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Surface Morphology and Molecular Organization of Lignins in Langmuir-Blodgett Films Daniel Pasquini,‡ D. T. Balogh,*,† P. A. Antunes,| C. J. L. Constantino,§ A. A. S. Curvelo,‡ R. F. Aroca,| and O. N. Oliveira, Jr.† Instituto de Fı´sica de Sa˜ o Carlos, Universidade de Sa˜ o Paulo, CP 369, 13560-970 Sa˜ o Carlos, SP, Brazil, Instituto de Quı´mica de Sa˜ o Carlos, Universidade de Sa˜ o Paulo, CP 780, 13560-970, Sa˜ o Carlos, SP, Brazil, Faculdade de Cieˆ ncias e Tecnologia, Universidade Estadual Paulista, CP 467, 19060-900 Presidente Prudente, SP, Brazil, and Materials and Surface Science Group, School of Physical Science, University of Windsor, N9B 3P4 Windsor, Ontario, Canada Received March 13, 2002. In Final Form: May 17, 2002 Atomic force microscopy (AFM) and Fourier transform infrared spectroscopy (FTIR) are used to investigate molecular organization in Langmuir-Blodgett (LB) films of two kinds of lignins. The lignins were extracted from sugar cane bagasse using distinct extraction processes and are referred to here as ethanol lignin (EL) and saccharification lignin (SAC). AFM images show that LB films from EL have a flat surface in comparison with those from SAC. For the latter, ellipsoidal aggregates are seen oriented perpendicularly to the substrate. This result is confirmed by a combination of transmission and reflection FTIR measurements, which also point to lignin aggregates preferentially oriented perpendicularly to the substrate. For LB films from EL, on the other hand, aggregates are preferentially oriented parallel to the substrate, again consistent with the flat surface observed in AFM data. The vibrational spectroscopy data for cast films from both lignins show random molecular organization, as one should expect.
Introduction The surface properties, including molecular arrangements and rheological properties, of lignin macromolecules have been investigated mostly employing the LangmuirBlodgett (LB) technology and microscopy methods such as scanning electron microscopy (SEM), environmental scanning electron microscopy (ESEM), and atomic force microscopy (AFM).1-15 The use of spectroscopic methods is less common, and only a few reports are encountered * To whom correspondence should be addressed. E-mail: balogh@ if.sc.usp.br. † Instituto de Fı´sica de Sa ˜ o Carlos, Universidade de Sa˜o Paulo. ‡ Instituto de Quı´mica de Sa ˜ o Carlos, Universidade de Sa˜o Paulo. § Faculdade de Cie ˆ ncias e Tecnologia, Universidade Estadual Paulista. | School of Physical Science University of Windsor. (1) Constantino, C. J. L.; Dhanabalan, A.; Cotta, M. A.; Pereira da Silva, M. A.; Curvelo, A. A. S.; Oliveira, O. N., Jr. Holzforschung 2000, 54, 55-60. (2) Micic, M.; Benitez, I.; Ruano, M.; Mavers, M.; Jeremic, M.; Radotic, K.; Moy, V.; Leblanc, R. M. Chem. Phys. Lett. 2001, 347, 41-45. (3) Radotic, K.; Simic-Krstic, J.; Jeremic, M.; Trifunovic, M. Biophys. J. 1994, 66, 1763-1767. (4) Micic, M.; Jeremic, M.; Radotic, K.; Macers, M.; Leblanc, R. M. Scanning 2000, 22, 288-294. (5) Shevchenko, S. M.; Bailey, G. W.; Shane Yu, Y.; Akim, L. G. Tappi J. 1996, 79, 227-237. (6) Goring, D. A. I.; Voung, C.; Gancet, C.; Chanzu, H. J. Appl. Polym. Sci. 1979, 24, 931-936. (7) Luner, P.; Kempf, U. Tappi 1970, 53, 2069-2076. (8) Luner, P.; Roseman, G. Holzsforschung 1986, 40 suppl., 61-66. (9) Gilardi, G.; Cass, A. E. G. Langmuir 1993, 9, 1721-1726. (10) Constantino, C. J. L.; Juliani, L. P.; Botaro, V. R.; Balogh, D. T.; Pereira, M. R.; Ticianelli, E. A.; Curvelo, A. A. S.; Oliveira, O. N., Jr. Thin Solid Films 1996, 284-285, 191-194. (11) Baumberger, S.; Aguie´-Beghin, V.; Douillard, R.; Lapierre, C.; Monties, B. Ind. Crops Prod. 1997, 6, 259-263. (12) Barros, A. M.; Dhanabalan, A.; Constantino, C. J. L.; Balogh, D. T.; Oliveira, O. N., Jr. Thin Solid Films 1999, 354, 215-221. (13) Cathala, B.; Aguie´-Beghin, V.; Douillard, R.; Monties, B. Polym. Degrad. Stab. 1998, 59, 77-80. (14) Cathala, B.; Lee, L. T.; Aguie´-Beghin, V.; Douillard, R.; Monties, B. Langmuir 2000, 16, 10444-10448. (15) Atala, R. H.; Agarwal, U. P. Science 1985, 227, 636-638.
in the lignin literature. Raman images of the cell wall of wood tissues indicated a certain degree of organization of the lignin molecules with the phenyl rings oriented preferentially along the cell wall surface.15 Theoretical modeling using computer simulations showed that extracted lignin molecules also form planes.16 On the other hand, in condensed Langmuir films, the phenylpropane units do not lie flat on the plane but rather tend to form an arrangement in which the aromatic rings are somehow vertical to the air-water interface.5,7-9,12 The “picture” for lignin emerging from these studies resembles an almost spherical or disklike molecule that behaves as a sphere in solution.6 The lignin molecules have a tendency to self-organize on substrates, and the aggregates formed depend on the type of substrate and its interactions with the sample.4 The molecular aggregates have some mobility and may grow to form spherical or ellipsoidal structures (“onionlike”) in multilayers or in thin cast films.4 In the films, some “space” between the layers may exist, which is necessary to accommodate the functional groups, and may be filled with gas or solvent.2 Such porous structures were denoted in ellipsometric experiments performed on lignin LB films, for which low refraction indexes were measured.10 On one hand, the tendency to form planes makes the lignin macromolecules capable of “covering” rough surfaces, which can be useful in nanotechnology applications where a very smooth surface is required.16 On the other hand, rough structures formed for other lignins may also be useful in cases where large effective areas are of interest. The reasons why lignin molecules form very flat structures in some cases but not in others are not yet clear. One possible explanation could lie in the different chemical structures of the lignins investigated.17 Natural lignins cannot be isolated in an unaltered form,4 since linkages have to be cleaved in the delignification process.18 (16) Paterno, L. G.; Mattoso, L. H. C. Polymer 2001, 42, 5239-5245.
10.1021/la025729v CCC: $22.00 © 2002 American Chemical Society Published on Web 07/25/2002
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Thus, all lignin extraction processes lead to some degree of chemical structural modifications. Understanding how the extraction process affects the surface properties and spatial organization of extracted lignins may be useful for devising novel nanotechnological applications. In this paper, we show that even for lignins extracted with very similar procedures the surface morphology may vary considerably. This is illustrated with the investigation of LB and cast films of two kinds of lignin, referred to as ethanol lignin and saccharification lignin. They were extracted from the same source, sugar cane bagasse, using different extraction processes. The differences in chemical structure in the lignin fragments are therefore due only to the isolation method employed. AFM was used to investigate the surface morphology, and Fourier transform infrared (FTIR) spectroscopy techniques, in transmission and reflection-absorption modes, were used to probe the molecular organization in the lignin LB films. Experimental Section
Pasquini et al. Isotherm experiments were carried out in a KSV-5000 LB system placed in a class 10 000 clean room. Surface pressure isotherms were obtained with the Wilhelmy plate method. Ultrapure water (18.2 MΩ cm) supplied by a R060 Millipore filter followed by a Millipore-Q system was used as the subphase. The spreading solutions (0.2 mg mL-1) were prepared using THF. For LB deposition, 700 µL quantities of spreading solutions were employed. The lignin monolayers were transferred as Y-type LB films with a dipping speed of 0.5-3 mm min-1. Transfer was carried out at a constant surface pressure of 25 mN m-1. All depositions were carried out at room temperature (22 °C). The monolayers were transferred onto freshly cleaved mica substrates (5 monolayers) for AFM measurements. Mica has been selected as the substrate since it does not interact strongly with lignin. For reflection-absorption infrared spectroscopy (RAIRS) and transmission FTIR measurements, 30-layer LB films were deposited on Ag mirrors (100 nm) and on Si wafers, respectively. Cast films were also prepared on mica for AFM measurements and on ZnSe for FTIR transmission using solutions of lignin in THF with concentration of 0.2 mg mL-1. Lignin was dispersed in KBr powder to produce pellets (bulk sample). The UV-vis spectra for the bulk lignin were recorded on a Cary 50 Scan UV-vis spectrophotometer. For the LB and cast films, the spectra were recorded with a Hitachi U-2001 spectrophotometer. The transmission FTIR experiments were carried out using the bulk lignin, LB films deposited onto Si, and cast films deposited onto ZnSe. RAIRS measurements were recorded for LB films deposited onto Ag mirrors at an 80° incident angle. The spectra were recorded at the midinfrared region using a BOMEM DA3 Fourier transform infrared spectrometer equipped with a MCT detector, using 1024 scans and resolutions of 1 cm-1 for the pellet and 4 cm-1 for the LB and cast films. In these measurements, the sample chamber was evacuated to 0.8 Torr. Data acquisition and analysis were carried out using the WiRE software for Windows and Galactic Industries GRAMS/32 C software including the 3D package. AFM measurements were obtained with an atomic force microscope from Digital Instruments, Nanoscope IIIA, using the tapping mode.
The lignins were isolated from sugar cane bagasse using two extraction processes: ethanol-water and acetone-water (saccharification). Prior to the lignin extraction, the bagasse was wet-screened to remove the pith and pre-extracted with hot water, to remove low molecular weight polysaccharides and inorganic compounds, and with hot cyclohexane/ethanol to remove mainly polyphenols. The ethanol-water process was carried out in a pressurized autoclave at 180 °C, with a reaction time of 60 min, using an ethanol-water mixture (1:1, v/v) and a liquor-to-bagasse ratio of 20:1. The liquor resulting from the delignification process was acidified with concentrated H2SO4 until reaching pH ) 2. The lignin was then precipitated, filtered, and dried. The lignin extracted via the saccharification process was supplied by DEDINI S.A. Henceforth, the samples extracted with the ethanol/ water process will be referred to as ethanol lignins (EL), and those obtained from the acetone/water process will be referred to as saccharification lignins (SAC). To reduce polydispersity and differences in molecular weight, both lignins were submitted to a successive solvent fractionation with dichloromethane and acetone. In this fractionation process, the lignins were suspended first in dichloromethane, stirred for 30 min, and filtered. The solids were resuspended in the same solvent, stirred for an additional 30 min, and filtered. The filtrates were mixed, and the solvent was evaporated under reduced pressure, yielding the lignin fraction soluble in dichloromethane. The typical yield obtained for this fraction was around 5%. This first fraction obtained for all lignins was discarded, since the chemical analysis showed that the major part of the sample contained nonlignin products, probably polyphenols and other extractives that were not removed in the pre-extraction process. The remaining solids were dried under reduced pressure and treated with acetone in the same way, yielding the acetone lignin fraction (around 15% yield). In this work, results will be presented only for the fractions obtained with acetone, since they presented mean molecular weights closer to the one used in ref 12. The molecular weight of the lignins was determined by size exclusion chromatography (SEC) using three Plgel columns (500, 1000, and 10 000 Å), tetrahydrofuran (THF) as the solvent at 1 mL min-1, and polystyrene standards. EL presented Mw ) 2400 g mol-1, Mn ) 1360 g mol-1, and polydispersity of 1.8. SAC presented Mw ) 1900 g mol-1, Mn ) 1300 g mol-1, and polydispersity of 1.5. The content of functional groups per C9 units was calculated based on C, H, O elemental analysis, on methoxyl and hydroxyl contents determined by 13C NMR spectroscopy of acetylated samples in a Bruker AC200 at 50 °C using dimethyl sulfoxide (DMSO) as the solvent, and on the contents of carboxyl determined by the chemisorption method.20
EL and SAC lignins have the same hydroxyl content (1.1/C9 unit) and methoxyl content (0.93/C9 unit), slightly different carboxyl contents (0.4 and 0.3/C9 unit, respectively), and total oxygen contents of 3.0 and 2.2/C9 unit. The total carbonyl content was estimated from the areas of the FTIR spectra, relative to the aromatic ring absorption, leading to ratios of 2.1 and 1.6 for EL and SAC lignins, respectively. The detailed chemical characterization of these lignins along with other ones is discussed elsewhere.20 In summary, the major differences in the functional groups for EL and SAC lignins appear in the content of oxygen and carbonyl (including the carboxyl) groups. The influence of extraction methods on the Langmuir film characteristics was investigated in previous studies,12,17,19,20 where the presence of strong polar groups appeared to determine molecular packing. The surface potential is particularly sensitive to the polar groups. In the studies cited above, lignins obtained with extraction methods that yielded more carbonyl groups, such as EL (more than SAC), had lower values of surface potential. This means that carbonyl groups contribute negatively to the surface potential, that is, these groups have the oxygen in the carbonyl linkage pointing to the air when the film is in the compressed state. EL and SAC displayed extrapolated areas of 194 and 182 Å2 and maximum surface potentials of 110 and 200 mV, respectively, for
(17) Pasquini, D.; Dhanabalan, A.; Balogh, D. T.; Oliveira, O. N., Jr.; Curvelo, A. A. S. Proceedings of the 6th Brazilian Symposium on the Chemistry of Lignins and Other Wood and Components, Lorena, Sa˜o Paulo, Brazil, 2000. (18) Balogh, D. T.; Curvelo, A. A. S.; DeGroote, R. A. M. C. Holzforschung 1992, 46, 343-348.
(19) Pasquini, D.; Balogh, D. T.; Oliveira, O. N., Jr.; Curvelo, A. A. S. Proceedings of the 7th Brazilian Symposium on the Chemistry of Lignins and Other Wood and Components; Belo Horizonte, Minas Gerais, Brazil, 2001. (20) Pasquini, D.; Balogh, D. T.; Oliveira, O. N., Jr.; Curvelo, A. A. S. Manuscript in preparation.
Results and Discussion
Molecular Organization of Lignins in LB Films
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Figure 1. AFM images for 5-layer LB films from SAC lignin on mica.
films on pure water. Therefore, the carbonyl groups in EL again contributed negatively, which explains the lower surface potential compared to that of the SAC Langmuir film. The presence of polar groups led to films with a more uniform spatial arrangement, which seemed to be maintained during film transfer, yielding LB films with a lower roughness, as will be shown in the following. Figure 1a displays the AFM image for a 5-layer LB film of SAC lignin deposited onto mica. The root-mean-square (rms) roughness value is 3.8 nm, which is much higher than for LB films with the same number of layers fabricated with lignins extracted with the acetosolv process.1 The large roughness value is probably due to the ellipsoidal aggregates in the LB film of Figure 1, which are perpendicular to the substrate. This perpendicular arrangement has been so important that the surface roughness of the LB film is higher than for a cast film. Figure 1b shows a lower roughness for the cast film compared to the LB film (Figure 1a), which may be attributed to the absence of molecular orientation in the cast film. For EL, on the other hand, Figure 2a shows an AFM image of a more homogeneous 5-layer LB film. The rms roughness is only 0.24 nm, and large aggregates are not to be seen. Now, the roughness of the LB film is lower than for a cast film of EL (Figure 2b), as one should expect. It may be inferred that the tendency to form aggregates and consequently less smooth surfaces depends on the type of lignin, or more specifically in this case, on the isolation method employed.
Figure 2. AFM images for 5-layer LB films from EL on mica.
The conclusions from the analysis of morphology data above are verified by probing the molecular orientation in the films with a combination of transmission FTIR spectroscopy and RAIRS through the surface selection rules.21-24 Basically, the absorption is proportional to the scalar product between the dynamic dipole moment (µ′) of each normal mode and the electric field (E) of light at the surface (µ′‚E). In the transmission infrared experiment, the electric field lies on the plane of the substrate surface, while in RAIRS the main component of E is polarized perpendicular to the substrate surface. Here, these techniques are applied to LB films of lignins using the FTIR spectra from samples with random molecular orientation as a reference. The main goal is to check anisotropy in LB films rather than the specific molecular orientation in the films since lignins do not even have their molecular structures completely determined. Figure 3 presents the FTIR spectra for the bulk (transmission), cast film (transmission), and 30-layer LB film (RAIRS) of SAC lignin. The transmission spectrum for the 30-layer (21) Debe, M. K. Prog. Surf. Sci. 1987, 24, 1. (22) Decius, J. C.; Hexter, R. M. Molecular Vibrations in Crystals; McGraw-Hill: New York, 1977. (23) Born, M.; Wolf, E. Principles of Optics, 5th ed.; Pergamon Press: Oxford, 1975. (24) Greenler, R. G. J. Chem. Phys. 1966, 44, 310.
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Figure 3. FTIR spectra of SAC lignin for bulk (transmission), a cast film (transmission), and a 30-layer LB film (RAIRS).
Figure 4. FTIR transmission spectra for EL in bulk, a cast film, and a 30-layer LB film.
LB film could not be recorded at a reasonable signal/noise ratio. Four bands dominate the RAIRS spectra for the LB film: the vibrational mode at 1049 cm-1 attributed to the in-plane bending of the C-H aromatic group, the modes at 1150 and 1209 cm-1 assigned to C-O-C ether bending, and the 1725 cm-1 band attributed to CdO stretching. On the other hand, the vibrational modes at 1269 cm-1 assigned to the guaiacyl ring deformation and at 1511 cm-1 attributed to the CdC benzene ring vibrations have their relative intensities greater for the bulk spectra.25 The SAC cast film displays a spectrum closer to that of the SAC bulk sample. Considering the surface selection rules and the differences of relative intensities, it can be concluded that the LB films of SAC lignin exhibit an anisotropic molecular organization. The aggregates (ellipsoids) are preferentially oriented with the longer axis perpendicular to the substrate surface, while the cast film appears to have a random molecular orientation. Figure 4 shows the FTIR transmission spectra for the bulk sample, cast film, and 30-layer LB film from EL. The vibrational modes at 1032 cm-1 assigned to in-plane bending of the aromatic C-H and at 1156 and 1205 cm-1 due to C-O-C ether bendings dominate the transmission (25) Methods in Lignin Chemistry; Lin, S. Y., Dence, C. W., Eds.; Springer-Verlag: Berlin, 1992.
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Figure 5. UV-vis spectra for bulk samples of SAC and EL. The electronic spectrum for the KBr pellet is also presented.
spectrum in the LB film. However, the vibrational modes at 1511 and 1602 cm-1 assigned to the benzene ring vibrations26 have greater relative intensities in the bulk spectra. Analogous to what was observed for the SAC lignin, the spectrum of an EL cast film is closer to the spectrum of EL bulk. The differences in relative intensities for EL in bulk and in the LB film indicate that the latter is anisotropic. However, in contrast to the case of SAC, the aggregates are now preferentially oriented with the longer axis parallel to the surface of the substrate. The cast film again displays a random molecular organization. In summary, LB films from EL and SAC have aggregates preferentially oriented perpendicularly and parallel to the substrate, respectively. This is consistent with the larger roughness of the SAC LB film and the very flat surface observed for the EL LB film. We also tried to investigate the surface properties using Raman spectroscopy. However, during trial experiments with the lignin samples using a 633 nm laser line, a strong fluorescence was observed, which precluded a detailed analysis of the Raman data. Complementary, the UVvis spectra for bulk samples of SAC and EL are shown in Figure 5. A strong absorption band appears at ca. 315 nm, which is assigned to benzene rings in the lignin structures.26 Figure 5 also shows the spectrum of a neat pellet of KBr (no lignin) to confirm that there is no influence of KBr on the spectra of the lignins in the range considered. Conclusions LB films from EL and SAC lignins were found to exhibit surface morphologies that were distinctively different, even though the lignins were obtained under very similar procedures. The LB films were anisotropic in both cases, with ellipsoid aggregates being preferentially oriented perpendicularly and parallel to the substrate for the SAC and EL lignins, respectively. This anisotropy was reflected in rougher LB films for SAC, as demonstrated by AFM studies. These conclusions were corroborated by transmission and reflection FTIR results. Acknowledgment. This work was supported by FAPESP and CNPq (Brazil) and NSERC (Canada). LA025729V (26) Albinsson, B.; Li, S.; Lundquist, K.; Stomberg, R. J. Mol. Struct. 1999, 508 19-27.
