Lignocellulose biotransformation with immobilized cellulase, d -glucose oxidase and fungal peroxidases

Lignocellulose biotransformation with immobilized cellulase, D-glucose oxidase and fungal peroxidases J. ,Lobarzewski and A. Paszczyfiski Department o f Biochemistry, University o f Maria Curie-Sklodowska, 20-031 Lublin, Poland

(Received 21 November 1984; revised 29 April 1985) Three enzymes, cellulase [see 1,4-(l,3;1,4)-(3-D-glucan 4-glucanohydrolase, EC 3.2.1.4], D-glucose oxidase (fl-D-glucose : oxygen 1-oxidoreductase, EC 1.1.3.4) and peroxidase (donor:hydrogen peroxide oxidoreductase, EC 1.11.1.7) immobilized on glass beads, have been incubated with lignoeellulose. Fungal peroxidases from Trametes versicolor and Inonotus radiatus when mixed with cellulase and D-glucose oxidase were able to liberate phenolic compounds and D-glucose from lignocellulose. Three lignin monomers were identified. When the immobilized enzymes were incubated individually with lignocellulose they did not degrade lignin.

Keywords: Enzymes; immobilized enzymes; lignocellulose; cellulase; D-glucose oxidase; peroxidases

Introduction Enzymatic biodegradation of lignocellulose is a very complex biochemical process and is still not completely understood) '2 In naturally occurring lignocellulose, lignin protects cellulose and hemicelluloses against enzymic degradation. According to Crawford 1 three methods can be used to study lignocellulose biodegradation: the classical non-isotopic method; tile radioisotopic method where lignocelluloses are labelled with 14C;3-13 and the use of lignin model compounds. All three methods have thus far involved whole fungal cultures. The use of free enzymes has never been described.3-13 The present paper shows, for the first time, the possibility of lignocellulose biotransformation using three immobilized enzymes: cellulase [see 1,4-(I,3:I,4)@Dglucan 4-glucanohydrolase, EC 3.2.1.4], D-glucose oxidase (/3-D-glucose:oxygen 1-oxidoreductase, EC 1.1.3.4) and fungal peroxidase (donor: hydrogen peroxide oxidoreductase, EC 1.11.1.7). The proposed interaction of these enzymes in the process of lignocellulose biotransformation is shown in Figure 1.

Materials and methods E n z y m e source Trametes versicolor (L. ex Fr/Quel) fungus was grown in submerged cultures (5 litre fermenter) as described previously. 14 The mycelium was homogenized in a Waring blender and the homogenate was salted out using (NH4)2SO4 at 80% saturation. After dialysis the cell free extract was lyophilized. The cell free extract was further purified by affinity chromatography according to the procedure used in previous studies. 1,* In this procedure phenol oxidase activity was removed and a homogeneous peroxidase preparation (isoelectric point, pI = 3.8) was obtained. 14' is The second

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source of fungal peroxidase was Inonotus mdiatus (Sow. ex Fr. P. Karst HMJPC No. 4335) which was grown in stationary cultures as described previously) 6 Inonotus radiatus produces extracellular peroxidase with no phenol oxidase activity. Peroxidase was isolated from the lnonoms radiatus culture filtrate after two weeks of growth using twice the filtrate volume of cold acetone. The precipitate was dissolved in 0.05 M phosphate buffer, pH 7, dialysed and lyophilized before use. Cellulase complex was obtained from Novo (Denmark) and D-glucose oxidase from Aspergillus niger produced by Sigma (USA). Enzyme immobilization

All the enzymes used in the experiments were immobilized separately on silanized porous glass beads (Pierce Chemical Co., USA) with pore diameter 500 )~ and particle size 125-177 ~m, after their activation with glutaraldehyde (Merck, FRG) using the procedure described previously./7

0141--0229/85/110564--03$03.00 © 1985Butterworth& Co. (Publishers)Ltd

Lignocellulose biotransformation: J. Zobarzewski and A. Paszczy4ski

Lignocellulose Wheat straw was milled in a ball grinder and sieved to obtain the 0.01 mm fraction. This fraction was further purified according to the procedure used by Haider and Trojanowski) 8

E n z y m e activity determinations Peroxidase and phenol oxidase activities were determined by the method of,Lobarzewski. 14 Cellulase (including ~-D-glucosidase [13-D-glucoside glucohydrolase, EC 3.2.1.21] activity) and D-glucose oxidase activities were determined according to Rogalski et al. 19 and Bright and Appleby, 2° respectively. Protein was determined according to the method of Schacterle and Pollack? 1

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Experimental conditions The immobilized enzymes were dispersed in 0.05 M phosphate buffer, pH 7, and incubated for 48 h on a shaker at 35°C with the lignocellulose preparation. The control sample contained 1 g silanized glass beads and 300 mg lignocellulose preparation mixed with 30 ml 0.05 M phosphate buffer, pH 7. Subsequent control samples contained 1 g immobilized cellulase, D-glucose oxidase or peroxidase, and each enzyme was separately mixed with 300 mg lignocellulose preparation and 30 ml 0.05 M phosphate buffer, pH 7. The supernatant of these mixtures was used for spectrophotometric measurements and, after acidified ethyl ether extraction, for thin layer chromatography (t.l.c.). In the main experiments the mixture contained the three immobilized enzymes, i.e. cellulase, D-glucose oxidase and a fungal peroxidase preparation. In such cases 1 g of each immobilized enzyme and 300 nag lignocellulose were suspended together in 30 ml 0.05 M phosphate buffer and then incubated for 48 h on a shaker at 35°C. Assays were performed after 0 and 48 h of incubation. The u.v. spectrum was recorded in a double beam u.v. vis spectrophotometer. After 48 h of incubation each material was centrifuged for 20 rain at 10 000g. The supernatants were lyophilized and dissolved in 5 ml water. All these samples were analysed by t.l.c, and column gel chromatography with Sephadex G-15, G-150. T.l.c. was performed on t.l.c. glass silica gel 60 with a concentration zone. The layer was 0.25 mm thick (Merck, FRG). After extraction from water solution by acidified ethyl ether the phenolic compounds were put on t.l.c, plates and developed in benzeneethanol-propionic acid ( 8 8 : 8 : 2 v/v) or benzenemethanol-propionic acid (11:2:1 v/v) systems. The phenolic compounds on the chromatograms were viewed in u.v. light and after spraying the plates with a mixture containing 5 ml 1% sulphanilamide in 10% HC1, 5 ml 5% NaNO2 and 40 ml butan-l-ol.

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The theoretical basis of our experiments was the assumption that, under the experimental conditions described, wheat straw lignocellulose could be biotransformed in the presence of three enzymes. This means that cellulase (containing ~-D-glucosidase 19) could release D-glucose from cellulose, D-Glucose oxidase could catalyse the H202 generation necessary for peroxidase action. This scheme is presented in Figure 1. The enzymes were immobilized on glass beads for easier removal of soluble products of the

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Papers enzymatic reaction on lignocellulose and to stabilize enzyme activity. 22' 23 In control experiments each of the immobilized enzymes was incubated with lignocellulose separately. The u.v. spectrum of the post-incubation solution was mainly flat, with the exception of curve 4 in Figure 2. This indicates the presence of D-glucose, which was identified by the reduction and also by a positive D-glucose oxidase test. The main results of the experiment are shown in Figure 3. After incubating lignocellulose with the three enzymes, the presence of D-gluconic acid in the solution was established spectrophotometrically from the increase in maximum absorbance at 220 nm and confimled by gel chromatography on Sephadex G-15 (Figures 3 and 4). In addition to the above observations, the increase in absorbance peaks at 280 and 320 nm indicated the action of peroxidase and H20= on lignin (Figure 3). The products of lignocellulose biodegradation, i.e. carbohydrates and phenolic compounds, were further analysed on Sephadex G-150 and G-15 columns. The Sephadex G-150 column analysis revealed only substances of low molecular weight. Results were confirmed by Sephadex G-15 column analysis as shown in Figure 4; among the soluble products of lignocellulose biotransformation, twelve phenolic compounds as well as D-gluconic acid were discovered. The phenolic compounds were separated and identified using t.l.c. (Figure 5). Three of them were identified as caffeic, sinapic and COUlnaric acids (Figure 5). These phenolic compounds are monomers of the lignin molecule. The results confirm our earlier theoretical assumption that the three enzymes used in our experiments are able to biotransform lignocellulose molecules. These observations were only possible because the enzymes used were immobilized and also imitated the function of enzymes bound to the cell wall. The process of lignocellulose biotransformation in the presence of the immobilized enzyme system was repeatable after removal of the reaction products, i.e. D-glucose, D-gluconcic acid and phenolic compounds. All these compounds decreased the enzymatic reaction velocity as a result of product repression. The proposed interaction of enzymes degrading cellulose and lignin (Figure 1) could take place in nature and play a very important role in lignocellulose degradation.

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References 1 Crawford, R. L. Lignin Biodegradation and Transformation J. Wiley and Sons, New York, 1981 2 Kirk, K. T. in Recent Advances in Lignin Biodegradation Research (Higuchi, T., Chang, H. and Kirk, T. K.. eds), Uni Publishers Co., Ltd, Tokyo, 1983, pp. 1 11 3 Crawford, D. L. and Crawford, R. L. Enzyme Microb. Technol. 1980, 2, 11 4 Haider,K. and Trojanowski, J. Arch. Microbiol. 1975, 105, 33 5 Haars, A. and Htlttermann, A. :Vaturwissenschaften 1980, 67, 39 6 Haider, K. and Trojanowski, J. tlolz/orschung 1981, 35, 33 7 Highley,T. L. Can. J. For. Res. 1982, 12,435 8 ltllttermann, A., Gebauer, M.. Volger, Ch. and R~sger, Ch. Holzforschung 1977, 31, 83 9 Kaplan, D. L. and Hartenstein, R. Soil Biochem. 1980, 12, 65 10 Kirk, T. K. in Biological Transformation of Wood by Microorganisms (Licse, W., ed.), Springer-Verlag, Berlin Heidelberg New York, 1975, pp. 153 11 Kirk, T. K. and Chang, tt. M. tlolzJbrschung 1975, 29, 56 12 Kirk, T. K. and Nakatsubo, I' Biochim. Biol)hyx..Iota 1983, 756,376 13 Leyonen-Munoz, E., Bone, D. El. and Daugulis, A. J. Eur. ,I. Appl. Microbiol. Biotechnol. 1983, 18, 120 14 Lobarzewski, J. Int. J. Biol. Macromol. 1981, 3, 77 15 Lobarzewski, J. and Trojanowski, J. Acta Biochim. Polon. 1979, 26, 309 16 Lobarzewski,J. Acta Microbiol. Polon. 1974, 6, 1 17 Lobarzewski,J. and Paszczyfiski, A. Biotechnol. Bioeng. 1983, 25, 3207 18 ttaider, K. and Trojanowski, J. Arch. Microbiol. 1975, 105, 33 19 Rogalski, J., Szczodrak, J. and llczuk, Z. Acta Microbiol. Polon. 1983, 32, 363 20 Bright,tt. J. and Appleby, M. J. Biol. Chem. 1969, 244, 3625 2t Schactcrle, G. R. and Pollack, R. L. Anal. Biochem 1973.51, 654 22 Lobarzewski, J. Holz/orschung 1984, 38, 105