Natural Cellulose Fibers: Heterogeneous Acetylation Kinetics and

Biomacromolecules 2001, 2, 476-482

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Natural Cellulose Fibers: Heterogeneous Acetylation Kinetics and Biodegradation Behavior Giovanna Frisoni, Massimo Baiardo, and Mariastella Scandola* University of Bologna, Department of Chemistry “G. Ciamician” and Centro di Studio per la Fisica delle Macromolecole del C.N.R., Via Selmi 2, 40126 Bologna, Italy

Denisa Lednicka´ ,† Margo C. Cnockaert, Joris Mergaert, and Jean Swings Laboratorium voor Microbiologie, Vakgroep Biochemie, Fysiologie en Microbiologie, Universiteit Gent, K.L. Ledeganckstraat 35, B-9000 Gent, Belgium Received November 23, 2000; Revised Manuscript Received March 19, 2001

Steam-exploded fibers from flax (Linum usitatissimum) are heterogeneously acetylated using acetic anhydride and sulfuric acid as catalyst, with the aim to modify the surface properties without changing fiber structure and morphology. The acetylation reaction follows first-order kinetics up to a reaction time that depends on catalyst concentration (15 h when using 0.4 vol % of H2SO4 or 50 h with 0.1 vol %). The fibers undergo no structural and/or morphological changes under either reaction condition. On the contrary, surface damage and structural modifications appear after longer reaction times, when the reaction kinetics change. The extent of biodegradation of acetylated fibers, evaluated from the weight percent remaining after 13 days of exposure to previously isolated cellulolytic bacteria CellVibrio sp., decreases with increasing acetylation degree. After biodegradation the fibers show a higher acetyl content than before the experiment, indicating that the bacteria preferentially biodegrade unsubstituted cellulose, though also acetylated chains are cleaved. Biodegradable acetylated cellulose fibers with modified surface chemistry and unchanged structure are obtained for applications as polymer composite reinforcements. Introduction

Experimental Part

Natural cellulose fibers, characterized by low cost, biodegradability, and high specific mechanical properties can be employed as reinforcing agents in polymer composites. The interest toward natural-fiber-reinforced polymers has considerably grown in recent years.1-5 The mechanical properties of the final composite strongly depend on the degree of adhesion between matrix and fibers. The very high polarity of cellulose fibers6 conflicts with the hydrophobic character of most synthetic polymers,7 and the obtained composites often show poor properties. A means to improve surface adhesion between the continuous and the dispersed phases is to modify (by chemical or physical means) either the matrix8-10 or the fibers11-14 or both components.15 In this work cellulose fibers obtained from a typical European5 bast fiber crop, flax, are subjected to heterogeneous acetylation reactions. The aim is to decrease the high surface polarity of the natural fibers in view of their use as composite reinforcements.16 The chemical modification reactions are performed on fibers obtained from steam explosion, a fiber isolating technique that minimizes noncellulosic impurities. This paper describes the acetylation reaction kinetics for two different catalyst concentrations. The biodegradation behavior of the chemically modified fibers is also investigated by using previously isolated cellulolytic bacterial strains.17

Materials. Natural cellulose fibers derived from flax (Linum usitatissimum) and purified by steam explosion18-20 were supplied by the Institute for Applied Research (IAF, Reutlingen, Germany). The amount (in wt %) of noncellulosic substances in the fibers after steam explosion was as follows: lignin < 1, pectin < 1, hemicellulose < 4.21 All chemicals were from Sigma-Aldrich (reagent grade) and were used without further purification. Fiber Modification. The acetylation reactions were performed at constant temperature (30 °C) using 2 g of cellulose fibers (dried overnight at 80 °C under reduced pressure) and 10 mL of acetic anhydride containing sulfuric acid as reaction catalyst. Two different H2SO4 concentrations were used: 0.1 and 0.4 vol %, corresponding respectively to 0.9 and 3.6 wt % of the fiber weight. These reaction conditions are indicated throughout the paper as LC (low catalyst concentration) and HC (high catalyst concentration), respectively. A 40-mL glass reaction vessel was used, and the fibers were totally immersed in the liquid reagent. After selected reaction times, small amounts of fibers were withdrawn from the reaction mixture and washed to neutrality with distilled water and subsequently with ethyl ether. To collect a large enough number of samples over a broad reaction time range, several replicate reactions were run in parallel. The acetylated fibers were dried overnight at 80 °C under reduced pressure. The degree of substitution (DS) was determined by saponification and titration according to a previously described procedure.22

† On leave from Department of Biology and Ecology, University of Ostrava, Bra´fova 7, CS-70103 Ostrava 1, Czech Republic.

10.1021/bm0056409 CCC: $20.00 © 2001 American Chemical Society Published on Web 04/13/2001

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Natural Cellulose Fibers

Elemental analysis (by inductively coupled plasma atomic emission spectrometry) on the acetylated fibers showed no significant increase of sulfur (potential sulfate esters deriving from the use of sulfuric acid as reaction catalyst) compared with unmodified fibers. Characterization Methods. Transmission Fourier transform infrared spectroscopy (FT-IR) was performed by means of a Nicolet 210 spectrometer, taking 32 scans for each sample with a resolution of 4 cm-1. The fibers were pounded in a mortar cooled with liquid nitrogen and 1 mg of the obtained powder was dispersed in 150 mg of potassium bromide. Fibers and KBr were carefully dried before disk preparation and were subjected to FT-IR analysis immediately afterward. Powder wide-angle X-ray spectra were recorded from 2Θ ) 5° to 2Θ ) 60° with a Philips PW 1050/81 diffractometer, equipped with a graphite monochromator in the diffracted beam and using Cu KR radiation at λ ) 0.1542 nm (40 KV, 40 mA). The fibers (about 60 mg) were pressed into a small rectangular felt using a Carver laboratory press and an appropriate spacer. Scanning electron microscope (SEM) observations on the fibers before and after chemical modification as well as after biodegradation were carried out using a Philips 515. The fibers were laid down on the aluminum stub using a conductive adhesive tape and were sputter-coated with gold prior to measurements. Biodegradation Experiments. Biodegradation was quantified gravimetrically as percent of initial mass remaining after exposure (13 days) of fibers to cultures of selected bacteria. Previously isolated cellulolytic bacteria CellVibrio sp. strains R4001 and R407917 were used. The tests were performed in 20 mL of minimal medium (0.1% NH4Cl, 0.05% MgSO4‚7H2O, 0.005% ferriammonium citrate, in KH2PO4-Na2HPO4 buffer, 33 mM, pH 6.8)23 with 0.1 g of fibers, inoculated with about 106 colony forming units (CFU) per milliliter of either strain. Medium and fibers were preliminary autoclaved together. Incubation was conducted at 28 °C with reciprocal shaking during 13 days. After this incubation period the fibers were recovered by filtration over filter paper, rinsed with distilled water, and dried under vacuum to constant weight. Five replicate samples were run in each experimental condition and measurements were averaged. Blank experiments in 0.033 M phosphate buffer, pH 6.8, were run in parallel. Results Figure 1 compares the FT-IR spectrum of the unmodified fibers (UF) with the spectra of two acetylated samples obtained in HC reaction conditions after different reaction times: 7 h (HC7) and 30 h (HC30). The spectrum of native flax (curve UF) is very similar to that of pure cellulose reported in the literature,24-26 except for the presence of a weak shoulder at 1745 cm-1, associated with the CdO stretching vibration of residual noncellulosic impurities present in the flax fibers used in this work. The spectra of the acetylated samples show the expected acetyl group vibrations27 at 1745 cm-1 (ν CdO), 1375 cm-1 (ν C-CH3),

Figure 1. FT-IR spectra of unmodified cellulose fibers (UF) and of fibers esterified in HC conditions for 7 h (HC7) and 30 h (HC30). Arrows indicate the stretching bands typical of the acetyl group. The insert shows the method used to evaluate the extent of esterification, r ) A/B (see text).

Figure 2. Acetylation degree (r) of cellulose fibers as a function of time in HC (O) and LC (0) reaction conditions. Each r value is the average of five FT-IR measurements per sample (standard deviation 0.02). Full symbols (b) indicate samples subjected to biodegradation tests.

and 1235 cm-1 (ν C-O). The intensity of the acetyl bands increases with reaction time. Also the spectrum of UF cellulose shows an absorption at 1375 cm-1, attributed to the C-H bending vibration.28 After esterification, the added contribution of the acetyl (C-CH3) stretching vibration intensifies this absorption peak (see HC7 and HC30). As illustrated in the insert of Figure 1, the extent of acetylation is estimated in this work by the ratio (r) between the intensity (A) of the CdO stretching band at 1745 cm-1 and the intensity (B) of C-O stretching vibration of the cellulose backbone at 1058 cm-1. The r values of cellulose fibers esterified in LC and HC conditions are reported as a function of reaction time in Figure 2. The plotted r value is the average of five FT-IR measurements on each acetylated fiber sample (standard deviation 0.02). The scatter of the data points reflects

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Figure 3. WAXS spectra of unmodified fibers (0) and of fibers HC21 (b), HC28 (4), and HC30 ([). The insert shows a magnification of the region 5° < 2θ < 12°.

differences of acetylation degree in fibers recovered from replicate reactions. The curves drawn in Figure 2 represent the first-order kinetic law rt ) r∞ + Ae-kt

(1)

where r∞ is the plateau value (t ) ∞) and the term (r∞ + A) is the initial r value (t ) 0), i.e., the r value of the unmodified fibers (r ) 0.06, full square in Figure 2). Equation 1 very satisfactorily fits the experimental results obtained in both HC and LC conditions, up to given reaction times (HC, 15 h; LC, 50 h). Throughout the paper the acetylation extent of the fibers is quantified by r rather than by the more common DS parameter. The reason for this choice is the much easier experimental determination of r (by FT-IR) than of DS (by saponification and titration). For acetylated fibers with r e 0.40, the following linear relationship between r and DS has been found: DS ) 0.7r. Some of the HC acetylated fibers have been selected for the biodegradation experiments. They are shown in Figure 2 by full circles and are identified throughout the paper by their reaction time (HC1, HC7, HC28). Figure 3 summarizes the main structural changes observed in the wide-angle X-ray scattering (WAXS) spectra of the chemically modified fibers. Only the spectral region where such changes appear (5° < 2Θ < 30°) is shown for the sake of clarity. Unmodified flax fibers display the typical X-ray diffraction pattern of Cellulose I.29 Identical spectra are shown by all cellulose fibers acetylated in HC conditions up to 15 h and in LC conditions up to 50 h. This means that fibers whose acetylation reaction follows the first-order kinetic law (curves in Figure 2) maintain the original structure of unmodified flax. On the contrary, after longer reaction times in either experimental condition (HC or LC) the X-ray spectra of acetylated fibers gradually change: the intensity of the 2Θ ) 22° reflection decreases and a broad

reflection in the 2Θ range from 5° to 10° appears. As an example, Figure 3 shows three X-ray spectra of fibers obtained in HC conditions after 21, 28, and 30 h of reaction. Analogous spectral changes are shown by acetylated fibers obtained in LC conditions with reaction times longer than 90 h. Figure 4 collects SEM pictures of the unmodified fibers and of the three esterified samples selected for biodegradation tests. Native fibers have a rather smooth surface (micrograph a), a diameter in the range from 15 to 30 µm, and an average fiber length of 14 mm. Arrows indicate the characteristic pits at the surface of native flax fibers.30,31 After the acetylation treatments no significant morphological changes are observed by SEM in fibers HC1 (b) and HC7 (c), whereas the sample with the highest acetylation degree (HC28) shows fiber surface damage (d). The results of the biodegradation experiments are summarized in Table 1, which reports the fiber weight percent remaining at the end of the experiments and the change of acetylation degree upon biodegradation. After 13 days in buffer, all samples undergo almost the same weight loss (45%). Conversely, after 13 days of incubation with both bacterial strains the extent of biodegradation (UF > HC1 ≈ HC7 > HC28) depends on the initial acetylation degree of the fibers. The most extensively acetylated fibers investigated (HC28) show a small but still appreciable weight loss (about 20%) after 13 days of bacterial exposure. The results of Table 1 show that upon buffer exposure the degree of acetylation does not change, indicating no chemical hydrolysis of the ester group over the time scale of the experiments. The acetylated fibers exhibit higher r values after biodegradation than before the experiment: hence the fibers remaining after 13 days have a higher ester group content than before bacterial exposure. If the cellulolytic strains used in this work were able to cleave only unmodified cellulose chains, the product r(final) × wt % remaining (from Table 1) should yield a number comparable with the r value of the fibers before biodegradation. Figure 5 shows that this is not the case for the acetylated fibers tested in this work, independent of the strain used: the calculated value is always lower than the initial acetylation degree, implying that also acetylated cellulose chains are involved in the biodegradation process. Figure 6 shows the SEM pictures of unmodified cellulose fibers and of one of the acetylated samples (HC1) after 13 days of exposure to strain R4001. Comparison of Figure 6a-c with Figure 4a, shows that after bacterial exposure native flax fibers are visibly shortened but maintain both their cylindrical appearance and a rather smooth surface. On the contrary, in acetylated fibers HC1 (compare Figure 6d-f with Figure 4b) the surface is much rougher than before bacterial exposure and no significant changes in fiber length appear after biodegradation. Comparison of micrographs b and e in Figure 6 shows that after biodegradation unmodified fibers exhibit rather regularly spaced transverse failures, whereas HC1 fibers display longitudinal cracks. This last observation might suggest different routes of microbial attack to cellulose fibers before and after chemical modification.



Figure 4. SEM pictures of unmodified fibers (a) and of fibers HC1 (b), HC7 (c), and HC28 (d) before biodegradation. Same magnification. Arrows in (a) indicate the pits at the fiber surface. Table 1. Biodegradation Results after 13 Days of Exposure before biodegradation sample UFb HC1 HC7 HC28 a

ra 0.06 0.28 0.40 0.62

buffer wt remaining (wt %) 94.8 ( 0.4 96.8 ( 0.4 96.1 ( 0.5 96.2 ( 0.5

ra 0.06 0.30 0.40 0.61

strain R4001 wt remaining (wt %) 33.8 ( 2.8 47.8 ( 2.2 52.6 ( 0.3 80.1 ( 0.3

strain R4079 wt remaining (wt %)

ra n.d.c 0.45 0.49 0.65

24.0 ( 2.2 40.8 ( 5.9 40.5 ( 1.4 78.2 ( 1.7

ra n.d.c 0.48 0.67 0.73

Extent of acetylation by FT-IR (see text). b Unmodified fibers. c Not determined.

increases and a weak shoulder appears at 1540 cm-1 (black arrow). The absorption bands at 1650 and at 1540 cm-1 are characteristic of the amide (CO-NH) group and their appearance in the spectra of all fibers after bacterial exposure suggests the presence of residual proteins bound to the fibers.

Discussion

Figure 5. r value of fibers HC1, HC7, and HC28 before biodegradation (black bars) compared with the value of the product r(final) × wt % remaining after 13 days of exposure to strain R4079 (gray bars) and strain R4001 (white bars), see text.

The WAXS spectra of fibers HC1, HC7, and HC28 recovered after 13 days of exposure to strains R4001 and R4079 (not shown) are the same as spectra obtained before biodegradation. On the contrary, the FT-IR spectra of all biodegraded fibers show marked changes, exemplified in Figure 7 for fibers HC7 incubated with strain R4079. In addition to the intensification of the acetyl stretching bands (dots in Figure 7) leading to the mentioned r changes (Table 1), the intensity of the 1650 cm-1 band (broken arrow) also

In this work acetylation of cellulose fibers is carried out in a medium (acetic anhydride) that is the acetylating reactant and also the solvent for both catalyst and soluble reaction products. This strategy is adopted to minimize the chemicals used. Moreover, compared with classical cellulose acetylation reactions,32 mild reaction conditions are applied in order to modify only the fiber surface, while preserving the structure and good mechanical properties of native fibers. This point is very important for fibers to be used as reinforcement in polymer composites, where deterioration of fiber structure can severely impair the reinforcing efficiency. The results of this investigation show that over a reaction time that depends on catalyst concentration, the heterogeneous acetylation reaction follows a first-order kinetic law (Figure 2). In the presence of a great excess of acetic

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Figure 6. SEM pictures of unmodified fibers (a-c) and of acetylated fibers HC1 (d-f), after 13 days of incubation with strain R4001. Same magnification in micrographs a and d, in micrographs b and e, and in micrographs c and f.

Figure 7. FT-IR spectra of fiber HC7 before (a) and after (b) biodegradation (13 days) with strain R4079. Dots and arrows indicate the acetyl and amide stretching bands respectively (see text).

anhydride the reaction proceeds toward consumption of the other reagent, i.e., of the easily available hydroxyls of cellulose. Such groups are located at the fiber surface and represent only a small fraction of the total OH groups present in the cellulose fibers. This point is clearly demonstrated by the FT-IR spectrum of HC7 in Figure 1, where the intensity of the OH stretching band at 3400 cm-1 remains practically unaltered after acetylation.

In Figure 2 the curves represent the best fit of eq 1 to the experimental results, using the same plateau value (r∞ ) 0.4) in both HC and LC conditions. The following rate constants for the two catalyst concentrations are obtained: kHC ) 1.2 h-1; kLC ) 4.0 × 10-2 h-1. The maximum reaction rate, observed at the beginning of the esterification reaction, increases as expected with catalyst concentration, being 22 × 10-2 h-1 in HC conditions and 16 × 10-3 h-1 in LC reaction conditions. The plateau value is reached after 7-15 h in HC conditions, whereas after 50 h in LC conditions the r values are still lower than r∞. The fact that in both reactions a first-order equation with the same r∞ value satisfactorily fits the experimental data up to given reaction times supports the suggestion that the readily available OH groups at the fiber surface are the limiting factor in both cases. In neither reaction condition structural changes are observed by WAXS and SEM analysis of modified fibers with r e 0.40. Figure 2 shows that for reaction times longer than 15 h in HC conditions and 60-70 h in LC conditions the r values diverge from the first-order kinetic. The change of behavior observed suggests that new hydroxyl groups, likely to be located deeper into the cellulose fibers, become available for the acetylation reaction. It seems reasonable that at this stage diffusion mechanisms play an important role in making inner OH groups available to the reaction with acetic


anhydride. The WAXS spectra of modified fibers with r > 0.40, especially those obtained in HC conditions, show (Figure 3) a moderate but appreciable decrease of crystallinity (χc changes from 60% in UF to about 52% in HC30) and the appearance of a shoulder at 2Θ < 10° that grows with increasing substitution. The latter WAXS pattern change can be tentatively attributed to cellulose triacetate formation33,34 in some areas of the sample. SEM observations of the chemically modified fibers show surface damage only at high r values (Figure 4). In view of employing acetylated fibers as reinforcing agents in composites, reaction conditions that produce structural or morphological changes should be obviously avoided. This work shows that acetylated fibers free from unwanted structural modifications can be obtained in either LC or HC reaction conditions, up to substitution levels around r ) 0.30-0.40. This statement is supported by measurements of the mechanical properties of the fibers showing that the tenacity after acetylation is comparable to that of untreated fibers.35 The effect of acetylation on cellulose fiber biodegradation behavior is one of the aims of this investigation. This point may be relevant in composite applications where the matrix is a biodegradable polymer and biorecycling is envisaged as a suitable end-use disposal method. After 13 days of incubation the cellulolytic bacterial strains used in this work degrade unmodified fibers down to 25-35% of their initial weight (Table 1). None of the acetylated fibers investigated degrades so fast. For both cellulolytic strains, the extent of biodegradation decreases with increasing acetylation degree, in agreement with earlier results on the biodegradation of cellulose acetates with different degrees of substitution.36,37 It is recalled that cellulose in the present fibers is not homogeneously acetylated throughout the bulk of the fiber but that the acetyl groups are mainly located at the fiber surface. The bacteria used in this work seem to go for unmodified cellulose, whenever possible. A strong preference for unmodified cellulose is demonstrated by the r data in Table 1, where the acetylation degree of all fibers increases after bacterial exposure. However, it is interesting to note that the CellVibrio strains used in this investigation also cleave acetylated cellulose chains, as shown in Figure 5. SEM observations of the degraded fibers suggest that unmodified cellulose is attacked by the microorganisms at surface pits rather than at random along the fiber (the length of the fiber fragments in Figure 6b roughly corresponds to the distance between the transverse pits30,31 in the fibers before exposure, Figure 4a). Degradation of damaged cellulose fibers by “CellVibrio fulVus” (invalid name) at the damaged ends was earlier reported for cotton.38 In the case of acetylated fibers, the absence of apparent shortening suggests that the bacteria tend to avoid the acetylated surface looking for the unmodified internal cellulose chains, possibly attained through the fiber lumen. Indeed, it has been shown earlier by transmission electron microscopy that cells of “CellVibrio fulVus” are preferentially accumulating in the lumen of damaged cotton fibers.38 No appreciable variations of crystallinity degree are found by WAXS analysis of the


fibers after bacterial exposure. This indicates that the CellVibrio strains used in this work have similar activity toward the amorphous and crystalline phases of the substrate. This work shows that it is possible to modify the surface chemistry of cellulose fibers without changing their structural characteristics and that after a limited exposure time to cultures of selected cellulolytic bacteria the chemically modified fibers show considerable weight losses. The acetylated fibers described in this paper can find interesting applications as reinforcements in biodegradable composites. Acknowledgment. This paper has been carried out with financial support from the Commission of the European Communities, Agriculture and Fisheries (FAIR) specific RTD program, CT98-3919, “New functional Biopolymer-natural fiber composites from agricultural resources”. It does not necessarily reflect its views and in no way anticipates the Commission’s future policy in this area. References and Notes (1) Herrmann, A. S.; Nickel, J.; Riedel, U. Polym. Degrad. Stab. 1998, 59, 251. (2) Bledzki, A. K.; Gassan, J. Prog. Polym. Sci. 1999, 24, 221. (3) Glasser, W. G.; Taib, R.; Jain, R. K.; Kander, R. J. Appl. Polym. Sci. 1999, 73, 1329. (4) Nabi Saheb, D.; Jog, J. P. AdV. Polym. Technol. 1999, 18 (4), 351. (5) Agriculture and Fisheries specific RTD programme of the European Community, “New functional Biopolymer-natural fiber composites from agricultural resources”, FAIR CT98-3919. (6) Westerlind, Bo S.; Berg, J. C. J. Appl. Polym. Sci. 1988, 36, 523. (7) Wu, S. Surface and interfacial tensions of Polymers, Oligomers, Plasticizers and Organic Pigments. In Polymer Handbook, 4th ed.; Brandrup, J., Immergut, E. H., Grulke, E. A., Eds.; J. Wiley & Sons: New York, 1999; p VI/524 ff. (8) Takase, S.; Shiraishi, N. J. Appl. Polym. Sci. 1989, 37, 645. (9) Hedenberg, P.; Gatenholm, P. J. Appl. Polym. Sci. 1996, 60, 2377. (10) Hoffman, A. S. Macromol. Symp. 1996, 101, 443. (11) Trejo-O’Reilly, J. A.; Cavaille, J. Y.; Gandini, A. Cellulose 1997, 4, 305. (12) Bledzki, A. K.; Reihmane, S.; Gassan, J. J. Appl. Polym. Sci. 1996, 59, 1329. (13) Zadorecki, P.; Flodin, P. J. Appl. Polym. Sci. 1985, 30, 2419. (14) Raj, R. G.; Kokta, B. V.; Maldas, D.; Daneault, C. J. Appl. Polym. Sci. 1989, 37, 1089. (15) Joly, C.; Gauthier, R.; Escoubes, M. J. Appl. Polym. Sci. 1996, 61, 57. (16) Baiardo, M.; Frisoni, G.; Scandola, M.; Licciardello, A. Submitted for publication in J. Appl. Polym. Sci. (17) Lednicka´, D.; Mergaert, J.; Cnockaert, M. C.; Swings, J. Syst. Appl. Microbiol. 2000, 23, 1. (18) Kessler, R. W.; Becker, U.; Kohler, R.; Goth, B. Biomass Bioenergy 1998, 14 (3), 237. (19) Kessler, R. W.; Kohler, R. Chem. Technol. 1996, December, 34. (20) Vignon, M. R.; Garcia-Jaldon, C.; Dupeyre, D. Int. J. Biol. Macromol. 1995, 17 (6), 395. (21) Kohler, R. (IAF, Reutlingen, Germany), private communication. (22) Samios, E.; Dart, R. K.; Dawkins, J. V. Polymer 1997, 38 (12), 3045. (23) Delafield, P.; Doudoroff, M.; Palleroni, N. J.; Lusty, J.; Conopoulus, R. J. Bacteriol. 1965, 90, 1455. (24) O’Connor, R. T.; Dupre`, E. F.; McCall, E. R. Anal. Chem. 1957, 29, 998. (25) Hurtbise, F. G.; Kra¨ssig, H. Anal. Chem. 1960, 32, 177. (26) Higgins, H. G.; Stewart, C. M.; Harrington, K. J. J. Polym. Sci. 1961, 51, 59. (27) Rensch, H. P.; Riedl, B. J. Wood Chem. Technol. 1993, 13 (2), 167. (28) Liang, C. Y.; Marchessault, R. H. J. Polym. Sci. 1959, 39, 269. (29) Marchessault, R. H.; Sundararajan, P. R. Cellulose. In The Polysaccharides; Aspinall, G. O., Ed.; Academic Press: New York, 1982; Vol. II, Chapter 2, p 11. (30) Esau, K. Anatomy of Seed Plants; Wiley: New York, 1997; p 65.

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(31) Bowes, B. G. A colour Atlas of plant structure; Manson Publishing: 1996; Chapter 4, p 49. (32) Klemm, D.; Philipp, B.; Heinze, T.; Heinze, U.; Wagenknecht, W. ComprehensiVe Cellulose Chemistry; Wiley-VCH: Weinheim, 1998; p 164. (33) Watanabe, S.; Takai, M.; Hayashi, J. J. Polym. Sci.: Part C 1968, 23, 825. (34) Roche, E.; Chanzy, H.; Boudeulle, M.; Marchessault, R. H.; Sundararajan, P. Macromolecules 1978, 11 (1), 86. (35) Lips, D.; Feuz, L.; Ruffieux, K.; Wintermantel, E. Proceedings of the 7th Internationale TagungsStoffliche Verwertung nachwach-

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Natural Cellulose Fibers: Heterogeneous Acetylation Kinetics and

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