Nanostructural Reorganization of Bacterial ... - ACS Publications

Biomacromolecules 2010, 11, 1217–1224

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Nanostructural Reorganization of Bacterial Cellulose by Ultrasonic Treatment Paula C. S. Faria Tischer,*,† Maria Rita Sierakowski,† Harry Westfahl, Jr.,*,‡ and Cesar Augusto Tischer† Laborato´rio de Biopolı´meros, Universidade Federal do Parana´ (UFPR), Caixa Postal 19081, 81531-990, Curitiba, Parana´, Brazil, and Laborato´rio Nacional de Luz Sıńcrotron (LNLS), CEP 13083-970, Caixa Postal. 6192, Campinas, SP, Brazil Received December 4, 2009; Revised Manuscript Received March 16, 2010

In this work, bacterial cellulose was subjected to a high-power ultrasonic treatment for different time intervals. The morphological analysis, scanning electron microscopy, and atomic force microscopy revealed that this treatment changed the width and height of the microfibrillar ribbons and roughness of their surface, originating films with new nanostructures. Differential thermal analysis showed a higher thermal stability for ultrasonicated samples with a pyrolysis onset temperature of 208 °C for native bacterial cellulose and 250 and 268 °C for the modified samples. The small-angle X-ray scattering experiments demonstrated that the treatment with ultrasound increased the thickness of the ribbons, while wide-angle X-ray scattering experiments demonstrated that the average crystallite dimension and the degree of crystallinity also increased. A model is proposed where the thicker ribbons and crystallites result from the fusion of neighboring ribbons due to cavitation effects.

Introduction Cellulose is one of the most important biopolymers and is widely used based on its availability, biocompatibility, biological degradability, and sustainable production.1 In particular, bacterial cellulose is a pure biopolymer, without collateral biogenic compounds like lignin, hemicelluloses, and pectin, and it is an attractive biopolymer due to its facile production and purification.2 The structure of bacterial cellulose (BC) was described more than a century ago,3,4 and the material has been extensively analyzed and characterized.5-9 Bacterial cellulose has been produced from Acetobacter xylinum by fermentation of different substrates as glucose from corn syrup,10-12 and more recently, rice bark supplemented with glucose was used as a substrate to produce cellulose nanospheres.13 Cellulose molecules form long chains in polycrystalline fibrous bundles, which contain crystalline and amorphous regions, and the degree of crystallinity and crystal dimensions are dependent on the origin of the cellulose. The biosynthetic process is the same in all organisms, but there are some differences in the cellulose synthase complexes that determine the size and thickness of the cellulose microfibrils, and the great interest in cellulose macromolecules is due to their crystalline orientation.14 The microstructures formed by the ultrafine microfibrils of bacterial cellulose have lengths varying from 1 to 9 µm and create a dense reticulated structure stabilized by various hydrogen bonds. These networks show a high index of crystallinity (above 60%) and a higher degree of polymerization, normally around 16.000 or 20.000,15 in comparison with vegetal cellulose. * To whom correspondence should be addressed. Phone: +55 41 33613260 (P.C.S.F.); +55 19 3512-1034 (H.W.). E-mail: [email protected] (P.C.S.F.); [email protected] (H.W.). † Universidade Federal do Parana´ (UFPR). ‡ Laborato´rio Nacional de Luz Sıńcrotron (LNLS).

The high crystallinity of bacterial cellulose is the result of a high number of inter- and intrahydrogen bonds between adjacent chains of glucan. These bonds create a regular crystalline arrangement between the glucan molecules,16 resulting in the distinct diffraction pattern, swelling, and reactivity of cellulose. One of the methods used to obtain microcrystals of cellulose is to submit cellulose to controlled acid (H2SO4) hydrolysis conditions. The hydronium ions can penetrate the cellulose chains in the amorphous domains, promoting the hydrolytic cleavage of the glycosidic bonds and releasing individual crystallites. In this process, sulfate groups are randomly distributed on the cellulose microfibril surface, inducing a negative electrostatic layer covering the microfibers.20-22 The cellulose microcrystals, because of their stiffness, thickness, and length, are commonly called “whiskers”.17-19 For pursuing applications that require extensively entangled networks and higher strength, chemically less aggressive hydrolysis concepts have been found to maintain a high aspect ratio of the cellulose I fibrils or fibril aggregates, potentially allowing strong or even permanent junction points for the networks. A classic example is provided by omitting the hydrolysis step and by solely imposing high shearing forces for disintegration. This process yields a highly entangled network, which typically consists of elements with a wide size distribution down to the nanoscale. The resulting material has been denoted microfibrillated cellulose (MFC), originally introduced by Turback et al.23 and Herrick et al.24 By this method, the cellulose forms a highly entangled network with a low degree of crystallinity, and some of the noncrystalline domains remain intact. This method has the same problems, and the increased energy of tension necessary to cause the disintegration of fibers requires several steps of treatment. Recently, a novel route was demonstrated that combines enzymatic hydrolysis and mechanical shearing to form a long and highly entangled network of cellulose I elements.25 Another method cited to prepare spherical nanoparticles of cellulose is based on a hard pretreatment of cellulose: heating

10.1021/bm901383a  2010 American Chemical Society Published on Web 04/06/2010

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at 80 °C in 5.0 M NaOH for 3 h, followed by the resuspension of fibers in DMSO in a water bath at 80 °C for 3 h. Next, the pretreated fibrils are suspended in an acid solution and sonicated with heating at 80 °C for 8 h. By this process, nanospheres of 60-570 nm composed of cellulose II are obtained.26 All these methods were realized with vegetal cellulose, and hard conditions for hydrolysis were used, which caused intense degradation of the polymer and a consequent reduction in crystallinity. The effect of ultrasound in degrading polysaccharide linkages is well described. Studies with chitosan,27-30 dextran,31-33 agarose and carrageenans,34 xyloglucan,35 cellulose derivatives, and carboxymethylcellulose36-38 have been described, and in the majority of cases, the treatment with ultrasound has been used in acid solutions with relatively low temperature and in alkaline39 or acidic conditions.40 The energy of ultrasound is transferred to the polymer chains through a process called cavitation, which is the formation, growth, and violent collapse of cavities in the water. The energy provided by cavitation in this so-called sonochemistry is approximately 10-100 kJ/mol,41 which is within the hydrogen bond energy scale.42 The main goal of this work was to analyze the effect of ultrasonic treatment on the reorganization of bacterial cellulose microfibrils using suitable conditions (aqueous medium, without acid or heating). Such results are needed to develop an easy method to obtain films with different nanostructures and characteristics (porosity, roughness, and crystallinity) and to develop a process in nanotechnology that permits the formation of new scaffolds for tissue engineering.

Materials and Methods Microorganism, Culture Media, and Growth Conditions. The bacterial strain Acetobacter xylinum ATCC 23769 (reclassified as the genus Gluconacetobacter), supplied by Foundation Andre´ Tosello from Campinas, Saõ Paulo, was grown in a glucose medium based on the Hestrin-Schramm’s medium culture.43 All media were autoclaved at 121 °C and 1.02 atm for 20 min. The inoculum was prepared in a nonagitated 200-mL Erlenmeyer flask containing about 5 mL of the thawed culture and 40 mL of the above medium. The flask was incubated at 28 ( 1 °C for 48 h. The obtained cell suspension was used as the inoculum. The static fermentation was carried out for 10 days at 28 °C, and the pH was adjusted to 5.25 with citric acid (5%, w/v). Treatment of Bacterial Cellulose. The bacterial cellulose (BC) pellicle that formed at the liquid-air interface of the fermentation medium was removed, rinsed thoroughly with deionized water, and purified by immersion in an aqueous solution of 0.1 M NaOH for one day. The films were washed repeatedly with deionized water and then vacuum-dried at 40 °C. This treatment was controlled to avoid the mercerization of the BC. Ultrasonic Process. The pellicles of the BC after vacuum drying at 40 °C were cut with scissors. The cut films (200 mg, immersed in 200 mL Mili-Q water) were shaken vigorously for 15 min. This material was submitted to ultrasonic treatment for 15, 30, 60, and 75 min using an ultrasonic processor, Ultrasound SONICS (200 W, 20 kHz). The ultrasonic treatment was carried out in an ice bath, and the ice was maintained throughout the entire sonication time. After this process, the ultrasonicated BC was reconstituted in the form of films and vacuum-dried at 40 °C. In all of the analysis techniques, the stable films formed after ultrasonication were used in the solid state as whole films or cut films. Small-Angle X-ray Diffraction (SAXS). The SAXS patterns were used to determine the dimensions of the microfibrils in the cellulose. SAXS diagrams were recorded on the D02A beamline using a fixed

Tischer et al. radiation wavelength (λ ) 1.488 Å) at the Brazilian Synchrotron Light Laboratory (LNLS) also in the transmission geometry. The samples were thin films, supported in the sample holder and kept in a 10-2 mbar vacuum. Scattering patterns were recorded during 300 s exposures on a marCCD 165 detector (8 × 8 binning), placed first at 2507 mm and then at 1484 mm away from the samples. Calibration of the distances was achieved using a silver Behenate pattern.44 The SAXS profiles from the samples consisted of isotropic scattering patterns that were subtracted from the CCD bias and darknoise according to a reference.45 The SAXS chamber parasitic scattering was also recorded (with bias and dark-noise subtraction) and subtracted from the sample pattern after sample attenuation correction. The final patterns were normalized by the phase space volume and azimuthally averaged for a sequence of wave-vectors q (q ) (4π/λ) sin(θ); 2θ ) scattering angle) ranging from 0.04 to 1.14 nm-1 when the detector was at the longer distance and 0.1 to 2.3 nm-1 for the shorter one. Wide-Angle X-ray Diffraction (WAXS). The WAXS patterns were used to determine the relative crystallinity. WAXS diagrams were recorded on the D03B beamline using a fixed radiation wavelength (λ ) 1.433 Å) at the LNLS in the transmission geometry. The samples were thin films supported in 1 cm aluminum rings aligned perpendicularly to the beam. Diffraction patterns were recorded during 120 s exposures on a MarCCD 165 detector (2 × 2 binning), placed 100.0 mm away from the samples. Calibration of the distances was achieved using a LaB6 pattern. The WAXS profiles from the samples consisted of circular Debye-Scherrer rings and were subtracted from the CCD bias and dark-noise according to a reference.45 The air scattering was also recorded (with bias and dark-noise subtraction) and subtracted from the sample pattern after sample attenuation correction. The final patterns were normalized by the phase space volume and azimuthally averaged for a sequence of wave-vectors q (q ) (4π/λ) sin(θ); 2θ ) scattering angle) ranging from 1.4 to 28.6 nm-1. Scanning Electron Microscopy (SEM) Analysis. Scanning electron microscopy (SEM) was conducted to observe the surface of the bacterial cellulose films before and after ultrasonic treatment. Samples were sputter coated with gold and examined using a JEOL JSM-6360LV in the CEM (Centre of Electron Microscopy) at the UFPR (Universidade Federal do Parana). Atomic Force Microscopy (AFM) Analysis. The topography and surface roughness of the cellulose films were determined using a Shimadzu FPM-9500J3 operating in air. Small pieces were cut from each membrane, glued onto metal disks and attached to a magnetic sample holder located on top of the scanner tube. The membrane surface was scanned in tapping mode. All of the AFM images were taken at 25 °C. Differential Thermal Analysis. This analysis was carried out to determine the thermal stability of our materials by employing a Mettler Toledo TGA/SDTA851. Samples were heated from ambient temperature to 1000 °C in an oxygen current of 50 mL/min at a heating rate of 10 °C/min.

Results and Discussion SEM Analysis. After ultrasonic treatment for different lengths of time in an aqueous medium, the cellulose was vacuum-dried at 40 °C, and the morphology of new films was initially observed by scanning electron microscopy (SEM). Figure 1a shows the 3-D reticulated structure of the cellulose fibrils with interconnected pores of different sizes, a typical structure of cellulose produced by A. xylinum in a glucose medium by static fermentation.16 The bacteria Acetobacter xylinum synthesize primary nanofibrils with lateral sizes in the range of 7-13 nm, which aggregate into thin and flat bands or ribbons having widths of 70-150 nm.46 The SEM micrograph of the native bacterial cellulose showed bands of microfibrils (ribbons) with widths of 90-140 nm. After

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Figure 1. Scanning electron micrographs of bacterial cellulose, native (a) and ultrasonicated for 15 (b), 30 (c), 60 (d), and 75 min (e).

the shorter ultrasonication time, we can observe that there is a reduction in the width of the ribbons and a decrease in the number/density of pores (Figure. 1b). Apparently, a change in the morphology causes the surface to become more compact with fewer pores. Increasing the length of the treatment reduces the ribbons even further, and the surfaces become more uniform (Figure 1c-e). The width of the ribbons cannot be determined by SEM analysis because of their reduced size, so it was determined by AFM analysis and by SAXS instead. AFM Analysis. The surface morphology and surface characteristics, such as roughness, are important factors in interfacial interactions; thus, the surface changes in bacterial cellulose, after ultrasonic treatment, were determined. The tapping-mode height images of the ultrasonic cellulose surfaces are shown in Figure 2. The width of the ribbons in the native BC was determined to be between 110-140 nm, values near those observed by SEM. In the BC that was ultrasonicated for 15 min, there was, apparently, a reduction in the width (70-110 nm) and an increase in the thickness (39-42 nm) of the ribbons; there seems to be a rather open fibrillar network on the surface of this film.

Apparently, the ultrasonic treatment changed the microfibrillar arrangement, leading to a film with a different nanostructure. From the height obtained with AFM, we observed that after 15 min of ultrasonic processing, there is an increase in the ribbons’ thickness (Table 1). Apparently, increasing the time of ultrasonic application to 60 min or more caused a reduction of thickness of the ribbons. The image presented in Figure 2c shows the surface of the film that was treated for 60 min with ultrasound, showing changes in the morphology of the ribbons. The surface of the BC film that was ultrasonicated for 30 min has similar features (data not shown). After 75 min, individual ribbons with widths of 40-70 nm could be observed. The AFM analysis allowed us to observe that on the surface of the cellulose films that were ultrasonicated for 30 and 60 min the shape of this nanomaterial was irregular. Determining the thickness of the ribbons from AFM is prone to imprecision due to the arrangement of the ribbons on a flat surface. The determination of the variation in the thickness and

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Figure 2. AFM tapping-mode height images in air of a bacterial cellulose thin film: native (a) and ultrasonicated for 15 (b), 60 (c), and 75 min (d). Table 1. Width, Height, and Roughness (rms) of the Native BC and the Ultrasonicated BC at Different Times, Analyzed by Atomic Force Microscopy (AFM)a rms (nm)

width (nm)

height (nm)

BC

38.5

BC15

37.9

BC60

29.3

BC75

17.3

110.0 120.0 140.0 51.0 64.6 70.4 30.9 49.9 52.9 26.4 36.2 39.5

25.2 20.6 20.4 39.4 42.3 40.3 12.6 12.0 22.1 22.0 26.4 20.0

a

Area: 2 × 2 µm.

width of the ribbons after ultrasonic treatment is performed by SAXS analyses in the next section. The corresponding mean roughness values (rms) were different for all of the films reconstituted after the ultrasonication process, varying from 38.5 to 17.3 nm (Table 1). At least two films of the same composition were analyzed at different areas of the surface. It is difficult to compare the different roughness values reported in the literature because of a lack of uniformity in the methods applied to determine surface roughness; moreover, the size of the scan area is highly variable. After the ultrasonication process, we can see evidence of homogeneous profiles and low roughness values due to more homogeneous substrates, opening the possibility for a future study of biomolecular adsorption.

Figure 3. Differential thermal analysis of bacterial cellulose, native and ultrasonicated.

Differential Thermal Analysis. Ciolacu and Popa47 showed that the lower the crystallinity of the polymer, the lower is its thermal stability. Differential thermal analysis of native bacterial cellulose and ultrasonicated samples is shown in Figure 3. The thermal decomposition pattern of cellulose can be explained mainly by two different mechanisms.48 The first one involves the degradation due to the breaking of internal bonds, dehydration or formation of free radicals, and reactive carbonaceous char. The second mechanism involves the cleavage of secondary bonds and the formation of intermediate products,



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Figure 5. SAXS plots from the three samples along with the extrapolated areas (light dots). Table 2. Dimensions of the Ribbons Obtained from SAXS

BC BC15 BC30

Figure 4. Guinier plots for native cellulose (a), cellulose treated for 15 min (b), and cellulose treated for 30 min (c).

such as anhydromonosaccharides, which are converted into low molecular weight polysaccharides and finally carbonized products.49 The results obtained from differential thermal analysis showed us the onset temperature of the pyrolysis was 208, 250, and 268 °C for the native BC and the BC ultrasonicated for 15 and 30 min, respectively. In addition, after comparing the three samples, it was observed that the weight losses of the two degradation events were similar, indicating that the ultrasonic process did not change the cellulose chemically; apparently, only structural changes occurred. More data from differential thermal analysis can be seen in the Supporting Information. SAXS Analysis. Previous X-ray scattering studies on BC50,51 have already evidenced an ultrastructure composed of nanofibrils (in the nm range) aggregated into microfibrils (in the 10 nm range), attached together through interconnecting cellulose chains, to produce ribbons (in the 100 nm range) that are the basic units of the 3D network observed in bacterial cellulose. The BC ribbons are the basic units that contribute most for the X-ray scattering at small angles. However, due to their extensive length (L), the full radius of gyration of the ribbons is way beyond the detection limit of conventional SAXS. Nevertheless, it is still possible to observe an intermediate scattering region where a cross sectional radius of gyration,rc, can be identified and related52 to the thickness a and the width

a (nm)

b (nm)

7.6 21 15

75 86 81

b of the ribbons byrc ) (a2 + b2)/(12). In this region, which in our experiment belongs to the lowest wave vectors, the cross sectional radius of gyration is determined53 by the linear coefficient of the curve ln(qI(q)) × q2 as in Figure 4. It is clear from Figure 4 that rc increases with ultrasonic treatment, indicating that one or both cross sectional dimensions increased with ultrasonic treatment. However, this does not necessarily imply a thickening of the ribbon. In fact, because the cross sectional radius of gyration alone cannot determine a and b independently, we also need to evaluate the Porod cross sectional area of the ribbons,S ) ab, given by the SAXS curves as52,53 S ) 2π limqf0(qI(q))/Q. The scattering invariant Q can be determined by the total area of the scattering curve,53 extrapolated into the Guinier and Porod regions (see Supporting Information for details of this calculation). From Figure 5, we can see that the normalized scattering curve, I(q)/Q, is shifted upward with ultrasonic treatment, indicating that the cross sectional area of the ribbons has also increased. Finally, combining the data for S and rc, one obtains the cross sectional dimensions a and b of the ribbons, as presented in Table 2. The dimensions of the native microfibril bands obtained here are similar to the ones reported previously;52 however, as suggested by the SAXS data analysis, the treatment with ultrasound seems to increase their thickness, resulting in a modification of the nanostructure of the BC. Note, however, that this observation does not necessarily imply an increase in crystallinity because the ribbons are composed of crystalline microfibrils and amorphous cellulose. The crystallite dimensions and crystalline fractions of the samples are determined from the WAXS data analysis in the next section. WAXS Analysis. The great interest in the mechanical and medical properties of bacterial cellulose is due to its crystalline orientation. The physical properties, as well as the accessibility in the chemical reactions, swelling and adsorption, are strongly influenced by the crystallinity.54 Due to the linearity of the cellulose backbone, adjacent chains of cellulose form a framework of water-insoluble aggregates of varying length and width. These microfibrils consist of both highly ordered (crystalline) and less ordered (amorphous) regions. Depending on the source and the conditions used in cellulose treatment, different degrees of crystallinity can be obtained.55

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Tischer et al. Table 3. Crystallinity (xc) and Widths of the Crystallites in the Directions Normal to the Corresponding Bragg Planes

native BC 15 BC 30

xc

D010

D001

D011

D-411

0.44 0.55 0.63

3.4 6.4 5.4

10.1 9.6 8.5

7.5 6.5 6.0

10.9 19.4 17.6

where K is a constant that depends on the shape of the crystallites (K ≈ 0.94 for cubic and K ≈ 1.1 for a spherical shape), and ∆qhkl is the fwhm of the hkl reflection. Although subtle, the differences in the crystalline structure of plant and bacterial cellulose have been a subject of great debate.54 In bacterial cellulose, the major fraction is usually ascribed to IR crystallinity. For the present data, a Rietveld or linked atoms least squares (LALS) fit would have to be performed to determine precisely the relative amounts of IR and Iβ components.55,56 However, our main concern here is only the relative amount of crystallinity between the native and the ultrasonically treated samples. We, therefore, index the reflections according to IR crystallinity. In Table 3, we present the average crystallite lengths along selected directions, according to the Scherrer relation. As we can see, due to the ultrasonic treatment, the average crystallite dimension increases in the (010) direction but shrinks in the other directions. These changes in crystalline dimensions are compatible with the increase in the ribbon dimensions obtained from the SAXS analysis. The crystallinity of the native bacterial cellulose and the samples ultrasonicated for 15 and 30 min was calculated from the invariant integral of the crystalline and total scattering contributions as Figure 6. Lorenzian fits to the WAXS reflections in the Debye-Scherrer mode for the native BC (a), BC treated for 15 min (b), and BC treated for 30 min (c).

The dimensions of the crystalline regions can also be obtained from the line widths of the crystalline reflections using the Scherrer method. Although not accurate, as a result of neglecting the contributions due to paracrystallinity defects, this method can be useful for comparing the crystallite sizes of the three samples. Thus, we fit Lorenzians to the main crystalline reflections with different positions and widths, along with a smooth amorphous background, as presented in Figure 6. The positions of the crystalline peaks are fixed for the three samples, using the indexing of the main IR reflections according to Nishiyama et al.56 Note also that the Miller indices of the reflections are from a unit cell with the chains aligned along the a-axis. The widths and heights of the Lorenzian peaks are fitted simultaneously to the smooth amorphous background, which we represent by four broad Lorenzians spread along the entire range of measured wave-vectors and centered at approximately q ) 0, 7, 12, and 26 nm-1. The fwhm (full width at half maximum) of the Lorenzian peaks are determined by the widths of the crystallites in the direction normal to the Bragg planes given by the Scherrer relation as follows:

Dhkl )

2πK ∆qhkl

xc )

∫ Icr(q)q2dq ∫ I(q)q2dq

where Icr(q) is the contribution to the scattering curve due to only the crystalline peaks and I(q) is the full (amorphous + crystalline) WAXS curve. The crystallinity of these samples, native and ultrasonically treated for 15 and 30 min, is 0.44, 0.55, and 0.63, respectively. Although this method is not as reliable for obtaining the crystallinity57 as the Rietveld method, the relative increase in xc is clear and appears to be due to the conversion of the amorphous halo background directly into the (010) facets of the new crystallites. The transformation of amorphous cellulose into cellulose I is expected because polymerization and crystallization are coupled in Acetobacter xylinum. It has been shown by Benziman et al.58 that by preventing crystallization using Calcofluor, the amorphous cellulose is formed as bundles of parallel glucan chains that crystallize into cellulose IR after drying. This suggests that, in normal biogenic conditions, the amorphous parts from bacterial cellulose are cell-directed into parallel bundles ready to crystallize into cellulose I. The activation energy required for such a process is provided by cavitation from the ultrasonic treatment. This process occurs primarily in the amorphous regions, where water can more effectively penetrate. It is clear that part of the amorphous background corresponding to the broad features around q ) 7 and 12 nm-1 (the blue curve in Figure 6), which may come from the intermicrofibrillar cellulose chains, is being converted into crystallites. However, if the conversion of amorphous material into crystalline material



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thank Mateus B. Cardoso for insightful discussions and help with Figure 7. Supporting Information Available. Details of the SAXS data analysis and additional data obtained by differential thermal analysis. This material is available free of charge via the Internet at http://pubs.acs.org.

References and Notes

Figure 7. Scheme illustrating the changes after ultrasonic treatment and the creation of new crystallites.

occurred in each ribbon independently, we would expect a decrease in their thickness rather than an increase because the density of crystalline cellulose is higher than that of amorphous cellulose. Therefore, we propose that the thicker crystallites result from the fusion of neighboring ribbons, as shown in Figure 7. Indeed, it is expected that the surface of the ribbons will be much more susceptible to the effects of cavitation than the crystalline core. This scenario supports the conclusions obtained from SEM, AFM, SAXS, and WAXS.

Conclusions In the present work, we have demonstrated that ultrasonic processing in mild conditions was effective in changing the microfibrillar structure of bacterial cellulose. The ultrasound energy is transferred through shearing and cavitation to the glucan chains, promoting the conversion of amorphous material into crystalline material. The energy scale of the cavitation processes is within the hydrogen bond energy scale and occurs primarily in the amorphous regions because these are the regions where water can more effectively penetrate. Moreover, due to the intrinsic nature of polymerization and crystallization promoted by Acetobacter xylinum, the crystalline material formed is of type I and not of type II, as would be expected by random crystallization of the amorphous phase. A scheme illustrating this effect is depicted in Figure 7. The shape of these crystallites changed, and this process created films with higher crystallinity and lower surface roughness. Acknowledgment. We thank CNPq for a postdoctoral fellowship (P.C.S.F.T.), CAPES/Brazil for financial support, FINEP for financial support for the use of electronic microscopy equipment (CEM), Prof. Paulo Cesar Camargo for the use of AFM microscopy, and LAMIR (Laborato´rio de Ana´lises de Minerais e Rochas) for Differential Thermal Analysis. We also

(1) Saxena, I. M.; Kudlicka, K.; Okuda, K.; Brown, R. M., Jr. J. Bacteriol. 1994, 176, 5735–5752. (2) Hornung, M.; Ludwig, M.; Schmauder, H. P.; Gerrard, A. M. Eng. Life Sci. 2006, 6, 546–551. (3) Gardner, D. J.; Oporto, G. S.; Mills, R.; Samir.; Azizi, M. A. S. J. Adhes. Sci. Technol. 2008, 22, 545–567. (4) Brown, A. J. J. Chem. Soc. 1886, 49, 172–187. (5) Brown, A. J. J. Chem. Soc. 1886, 49, 432–439. (6) Iguchi, M.; Yamanaka, S.; Budhiono, A. J. Mater. Sci. 2000, 35, 261– 270. (7) Brown, R. M., Jr.; Willison, J. H. M.; Richardson, C. L. Proc. Natl. Acad. Sci. U.S.A. 1976, 73, 4565–4569. (8) Yamanaka, S.; Watanabe, K.; Kitamura, N. J. Mater. Sci. 1989, 24, 3141–3145. (9) Cousins, S. K.; Brown, R. M., Jr. Polymer 1997, 38, 897–902. (10) Son, H. J.; Heo, M. S.; Kim, Y. G.; Lee, S. J. Biotechnol. Appl. Biochem. 2001, 33, 1–5. (11) Othmer, K. Encyclopedia of Chemical Technology: Cellulose; John Wiley & Sons, New York, 1993. (12) Okiyami, A. M.; Motoki, M.; Yamanaka, S. Food Hydrocolloids 1993, 6, 503–511. (13) Goelzer, F. D. E.; Faria-Tischer, P. C. S.; Vitorino, J. C.; Sierakowski, M.-R.; Tischer, C. A. Mater. Sci. Eng., C 2009, 29, 546–551. (14) Sarko, A.; Muggli, R. Macromolecules 1974, 7, 486–494. (15) Watanabe, K.; Tabuchi, M.; Ishikawa, A.; Takemura, H.; Tsuchida, T.; Morinaga, Y.; Yoshinaga, F. Biosci., Biotechnol., Biochem. 1998, 62, 1290–1292. (16) Gardner, K. H.; Blackwell, J. Biopolymer 1974, 13, 1975–2001. (17) Lima, M. M. D.; Borsali, R. Macromol. Rapid Commun. 2004, 25, 771–787. (18) Araki, J.; Wada, M.; Kuga, S.; Okano, T. Colloids Surf., A 1998, 142, 75–82. (19) Battista, O. A. Ind. Eng. Chem. Res. 1950, 42, 502–507. (20) Marchessault, R. H.; Morehead, F. F.; Walter, N. M. Nature 1959, 184, 632–633. (21) Marchessault, R. H.; Morehead, F. F.; Joan, K. M. J. Colloid Sci. 1961, 16, 327–344. (22) Dong, X. M.; Kimura, T.; Revol, J. F.; Gray, D. G. Langmuir 1996, 12, 2076–2082. (23) Turbak, A. F.; Snyder, F. W.; Sandberg, K. R. J. Appl. Polym. Sci. 1983, 37, 815–827. (24) Herrick, F. W.; Casebier, R. L.; Hamilton, J. K.; Sandberg, K. R. J. Appl. Polym. Sci. 1983, 37, 797–813. (25) Paakko, M.; Ankerfors, M.; Kosonen, H.; Nykanen, A.; Ahola, S.; Osterberg, M.; Ruokolainen, J.; Laine, J.; Larsson, P. T.; Ikkala, O.; Lindostrom, T. Biomacromolecules 2007, 8, 1934–1941. (26) Zhang, J.; Elder, T. J.; Pu, Y.; Ragauskas, A. J. Carbohydr. Polym. 2007, 69, 607–611. (27) Liu, H.; Bao, J.; Du, Y.; Zhou, X.; Kennedy, J. F. Carbohydr. Polym. 2006, 64, 553–559. (28) Liu, H.; Du, Y.-M.; Kennedy, J. F. Carbohydr. Polym. 2007, 68, 598– 600. (29) Wu, T.; Zivanovic, S.; Hayes, D. G.; Weiss, J. J. Agric. Food Chem. 2008, 56, 5112–5119. (30) Kasaai, M. R.; Arul, J.; Charlet, G. Ultrason. Sonochem. 2008, 15, 1001–1008. (31) Cote, G. L.; Willet, J. L. Carbohydr. Polym. 1999, 39, 119–126. (32) Lorimer, J. P.; Mason, T. J.; Cuthbert, T. C.; Brookfield, E. A. Ultrason. Sonochem. 1995, 2, 55–57. (33) Portenlanger, G.; Heusinger, H. Ultrason. Sonochem. 1997, 4, 127– 130. (34) Lii, C.-Y.; Chen, C.-H.; Yeh, A.-I.; Lai, V. M. F. Food Hydrocolloids 1999, 13, 477–481. (35) Vodenicarova, M.; Driı´malova´, G.; Hroma´dkova´, Z.; Malovı´kova´, A.; Ebringerova´, A. Ultrason. Sonochem. 2006, 13, 157–164. (36) Imai, M.; Ikari, K.; Suzuki, I. Biochem. Eng. J. 2004, 17, 79–83. (37) Gronroos, A.; Pirkonen, P.; Ruppert, O. Ultrason. Sonochem. 2004, 11, 9–12.

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(38) Aliyu, M.; Hepher, M. J. Ultrason. Sonochem. 2000, 7, 265–268. (39) Mislovicˇova´, D.; Masarova, J.; Bebdzalova, K.; Soltes, L.; Machova, E. Ultrason. Sonochem. 2000, 7, 63–68. (40) Chen, R. H.; Chang, J. R.; Shyur, J. S. Carbohydr. Res. 1997, 299, 287–294. (41) Suslick, K. S. Science 1990, 247, 1439–1445. (42) Li, S. Y.; Liu, X. F.; Zhuang, X. P.; Guan, Y. L.; Cheng, G. X. Chin. J. Appl. Chem. 2003, 20, 1030–1035. (43) Hestrin, S.; Schramm, M. Biochem. J. 1954, 58, 345–352. (44) Blanton, T. N.; Barnes, C. L.; Lelental, M. J. Appl. Crystallogr. 2000, 33, 172–173. (45) Narayanan, T.; Diat, O.; Bosecke, P. Nucl. Instrum. Methods Phys. Res.,Sect. A 2001, 467, 1005–1009. (46) Klemm, D.; Schumann, D.; Kramer, F.; Hessler, N.; Hornung, M.; Schmauder, H.-P.; Marsch, S. AdV. Polym. Sci. 2006, 205, 49–96. (47) Ciolacu, D; Popa, V. On the thermal degradation of cellulose allomorphs. Cellul. Chem. Technol. 2006, 40, 445–449. (48) Rowell, R. M., Le Van-Green, S. L. In Handbook of Wood Chemistry and Wood Composites; Rowell, R. M., Ed.; CRC Press: Gainesville, FL, 2005; p 121.

Tischer et al. (49) Kawamoto, H.; Murayama, M.; Saka, S. J. Wood Sci. 2003, 49, 469– 473. (50) Klemm, D.; Heublein, B.; Fink, H.-P.; Bohn, A. Angew. Chem., Int. Ed. 2005, 44, 3358–3393. (51) Nishiyama, Y. J. Wood Sci. 2009, 55, 241–249. (52) Astley, O. M.; Chanliaud, E.; Donald, A. M.; Gidley, M. J. Int. J. Biol. Macromol. 2001, 29, 193–202. (53) Glatter, O. In Small Angle X-ray Scattering; Glatter, O., Kratky, Eds.; Academic Press: London, 1982; p 167. (54) Klemm, D.; Schumann, D.; Udhardt, U.; Marsch, S. Prog. Polym. Sci. 2001, 26, 1561–1603. (55) Iwata, T.; Indrarti, L.; Azuma, J. Cellulose 1998, 5, 215–228. (56) Nishiyama, Y.; Sugiyama, J.; Chanzy, H.; Langan, P. J. Am. Chem. Soc. 2003, 125, 14300–14306. (57) Thygesen, A.; Oddershede, J.; Lilholt, H.; Belinda, A. T.; Stahl, K. Cellulose 2005, 12, 563–576. (58) Benziman, M.; Haigler, C. H.; Brown, M. R.; White, R. A.; Cooper, M. K. Proc. Natl. Acad. Sci. U.S.A. 2007, 77, 6678–6682.

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