Acetyl xylan esterases in fungal cellulolytic systems

Volume 186. number

FEBS 2661

1

July 198.5

Acetyl xylan esterases in fungal cellulolytic systems Peter Biely’, Jiirgen Puls” and Henry Schneider* Division qf Biological Sciences, National Research Council of Canada, Ottawa KIA ORb, Canada, +Jnstitute of Chemistry, Siovak Academy of Sciences, 842 38 ~ratislava, Czechoslovakia and “Institute 01 Wood C~~e~istry and Chemical

Technology of Wood, BFH, Hamburg, FRG Received 1 March 1985, revised version received 30 April 1985 Xylan of several tree and other plant species is acetylated, and the enzymology of its degradation is unknown. The present study shows that enzymes liberating acetic acid from an acetylated xylan occur in several fungal eelIulolytic systems. Consequently, an additional step will have to be considered in current concepts for the degradation of hemicellulose by xylanases and xylosidases. Ace~yl group

Xyian

Esterase

I. INTRODUCTION The xylan of several species of trees [l-4] and other plants [5] is partially acetylated. The high content of acetic acid in hydrolysates of many wood and cereal species [6,7] suggests that xylan could be acetylated generally. Consequently, in at least some natural habitats, microorganisms are faced with the task of degrading an acetylated rather than a non-acetylated polymer. Although this aspect of microbial degradation of acetylated polysaccharides has been raised earlier by Bacon et al. IS], the enzyme systems that degrade xylan have been investigated using only the deacetylated material obtained on alkaline extraction of delignified holocellulose, hence a polymer that does not occur naturally. The sequence of reactions in the enzymic degradation of acetyl xylan is unknown. It has been reported that acetyl substituents inhibit the digestion of plant polysaccharides in ruminants [S,S]. This suggested that enzymic deacetylation may be prerequisite for the breakdown of acetyl xylan, or may enhance the rate of its hydrolysis by

* To whom correspondence

should be addressed

Issued as NRCC publication

no.24414

80

Hemicellulose

Fungi

gfycanases . The existence of enzymes that deacetylate xylan might have been anticipated, particularly in view of several papers demonstrating the occurrence of microbial esterases that act on various synthetic acetyi derivatives of monoand disaccharides [9- 131. However, esterases that act on acetyl xylan do not seem to have been described. Elucidation of the enzymology of degradation of acetyl xylan is of interest in efforts to understand the microbial decomposition of lignocellulosics, It is also of interest for technological reasons. Steam treatment of wood, which can be used as an initial step in separating cellulose from other components, produces a soluble hemicellulose fraction containing xylan [ 141. This fraction is a potential substrate for microbiological conversion to single cell protein and chemical fuels. This report shows that a xylan component in the hemicellulose fraction produced by steaming of wood is acetylated and also demonstrates the occurrence of fungal esterases that liberate acetic acid from acetyl xylan. 2. MATERIALS

AND METHODS

2.1. Enzymes The cellulolytic system of Schizophyilum

com-

Pubisbed by Ekevier Scrence Publishers 3. V. (Btomedrcal Division) 00145793/85/$3.30 Gj 1985 Federation of European Biochemical Societies

Volume 186, number 1

FEBS LETTERS

mune (ATCC 38548) was used as the crude culture

filtrate. It was prepared by growing the organism on 1% Solka Floe (Brown Co., Berlin, NH) for 9 days as described by Willick et al. [15]. The extracellular xylan-degrading enzyme system of Aureobasidium pullulans (CCY 27-l-32) was the spent culture medium containing 1% larchwood xylan (Sigma) and 0.67% yeast nitrogen base (Difco). The organism was grown for 4 days. Partially purified xylanase (EC 3.2.1.8, 1,4-,0-D-xylan xylanohydrolase) of S. commune (ATCC 38548) was a generous gift from Dr M. Yaguchi (National Research Council of Canada, Ottawa). Other sources of enzymes from organisms that degrade wood components included several commercial preparations of fungal cellulases: Trichoderma reesei RUT C-30 (Celluclast, Nova, Denmark), Aspergillus niger (Sigma) and Trichoderma viride (Onozuka, Japan). Esterases from non-microbial sources tested were porcine liver esterase, type II (EC 3.1.1.1), partially purified orange peel acetylesterase (EC 3.1.1.6), orange peel pectinesterase (EC 3.1.1.11) both as a suspension in 2.5 M ammonium sulfate and a lyophilized powder, and tomato pectinesterase. All were purchased from Sigma. 2.2. Acetylated and deacetylated xylan An acetylated xylan was obtained as a nondialyzable fraction of water-soluble, non-cellulosic polysaccharide produced by steaming birch at 200°C for 10 min. Deacetylation was accomplished using 0.2 M NaOH for 16 h at 22°C. After neutralization with HCl, the product was desalted by dialysis and lyophilized. Yields were usually about 40% indicating considerable degradation of the original material. 2.3. Assay of acetyl xylan esterase A 20% (w/v) solution of the non-dialyzable fraction of birch hemicellulose in 0.4 M phosphate buffer, pH 6.5, was mixed with an equal volume of appropriate diluted enzyme solution in distilled water and, after addition of a few drops of toluene as an antimicrobial agent, incubated at 30°C. The reaction was terminated by centrifugation followed by immediate analysis of the supernatant for acetic acid. In some experiments, reactions were terminated by freezing prior to analysis. One unit of acetyl xylan esterase is defined as the amount of

July 1985

enzyme liberating 1 pmol acetic acid from 10 mg of the substrate present in 0.1 ml of 0.2 M phosphate buffer in 1 min. Acetic acid was determined by HPLC using a Bio-Rad Aminex HPX-87H column and elution with 0.01 N HzS04. 2.4. Assay of acetylesterase Enzyme solution (lo-50 ~1) was mixed with 1 ml of a freshly prepared saturated solution of 4-nitrophenyl acetate in 0.2 M phosphate buffer, pH 6.5, and incubated at 22°C. Liberation of 4-nitrophenol was followed photometrically at 410 nm as a function of time. One unit of acetylesterase activity hydrolyzes 1 pmol of the substrate in 1 min. 2.5. Other methods Thin-layer chromatography of fragments of acetyl xylan liberated by enzymic action was carried out on cellulose (Merck, FRG) using ethyl acetate/acetic acid/water (18 : 7 : 10, v/v) as solvent. Reducing sugars were detected using anilinephthalate [ 161. Protein was determined according to Lowry et al. [17]. Enzymic liberation of methanol from citrus pectin (Sigma) was followed by gas chromatography. i3C-NMR spectra were recorded in D20 on a Varian CFT-20 spectrophotometer in the FT mode by a routine technique. 3. RESULTS AND DISCUSSION 3.1. Partial

characterization

of

acetyl

xylan

fraction

Since the non-dialyzable hemicellulose fraction had not been used previously as a substrate, some characterization was necessary. It contained 60% D-xylose and 10% acetic acid by weight, hence the molar ratio of D-xylose to acetic acid was -2: 1. The 13C-NMR spectrum (fig.1) revealed the presence of the carbon atoms of the acetyl group plus signals for sugar carbons that could not be interpreted unequivocally. However, after alkaline deacetylation, the signals arising from the acetyl group disappeared and the number of signals attributable to sugar carbons decreased, yielding a simple spectrum characteristic of a 1,4-P-linked polyxylose chain [18,19]. Additional evidence for the presence of acetylated xylose residues was provided by two81

Volume 186, number 1

FEBS LETTERS

July 198.5

XYl XYl2 XY’3 XYl4

1

Fig. 1. 13C-NMRspectra of non-dialyzable fraction of birch hemicellulose before (A) and after chemica1 deacetylation (B). The signals at about 21 and 174 ppm (arrows) correspond to the carbon atoms of the acetyl group.

dimensional thin-layer ~hromatographic studies of the components produced by digestion with partiaily purified xylanase from S. commune. Chromatograms run in one direction and stained for reducing sugars revealed the presence of xylose, xylobiose, xylotriose as well as several compounds with mobility different from that of 1,4+3xylooligosaccharides (fig.2A). These components contained acetyl groups, as shown by the changes in their mobility on chromatography in the second direction after deacetylation with ammonia vapour (fig.ZB). Several of the components were converted to the same xylooligosaccharide, indicating that the enzymic hydrolysate contained several xylooligosaccharides of the same chain length, but differing in the number of acetyl groups. In addition, some of the components appeared to be isomeric with respect to the position of the acetyl groups. The assignments are presented in table 1. It is to be noted that the above results indicate only that the fractions produced contained acetylated xylooligosaccharides, and not that the enzyme can break the bonds between two adjacent sugar residues that bear acetyl groups. The site of cleavage and the location of acetyl groups on the 82

234

567

89

Fig.2. Thin-layer chromatography of products of xylanase hydrolysis of acetyl xylan. (A) Development of the first dimension (fractions 1, 2 and 4 contain xylotriose, xylobiose and D-xylose, respectively), (B) development in the second dimension after deacetylation of the compounds on a dry chromatogram with ammonia vapour in a sealed chamber for 14 h at 22°C.

oligomers produced was unknown. In addition, the xylanase preparation was only partially purified and contained traces of acetyl xylan esterase activity.

Table 1 Assignments made to the reducing sugars liberated from acetyl xylan by xylanase from S. commune Fraction no.

Assignment Xyb, AcXyl4 XYlz AcXyl3 Xyl, diAcXyl4 AcXyl2 diAcXyl3 triAcXyl4 diAcXylza, triAcXylxb, tetraAcXyl4 diAcXyIza, triAcXylsb

ab Isomeric with respect to position of acetyl groups The

fractions

separated by were chromatography (fig.2)

thin-layer

FEBS LETTERS

Volume 186, number 1

Acetyl xylan esterase activity in fungal cellulolytic systems Incubation of non-dialyzable, water-soluble polysaccharide from birch with crude celluloseand xylan-hydrolyzing enzymes under nonbuffered conditions resulted in a drop of pH. The acidification was due to the liberation of acetic acid and showed characteristics of an enzymecatalyzed reaction. The liberation of acetic acid failed to occur with heat-denatured enzymes. In addition, the reaction in the presence of cellulase preparations of T. reesei and of A. niger had an optimum pH of 6.5. Were the reaction catalyzed simply by H+ or OH-, the rates would have been expected to increase as the pH deviated from neutrality. Other evidence supporting the enzymic nature of the deacetylation was the dependence of the amount of acetic acid liberated on the amount of protein and on the time of incubation (fig.3). Similar dependencies were obtained with the enzyme preparations of T. viride and A. pullulans (not shown). Analysis for acetic acid in this study employed HPLC. Gas chromatography was found un3.2.

A

Liberation

T. reesei cellulase;

suitable, since acetyl xylan decomposed thermally, presumably in the injector, to yield acetic acid. The fungal systems differed from the plant and animal esterases in specific activities towards two test substrates. The specific activities on acetyl xylan were invariably greater with fungal preparations (table 2) even though these preparations were crude mixtures of proteins. In addition, the fungal preparations exhibited relatively lower activity on 4-nitrophenyl acetate, as shown by the ratios of specific activities on 4-nitrophenyl acetate and acetyl xylan (table 2). However, the fungal preparations also contained xylanases which could depolymerize acetyl xylan. In principle, the specific activity of acetyl xylan esterases can depend on the molecular mass of the substrate. The extent to which the specific activities of fungal enzyme preparations were enhanced by depolymerization of the substrate is a matter that will require resolution. Acetyl xylan esterase activity was not associated with pectinesterase activity. Pectinesterase activity was low in the fungal preparations relative to that of partially purified pectinesterases.

B

Protein Fig.3.

July 1985

(mg)

Time (h)

of acetic acid from acetyl xylan as (A) a function of protein present in the incubation mixture 0, S. commune cellulase) and (B) a function of time of incubation with T. reesei cellulase 0.25 mg protein per 0.1 ml; 0, 0.5 mg protein per 0.1 ml).

Volume 186, number

1

July 1985

FEBS LETTERS Tabte 2

Specific esterase activities (U/mg protein) of various enzyme preparations on 4”nitrophenyl acetate and acetyl xylan, and their ratio Enzyme preparation

T. 7: A. S. A.

4-Nitrophenyl acetate esterase

reesei cellulase viride cellulase niger ~eilulase ~~~~~~~ cellulase ~~1~~~~~xylanase

Porcine Iiser Acetylesterase Pectinesterase Pectinesterase Pectinestexase

esterase (orange) (orange, susp.) (orange, solid) (tomato)

ACKNOWLEDGEMENTS We are indebted to Dr iM, Yaguchi for the sample of 3, co~~~~e xytanase, to Dr CR. cachedzie for providing the cellulotytic system of T. reesei and to Dr R. Roy for measuring the 13C-NMR spectra. We also thank Mr J. Labelfe for assistance in determining acetic acid. Critical reading of the manuscript by Dr H. Lee is greatly appreciated.

REVERENCED fl] Time& T.E. (f970) Wood Sci. Technol. I, 45-70. 121 Bouveng, H.O., Garegg, P,.J. and Lindberg, B. (1960) Acta C&em. Stand. 14, 742-748, 131 Sharkov, V.J., Kuibina, N.I. and Solovjeva, Y.P. (1960) Zh. Prikl. Khim. 33, 2571-2575. [4] Kardcsonyi, S., AlfGldi, J., KubaEkovd, M. and Stupka, i. (1983) Cellulose Chem. Technol. 17. 637-645. [S] Chesson, A., Gordon, A.H. and Lomax, J.A. (1983) J. Sci. Food Agric. 34, 1330-1340. [6] Rydholom, S. (1965) in: Pulping Process, pp.94-96, Interscience, New York.

84

0.061 0.027 0.135 0.363 I.75 49.7 1.47 0.14 0.16 0.39

Acetyl xylan esterase 0.043 0.08 0.04I 0.072 0.39 0.01 I 0.022 0.008 0.004 0.025

Ratio

I.4 0.34 3.3 5.0 4.5 4518.2 66.8 17.5 40.0 15.6

[71 Puls, J. (1983) Comm. Eur. Communit~es~ [Rep.] EUR 8245, Energy Biomass, 863-867. [8] Bacon, J.&D., Gordon, A.H. and Morris, E.A. f1975) B&hem. J. 149, 485-487. f9f Mandels, M. and Reese, E.T. (1960) J. Bacterial. 79, 816-826. flO] Frohwein, Y.Z., Zori, U. and Leibowitz, J. (1963) Enzymologia 26, 193-200. [ll] Reese, E.T., Lola, J.E. andparrish, 1F.W. (1969) J. Bacterial. 100, 1151-1154. [12] Renter, G. and Huettner, R. (1977) Z. Allot. Mikrobiol. 17, 149-151. I I131 Hrmovii, M., &urdik, E., KoSik, M., Gemeiner, I?, and PetruS, L. (1983) Z. Allg. Mikrob~oI. 23, 303-312. 1141 Dietrichs, H.H., Sinner, M. and Puls, .I. (1978) Holzforschung 32, 193- 199. WI Willick, GE, Morosoli, R., Sdigy, V.L., Yaguchi, M. and Desrochers, M. (1984) J. Bacterial. 159, 294-299. I161 Patridge, S.M. (1949) Nature 164, 443. 1171 Lowry, O.H., Rosebrough, N.J., Farr, A.L. and Randall, R.J. (1951) J. Biol. Chem. 193, 265-275. [la Gast, J.C., Atalla, R.H. and McKelvey, R.D. (1980) Carbohydr. Res. 84, 137-146. WI KovriE, P. and Hirsch, J. (1982) Carbohydr. Res. 100, 177-193.