Nanostructural Reorganization of Bacterial Cellulose byUltrasonic Treatment

Paula C. S. Faria Tischer,*,† Maria Rita Sierakowski,† Harry Westfahl, Jr.,*,‡ andCesar Augusto Tischer†

Laboratorio de Biopolımeros, Universidade Federal do Parana (UFPR), Caixa Postal 19081,81531-990, Curitiba, Parana, Brazil, and Laboratorio Nacional de Luz Sıncrotron (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 treatmentchanged the width and height of the microfibrillar ribbons and roughness of their surface, originating films withnew nanostructures. Differential thermal analysis showed a higher thermal stability for ultrasonicated sampleswith a pyrolysis onset temperature of 208 °C for native bacterial cellulose and 250 and 268 °C for the modifiedsamples. The small-angle X-ray scattering experiments demonstrated that the treatment with ultrasound increasedthe thickness of the ribbons, while wide-angle X-ray scattering experiments demonstrated that the average crystallitedimension and the degree of crystallinity also increased. A model is proposed where the thicker ribbons andcrystallites result from the fusion of neighboring ribbons due to cavitation effects.

Introduction

Cellulose is one of the most important biopolymers and iswidely used based on its availability, biocompatibility, biologicaldegradability, and sustainable production.1 In particular, bacterialcellulose is a pure biopolymer, without collateral biogeniccompounds like lignin, hemicelluloses, and pectin, and it is anattractive biopolymer due to its facile production and purifica-tion.2 The structure of bacterial cellulose (BC) was describedmore than a century ago,3,4 and the material has been extensivelyanalyzed and characterized.5-9

Bacterial cellulose has been produced from Acetobacterxylinum by fermentation of different substrates as glucose fromcorn syrup,10-12 and more recently, rice bark supplemented withglucose was used as a substrate to produce cellulose nano-spheres.13

Cellulose molecules form long chains in polycrystallinefibrous bundles, which contain crystalline and amorphousregions, and the degree of crystallinity and crystal dimensionsare dependent on the origin of the cellulose. The biosyntheticprocess is the same in all organisms, but there are somedifferences in the cellulose synthase complexes that determinethe size and thickness of the cellulose microfibrils, and the greatinterest in cellulose macromolecules is due to their crystallineorientation.14

The microstructures formed by the ultrafine microfibrils ofbacterial cellulose have lengths varying from 1 to 9 µm andcreate a dense reticulated structure stabilized by varioushydrogen bonds. These networks show a high index of crystal-linity (above 60%) and a higher degree of polymerization,normally around 16.000 or 20.000,15 in comparison with vegetalcellulose.

The high crystallinity of bacterial cellulose is the result of ahigh number of inter- and intrahydrogen bonds between adjacentchains of glucan. These bonds create a regular crystallinearrangement between the glucan molecules,16 resulting in thedistinct diffraction pattern, swelling, and reactivity of cellulose.

One of the methods used to obtain microcrystals of celluloseis to submit cellulose to controlled acid (H2SO4) hydrolysisconditions. The hydronium ions can penetrate the cellulosechains in the amorphous domains, promoting the hydrolyticcleavage of the glycosidic bonds and releasing individualcrystallites. In this process, sulfate groups are randomlydistributed on the cellulose microfibril surface, inducing anegative electrostatic layer covering the microfibers.20-22 Thecellulose microcrystals, because of their stiffness, thickness, andlength, are commonly called “whiskers”.17-19

For pursuing applications that require extensively entanglednetworks and higher strength, chemically less aggressive hy-drolysis concepts have been found to maintain a high aspectratio of the cellulose I fibrils or fibril aggregates, potentiallyallowing strong or even permanent junction points for thenetworks. A classic example is provided by omitting thehydrolysis step and by solely imposing high shearing forcesfor disintegration. This process yields a highly entanglednetwork, which typically consists of elements with a wide sizedistribution down to the nanoscale. The resulting material hasbeen denoted microfibrillated cellulose (MFC), originally in-troduced by Turback et al.23 and Herrick et al.24

By this method, the cellulose forms a highly entanglednetwork with a low degree of crystallinity, and some of thenoncrystalline domains remain intact. This method has the sameproblems, and the increased energy of tension necessary to causethe disintegration of fibers requires several steps of treatment.Recently, a novel route was demonstrated that combinesenzymatic hydrolysis and mechanical shearing to form a longand highly entangled network of cellulose I elements.25

Another method cited to prepare spherical nanoparticles ofcellulose is based on a hard pretreatment of cellulose: heating

* To whom correspondence should be addressed. Phone: +55 41 3361-3260 (P.C.S.F.); +55 19 3512-1034 (H.W.). E-mail: paula.tischer@pq.cnpq.br(P.C.S.F.); westfahl@lnls.br (H.W.).

† Universidade Federal do Parana (UFPR).‡ Laboratorio Nacional de Luz Sıncrotron (LNLS).

Biomacromolecules 2010, 11, 1217–1224 1217

10.1021/bm901383a 2010 American Chemical SocietyPublished on Web 04/06/2010

at 80 °C in 5.0 M NaOH for 3 h, followed by the resuspensionof fibers in DMSO in a water bath at 80 °C for 3 h. Next, thepretreated fibrils are suspended in an acid solution and sonicatedwith heating at 80 °C for 8 h. By this process, nanospheres of60-570 nm composed of cellulose II are obtained.26

All these methods were realized with vegetal cellulose, andhard conditions for hydrolysis were used, which caused intensedegradation of the polymer and a consequent reduction incrystallinity.

The effect of ultrasound in degrading polysaccharide linkagesis 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 inthe majority of cases, the treatment with ultrasound has beenused in acid solutions with relatively low temperature and inalkaline39 or acidic conditions.40

The energy of ultrasound is transferred to the polymer chainsthrough a process called cavitation, which is the formation,growth, and violent collapse of cavities in the water. The energyprovided by cavitation in this so-called sonochemistry isapproximately 10-100 kJ/mol,41 which is within the hydrogenbond energy scale.42

The main goal of this work was to analyze the effect ofultrasonic treatment on the reorganization of bacterial cellulosemicrofibrils using suitable conditions (aqueous medium, withoutacid or heating). Such results are needed to develop an easymethod to obtain films with different nanostructures andcharacteristics (porosity, roughness, and crystallinity) and todevelop a process in nanotechnology that permits the formationof new scaffolds for tissue engineering.

Materials and Methods

Microorganism, Culture Media, and Growth Conditions. Thebacterial strain Acetobacter xylinum ATCC 23769 (reclassified as thegenus Gluconacetobacter), supplied by Foundation Andre Tosello fromCampinas, Sao Paulo, was grown in a glucose medium based on theHestrin-Schramm’s medium culture.43 All media were autoclaved at121 °C and 1.02 atm for 20 min.

The inoculum was prepared in a nonagitated 200-mL Erlenmeyerflask containing about 5 mL of the thawed culture and 40 mL of theabove medium. The flask was incubated at 28 ( 1 °C for 48 h. Theobtained cell suspension was used as the inoculum. The staticfermentation was carried out for 10 days at 28 °C, and the pH wasadjusted 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 fermentationmedium was removed, rinsed thoroughly with deionized water, andpurified by immersion in an aqueous solution of 0.1 M NaOH for oneday. The films were washed repeatedly with deionized water and thenvacuum-dried at 40 °C. This treatment was controlled to avoid themercerization of the BC.

Ultrasonic Process. The pellicles of the BC after vacuum drying at40 °C were cut with scissors. The cut films (200 mg, immersed in 200mL Mili-Q water) were shaken vigorously for 15 min. This materialwas submitted to ultrasonic treatment for 15, 30, 60, and 75 min usingan ultrasonic processor, Ultrasound SONICS (200 W, 20 kHz). Theultrasonic treatment was carried out in an ice bath, and the ice wasmaintained throughout the entire sonication time. After this process,the ultrasonicated BC was reconstituted in the form of films andvacuum-dried at 40 °C. In all of the analysis techniques, the stablefilms formed after ultrasonication were used in the solid state as wholefilms or cut films.

Small-Angle X-ray Diffraction (SAXS). The SAXS patterns wereused to determine the dimensions of the microfibrils in the cellulose.SAXS diagrams were recorded on the D02A beamline using a fixed

radiation wavelength (λ ) 1.488 Å) at the Brazilian SynchrotronLight Laboratory (LNLS) also in the transmission geometry. Thesamples were thin films, supported in the sample holder and keptin a 10-2 mbar vacuum. Scattering patterns were recorded during300 s exposures on a marCCD 165 detector (8 × 8 binning), placedfirst at 2507 mm and then at 1484 mm away from the samples.Calibration of the distances was achieved using a silver Behenatepattern.44 The SAXS profiles from the samples consisted of isotropicscattering patterns that were subtracted from the CCD bias and dark-noise according to a reference.45 The SAXS chamber parasiticscattering was also recorded (with bias and dark-noise subtraction)and subtracted from the sample pattern after sample attenuationcorrection. The final patterns were normalized by the phase spacevolume and azimuthally averaged for a sequence of wave-vectors q(q ) (4π/λ) sin(θ); 2θ ) scattering angle) ranging from 0.04 to1.14 nm-1 when the detector was at the longer distance and 0.1 to2.3 nm-1 for the shorter one.

Wide-Angle X-ray Diffraction (WAXS). The WAXS patterns wereused to determine the relative crystallinity. WAXS diagrams wererecorded on the D03B beamline using a fixed radiation wavelength (λ) 1.433 Å) at the LNLS in the transmission geometry. The sampleswere thin films supported in 1 cm aluminum rings aligned perpendicu-larly to the beam. Diffraction patterns were recorded during 120 sexposures on a MarCCD 165 detector (2 × 2 binning), placed 100.0mm away from the samples. Calibration of the distances was achievedusing a LaB6 pattern. The WAXS profiles from the samples consistedof circular Debye-Scherrer rings and were subtracted from the CCDbias and dark-noise according to a reference.45 The air scattering wasalso recorded (with bias and dark-noise subtraction) and subtracted fromthe sample pattern after sample attenuation correction. The final patternswere normalized by the phase space volume and azimuthally averagedfor a sequence of wave-vectors q (q ) (4π/λ) sin(θ); 2θ ) scatteringangle) ranging from 1.4 to 28.6 nm-1.

Scanning Electron Microscopy (SEM) Analysis. Scanning electronmicroscopy (SEM) was conducted to observe the surface of the bacterialcellulose films before and after ultrasonic treatment. Samples weresputter coated with gold and examined using a JEOL JSM-6360LV inthe CEM (Centre of Electron Microscopy) at the UFPR (UniversidadeFederal do Parana).

Atomic Force Microscopy (AFM) Analysis. The topography andsurface roughness of the cellulose films were determined using aShimadzu FPM-9500J3 operating in air. Small pieces were cut fromeach membrane, glued onto metal disks and attached to a magneticsample holder located on top of the scanner tube. The membrane surfacewas scanned in tapping mode. All of the AFM images were taken at25 °C.

Differential Thermal Analysis. This analysis was carried out todetermine the thermal stability of our materials by employing a MettlerToledo TGA/SDTA851. Samples were heated from ambient temperatureto 1000 °C in an oxygen current of 50 mL/min at a heating rate of10 °C/min.

Results and Discussion

SEM Analysis. After ultrasonic treatment for different lengthsof time in an aqueous medium, the cellulose was vacuum-driedat 40 °C, and the morphology of new films was initially observedby scanning electron microscopy (SEM). Figure 1a shows the3-D reticulated structure of the cellulose fibrils with intercon-nected pores of different sizes, a typical structure of celluloseproduced by A. xylinum in a glucose medium by staticfermentation.16 The bacteria Acetobacter xylinum synthesizeprimary nanofibrils with lateral sizes in the range of 7-13 nm,which aggregate into thin and flat bands or ribbons havingwidths of 70-150 nm.46

The SEM micrograph of the native bacterial cellulose showedbands of microfibrils (ribbons) with widths of 90-140 nm. After

1218 Biomacromolecules, Vol. 11, No. 5, 2010 Tischer et al.

the shorter ultrasonication time, we can observe that there is areduction in the width of the ribbons and a decrease in thenumber/density of pores (Figure. 1b). Apparently, a change inthe morphology causes the surface to become more compactwith fewer pores. Increasing the length of the treatment reducesthe ribbons even further, and the surfaces become more uniform(Figure 1c-e). The width of the ribbons cannot be determinedby SEM analysis because of their reduced size, so it wasdetermined by AFM analysis and by SAXS instead.

AFM Analysis. The surface morphology and surface char-acteristics, such as roughness, are important factors in interfacialinteractions; thus, the surface changes in bacterial cellulose, afterultrasonic treatment, were determined. The tapping-mode heightimages of the ultrasonic cellulose surfaces are shown in Figure2. The width of the ribbons in the native BC was determined tobe 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 anincrease in the thickness (39-42 nm) of the ribbons; there seemsto be a rather open fibrillar network on the surface of this film.

Apparently, the ultrasonic treatment changed the microfibrillararrangement, leading to a film with a different nanostructure.From the height obtained with AFM, we observed that after 15min of ultrasonic processing, there is an increase in the ribbons’thickness (Table 1). Apparently, increasing the time of ultrasonicapplication to 60 min or more caused a reduction of thicknessof the ribbons.

The image presented in Figure 2c shows the surface of thefilm that was treated for 60 min with ultrasound, showingchanges in the morphology of the ribbons. The surface of theBC film that was ultrasonicated for 30 min has similar features(data not shown). After 75 min, individual ribbons with widthsof 40-70 nm could be observed. The AFM analysis allowedus to observe that on the surface of the cellulose films that wereultrasonicated for 30 and 60 min the shape of this nanomaterialwas irregular.

Determining the thickness of the ribbons from AFM is proneto imprecision due to the arrangement of the ribbons on a flatsurface. The determination of the variation in the thickness and

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

Nanostructural Reorganization of Bacterial Cellulose Biomacromolecules, Vol. 11, No. 5, 2010 1219

width of the ribbons after ultrasonic treatment is performed bySAXS analyses in the next section.

The corresponding mean roughness values (rms) were dif-ferent for all of the films reconstituted after the ultrasonicationprocess, varying from 38.5 to 17.3 nm (Table 1).

At least two films of the same composition were analyzed atdifferent areas of the surface. It is difficult to compare thedifferent roughness values reported in the literature because ofa lack of uniformity in the methods applied to determine surfaceroughness; moreover, the size of the scan area is highly variable.After the ultrasonication process, we can see evidence ofhomogeneous profiles and low roughness values due to morehomogeneous substrates, opening the possibility for a futurestudy of biomolecular adsorption.

Differential Thermal Analysis. Ciolacu and Popa47 showedthat the lower the crystallinity of the polymer, the lower is itsthermal stability. Differential thermal analysis of native bacterialcellulose and ultrasonicated samples is shown in Figure 3.

The thermal decomposition pattern of cellulose can beexplained mainly by two different mechanisms.48 The first oneinvolves the degradation due to the breaking of internal bonds,dehydration or formation of free radicals, and reactive carbona-ceous char. The second mechanism involves the cleavage ofsecondary bonds and the formation of intermediate products,

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 BCand the Ultrasonicated BC at Different Times, Analyzed by AtomicForce Microscopy (AFM)a

rms (nm) width (nm) height (nm)

BC 38.5 110.0 25.2120.0 20.6140.0 20.4

BC15 37.9 51.0 39.464.6 42.370.4 40.3

BC60 29.3 30.9 12.649.9 12.052.9 22.1

BC75 17.3 26.4 22.036.2 26.439.5 20.0

a Area: 2 × 2 µm.

Figure 3. Differential thermal analysis of bacterial cellulose, nativeand ultrasonicated.

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such as anhydromonosaccharides, which are converted into lowmolecular weight polysaccharides and finally carbonized prod-ucts.49

The results obtained from differential thermal analysis showedus the onset temperature of the pyrolysis was 208, 250, and268 °C for the native BC and the BC ultrasonicated for 15 and30 min, respectively. In addition, after comparing the threesamples, it was observed that the weight losses of the twodegradation events were similar, indicating that the ultrasonicprocess did not change the cellulose chemically; apparently, onlystructural changes occurred. More data from differential thermalanalysis 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 nmrange), attached together through interconnecting cellulosechains, to produce ribbons (in the 100 nm range) that are thebasic units of the 3D network observed in bacterial cellulose.

The BC ribbons are the basic units that contribute most forthe X-ray scattering at small angles. However, due to theirextensive length (L), the full radius of gyration of the ribbonsis way beyond the detection limit of conventional SAXS.Nevertheless, it is still possible to observe an intermediatescattering region where a cross sectional radius of gyration,rc,can be identified and related52 to the thickness a and the width

b of the ribbons byrc ) (a2 + b2)/(12). In this region, which inour experiment belongs to the lowest wave vectors, the crosssectional radius of gyration is determined53 by the linearcoefficient of the curve ln(qI(q)) × q2 as in Figure 4.

It is clear from Figure 4 that rc increases with ultrasonictreatment, indicating that one or both cross sectional dimensionsincreased with ultrasonic treatment. However, this does notnecessarily imply a thickening of the ribbon. In fact, becausethe cross sectional radius of gyration alone cannot determine aand b independently, we also need to evaluate the Porod crosssectional area of the ribbons,S ) ab, given by the SAXS curvesas52,53 S ) 2π limqf0(qI(q))/Q. The scattering invariant Q canbe determined by the total area of the scattering curve,53

extrapolated into the Guinier and Porod regions (see SupportingInformation for details of this calculation). From Figure 5, wecan see that the normalized scattering curve, I(q)/Q, is shiftedupward with ultrasonic treatment, indicating that the crosssectional area of the ribbons has also increased.

Finally, combining the data for S and rc, one obtains the crosssectional dimensions a and b of the ribbons, as presented inTable 2. The dimensions of the native microfibril bands obtainedhere are similar to the ones reported previously;52 however, assuggested by the SAXS data analysis, the treatment withultrasound seems to increase their thickness, resulting in amodification of the nanostructure of the BC. Note, however,that this observation does not necessarily imply an increase incrystallinity because the ribbons are composed of crystallinemicrofibrils and amorphous cellulose. The crystallite dimensionsand crystalline fractions of the samples are determined fromthe WAXS data analysis in the next section.

WAXS Analysis. The great interest in the mechanical andmedical properties of bacterial cellulose is due to its crystallineorientation. The physical properties, as well as the accessibilityin the chemical reactions, swelling and adsorption, are stronglyinfluenced by the crystallinity.54

Due to the linearity of the cellulose backbone, adjacent chainsof cellulose form a framework of water-insoluble aggregatesof varying length and width. These microfibrils consist of bothhighly ordered (crystalline) and less ordered (amorphous)regions. Depending on the source and the conditions used incellulose treatment, different degrees of crystallinity can beobtained.55

Figure 4. Guinier plots for native cellulose (a), cellulose treated for15 min (b), and cellulose treated for 30 min (c).

Figure 5. SAXS plots from the three samples along with theextrapolated areas (light dots).

Table 2. Dimensions of the Ribbons Obtained from SAXS

a (nm) b (nm)

BC 7.6 75BC15 21 86BC30 15 81

Nanostructural Reorganization of Bacterial Cellulose Biomacromolecules, Vol. 11, No. 5, 2010 1221

The dimensions of the crystalline regions can also beobtained from the line widths of the crystalline reflectionsusing the Scherrer method. Although not accurate, as a resultof neglecting the contributions due to paracrystallinitydefects, this method can be useful for comparing thecrystallite sizes of the three samples. Thus, we fit Lorenziansto the main crystalline reflections with different positionsand widths, along with a smooth amorphous background, aspresented in Figure 6.

The positions of the crystalline peaks are fixed for the threesamples, using the indexing of the main IR reflections accordingto Nishiyama et al.56 Note also that the Miller indices of thereflections are from a unit cell with the chains aligned alongthe a-axis. The widths and heights of the Lorenzian peaks arefitted simultaneously to the smooth amorphous background,which we represent by four broad Lorenzians spread along theentire range of measured wave-vectors and centered at ap-proximately q ) 0, 7, 12, and 26 nm-1.

The fwhm (full width at half maximum) of the Lorenzianpeaks are determined by the widths of the crystallites in thedirection normal to the Bragg planes given by the Scherrerrelation as follows:

where K is a constant that depends on the shape of thecrystallites (K ≈ 0.94 for cubic and K ≈ 1.1 for a sphericalshape), and ∆qhkl is the fwhm of the hkl reflection.

Although subtle, the differences in the crystalline structureof plant and bacterial cellulose have been a subject of greatdebate.54 In bacterial cellulose, the major fraction is usuallyascribed to IR crystallinity. For the present data, a Rietveld orlinked atoms least squares (LALS) fit would have to beperformed to determine precisely the relative amounts of IR andI� components.55,56 However, our main concern here is onlythe relative amount of crystallinity between the native and theultrasonically treated samples. We, therefore, index the reflec-tions according to IR crystallinity. In Table 3, we present theaverage crystallite lengths along selected directions, accordingto the Scherrer relation.

As we can see, due to the ultrasonic treatment, the averagecrystallite dimension increases in the (010) direction but shrinksin the other directions. These changes in crystalline dimensionsare compatible with the increase in the ribbon dimensionsobtained from the SAXS analysis.

The crystallinity of the native bacterial cellulose and thesamples ultrasonicated for 15 and 30 min was calculated fromthe invariant integral of the crystalline and total scatteringcontributions as

where Icr(q) is the contribution to the scattering curve due toonly the crystalline peaks and I(q) is the full (amorphous +crystalline) WAXS curve.

The crystallinity of these samples, native and ultrasonicallytreated for 15 and 30 min, is 0.44, 0.55, and 0.63, respectively.Although this method is not as reliable for obtaining thecrystallinity57 as the Rietveld method, the relative increase inxc is clear and appears to be due to the conversion of theamorphous halo background directly into the (010) facets ofthe new crystallites.

The transformation of amorphous cellulose into cellulose Iis expected because polymerization and crystallization arecoupled in Acetobacter xylinum. It has been shown by Benzimanet al.58 that by preventing crystallization using Calcofluor, theamorphous cellulose is formed as bundles of parallel glucanchains that crystallize into cellulose IR after drying. This suggeststhat, in normal biogenic conditions, the amorphous parts frombacterial cellulose are cell-directed into parallel bundles readyto crystallize into cellulose I. The activation energy requiredfor such a process is provided by cavitation from the ultrasonictreatment. This process occurs primarily in the amorphousregions, where water can more effectively penetrate.

It is clear that part of the amorphous background correspond-ing to the broad features around q ) 7 and 12 nm-1 (the bluecurve in Figure 6), which may come from the intermicrofibrillarcellulose chains, is being converted into crystallites. However,if the conversion of amorphous material into crystalline material

Figure 6. Lorenzian fits to the WAXS reflections in the Debye-Scherrermode for the native BC (a), BC treated for 15 min (b), and BC treatedfor 30 min (c).

Dhkl )2πK∆qhkl

Table 3. Crystallinity (xc) and Widths of the Crystallites in theDirections Normal to the Corresponding Bragg Planes

xc D010 D001 D011 D-411

native 0.44 3.4 10.1 7.5 10.9BC 15 0.55 6.4 9.6 6.5 19.4BC 30 0.63 5.4 8.5 6.0 17.6

xc )∫ Icr(q)q2dq

∫ I(q)q2dq

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occurred in each ribbon independently, we would expect adecrease in their thickness rather than an increase because thedensity of crystalline cellulose is higher than that of amorphouscellulose.

Therefore, we propose that the thicker crystallites result fromthe fusion of neighboring ribbons, as shown in Figure 7.

Indeed, it is expected that the surface of the ribbons will bemuch more susceptible to the effects of cavitation than thecrystalline core. This scenario supports the conclusions obtainedfrom SEM, AFM, SAXS, and WAXS.

Conclusions

In the present work, we have demonstrated that ultrasonicprocessing in mild conditions was effective in changing themicrofibrillar structure of bacterial cellulose. The ultrasoundenergy is transferred through shearing and cavitation to theglucan chains, promoting the conversion of amorphous materialinto crystalline material. The energy scale of the cavitationprocesses is within the hydrogen bond energy scale and occursprimarily in the amorphous regions because these are the regionswhere water can more effectively penetrate. Moreover, due tothe intrinsic nature of polymerization and crystallization pro-moted by Acetobacter xylinum, the crystalline material formedis of type I and not of type II, as would be expected by randomcrystallization of the amorphous phase. A scheme illustratingthis effect is depicted in Figure 7. The shape of these crystalliteschanged, and this process created films with higher crystallinityand lower surface roughness.

Acknowledgment. We thank CNPq for a postdoctoralfellowship (P.C.S.F.T.), CAPES/Brazil for financial support,FINEP for financial support for the use of electronic microscopyequipment (CEM), Prof. Paulo Cesar Camargo for the use ofAFM microscopy, and LAMIR (Laboratorio de Analises deMinerais e Rochas) for Differential Thermal Analysis. We also

thank Mateus B. Cardoso for insightful discussions and helpwith Figure 7.

Supporting Information Available. Details of the SAXSdata analysis and additional data obtained by differential thermalanalysis. This material is available free of charge via the Internetat http://pubs.acs.org.

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