196. (2) Li, Q.; Zhou, J.; Zhang, L. J. Polym. Sci., Part B: Polym. Phys. 2009, 47, 1069–1077. (3) Lu, Y.; Larock, R. C. Biomacromolecules 2006, 7, 2692–2700. (4) Lu, J.; Hong, C. K.; Wool, R. P. J. Polym. Sci., Part B: Polym. Phys. 2004, 42, 1441–1450. (5) Miyagawa, H.; Misra, M.; Drzala, L. T.; Mohanty, A. K. Polymer 2005, 46, 445–453. (6) Petersson, L.; Kvien, I.; Oksman, K. Compos. Sci. Technol. 2007, 67, 2535–2544. (7) Jiang, L.; Morelius, E.; Zhang, J.; Wolcott, M.; Holbery, J. J. Compos. Mater. 2008, 42, 2629–2645. (8) Samir, M. A. S. A.; Alloin, F.; Dufresne, A. Biomacromolecules 2005, 6, 612–626. (9) Oksman, K.; Mathew, A. P.; Bondeson, D.; Kvien, I. Compos. Sci. Technol. 2006, 66, 2776–2784. (10) Lima, M. M. D. S.; Wong, J. T.; Paillet, M.; Borsali, R.; Pecora, R. Langmuir 2003, 19, 24–29. (11) Braun, B.; Dorgan, J. R. Biomacromolecules 2009, 10, 334–341. (12) Sy¨turcova, A.; Davies, G. R.; Eichhorn, S. J. Biomacromolecules 2005, 6, 1055–1061. (13) Iwamoto, S.; Kai, W.; Isogai, A.; Iwata, T. Biomacromolecules 2009, 10, 2571–2576. (14) Rusli, R.; Eichhorn, S. J. Appl. Phys. Lett. 2008, 93, 033111.1033111.3. (15) Heux, L.; Chauve, G.; Bonini, C. Langmuir 2000, 16, 8210–8212. (16) Ljungberg, N.; Bonini, C.; Bortolussi, F.; Boisson, C.; Heux, L.; Cavaille, J. Y. Biomacromolecules 2005, 6, 2732–2739. (17) Ljungberg, N.; Cavaille, J. Y.; Heux, L. Polymer 2006, 47, 6285– 6292. (18) Yuan, H.; Nishiyama, Y.; Wada, M.; Kuga, S. Biomacromolecules 2006, 7, 696–700. (19) Nair, K. G.; Dufresne, A.; Gandini, A.; Belgacem, M. N. Biomacromolecules 2003, 4, 1835–1842. (20) Angellier, H.; Boisseau, S. M.; Belgacem, M. N.; Dufresne, A. Langmuir 2005, 21, 2425–2433. (21) Andresen, M.; Johansson, L. S.; Tanem, B. S.; Stenius, P. Cellulose 2006, 13, 665. (22) Angle`s, M. N.; Dufresne, A. Macromolecules 2000, 33, 8344–8353. (23) Mathew, A. P.; Dufresne, A. Biomacromolecules 2002, 3, 609–617. (24) Kristo, E.; Biliaderis, C. G. Carbohydr. Polym. 2007, 68, 146–158. (25) Sriupayoa, J.; Supaphola, P.; Blackwell, J.; Rujiravanita, R. Carbohydr. Polym. 2005, 62, 130–136. (26) Podsiadlo, P.; Choi, S. Y.; Shim, B.; Lee, J.; Cuddihy, M.; Kotov, N. A. Biomacromolecules 2005, 6, 2914–2918. (27) Prashanth, K. V. H.; Tharanathan, R. N. Trends Food Sci. Technol. 2007, 18, 117–131. (28) Laranjeira, M. C. M.; Fa´vere, V. T. Quim. NoVa 2009, 32, 672–678. (29) Azeredo, H. M. C. Food Res. Int. 2009, 42, 1240–1253. (30) Liu, Y. L.; Hsu, C. Y.; Su, Y. H.; Lai, J. Y. Biomacromolecules 2005, 6, 368–373. (31) Tang, C.; Chen, N.; Zhang, Q.; Wang, K.; Fu, Q.; Zhang, X. Polym. Degrad. Stab. 2009, 94, 124–131. (32) Darder, M.; Colilla, M.; Hitzky, E. R. Chem. Mater. 2003, 15, 3774– 3780. (33) Darder, M.; Lo´pez-Blanco, M.; Aranda, P.; Aznar, A. J.; Bravo, J.; Hitzky, E. R. Chem. Mater. 2006, 18, 1602–1610. (34) Wang, S. F.; Shen, L.; Zhang, W. D.; Tong, Y. J. Biomacromolecules 2005, 6, 3067–3072.

480


(35) Carson, L.; Brown, C. K.; Stewart, M.; Oki, A.; Regisford, G.; Luo, Z.; Bakhmutov, V. I. Mater. Lett. 2009, 3, 617–620. (36) Shieh, Y. T.; Yang, Y. F. Eur. Polym. J. 2006, 42, 3162–3170. (37) Liu, Y. L.; Chen, W. H.; Chang, Y. H. Carbohydr. Polym. 2009, 76, 232–238. (38) Decher, G. Science 1997, 277, 1232–1237. (39) Podsiadlo, P.; Kaushik, A. K.; Arruda, E. M.; Waas, A. M.; Shim, B. S.; Xu, J.; Nandivada, H.; Pumplin, B. G.; Lahann, J.; Ramamoorthy, A.; Kotov, N. A. Science 2007, 318, 80. (40) Jiang, C.; Rybak, B. M.; Markutsya, S.; Kladitis, P. E.; Tsukruk, V. V. Appl. Phys. Lett. 2005, 86 (1-3), 121912. (41) Markutsya, S.; Jiang, C.; Pikus, Y.; Tsukruk, V. V. AdV. Funct. Mater. 2005, 15, 771–780. (42) Krasteva, N.; Besnard, I.; Guse, B.; Bauer, R. E.; Mu¨llen, K.; Yasuda, A.; Vossmeyer, T. Nano Lett. 2002, 2, 551–555. (43) Kidambi, S.; Bruening, M. L. Chem. Mater. 2005, 17, 301–307. (44) Lu, Z.; Prouty, M. D.; Guo, Z.; Golub, V. O.; Kumar, C. S.; Lvov, Y. M. Langmuir 2005, 21, 2042–2050. (45) Podsiadlo, P.; Sui, L.; Elkasabi, Y.; Burgardt, P.; Lee, J.; Miryala, A.; Kusumaatmaja, W.; Carman, M. R.; Shtein, M.; Kieffer, J.; Lahann, J.; Kotov, N. A. Langmuir 2007, 23, 7901–7906. (46) Cranston, E. D.; Gray, D. G. Sci. Technol. AdV. Mater. 2006, 7, 319– 321. (47) Cranston, E. D.; Gray, D. G. Biomacromolecules 2006, 7, 2522–2530. (48) Jean, B.; Dubreuil, F.; Heux, L.; Cousin, F. Langmuir 2008, 24, 3452– 3458. (49) Jean, B.; Heux, L.; Dubreuil, F.; Chambat, G.; Cousin, F. Langmuir 2009, 25, 3920–3923. (50) Podsiadlo, P.; Tang, Z.; Shim, B. S.; Kotov, N. A. Nano Lett. 2007, 7, 1224–1231. (51) Candanedo, S. B.; Roman, M.; Gray, D. G. Biomacromolecules 2005, 6, 1048–1054.

de Mesquita et al. (52) Masini, J. C.; Abate, G.; Lima, E. C.; Hahn, L. C.; Nakamura, M. S.; Lichtig, J.; Nagatomy, H. R. Anal. Chim. Acta 1998, 364, 223–233. (53) Masini, J. C. Anal. Chim. Acta 1993, 283, 803–810. (54) Mesquita, J. P.; Martelli, P. B.; Gorgulho, H. F. J. Braz. Chem. Soc. 2006, 17, 1133–1143. (55) Gorgulho, H. F.; Mesquita, J. P.; Goncalves, F.; Pereira, M. F. R.; Figueiredo, J. L. Carbon 2008, 46, 1544–1555. (56) Chapra, S. C., Cnale, R. P. Numerical Methods for Engineers: With Programming and Software Applications; New York: McGraw Hill, 1998. (57) Benedetti, T. M.; Bazito, F. F. C.; Ponzio, E. A.; Torresi, R. M. Langmuir 2008, 24, 3602–3610. (58) Elazzouzi-Hafraoui, S.; Nishiyama, Y.; Putaux, J. L.; Heux, L.; Dubreuil, F.; Rochas, C. Biomacromolecules 2008, 9, 57–65. (59) Bondeson, D.; Mathew, A.; Oksman, K. Cellulose 2006, 13, 171– 180. (60) Araki, J.; Wada, M.; Kuga, S.; Okano, T. Colloids Surf., A 1998, 142, 75–82. (61) Hasani, M.; Cranston, E. D.; Westman, G.; Gray, D. G. Soft Matter 2008, 4, 2238–2244. (62) Dong, X. M.; Kimura, T.; Revol, J. F.; Gray, D. G. Langmuir 1996, 12, 2076–2082. (63) Liu, D.; Wei, Y.; Yao, P.; Jiang, L. Carbohydr. Res. 2006, 341, 782– 785. (64) Wu, T.; Zivanovic, S. Carbohydr. Polym. 2008, 73, 248–253. (65) Pavia, D. L., Lampman, G. M., Kriz, G. S. Introduction to Spectroscopy. A Guide for Students of Organic Chemistry; Thomson Learning: Mexico, 2001. (66) Shigemasa, Y.; Matsuura, H.; Sashiwa, H.; Saimoto, H. Int. J. Biol. Macromol. 1996, 18, 237–242.

BM9011985

Biobased Nanocomposites from Layer-by-Layer Assembly of

Recommend Documents