Biodegradation of Chemically Modified Flax Fibers ... - ACS Publications

Biomacromolecules 2004, 5, 596-602

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Biodegradation of Chemically Modified Flax Fibers in Soil and in Vitro with Selected Bacteria Alberto Modelli,†,‡ Giampaolo Rondinelli,‡ Mariastella Scandola,*,† Joris Mergaert,§ and M. C. Cnockaert§ Dipartimento di Chimica “G. Ciamician”, Universita` di Bologna, Via Selmi 2, 40126 Bologna, Italy, Centro Interdipartimentale di Ricerca in Scienze Ambientali (CIRSA), Universita` di Bologna, Piazza Kennedy 12, 48100 Ravenna, Italy, and Laboratorium voor Microbiologie, Vakgroep Biochemie, Fysiologie en Microbiologie, Universiteit Gent, K.L. Ledeganckstraat 35, B-9000 Gent, Belgium Received October 21, 2003; Revised Manuscript Received December 22, 2003

The extent and rate of degradation of flax (Linum usitatissimum) fibers, both in the native state and after surface chemical modification (acetylation or poly(ethylene glycol), PEG, grafting), was investigated under laboratory conditions in two different biodegrading environments. Degradation of the fibers under aerobic conditions by the action of the microorganisms present in soil is assessed with the ASTM 5988-96 method by monitoring carbon dioxide evolution. In vitro biodegradation experiments were carried out by exposing the fibers to a pure culture of CellVibrio fibroVorans bacteria and measuring the mass loss as a function of time. Despite the complexity of the system, the results of degradation in soil were satisfactorily reproducible, although the absolute rates were found to change in different experiments using the same soil. The degradation rate of acetylated fibers in soil nearly equals that of unmodified fibers, whereas in the pure culture, acetylated fibers biodegrade slower than native fibers. The opposite happens with the PEG-grafted fibers, which degrade slower than unmodified flax in soil and at a comparable rate upon in vitro exposure to the bacterial culture. The different biodegradation kinetics observed in the two biodegrading environments were attributed to differences of biocenoses, abiotic factors, and biodegradation assessing methods. Nevertheless, the final extent of biodegradation was the same for modified and unmodified fibers both in soil and in the pure culture, showing that the surface chemical modifications applied do not significantly affect biodegradability of the flax fibers. Introduction The widespread and ever growing use of plastics as a convenient alternative to a variety of other materials is based on the successful formulation of a wide range of polymer materials with mechanical and physical properties tunable to the requirements of the specific application. However, because of the increasing attention being paid to environmental problems, in many instances polymer resistance to biological degradation is regarded as a limit to be overcome, especially in plastics applications where a restricted servicetime is required and disposal after use is a concern. Hence, the search for materials that associate suitable properties with eco-compatibility. Because of their low cost, biodegradability, and high specific mechanical properties, natural vegetable fibers are good candidates as reinforcing agents in polymer composites.1-3 The mechanical properties of fiber-reinforced polymer composites depend on the efficiency of load transfer from continuous to dispersed phase, i.e., on the degree of adhesion between matrix and fibers. However, the high polarity of * To whom correspondence should be addressed. Phone: +39 051 2099577. Fax: +39 051 2099456. E-mail: [email protected]. † Dipartimento di Chimica “G. Ciamician”, Universita ` di Bologna. ‡ CIRSA, Universita ` di Bologna. § Universiteit Gent.

cellulose, which is the main component of the fibers, prevents good adhesion with most synthetic polymers that are essentially hydrophobic.4 The strength of the fiber-matrix interface can be improved by physical or chemical modification of the fibers.5-7 In particular, fiber surface polarity can be lowered by substituting the polysaccharide hydroxyls with a variety of less polar groups.6,8-11 An interesting development of the use of natural fibers as composite reinforcement regards their potential applications in association with biodegradable polymers. Composites for temporary applications that biodegrade in the environment after use can be developed. In this perspective, if the reinforcement is constituted of surface modified vegetable fibers, it is important to verify that after chemical modification the fibers maintain a sufficient degree of biodegradability. In this work, cellulose fibers obtained from flax (Linum usitatissimum) and carrying acetate groups8 or grafted poly(ethylene glycol) (PEG) chains11 at the surface are investigated with the aim of determining the extent of biodegradability in two different biodegrading environments (soil and selected bacteria in vitro). The effect of the chemical modification on the biodegradation kinetics is also analyzed. Degradation of the fibers in soil is monitored using the ASTM D 5988-96 method,12 devised for determining the

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Biodegradation of Modified Flax Fibers

degree and rate of aerobic biodegradation of plastic materials under laboratory conditions by the action of microorganisms present in soil. In an earlier work,13 this test method was shown to be reliable for determining the degree and rate of biodegradation of poly(3-hydroxybutyrate) and poly(-caprolactone), provided that KOH instead of Ba(OH)2 is used for the CO2 trapping solutions. The in vitro experiments are carried out by exposing the fibers to a previously isolated cellulolyticbacterium,14 nowclassifiedasCellVibriofibriVorans.15 Experimental Part Materials. Natural cellulose fibers from flax (Linum usitatissimum), bleached with hydrogen peroxide in the presence of NaOH and Na2CO3, were supplied by Linificio e Canapificio Nazionale S.p.A. (BG, Italy). The fibers were subjected to heterogeneous acetylation reaction using acetic anhydride.8 The reaction was carried out for 3.5 h at 30 °C, and 0.4% v/v of sulfuric acid was used as catalyst. A procedure described elsewhere11 was applied to graft at the fiber surface poly(ethylene glycol) monomethyl ether (PEG) with average molar mass of 750 g mol-1. Starch (soluble, A. C. S. reagent) was purchased from Sigma-Aldrich. Biodegradation in Soil. Desiccators of approximately 2-L internal volume, sealed airtight, were used for the ASTM D 5988-96 test method.12 In each test, 0.5 g of fibers (cut to 2 cm length) were mixed with 300 g of soil, sieved to 2-mm particle size. The soil was collected from the coastal area called Piallassa Baiona, close to Ravenna, Italy, and taken only from the surface (maximum depth 10 cm). The C:N ratio was adjusted to a value of 15:1 with ammonium phosphate solution. Distilled water was also added to bring the moisture content of the soil to about 90% of the moisture holding capacity, determined with test method ASTM D 425.16 The amount of CO2 produced was determined by titrating 0.4 M KOH solutions placed in the test and the blank (same conditions, but without fibers) desiccators with 0.25 M HCl to a phenolphthalein end-point. The frequency of titrations ranged from daily to weekly, depending on the degradation rate. The O2 content of the vessel (about 19 mmol) never fell by more than 10% in the interval between subsequent titrations. In addition to the fibers, a control substance (0.5 g of powder starch) was degraded in each experiment. Blank experiments were carried out in the same conditions, using soil without any additional carbon source. Each test (and blank) was run in triplicate. The temperature was 26 ( 2 °C for the first and second degradation experiment, and 20( 2 °C for the third experiment. In Vitro Bacterial Degradation. Biodegradation was quantified gravimetrically as percent of initial mass remaining after exposure (for a given time) of fibers to pure cultures of cellulolytic bacterium CellVibrio fibriVorans LMG 18561T () strain R4079; T, type strain).14,15 Members of this species were isolated as part of the dominant cellulolytic bacteria from soil. The strain was selected because of its rapid degradation potential of cellulose fibers (native as well as modified) in a standardized test system, yielding partially degraded material suitable for further analysis, and has been used earlier for similar degradation experiments.14,15

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The tests were performed in 20 mL minimal medium (0.1% NH4Cl, 0.05% MgSO4‚7H2O, 0.005% ferriammonium citrate, in KH2PO4-Na2HPO4 buffer, 33 mM, pH 6.8) with 0.1 g of fibers, inoculated with about 106 colony forming units (CFU) per mL. Medium and fibers were autoclaved together before inoculation. Incubation was conducted at 28 °C with reciprocal shaking. After selected incubation periods (13, 20, and 27 days), 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 noninoculated sterile 33 mM phosphate buffer, pH 6.8, were run in parallel. Results and Discussion Degradation in Soil. Three separate degradation experiments in soil were carried out using the ASTM D5988-96 method. The following flax samples (0.5 g) were analyzed: unmodified and acetylated fibers (first experiment, T ) 26 ( 2 °C), PEG-grafted fibers (second experiment, T ) 26 ( 2 °C), and unmodified and PEG-grafted fibers (third experiment, T ) 20 ( 2 °C). In addition to fiber samples, in each experiment, 0.5 g of starch powder were also degraded, in triplicate, as a control and reference material. The second experiment started 20 days after the first one, using the same soil stored at 4 °C. The third experiment was carried out five months later with a new sample of soil, taken from the same area. All degradations were followed during six months. The flax fibers used in this work contain 87.2% of carbohydrates, 92.5% of which are glucose.10 The main noncarbohydrate components are waxes and lignin. To calculate the expected total CO2 production during the biodegradation experiments, the simplifying assumption that the fibers are constituted of pure cellulose is introduced. The empirical formula of the repeating unit of both starch and cellulose is C6H10O5, with a carbon content of 44.44 wt %. Both polysaccharides are hydrophilic, and thermogravimetric analysis shows a water content of 5.0% in flax fibers and 12.1% in starch. Hence, complete degradation of 0.5 g of fibers (assimilated to plain cellulose) and starch is expected to give rise to 17.58 and 16.28 mmol of CO2, respectively. The total CO2 production, as a function of time, measured in the first experiment for unmodified flax fibers is reported in Figure 1A, where the rate of degradation (mmol of carbon per day) is given by the slope of the tangent to each curve. Despite the complexity of the heterogeneous system analyzed, the results of the triplicate experiments are quite reproducible. The induction time (if any) to start the degradation process is less than 1 day, indicating that the microorganisms contained in the soil can promptly produce suitable enzymes to degrade the fibers. Enzymatic degradation of solid substrates is a surface phenomenon. In agreement, there is a clear trend in the gradually decreasing rate with time, due to the decreasing surface of the fibers. A quantitative analysis of the data reveals that (i) the maximum degradation rate (0.48 mmol/ day (averaged on the three tests), corresponding to 2.7% of initial polymer degraded per day, is reached after 3 days;

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Modelli et al. Table 1. Maximum Rate (vmax, %/day) of Degradation, Half-Life Periods (t1/2, days) Relative to Production of 50% of the Total Amount of CO2 Actually Found or (in Parentheses) of the Expected Final Amount, Rate of Degradation Relative to Starch (vrel, Evaluated from the t1/2 Values), and Measured Final % of Degradation ( Relative Error (95% Confidence Limit) sample starch

unmodified flax fibers

acetylated flax fibers

1st expt (T ) 26 °C)

vmax 5.6 t1/2 15.5 (14.1) 106.3 ( 12.2% vmax 2.7

2nd expt (T ) 26 °C)

vmax 8.1 t1/2 7.0 (8.2) 90.0 ( 7.7%

t1/2 20.1 (24.3) vrel) 0.77 (0.58) 89.8 ( 7.4% vmax 2.4

3rd expt (T ) 20 °C)

vmax 5.1 t1/2 12.9 (12.7) 100.9 ( 6.2% vmax 4.6 t1/2 16.3 (18.1) vrel) 0.79 (0.70) 94.4 ( 4.4%

t1/2 21.7 (29.4) vrel) 0.71 (0.48) 82.1 ( 10.0% PEG-grafted flax fibers

vmax 3.8

vmax 3.0

t1/2 15.4 (18.8) t1/2 30.1 (31.9) vrel) 0.45 (0.43) vrel ) 0.43 (0.40) 87.4 ( 5.4% 97.2 ( 4.2%

Figure 1. Example of curves showing the total CO2 production as a function of time: (A) unmodified flax fibers (first experiment, triplicates), (B) PEG-grafted fibers (second experiment, triplicates).

(ii) the production of 8.79 mmol of CO2 (50% of the expected final production, assuming that also the non-carbohydrate components are degraded) is measured after about 24 days (half-life period, t1/2); (iii) after 40 days, when the degradation rate starts to decline significantly, the CO2 production corresponds to about 70% of degradation; (iv) after 6 months, the degradation rate is less than 0.01 mmol/day, and the measured total quantity of CO2 (averaged over the three tests) is 15.78 mmol. The final percentage degradation is thus 89.8%, with a standard error of (7.4% (95% confidence limit, calculated by evaluating the standard deviations of the tests and blanks, according to the procedure described in ASTM D 5988-96). Even considering the standard error, the evaluated percent degradation is less than 100%. The possible occurrence of anaerobic degradation mechanisms with methane production, which would escape detection by means of the ASTM method, cannot be ruled out. However, this finding is more likely to be ascribed to the presence of the non-carbohydrate components (12.8%) of the fibers, that might have undergone only partial degradation during the observation period. Moreover, some of the carbon is likely to have been incorporated in microbial biomass. Such carbon might not be accounted for when the CO2 evolved in the biodegradation tests is corrected relative to the blanks. For these reasons,

evaluation of the half-life period relative to the CO2 actually measured after six months (15.78 mmol instead of the expected 17.58 mmol) is probably more significant. This leads to a reduction of t1/2 from about 24 to 20 days. Table 1 collects the relevant results averaged over triplicate experiments obtained in the three series of degradation tests, i.e., initial (maximum) percent rate (Vmax), half-life period (t1/2) relative to the total CO2 actually produced or (in parentheses) to the expected amount, degradation rate of the fibers relative to the reference starch run in the same experiment (Vrel), and final percent degradation with corresponding error. In the first experiment, the tests with starch and acetylated flax fibers yielded a CO2 production as a function of time with trends very similar to those observed for unmodified fibers in Figure 1A. However, the absolute rate (as evaluated from the half-life period) is higher for starch and slightly lower for acetylated fibers compared to unmodified flax (Table 1). For PEG-grafted fibers, the CO2 evolution showed an induction period of about 2 days, barely visible in Figure 1B. The degradation rate of PEG-grafted fibers relative to the pertinent reference starch is slightly higher than 0.4 (Table 1). The third experiment (Table 1) allows for direct comparison between unmodified and PEG-grafted fibers. Their degradation rates relative to starch obtained in the third experiment are in good agreement with those measured separately in the first and second experiment (unmodified flax and PEG-grafted fibers respectively). A recent study13 has demonstrated that the kinetics of degradation of powder starch in soil are closely reproduced by simple first-order kinetics for solution reactions, in line with the classical Michaelis-Menten model for enzymatic catalysis, once the heterogeneity of the system is accounted



Figure 2. Plots of the ln(1 - DC) vs time (see text) for starch (diamonds) and unmodified (filled circles) and acetylated (squares) flax fibers, obtained from the first experiment.

for. Here we test whether the present data obtained with powder starch confirm the previous results, and we apply the same procedure to the data supplied by the fibers, for the sake of comparison. The integrated kinetic equation ln([C]/[C0]) ) -kt

(1)

for first-order reactions in solution can be formulated in terms of the degree of completion (DC) as ln(1 - DC) ) -kt

(2)

where DC ) ([C0] - [C])/[C0]. In the present case, the quantities corresponding to the concentration [C] at any given time and the initial concentration [C0] are the remaining and the initial moles of polymer carbon atoms, respectively, the difference [C0] - [C] being the moles of CO2 evolved. As mentioned above, the finding of a final percent degradation somewhat smaller than 100% is likely to be ascribed to the non carbohydrate components of the fibers and incorporation in biomass. Therefore, for each test, the DC has been evaluated as a function of time after normalization of the final percent degradation to 100%, that is, DC ) 1. Figure 2 shows the experimental results for starch, unmodified flax, and acetylated flax fibers (first experiment), plotted according to eq 2 up to DC > 0.7. Good linear correlations are found, with squared correlation coefficients of 0.9903, 0.9982, and 0.9989, respectively. Good linear regressions are obtained also from elaboration of the results of the second and third biodegradation experiments (not shown). The present results confirm earlier evidence13 that the degradation of powder starch in soil tends to follow the kinetics of first order reactions in solution. Moreover, noteworthy, the same rate law also describes the degradation of flax fibers (unmodified and modified). From the slopes of the linear regressions of Figure 2, the following half-life periods (days) can be evaluated: starch 14.9, cellulose 19.1, acetylated cellulose 20.5. These values are close to those

directly measured in correspondence with 50% of the final production of CO2, as calculated for the data obtained in the first experiment (see Table 1). Inspection of Table 1 reveals the complexity of the factors which may influence the degradation process in soil. Despite a good reproducibility of the results within the same experiment, notably different absolute degradation rates (evaluated from the half-life periods) for the same substrate are measured in different experiments. For instance, the biodegradation rate of starch in the second experiment is almost twice that obtained in the first one, although the temperature was 26 ( 2 °C in both cases, whereas in the third experiment the degradation rate of starch is slightly higher than in the first one, despite the lower temperature (20 ( 2 °C). These findings clearly prevent the possibility of direct comparison between the rates measured in different experiments for the same sample. However, the degradation rate (Vrel) of the fibers relative to the corresponding starch reference can be calculated (as the ratio between the halflife periods) within each experiment. Table 1 shows that the Vrel values found in the three different experiments are satisfactorily reproducible. This allows the use of all collected data to evaluate the degradation rate of the modified fibers with respect to natural flax. Comparison of the Vrel values shows lower degradation rates of the acetylated and PEGgrafted fibers compared to unmodified fibers (0.9 and 0.6, respectively). Although the rate difference between the unmodified and acetylated fibers could fall within experimental error, the lower degradation rate found for the PEGgrafted fibers seems to be significant. It is worth pointing out, however, that according to the present results the surface modifications do not cause a reduction of the final percent degradation. In Vitro Degradation. Unmodified flax fibers, acetylated, and PEG-grafted fibers were incubated in cultures of CellVibrio fibriVorans LMG 1856114,15 for different periods of time up to 27 days. Experiments in sterile buffer were run in parallel, and the weight remaining after certain time intervals is plotted in Figure 3 for the three types of fibers investigated. For all fibers, the maximum weight loss after 27 days in sterile buffer is around 7%. This decrease of weight is attributed to dissolution in the buffer solution of noncellulosic substances present in the flax fibers. Figure 3 also shows the results of the in vitro biodegradation experiments. All data are corrected by subtracting the observed weight loss in the corresponding blank experiment. The degradation behavior of unmodified and of PEG-grafted flax fibers is quite similar. The observed differences (PEG-grafted fibers degrading slightly faster than plain flax) lay within the experimental error. Conversely, acetylated fibers seem to follow slower biodegradation kinetics. Compared with unmodified fibers, the difference is most notable at the beginning of the experiment and tends to diminish with increasing incubation time. After 27 days of exposure, the weight (%) remaining is 28.1 ( 5.9, 17.2 ( 4.6, and 11.2 ( 2.0 for acetylated, unmodified, and PEG-grafted fibers, respectively.

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Figure 3. Fraction of initial weight remaining (%) after different exposure times to buffer (open symbols) and to Cellvibrio sp. R4079 strain (black symbols). Circle, unmodified fibers; triangle, acetylated fibers; diamond, PEG-grafted fibers.

Figure 4. Extent (%) of biodegradation (b), degree of acetylation r (4), and product [r × remaining fiber weight fraction] (0), as a function of incubation time (in vitro studies, see text).

In the case of acetylated fibers, the change of acetylation degree during the degradation experiment was investigated by FT-IR as previously described.8 From the FT-IR spectra of the fibers collected after the biodegradation experiments, the parameter r, defined as the ratio of the intensities of the CdO stretching band at 1745 cm-1 (associated with the ester group) and of the C-O stretching vibration at 1058 cm-1 (due to the cellulose backbone), was calculated and was compared with the initial r value. Figure 4 shows a plot of r (triangles) as a function of incubation time together with the increase of the extent of biodegradation (100 - % fibers remaining) with time (circles). In agreement with earlier in vitro experiments8 r is seen to increase with incubation time, implying an increase of ester group content in the fibers as biodegradation proceeds. This suggests that unmodified polysaccharide chains are preferentially degraded. To ascertain if the enzymes only cleave nonsubstituted cellulose chains or also attack, though to a lesser extent, acetylated cellulose, the product (r × remaining fiber weight fraction) was calculated. The obtained values, which should remain constant and equal to the initial value of r if biodegradation

Modelli et al.

were restricted to cleavage of unmodified cellulose, are plotted in Figure 4 (squares) and are seen to decrease with increasing incubation time. This result indicates that the CellVibrio strain used in this work shows a preference for plain cellulose but also cleaves acetylated sugar sequences. Carbohydrate esterases that are involved in the deacetylation of cellulose acetates have been described for several microorganisms, mainly fungi,17 but to our knowledge have not been reported for CellVibrio strains. The results observed with CellVibrio fibroVorans seem to support the idea that the bacterium has no or little esterase activity. The acetylated fibers, besides degrading slower than unmodified and PEG-grafted ones, also show a different morphology after intermediate bacterial exposure times. Figure 5 compares the scanning electron (SEM) micrographs of the three different flax fibers after 13 days (micrographs b, c, and d). A micrograph representative of fiber morphology before biodegradation (no differences are observed upon surface chemical modification) is also included (Figure 5a). As earlier reported8 under microbial attack, the acetylated fibers tend to show longitudinal cracks without evident shortening (Figure 5d), whereas unmodified fibers (Figure 5b) maintain the cylindrical shape and are remarkably shortened to a length that reflects the average distance (80120 µm) between adjacent pits.18 PEG-grafted fibers (Figure 5c) after 13 days of biodegradation look very similar to unmodified fibers, suggesting that in both cases the microorganisms attack the fiber surface preferentially at the pit locations causing the observed shortening. Conversely, the bacteria seem to avoid whenever possible the surface of acetylated fibers (Figure 5d), trying to penetrate the fiber interior through longitudinal cracks or through the fiber lumen. It has been shown earlier that CellVibrio fulVus preferentially accumulates in the lumen of damaged cotton fibers.19 However, after prolonged exposure, also acetylated flax fibers shorten and loose their cylindrical shape, as shown in Figure 5e. Conclusions The present results show that new types of chemically modified cellulosic plant fibers (in casu flax) show a biodegradability quite similar to those of unmodified fibers. However, different approaches in biodegradation experiments lead to different final degradation degrees and degradation rates. In general, experimental approaches range from exposure to natural in situ environments, containing an unknown biocenosis and without abiotic monitoring, to testing in highly defined synthetic media with selected cultures under controlled abiotic conditions. Also, different biodegradation assessment methods are applied, depending either on the measurement of degradation products (CO2 released in aerobic conditions of exposure) or of changes in the original material (gravimetry, molecular mass and composition, morphological changes, and surface and mechanical characteristics).20,21 It has been shown for other polymeric materials (e.g., bacterial plastics) that in situ degradation varies widely between different soils.22 Even when using standard methods, as used in our first approach



Figure 5. SEM micrographs ofj (a) fibers before biodegradation, (b) unmodified fibers after 13 days of exposure, (c) PEG-grafted fibers after 13 days of exposure, (d) acetylated fibers after 13 days of exposure, and (e) acetylated fibers after 27 days of exposure (in vitro studies).

(degradation in soil), different absolute results are obtained. Therefore, it is essential to either run the experiments on different materials in parallel or at least to include a wellcharacterized reference material in each of the independent experiments, as has been done and demonstrated in our soil experiments. Also when using the pure-culture approach, results depend on the bacterial strain used, as shown for different unmodified cellulosic fibers14 as well as acetylated flax fibers.8 The different relative degradation rates obtained in our experiments in soil versus in pure culture can be attributed to different biocenoses (complex biocenosis versus a pure culture of a CellVibrio strain), as well as differences in abiotic factors (temperature, oxygen, water content, pH, and minerals and other nutrients) and methods to assess biodegration (product formed versus substrate remaining). Despite the different biodegradation kinetics, our results clearly demonstrate that the chemical modifications of flax fibers do not significantly affect their total biodegradablity. These modified fibers offer new opportunities for applications in which biodegradability is a desirable feature. Limited lifetime polymer composites designed to biodegrade after use represent a new area of application of natural fibers, in addition to their well-established employment in association with polypropylene in interior paneling for the automotive industry. New bio-composites comprising surface modified vegetable fibers and a biodegradable polymer matrix may

find interesting applications as temporary structural materials in agriculture and construction. Acknowledgment. This work was supported by the Italian Ministry for University and Research (MIUR) and by the Commission of the European Communities, Agriculture and Fisheries (FAIR) specific RTD program, CT983919, “New Functional Biopolymer-Natural-Composites from agricultural resources”. The study does not necessarily reflect the views of the Commission and in no way anticipates the Commission’s future policy in this area. References and Notes (1) Hermann, A. S.; Nickel, J.; Riedel, U. Polym. Degrad. Stab. 1998, 59, 251. (2) Bledzki, A. K.; Gassan, J. 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) Wu, S. In Polymer Handbook, 4th ed.; Brandrup, J., Immergut, E. H., Grulke, E. A., Eds.; J. Wiley and Sons: New York, 1999. (5) Trejo-O’Reilly, J. A.; Cavaille, Y. C.; Gandini, A. Cellulose 1997, 4, 305. (6) Bledzky, A. K.; Reihmane, S.; Gassan, J. J. Appl. Polym. Sci. 1996, 59, 1329. (7) Raj, R. G.; Kokta, B. V.; Maldas, D.; Daneault, C. J. Appl. Polym. Sci. 1989, 37, 1089. (8) Frisoni, G.; Baiardo, M.; Scandola, M.; Lednicka`, D.; Cnockaert, M. C.; Mergaert, J.; Swings, J. Biomacromolecules 2001, 2, 476. (9) Baiardo, M.; Frisoni, G.; Scandola, M.; Licciardello, A. J. App. Polym. Sci. 2002, 83, 38. (10) Zini, E.; Scandola, M.; Gatenholm, P. Biomacromolecules, in press.

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(11) Scandola, M.; Sandri, S.; Baiardo, M.; Frisoni, G., European Patent Application EP 1 170 415 A1. (12) Annual Book of ASTM Standards; American Society for Testing and Materials: Philadelphia, PA, 1997; Vol. 08.03.A. (13) Modelli, A.; Calcagno, B.; Scandola, M. J. EnViron. Polym. Degrad. 1999, 7, 109. (14) Lednicka´, D.; Mergaert, J.; Cnockaert, M. C.; Swings, J. J. Syst. Appl. Microbiol. 2000, 23, 1. (15) Mergaert, J.; Lednicka´, D.; Goris, J.; Cnockaert, M. C.; De Vos, P.; Swings, J. Int. J. Syst. EVol. Microbiol. 2003, 53, 465. (16) Standard Methods for the Examination of Water and Wastewater, 17th ed.; American Public Health Association (APHA): Washington, DC, 1989.

Modelli et al. (17) Altaner, C.; Saake, B.; Tenkanen, M.; Eyzaguirre, J.; Faulds, C. B.; Biely, P.; Viikari, L.; Siika-aho, M.; Puls, J. J. Biotechnol. 2003, 105, 95. (18) Esau, K. Anatomy of Seed Plants; Wiley: New York, 1997. (19) Berg, B.; Hofsten, B. V.; Pettersson, G. J. Appl. Bacteriol. 1972, 35, 215. (20) Mergaert, J.; Ruffieux, K.; Bourban, C.; Storms, V.; Wagemans, W.; Wintermantel, E.; Swings, J. J. Polym. EnViron. 2000, 8, 17. (21) Augusta, J.; Mu¨ller, R.-J.; Widdecke, H. Chem. Ing. Technol. 1992, 64, 410. (22) Mergaert, J.; Webb, A.; Anderson, C.; Wouters, A.; Swings, J. Appl. EnViron. Microbiol. 1993, 59, 3233.

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