Biotechnology in the Pulp and Paper Industry

57 Advances in Biochemical Engineering Biotechnology Managing Editor: T. Scheper Springer Berlin Heidelberg New York...

Author: K.E.L. Eriksson

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57

Advances in Biochemical Engineering Biotechnology Managing Editor: T. Scheper

Springer Berlin Heidelberg New York Barcelona Budapest Hong Kong London Milan Paris San ta Clara Singapore Tokyo

Biotechnolgy in the Pulp and Paper Industr~Y Volume Editor: K.-E. L. Eriksson

With Contributions by M. Akhtar, D.S. Argyropoulos, P. Bajpai, P. K. Bajpai, R. A. Blanchette, J. Buchert, J. F. D. Dean, K.-E. L. Eriksson, R. L. Farrell, M. Guenette, K. Hata, S. C. Johnsrud, T. K. Kirk, R. C. Kuhad, P. R. LaFayette, S. B. Menachem, S. A. Merkle, A. Singh, A. Suurn~ikki, M. Tenkanen, J. S. Tolan, L. Viikari, M. B. Wall

With 41 Figures and 52 Tables

Springer

ISSN 0724-6145 ISBN3-540-61868-6 Springer-Verlag Berlin Heidelberg NewYork

Library of Congress Catalog Card Number 72-152360 This work is subject to copyright. All rights are reserved, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, re-use of illustrations, recitation, broadcasting, reproduction on microfilms or in other ways, and storage in data banks. Duplication of this publication or parts thereof is only permitted under the provisions of the German Copyright Law o f September 9,1965, in its current version, and a copyright fee must always be paid. Springer-Verlag Berlin Heidelberg 1997 Printed in Germany The use of registered names, trademarks, etc. in this publication does not imply, even in the absence o f a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. Typesetting: Macmillan India Ltd., Bangalore-25 SPIN: 10550104 02/3020 - 5 4 3 2 1 0 - Printed on acid-free paper

Managing Editor Professor Dr. T. Scheper Institute of Technical Chemistry, University of Hannover Callinstral3e 3, D - 30167 Hannover, FRG Volume Editor Professor Dr. K.-E. L. Eriksson Center for Biological Resource Recovery, The University of Georgia, A214 Life Sciences Building, Athens, GA 3 0602-7229/USA

Editorial Board Prof. Dr. W. Babel

Center of Environmental Research Leipzig-Halle GmbH Section of EnvironmentalMicrobiology PeermoserstraBe 15 D - 04318 Leipzig/FRG

Prof. Dr./4~ W. Blanch

University of California Department of Chemical Engineering Berkely, CA 94720-9989/USA

Prof. Dr. Ch. L. Cooney

Massachusetts Institute of Technology Department of Chemical Engineering 25 Ames Street Cambridge, MA 02139/USA

Prof. Dr. S.-O. Enfors

Department of Biochemistry and Biotechnology Royal Institute of Technology Teknikringen 34, S - 100 44 Stockholm/Sweden

Prof. Dr. K.-E. L. Eriksson Center for Biological Resource Recovery The University of Georgia A214 Life Sciences Building Athens, GA 30602-7229/USA Prof. Dr. A. Fiechter

Institute of Biotechnology EidgenOssische Technische Hochschule ETH-HOnggerberg,CH-8093 Ztirich/Switzerland

Prof. Dr. A. Iv[ Klibanov Massachusetts Institute of Technology Department of Chemistry Cambridge, MA 02139/USA Prof. Dr. B. Mattiasson

Department of Biotechnology Chemical Center, Lund University P.O. Box 124, S -221 00 Lund/Sweden

Prof. Dr. S. B. Primrose

21 Amersham Road

High Wycombe, Bucks HP13 6QS/UK

VI

Editorial Board

Prof. Dr. H. d. Rehm

Westf'alische Wilhelms Universit/it Institute of Microbiology Corrensstr. 3, D - 48149 Miinster/FRG

Prof. Dr. P. L. Rogers

Department of Biotechnology Faculty of Applied Science The University of New South Wales Sydney2052/Australia

Prof. Dr. H. Sahm

Institute of Biotechnology ForschungszentrumJiilich GmbH D - 52428 Jtilich/FRG

Prof. Dr. K. Schi~gerl

Institute of Technical Chemistry University of Hannover Callinstr. 3, D - 30167 Hannover/FRG

Prof. Dr. G. T. Tsao

Director, Lab. of Renewable Resources Eng. A. A. Potter Eng. Center, Purdue University West Lafayette, IN 47907/USA

Prof. Dr. K. Venkat

Phyton Inc., 125 Langmuir Lab. 95 Brown Road, Ithaca, NY 14850-1257/USA

Prof. Dr. John Villadsen

Department of Biotechnology Technical University of Denmark Bygning223, DK-2800 Lyngby/Denmark

Prof. Dr. U. yon Stockar

Swiss Federal Institute of Technology Lausanne Institut de G6nie Chimique CH- 1015 Lausanne/Switzerland

Prof. Dr. C. Wandrey

Institute of Biotechnology Forschungszentrum Jtilich GmbH P.O.Box 1913, D - 52428 Jtilich/FRG

Preface

One of natures most important biological processes is the conversion of wood and other lignocellulosis to carbon dioxide, water and humic substances. Biotechnology, by definition, is the technical utilization of biological reactions. Since wood and other lignocellulosics constitute the raw material for the forest industries there should be ample opportunity for biotechnology in manipulating these resources. There are, however, difficulties for implementation o f such technologies in this industry. While biotechnology in areas like medicine and pharmacology concerns production o f expensive products on a small scale, biotechnical utilization and conversions of lignocellulosics means production of inexpensive products on a large scale. Biotechnological utilization of lignocellulosic materials is therefore a very difficult task, and the commercial utilization of this technology has, therefore, only recently gained momentum. One reason for this was the lack of basic knowledge about enzyme mechanisms involved in the degradation and conversion of wood, other lignocellulosics and their individual components. However, the worldwide efforts devoted over the past few decades to research for a better understanding of these mechanisms now provide a solid base for successful development of biotechnology for the pulp and paper industry. For those deeply involved in these investigations, it is obvious that the investments have been a dazzling success. This issue of Advances in Biochemical Engineering/Biotechnology presents, in great detail, recent findings about microorganisms and their enzymes involved in the degradation of wood and wood components, cellulose, the hemicelluloses and lignins. The issue covers not only biotechnology in the pulp and paper industry, but also in forestry. The chapter on Forest Tree Biotechnology demonstrates how trees, unlike such inanimate resources as metallic ores, have the potential to be modified genetically, essentially transforming lead into gold. The new methodologies being used to address problems in forest biotechnology are described with respect to their potential impact on forest tree improvement. A whole chapter is devoted to ,,Lignin", it's role in wood, it's biosynthesis and structure as well as methods for it's analysis. The chapter on the use of white-rot fungi for a specific delignification of wood chips, i.e .... Biomechanical Pulping" describes how this technique is now being run in pilot plant scale and how it approaches commercialization. Pitch problems in pulping processes are addressed with respect to how they can be minimized by the use of enzymes or fungi. The technique for how to do this is described in great depth.

VIII

Preface

No less than three chapters are devoted to pulp bleaching. One is concerned with purification of effluents containing organochlorine compounds using biotechnology. Two other chapters are devoted to the use of hemicellulases for bleaching, one of which describes the basic research in this area, while the other presents results from the use of enzymes in the pulp mills. The last chapter focuses on biotechnology to solve slime problems caused by microorganisms growing in the water systems of pulp and paper mills. It is inevitabte that there will be an increasing need for closing of these systems, thus there will likely be increasing slime problems in these systems in the future. These papers convey the current and developing applications ofbiotechnology in the pulp and paper industry. Wherever possible, the authors have attempted to peer into their crystall balls to see what lies ahead. However, the age of biotechnology is only just dawning, but I have no doubt that surprises that lie yet ahead for us will only serve to further enthuse this field of research. September, 1996

Karl-Erik L. Eriksson

Table of Contents

Forest Tree Biotechnology J. F. D. Dean, P. R. LaFayette, K-E. L. Eriksson, S. A. Merkle. Microorganisms and Enzymes Involved in the Degradation of Plant Fiber Cell Walls R. C. Kuhad, A. Singh, K.-E. L. Eriksson . . . . . . . . . . . . . . . . . . Lignin D.S. Argyropoulos, S.B. Menachem . . . . . . . . . . . . . . . . . . . . . .

1

45

127

Fungal Delignification and Biomechanical Pulping of Wood M. Akhtar, R. A. Blanchette, T. K. Kirk . . . . . . . . . . . . . . . . . 159 Solving Pitch Problems in Pulp and Paper Processes by the Use of Enzymes or Fungi R. L. Farrell, K. Hata, M. B. Wall . . . . . . . . . . . . . . . . . . . . . . . .

197

Reduction of Organochlorine Compounds in Bleach Plant Effluents P. Bajpai, P. K. Bajpai . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

213

Hemicellulases in the Bleaching of Chemical Pulps A. Suurn~ikki, M. Tenkanen, J. Buchert, L. Viikari . . . . . . . . . . .

261

Using Enzymes in Pulp Bleaching: Mill Applications J. S. Tolan, M. Guenette . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

289

Biotechnology for Solving Slime Problems in the Pulp and Paper Industry S. C. Johnsrud . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

311

Author Index Volumes 51 - 57 . . . . . . . . . . . . . . . . . . . . . . . . .

329

Subject Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

333

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Forest Tree Biotechnology Jeffrey F.D. Dean 1, Peter R. LaFayette 2, Karl-Erik L. Eriksson z, and Scott A. Merkle 1 Daniel B. Warnell School of Forest Resources and Department of Biochemistry and Molecular Biology, University of Georgia, Athens, GA-30602, USA

List of S y m b o l s a n d A b b r e v i a t i o n s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 The C h a l l e n g e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ............... 2 Key P r o b l e m s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 W o o d and P a p e r P r o d u c t s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Breeding a n d Life-Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 M a n - M a d e a n d E n v i r o n m e n t a l . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Propagation ............................................. 3.1 Cell a n d Tissue C u l t u r e of F o r e s t Trees . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 In V i t r o P r o p a g a t i o n T e c h n i q u e s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 A x i l l a r y S h o o t M u l t i p l i c a t i o n . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.40rganogenesis ......................................... 3.5 S o m a t i c E m b r y o g e n e s i s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6 P r o t o p l a s t C u l t u r e . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.7 In Vitro Screening a n d S o m a c l o n a l V a r i a t i o n . . . . . . . . . . . . . . . . . . . . . . . 3.8 C r y o p r e s e r v a t i o n . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.9 Artificial Seeds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Genetic Engineering ........................................ 4.1 Tissue C u l t u r e C o n s i d e r a t i o n s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Biological G e n e Transfer ( A g r o b a c t e r i a - M e d i a t e d T r a n s f o r m a t i o n ) . . . . . . . . . . 4.3 Physical G e n e Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3.1 T r a n s f o r m a t i o n by Microprojecti,~e B o m b a r d m e n t . . . . . . . . . . . . . . . . 4.3.2 T r a n s f o r m a t i o n of P r o t o p l a s t s . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3.3 A l t e r n a t i v e T r a n s f o r m a t i o n T e c h n i q u e s . . . . . . . . . . . . . . . . . . . . . . . 4.4 Selection Systems, P r o m o t e r s , a n d R e g u l a t o r y E l e m e n t s . . . . . . . . . . . . . . . . 4.5 C u r r e n t a n d F u t u r e T a r g e t s for G e n e t i c E n g i n e e r i n g in F o r e s t Trees . . . . . . . . . 4.5.1 L i g n i n C o n t e n t a n d C o m p o s i t i o n . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.2 Sterility a n d E a r l y F l o w e r i n g . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.3 H e r b i c i d e Resistance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.4 Insect a n d P a t h o g e n Resistance . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.5 Tree F o r m a n d F i b e r M o r p h o l o g y ......................... 4.5.6 N o v e l T r a i t s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 R e g u l a t o r y C o n s i d e r a t i o n s . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 M o l e c u l a r Breeding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 D N A M a r k e r T e c h n i q u e s .................................. 5.1.1 R e s t r i c t i o n F r a g m e n t L e n g t h P o l y m o r p h i s m ( R F L P ) .............. 5.1.2 R a n d o m Amplified P o l y m o r p h i c D N A ( R A P D ) . . . . . . . . . . . . . . . . . . 5.1.3 B u l k e d Segregant Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1.4 M i c r o s a t e l l i t e R e p e a t P o l y m o r p h i s m s . . . . . . . . . . . . . . . . . . . . . . . . 5.1.5 Amplified F r a g m e n t L e n g t h P o l y m o r p h i s m s ( A F L P ) . . . . . . . . . . . . . . . 5.1.6 Microsatellite H y b r i d i z a t i o n . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 A p p l i c a t i o n s of D N A M a r k e r s in F o r e s t R e s e a r c h . . . . . . . . . . . . . . . . . . . . 5.2.1 G e n e t i c L i n k a g e M a p s ................................ 5.2.2 M a p p i n g Projects ................................... 5.2.3 M a r k e r - A s s i s t e d Selection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

3 4 4 4 6 7 7 7 8 9 10 11 13 14 14 14 15 16 17 20 20 21 21 22 23 23 24 25 25 26 26 26 27 28 28 28 29 29 30 30 31 31 32 32

Advances in Biochemical Engineering/ Biotechnology, Vol. 57 Managing Editor: T. Scheper 9 Springer-Verlag Berlin Heidelberg 1997

2 5.3 Other Applications of D N A Markers . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3.1 Quantification of Genetic Diversity . . . . . . . . . . . . . . . . . . . . . . . . . 5.3.2 Genotype Verification and Delineation . . . . . . . . . . . . . . . . . . . . . . . 6 Electronic and Computational Resources . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Jeffrey F.D. Dean et al. 34 34 34 35 36 37

The forest products industry has traditionally viewed trees as merely a raw, and more or less immutable, natural resource. However, unlike such inanimate resources as metallic ores, trees have the potential to be modified genetically, essentially transmuting lead into gold. Increasingly, m o d e r n alchemists are applying the tools of biotechnology in efforts to reduce the biological constraints on forest productivity. Several new methodologies being used to address problems in forest biology are described with respect to their potential impact on forest tree improvement. In addition to addressing problems inherent to the current use of trees for production of pulp and paper or solid wood products, genetic manipulation of trees brings with it the potential to create new industries based on the novel characteristics of transgenic trees, e.g. trees containing transgenes to detoxify specific pollutants could be used in the remediation of sites contaminated with hazardous wastes. Efforts to modify trees through biotechnology are in their infancy, and this review seeks to outline the underpinnings of what will undoubtedly be an area of increased emphasis in the next millennium.

Forest TreeBiotechnology

List of Symbols and Abbreviations Symbol 2,4-D AFLP BAP Bt CAD CaMV CAT EPSP F5H GFP GUS IAA NAA OMT PCR PEG QTL RAPD RFLP STFI vir

Description 2,4-dichlorophenoxyacetic acid amplified fragment length polymorphism benzylaminopurine Bacillus thuringiensis endotoxin cinnamyl alcohol dehydrogenase cauliflower mosaic virus chloramphenicol acetyltransferase 5-enolpyruvylshikimate-3-phosphate ferulate-5-hydroxylase green fluorescent protein [3-glucuronidase indoleacetic acid naphthylacetic acid o-methyltransferase polymerase chain reaction polyethylene glycol quantitative trait locus random amplified polymorphic DNA restriction fragment length polymorphism Swedish Pulp and Paper Research Institute a class of genes controlling the virulence of Agrobacterium tumefaciens

4

Jeffrey F.D. Dean et al.

1 The Challenge Since the advent of agriculture, one of the principal challenges for mankind has been to increase the ability of plants to capture incoming solar energy for the production of food, fuel, and fibers. As the human population continues to burgeon, it becomes clear that the demand for increased production efficiency in crops and trees will not diminish. While the most significant advances of the "green revolution" have resulted in a doubling and tripling of yields from agronomic crops, the potential for increasing forest productivity, for instance through reduced rotation times, could be much greater given the minuscule effort to date in applying the techniques of modern biotechnology to potentials in forest tree biology. The increasing use of plant polysaccharides for the production of fuel-grade ethanol is one example suggesting that the future products derived from forest trees may not remain just paper and timber. Thus, new products derived from wood also have the potential to drive demand for greater forest productivity. However, the greatest pressure for increased tree productivity is likely to result from preservation efforts that remove forests on public lands from the role of harvestable inventory. Regardless of corporate sentiment, current trends in this area are unlikely to be reversed, and, as a consequence, plantation forestry is likely to become a major production paradigm for the next century. Tree productivity will need to be increased significantly if we are to have any chance at meeting projected fiber demands using the diminished acreage allotted to production forestry. Forestry has never faced challenges as great as those before it now. In some parts of the world misguided land use policies have led to extensive deforestation, while, elsewhere, productivity has stagnated due to the detrimental effects of environmental pollutants such as acid rain. Population growth in every corner of the globe increases pressure to produce more fodder and fuel wood. In the past, answers to these complex issues usually centered on social, political, or economic redress, and such is likely to remain the case for the near future. However, plant biotechnology has advanced to a point where we can now envisage biological solutions to some of the more intractable forestry problems. This chapter is intended to convey the optimism that currently pervades this field by highlighting a few of the most exciting areas in which biotechnology is being applied to problems of forest biology.

2 Key Problems 2.1 Wood and Paper Products The term "wood quality" is often used to describe the overall suitability of a particular wood source for a specific end use. Thus, wood considered to be of

Forest Tree Biotechnology

5

excellent quality for one type of product may be useless for another application and vice versa. For example, spruce fibers generally tend to have thinner walls than those from pine, and, as a result, spruce pulps form more compact and denser sheets of paper. Similarly, softwood fibers, which are always longer than hardwood fibers, increase the tear strength index of paper, but the tight packing of short hardwood fibers yields a smoother, more uniform writing surface. Solid wood products are dependent on a completely different set of parameters to define wood quality. Zobel and Van Buijtenen [1] have reviewed the ways in which variations in wood properties influence the quality of wood and paper products, while Karenlampi [-2] has discussed the role of fiber properties in product quality. Wood quality is a function of the morphology and chemistry of all the various cell types and the ratio of their occurrence in a particular species of tree. Thus, wood quality is constrained by the genetics, age, environmental conditions, and cultivation practices imposed on the individual tree. G r o w t h conditions can significantly alter such cellular parameters as fiber volume and morphology as well as cell wall architecture and chemistry. Cell walls are actually a composite material, and their ultrastructure thus derives from the ways in which the various molecular components are arranged and oriented to form the fiber wall. In general, embedded cellulose microfibrils give the walls tensile strength, while the surrounding matrix of hemicelluloses and lignin bind and rigidities the composite. Lignin also serves to hold the individual fiber cells together. The carbohydrate components, i.e. cellulose and hemicelluloses, are hydrophilic materials, while lignin is hydrophobic, and the relative proportions of these two classes of material in a given fiber wall define many of the bulk properties of the fiber. Structural features of importance for paper production include fiber length, diameter, wall thickness and ultrastructure. Softwood fibers used in pulp production can have a mean length anywhere from 2.5 mm to more than 10 mm; however, the majority of softwoods have an average fiber length of 3-5 mm. Most softwood pulp fibers have a diameter of less than 0.1 ram. Hardwood fibers are, on the average, about 1/3 the length and about 1/2 the width of softwood fibers. It seems likely that the fibers used for paper manufacturing in the future will increasingly come from plantations of fast-growing trees where juvenile fibers will predominate [3]. It is relatively easy to predict the effect this will have on products made with hardwood fibers since the morphology and chemistry of juvenile fibers of hardwoods are more similar to mature fibers than is the case for softwood fibers [4, 5]. However, juvenile softwood fibers are significantly shorter and weaker than mature fibers, and, as a consequence, they do not have the superior strength characteristics needed for making strong paper using current technology. Thus, the long fibers currently obtained from mature temperate softwoods may well become a limiting factor for future paper production. There is currently a fundamental lack of knowledge about the relationship between fiber quality and pulp product properties. One reason for this is that the

6

JeffreyF.D. Dean et al.

parameters which are used to characterize the fibers at different steps in the processing chain are non-uniform. Foresters describe wood quality with respect to tree growth and density, pulp makers focus on measurements of viscosity and kappa number, while papermakers test freeness and handsheet properties. All of these players are looking at the bulk properties of fiber mixtures, but no one has studied purified, uniform fiber preparations in sufficient detail to explain how individual fiber types might influence the bulk properties of the mixture. An understanding of individual fiber properties should make it possible to model how different combinations of fiber mixtures might improve paper processing. To collect such information on fiber mixtures, the Swedish Pulp and Paper Research Institute (STFI) has developed a measurement system called STFI Fiber Master. This equipment can rapidly determine fiber form, length, width, and flexibility for more than 1000 particles per second in a diluted fiber suspension. When combined with information on the chemistry and binding parameters for the fibers, this new analytical tool provides a powerful technique for optimizing fiber mixtures to suit the production of a particular product.

2.2 Breeding and Life-Cycle Although they have been exploited by man for millennia, forest trees constitute one of the last major plant groups that has not been subjected to significant genetic manipulation with an eye toward improvement. The reason for the lack of progress with forest trees compared to agronomic and horticultural species can be attributed to the fact that these plants are characterized by a number of unique biological features which have made their breeding a slow and difficult process. The most obvious of these is the relatively large size of tress of reproductive age, which makes controlled breeding in the greenhouse difficult. However, two facets of their long life histories currently present major barriers to rapid improvement of trees via traditional breeding: (1) the long lag period between seed germination and flower production, and (2) the lengthy interval between seedling and mature phenotype. Average time to flower production varies greatly among species, from as short as 5 years for some species of Pinus to as long as 25 years for some species of Quercus [6] Consequently, the tree breeder may have to wait more than two decades between breeding cycles. Furthermore, the major traits for which improvement is desired, such as volume, wood specific gravity, and form or growth habit, usually cannot be assessed until the tree assumes its mature phenotype, perhaps 20-30 years from the seedling stage. Attempts to predict mature tree performance from that of seedlings has so far met with limited success [e.g. 7, 8]. To make matters worse, many of these desirable traits are not inherited in a simple fashion, but instead result from complex interactions of genes at multiple loci. As a consequence, improvement of quantitative traits requires breeding programs designed to make incremental advances in a population mean for a desired trait using such tools as mass selection. While this approach has led to measurable improvement in some


7

commercial tree species such as Pinus taeda [9], these gains have required massive cooperative efforts over multiple decades. For the majority of trees, traditional breeding approaches are simply not a realistic means for achieving genetic improvement.

2.3 Man-Made and Environmental In addition to the natural constraints that limit forest tree productivity, man has added new problems as a consequence of population growth and industrialization. One of these problems is pollution of the soil, air and water. While some forest trees seem able to tolerate high levels of air pollution, surviving well, for example, in urban environments, others are severely damaged by high levels of pollutants such as ozone. Unless we are satisfied with being limited to a few species of trees for growth in polluted areas, trees that currently cannot survive airborne toxins will have to be engineered to tolerate certain levels of these pollutants. Population growth in some countries has led to increasing competition for land. For example, in the south eastern United States, old agricultural fields were once plentiful enough to be employed for pine plantation forestry. Today, with the growth of cities and suburbs, these lands are no longer available for agriculture, let alone forestry. Thus, forests will probably have to be planted on lands that are less than optimal with regard to fertility and water availability. In these cases, we will need trees that are specifically adapted for growth on sites where trees available today would be unable to survive. An extreme case might be the need to produce populations of trees adapted to growing on sites which are unsuitable for any other use, such as those contaminated by heavy metal residues.

3 Propagation 3.1 Cell and Tissue Culture o f Forest Trees The goal most often cited for in vitro culture of forest trees is propagation. As with conventional vegetative propagation (macropropagation) techniques such as rooted cuttings, the primary objective of in vitro propagation (micropropagation) is to capture the total genetic superiority of the parent material, including both additive and nonadditive genetic components. In addition, clonal propagation systems allow the application of a very high selection differential, since whole new populations of plants can be cloned from just a few elite individuals. Theoretically, since somatic tissues are used as the starting material, each tree regenerated from these tissues should be an exact clonal copy of the ortet. The ideal result should therefore be a highly uniform clonal population that

8


replicates the phenotype of the source tree. However, unlike macropropagation systems, some in vitro propagation methods have the potential to introduce significant new genetic or epigenetic changes into propagules regenerated from them, so that the parent material is not truly replicated (see section on somaclonal variation). Furthermore, the nature of much in vitro propagation technology for forest species excludes clonal propagation of proven genotypes, since many of these systems are limited to employing explants from juvenile genetically unproven material. A second principal goal of micropropagation that parallels conventional vegetative propagation is simply to mass propagate plants, bypassing sexual reproduction, which for a particular species may be too expensive, inefficient or impractical. In vitro propagation can also be used to overcome problems associated with macropropagation, including limitations on available parent material or space. In addition, just as propagation through seeds may be limited by biological factors, certain species or even individual genotypes may be recalcitrant to conventional vegetative propagation techniques such as rooting of stem cuttings. In these cases, in vitro techniques may succeed where macropropagation fails. In addition to the advantages shared with conventional vegetative propagation, in vitro culture has a number of associated potential and real applications that are unique. These include protoplast culture and fusion for generation of somatic hybrids, in vitro screening, generation of useful somaclonal variants, generation of artificial seeds, and long-term storage of germplasm using cryopreservation. Finally, in vitro culture currently provides the only route for generation of genetically engineered genotypes of forest trees. Gene transfer technologies including Aorobacterium-Ti plasmid mediated gene transfer, electroporation, microinjection, and microprojectile bombardment all depend on the ability to culture cells in vitro in order to select the transformed cells, usually by employing drug-resistance marker genes. Once transformed cells are obtained, these can be cultured to regenerate transformed plantlets (see section 4.2 on gene transfer).

3.2 In Vitro P r o p a g a t i o n T e c h n i q u e s

Micropropagation systems fall into three broad categories: axillary shoot (or bud) multiplication, organogenesis, and somatic embryogenesis. Axillary shoot methods rely on multiplication of preformed structures, while organogenesis and somatic embryogenesis rely on de novo generation of either plant organs or embryos, respectively (i.e. morphogenesis). All these regeneration systems have great potential to be applied for mass propagation of forest tree species. As the examples below show, the success of a given method appears to be highly species-specific, with shoot multiplication or organogenic systems working well for some species, while embryogenic systems are clearly superior for others.


9

Forest trees were among the first plants cultured in vitro. Gautheret [10] reported formation of adventitious buds in cultured cambial explants of Ulmus campestris, demonstrating that an exogenous source of sugar was required for bud production, while high levels of indoleacetic acid (IAA) in the medium inhibited bud formation. Jacquiot [11] extended the research with this species, using trees up to 180 years old as tissue sources. Mathes [12] reported in vitro production of both roots and shoots in callus cultures of Populus tremuloides, although apparently no plantlets were produced. It was not until four years later that Wolter [13] produced entire P. tremuloides plantlets in vitro by inducing shoot formation with benzylaminopurine (BAP) and subsequently rooting the shoots in vitro. Winton [14], working with triploid P. tremuloides callus, also obtained complete plantlets, demonstrating for the first time in vitro propagation of a forest tree with a superior genotype. Among coniferous species, adventitious shoots were first produced in callus cultures of Sequoia sempervirens [15]. However, it was not until 25 years later that complete plantlets of a conifer, Pinus palustris, were regenerated in vitro [16]. Since the pioneering work with Populus and P. palustris, hundreds of tree species have been propagated in vitro, although most of this work has been done on an experimental rather than an operational scale. It would be unrealistic to attempt to review here all of the in vitro propagation research with forest trees reported to date. The most recent complete review of forest tree micropropagation is that of Thorpe et al. [17]. In addition, a number of books have been published in the past 10 years containing chapters describing in vitro propagation of many of the important woody angiosperm and gymnosperm genera. We would refer those desiring information on manipulation of individual woody plant genera in vitro to books edited by Bajaj [18-20] or Bonga and Durzan [21]. Here, we will briefly review the basics of each of the three systems along with their advantages and disadvantages, citing examples of some of the more advanced systems developed for commercially important forest species.

3.3 A x i l l a r y S h o o t M u l t i p l i c a t i o n

Production of plantlets from axillary shoots is similar to vegetative propagation via rooted cuttings in that shoots are excised from source tissue and rooted individually to multiply the original genotype. The differences are that, with axillary shoot systems, the procedures are performed in vitro under aseptic conditions, and subsequent manipulations in vitro can be used to greatly multiply the number of propagules obtained. These manipulations may include treatment with a cytokinin such as BAP or zeatin, or culture conditions which induce axillary buds in the cultured tissues to elongate into shoots. Such shoots can be excised and either rooted in vitro or placed back into culture for further enhancement of axillary branching. Thus, unlike rooted cutting macropropagation, treatments can be applied in vitro which promote multiplication of new shoot material ad infinitum.

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One of the principal advantages of axillary shoot methods for forest tree multiplication is that all propagules are derived from preformed buds, thereby enhancing the likelihood that propagules will be true to type. Because shoots arise from meristems present in the explant, there is little chance of introducing somaclonal variation into the propagules. The greatest advantage of axillary shoot multiplication systems, however, is the fact that this method has to date proven to be the most effective technique for multiplying mature selected trees [17]. Some examples of mature hardwood forest species propagated using this method are Robinia pseudoacacia [22], Liquidambar styraciflua [23], Eucalyptus citriodora [24] and Acer saccharinum [25, 26]. While axillary shoot multiplication systems have been the primary in vitro propagation method applied to hardwoods, the successful application of this technique to conifers has been less frequently reported. In pines, this appears to be due to difficulty in stimulating elongation of the axillary buds found at the bases of needle fascicles [27]. An exception to this occurs with Pinus radiata, for which workers in New Zealand have developed an axillary shoot multiplication system which has enabled them to plant thousands of clonal trees in the field since the mid-1980s [28]. Given that production of plants from axillary shoots has some similarities to propagation from rooted cuttings, it might be expected that these techniques would share some common disadvantages. One such disadvantage is a relatively low frequency of propagule production. Although repeated cycles of in vitro culture can ultimately produce thousands of propagules, there is often a substantial lag phase before operational numbers of plants can be produced regularly. Axillary shoot multiplication methods are also relatively labor intensive, requiring significant amounts of handling, for both cycling of cultures and production of plantlets. Plantlet production can require multiple steps, including induction, shoot elongation, shoot excision, rooting, and acclimatization of plantlets, each of which may require significant inputs of labor. Therefore, although axillary shoot multiplication remains the major route for the regeneration of woody angiosperms in vitro, its labor-intensive nature has limited its application mainly to woody ornamentals which have relatively high single-tree values compared to forest species. McCown and McCown [29], who described axillary shoot multiplication systems for Quercus, Amelanchier, Ulmus, Betula, and Populus, argued that tree microculture must become more automated to reduce labor costs before this technique can be economically applied to forest species. However, because of multiple handling steps, axillary shoot multiplication methods may not be as amenable to scale-up or automation as other methods for forest tree propagation, although advances in robotics may change this potential [30].

3.40rganogenesis Organogenesis is the de novo production of plant organs (buds, shoots, and roots) from organized tissues or callus. As with axillary shoot multiplication


11

systems, organogenic cultures are usually stimulated to produce buds by addition of a cytokinin to the medium. In many cases, initial explants are induced to form callus and subsequently buds. Then, as with axillary shoot methods, the buds are elongated into shoots, excised, and rooted individually. The primary advantage of organogenic regeneration methods over axillary shoot methods is a potential for higher frequency plantlet production in a shorter period of time. Callus can be grown in large quantities with less demand for labor and space, and adventitious shoot production can be used to achieve high multiplication rates. However, like axillary shoot methods, organogenesis-based methods may require the labor-intensive steps of shoot elongation, excision, rooting, and plantlet acclimatization to produce field-plantable stock. In addition, unlike axillary shoot methods, organogenic methods, especially those requiring an intermediate callus, may be associated with the production of significant amounts of somaclonal variation in the regenerated plantlets. Thus, the resulting plantlets from a given clone may display unacceptable phenotypic variation. Lester and Berbee [31], for example, observed a wide range of variation in plantlets derived from callus cultures of Populus. Organogenic regeneration systems have been developed for a number of coniferous genera, including Pinus [e.g. 16], Picea [e.g. 32], Pseudorsuga [33] and Abies [e.g. 34]. In most cases, adventitious buds were induced to form directly from tissues such as seedling cotyledons. Among conifers, only Pinus eldarica has produced long-term callus cultures that could be induced to form adventitious buds from which plants were successfully regenerated [35]. In a few cases, plantlets produced from adventitious conifer buds have been tested in the field. In the cases of Pinus taeda and Pseudotsuga menziesii, plantlets derived from adventitious buds survived well, but lagged behind seedlings in height growth [36, 37]. However, over the past 10 years the emphasis of conifer micropropagation research has shifted to somatic embryogenesis, since it is believed by many working in the area that this technique offers the best potential for scale-up to commercially viable levels. As of 1987, McCown and McCown [29] were unable to cite a single case of commercial application of a micropropagation system based on adventitious bud generation for a North American hardwood. Although to our knowledge this situation remains unchanged, a number of promising adventitious bud systems have been reported for hardwoods in the past few years. Among these, high-frequency adventitious bud regeneration systems have been reported for Liquidambar styraciflua [-38~41], several Populus species and hybrids [e.g. 42-45], Ulmus spp. [46], and Robinia pseudoacacia E47-50].

3.5 Somatic Embryogenesis Somatic embryogenesis is the de novo production of structures resembling zygotic embryos, either from organized tissues or from callus. Structures

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classified as somatic embryos must be bipolar (possessing both root and shoot poles) and have no vascular connection to the source tissue. Somatic embryos may be derived either through direct or indirect embryogenesis. In direct embryogenesis, embryos are formed essentially by multiplication of a zygotic embryo explant, i.e. by embryo cloning [51]. Indirect embryogenic systems involve a dedifferentiation of non-embryonic tissue to form a callus from which somatic embryos arise [51]. As with most other plant species, somatic embryogenesis in forest trees is usually induced by exposure to an auxin such as NAA or 2,4-D, although there are a few cases of embryogenesis being induced by cytokinins [e.g. 52, 53] or no exogenous growth regulators at all [e.g. 54]. Since continuous exposure to auxin usually results in repetitive cycles of embryo production, growth regulators are usually removed from the medium following induction to allow the somatic embryos to complete development. In some cases, only a few weeks or even days of exposure to auxin is required to induce repetitive embryogenesis [e.g. 55, 56]. The first report of somatic embryogenesis in a hardwood forest species was for Santalum album [57]. Since that time, somatic embryogenesis has been reported in hundreds of angiosperm trees (see reviews by Tulecke [-58] and Warm [59]). However, it was not until 1985 that somatic embryogenesis was reported in a coniferous species, Picea abies [60]. In the last decade, somatic embryogenesis has been reported for most commercially important conifers, including Pinus [e.g. 61-63], Abies [52], Larix [e.g. 64] and Pseudotsuga [65]. For a full summary of somatic embryogenesis in coniferous trees, the reader is referred to reviews by Attree and Fowke [66] and Tautorus et al. [67]. The most advanced systems with regard to high-frequency plantlet production have been developed for Picea. Canadian researchers working with Picea 91auca have used abscisic acid, osmotica and desiccation treatments to obtain high levels of vigorous plant production from somatic embryos [68]. Workers at BC Research planted the first large-scale field tests of somatic embryo-derived conifers from cultures of interior spruce (Picea 91auca engelmannii complex) [69]. Somatic embryogenesis has been cited by many authors as the in vitro regeneration system of choice for economical production of clonal populations of forest trees [e.g. 70]. Certainly, this type of system has a number of powerful advantages over axillary shoot multiplication methods and organogenesis. A major advantage is the potential for very high frequency regeneration. Depending on the species, virtually unlimited numbers of embryos can be generated from a single explant. In addition, embryogenic cultures of many species can be grown in liquid, allowing production and handling of thousands of embryos at one time. Thus, in comparison to axillary shoots and organogenesis, somatic embryogenesis offers the potential for high-volume large-scale propagation which can translate into significant labor savings. Greater economies of scale may be possible if bioreactor and continuous culture technologies can be applied to embryogenic systems [e.g. 71,72]. The feature of somatic embryogenesis which may ultimately have the largest impact for mass propaga-


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tion of forest trees is the fact that the product is an embryo. The morphological and physiological similarity of somatic embryos to zygotic embryos means that they are complete propagules in themselves, with embryonic roots, shoots and leaves, and, most importantly, the "program" to make a complete plant. As a consequence, no separate shoot elongation or rooting steps are required for plantlet production. This characteristic further lowers labor inputs and gives somatic embryos the potential for direct delivery to the greenhouse or field (see section 3.9 on artificial seeds), thereby eliminating the need for labor-intensive transplanting. One drawback of current forest tree embryogenic systems is the low frequency of plantlet production or "conversion" of somatic embryos to plantlets. Although numerous systems have been reported, and some of these produce embryos at high frequencies, a major bottleneck has been induction of embryo maturation and subsequent production of field-plantable stock. Currently, only a handful of tree species such as Liriodendron tulipifera and some Picea species can be propagated via somatic embryogenesis in sufficient numbers to enable establishment of useful field tests [69, 733. Another major disadvantage of embryogenic propagation methods for forest species is that the bulk of the systems reported to date rely on immature tissues (i.e. from seeds or seedlings) as explanting material. Thus, the material being propagated is of unproven genetic value. Most reports of somatic embryogenesis in tree species are in reality reports of "embryo cloning," in which the zygotic embryo is induced to replicate itself indefinitely. However, in the past few years, a few reports have appeared in which embryogenic cultures were initiated from mature tissues. For instance, embryogenesis cultures have been initiated from the male flower parts of Quercus [743 and Aesculus [75], while Michler and Bauer [76] used leaf tissues of a Populus hybrid of known genetic value to obtain somatic embryos.

3.6 Protoplast Culture Another potential application of in vitro culture which may eventually succeed for forest tree improvement is generation of somatic hybrids via protoplast fusion. For example, protoplasts of Citrus species from different genera which were unable to hybridize sexually were fused, and somatic hybrid plantlets were regenerated [77]. Although regeneration of somatic hybrids has not been reported to date for a forest tree, protoplasts have been isolated from a number of forest species and induced to regenerate cell walls and eventually divide to produce protoplast-derived plantlets. Angiosperms regenerated from protoplasts include Populus spp. [78], Ulmus spp. [79] and Liriodendron tulipifera [80]. Among gymnosperms, embryogenic cultures have provided a reliable source of totipotent protoplasts, and many conifers have been regenerated from protoplasts derived from embryogenic cultures. These include Pinus taeda [62], Pinus caribaea [81], Picea glauca [82], Abies alba [83], and Larix species [84, 85],

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3.7 In Vitro Screening and Somaclonal Variation As mentioned earlier, organogenic and embryogenic regeneration systems may be characterized by significant levels of somaclonal variation. This phenomenon may be regarded as a nuisance if the goal is production of truly clonal propagules. However, somaclonal variation may also be applied as a tool for generating useful variants. Although not frequently reported in forest trees, screening of genotypes for such traits as disease resistance can be accomplished in vitro, and this can be followed by production of propagules from the selected material. This technique has been applied to the selection of some coniferous trees [86]. It has also been used to generate hybrid Populus variants that were resistant to the herbicide glyphosate [87] as well as eastern cottonwood (Populus deltoides) variants that displayed increased or decreased resistance to leaf rust caused by Metampsora medusae [88].

3.8 Cryopreservation Another technique that is likely to come into widespread use in operational propagation of forest trees is long-term cryopreservafion of culture tissues. The ability to store germplasm for long periods will become critical if in vitro propagation methods are to be used commercially. This is because most of the forest tree embryogenic systems reported to date depend on materials of unknown genetic value. Consequently, trees derived from these cultures will need to be tested for field performance prior to their release into production programs. Even if researchers are eventually able to clone mature tissues from elite genotypes of particular forest species, there will still be a need for field evaluation of trees derived from these cultures. In addition, under continuous culture, the useful life of most embryogenic cultures does not exceed a few years. Therefore, embryogenic cultures must be held in a suspended state while trees derived from them are tested. Once field testing is complete, cultures of those clones showing the best field performance can be scaled up for production of propagules. Fortunately, embryogenic cultures of both gymnosperms and angiosperms seem to be relatively amenable to cryopreservation. This technique has already been applied to the long-term storage of embryogenic cultures of Betula [89], Picea glauca [90], Pinus caribaea [91] and Abies nordmanniana [92].

3.9 Artificial Seeds One of the advantages of somatic embryogenesis over the other in vitro propagation systems noted above is the potential of somatic embryos to be directly delivered to the greenhouse or field as synthetic seeds. Efforts have concentrated on preparing somatic embryos to emulate seeds with regard to such characteristics as desiccation tolerance, resistance to mechanical damage and ability to


15

sustain ex vitro post-germinative growth. These features would facilitate mechanical handling and automated planting of somatic embryos. A number of encapsulation techniques have been employed, including the use of such materials as hydrated gels and water-soluble resins [93]. While development of artificial seed technology has mainly focused on species of agronomic importance, such as alfalfa, the same technology has been tested on somatic embryos of a few forest trees. Artificial seeds have been generated by alginate gel encapsulation of somatic embryos of Pinus taeda [61, 62], Picea species [94], Santalum album [95] and Robinia pseudoacacia [96]. Although plantlets have been regenerated from some of these artificial seeds, none of these systems has been developed to the point where the somatic embryos could be employed as if they were true seeds.

4 Genetic Engineering As illustrated by the wide variety of specialized cultivars available within agronomic and horticultural crop species, traditional plant breeding based on controlled sexual crosses has the potential to tailor the traits of virtually any plant to be more suitable for its end use. However, this process generally requires selection and back-crossing through nine or more generations just to introduce a single trait into a plant line and, thus, the life-cycle characteristics of long-lived and slow-maturing forest trees make such breeding efforts difficult at best. Additionally, such plant improvement schemes are limited to modification of traits already available within the species or within closely-related hybridizing species. Fortunately, recent advances in biochemistry and molecular biology have provided techniques for the direct transfer of foreign genes to plant tissues, thereby circumventing some of the most troublesome bottlenecks inherent to traditional plant improvement strategies. Only slightly more than a decade ago, researchers in several laboratories accomplished the first controlled introductions of foreign genes into plants [97-100]. This process, commonly referred to as transformation, has been used to introduce into plants a variety of foreign genes whose expression confers novel traits to the transformants [101, 102]. A wide variety of genetic constructs designed to modulate the expression of existing plant genes in such a way as to enhance the properties of the desired plant products have also been tested, and the product of one such transformation, the Flavr-SavrTM tomato, recently became the first genetically engineered plant product approved for human consumption [103]. Most plant transformation work has so far focused on agricultural crops [104], but efforts to produce genetically engineered trees are on the increase and several reviews of the work to date are available [105-111]. The treatise by Haines [112] is a particularly useful guide to the opportunities presented by biotechnology with respect to tree improvement. As discussed in greater detail in the following sections, the minimum requirements for successful gene transfer into forest trees are: (1) a tissue culture system

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Table 1. Tree species that have been successfully transformed and regenerated.

Species

Method

Reference

Agrobacterium Agrobacterium Microprojection Agrobacterium Agrobacterium Protoplast/Chemical Electroporation Agrobacterium Agrobacterium Microprojection Microprojection Agrobacterium Agrobacterium Microprojection Agrobacterium Agrobacterium Agrobacterium

[114] [115] [116] [117] [118] [1 I9] [120] [121] [122, 123] [ 124] [125] [126] [127] [128] [129] [130] [49]

Agrobacterium Microprojection

[131] [ 132]

Angiosperms Allocasuarina verticillata Azadirachta indica Carica papaya Carya illinoensis Citrus jambhiri Eucalyptus sp. Juglans regia Liquidambar styraciflua Liriodendron tulipifera Malus Pumila Populus sp. Prunus domestica Prunus persica Robinia pseudoacacia

Cymnosperms Larix decidua Picea glauca

allowing for the regeneration of intact plants (see the preceding section); (2) a delivery system for stable introduction of genes into the cultured cells; and (3) a selection system for recovering those cells that have received the introduced genes. It then remains for the researcher to identify the appropriate foreign gene to be introduced or endogenous gene requiring altered expression, while of final concern are the promoter and regulatory elements that will serve to express the introduced gene construct in the correct tissues at the proper developmental stage. Table 1 lists a variety of horticultural and forest trees that have been successfully transformed and regenerated, although in most cases the inserted genes served only as markers of transformation and not as sources of new traits. It should be noted that, owing to the ease with which they may be transformed and regenerated, members of the genus Populus have been the predominant model system to date for demonstrating new transformation techniques for trees [e.g. l 13]. Thus, numerous reports of transformation and regeneration of species and hybrids in this genus appear in the literature, but no attempt has been made to present an exhaustive list of that work here.

4.1 Tissue Culture Considerations All of the considerations previously discussed for the regeneration of intact plantlets from in vitro tissue culture are important when considering a system for


17

gene transfer work. However, the two most important considerations are that the culturing system should be one in which the new plantlet can be derived from a single cell (i.e. organogenic, embryogenic or protoplast culture), and that the frequency with which plantlets can be generated from the system should be as high as possible. Regeneration from single cells ensures that all cells in the resulting plantlet will carry the introduced transgene at the same position in the genome, thereby standardizing its expression and inheritance in subsequent progeny. A high frequency of regeneration is required, particularly when using physical gene transfer techniques (see below), because as few as 1:1000 cells receiving the engineered gene will insert it into their genetic material in such a way that it will be stably retained and expressed in subsequent generations of cells. Unfortunately, tissue culture systems that are highly regenerable are not always highly transformable [133, 134], and even when transgenes are stably inserted into the recipient genome, a variety of positional and cell state effects may lead to a loss of expression in regenerated plantlets [135]. Although tissue culture systems meeting all these requirements are still relatively rare for forest tree species, the increasing number of successes noted in the foregoing section suggest that the availability of competent systems will not be a limiting factor for long.

4.2 Biological Gene Transfer (Agrobacteria-Mediated Transformation) The techniques by which exogenous DNA is transferred into plant cells may be classified as either physical or biological, depending upon whether the DNA is "naked" or passed along by an intermediary organism. Transformation systems based on plant viruses were at one time thought to hold great potential for genetic engineering and some effort has continued in this area [136, 137], but results for the most part have been disappointing [101]. In contrast, the best characterized and most accessible plant transformation systems available today are biological and use domesticated varieties of the plant pathogens, Agrobacterium tumefaciens or A. rhizogenes, to mediate transfer of DNA to the host tissues. Manuals describing comprehensive protocols for performing such work are available [138, 139], and the book edited by Croy [140] provides an invaluable resource enumerating a vast number of vectors, marker genes, regulatory elements and other tools for gene transfer into plants. In their pathogenic forms, A. tumefaciensand A. rhizogenes cause crown-gall and hairy-root diseases, respectively [141]. The diseases appear primarily in woody and herbaceous dicots, and actually represent unusual parasitic interactions that are initiated when the bacteria colonize wounded tissues. The bacteria locate these wounds through a chemotactic response to phenolic compounds (e.g. acetosyringone or sinapic acid) released into the soil from the wound [142]. Once inside the wound, the bacteria use a specialized process resembling bacterial

18


conjugation to transfer into the plant cells DNA copied from an extra-chromosomal element referred to as the Ti-plasmid [143]. A pair of 25-base-pair direct repeats on the Ti-plasmid, the so-called T-DNA borders, facilitate transfer and integration of any genes lying between the borders into the host plant genome where they can subsequently be replicated and expressed. Only that DNA lying between the T-DNA borders is copied and transferred as single stranded DNA to the host plant cells. Until recently, this process was considered the only example of inter-kingdom DNA transfer. However, a related system appears to be capable of transferring DNA between bacteria and yeast [144]. In the wild-type disease-state interaction, some of the genes carried between the T-DNA borders encode enzymes that alter phytohormone (auxin or cytokinin) levels in the transformed tissues, and thereby cause tissue deformation (i.e. galls or hairy roots). Another set of genes transferred from the wild-type Ti-plasmid catalyze the production of opines, a class of amino acid derivatives unique to Agrobacterium infections, which are utilized by the bacteria as a source of both carbon and nitrogen [145]. Several other (vir) genes Iie in regions of the Ti-plasmid outside the T-DNA borders, and are expressed only in the bacterium. The vir gene products control such aspects of the interaction as conjugative transfer of T-DNA into the plant cell, integration of the DNA into the host genome, and the range of plant species recognized as hosts for the bacterium [146-148]. Several detailed reviews of the molecular basis for Agrobacterium-mediated transformation are available [143, 148, 149]. The realization that any DNA flanked by T-DNA sequences could be transferred from the Ti-plasmid and integrated into the genome of infected plant tissues was the key to development of transformation systems based on Agrobacteria. Of course, crown galls and hairy roots are not desirable features for trees of the future, so the Ti-plasmids used for routine plant transformation have been domesticated by removal of the "oncogenes" (phytohormone and opine biosynthesis genes) which cause these tissue deformations. The oncogenes have been replaced in the T-DNA by a variety of selectable marker genes, such as those whose products confer resistance to antibiotics or herbicides (see below). To facilitate recombinant gene manipulations, the T-DNA and vir gene regions of the Ti-plasmid have, in many cases, been placed on separate plasmids to yield what are commonly referred to as binary vector systems [99, 150]. The shuttle plasmid of a typical binary system contains the T-DNA, and is often simply referred to as the binary vector. These shuttle plasmids generally contain a selectable marker gene, a multiple cloning site for insertion of the genetic construct, and a variety of gene expression modulators (promoters, enhancing elements, and terminators), all of which are flanked by the T-DNA sequences [149]. Elsewhere on the shuttle plasmid are replication origins competent for maintaining the plasmid in either Agrobacterium or E. coli - - the latter primarily to facilitate access to the well-developed molecular biology systems available for this organism. A wide variety of shuttle vectors are available, and improved or specialized versions are continually being announced [151-156]. The vir genes required for T-DNA transfer and integration with the binary system are


19

maintained in Agrobacterium on so-called helper plasmids, and a variety of these plasmids have been developed in an effort to improve the efficiency of T-DNA transfer and integration and to expand the range of host plants which may be transformed using the Agrobacterium system [157, 158]. This last point, in particular, is significant with respect to efforts at transforming commercial gymnosperm tree species. The range of host plants suitable for Agrobacterium transformation was long thought to be limited to the dicotyledonous angiosperms, since tumors are seldom formed when monocots or gymnosperms are treated with these bacteria [159 161]. However, tumor formation is a result of a complex series of events that occur after DNA transfer and integration, and, consequently, transformation may occur in the absence of tumor formation [157]. Since recognition of the separate nature of these events, a significant number of monocots have been verifiably transformed using Agrobacterium [162]. In some cases where DNA was not transferred to any cells in treated monocots, specific strains of Agrobacterium were shown to have difficulty in attaching to the cells in the wound [163], while in other cases the intended hosts were found to secrete compounds that blocked expression of the vir genes [164]. An increasing array of bacterial strains and plasmids have provided ways to circumvent some of these problems [165, 166]. As a consequence, various combinations of broad host range Agrobacterium strains and vir genes are now routinely tested against different genetic lines of recalcitrant host species during the development and optimization of appropriate transformation systems [167]. The observation that vir gene expression can be induced by tissue explants from a variety of algae, bryophytes, pteridophytes, and gymnosperms, as well as the monocotyledonous and dicotyledonous angiosperms, suggests that the host range for Agrobacterium transformation may yet be extended far beyond its current use [168]. However, it seems unlikely that Agrobacteriummediated transformation could ever become universally applicable to all species of plants. As can be seen from Table 1, a variety of woody dicots, including species important for the wood and wood-product industries, have been regenerated from tissues transformed using Agrobacterium. Sadly, amongst the conifers, Agrobacterium-mediated transformation followed b y regeneration has so far only been accomplished with larch [-131,169]. Although Agrobacterium has been shown to be capable of introducing DNA into the cells of other conifer species [ 170-173], regeneration of plants from such transformed tissues remains problematic. It seems that those conifer cells predisposed to regenerate are difficult to transform with Agrobacterium, while easily transformed tissues are recalcitrant to regeneration [174]. It appears that a variety of factors, including the genetic background and metabolic state of the host cells, play a significant role in governing the susceptibility of these plants to Agrobacterium-mediated transformation [135, 175]. As a consequence, it is difficult to determine at this time whether the lack of progress in this area could best be addressed through focused research into improved Agrobacterium systems or more reliable regeneration protocols for the commercial gymnosperms.

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4.3 Physical Gene Transfer 4.3.1 Transformationby Microprojectile Bombardment Increasingly, microprojectile bombardment is becoming the transformation technique of choice for plants that are recalcitrant to Agrobacterium-mediated systems [176-179]. In this method, DNA containing the transgene is coated onto small (1-5 ~tm) particles of gold or tungsten which are subsequently shot into the target tissues with sufficient force to pierce the plant cell walls. The particles lodge within the cells without killing them, and the DNA is released for incorporation into the recipient genome. This technique is not restricted to particular plant species or even to plants in general, as it has also been used to transform a wide variety of animal cells [180-182], yeast [183, 184], fungi [185, 186] and even bacteria [187]. Microprojectile bombardment can even be used to transform the genomes of chloroplasts [188, 189] and mitochondria [183]. In the original instrument, tungsten microcarriers were accelerated by a gunpowder charge (hence the common reference to "gene guns") [190], but the commercial instrument developed from that early work uses a controlled burst of pressurized helium for improved consistency [191]. Subsequent variations on this general theme have resulted in the development of instruments optmized for increased control over the bombardment conditions [192, 193] or decreased cost of the instrumentation [194, 195]. The equipment designs described in the latter papers put this technology well within reach of any moderately wellfunded laboratory contemplating plant transformation experiments. As is the case for binary vectors used in Agrobacterium-mediated transformation, vectors containing various selectable markers, multiple cloning sites, and expression modulating sequences (promoters/terminators) have been designed specifically for microprojectile transformation work [196]. The principal difference between these vectors and those used in Agrobacterium-mediated systems is the elimination of DNA required by Agrobacterium, such as the T-DNA borders and the Agrobacterium-competent origin of replication. The smaller vectors that result from omission of these sequences are more tractable to many of the molecular biological manipulations used to prepare transgene constructs, i.e. plasmid preparation, restriction digests, ligations, and bacterial transformation, but a further reason for minimizing extraneous DNA in these vectors lies in the nature of the integration event. With Agrobacterium-mediated transformation, only the DNA lying between the T-DNA borders is transferred into the recipient cell, and the number of copies of T-DNA integrated into the host genome is usually very low [149]. However, with microprojectile transformation, many copies of the entire plasmid containing the transgene are introduced into every cell, and it is often the case that many copies of this DNA are integrated in the recipient genome [149]. Given the current regulatory concerns with regard to characterization of the integrated DNA prior to field release of transgenic plants, it is surprising that more effort has not gone into developing minimal DNA vectors specialized for microprojectile transformation


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similar to those developed by During [155] for Agrobacterium-mediated transformation. On the other hand, efforts are underway to develop innovative systems for site-specific transgene insertion [197, 198], and, in this respect, elements from both the Saccharomyces FLP/FRT system [199, 200] and the bacteriophage P1 Cre-loxP system [201,202] have been successfully tested in plants. The specificity inherent in these systems would appear to be ideal for the development of transgenic plants with precise insert structure and location. Further flexibility in future transgenic plant development may result from using derivatives of the Ac/Ds transposable element system from maize to remove specific sequences, such as selectable marker genes, after transformation has been accomplished [203].

4.3.2 Transformation of Protoplasts Direct uptake of foreign DNA by plant cells can potentially avoid the difficulty presented by limited susceptibility of some woody plants to infection with Agrobacteria, as well as the high equipment costs usually associated with microprojectile transformation. Protoplasts can be induced to efficiently take up naked DNA under the influence of polyethylene glycol (PEG) [204-206] or a polarizing electric field (electroporation) [207-209]. Several forest tree species of commercial importance having limited transformability by Agrobacterium, including various spruces and eucalypts [109, 174], have demonstrated at least transient expression of foreign genes introduced by one or more of these techniques. Systems for regenerating spruce, pine, and eucalyptus trees from protoplasts have also been reported [210]. However, the high labor demand required to isolate and culture protoplasts, as well as the need for a reliable regeneration system, remain significant drawbacks to widespread use of this transformation technique. In addition, the increased risk of isolating somaclonal variants from transformed protoplasts limits the usefulness of this system for developing commercial products [174]. On the other hand, the number of woody species for which protoplast regeneration systems are available is constantly increasing [109, 210], and in some situations protoplast transformation may provide unique opportunities, for example, as an alternative to microprojectile bombardment in organelle transformation [211,212].

4.3.3 Alternative Transformation Techniques The transformation techniques described above are by far the most widely used and accepted. However, each has its drawbacks. Consequently, a variety of alternatives have been proposed, but most have met with marginal success at best [101]. Sawahel and Cove [213] have provided a fairly concise catalog of most of these alternative techniques, but two interesting techniques for transforming intact cells, sonication [214] and silicon carbide fiber microinjection

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[215], were developed too recently to be covered by these authors. The latter technique seems particularly simple and inexpensive, and has been used successfully in several different laboratories [216-219]. The essence of the technique is that DNA is coated onto minute (0.6 ~tm diameter, 10-80 lain length), rigid fibers of silicon carbide, and these are subsequently vortexed with suspension-cultured cells of the recipient plant. Like tiny syringe needles, the fibers penetrate the cell walls without killing the cells and deliver the DNA for subsequent integration. This technique would appear to have the potential to gain widespread acceptance once its less precise aspects, e.g. how best to immobilize the DNA on the fibers, are more rigorously optimized. In terms of cost efficiency and ease of use, silicon carbide fiber transformation appears to hold great potential for enabling any laboratory equipped for plant tissue culture to successfully embark on plant transformation projects.

4.4 Selection Systems, Promoters, and Regulatory Elements To identify those cells that have received the transgene, transformation vectors carry either screenable or selectable markers [104, 220]. Selectable markers encode enzymes that enable the transformed cells to grow under conditions that kill non-transformed cells; thus, the most commonly used selectable markers encode enzymes that detoxify antibiotics or herbicides added to the culture medium. Neomycin phosphotransferase (kanamycin-resistance) [221], hygromycin phosphotransferase [222], gentamicin acetyltransferase [223, 224], and the Tn5 gene encoding bleomycin resistance [225] are the most widely used antibiotic resistance markers, while phosphoinothricin acetyltransferase (biaphalos-resistance) [226], modified 5-enolpyruvylshikimate-3-phosphate (EPSP) synthase (glyphosate-resistance) [227], modified acetolactate synthase (chlorsulfuron-resistance) [228], and bromoxynil nitrilase (bromoxinyl-resistance) [229] are commonly used herbicide resistance markers. It is not unusual to find that cell lines recovered from selective media have lost their regenerative capacity, and this is a principal reason for using screenable markers. The most widely used screenable markers have been chloramphenicol acetyltransferase (CAT) [230] and/~-glucuronidase (GUS) [231], but these have the drawback of requiring destructive sampling of the putatively transformed tissues in order to assay the enzyme activity. Alternatively, a gene from the jellyfish, Aequoria victoria,which encodes a protein (GFP) that fluorescences green under UV irradiation, is showing great promise as a non-destructive screenable marker for transgenic organisms [-232]. Modified versions of the gene have already been developed to express proteins with altered emission spectra so that the expression from multiple gene constructs may be monitored simultaneously [233]. GFP has been used as a reporter gene in several plant systems, including suspension-cultured Citrus cells [234, 235], after the gene was modified to eliminate problematic sequences, such as a cryptic splice site [236]. However, its use in woody species may be limited, since transformed cells of


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L. tulipifera or L. stryraciflua which expressed the G F P polypeptide did not fluoresce (Kim and Merkle, unpublished observations), possibly because these particular cells could not catalyze the post-translational event necessary to form the fluorophore. The promoter and terminator sequences most widely used for expression of transgenes in plants have been derived from one of three sources: the 35S RNA of cauliflower mosaic virus (CaMV), the nopaline synthase (hop, nos) gene from one class of A. tumefaciens Ti-plasmids, or the octopine synthase (ocs, oct) gene from a second class of A. tumefaciens Ti-plasmids. These promoters all lead to expression of chimeric genes at high levels in a wide variety of plant tissues, and various combinations have been developed to improve constitutive expression of gene constructs [220]. To efficiently modify characteristics of specific plant tissues, e.g. wood, increased efforts need to be made to identify promoters that can drive gene expression to high levels in the specific tissues of interest (106). Relatively few such promoters have been identified in plants, and woody plants, in particular, have received only cursory examination with respect to the promoters that control gene expression. To some extent this may be due to a perception that research in this direction should be supported by those commercial enterprises that stand to gain the most from application of tissue-specific promoters. 4.5 Current and Future Targets f o r Genetic Engineering in Forest Trees 4.5.1 Lignin Content and Composition Lignin makes up 20~30% of the total dry weight of wood and constitutes the principal barrier to production of pulp and paper. Estimates suggest that altering the composition of lignin in gymnosperms so that it resembles the more easily extracted lignin in angiosperms could provide the US industry alone with an annual saving in excess of $6 billion [237]. The potential savings that would accrue from reductions in total lignin content of the order of 10-15% have been suggested to be of a similar magnitude. Because the lignin biosynthetic pathway is relatively well understood, it has provided an opportune target for early experiments in genetic engineering of forest trees, and detailed reviews of efforts to date are available [107, 238]. Reduced cinnamyl alcohol dehydrogenase (CAD) activity in transgenic plants leads to incorporation of hydroxycinnamoyl aldehydes into lignin, and the resultant red-brown polymer is much easier to remove from fibers using chemical pulping techniques [239,240]. Transgenic plants having reduced hydroxycinnamoyl O-methyltransferase (OMT) activity have shown a variety of effects in terms of both lignin content and composition [241-244]. However, it is hoped that co-expression of an angiosperm OMT along with a ferulate-5hydroxylase (F5H), such as was recently cloned from Arabidopsis [245, 246], will

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lead to the production of sinapyl alcohol in transgenic gymnosperms. A successful outcome would result in the conversion of the softwood lignin from the less easily degraded guaiacyl-rich polymer to a more easily degradable guaiacylsyringyl copolymer, such as that which distinguishes hardwood species. Extracellular glucosidases with high specificity for monolignol glucosides have been purified from conifer cambium, and these would appear to provide excellent targets for efforts to alter lignin deposition [247, 248]. The oxidative polymerization step in lignin biosynthesis presents another interesting possibility for manipulating lignin via genetic engineering [-107]. Attempts to down-regulate lignification by reducing peroxidase expression in transgenic plants have not yet yielded positive results [249], and efforts to use antisense laccase genes in a similar manner have so far been inconclusive E250, 251].

4.5.2 Sterility and Early Flowering The release of genetically engineered trees into environments in which they can interbreed with wild populations is unlikely to be approved by regulatory agencies unless mechanisms to control sexual reproduction are in place [-252]. Thus, if the value of genetically engineered trees is to be realized, it is essential that sterile tree lines be created. In addition to solving problems associated with release, sterile tree lines will have the added benefit that proprietary genetic materials will be better protected from acquisition by competitors. It is also possible that by blocking formation of reproductive structures, energy resources will be redirected into vegetative growth, thereby increasing growth yields. The production of cellular toxins under the control of promoters specific for gene expression in pollen-producing tissues has been shown to result in male sterility [253, 254], and such a system should function equally well in trees. Tree breeding could be greatly accelerated if trees could be induced to flower while they were still seedlings, i.e. within 1 or 2 years following germination. Not only would this dramatically shorten the breeding cycle, but it would also make breeding in the controlled environment of the greenhouse possible. Finally, induction of early flowering in forest trees would enable rapid characterization of inheritance patterns of transgenes in the progeny of transgenic trees. Recent research with a group of flower-meristem-identity genes from the herbaceous model plant, Arabidopsis, has culminated in the production of transgenic Arabidopsis, in which precocious flower development was induced by overexpression of the inserted transgene [255]. Hybrid aspen transformed with the same Arabidopsis LEAFY gene (LFY) under the control of the cauliflower mosaic virus 35S promoter produced flowers after only five months in the greenhouse [-256]. This work has obvious potential to completely revolutionize the entire field of tree breeding.


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4.5.3 Herbicide Resistance Herbicides are widely used in site preparation to minimize competition from weedy species in newly replanted forests. However, these chemicals often cannot be applied directly to the trees themselves, or even to the area immediately around them, without causing significant damage or even death. Glyphosate is a broad-spectrum low-toxicity herbicide that blocks the aromatic amino acid biosynthetic pathway at EPSP synthase, and genes encoding mutant forms of this enzyme have been shown to confer resistance to the herbicide when expressed in transgenic plants [227]. Resistance to glyphosate has been introduced into commercial hardwood [127, 257] and softwood [169] tree species, and resistance to other herbicides has been conferred to various other transgenic plants and trees [258-260]. As shading and competition for nutrients are known to be significant factors limiting tree growth rates in young plantations, development of herbicide resistance would appear to be a logical target for tree improvement. However, widespread application of these compounds can have a profoundly detrimental effect on the environment, and concern for this was highlighted in a recent survey that showed a widespread understanding among forestry professionals that aerial spraying of herbicides in forests would not easily be accepted by the general public [261]. On the other hand, as noted by Jouanin et al. [110], herbicide-resistant trees would probably be most helpful in a nursery setting where herbicide application can be more rigorously controlled.

4.5.4 Insect and Pathogen Resistance Endotoxin (Bt) produced by the Bacillus thuringiensis soil bacterium has enjoyed widespread forest application as a non-toxic biopesticide that is highly specific for phytophageous lepidopteran, coleopteran, and dipteran insects [262]. The biology and mode of action of this biopesticide has been reviewed [263-265], and various studies have examined factors influencing its use in forest settings as well as its persistence in forest ecosystems [266, 267]. Transgenic tobacco and tomato plants expressing this protein were shown to be protected from feeding by lepidopteran larvae [268, 269], and similar protection from gypsy and forest tent moth caterpillars was demonstrated in transgenic poplar [270-272]. Transgenic larch trees expressing the Bt endotoxin have been recovered by Shin et al. [169], but there are no reports, as yet, regarding the resistance of these softwoods to insect attack. Although it is anticipated that such genetically engineered trees will eventually be deployed in commercial plantings, insects have been shown to be capable of developing resistance to these toxins [273]. There is thus a need to develop appropriate deployment strategies [274]. Protease inhibitor proteins are produced in many plants as a response to wounding or insect feeding, and the role of these proteins is to interfere with digestion in the insect gut [275]. A trypsin inhibitor protein from cowpea was

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used to protect transgenic tobacco from Heliothis virescens larvae [276], and a cysteine protease inhibitor from rice was recently shown to protect poplar from a species of boring beetle [277]. Schuerman and Dandekar [111] reviewed efforts to date to engineer plants for viral resistance, but work with woody plants has so far been limited to horticultural species [278, 279]. A variety of strategies using transgenes to control fungal diseases have been discussed and tested [280, 281]. However, none of these techniques have yet been tested in trees.

4.5.5 Tree Form and Fiber Morphology Wood formation requires carbon partitioning to favor the plant stem, and Timmis and Trotter [237] noted the potential for increasing carbon deposited in the tree bole by altering the growth habit of the tree. They used several lines of evidence to suggest that tree growth habit might be controlled by a limited number of genes, and speculated on the possible significance of work performed by Klee and co-workers [282] in which transgenic petunias having altered auxin metabolism showed significant increases in xylem and phloem formation. Recent work has shown that alteration of auxin metabolism in transgenic poplar leads to a variety of changes in tree form and wood characteristics [283].

4.5.6 Novel Traits There has recently been significant interest in using plants to clean up sites contaminated with heavy metals and toxic organic wastes [284 288]. Stomp et al. [289] noted a variety of tree growth habit characteristics that would make these plants very good candidates for use in phytoremediation.

4.6 Regulatory Considerations Like all other technological advances, the production of transgenic plants brings with it the potential for unforeseen consequences [290-292]. To minimize the possibility that such consequences could lead to irreversible problems when these organisms are introduced into the environment, all transgenic plants are subjected to a process of risk assessment prior to field release. The regulations governing such risk assessments vary from country to country, but Raffa [293] presents a set of guiding principles that are valid in most cases. In addition to the explicit governmental regulations on the subject, several recent reviews discussing the possible impacts of transgenic plant release should be considered by researchers attempting to create transgenic plants for use in the commercial sector [294-299]. Although most of these guidelines and rules were developed to address situations encountered in the cultivation of agricultural crops, it can


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readily be argued that these considerations are even more critical in the case of genetically engineered trees, since these organisms have a longer life-span and hence a much longer period in which to pass their transgenes to wild populations.

5 Molecular Breeding Classical plant breeding strategies are generally untenable for most forest tree species. Long generation times (5 to 20 years), coupled with the fact that many traits important to forest product industries can only be fully assessed after the tree has reached maturity, preclude rapid analyses of test crosses and have, thus, effectively limited breeding programs to the most economically important tree species. Even in these cases, family pedigrees of more than three or four generations are rare. To avoid the delay of having to score mature phenotypes, breeding experiments have increasingly relied on biochemical markers whose expression has been correlated with particular mature phenotypes. For example, isozymic variation has been widely used in agronomic as well as forest breeding programs [e.g. 112, 300]. For isozyme analyses, proteins contained in tissue extracts are resolved in a gel matrix on the basis of their physical properties (size, shape, or electrical charge). Subsequent staining based on the catalytic activity of the enzyme results in specific visualization of the isozymes, even in the presence of numerous other unrelated enzymes. An advantage of this method is that it yields co-dominant markers, since both alleles of a heterozygous locus can be detected. Unfortunately, there are relatively few biochemical markers whose expression has been matched with desirable phenotypes, and the expression of these markers is in many cases highly dependent upon environmental factors. It is also important to note that only a small amount of the genetic variation in a population is displayed phenotypically. Furthermore, since only about one-third of amino acid substitutions effect changes in the protein that can be detected by electrophoretic techniques, no more than one third of the total genetic variation is discernible through the use of isozyme analysis. With respect to the potential variation contained in the entire genome, isozymic markers are even more limited, since only about 0.5% of the typical eukaryotic genome consists of coding sequences [301]. Using DNA-based markers, researchers can potentially access all of the variation contained within a given genome, thereby increasing their chances of finding a marker that segregates with the specific phenotype of interest. Thus, the main advantage of molecular markers is that they are based on the polymorphisms occurring naturally in the DNA of a given species, and, as forest trees are among the most genetically variable organisms known [302], molecular markers useful for tree breeding programs should not be difficult to identify. In

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addition to constituting a larger pool of potential markers, DNA-markers have the advantage that they do not change in response to environmental factors or the developmental stage of a particular plant tissue. For a general review on the use of molecular genetic markers in plants see Rafalski and Tingey [303], and for comprehensive discussions of the ways in which molecular markers may be applied to forest tree improvement programs consult Neale et al. [304] and Haines [112].

5.1 DNA Marker Techniques 5.1.l Restriction Fragment Length Polymorphism (RFLP) RFLP (restriction fragment length polymorphism) analysis was first described by Botstein et al. [305], and has been reviewed elsewhere [306-308]. Briefly, DNA is extracted and cut into discrete fragments using restriction endonucleases. The DNA fragments are then separated by gel electrophoresis and transferred to a membrane filter so that they can be detected using specific RNA or DNA probes in a process known as Southern blotting. Polymorphisms are then detected by the presence or absence of labeled probe in bands on the blot [309]. Advantages of this method are that RFLPs are multi-alleleic and co-dominant (i.e. heterozygotes can be distinguished from either homozygote), and there are a virtually unlimited number of potential probes. Drawbacks include the relatively large amount of DNA required (2-15 pg per gel lane, depending on the genome size of the species, although it should be noted that a single blot can be reprobed many times), as well as the length of time and high cost required for each analysis. More problematic is the potential for artifacts which arise from incomplete digestion of the DNA by restriction endonucleases. In order to maximize sample throughput, procedures for preparing DNA are often streamlined, and, as a result, the DNA can be impure. Impurities in turn lead to partial digestion which becomes manifest as extra bands on the Southern blots.

5.1.2 Random Amplified Polymorphic DNA (RAPD) RAPDs (random amplified polymorphic DNA) are dominant molecular markers pioneered by Williams et al. [310] and Welsh and McClelland [311]. The method is based on PCR (polymerase chain reaction) and uses random oligonucleotide primers (usually 10-mers) to amplify genomic DNA. The use of short primers and low annealing temperatures leads to the generation of PCR products from multiple sites in the genome, and polymorphisms become apparent when sequence differences affecting primer binding sites give rise to different banding patterns when the reaction products are analyzed by gel electrophoresis.


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By way of advantages, RAPD analyses do not require prior sequence information, and none of the plant-derived DNA requires passage through bacteria. In addition, the technique requires very little DNA (10-25 ng), radioisotopes are unnecessary, and once the DNA has been prepared, the experimental procedure is relatively simple and amenable to automation [312]. The principal disadvantage with RAPDs is that only one strand of the DNA is amplified, and, as a consequence, dominant alleles are visualized whether the individual is heterozygous or homozygous at the particular locus. Haploid tissues, such as occur in conifer megagametophytes, can be used to circumvent this limitation [313]. A further disadvantage is that PCR-based technologies on the scale required for plant breeding are relatively costly because of the expense of the polymerase.

5.1.3 Bulked Segregant Analysis Bulk segregant analysis, described by Michelmore et al. [314], is a useful technique for finding new markers, typically RAPDs, that are linked to a particular locus or to an area that is sparsely populated with markers. The method uses two groups of individuals that arise from a single cross and are homozygous for alternate alleles governing a particular trait. If members of a segregating population are pooled on the basis of their phenotype, then the groups can be expected to be heterozygous at all unlinked loci but markedly skewed in the direction of one or other parental allele for a linked locus. Advantages of the procedure are that arbitrary primers can be used as probes, and the ability to pool individuals speeds analysis. One particular disadvantage of this technique for the purposes of forest tree breeding is linkage equilibrium, which is highly outbred species may mask the segregation of loci. In the case of conifers, this particular problem can again be minimized by analyzing haploid megagametophyte tissues from a sufficient number of individuals within a family [3133.

5.1.4 Microsatellite Repeat Polymorphisms Microsatellite repeats are highly polymorphic regions of DNA that occur frequently in eukaryotic genomes and are most commonly represented by the dinucleotide repeats [AC]n, [AG]n, and [AT]n [315-317]. PCR primers based on DNA sequences from regions flanking microsatellite sequences may be used to amplify the genomic DNA spanning the repeat region. The resultant products may then be resolved by gel electrophoresis to reveal allelic variations in the lengths of repeats. Advantages of this technique are that microsatellite repeats occur with moderate abundance, are highly polymorphic, and provide codominant markers. In addition, because the technique is based on PCR, it requires very little input DNA, and no radioisotopes are required.

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Disadvantages include a requirement for cloning and sequencing to characterize multiple microsatellite loci, not all of which will prove to be useful for analysis, as well as a need to use high resolution gels in order to differentiate polymorphisms as slight as two or three nucleotides.

5.1.5 Amplified Fragment Length Polymorphisms (AFLP) A DNA fingerprinting technique referred to as amplified fragment length polymorphism (AFLP) uses genomic DNA digested with restriction endonucleases as a PCR template [318]. The technique is similar to RFLP analysis, but relies on the power of PCR, rather than hybridization, to reveal subtle differences in the DNA from different sources. The binding between short PCR primers and DNA restriction fragments is very specific, and may not even occur between sequences that differ by as little as one or two nucleotides. Advantages of the method include no need for prior sequence data and the ability to selectively amplify sequences from a large number of restriction fragments. In addition, AFLP mapping can be used to detect corresponding genomic clones in large fragment libraries, e.g. cosmid or yeast artificial chromosomes. On the other hand, the technique is limited in practice to the resolving power of the gel electrophoresis system. Thus, reactions that generate more than 50 to 100 fragments are generally too complex to analyze. Also, the method functions best when used in conjunction with established breeding programs and documented multigenerational lines. The procedure also shares the high cost inherent in RAPD and RFLP analyses as well as the lengthy time requirements of RFLP mapping.

5.1.6 Microsatellite Hybridization Another recently developed method takes advantage of the fact that, in a typical RAPD analysis, abundant PCR products may be visualized by staining, but additional products in quantities below the staining threshold of ethidium bromide are also resolved in the gel. These minor products can be visualized by hybridization with microsatellite probes (e.g. [GT]8, [CT]8, [CC]12, [GA]12) to produce multiple independent and polymorphic fingerprints [319,320]. Advantages of this combined method include no need for prior sequence information, low input of template DNA, and when using non-isotopic detection methods, the possibility of using probes repeatedly. The method is especially useful for displaying polymorphisms in species where little variation is revealed by RAPD analyses alone. However, the method also suffers from the relatively high cost associated with RAPD analyses, as well as the time required for RFLP analyses.


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5.2 Applications o f DNA Markers in Forest Research The long generation times of forest trees places a premium on the development of methods that will allow for early genetic selection of the desired phenotype. Indirect selection based on DNA markers can be practiced at a very early age once linkages to important traits are identified. The advantages are obvious in that cost would be much lower than with progeny testing, selection intensities would be much higher, and selection would be potentially more efficient because of the higher heritabilities of the markers. One recent development that has great potential for forest tree breeders came from studies of the genes that regulate floral development in the model herbaceous plant, Arabidopsis thaliana. By introducing the LEAFY gene from Arabidopsis into aspen it was possible to induce flowering in transgenic aspen during their first year of growth as opposed to waiting the typical 10-20 years [-256]. Undoubtedly, this technology will find widespread application in forest tree improvement programs, particularly when coupled with marker-assisted selection.

5.2.1 GeneticLinkage Maps Genetic linkage maps are constructed by analyzing RFLP and/or RAPD markers and monitoring the segregation of a marker among progeny of a test cross. The high degree of polymorphisms allows for a virtually unlimited number of markers to be mapped using a single segregating population. A database of markers is made from the same set of F2 or backcross plants. When a map containing 100-200 well-dispersed markers has been constructed, virtually any new marker can be linked to one that has been previously mapped. The accumulated markers for a segregating population form a database that becomes a valuable resource for placement of new markers, and the database can be expanded by distributing the mapping data to other research groups. The practical strategies for building genetic linkage maps in plants was recently reviewed [321]. A major obstacle to the assembly of genomic maps for marker-aided tree breeding is that most tree species are highly outcrossed, resulting in significant linkage equilibrium between marker loci and the genetic locus of interest [322]. As a consequence, relationships established from the analysis of one cross do not necessarily hold for a second cross and, therefore, conventional mapping of forest trees in natural populations is extremely difficult. It has been argued that these hurdles severely limit the utility of marker-assisted selection in forest tree breeding. However, an opposing view, based on recent technological and theoretical advances, has been offered by O'MaUey et al. [323]. These authors note that by screening a sufficient number of potential RAPD markers and choosing the most reliable, i.e. markers that are consistently easy to score, one is able to quickly map the genome of individual trees. With maps of individual trees, breeders can minimize the difficulties brought about by linkage equilibrium and

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circumvent the need for the extended pedigrees commonly used for agronomic crop improvement. Of special interest to forest tree geneticists is the advantage provided by using conifer megagametophyte tissue as a DNA source. The haploid nature of this DNA allows for recombination and segregation events to be followed among open-pollinated seeds from a single tree, and circumvents the dominant allele problem inherent in using RAPDs with diploid samples. One is able to search for markers that are consistently found together in the same seeds, indicating that they are on the same chromosome [313]. This unique advantage of gymnosperms serves to partially offset the difficulties inherent in the large genome sizes typical of many softwood tree species. The lack of a suitable haploid stage in angiosperms led to the development of a "pseudo-test cross" method, which, in conjunction with RAPD technology, allows construction of single-tree genetic linkage maps [324]. The method ~elies on the observation that a cross between two heterozygous individuals results in many single-dose RAPD markers segregating 1 : 1 in their F1 progeny. Although this mapping strategy requires a controlled genetic cross to be made, the additional effort enables one to survey twice the heterozygosity, i.e. that from each parent. The method may also be applied to conifers to quickly generate single-tree linkage maps [-324].

5.2.2 Mapping Projects DNA-based molecular marker maps are being made for a number of hardwood and softwood species. RAPDs of megagametophyte tissue have been used to build a map covering 90% of the maritime pine (Pinus pinaster) [325] and 85% of the longleaf pine (Pine palustris Mill.) genomes [326]. Maps are being constructed for Norway spruce (Picea abies Karst.) [327], slash pine (Pinus elliotti Engelm. var. elliotti) [328], and white spruce (Picea 91auca (Moech) Voss) [313]. RFLPs were used to construct a map for loblolly pine (Pinus taeda L.) in a three-generation pedigree strategy [329]. More recently it was shown that RFLP probes used in building a loblolly pine map could be used in related species for the purpose of comparative genome mapping [330]. For eucalyptus, an outbred third-generation pedigree was used to build a map using RAPDs, RFLPs, and isozyme markers [331], and the pseudo-test cross strategy was used for Eucalyptus 9randis and E. urophylla [332]. A genomic map was made for peach [333] using bulk segregant analysis and RAPDs.

5.2.3 Marker-Assisted Selection Molecular markers provide a powerful tool for monitoring traits difficult to select for using traditional techniques, as well as for the simultaneous selection


33

of multiple traits. To use this technique, linkages between marker loci and phenotypically (economically) important traits must be identified. Then, instead of waiting for the tree to reach maturity before it displays the trait of interest, the breeder can select the offspring of a cross that carry the specific linked DNA marker(s). With forest trees, marker-aided selection is best applied to single traits that are relatively well characterized, e.g. wood specific gravity [334]. Neale and co-workers have identified quantitative trait loci (QTLs) influencing wood specific gravity in an outbred pedigree of loblolly pine [335]. RAPDs have been used to identify QTLs which influence early height growth in pines [336], while RAPDs and the pseudo-test cross strategy were used to identify QTLs affecting vegetative propagation in Eucalyptus [332]. Bradshaw and Stettler [337] mapped QTLs for stem growth and form as well as spring leaf flush in poplar. The molecular genetics of rust resistance in poplars was studied by Villar et al. [338] who used RAPDs and bulk segregant analysis to quickly identify suitable markers which were subsequently validated in a 2 x 2 factorial mating design. Markers may also be used to locate valuable genes to facilitate their cloning by map-based or positional cloning. The technique is based on the premise that any gene consistently inherited with a marker must lie near it on the same chromosome. Once isolated and characterized, the genes could be used to transform recipient forest trees or used as markers for the early selection of desirable phenotypes in wild-type (non-transformed) trees. The approach shows the greatest immediate potential when applied to traits governed by single genes. Devey et al. [339] have combined the power of RAPDs and bulked segregant analysis of haploid megagametophyte tissues to make significant progress with this approach by identifying ten RAPD markers that map close to a sugar pine (Pinus lambertiana Dougl.) gene for resistance to white pine blister rust, one of the most damaging pathogens of Southern pines. Fusiform rust is a devastating disease of loblolly pine for which resistance was for many years assumed to have a polygenic basis [9]. However, RAPDs and bulk segregant analysis were recently used to identify a region of the host genome that behaves as a single dominant gene and is responsible for resistance to this disease [340]. Narrowcrown growth habit is a desirable phenotype for high-density forest plantations, and RAPDs, in conjunction with bulked segregant analysis, have been used to identify a locus linked to pendula, a single gene controlling the narrow-crown phenotype in Norway spruce (Picea abies L.) [341]. Three markers linked to resistance to black leaf-spot disease in Chinese (Ulmus parvifolia) and Siberian (U. pumila) elms [342], and two markers linked to scab resistance in apple (Malusfloribunda) [343] have been identified using a similar strategy. Considering the difficulties (transformation, vegetative propagation, government regulations) currently besetting genetic engineering of forest trees, it is conceivable that marker-assisted selection may have greater near-term impact on forest tree improvement [323].

34


5.3 Other Applications o f DNA Markers 5.3.1 Quantification of Genetic Diversity RAPD markers have been used to investigate the distribution of variability in natural populations of Eucalyptus 91obulus and provide the foundation for effective breeding and gene conservation strategies [344]. RAPDs have revealed an extensive amount of genetic diversity in trembling aspens (Populus tremuloides) in a study that also highlighted the benefits of careful PCR primer selection [345]. The authors identified a single RAPD primer that was able to separate 83% of the trees sampled in any given population. Microsatellite markers have also been used to determine the level of genetic variation in tropical rain forest species where they proved to be powerful tools for analyzing population structure with respect to gene flow and paternity [346].

5.3.2 Genotype Verification and Delineation RAPD markers have proven useful in identifying the parentage of progeny. In European white birch (Betula pendula Roth), a commercial set of random 10-mer primers was used to confirm lineages [347]. The authors pointed out that although robotics are becoming more commonplace in repetitive PCR routines, there is still considerable labor involved in the preparation of amplificationquality DNA templates. To minimize this problem, they developed a pooledprogeny strategy similar to that used in bulk segregant analysis [314], in which leaf samples of equivalent fresh weight from different individuals were pooled and used as the source of DNA for subsequent RAPD analyses. RAPDs have also been used to analyze somatic embryos and plants regenerated from embryogenic lines of Norway spruce (Picea abies (L.) Karst) to determine the level of somaclonal variation [348]. It was shown that material from the same cell line shared identical banding patterns, whereas regenerants from different cell lines were clearly distinguishable. RAPD patterns were reported to be somatically stable, thereby enabling researchers to trace clone identity from the laboratory to the field [349]. However, because of the dominant nature of RAPD markers, as well as the difficulty in reproducing banding patterns both in different laboratories and between pedigrees, the authors concluded that RAPD markers were not likely to replace isozyme analyses for this purpose. RAPD mapping of peach (Prunus persica) was reported by Pooler and Scorza [350], who cautioned that spontaneous somatic rearrangements (bud sports) could lead to unexpected inheritance of RAPD markers. RAPDs were used to identify mislabeled clonal material (ramets) in a sitka spruce (Picea stichensis) breeding program [351], and were further used to identify and assess the extent of natural introgression between populations of interior spruce, white spruce (Picea 9lauca (Moench) Voss) and Engelmann spruce (Picea engelmannii Parry) [352].

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6

Electronic and Computational Resources

Although not a biotechnology, sensu stricto, the Internet hosts a wide variety of information resources that are of increasing use to researchers in all areas of forestry. As with all other aspects of the Internet, forestry-related resources are being added at an exponential rate, and providing a current and comprehensive list of useful sites is becoming impossible. However, Table 2 contains a highly subjective list of relatively stable sites that should act as excellent starting points for those wishing to explore and sample the wide variety of forestry resources online. Table 2. A Selection of Internet Resources Related to Forest Biotechnology (5/1/96) Server Page Title General Starting Points Yahoo Forestry List Galaxy Forestry List TAPPIWeb List of Forestry Servers Natural Resources: Institutions and Organizations More Forestry Stuff The Pulp and Paper Jumplist Other Related Forestry Servers The WWW Virtual Library - Forestry Governmental Organizations CSIRO Forestry and Forest Products Dendrome-Tree Genome Mapping Project Finnish Forest Research Institute (METLA) Japanese Forest Research Institute National Center for Biotechnology Information Natural Resources Canada US Forest Service Educational Organizations Auburn University Institute for Paper Science and Technology Louisiana State University NYForest Online Oxford University Poplar Molecular Network Texas A&M University University of Alberta University of Illinois University of Maine University of Minnesota University of Washington

Internet Address http://www.yahoo.com/science/agriculture/ forestry/ http://galaxy.einet.net/galaxy/Engineering-andTechnology/Agriculture/Forestry.html http://www.tappi.org/resource.htm http://sfbox.vt.edu: 10021/Y/yfleung/forestry. html ht tp://forestry.bangor.ac.uk/forst uff.htm http://www.nlbbs.com/~ dc001/paper/j umplist. html http://www.chem.csiro.au/extpulp.htm http://www.icfrnet.nnp.ac.za/mirrors/Forestry. html http://www.chem.csiro.au/fordiv2.htm http://s27w007.pswfs.gov/ http://www.metla.fi/ ht tp://ss.ffpri.affrc.go.jp/ http://www.ncbi.nlm.nih.gov/ http://www.emr.ca/ ht tp://www.fs.fed.us/Homepage,html http://www.forestry.auburn.edu/ http://www.ipst.edu/ ht tp://wwwlfpl.forestry.Isu.edu/ http://149. 119.1.26/default.html http://ifs.plants.ox.ac.uk/ gopher://poplar 1,cfr.washington.edu:70/1 http://165.91.48.43/ http://www.rr.ualber ta.ca/ http://gopher.ag.uiuc.edu:80/NRES/forest ry-research.html http://www.ume.maine.edu/~ nfa/for mgt/ wse.htm http://mercury.forestry.umn.edu/FP/ForP.html http://weber.u.washington.edu: 80/--,cfrwww/


36 Table 2. (Continued)

Server Page Title

Internet Address

Independent Organizations

American Forest & Paper Association Technical Association of the Pulp and Paper Industry United Nations - FAO Corporations ForestNet ForestPro Lej6 International

http://www.infobahn.com.tw/adv/afpa/afpa.htm http://www.tappi.org/ http://www.fao.org/WAICENT/Forestry.htm http://www.forestnet.com/ http://www.forestpro.com/ http://www.transport.com/~ leje/homepg.html

Individuals

Jeff Lindsay Knut's Pages Steve Shook's Directory

http://www.athenet.net/~ jlindsay/Paper.shtm 1 :~top http://www.carleton.ca/~kmenard/forest.html http://weber.u.washington.edu/~ esw/fpm.htm

Miscellaneous

BiotechnologyLaw Web Server BiotechnologyPermits Home Page

http://biotechlaw.ari.net/ http://www.aphis.usda.gov/BBEP/BP/

Usenet Newsgroups

Agroforestry Pulp and Paper

bionet.agroforestry misc.industry.pulp-and-paper

Listservs Wood Net Wood Science

wood-net@ esusda.gov wood-science @unixg.ubc.ca

7 Conclusions The true innovator is often characterized by an ability to see the far-reaching trends and changes in technology and economy as well as their connections to global, social, and industrial patterns. The long rotation times required for forest tree production demand that the forester should have a particularly good crystal ball in order to anticipate future trends and needs for the trees being planted. When rotation times are short, e.g. Brazilian eucalyptus cut after seven years, it is relatively easy to predict that a newly planted tree will probably be used for fibers. Even with rotation times of the order of 25 to 30 years, as is the case for southern pine in the United States, one may be able to make a reasonably correct prediction regarding the future use of the tree. However, when rotation times run to 100 years or more, as is the case in Scandinavia, you need a crystal ball of increasingly high quality to make any reasonable predictions. Regardless of what will be the particular end use of trees in the next century, there is no denying that there will be an urgent need for them to grow faster.


37

Demand for fibers and solid wood will continue to grow even as harvestable forest acreage decreases due to population increases and set-asides that reserve forested areas for recreational purposes. In addition, advances in wood chemistry and biotechnology may also make trees the ideal feedstock for "biorefineries" that could produce alternatives to most of the current petrochemical products of oil-refineries. It is unlikely that we will forever be stuck with the dogma that forestry is only for the production of paper and timber. Thus, we anticipate that trees will eventually be cultivated like other plants for faster growth, greater yield, and a more diversified and efficient use. New silvicultural practices will certainly be needed to speed the growth of trees, and as a consequence, biotechnological techniques are likely to become commonplace in forestry. In vitro propagation will enable us to produce large numbers of genetically improved trees without having to establish large breeding orchards. Molecular markers will help in identifying superior genotypes as well as in monitoring the products of improved breeding schemes. Genetic engineering will provide the means to rapidly bring new genetic materials to bear on problems such as acid rain and attack by exotic insects. By combining traditional practices with the techniques of biotechnology, tomorrow's foresters will likely produce a revolution in the way forestry is practiced, as well as in the products which can be derived from trees. There is no doubt that there is a way ahead if only we have a head for the way.

8 References

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Microorganisms and Enzymes Involved in the Degradation of Plant Fiber Cell Walls Ramesh Chander Kuhad 1, Ajay Singh 2, and Karl-Erik L. Eriksson 3 Department of Microbiology, University of Delhi South Campus, BenitoJuarez Road, New Delhi-ll0021, India 2 Department of Biology, University of Waterloo, Ontario, Canada N2L 3G1 3 Center for Biological Resource Recovery, Department of Biochemistry and Molecular Biology, University of Georgia, Athens, GA 30602-7229, USA

1 Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1 Composition of W o o d and Other Plant Fibers . . . . . . . . . . . . . . . . . . . . . . 1.2 Structure and Composition of Wood and Other Plant Cell Walls . . . . . . . . . . . 1.2.1 The Cellulose C o m p o n e n t . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.2 The Hemicellulose Components . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.3 The Lignin C o m p o n e n t . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.4 Cell Wall Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.5 Other Cell Wall Components . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Microorganisms and Their Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Microorganisms Involved in the Degradation of Lignocellulosic Materials . . . . . . 2.1.1 Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.2 Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Enzymes Involved in the Degradation of Plant Fiber Cell Wall C o m p o n e n t s . . . . 3 Degradation of Cellulose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Microorganisms Producing Cellulose-Degrading Enzymes . . . . . . . . . . . . . . . 3.2 Cellulolytic Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.1 Regulation of Cellulase Production . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.2 Molecular Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.3 Cellulases are Organized in D o m a i n s . . . . . . . . . . . . . . . . . . . . . . . . 3.2.4 Catalytic Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Assay of Cellulolytic Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Possibilities for Biotechnology Based on Cellulolytic Enzymes . . . . . . . . . . . . . 4 Degradation of Hemicelluloses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Microorganisms Producing Hemicellulose-Degrading Enzymes . . . . . . . . . . . . 4.2 Hemicellulolytic Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.1 Xylan-Degrading Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 94.2.2 M a n n a n - D e g r a d i n g Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Assay of Hemicellulose-Degrading Enzymes . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Possibilities for Biotechnology Based on Hemicellulolytic Enzymes . . . . . . . . . . 5 Degrad ation of Lignin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Microorganisms Involved in Lignin D e g r a d a t i o n . . . . . . . . . . . . . . . . . . . . . 5.2 Ligninolytic Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.1 Lignin Peroxidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.2 Manganese Peroxidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.3 Laccase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.4 HzOz-Producing Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.5 Oxidoreductases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3 Assay of Ligninolytic Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5,4 Possibilities for Biotechnology Based on Ligninolytic Enzymes . . . . . . . . . . . . 6 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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One of natures most important biological processes is the degradation oflignocellulosic materials to carbon dioxide, water and humic substances. This implies possibilities to use biotechnology in the pulp and paper industry and consequently, the use of microorganisms and their enzymes to replace or supplement chemical methods is gaining interest. This chapter describes the structure of wood and the main wood components, cellulose, hemicelluloses and lignins. The enzyme and enzyme mechanisms used by fungi and bacteria to modify and degrade these components are described in detail. Techniques for how to assay for these enzyme activities are also described. The possibilities for biotechnology in the pulp and paper industry and other fiber utilizing industries based on these enzymes are discussed.

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1 Background Because of growing consumer demand for environmentally safe processes and products faced with market and legislative pressures, the pulp and paper industry is modifying its pulping, bleaching and effluent treatment technologies to reduce the environmental impact. However, new approaches to pulp and paper manufacture are rare because of technical and economical constraints [1]. Biotechnology has received increasing attention during the past two decades because of its commercial potential in many fields. The virtue of biotechnology lies in its potential for more specific reactions, to provide less environmentally deleterious processes, to save energy, and to be used where non-biological processes are impractical. The raw material in the forest product industries is wood and its components. Thus, possibilities for employing biotechnology in these industries must be numerous since one of nature's most important biological processes is the degradation of lignocellulosic materials to carbon dioxide, water, and humic substances. Consequently, the use of microorganisms and their enzymes to replace or supplement older chemical methods in the pulp and paper industry is gaining interest. The accessibility of enzymes to wood and fibers is limited due to factors such as adsorption to surface areas, low fiber porosity, and low median pore size of fibers [1]. In addition, the molecular organization of the different components of the plant fiber cell wall, i.e. cellulose, hemicellulose, and lignin, which are jointly termed the lignocellulose complex, also limits the accessibility of microorganisms and their enzymes to wood and its fiber components. Thus, the proper and useful application of microorganisms and their enzymes in improving pulp and paper production processes requires a more complete understanding of the molecular architecture of the substrates, mechanisms of enzyme action, and methods for the study of the specific degradation of each component of the fiber cell wall. This article reviews the composition of wood and other plant fibers, microorganisms, and their enzymes involved in the degradation of wood and its components cellulose, hemicellulose, and lignin, techniques for the assay of enzymes modifying and degrading these components, and the possibilities for biotechnology based on these enzymes.

1.1 Composition of Wood and Other Plant Fibers Lignocellulosics in the form of wood and agricultural residues are virtually inexhaustible, since their production is based on the photosynthetic processes. They account for more than 60% of total biomass produced. The net photosynthetic production of the dry biomass by plants on earth has been estimated to be 155 billion tons per year [2]. Regardless of source, lignocellulosic materials contain cellulose, hemicellulose and lignin as major components. Table 1 shows

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Table 1. Composition of some wood and agricultural lignocellulosic residues Residue

% Dry weight Hexosans

Pentosans

Lignin

Ash

Hardwoods Aspen Beech Birch Cottonwood Oak Poplar Red maple

50 47 41 46 48 45 39

28 20 26 19 18 19 33

15 23 25 24 28 20 23

0.3 0.2 1.0 0.6 0.4 0.1 1.0

Softwoods Douglas fir Eastern hemlock Jack pine White pine Red spruce White spruce

57 43 41 44 43 44

8 10 10 11 12 I0

24 32 27 28 27 27

0.4 0.4 0.1 0.1 0.2 0.3

Agricultural Residues Bagasse Barley straw Corn cob Cotton stalks Groundnut shells Oat straw Rice straw Rye straw Wheat straw

33 40 42 42 38 41 32 37 30

30 20 39 12 36 16 24 30 24

29 15 14 15 16 11 13 19 18

4 11 2 6 5 12 18 4 10

the composition of several hardwoods, softwoods and agricultural residues. The compositions of hardwoods and softwoods are significantly different. The lignin content of softwood is generally higher than that of hardwoods, whereas the hemicellulose content of hardwoods is higher than that of softwoods. With a few exceptions, straw species are more uniform in composition than wood species. Generally straws have lower cellulose content than wood, but, in spite of this, the holocellulose (cellulose + hemicellulose) fraction is approximately equal to that of wood.

1.2 Structure and Composition of Wood and Other Plant Cell Walls W o o d is m a d e u p o f cells o r fibers. A m a t u r e tree s t e m h a s a n o u t e r m o s t l a y e r o f b a r k ( p h l o e m ) s e p a r a t e d f r o m t h e w o o d (xylem) b y a t h i n layer, t h e c a m b i u m . T h e c a m b i u m is t h e l a y e r w h e r e t r a c h e i d s ( w o o d fibers) a r e p r o d u c e d [3]. T h e fibers a r e p r o d u c e d in a n a q u e o u s e n v i r o n m e n t a n d exist in a l i v i n g t r e e in a m a x i m a l l y s w o l l e n state. T h e g r o w i n g tree f o r m s l a r g e d i a m e t e r t h i n - w a l l e d


49

fibers in the early part of the growing season, small diameter thick-walled fibers in the late growing season. Thin-walled early wood fibers are much more flexible than the thick-walled late wood fibers. Wood is a porous material consisting of a matrix of fiber walls and air space. The air space exists mostly in the form of fiber cavities (lumens) and to a much lesser extent as voids within fiber walls. The wood fiber wall has three major constituents: cellulose, hemicelluloses, and lignin. Juvenile wood, heartwood/sapwood proportion, wood density, and fiber length are the wood quality attributes important to the pulp and paper industry. Among these, fiber length is of particular importance in pulp and paper quality. Wood is classified in two major groups: softwoods, i.e., gymnosperms (pine, spruce, larch, etc.) and hardwoods, i.e., angiosperms (birch, aspen, beech, maple, oak, etc.). They differ considerably in cell type. Softwoods have mainly two types of cells, the long (2-5 mm) tracheids, which give strength to the wood and are responsible for vertical water transport, and the smaller ray cells, which transport water in the horizontal direction. In addition, there are resin channels in both horizontal and vertical directions [3]. Softwood tracheids are connected by bordered pits [-4]. Hardwoods possess more diverse types of cells. In general, the annual growth ring of hardwoods is composed of an array of vessels, fibers, and ray parenchyma cells in various arrangements. These cells provide for water conduction, strength support, and transport and storage of nutrients. The thickwalled fiber tracheids and libriform fibers (fiber length 0.64-2.30mm) are located around the vessels and are connected to other cells via bordered or simple pits [4]. Vessels are thin and short (0.03-0.13 ram), but when located on top of each other they form tubes up to several meters in length. These vessels are more effective for water transport than the softwood tracheids [3]. Each tracheid cell is initially surrounded by a primary cell wall low in lignin content. As growth progresses, a secondary cell wall, into which lignin is successively incorporated, develops centripetally to the primary wall. The secondary wall constitutes the largest proportion of the total cell wall, and most of cell wall lignin (60-80%) is located here [-5-9]. Lignification is always preceded by deposition of cell wall polysaccharides, and polymerization of monolignols occurs within the carbohydrate gel. However, the type of carbohydrate changes during the course of cell wall development, i.e., the formation of cell wall layers [10]. A model illustrating the arrangement of lignin, hemicellulose, and cellulose within the cell wall has been proposed by Kerr and Goring [11]. Terashima and Fukushima [10] have schematically pictured the deposition of cell wall components and their irreversible assembly to form a lignified cell wall in tree xylem (Fig. 1). Every supporting cell (dead) has a lumen, which can be more or less empty or filled with water. In all softwoods and many hardwoods, the inner or warty layer is a heterogenous mixture of components of unknown composition. The secondary cell wall outside the warty layer is made up of three layers, $1, $2, and $3, with $3 closest to the lumen, Sz the middle layer, and $1 the outermost layer.

50

R.C. Kuhad et al. Cambium Deposition of celt walt components 9ML,CC-" ( Laps of time )

Formation of cell wall layers

w p --.. -S1--

w

, ,~Mature cell wall

52-" ~S3~

Deposition of carbohydrates

Deposition of [lgnin and type of monolignol

/Gymnosperm Monomer / composition |

Tracheid

. and . ~ Angiosperm condensation I Vessel

, Pectic substances ------ Arabinogalactan 1Xylan - - MannQn Ceilulo'~e

'

Llgnin , p-Coumary| alcohol 1 Conlferyl alcohol ~ G $1napyt alcohol H ~ ' ~ I ~ S ~ I~176

{ ;C 7.,ML ~ SW

{ CCML H

~ H

_

.

~

r

~ ~

,

_

~

i[Ageing .!! 'I 1

~

condensation

: I

I

Fig. 1. Schematic representation of the process of deposition of cell wall components and the middle lamella in gymnosperms and angiosperms. M L middle lamella, CC cell corner, P primary wall, C M L compound middle lamella, $1 outer, $2 middle, and $3 inner layer of secondary wall, H, G, and S, p-hydoxy-, guaiacyl-, and syringyl-propane units [10]

These layers consist of cellulose microfibrils embedded in an amorphous mixture of different hemicelluloses and lignin. The microfibrils form helices around the cell with different angles and directions for the different layers. The Sz layer is the thickest (1-5 gm) and the $3 layer the thinnest (0.1 gm). The width of $1 layers varies from 0.2-0.3 gm [3] It is important to note that cellulose, hemicelluloses, and lignin are present in each layer of the cell wall. The concentration of cellulose is highest in sub-layer $2 and decreases toward the middle lamella, where it is present only in small amounts. The $3 layer is rich in hemicelluloses, while lignin is the dominant compound in middle lamellae [12]. While lignin concentration is high in the middle lamella, this layer is extremely thin, and most of the lignin is in the secondary wall. Straw is a much more heterogenous material than wood. However, cell wall structures of straws have been less studied than wood fiber walls. Straw fibers, principally derived from cells and internodes, are fairly long and slender with sharply pointed ends. In addition to fibers, straw contains short nonfibrous tissues consisting of epidermal cells, platelets, serrated cells, and spirals which are derived from the pitch, nodes, chaffs, and rachises. While 96% of softwood cells may be considered as fibers, only about 35-39% of the cells in straw are fibers [12].


51

1.2.1 The Cellulose Component Cellulose is considered to be a linear homopolymer of hydroglucose units linked with [3-1,4-glucosidic units, although the true repeating stereochemical unit of cellulose is cellobiose ([3-1,4-D-glucosyl-D-glucose) [13]. Glucose and cellodextrins are the products when cellulose is hydrolyzed. Some natural materials, e.g., cotton, are almost pure cellulose. Cotton is mostly a-cellulose, the form of cellulose insoluble in 17.5% NaOH. Wood and other plant celluloses generally contain [3-cellulose as well, a material soluble in the above NaOH solution. The and [3 forms of cellulose also differ in their intermolecular hydrogen bonding patterns. The cellulose molecule is a polymer with a degree of polymerization (DP) of up to about 15 000. However, DPs as high as 25 000 have been reported for the algae Valonia. The cellulose molecule is thread-like, existing as bundles of molecules forming fibrils stabilized laterally by hydrogen bonds between hydroxyl groups of adjacent chains. The arrangement of cellulose molecules in the fibrillar bundles is so regular that cellulose has a crystalline X-ray diffraction pattern. The individual cellulose chains are packed in groups of about 30 to form elementary fibrils (microfibrils), approximately 100 of which are packed to form fibrils. These fibrils are further packed to form the cellulose fiber [14]. The microfibril has been considered to be stronger than steel of corresponding size and thus significantly contributes to the strength of wood [15]. Cellulose is known to exist in four structures (cellulose I-IV) as revealed by X-ray diffraction patterns [16, 17]. Cellulose I is the native form of cellulose in the secondary cell wall. Cellulose I can be converted into types II and III by chemical or physical treatment. The transition from cellulose I to cellulose II is irreversible. Cellulose II may, however, be the thermodynamically most stable form. Cellulose IV, the dominant structure in the primary cell wall, is believed to have a form similar to but more disordered than that of cellulose I. Cellulose I can be transformed to cellulose IV via cellulose III [-18-20]. In cellulose I the chains are arranged partially in flat sheets. These sheets are thought to be kept together by van der Waals forces, hydrophobic interactions, and interchain hydrogen bonds. The degree of crystallinity of cellulose depends very much on origin and type of pretreatment. It may vary from 0% (amorphous and acid-swollen cellulose) to approximately 100% (cellulose from Valonia macrophysa) [21]. The degree of crystallinity of cotton cellulose is about 70%, while the figures vary from 30 to 70% for most other commercial celluloses [22]. However, the crystalline forms of cellulose are still not completely understood. Crystalline cellulose is highly resistant to microbial attack and enzymatic hydrolysis, whereas amorphous cellulose is degraded at a much faster rate [23]. As its crystallinity increases, cellulose becomes increasingly resistant to further hydrolysis.

52

R.C. Kuhad et al.

1.2.2 The HemicelIulose Components Hemicelluloses are easily hydrolyzed [24] short chains of branched heteropolysaccharides composed of both hexoses and pentoses. D-Xylose and L-arabinose are the major constituents of the pentosans (xylans), while D-glucose, D-galactose, and D-mannose are the constituents of the hexosans (mannans). The major hemicellulose components in softwood are mannan-based, and those in hardwood xylan-based. The close association of the hemicelluloses with cellulose and lignin in the fiber cell walls contributes both to rigidity and flexibility. Hemicelluloses are composed both of neutral sugars, all present as their respective anhydrides, i.e. xylan, araban, glucan, galactan, and mannan (substituted with acetyl groups) and of uronic acids. Hemicelluloses constitute on average about 26% of hardwood, 22% of softwood, and 30% of various agricultural residues [25-28]. They usually have DPs of 100 to 200. Apart from galactose-based hemicelluloses, which are characterized by 13-1,3-1inkages, most of the hemicelluloses are built up by ~-l,4-1inkages between their backbone sugars. Besides wood, hemicelluloses are also found in grasses, cereals, and some very primitive plants [24]. The type and amount of hemicellulose varies widely, depending on plant materials, type of tissues, growth stage, growth conditions, storage, and method of extraction [29-31]. The mannan hemicelluloses, galactoglucomannans and glucomannans in softwoods and hardwoods, are both branched heteropolysaccharides. At least two galactoglucomannans, often termed "glucomannans" with different sugar ratios are predominant in softwoods. The less soluble polysaccharide and the major (water-soluble) polysaccharide, consist of galactose: glucose: mannose in the ratio of 0.1 : 1 : 3 and 1 : 1 : 3, respectively. Their backbones are built up of 1,4-1inked [3-D-glucopyranose and 13-D-mannopyranose units, largely distributed at random. The mannose and glucose units in the backbone are partially substituted in C-2 and C-3 positions by acetyl groups, approximately 1 per 3-4 hexose units [23, 32]. Depending on the wood species, the glucose: mannose ratio has been reported to vary from 1 : 1 to 1 : 2. Galactose is not present in hardwood mannans [23, 32]. The major class of hemicelluloses is xylans, which are found in large quantities in annum plants and deciduous trees, and in smaller quantities in conifers. Xylans in grasses and cereals are generally characterized by the presence of L-arabinose linked as a single unit substituent to a D-xylose backbone. Substantial differences in substituents are found in wood xylans. These xylans are characterized by the presence of 4-O-methyl-D-glucuronic acid, L-arabinose, or acetyl groups as substituents on the D-xylose backbone. In general, the content of 4-O-methyl-D-glucuronic acid is higher in softwood than in hardwood. Unbranched linear xylan homopolysaccharides are rare [33]. Xylans comprise 15-30% of annual plants, 20-25% of hardwoods, and 7 12% of softwoods. Xylans appear to be a major interface between lignin and other carbohydrate components in many isolated phenolic-carbohydrate complexes [-34-38], where they are probably covalently linked to phenolic residues via the arabinosyl [39]

Microorganisms and EnzymesInvolvedin the Degradation of Plant Fiber Cell Walls

53

and/or glucuronosyl residues [-40]. Xylans tend to adsorb onto cellulose and to aggregate with other hemicellulosic components, probably as a result of hydrogen-bonding interactions [41, 42]. Xylan may play a major role in cell wall cohesion since its selective removal from delignified wood fiber results in a substantial increases in fiber porosity [43]. There have been observations which suggest that cellulose is protected from enzymatic attack by xylan and mannan [44, 45]. Naturally occurring hemicelluloses differ from isolated hemicelluloses. Besides impurities of other cell wall materials, isolated hemicelluloses are altered during oxidative delignification, which may lead to reduction in the chain length of the polysaccharides. The structure of xylan isolated from wood is dependent upon the type of polymer originally present in the wood and also on the pH of the cooking liquor used in pulping. In Kraft cooking, with its high pH, xylans or arabinoxylans are found in good yield dependent upon whether the wood contains 4-O-methylglucuronoxylan or arabino-4-O-methylglucuronoxylan. The Kraft process begins with extreme alkaline conditions which causes losses of hemicelluloses. Two-thirds of the glucomannans are dissolved very quickly and degraded by alkaline peeling reactions [46]. The peeling reaction in xylans is much slower than those occuring with cellulose or glucommannans because of the unique sequence of sugar units at the reducing end of xylans [46].

1.2.3 The Lignin Component Lignin is not a definite uniform compound, but is a collective form for substances that have similar chemical properties but very different molecular weights. The molecular weight of lignins may be 100 kDa or greater [47]. A considerable part of the photosynthetic activity in plants is devoted to the conversion of atmospheric carbon dioxide to lignin. Lignin is found in the highest concentration in the middle lamella, but is most abundant in the secondary walls of vascular plants [48]. It performs important functions in the life of plants as a permanent bonding agent between cells, making a wood composite material, and is a UV light stabilizer, antioxidant, and water-proofing agent. It also protects plants, wood in particular, from attack by microorganisms. The water permeation-reducing property of lignin plays an important role in the internal transport of water, nutrients, and metabolites in the plant. Lignins are highly branched polymeric molecules consisting of phenylpropane-based monomeric units linked together by different types of bonds, including alkyl-aryl, alkyl-alkyl, and aryl-aryl ether bonds. The relative proportions of three cinnamyl alcohol precursors incorporated into lignin, i.e., p-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol, vary not only with the plant species but also with the plant tissues and location of the lignins within the plant cell wall. Ecological factors such as age of the wood, climate, plant sustenance, and amount of sunlight also affect the chemical structure of lignins. A major problem in studying the chemistry of lignins has been the difficulty in

54

R.C. Kuhad et al.

isolating intact lignins from plant materials. The hydrolyzable linkages in lignins are suggested to be of two types: I]-aryl ether and 0~-aryl ether [49, 50]. The predominant [3-aryl ether type bond is more resistant to cleavage. Under mild hydrolytic conditions, the cleavage of the ether bond is exclusively restricted to the a-aryl ether type [51, 52]. In terms of physical properties, lignins are amorphous polymers that have no crystallinity. The amorphous nature of lignin has been studied using various techniques such as broad-line nuclear magnetic resonance, differential scanning calorimetry, viscoelasticity, and X-ray diffractometry [53]. The mode of polymerization during lignin biosynthesis makes it optically inactive. Lignins are insoluble in water and difficult for microorganisms to penetrate and degrade. They are generally acid stable but can be solubilized under alkaline conditions. Lignins are closely associated to cellulose and hemicelluloses in plant cell walls, and it has been shown that some hemicelluloses are linked by covalent bonds to lignin [54]. Recently, Joseleau et al. [33] have discussed covalent bonding between lignin and xylan. The best documented are ester linkages between glucuronoxylans and lignin via benzyl ester bonds with the carboxyl group of 4-O-methylglucuronic acid [40, 55, 56]. This bond is likely to be established between quinonemethide intermediates and D-glucuronic acid during the polymerization processes [57]. The second most reported covalent bonds between xylans and lignins are ether linkages involving the L-arabinose side chains [58] or xylose units [59, 60]. Lignin-xylan complexes have been isolated from hardwoods, while from softwoods both lignin-mannan and lignin-xylan complexes were obtained [54]. Several studies have suggested substituents of the backbone of the hemicelluloses, such as arabinose, galactose and 4-O-methylglucuronic acid, are the connecting links to lignin [61-63]. Possible links between hemicelluloses and lignins are demonstrated in Fig. 2. Lignin isolates from woody materials contain, beside carbohydrates, significant amounts of protein [64].

Fig. 2. Schematic representation of connections betweensoftwoodhemicellulosesand lignin [54]


55

1.2.4 Cell Wall Proteins

The plant cell wall contains, besides cellulose, hemicelluloses and lignins, structural proteins called extensins. Three kinds of cell wall proteins, which differ in amino acid composition, have been found in plants [65]. These are hydroxyproline-rich glycoproteins, glycine-rich proteins and proline-rich proteins [65, 66], all considered to be important structural components of plant cell walls. Recently Bao and co-workers [67] found an extensin-like protein in mature wood of loblolly pine (Pinus taeda L). These authors suggested that such structural proteins play important roles in the differentiation of xylem, and could thereby affect the properties of wood. However, the existence of lignin-protein complexes suggests a role for proteases in the degradation of woody and other plant materials, particularly in delignification processes such as pulp bleaching. Use of such enzymes might be beneficial for the improvement of pulp and paper manufacturing processes.

1.2.5 Other Cell Wall Components

Plant cell walls also contain extraneous materials, including extractives and non-extractives. Depending upon species, wood contains from 0.4 to 8.3% extractives on a dry weight basis, whereas agricultural residues may contain even greater amounts [68, 69]. The extractives can be broadly divided into three groups, i.e., terpenes, resins, and phenols. The terpenes can be regarded as isoprene polymers, with terpene alcohols and ketones as building blocks. The resins include a wide variety of non-volatile compounds, ineluding fatty acids, alcohols, resin acids, phytosterols, and some less known neutral compounds. The most important phenols are tannins, heartwood phenols, and related substances. In addition, low molecular weight carbohydrates, alkaloids, gums, and various other cytoplasmic constituents are present [70-73]. The non-extractives make up 0.2q18% of the dry weight and include inorganic components such as silica, carbonates, oxalates, and non-cell wall substances such as starch, pectin, and protein [69]. In agricultural residues, the non-extractives make up about 10% of the dry weight. Silica, deposited as crystals, is especially abundant in straw. In spite of the small quantity of extraneous materials their role is significant in that they provide an obstacle to the hydrolysis of cellulosic materials. Pectic substances (protopectin), though in small proportions, occur notably in the middle lamellae of the parenchymatous tissues, where they impart a structural function in binding and supporting cells.

56

R.C. Kuhad et al.

2 Microorganisms and their Enzymes 2.1 Microorganisms Involved in the Degradation o f Lignocellulosic Materials Wood and other lignocellulosic materials are degraded by a variety of fungi and bacteria. The structural architecture and chemical composition of wood play a significant role in its resistance to degradation by microorganisms.

2.1.1 Fungi Most of the fungi able to produce the necessary enzymes for the degradation of lignocellulosic materials belong to the Ascomycetes, Deuteromycetes, or Basidiomycetes groups. Fungi living on dead wood that preferentially degrade one or more of the wood components cause three types of wood rot, i.e., soft rot, brown rot and white rot [23, 74]. Important fungi causing soft-rot include Chaetomium cellulolyticum, Asper-

gillus niger, Trichoderma viride (reesei) , Fusarium oxysporum, Thielavia terrestris, Penicillium jenthillenum, Dactylomyces crustaceous and different species of Paecilomyces, Papulaspora, Monodictys, Allescheria, Hypoxylon, Xylaria, and Graphium [23, 75, 76]. They all efficiently attack wood carbohydrates but modify lignins only to a limited extent. A unique feature of soft-rot attack is the production of chains of biconical and cylindrical cavities within the secondary wall. This unusual microscopic pattern of decay in the $2 layer is characteristic of the soft-rot type of deterioration. Two patterns of soft-rot attack have been identified. One form of attack comprises cavities formed within the secondary cell wall, and the second form causes an erosion of the entire secondary wall originating from hyphae in cell lumina and progressing towards the middle lamella [77, 78]. The rate and extent of decay by soft-rot fungi depend on the type of wood they attack. In general, hardwoods are degraded to a greater extent than softwoods [79]. The ability of fungi to cause soft-rot attack of aromatic moieties of lignins has also been reported [79, 80]. However, they mainly cause demethylation of lignin and degrade the side chains and aromatic rings to a lesser extent. Fungi causing the brown-rot type of decay include Poria placenta, Tyromy-

ces balsemeus, Gloeophyllum trabeum, Lentinus lepidius, Lenzites trabeam, Coniophora puteana, Laetiporus sulphureus and Fomitopsis pinicola [23, 76, 81, 82]. These also exhibit preference for cellulose and hemicelluloses, lignins being degraded only to a limited extent. These fungi can cause rapid and extensive degradation of cellulose early in the decay process [23, 83]. The hyphae of brown-rot fungi are normally localized in the wood cell lumen and penetrate adjacent cells either through existing openings or by producing boreholes in wood cell walls. During the decay process, brown-rot fungi remove cell wall


57

substances first from the S 2 layer of the secondary wall, while the $3 layer, adjacent to the lumen, remains virtually unchanged. The $1 layer may also be attacked, but the primary wall and the middle lamella are very resistant because of their high lignin content [84]. Although hyphae are in direct contact with the S 3 layer, the ultrastructural changes are most obvious deep within the secondary wall. For a long time, involvement of brown-rot fungi in lignin degradation eluded clear demonstration. However, there is much evidence that brown-rot fungi can cause substantial degradation of lignins in wood [23, 85, 86]. The efficient demethylation of lignins or of simple lignin-related model compounds by G. trabeum has been reported. This fungus was also found to degrade [O14CH3]-labelled dimeric lignin model compounds in the presence of spruce sapwood relatively soon after inoculation compared to the white-rot fungus Phanerochaete chrysosporium [81]. In spite of the fact that brown-rot fungi primarily degrade wood carbohydrates, most of them are unable to degrade and utilize pure cellulose, particularly in submerged liquid cultures [87-89]. However, a few species, several of which are important degrades of forest products, do degrade pure cellulose [90] when in contact with inoculated pinewood blocks [87]. Lignified jute fibers were also found to be easily degraded by brown-rot fungi, whereas delignified ones were not [87]. Moreover, Highley [91] reported the degradation of cotton cellulose in contact with wood. It seems that the degradation of cellulose by brown-rot fungi is activated by the presence of lignin or lignin-like compounds [91]. It may be dependent upon prior or concomitant removal of the hemicellulose components [92, 93]. However, conclusive evidence does not yet exist that lignin activates the cellulolytic system of brown-rot fungi. Recently, the presence of spruce sapwood has been reported to greatly stimulate the demethoxylation of labelled lignin by brown-rot fungi [81]. This finding suggests that wood has some stimulatory effect on these fungi to utilize cellulose or lignin. A great deal of research seems to be necessary to establish the exact relationships. Brown-rot fungi differ substantially from white-rot fungi with respect to the cellulolytic enzymes produced and the pattern of cellulose degradation. The involvement of oxidative systems in the depolymerization of cellulose during the early stages of decay by brown rot has been demonstrated [88]. Considering that most of the pore sizes in sound wood are too small to allow cellulolytic enzymes to penetrate the wood, this might offer an advantage. A non-enzymatic hypothesis involving the generation of oxygen-derived radical species has been suggested to be involved in the initial attack on the cellulose polymer [94, 95]. However, this hypothesis has not been satisfactorily explored. White-rot fungi are the only wood-rotting fungi which, to any extent, can attack all the components of plant cell walls. The most studied fungi of this group are Phanerochaete chrysosporium, Trametes versicolor, Dichomitus squalens, Phlebia radiata, Heterobasidium annosum, Phellinus pini, C yathus stercoreus, Pleurotus ostreatus, Ceriporiopsis subvermispora, Polyporus anceps and Ustulina vulgaris [23, 96, 97]. Most of the fungi belong to Basidiomycetes except

58

R.C. Kuhad et al.

for Ustulina vulgaris, which is classified as an Ascomycete. The normal pattern of wood decay by these fungi involves simultaneous attack on both polysaccharides and lignin [23]. The ability of the white-rot group to efficiently remove lignin from wood makes them suitable for industrial applications where lignin or other phenolic compounds need to be modified or removed [23, 98]. Some white-rot fungi such as P. chrysosporium and T. versicolor have an unselective mode of wood degradation, i.e., they degrade cellulose, hemicellulose, and lignin simultaneously, while others such as Phlebia tremellosa, C. subvermispora and P. pini degrade the lignin component more selectively [99]. Some white-rot fungi have been reported to attack both selectively and non-selectively in different regions of the same piece of wood [100]. Several strains of P. chrysosporium and C. subvermispora degrade the different components of plant fiber cell walls with a different mode of attack and to a different extent [98]. The factors responsible for the different modes of degradation, i.e., a rather specific delignification or a simultaneous decay of lignin and carbohydrates, remain unexplained [23]. Dill and Kraepelin [101] suggested that environmental factors such as nitrogen concentrations in wood may be responsible in governing white-rot decay type. Also, white-rot fungi have a different production pattern with respect to the extracellular lignin-degrading enzymes - a trait likely to affect the degradation pattern [102]. White-rot fungi grow in all the different types of cells of hardwoods and softwoods. Fungal hyphae enter the cell lumen, first colonize the ray parenchyma cells, and then penetrate from cell to cell either via pits or by the development of boreholes directly through the cell walls [23, 84, 103]. Soon after the easily metabolizable nutrients are depleted, cell wall degradation starts. A separation between cells within or adjacent to the compound middle lamella has been reported both at early [103] and late [104] stages of wood decay. White-rot fungi that simultaneously attack cell wall components erode localized regions of cell wall layers, and these further extend through the secondary cell wall and middle lamella. The extensive erosion of cell walls and numerous holes in adjacent walls as later stages of decay have been observed in white-rot fungi that selectively degrade lignin. Lignin degradation by fungal hyphae in cell lumen progresses from the lumen edge of the secondary wall towards the middle lamella. Several reports suggest that white-rot fungi degrade lignin from the secondary wall before the middle lamellae between cells is degraded [105, 106]. The separation of cell occurs as a result of degradation of the middle lamella. It is well established that white-rot fungi often cause a progressive thinning of the cell walls, starting from the lumen and continuing towards the middle lamella [103, 107]. P. chrysosporium, when growing on aspen wood chips supplemented with nutrients, either produced boreholes or hyphae that entered and advanced through cell walls using natural wood pits [107, 108]. P. chrysosporium was frequently found to progress along the axis of vessels or fibers, with secreted enzymes etching the $3 and $2 layers [108, 109]. Likely, fungal decay


59

involves enzymatic softening and swelling of wood cell wall fibers as well as thinning and fragmentation of the wood cell walls. White-rot fungi produce an extracellular slime sheath around the hyphal cells which establishes close contact between hyphae and wood cell walls [110-112]. With the progress of the decay, the fibrillar slime materials become intrinsically associated with the swollen and delignified wood cell wall and form a connection between the fungal hyphae and wood substrates [113]. Several suggestions have been put forward for the roles of the fungal slime sheath. For example, studies have revealed the localization of lignin peroxidase and manganese peroxidase to be associated with the slime during decay of wood [113]. A more detailed investigation concerning the role of fungal slime would possibly contribute to the knowledge of mechanisms of wood attack by white-rot fungi. Recently, Akin et al. [72] studied the chemical and structural modifications of Bermuda grass (Cynodon dactylon) cell walls caused by the two white-rot fungi C. stercoreus and C. subvermispora. Both fungi were observed to extensively colonize the cut end of the stem section, disrupt the parenchyam cell walls, and partially degrade sclerenchyma cells. UV absorption microspectrophotometry indicated that ester-linked phenolic acids were totally removed from the parenchyma cell walls, and these cells were readily and completely degraded by both fungi. However, aromatic constituents were only partly removed from the recalcitrant sclerenchyma cell walls. For more extensive details about the various patterns of wood decay by white-rot fungi, readers are referred to Eriksson et al. [-23]. In addition to aerobic fungi, anaerobic fungi, inhabitants of the alimentary tract of herbivorous animals, play key roles in the degradation of plant cell wall material. Anaerobic fungi degrade the major structural polysaccharides, but cannot utilize the lignin moieties. Some anaerobic (rumen) fungi have the ability to solubilize small amounts of phenolic compounds [114-118]. These fungi colonize materials such as soybean hulls and grasses [119, 120]. Moreover, highly recalcitrant plant materials like mestome sheath of leaf blades [121] and palm press fibers and wood [-122] have been reported to be colonized by the anaerobic fungi. Among anaerobic fungi, the most studied are Neocallimastix frontalis, N. Patriciarum, Piromyces (Piromonas) communis, and Caecomyces (Sphaeromonas) communis [118, 123]. Electron microscopic studies have revealed that rumen fungi preferentially attach to specific regions of plant particles, i.e., the cut ends or stomata [124]. The fungal rhizoids readily invade the plant cell wall lumen and the middle lamellae between individual plant cell walls [14, 118]. Ho and co-workers [125] showed that some anaerobic fungi produced appressorial-like structures "penetration pet" which penetrated undamaged cell walls of guinea grass and rice straw. The fungal hyphae grow extensively and ramify throughout the interior of plant cell walls, which causes physical disruption of the cell wall [121]. Furthermore, Joblin [126] has also suggested the physical disruption of the plant fibers by Caecomyces spp. The ability of anaerobic fungi to disrupt plant tissues could well be attributed to their ability to produce an array of

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extracellular enzymes, i.e., cellulases, hemicellulases, proteases, amylases, phenolic acid esterases, and pectinases [120]. For more details about rumen fungi, the reader is referred to the excellent reviews by Weimer [,14], Wubah et al. [118] and Trinci et al. [120].

2.1.2 Bacteria Bacteria generally degrade wood slowly. Degradation takes place on wood surfaces with a high moisture content. Because of lack of penetrating ability, bacteria usually invade wood cells simultaneously with fungi. They appear to attack both softwoods and hardwoods by first colonizing the parenchyma cells. After utilizing the cell contents, they may also attack the parenchyma cell walls [73]. They move into adjacent cells and trachieds with fast disruption of pits [-127]. Although bacteria can directly attack fibers, vessels, and trachieds, few species or strains can degrade all the cell wall components [-23]. However, some bacteria have been found to degrade lignified wood cells, which was confirmed by ultrastructural investigations [128]. Cell wall erosion [129, 130], tunnel formation [131], and removal of lignin [132] by bacteria have also been reported. Different patterns of cell wall decay, cavitation, and tunneling by bacteria have been found both in natural and laboratory environments [131,133, 134]. In contrast, studies in some laboratories have shown that bacteria are unable to degrade lignified plant cell walls [135, 136], but they were able to do so after chemical pretreatment of the cells. Efficient bacterial degradation of wood already treated with cellulase-less mutants of P. chrysosporium and Phlebia 9igantea has been observed [137]. Rumen bacteria are major degraders of plant fiber cell walls by production of enzymes active against structural components of these cell walls [138]. Some of the most extensively studied rumen bacteria include Fibrobacter succinogenes, Ruminococcus albus, and R. flavifaciens. These bacteria have a complete set of polysaccharide-degrading enzymes and also the ability to adhere to fibers [138, 139]. These species adhere strongly to partially degraded cell walls, but erode the components only if they are adjacent and in direct contact with the bacteria. Often, the plant cell walls are totally degraded, but at other times digestion seems to be interrupted before the hydrolysis is completed. Some actinomycetes, the filamentous eubactria, also actively degrade lignocellulosic plant materials. Different species of Streptomyces have been reported to colonize vessels, fibers, and parenchyma cells [140]. S. flavovirens rapidly colonizes the phloem and degrades parenchyma cells as well as thickwalled, highly lignified sclereids [129]. In advanced stages of degradation, parenchyma cells were found to be completely destroyed, and sclereids showed evidence of eroded cell walls. Various studies have established that several actinomycetes attack grass lignocellulosics, leading to the partial solubilization of the substrate rather than to its mineralization [141]. Several species of


61

Streptomyces have the ability to degrade and remove lignin from softwood, hardwood, and graminaceous substrates [142, 143]. Streptomyces viridosporus oxidatively depolymerizes lignin as it degrades cellulose and hemicellulose components of plant residues [144]. Lignin degradation by actinomycetes is reviewed by Zimmerman [128], and the varying abilities of different species of Streptomyces to delignify lignocellulosics is discussed in detail.

2.2 Enzymes Involved in the Degradation of Plant Fiber Cell Wall Components Because of the complex nature of wood and other plant cell walls, a number of different enzymes are required to degrade the wall components. The generally accepted picture of enzymatic degradation of cellulose is that it proceeds by the synergistic action of at least three major types of hydrolytic enzymes. In addition, oxidative and phosphorolytic enzymes also participate in cellulose degradation in some organisms. The hydrolytic enzymes (Fig. 3) involved in degradation of native cellulose to glucose are: (1) endoglucanases (endo-l,4-[3D-glucan-4-glucanohydrolase, EC 3.2.1.4), which randomly attack the cellulose chains and split 1,4-[~-glycosidic linkages, (2) exoglucanases, generally cellobiohydrolases (exo-l,4-[3-D-glucan-4-cellobiohydrolase, EC 3.2.1.91), which release either cellobiose or, in some cases, glucose from the non-reducing end of cellulose, and (3) 1,4-[3-glucosidases (EC 3.2.1.21), which hydrolyze cellobiose and other water-soluble cellodextrins to glucose [23]. Two types of oxidative enzymes, cellobiose oxidase (CBO), now called cellobiose dehydrogenase (CDH) (EC 1.1.9.18), and cellobiose: quinone oxidoreductase (CBQ) (EC 1.1.5.1), which oxidize the reducing end group in cellobiose or higher cellodextrins in the presence of a suitable electron acceptor, have been identified in many wooddegrading fungi and may have a key role in cellulose degradation [23]. In addition, lactonase (EC 3.1.1.17) has been found to operate synergistically with cellulases in the degradation of cellulose [145]. Some aerobic and anaerobic bacteria lack [~-glucosidases and produce cellobiose phosphorylase (EC 2.4.1.20), which cleaves cellobiose but not cellotriose or higher cellodextrins [146, 147]. Because of the complex structure of hemiceUuloses, several different enzymes are needed for their enzymatic degradation. The two main enzymes responsible for the depolymerization of hemicellulose backbone are endo-l,4-[3-D-xylanase (EC 3.2.1.8) and endo-l,4-13-D-mannanase (EC 3.2.1.78). While small oligosaccharides are hydrolyzed by 1,4-]3-D-xylosidase, 1,4-[3-D-mannosidase, and 1,4-[3m-glucosidase, the side groups are split off by ~t-L-arabinosidase, a-Dglucuronidase and cx-D-galactosidase. Esterified side groups are released by acetylxylan esterase [23, 148-150]. Since the process of biodegradation of lignin by white-rot fungi is oxidative in nature, the role of phenoloxidases in lignin degradation has been extensively studied [23]. At least three phenoloxidases (Fig. 3) have been identified as important in the ligninolytic enzyme system, i.e., laccase [151,152],

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R.C. Kuhad et al.

~__CH3iH MeOXi= HCOH+ H202 ~ I Glucose~ Glu II Ox Araoino-2-hexulose . . I '~ I .r }, I ',

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Fig. 3. Extra- and intracellular enzyme mechanisms involved in degradation oflignin and cellulose by the white-rot fungus Phanerochaete chrysosporium. Enzymes involved: (1) glyoxal oxidase, (2) lignin peroxidase, (3) manganese peroxidase, (4) laccase, (5) cellobiose dehydrogenase, (6) cellobiose: quinone oxidoreductase, (7) Protease, (8) Lactonase, (9) endo-l,4-~-glucanase, (10) exo-l,4-~glucanase, (11) 1,4-[~-glucosidase,(12) glucose-l-oxidase, (13) catalase. Meox methanol oxidase, Glu II Ox glucose-2-oxidase, VA veratryl alcohol. Vanillic acid metabolism: VH vanillate hydroxylase, Ph.Ox phenol oxidase, Q-red NAD(P)H: quinone oxidoreductase, Diox dioxygenase, M-red maleylacetate reductase. Metabolic products from lignin degradation: A vanillin, B vanillic acid, C methoxyhydroquinone (MHQ), D hypothetical ortho-quinone (II), E hydroxyquinol, F maleyl acetate, G Jbketoadipate

Microorganisms and Enzymes Involved in the Degradation of Plant Fiber Cell Walls

63

lignin peroxidase (LIP), and manganese-dependent peroxidase (MnP) [153-155]. Both peroxidases belong to the class of horseradish peroxidases (EC 1.11.1.7) and oxidize their substrate by two consecutive one-electron oxidation steps with intermediate cation radical formation. LiP, because of its high redox potential, can attack non-phenolic methoxyl-substituted lignin subunits, whereas MnP acts exclusively as a phenol oxidase on phenolic substrates using Mn 2 § and Mn 3 § as an intermediate redox couple [156]. Laccase (EC 1.10.3.2), a true phenoloxidase, has a broad substrate specificity. It oxidizes phenols and phenolic lignin substructures with formation of radicals. The enzyme can cause both polymerization and depolymerization by production of phenoxy radicals [157]. However, laccase is generally considered to have a redox potential too low to allow for oxidation of non-phenolic lignin structures. Other enzymes of importance in lignin degradation are the H2Oz-producing ones, (Fig. 3). Such enzymes are glucose-l-oxidase and glucose-2-oxidase [158, 159]. Other oxidases producing HzO2 have been detected in P. chrysosporium, namely methanol oxidase and glyoxal oxidase [160, 161]. The oxidative action of methanol oxidase yields formaldehyde. Glyoxal oxidase is produced extracellularly and uses several aldehydes and hydroxycarbonyl compounds found as secondary metabolites in the fungal culture filtrate [161]. In addition, the enzymes reducing phenoxy radicals may also be involved in lignin degradation processes [1]. Such enzymes are CDH, CBQ, and NAD(P) H: quinone oxidoreductase [1,162]. The novel extracellular enzymes capable of releasing aromatics (feruloyl and p-coumaroyl groups) from isolated grass cell walls and from their methyl esters are feruloyl and p-coumaroyl esterases [163, 164]. An esterase which unlocks p-coumaroyl groups from lignocellulose has been reported from Streptomyces viridosporus T7A [165], while feruloyl esterase has been detected in cultures of Schizophyllum commune [-166], F. succinogenes [167, 168], and Butyrivibrio fibrisolvens [167]. Both feruloyl and p-coumaroyl esterases have been reported to be produced by anaerobic fungi [163, 164]. The effective attack and penetration of lignified tissues, although without mineralization of lignin per se, by the anaerobic fungi strongly suggest that phenolic acid esterases play an important role in the biodegradation of realcitrant cell walls in grasses [169]. Use of these enzymes, in combination with specific hemicellulolytic enzymes, may offer a strategy to remove aromatics and specific sugars from lignocellulosics [170].

3 Degradation of Cellulose 3.1 Microorganisms Producing Cellulose-Degrading Enzymes A large number of microorganisms have the ability to produce cellulose-degrading enzymes (Table 2). However, relatively few are capable of producing all the

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Table 2. Important microorganisms with respect to cellulolytic potential Soft-rot fungi Aspergillus niger Chaetomium cellulolyticum F usariurn ox ysporum Neurospora crassa Penicillium pinophilum Trichoderma reesei

Aerobic bacteria Bacillus circulans Bacillus subtilis Cellulomonas fimi Cellvibrio gilvus Microbispora bispora Pseudomonas fluorescens

Brown-rot fungi Coniophora puteana Lanzites trabeum Poria placenta Tyromyces palustris

Anaerobic bacteria Acetovibrio cellulolyticus Clostridium cellulovorans Clostridium thermocellum

White-rot fungi Phanerochaete chrysosporium Sporotrichum thermophile Coriolus versicolor

Rumen (anaerobic) bacteria Butyrivibrio fibrisolvens Fibrobacter succinogenes Ruminococcus albus

Rumen (anaerobic) fungi Caecomyces (Sphaeromonas) communis Neocallimastix frontalis Neocallimastix patriciarum Piromyces communis

Actinomycetes Streptomyces lividans rhermoactinomyces curvata Thermomonospora fusca

necessary enzymes for degradation of crystalline cellulose [171]. While bacteria degrade cellulose by cell-bound enzymes, fungi secrete most of these enzymes into the surrounding growth medium [172, 173]. All organisms able to degrade crystalline cellulose secrete more or less complex systems of cellulolytic enzymes with different specificities and modes of action, which act in synergism to hydrolyze or oxidize cellulose. Fungi are the most studied organisms with respect to degradation of cellulose and production of cellulolytic enzymes. Fungal cellulases are sometimes produced in very high concentrations in submerged cultures and do not seem to form physical complexes with each other as do bacterial cellulases. However, fungal cellulases often act in a strongly synergistic manner [174-180]. Fungi causing soft-rot mainly degrade the polysaccharides. The ability to produce all the necessary enzymes for degradation of cellulose varies among the different organisms in this group. The best known of those producing a complete set of cellulases is T. viride (reesei) [181, 182]. Trichoderma strains secrete large amounts of different cellulolytic enzymes which, by their combined action, degrade crystalline cellulose. T. reesei produces at least three endoglucanases, two exoglucanases, and one or two 13-glucosidases [183, 184]. Other well known fungi causing soft-rot are listed in Table 2. Fungi causing brown-rot degrade cellulose rapidly, but the enzyme system seems to operate differently from those of T. viride (reesei) and the white-rot fungus P. chrysosporium. P. placenta, L. trabeum, and Tyromyces palustris are brown-rot fungi among the most studied for their cellulolytic activities [23].


65

Brown-rot fungi have been suggested to employ a different mechanism of cellulose degradation than that operating in soft-rot and white-rot fungi. Brown-rot fungi depolymerize cellulose rapidly during the early decay of wood, whereas white-rot fungi do so more slowly and utilize the degradation products simultaneously [23,185]. Brown-rot fungi produce endoglucanases and ]3-glucosidases but no detectable levels of exoglucanases [23, 91]. In contrast to these reports, C. puteana (a wide spread brown-rot fungus causing severe damage to buildings) has recently been reported to secrete two exoglucanases (cellobiohydrolases) when growing on pure cellulose [186, 187]. The presence of an exoglucanase in a brown-rot fungus is certainly intriguing. These workers have further reported that C. puteana gives a positive Bavendamm reaction [188], a feature used to distinguish between brown-rot and white-rot fungi. According to this definition, C. puteana is not a brown-rot fungus. The induction and repression mechanisms of cellulases differ remarkably in brown-rot and white-rot fungi. While the brown-rot fungi produce ample amounts ofcellulases on glucose as carbon source [189], this is not the case with white-rot fungi, at least not with P. chrysosporium, whose cellulase production is induced by cellobiose and repressed even by low concentrations of glucose [190]. Since brown-rot fungi apparently lack exoglucanases, there is no synergistic cooperation between endo- and exoglucanases to degrade crystalline cellulose [23, 91, 191]. No other enzyme systems are known to substitute for these effects [23]. The mechanisms of cellulose degradation by brown-rot fungi remain a mystery and need to be clarified. However, one hypothesis includes the involvement of the Fenton reagent mechanism, i.e. H 2 0 2 and Fe 2 +, as critical for predisposing cellulose to attack by cellulolytic enzymes during early degradation [74, 192]. Contrary to earlier reports [191,192] on the accumulation of H202 in culture media, Veness and Evans [193] could not detect extracellular H202 in cultures of several brown-rot fungi. However, Fenton's reagent was observed to give rise to acid production from cellulose. The same acids were formed by a crude enzyme mixture from T. reesei. Recently brown-rot fungi have been observed to secrete low molecular mass glycoproteins which chelate ferrous ions [194]. It was proposed that these iron-containing glycoproteins are small enough to penetrate the $2 and Ss layers of wood cell walls to catalyze the oxidation of electron donors by molecular oxygen to produce oxygen species such as 02" and H202. According to the hypothesis, all reactants required for cell wall degradation by a Fenton-like reaction would be present simultaneously in the Sz layer. More recently, Backa et al. [195] have reported the generation of hydroxyl radicals by brown-rot fungi grown in liquid media, agar media, or wood. Their observations have indicated that hydroxyl radicals (HO.) contribute significantly to the initial degradation of wood. The hydroxyl radical is one of the few agents that can disrupt the crystalline structure of cellulose. In biological systems, most hydroxyl radicals are produced by Fenton reactions [196]. Kirk et al. [197] have reported that Fenton's reaction causes a cellulose degradation

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similar to that resulting from brown-rot decay. Other non-enzymatic agents such as siderophores [198] and oxalic acid [199] have been reported to be involved during degradation of wood by brown-rot fungi. The mechanism by which these agents work along with the enzymes in situ to degrade cellulose is not known. The brown-rot fungi form extensive hyphal sheaths, which appear to provide a functional interface between the fungal hyphae and the wood [200-202]. Degradation of wood by brown-rot fungi has been reported to occur at a distance from the fungal hyphae and to be characterized by early removal of the hemicellulose components. The following functions have been assigned to the extracellular sheath: (1) modification of the extracellular ionic environment and pH-level, (2) recognition of and adhesion to the substrate, (3) concentration, storage, and transport of decay agents, (4) protection against dehydration and other environmental impacts, (5) conditioning of the wood substrate to degradation, (6) storage of nutrients, (7) regulation of the decay process, (8) increase of the surface area for aerobic respiration, and transportation and presentation of wood-degrading enzymes during the degradation process [201,203-205]. Green et al. [-205] have revealed the localization of enzymes on fibrillar elements of the sheath structure, on the hyphal surface, and within the soluble sheath matrix. The group of fungi causing white-rot is rather heterogenous, but these fungi have in common the ability to degrade lignin as well as other lignocellulosic components [206]. The most studied white-rot fungus was first isolated from wood chip piles and was originally given the name Chrysosporium lignorum [207]. The name was later changed to Sporotrichum pulverulentum for its imperfect stage [208] and then to Phanerochaete chrysosporium for its perfect stage [209]. The fungus grows optimally at 38-39 ~ and hence is characterized as thermotolerant. At least five endoglucanases, three exoglucanases, and two [3-glucosidases have been characterized in cultures of P. chrysosporium [210-213]. The anaerobic rumen-inhabiting fungi secrete cellulases, often described as a complex of enzymes which, by acting together, solubilize cellulose efficiently [214]. An extracellular enzyme system from Neocallimastixfrontalis was found to have a several-fold higher specific activity in solubilizing cotton fibers per unit of endo-I~-glucanase activity than the endoglucanase of T. reesei mutant, Rut C-30, known to produce a very active cellulolytic system [215]. The major end product of the N.frontalis-cellulases, acting on Avicel cellulose, was glucose and not cellobiose, which also indicates an efficient 13-glucosidase activity [216]. Wilson and Wood [217] showed that a minor component (4% of the total extracellular protein) of the cellulolytic enzymes of N. frontalis is present in a cellulosome-like structure which is secreted and adsorbed on to cellulose. In this respect, N.frontalis resembles the bacterium Clostridium thermocellum. The component which degrades crystalline cellulose contains several enzymes arranged in a complex called the cellulosome, although the cellulosome of N.frontalis (700 kDa) is smaller than that of C. thermocellum (2 • 106 kDa) and also differs in other ways [120]. Recently, Wood and Wilson [218] reported on


67

Piromyces communis, another anaerobic rumen fungus capable of producing a highly active cellulase system. This enzyme system was capable of solubilizing highly crystalline cotton fibers at a rate greater than that of any other cell-free cellulase system reported in the literature. Cellulose is also degraded by both aerobic and anaerobic bacteria. However, the cellulolytic enzyme systems of bacteria are not directly comparable to those of fungi. Bacteria often produce cellulases in small amounts ( < 0.1 g/L), and degradation of cellulose seems to take place by a cluster of multienzyme complexes which are difficult to disrupt without loss of total activity as well as of the individual components [219]. The most studied bacteria with respect to their cellulase systems are species of Clostridium, Cellulomonas, Bacillus and Pseudomonas. The cellulase complex of C. thermocellum cellulosome, consists of several endoglucanases, an exoglucanase, a cellobiose phosphorylase that breaks down cellobiose to glucose and glucose-l-phosphate, a cellodextrin phosphorylase which phosphorolyzes 13-1,4-oligoglucans, and two 13-glucosidases [220]. The cellulosome produced by C. thermocellum and other Clostridia is a cellulose-binding and cellulase-containing complex, which is responsible for most of the cellulase and hemicellulase activities observed in culture broth of such bacteria [219]. During exponential growth on cellulose, the cellulosome is anchored to the bacterial cell surface, and anchors the cells to the insoluble substrate. When the cellulose is solubilized, the bound enzyme clusters are released into the culture medium [221]. Other bacteria which may produce cellulosome-like entities similar to those produced by C. thermocellum include Acetovibrio cellulolyticus, Bacterioides cellulosolvens, Butyrivibrio fibrisolvens, Cellulomonas sp., Clostridium cellobioparum, C. papyrosolven, Fibriobacter succinogenes, Ruminococcus albus, R. flavefaciens, and Thermomono spora curvata [222]. The extracellular cellulase systems of Pseudomonas fluorescens and Cellvibrio gilvus are also similar to those mentioned above, and degradation products of enzymatic action are mainly cellobiose and cellotriose [223]. Filamentous prokaryotes such as Actinomycetes are also important members of the microbial community responsible for the degradation of cellulose. Mesophilic species of Streptomyces and thermophilic species of Thermomonospora and Thermoactinomyces are the most studied of these organisms [224, 225]. Their extracellular cellulases degrade cellulose by mechanisms similar to those of fungi, i.e., non-associated enzymes excreted into the culture medium.

3.2 Cellulolytic E n z y m e s 3.2.1 Regulation of Cellulase Production The production of cellulases appears to be controlled by induction and repression mechanisms. In most microorganisms, cellulase biosynthesis is induced by

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cellulose degradation products [21]. The endoglucanases have been the most studied in this respect. P. chrysosporium [190] and G. trabeum [226] have been shown to form endoglucanases in media containing only carboxymethylcellulose (CMC), while this was not the case with T. reesei. Cellobiose is an inducer of endoglucanases in both P. Chrysosporium [190] and G. trabeum [226], but does not induce any endoglucanase activity in T. reesei [190]. In contrast, sophorose is the most potent cellulase inducer in T. reesei [227, 228]. However, cellobiose has been shown to induce cellulases in T. reesei, but only when cellobiose hydrolysis is artificially decreased by addition of nojirimycin [229]. Other cellulose degradation products such as cellobiono-l,5-1actone 1-230] or oxidized cellulose [231] have also been demonstrated to enhance the formation of cellulases in T. reesei. Glucose is the end product of cellulose hydrolysis and causes catabolite repression of endoglucanases in P. chrysosporium [190] and T. reesei [232]. In contrast, the brown-rot fungi P. placenta and G. trabeum produce endoglucanases with glucose or mannose as the sole carbon source, even if the expression of these enzymes was four to five times as high with cellulose or cellobiose as carbon sources [88,226]. The addition of glucose to induced cultures of G. trabeum to a concentration of 40 mM or higher does not repress the production of endoglucanases [226]. It appears from various studies that the regulation of cellulase synthesis in anaerobic fungi also involves both induction by cellulose and catabolite repression by glucose [233-237]. However, considering the low concentration of free glucose in the rumen, it is unlikely that glucose regulates cellulase synthesis in vivo [14]. Production of cellulases in fungi is also regulated by phenomena other than induction and catabolite repression by cellulose degradation products. Various phenols have thus been demonstrated to repress the production of cellulases and xylanases in S. commune and C. 91obosum. In addition, phenols repressed endoglucanase in phenoloxidase-less mutants of P. chrysosporium, but caused no significant repression in phenoloxidase-less revertants and in the wild type [147]. Moreover, cellulose activity in culture filtrates of fungi is dependent not only on regulation of cellulase biosynthesis but also on the presence of specific inhibitors in the culture. Gluconolactone is a powerful inhibitor of [3-glucosidase in P. chrysosporium and T. reesei [-211,238]. Inhibition of ~-glucosidase activity by gluconolactone or nojirimycin prevented induction by cellulose, but not by sophorose [238]. However, sophorose fails to induce all components of the cellulolytic system, indicating that other degradation products of cellulose might contribute to regulation of this system [239]. The true inducer of cellulase expression, in T. reesei as well as enzyme(s) responsible for its formation, remains unexplained. However, not only inhibition but also activation of cellulases seems to occur in culture solutions of fungi. Thus, two acidic proteases produced under cellulolytic conditions have been demonstrated to enhance endoglucanase activity (Fig. 3) in P. chrysosporium up to tenfold [23].


69

Induction of cellulase biosynthesis by cellulose degradation products is likely to occur in actinomycetes and bacteria as well. Addition of cellulases exogenously to a cellulose culture medium of Thermomonosporafusca prompted the induction of cellulases [240]. Cellobiose and sophorose have both been shown to act as inducers of cellulase synthesis in Cellulomonas mutants insensitive to catabolite repression [241]. Recently, Weimer [14] has discussed the regulation of the cellulase systems in rumen bacteria. The complete cellulase complexes in these bacteria do not seem to be highly regulated by the carbohydrate concentration. However, the regulation of individual proteins must not be ignored. The cellulase synthesis does not appear to be induced by cellulose and cellobiose, and glucose neither represses enzyme synthesis nor inhibits enzyme activity, except at concentrations well above those encountered in vivo [242, 243].

3.2.2 Molecular Properties

Most of the fungal and some of the bacterial cellulases are glycoproteins with sugars attached to asparagine (N-linked) or serine and threonine (O-linked) residues. The carbohydrate content of cellulases varies from 1 to about 10%, and the principal sugar is mannose. However, other sugars such as glucose, galactose, xylose, N-acetyl glucosamine, and galactosamine have also been detected [244]. Nitrogen-linked glycosylation appears to impart a specific conformation and stabilize the structure of the cellulases, thereby protecting them from proteolytic attack during secretion while O-linked glycosylation seems to be required for secretion of an active cellulase [-245]. Endoglucanases hydrolyze ]3-1,4-glucosidic bonds in a random fashion over a cellulose chain. As a result, there is a rapid decrease in chain length and a slow increase in reducing end groups. Three different endoglucanases, i.e EG I, EG II, and EGIII are produced by T. reesei [239, 246]. The major EG component, EG I, represents approximately 5-10% of the totally secreted proteins in T. reesei cultures [247]. In spite of clear differences in their mechanism of actions, the N-terminal sequences indicate considerable homology between EG I and CBH I [-248]. Multiple forms of EG III have been purified exhibiting Mr values of 48, 48 and 37 kDa with pIs corresponding to 5.4, 5.7, and 4.8 [-184]. A number of other endoglucanases have also been isolated from T. reesei cultures which could not be identified as either EG I or E G I I I [249]. Cellobiohydrolases (exoglucanase, EC 3.2.1.91) degrade cellulose by splitting off cellobiose from the non-reducing end of the chain. These enzymes have very limited action on substituted cellulose such as carboxymethyl cellulose (CMC) and hydroxyethyl cellulose (HEC). Heterogeneities of the cellobiohydrolase components from T. reesei have been studied in detail, and two immunologically distinct cellobiohydrolases, CBH I and CBH II, have been detected in the extracellular medium of T. reesei using polyclonal antibodies [-250]. Both are glycoproteins differing in the amount of carbohydrates attached to the protein.

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They lack any apparent homology in their amino acid sequences. The differences in the active centers of these enzymes are reflected by differences in their mode of action. While CBH I preferentially binds to crystalline regions, CBH II binds to both crystalline and amorphous regions [251]. CBH I comprises the major part (ca. 60%) of the cellulolytic enzymes synthesized by T. ressei [252]. ~-Glucosidases, secreted by cellulolytic organisms, represent a small portion of the total extracellular proteins. They catalyze the hydrolysis of water-soluble cellodextrins as well as alkyl- and aryl-13-D-glucosides. Intracellular and plasma membrane-bound 13-glucosidases have also been identified. However, the exact genetic and biochemical relationships between these different [~-glucosidases are not yet clearly understood. It has been suggested that extracellular enzymes may result from the release of intracellular or membrane-bound enzymes to the outside medium upon autolysis [253]. Species of Aspergillus and Phanerochaete seem to produce 13-glucosidases of higher molecular weights than does T. reesei [23]. From Sporotrichum thermophile, two distinctly different [3-glucosidases were isolated [254]. One with a Mr of 440 kDa had only aryl-[3-glucosidases activity, while the other, with a Mr of 40 kDa, had cellobiase activity and only low activity toward aryl-[3-glucosides. Purified [3-glucosidase from C. 9ilvus attacks cellobiose slowly as compared to its activity towards higher oligosaccharides [255]. Cellobiose dehydrogenase (CDH), previously termed cellobiose oxidase (CBO) [256-258], is produced by several cellulolytic fungi (soft-rots, white-rots, and one brown-rot, i.e., C. puteana). Moreover, recently, CDH production from a cellulolytic bacterium Cyotophaga sp. LX-7 has also been reported [259]. CDH carries both FAD and heme as prosthetic groups. Various investigations have revealed that a proteolytic cleavage product of CDH, previously known as cellobiose: quinone oxidoreductase (CBQ) [260], represents the flavin-containing domain of CDH [261-263]. Both CDH and CBQ, in the presence of an appropriate electron acceptor, oxidize the reducing end groups of cellobiose, higher cellodextrins, and even cellulose to the respective onic acids via the corresponding lactone [257, 258]. Both CDH and CBQ are known to utilize quinones and their analogs, such as 2,6-dichlorophenol-indophenol (DCPIP), as electron acceptors. Cytochrome c acts as an electron acceptor for CDH but not for CBQ [264, 265, 257]. This ability of CDH is used to differentiate between CDH and CBQ. The total activities of both CDH and CBQ can be determined using either ferricyanide DCPIP or other reducible quinones as electron acceptors. CDH is a monomeric protein in which the heme group has been shown to be of the cytochrome b type [266]. The enzyme reduces cytochrome c about 200 times as fast as it reduces oxygen [264], whereas Fe(III) compounds such as Fe(III) acetate and Fe(CN)~ are reduced 35-50 times as fast as molecular oxygen [267]. CDH could be split into the FAD and the heme domains by papain [261] and by proteases from P. chrysosporium [263], although, in the latter case, only when the CDH was bound to cellulose. Only the FAD domain


71

was reduced by the addition of cellobiose [261]. This indicates that the FAD domain is directly responsible for the oxidation of cellobiose and that the electrons must then be transferred from the reduced FAD to an appropriate acceptor, i.e the heme domain, quinone, Fe(III), or even molecular oxygen. Small-angle X-ray scattering studies have suggested that CDH as well as its fragments appear to be of prolate shape, and that the cross-section of the FAD (4.3-5.1 nm) is considerably larger than that of the heine fragment (3.3 nm) [268]. The observations further suggested a co-linear arrangement of the two domains of CDH. Thus, according to their model, CDH appears very much to be a linear particle. However, the possibility that CDH may have a bent shape cannot be ruled out. Based on the research conducted with these oxidative enzymes, some possible physiological roles of these in cellulose and lignin degradation have been suggested [257]: (1) oxidation of cellulose and introduction of carboxyl groups, resulting in a distortion of the crystalline structure of cellulose, (2) conversion of cellobiose, which inhibits hydrolytic enzymes, into cellobionic acid, (3) oxidation of reducing end groups in cellulose to prevent reformation of glycosidic bonds broken by cellulases, (4) direct energy uptake by coupling with electron transfer chains on the fungal cell wall, (5) generation of active species (superoxide anion radicals) to disrupt the crystalline structure of cellulose, thus facilitating hydrolysis of cellulose, and (6) reduction of phenoxy radicals to prevent repolymerization of lignin degradation products. Interestingly, CDH has also been demonstrated to act as a Fenton's reagent to generate hydroxyl ('OH) radicals (Fig. 3) in the presence of Fe(III) and H202 [267, 269]. The hydroxyl radicals are strong enough oxidants to attack not only cellulose but also other plant cell wall components in a non-specific manner [162]. However, a physiological operation of such a mechanism requires some special extracellular compartmentation arrangements since the production of these radicals could also be harmful to the fungus [270]. Bacterial cellulases seem to be quite a complex mixture. In Cellulomonasfimi cultures, up to 10 components with CMCase (endoglucanase) activity have been identified [271]. The glycosylated components bind strongly to Avicel and are stabilized by this substrate, whereas enzymes free in solution are unstable and are altered by proteolysis and deglycosylation [272]. The endoglucanase of B. subtilis exhibits almost twice the specific activity of the T. reesei endoglucanase [273]. Recently, a bifunctional cellulase was isolated from Bacillus sp. D 04 [274]. The enzyme was a single polypeptide (Mr 35 kDa) with both endo- and exoglucanase activities, each activity existing at a separate site. One cell-bound and two extracellular (A and B) endoglucanase components of P.fluorescens var. cellulosa have been identified [275]. All three of these enzymes hydrolyzed a variety of substrates, and the mode of action toward CMC substrates appears to be similar to that shown by fungal endoglucanases. The major endoglucanase ( > 80% of total endoglucanase activity) of T. curvata exhibits its highest activity on CMC with a high degree of polymerization [276]. Glutamic and aspartic acids constitute 24% of the total amino acid composition.

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Bacterial cellulase systems differ from fungal cellulase systems by forming aggregates of multienzymes (cellulosomes) rather than existing as individual extracellular enzymes [222]. Cellulosomes are known to be produced by several anaerobic cellulolytic bacteria, Clostridia in particular. The cellulase system of C. thermocellum is very active against crystalline cellulose, with a specific activity higher than that observed for the T. reesei cellulase system [277]. The cellulosomes have molecular masses ranging from 2 x 106 to 6.5 x 1 0 6 Da, diameters of about 18 nm, and contain 14 to 26 different polypeptides ranging in size from 37 to 210 kDa [219, 221,278]. Cellulosomes form larger complexes, polycellulosomes, with masses from 50 x 1 0 6 to 80 • 1 0 6 Da [279]. Protuberances covering the surface of the bacterial cells are packed with polycellulosomes, each of which seems to contain several hundred cellulosomes [280]. The cellulosome efficiently hydrolyzes both amorphous and crystalline cellulose, whereas individual peptides alone or as a mixture do not [281]. Many of the polypeptides in the cellulosome complex are catalytically active and can be characterized as endoglucanases, xylanases, and cellodextrinases. The occurrence of cellobiohydrolase in cellulosomes has been suggested [282, 283], but its presence still needs reconfirmation [281]. In addition, 13-glucosidase, [3-xylosidase, J3-galactosidase, and [3-mannosidase activities have also been observed in cellulosomes [284]. Several polypeptides in cellulosomes have been sequenced, including the largest (210-250 kDa), a cellulosome-integrating protein previously termed S1 or SL and now designated CelL or CipA [285]. CipA may act as a cellulosebinding factor and/or a scaffolding protein anchoring the various catalytic subunits of the cellulosome, which, in addition, binds to the cellulosic substrate and also anchors the complex to the bacterial cell surface [281,286]. Another important subunit from the cellulosome has been identified as CelS (82 kDa), termed the catalytic unit [286]. CipA and CelS are thus two of the components of the cellulosome of C. thermocellum [287]. An "anchor-enzyme model" proposed to explain the cooperative action between these two subunits [-288] was confirmed by DNA sequencing of the genes encoding these two proteins [289]. Based on these observations, the model was recently expanded into a more sophisticated one [290]. It is now evident that CipA provides the anchor for the other catalytic components and that CelS provides the cellobiohydralase or exoglucanase activity necessary for crystalline cellulose degradation. For details about the mechanism of cellulose degradation according to the anchor-enzyme model, readers are referred to Wu [-290] and Wang et al. [286]. Also, an excellent review on the cellulosome of the anaerobic thermophilic C. thermocellum has recently been published by Felix and Ljungdahl [281]. The cellulosomes in other Clostridia such as C. cellulovorans, C. cellulolyticum and C. jsui appear to have properties very similar to those of the C. thermocellum cellulosome [291]. The cellulosome of C. cellulovorans consists of three major subunits called CbpA, P100, and P70 and of at least six different enzyme subunits [292].


73

Some aerobic and anaerobic bacteria employ phosphorylases for cellulose degradation [-147]. Cellobiose and higher cellodextrins are metabolized by C. 9ilvus, and the cell yield per glucose equivalent increases with the degree of polymerization (DP) of the cellodextrins. This suggests that cleavage of glucosidic bonds is brought about by phosphorylation and thus conservation of energy of these bonds [23]. The physiological role of cellobiose phosphorylase (EC 2.4.1.20) is to convert cellobiose into glucose-l-phosphate, which is utilized more efficiently than glucose by these microorganisms. The presence of cellobiose phosphorylase was first demonstrated in C. thermocellum 1-293]. Two types of enzymes have been identified, namely, those specific for cellobiose and those utilizing higher cellodextrins for the formation of a-glucosyl phosphate [146, 294]. The major products of enzymatic action are glucose-l-phosphate and glucose. The purified cellobiose phosphorylase from C. 9ilvus was found to consist of four subunits and to have specificity for cellobiose, requiring Pi and Mg 2+ for phosphorylation [146].

3.2.3 CelluIases are Organized in Domains

Proteolytic cleavage of cellulases into separate catalytic and cellulose-binding regions demonstrated the presence of these two domains within these enzymes 1-295]. On the basis of existing amino acid sequence identities in their putative catalytic domains, fungal and bacterial cellulases have been classified into families of homologous proteins. This classification is confirmed by hydrophobic cluster analysis [296, 297]. For details of classification based on catalytic domains, their occurrence and properties, readers are referred to the excellent reviews available on this aspect [298, 299]. In brief, family A includes cellulases from a diverse range of organisms including Gram-negative and Gram-positive bacteria, aerobic and anaerobic bacteria, and the fungus T. reesei (Table 3). Besides typical patterns of hydrophobic amino acids, some of the conserved regions contain residues that are identical in most of the sequences. In family B, three clearly related endoglucanases from C. fimi, Streptomyces sp. KSM-9 and Microbispora bispora, and CBH II from T. reesei sharing significant similarity are included (Table 3). Family C comprises three cellulases of fungal origin, exhibiting at least 45% similarity (to each other). Among three bacterial cellulases in family D, Erwinia ehrysenthemi EGY and C. uda EG are more closely related. Families E, F, and G are also of bacterial origin, with most of the enzymes from F and G having xylanase activity. Out of nine families, A, B, F, and H contain both fungal and bacterial enzymes. Family C contains only fungal enzymes, and families D, G and I contain only bacterial enzymes. Family E contains both bacterial and plant enzymes (Table 3). Cellulose-binding domains (CBDs) form distinct functional units for most cellulolytic enzymes. Although these CBDs are not essential for catalytic activity, they do modulate the specific activities of the enzymes on soluble and insoluble cellulosic substrates. Table 4 summarizes the different CBDs identified

R.C. Kuhad et al.

74 Table 3. Cellulase and xylanase families of homologous catalytic domains Family organism

Enzyme

Terminus

Reference

A Bacillus sp. 1139 Bacillus sp. N-4 Bacillus sp. N-4 Bacillus sp. N-4 Bacillus polymyxa Bacteroides ruminicola Butyrivibriofibrisolvens Caldocellum saccharolyticum Clostridium acetobutylicum Clostridium cellulolyticum Clostridium thermocellum Clostridium thermocellum Clostridium thermocellum Clostridium thermoceUum Erwinia chrysenthemi Fibrobacter succinogenes Ruminococcus albus SY 3 Ruminococcus albus SY 3 Trichoderma reesei Xanthomonas campestris

EG EG A EG B EG C EG EG END 1 EG B EG 1 EG A Cel B Cel C Cel E Cel H EG Z EG 3 CEL A CEL B EG III EngXCA

N N N N

[637] [638] [638] [639] [640] [641] [642] [643] [644] [627] [645] [646] [647] [648] [628] [649] [650] [650] [635] [651]

B Cellulomonas fimi Microbispora bispora Streptomyces sp. KSM-9 Trichoderma reesei

Cen A Cel A Cas A CBH II

C Humicola grisea Neurospora crassa Phanerochaete chrysosporium Trichoderma reesei Trichoderma reesei Trichoderma viride

CBH CBH CBH CBH EGI CBH

D Bacillus circulans Cellulomonas uda Clostridium thermocellum Erwinia chrysenthemi

Bgc EG Cel A EG Y

E Butyrivibriofibrisolvens Cellulomonasfimi Cellulomonas fimi Clostridium thermocellum Clostridium stercorium Dictyostelium discoideum Persea americana Persea americana Persea americana Pseudomonas fluorescens Pseudomonasfluorescens Pseudomonas fluorescens

CED I Cen B Cen C Cel D Cel Z SGSP27(~6 EG Cel 1 Cel 2 Cel A Cel B Cel E

F Bacillus sp. C-125 Butyrivibrio fibrisolvens Caldocellum saccharolyticum Caldocellum saccharolyticum Caldocellum saccharolyticum Cellulomonas fimi

Xyn A Xyn A Cel B Xyn A ORF 4 Cex

I I I I

N N C N N N N C N C C N C N C

[623] [629] [652] [633]

N N N N N

[653] [636] [631] [632] [634] [654]

N

N Internal N N N

N C N N N N

[655] [656] [657] [658] [622] [624] [625] [659] [660] [661] [662] [663] [663] [630] [303] [304] [664] [665] [643] [666] [666] [626]


75

Table 3. (Continued) Family organism Clostridium therrnocellum Cryptococcus albidus Pseudomonasfluorescens Pseudomonasfluorescens Thermoascus aurantiacus

Enzyme

Terminus

Reference

Xyn Z Xyn Xyn A Xyn B Xyn

C

[667] [668] [669] [670] [671]

C C

G Bacillus circulans Bacillus pumilus Bacillus subtilis Clostridium acetobutylicum

Xyn Xyn A Xyn Xyn B

[672] [673] [674] [675]

H AspergiUus aculeatus Erwinia carotovora

EG Cel S

[676] [310]

I

Cel A

[311]

Ruminococcus albus

Table 4. Cellulose binding domains in bacterial and fungal cellulases Organism

Enzyme

Terminus

Reference

EG Cen A Cen B Cen C Cex EG A EG Z Cel A Cel A Cel B Cel E

C N C N C C C C C N C

[622] [623] [624] [625] [626] [627] [628] [629] [630] [303] [304]

CBH I CBH I CBH I CBH II EG I EG II

C C C N C N

[636] [631] [632] [633] [634] [635]

Bacteria Butyrivibrio fibrosolvens Cellulomonas fimi Cellulomonasfimi Cellulomonas fimi Cellulomonas fimi Clostridium cellulolyticum Erwinia chrysenthemi Microbispora bispora Pseudomonasfluorescens vat. cellulosa Pseudomonasfluorescens var. cellulosa Pseudomonasfluorescens var. cellulosa

Fungi Neurospora crassa Phanerochaete chrysosporium Trichoderma reesei Trichoderma reesei Trichoderma reesei Trichoderma reesei

in b a c t e r i a l a n d fungal cellulases. Sequence a n a l y s e s of four cellulases f r o m T. reesei reveal the presence of a s h o r t c o n s e r v e d r e g i o n (AB region) at either N - t e r m i n a l ( C B H I a n d E G III) o r C - t e r m i n a l ( C B H I a n d E G I) of these enzymes [-300]. T h e r e m o v a l of the A B region f r o m C B H s h a s been s h o w n to decrease significantly their affinity to m i c r o c r y s t a l l i n e cellulose. C B D of E G I has significantly h i g h e r c e l l u l o s e - b i n d i n g affinity t h a n t h a t of C B H I [301]. M o s t of the differences are a s c r i b e d to a r e p l a c e m e n t of a t y r o s i n e b y a t r y p t o p h a n in tile c e l l u l o s e - b i n d i n g d o m a i n .

76

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The most important features of bacterial CBD sequences are: low contents of charged amino acids, high contents of hydroxy amino acids, and conserved tryptophan, asparagine, and glycine residues. Din et al. [302] have demonstrated that the isolated CBD of CenA (endoglucanase) from C. fimi, while having no detectable hydrolytic activity, disrupts the structure of cellulose fibers and releases small cellulolytic particles. The CBD of an endoglucanase (CelB) from P. fluorescens var. cellulosa binds the enzyme to cellulose as does CBD of CenA from C. fimi [303]. Recently, a novel endoglucanase (CelE), containing N-terminal catalytic and C-terminal cellulose-binding domains, was characterized by the same group of scientists [304]. A truncated form of the enzyme, which lacked the CBD, displayed the same activity of a full-length CelE against soluble cellulose and acid-swollen cellulose, but substantially lower activity than the full-length enzyme against Avicel, a microcrystalline type of cellulose. The cellulosome of C. thermocelIum binds strongly to cellulose, and this binding may be mediated by a non-catalytic component of the cellulosome [-278]. A CBD has been reported to be present in a cellulosome-integrating protein A (CipA) [305]. The cellulose-binding protein (CbpA) has been shown to mediate the interaction between crystalline cellulose substrates and the cellulase enzyme complex of Clostridium cellulovorans [-306]. Mutation analysis revealed that the entire 163-amino acid region of CBD was required for the maximal binding to crystalline cellulose [307]. The cellulose binding of CDH from P. chrysosporium was discovered independently in two different laboratories [,261,308]. The binding was found to be comparable with that of T. reesei CBH I and to be associated with the FAD domain and probably independent of the catalytic site. A comparison of the binding isotherms of CDH and CBH I has shown that the dissociation constant of CDH is much lower than that of CBH I. The capacity (the amount of enzyme bound per unit mass of cellulose) is also lower, which indicates that CDH binds more sparsely than does CBH I [-309]. Recently, cDNA encoding CDH from P. chrysosporium has been cloned and characterized [310, 311]. The low sequence similarity with the conserved cellulose binding sequences of cellulases suggested that CDH might possess a specific sequence for cellulose binding which is different from that of cellulases [311].

3.2.4 Catalytic Properties It is now a well-established fact that degradation of crystalline cellulose is carried out by a multicomponent enzyme system wherein the individual components interact in a synergistic way to convert the insoluble substrate glucose [23, 180]. The synergistic action of cellulases depends on the ratio of individual enzymes, the degree of substrate saturation, and the type of substrate [-312]. The action of CBH I and CBH II and the EG I and EG III, purified from T. reesei, has been evaluated against different substrates [313]. Synergism between the EGs and CBH II follows the normal pattern for endo/exo type synergism,


77

whereas the synergism between the EGs and CBH I depends on the structural and ultrastructural features of the substrate. The activity of individual enzymes from T. reesei is greatest toward amorphous cellulose, whereas binary combinations CBH I/EG III and CBH I/CBH II exhibit a greater degree of synergism toward crystalline cellulose [314-3163. The synergistic action of two immunologically distinct cellobiohydrolases (I and II) from Penicillium pinophilum was at its maximum on Avicel when the two enzymes were mixed in the ratio 1 : 1 [317]. It was suggested that these may be two stereospecific enzymes concerned with the hydrolysis of two different configurations of non-reducing end groups, one being exposed on the cellulose surface and the other being buried as shown by Henrissat et al. [318]. While only CBH II of P. pinophilum acts synergistically with cellobiohydrolases of T. koningii and Fusarium solani, only CBH I shows synergistic action with the endoglucanases of T. koningii or F. solani [-317]. About 45 years ago, Reese et al. [319] postulated that microbial conversion of native cellulose to soluble sugars involved two enzymes that acted consecutively. The model proposed that an unidentified enzyme called C1 acted first, making the substrate more accessible to a hydrolytic enzyme called Cx. However, the precise action of C1 was not clear, and a strictly nonhydrolytic C~ has never been identified. Intramolecular synergism was recently shown in endoglucanase CenA from C. fimi, which is composed of a catalytic and a nonhydrolytic CBD that can function independently [320]. The individual domains interact synergistically in the disruption and hydrolysis of cellulose fibers. It was suggested that the catalytic domain corresponds to the hydrolytic Cx system, and the CBD corresponds to the nonhydrolytic C~ system postulated by Reese and his colleagues. The cleavage of [3-1,4-glycosidic linkages catalyzed by cellulases is commonly assumed to proceed with a lysozyme-type mechanism through protonation of the glycosidic oxygen by an acidic amino acid residue and stabilization of the resulting carbonium ion by another amino acid residue [321]. The active sites of many enzymes have been found to involve aspartic and glutamic acid residues. There are pronounced differences in the mode of hydrolysis even between CBH I and CBH II and EG I and EG IlL CBH I attacks at sites other than the non-reducing end in the higher homologs (cellodextrins), and hydrolyzes both cellobiosides and lactosides. On the other hand, CBH II shows strict substrate specificity, three to four contiguous ~-l,4-1inked glycosyl residues being required [322]. It has also been deduced that CBH I acts with overall retention of configuration, whereas CBH II acts with reversion of configuration [-323]. Endoglucanases act via retention of configuration. EG 1 belongs to the group of non-specific endoglucanases because it also hydrolyzes xylan [324]. Its hydrolytic reaction proceeds with transfer reactions, which indicates that it probably possesses several subsets, each capable of binding a glucose molecule, and the glucosyl intermediates formed during the hydrolytic reactions are then transferred to acceptor molecules [323]. The structure of some members of cellulase families is now known. Smallangle X-ray diffraction studies suggest that both CBH I and CBH II from

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T. reesei are essentially tadpole-shaped [325, 326]. The core is estimated to be 60 to 7OA long with a diameter of 40 A, and to be attached to a tail estimated to be 150 A long. Bergfors et al. [327] successfully crystallized the catalytic core of the T. reesei CBH II after removing 82 amino acids from the N-terminus by papain cleavage. It consists of a TIM-like a/[3-barrel with the active site residues (two aspartates) located in the center of a tunnel [328]. Later, the catalytic cores of CBH I and EG I from T. reesei were also crystallized using the hanging drop vapor diffusion method [329]. Joliff et al. [330] purified and crystallized enduglucanase D from C. thermocellum. Crystals were trigonal and diffracted X-rays to a resolution of 2.8 A. CelD of C. thermocellum consists of a 12-0t-helical barrel with the active site residues situated in a long groove across the surface of the enzyme [331]. It contains an extended NHz-terminal segment sticking out from the enzyme core to interact with a symmetry-related molecule through an intermediate salt bridge (Lys 38-Asp 201) [332]. EG I from Hurnicola isolens was crystallized in different forms using various precipitants by vapor diffusion methods [333]. Two forms obtained from ammonium sulfate precipitation grow as tetragonal bipyramids, while third and fourth forms obtained from PEG 8000 grow as monoclinic plates and long hexagonal rods, respectively. EG V from H. isolens consists of a six-stranded [~-barrel domain with interconnecting loops with a 40 A groove along the surface of the enzyme containing the catalytic residues Asp 10 and Asp 121

[334]. A comparison of the structures of the catalytic cores of CBH II from T. reesei [328] and the catalytic domain (E2cd) of Thermomonosporafusca revealed large differences in their active site accessibility and supported the hypothesis that the main differences between endo- and exoglucanases is the degree to which their active sites are accessible to substrates [335].

3.3 A s s a y o f Cellulolytic E n z y m e s A wide variety of substrates have been used for the assay of cellulose-degrading enzymes (Table 5). However, due to the lack of specific substrates and lack of standardization of activity determination, it has been difficult to compare cellulase production in one fungal or bacterial strain with another. The most commonly used assay of complete cellulase activity is based on the hydrolysis of filter paper [336]. The activity is defined as the amount of reducing sugars released in one hour when using 50 mg of Whatman No. 1 filter paper as substrate under standardized conditions. Released reducing sugars are quantified by the dinitrosalicylic acid (DNS) method [337], and the results are expressed as glucose equivalents. However, this method registers only the number of reducing end groups and is very much dependent upon the [3-glucosidase levels in the enzyme mixture [338]. The use of standardized dyed insoluble cellulosic substrates (dyed Avicel or filter paper) has also been


79

Table 5. Assay of cellulolytic enzymes Enzyme

Substrate

Measurement

Total cellulase

Filter paper/Avicel Dyed cellulose Cellulose agar 14C-cellulose

Reducing sugars Release of dye Agar clarification 14C_products

Endoglucanase

Carboxymethyl cellulose Hydroxyethyl cellulose

Reducing sugars viscosity

Exoglucanase

Amorphous cellulose p-Nitrophenyl-13-cellobioside p-Nitrophenyl-13-1actoside

Reducing sugars p-Nitrophenol

fI-Glucosidase

Cellobiose p-Nit rophenyl- ~-D-glucoside 4-Methylumbelliferyl-[3-glucoside

Glucose p-Nitrophenol 4-Methylumbelliferone

(Cellobiose: quinone oxidoreductase)

Reducible quinones such as 2,6-Dichlorophenol-indophenol, 3-Methyl-5-t-butyl-benzoquinone in the presence of cellobiose or lactose

Decrease in absorbance

Cellobiose phosphorylase

Cellobiose

Glucose- 1-P

recommended for the comparison of cellulase preparations [339]. An alternative is to use a X4C-labelled cellulose, where the amount of 14C set free in solution can be easily related to the total amount of 14C in the solid substrate [340]. The release of reducing sugars from water-soluble cellulose derivatives such as CMC and HEC has most commonly been used for the determination of endoglucanase activity, since these derivatives are not hydrolyzed by exoglucanases. Viscometric methods, in which the viscosity changes of a CMC solution can be calculated in absolute enzyme units, have also been developed [341,342]. However, this method requires thorough knowledge of the properties of the cellulose derivative to allow for accurate calculations. A zymogram technique using viscous solutions of CMC for the detection of endoglucanases in polyacrylamide gels after isoelectric focusing was also developed [343]. When CMC solutions are applied on blotting papers to be put on top of the gels for a short period of time, detectable reducing sugars are produced where the endoglucanases are located. Measurement of exoglucanase activity in a mixture of cellulolytic enzymes is more difficult. Deshpande et al. [344] used the heterobiosides, p-nitrophenyl-[3D-cellobioside (pNPC) or p-nitrophenyl-13-D-lactoside (pNPL) as selective substrates for the measurement of exoglucanase (cellobiohydrolase) activity. Exoglucanases, splitting off cellobiose units from the non-reducing end of cellulose chains, specifically act on the agluconic bond (between p-nitrophenyl and the disaccharide moiety) and not on the holosidic bond (between the two glucose units in cellobiose). Another procedure used for assay of exoglucanase activity

80

R.C. Kuhad et al.

takes advantage of the enzyme cellobiose: quinone oxidoreductase (CBQ), which in the presence of cellobiose or higher cellodextrins reduces quinones [345]. Exoglucanase activity in mixtures with endoglucanases can be determined using amorphous cellulose [346]. The number of endoglucanase units in the enzyme mixture is determined preferably by the viscometric method by Almin and Eriksson [341], and the amount of reducing sugars (A) produced by the endoglucanase is computed from a standard curve prepared by the use of known amounts of purified endoglucanase towards the same amorphous cellulose. The amount of reducing sugars (B) obtained with the enzyme mixture is then measured. The reducing sugars content (C) released due to exoglucanase activity is estimated as C = B - A . Any problem caused by the presence of [3glucosidase activity in the enzyme mixture can be overcome by addition of 6-gluconolactone, which effectively inhibits the [3-glucosidase activity. Crystalline cellulose cannot be used in this assay, since the synergistic effect obtained from the endo- and exoglucanases will conceal the results. The most commonly used assay methods for [3-glucosidases include measurement of the release of glucose from cellobiose or of p-nitrophenol from p-nitrophenyl-[3-D-glucophyranoside (pNPG) 1-347]. The most specific method used to study the production of glucose from cellobiose is the glucose oxidase/peroxidase method [348]. [3-Glucosidase activity can also be assayed by determining the release of 4-methylumbelliferone from the substrate 4-methylumbelliferyl-[3-glucoside [349]. The most commonly used substrates for the assay of CBQ are reducible quinones such as DCPIP or 3-methoxy-5-tert-butylbenzoquinone (DTBB) [260, 350-352]. The substrate reduction is monitored by the decrease in absorbance at a specific wavelength, 600 nm and 420 nm for DCPIP and DTBB respectively [351,263]. CDH can also reduce both of these substrates. However, only CDH can reduce cytochrome c. Cellobiose phosphorylase activity is commonly assayed spectrophotometrically by measuring the formation of glucose-l-phosphate from cellobiose [146].

3.4 Possibilities for Biotechnology Based on Cellulolytic Enzymes Cellulases have been commercially available for the last 15-20 years. They have attracted a lot of commercial interest because of their many industrial applications in areas such as bioconversion of cellulosic materials, food, textiles, pharmaceuticals, detergents, wastewater treatment, and fruit and vegetable processing (Table 6) [1, 2, 353]. Interest in the application of enzymes in the pulp and paper industry has increased dramatically in recent years. The main application now is to remove inks from recycled papers. More than 40% of the total paper used in the United States is now being recycled. In Europe and Japan this figure is about 55%. Production of market deinked pulp in the US has, over the past six years, increased from 210000 tons to about 1 million tons by the end of 1995 and is


81

Table 6. Applications of cellulolytic enzymes

Field

Applications

Bicconversion

Conversion of cellulosic materials to sugars for production of ethanol, other solvents, organic acids, and single cell protein Biostoning and biopolishing of jeans and other fibers

Textiles Detergents Food

Medical Livestock Pulp and paper Environment

Cellulasesas a componentin detergents Improvementofyieldsin starchand proteinextraction;macerationand colour extraction of fruits and vegetables Additivein digestiveenzymes Feed additiveand silagequalityimprovement Enzymaticdeinking,increaseof freenessof fiber suspensions Waste water treatment/purification

expected to reach 1.8 million tons by the end of 1997. It is estimated that approximately 90% of pre-consumer paper waste and 42% of post-consumer office waste paper (OWP) is likely to be recovered and reused to produce printing/writing paper [354]. The conventional chemical methods for deinking are less well suited for deinking of laser and xerox printed paper for cost-effective production of high quality pulp [355]. Cellulases alone, or in combination with a few other enzymes such as hemicellulases, have been found useful for deinking different types of paper waste. Several reports and patents describing enzymatic deinking have been issued [356-359]. Endoglucanases and endoxylanases have been demonstrated to effectively deink selected old newsprint waste [360] and to improve optical and strength properties of paper from enzymatically deinked pulp. Successful enzymatic deinking of mixed wastepapers, i.e., old newspapers/old magazines (ONP/OMG), using specially formulated mixtures of enzymes followed by flotation to separate the enzymatically released ink from the fibers, has been demonstrated [355]. The enzymatic deinking process also improves freeness (water drainage of the recycled fibers) compared to chemically deinked fibers. Enzymatic deinking represents the first promising biotechnological application of cellulases in the pulp and paper industry. In a short period of time, the use of cellulases has thus made an impressive leap forward. The cellulase-based deinking process offers an environmentally benign way to improve pulp and paper production from recycled waste papers. Cellulases have been successfully used in the textile industry for bio-stoning of jeans and bio-polishing of cotton and other cellulosic fabrics. Traditional stone-washing of jeans involves three steps. After removal of the starch coating of amylases (desizing), the jeans are abraded with pumice stone (1-2 kg/pair of jeans) in large washing machines (abrasion). After this treatment the jeans are washed to remove excess dye. However, the high loading of stones in the washing machines not only decreases the capacity of jean loading but also causes a lot of mechanical wear to both machines and jeans. Cellulases have been shown to effectively facilitate the removal of indigo dye

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from the fiber surfaces [361 363]. Enzymatic stone-washing allows for up to 50% higher jean load and provides the desired soft finish. Typically, 100 kg of stones are replaced by 1 kg of enzyme [363]. Two different types of cellulases, neutral and acidic, are generally used, but neutral cellulases are the enzymes of choice because of the decrease in indigo black staining and a wider working pH range. During the last few years, cellulases have also been used for quality improvement of cellulosic fabrics. This process is called "Bio-polishing." Enzymatic bio-polishing is a novel biological finishing process for textiles made from cellulosic fibers such as cotton, rayon, ramie, and tencel. The controlled treatment with acidic cellulases improves softness and water absorbency of the fibres, strongly reduces the tendency for pill formation, and provides a clearer surface structure with less fuzz [361,363]. Also, bio-polishing permanently enhances fabric look, feel and color without any chemical coating of the fibres. Enzymes like proteases and lipases are now commonly used in the detergent industries, but the use of cellulases in this industry is a more recent approach. Soil in the interfiber space of cotton is the most difficult contaminant to remove with common detergents [364]. An alkaline cellulase from a Bacillus sp. has been shown to interact selectively with cellulose in the interior of fibers and remove soil in the interfiber spaces in the presence of usual detergent ingredients [364]. The cleaning power of cellulase-containing detergents shows remarkable improvement. Cellulases are also used to improve silage of cattle feed, i.e., to produce high quality silage from grasses containing only small amounts of water-soluble carbohydrates. Addition of cellulases and hemicellulases helps to release low molecular mass sugars to promote fermentation by lactic acid-producing bacteria. Cellulases, used as feed additives alone or with proteases, have been shown to significantly improve body weight gain in piglets and to improve the quality of pork meat [365]. Enzymatic saccharification of lignocellulosic materials produces monosaccharides, which subsequently can be fermented to a variety of products such as ethanol, other alcohols, organic acids, single cell protein, and lipids. Among various pretreatment methods studied, steam explosion pretreatment at temperatures of 170-250~ was reported to be the most efficient method for preparing lignocellulosic substrates for enzymatic hydrolysis [366]. Steamexploded sugar cane bagasse was saccharified by cellulases produced by T. reesei C-30 [367]. Cellulase preparations, 20 FPU g of substrate, gave 70% saccharification, and could, if supplemented with 13-glucosidase, result in 90% saccharification. A similar effect by addition of [~-glucosidase was observed for saccharification of autohydrolyzed Eucalyptus regnans sawdust [368]. A certain amount of 13-glucosidase is thus critical for achieving efficient saccharification [366]. The cost of enzymes is an important factor for the economy of enzymatic saccharification of lignocellulosic materials. Therefore, the extent to which


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enzymes can be recycled has a significant impact on the production costs of sugar and alcohol [369]. Various recycling methods for cellulases have been discussed, some of which have certain drawbacks. However, the one proposed by Vallander and Eriksson [366], i.e., mixing already saccharified lignocellulosic materials to which enzymes are adsorbed with new substrate, seems to be an efficient way to recycle the enzymes. Bioconversion of glucose of ethanol is a well-documented technology. However, utilization of pentose sugars for ethanol production, (pentoses account for approximately 25% by weight of lignocellulosic materials [23]) is in its infancy. A suitable pentose fermentation technology needs to be developed using pentose (xylose) metabolizing microorganisms [366, 370, 371]. The bioconversion of pentose and hexose sugars to alcohol poses a challenging task for fermentation scientists. Fungal cellulases have been shown to improve the yield of sugars from unmodified malts in the brewing process [372]. In production of alcohol from cassava, addition of Trichoderma cellulases increases ethanol yields by conversion of cellulose to glucose [373]. The isolation of starch from tubers and cereals requires a release of the integrated particles as cleanly as possible. Cereal starches are released and improve yield when amylase-free cellulases and xylanases are applied to the starch slurry prior to centrifugation and washing. Cellulases used to improve and speed up color extraction by pectinases from the skin of fruits also have an important application as a part of an enzyme complex in the maceration of fruits and vegetables. Lignocellulosic wastes from pulp and paper industries, as well as wastes from dairy and agricultural industries, are potential substrates for the production of microbial protein. Two processes have been in commercial operation for such production from wood sugars. The Candida process, using the yeast Candida utilis, started in the beginning of this century, while the Pekilo process, using the fungus Paecilomyces variotii, is more recent [374, 375]. Two other processes for the direct conversion of solid lignocellulosic wastes to protein by fungi have been developed to pilot plant scale. One process using the white rot fungus S. pulverulentum (P. chrysosporium) was developed at the Swedish Pulp and Paper Research Institute (STFI) [376]. The other process, using the mold C. cellulolyticum, was developed at the University of Waterloo, Canada [377]. However, these processes were found not to be economically feasible, since the protein produced could not compete on a cost basis with soybean protein unless the substrate used in the fermentation had a negative value [378]. The STFI approach to protein production based on lignocellulosic substrates was later changed to a waste water treatment/purification process in which the dissolved substances in the white-water system from mechanical pulp production were used as substrates. The process was tested on a large pilot scale in a newsprint paper mill, and was found to have a good potential as a technique to be used for closed white-water systems of mechanical pulp and paper mills [378].

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4. D e g r a d a t i o n o f H e m i c e l l u l o s e s

4.1 Microorganisms Producing Hemicellulose-Degrading Enzymes Hemicellulases are widespread, since they are produced by fungi, bacteria from terrestrial and marine environments, rumen microorganisms (protozoa, fungi, and bacteria), yeasts, and marine algae. Fungi, yeasts, and some bacteria secrete hemicellulases extracellularly, although cell wall-bound and intracellular hemicellulolytic enzymes have also been reported [149, 379]. Table 7 lists some important microorganisms producing hemicellulose-degrading enzymes. They are produced both constitutively and inductively. Hemicellulolytic activity is generally associated with an enzyme complex consisting of a variety of enzymes

Table 7. Important microbial sources of hemicellulolytic enzymes

Enzyme

Bacteria

Fungi

[3-1,4-Xylanases

Bacillus pumilus Bacillus subtilis Clostridium acetobutylicum Cellulomonas ada Streptomyces xylophagus

Aspergillus niger Fusarium oxysporum Aspergillus wentii Trichoderma koningii Neurospora crassa

[3-1,4-Xylosidases

Clostridium thermocellum Bacillus pumilus Acetovibrio cellulolyticus

Aspergillus niger Corticium rolfsii Penicillium wortmanni Trichoderma reesei

et-Arabinosidase

Streptomyces pupurascens Bacillus subtilis Ruminococcus albus

Aspergillus niger Cortisium rolfsii Trochoderma reesei

a-Glucuronidase

Trichoderma reesei Agaricus bisporus Pleurotus ostreatus

Esterases

Fibrobacter succinogenes Bacteroides cellulosolvens Clostridium thermocellum

Aspergillus niger Aspergillus phoenicus Trichoderma reesei

[3-1,4-Mannanases

Aerobacter aerogenes Bacillus subtilis Caldocellum saccharolyticum Streptomyces lividans

Aspergillus niger Thielavia terrestris Trichoderma reesei Paecilomyces variotii

[3-1,4-Mannosidase

Bacillus subtilis

Aspergillus niger Aspergillus awamori Thielavia terrestris Polyporus sulphureus

et-Galactosidase

Bacillus subtilis Cellulomonas sp.

Aspergillus niger Sclerotium rolfsii Aspergillus tamarii Mortierella vinacea

Microorganismsand EnzymesInvolvedin the Degradationof Plant Fiber CellWalls

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splitting off substituents from the backbone of the polysaccharides as well as depolymerizing this backbone [-23, 148, 380a, 380b]. Most filamentous fungi secrete these complexes of hydrolytic enzymes in response to inductive stimuli [23, 381,382]. Xylanases, mannanases, cellulases and other related enzymes are usually secreted together into the culture media, and selective induction of individual enzymes is not a common mechanism in fungi [,190, 383-386]. Hemicellulolytic enzymes of fungal origin are well characterized and have been studied in detail [23, 387]. Production and characterization of xylanases from several Trichoderma, Aspergillus and Fusarium strains have been reported [149, 388]. Brown-rot fungi such as Tyromyces palustris, Poria placenta and Coniophora cerebella are also potential producers of these enzymes [389, 390]. Of the fungal hemicellulases, those from A. niger have been best characterized [-23, 150]. The three-dimensional structures of several low molecular mass xylanases have recently been reported ~391-393]. The Trichoderma xylanase is ellipsoidal with dimensions 32 A x 42 A. Only a few yeast species such as some species of Aureobasidium, Cryptococcus, and Trichosporon are known to produce xylan-degrading enzymes and are therefore identified as xylanase producers [-23]. Hemicellulases of bacterial origin are less studied, because eukaryotic microbes, such as the filamentous fungi, are better producers of these enzymes. Most of the bacterial studies are confined to species of Bacillus, Streptomyces, Cellulomonas, Thermomonospora, and Chainia sp. Among actinomycetes, the xylan-degrading enzyme systems of some Streptomyces have been studied in some detail, whereas little information is available on the nature and function of hemicellulolytic enzymes produced by other actinomycetes [-23, 394]. The anaerobic bacteria have been reported to be producers of hemicellulosedegrading enzymes. The major bacteria are of the genera Clostridium, Thermoanaerobacter, Acetovibrio, and Bacteroides [23, 395-397]. Khan and coworkers [397] have studied the production of endoxylanases, xylosidases, and esterases from C. thermocellum, A. cellulolyticus, A. cellulosolvens, and B. cellulosolvens. More recently, two acetyl-xylan esterases, two 13-xylosidases, an a-glucuronidase, and a cell-associated endoxylanase have been purified and characterized from the Thermoanaerobacterium sp. strain JW/SL-YS485 [-398-400]. Six distinct ~-xylosidase genes of C. stercorarium 9F-9 have been cloned. One of them (XylA) has been reported to encode a bifunctional protein (xylosidase A) with [3-D-xylosidase and a-L-arabinofuranosidase activities [-401]. However, the gene products do not seem to have two catalytic domains, as both activities resided in one catalytic site.

4.2 Hemicellulolytic Enzymes Xylose and mannose form the backbone of the hemicellulose polymers in wood. Because of complex structure of the hemicelluloses, several different types of enzymes are required for their enzymatic degradation and modification. Among

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these, the two major hemicellulose-degrading enzymes are endo-l,4-[3-Dxylanase and endo-l,4-[3-D-mannanase. The water-soluble oligosaccharides are then further hydrolyzed by 1,4-13-D-xylosidase. Moreover, enzymes splitting off side groups and notably, a-c-arabinosidase, a-D-glucuronidase, 0t-D-galactosidase, acetylxylan esterase, and acetylgalactoglucomannan esterases have been found to be of importance for a complete degradation of these heteropolymers [23, 402, 403].

4.2.1 Xylan-Degrading Enzymes A complete degradation of branched xylans requires the concerted action of several different enzymes (Fig. 4), i.e. endoxylanase, [3-xylosidase, e-glucuronidase, e-arabinosidase and acetylxylan esterase. Endoxylanase, the

H

.

H

u.

H

"

I

COOH

o

H

.

!

o.

OH

H

H

H

.Io. ! 0

.

O,e

OH

Araf OI

Ac

lO

- - 4XyN1--4XylOl--4Xyl/31--4Xyl.81--4Xyl/~ 1--4XylOl--4XylOl--4XyL81--4Xyl.81 --4XylO 1-2 2 2 Of

I~

MeGIcA

endo-l,4-O-xylanase

MeGlcA

(EC 3.2.1.8)

Xyl~1--4Xyl/31--

O-xylosidase (EC 3.2.1.37) 9E ~ =-glucuronidase (EC 3.2.1. ) '~

=-L-arabinofuranosidase (EC 3.2.1.55) acetylesterase (EC 3.1.1.6) or acetyl xylan esterase ?

Fig. 4. A hypothetical structure of plant xylan and the linkages attacked by microbial xylandegrading enzymes.The fragmentcomprising5 D-xylose(Xyl)monomersis shown in the upper part of the figure.Ac acetylgroup, ArafL-arabinofuranose, MeGlcA 4-O-methyl-D-glucuronicacid [148]


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most widely studied and best characterized xylanolytic enzyme, attacks the xylan backbone to produce both substituted and nonsubstituted shorter oligomers, xylobiose and xylose. [3-Xylosidase is employed to convert oligomers to xylose and acts in concert with endoxylanases, a-glucuronidase, a-arabinosidase and acetyl-xylan esterase to achieve total hydrolysis of xylans to monosaccharides [148]. Endoxylanases (1,4-[3-D-xylan xylanohydrolase, EC 3.2.1.8) hydrolyze the 1,4-[3-D-xylopyranosyl linkages of xylans such as L-arabino-D-xylan, L-arabinoD-glucurono-D-xylans and D-glucurono-D-xylans. The existence of an exoenzyme (1,4-[3-D-xylan xylohydrolase, EC 3.2.1.37), although much less studied, has also been reported [379]. It appears that the xylosidic linkages in lignocellulosics are not all equivalent and equally accessible to xylanolytic enzymes. The accessibility of some linkages also changes during the course of hydrolysis due to removal of substituents and shortening of the backbone chain [404]. The existence of two types of fungal endoxylanases has been demonstrated, i.e., debranching or arabinose-releasing xylanases and non-debranching or xylotriose-cleaving xylanases [405]. Both types of xylanases are capable of attacking glucuronoxylans and unsubstituted 1,4-[3-D-xylans [149]. A. niger van Tiegham produces three endoxylanases, Xyl I, Xyl II and Xyl III [406]. Hydrolysis of rice straw arabinoxylan by Xyl I resulted in the accumulation of arabinoxylotriose and arabinoxylobiose. These were degraded by Xyl II. However, Xyl III, which had both arabinose-releasing and xylotriose-cleaving activities, did not enhance the hydrolysis of arabinoxylan in addition to what was already accomplished by Xyl I and Xyl II. The non-debranching xylanases degrade heteroxylans randomly. Five xylanases of different specificities were isolated from a commercial enzyme preparation (Rhozyme HP-150 from A. niger) that did not debranch arabinoglucuronoxylans E407]. One of these (20.8 kDa, pI 6.7) degraded heteroxylans and xylose oligosaccharides to mainly xylobiose and xylose. The four other xylanases (1~28 kDa) degraded heteroxylan to xylose oligosaccharides (substituted heteroxylan) but without release of xylose. Vrsanska et al. [408] studied the quantitative binding and hydrolysis of xylose oligosaccharides by an A. niger xylanase. The substrate binding site of the enzyme had seven D-xylosyl binding subsites. Bond cleavage frequencies of oligosaccharides by this enzyme were dependent on substrate concentration. At low substrate concentration, hydrolysis of xylo-oligosaccharides occurred via a (unimolecular) mechanism that yielded xylose from xylotriose, xylobiose from xylotetrose, and xylobiose and xylotriose from xylopentose as major end products. At higher substrate concentrations, there was a deviation from a unimolecular mechanism to one approaching a termolecular shifted hydrolysis with subsequent change in product distribution. Thus, hydrolysis of xylotriose yielded xylobiose, while xylotetrose and xylopentose were prone to transglycosylation reactions resulting in xylooligosaccharides of higher DP, which in turn were subsequently degraded to mainly xylotriose and xylobiose. Both xylobiose and xylose could be utilized as glycosyl acceptors in transglycosylation reactions.

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Sporotrichum dimorphosporum degraded redwood arabinoglucuronoxylan mainly to xylose, xylobiose, arabinoxylobiose, arabinoxylotriose, glucarabinoxylotriose, and glucarabinoxylotetrose [409]. Although L-arabinose and D-glucuronic acid were identified at the branch points of the reducing ends of the D-xylosyl chains of the oligosaccharides, they were not released on hydrolysis. The wood-rotting fungus, Trametes hirsutus, was found to produce a xylanase (23 kDa), which degraded willow 4-O-methylglucuronoxylan mainly to xylotetrose, xylopentose and 4-O-methyl derivatives [410]. Species of Trichoderma have long been known to produce xylanases. These enzymes are generally produced along with cellulases during growth on lignocellulosic materials. Trichoderma xylanases have been found to be active on xylans from different sources, usually producing xylooligomers, xylobiose and xylose. Xylose is not the major product and it is typically produced after an accumulation of xylo-oligomers [411]. Partially deacetylated xylo-oligomers were found to be more accessible than acetylated xylo-oligomers to hydrolysis by xylanases. Most of the Trichoderma endoxylanases appear to lack activity for removing the arbinosyl substituents from arabinoxylan. However, the xylanase from T. koningii was found to release arabinose. Non-specific xylanases from Trichoderma spp. have also been reported. They not only hydrolyze xylan but may also attack cellulose, CMC, p-nitrophenyl-~-glucoside, cello-oligomers, and laminarin [411]. Multiple xylanases (electrophoretically distinct endoxylanases) of microbial origin have been found [404]. The production of a system of enzymes, each enzyme having a specialized function, is probably one strategy that a microorganism uses to achieve efficient substrate hydrolysis. More comprehensive studies are required before the basis of xylanase multiplicity can be properly understood. The regulation, cross-substrate specificity, and post-translational modifications of these enzymes must be considered [411]. In fungi some of the multi-enzymes may also be allozymes, products of different alleles of the same gene. On the other hand, each of the multiple xylanases may be a distinct gene product produced by a microorganism to enhance xylan utilization. Cooperative interactions among multiple endoxylanases (electrophoretically distinct) from N. crassa, S. exofoliatus, T. byssochlamydoides, and T. harzianum have been demonstrated [412]. They can increase the extent of hydrolysis of xylan. Thus, cooperative interactions involving all the three endoxylanases from T. harzianum are required to achieve maximal hydrolysis of aspen xylan [413]. The degree of cooperativity apparently increased with increased complexity of the substrate, from the form of the deacetylated polymer to that of the acetylated polymer and, most significantly, to the form of aspen holocellulose. Hydrolysis of corn cob arabinoxylan releases arabinoxylotriose containing an interposed L-arabinose residue attached to a terminal D-xylose unit [-414]. Yeasts belonging to the genera Aureobasidium, Cryptococcus, and Trichosporon have been recognized to produce xylanases [23, 415]. T. cutaneum produces a single xylanase (Mr 45 kDa), which degrades several xylans but not xylobiose or cellulose [416]. Degradation of oat husk arabinoxylan by


89

T. cutaneum xylanase yielded xylose, xylobiose, and xylotriose as major end products. Endoxylanase from C. albidus exhibits a low affinity for p-nitrophenyl-[3-D-xyloside [380a]. Xylobiose was attacked at a very slow rate by this xylanase and cleavage was detected only when radio-labelled [1-3H]xyl2 was used [417]. One of the most studied bacteria, an alkalophilic Bacillus sp. C-59-2, produces a xylanase which degrades rice straw arabinoxylan to xylobiose and xylotriose as major end products with smaller amounts of longer xylo-oligosaccharides [418]. A xylanase from an acidophilic Bacillus sp. 11-1S degraded xylans of larchwood and rice straw in an endo manner, liberating xylobiose, xylotriose, and xylose as the major end products of hydrolysis. Streptomyces endoxylanases preferentially attack the xylan chain where the degree of branching by arabinose is low [419]. These xylanases yield mainly xylotriose in the early stages of hydrolysis. As the hydrolysis proceeds, xylo-oligosaccharides with a degree of polymerization of 3 5 are further degraded into xylobiose and some xylose [420423]. A purified endoxylanase from Streptomyces sp. hydrolyzed corn hull and cob arabinoxylans and several other xylans to xylotriose, D-xylose and L-arabinose [414]. The arabinoxylans are not completely hydrolyzed by Strepromyces endoxylanase and a mixture of substituted oligosaccharides is obtained as products [424]. A Streptomyces isolate has been observed to exhibit a very low arabinosidase activity [-423]. For details about hemicellulases from actinomycetes see ref. [394]. Cellulose-binding domains have been conclusively demonstrated in xylanases of P. fluorescens var. cellulosa [425, 426]. The amino acid sequences have been determined by cloning and expression of the genes encoding two endoxylanases (XylA and XylB), one an arabinofuranosidase (XylC) and the other an acetyl esterase (XylD). The N-terminal sequence of XylA is remarkably similar to the C-terminal sequence of endoglucanase A [427] and to the N-terminal sequence of endoglucanase B from P. fluorescens var. cellulosa [428]. Within these regions of all three enzymes is a homologous sequence of about 100 amino acid residues followed by two serine-rich regions flanking a second smaller homologous region. XylA binds strongly to crystalline cellulose, but not to insoluble xylan. It can also hydrolyze soluble xylan while bound to cellulose [429]. The non-specific xylanases of Myrothecium verrucaria and Penicillium capsulatum also bind to cellulose and are active while still bound [430]. However, they are easily eluted with buffer, and it is not clear as yet whether they bind non-specifically or via cellulose-binding domains. Recently, an extracellular endoxylanase from F. oxysporum has been reported to contain a CBD, a peptide (--,2 kDa) consisting of 18 amino acids. The amino acid sequence showed no homology with any known CBD [431], and therefore could depict a new class of fungal CBDs. 13-D-Xylosidase (1,4-[~-D-xylan xylohydrolase, EC 3.2.1.37) hydrolyzes xylooligosaccharides to xylose and are essential for the complete hydrolysis of xylans. This enzyme is produced in appreciable amounts by A. niger, and xylosidases have also been characterized from T. reesei, Sclerotium rolfsii,

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Chaetomium trilaterale and B. pumilus. Two [3-xylosidase genes from B. pumilus have been isolated and their products compared [432]. The purified [3-xylosidases I and II were both dimers consisting of 65-70kDa subunits. Xylosidase I converted oligosaccharides to xylose, while xylosidase II had little activity on xylobiose. The xylosidase from B. pumilus does not transfer xylose residues to nucleophiles other than water and operates with inversion of product configuration, suggesting a single displacement mechanism without formation of an intermediate enzyme-xylosyl complex [433]. Fungal xylosidases exhibit several different properties compared to B. pumilus enzymes. The molecular masses of fungal enzyme proteins are generally higher, and they do exhibit transferase activity. Strong transferase activities have been observed in [3-xylosidases from A. niger [434] and Penicillium wortmanni [435]. Most of the reported enzymes show highest activity for xylobiose and no activity for xylan. The activity toward xylo-oligosaccharides decreases rapidly with increasing chain lengths. T. viride xylosidase hydrolyzed xylo-oligosaccharides in the rate order X2 > X3 > X4 > X5 [436]. The production of e-D-glucuronidase (EC 3.2.1.??) by the fungi Agaricus bisporus, Pleurotus ostreatus [437], and T. reesei [438] has been demonstrated. The importance of this enzyme for the hydrolysis of acidic xylo-oligomers was pointed out by Puls et al. [438, 439]. The enzyme was found to act in synergism with endoxylanases and to liberate 4-O-methylglucuronic acid from 4-0methylglucuronic acid-substituted xylo-oligomers. Among the different organisms tested, only T. reesei seemed to produce all the side-group cleaving activity [440]. F. oxysporum and S. olivochromogenes produced acetyl-xylan esterase, while other side-group cleaving activities were low in the culture filtrates of A. awamori and B. subtilis. The molecular masses of the purified aglucuronidases from T. reesei [440] and A. bisporus [439] were 70 and 45 kDa, respectively. a-L-Arabinofuranosidase (EC 3.2.1.55) hydrolyzes nonreducing ~-L-arabinofuranosyl groups of a-L-furanosides, arabinans, arabinoxylans, and arabinogalactans. Although several a-arabinosidases have been purified and partially characterized, relatively little has been reported about their role in the hydrolysis of xylan. The enzyme has been characterized from A. niger, Corticium rolfsii, Streptomyces purpurascens, B. subtilis, Thermoascus aurantiacus and Ruminocoecus albus [441~443]. All a-arabinosidases hydrolyze beet arabinan to arabinose. Arabinose is also liberated by purified xylanase from A. niger [444] and T. koningii [445]. However, these xylanases did not show any activity toward p-nitrophenyl-a-L-arabinofuranoside, arabinotriose, or arabinotetraose. Some a-arabinosidases have been reported to produce arabinose from arabinoxylan [441]. Although arabinose-substituted xylo-oligosaccharides, produced by endoxylanases, were the preferred substrates for T. reesei a-arabinosidase, the purified enzyme also released arabinose from long arabinoxylan chains [446]. The existence of acetyl-esterases in unpurified enzyme preparations was demonstrated by Frohwein et al. [447]. These enzymes were shown to hydrolyze


91

acetylated carbohydrates, i.e. acetyl-mannose, acetyl-glucose, acetyl-maltose and acetyl-cellobiose. Williams and Withers [448] isolated more than 100 bacterial cultures from bovine rumen and observed acetyl-esterase activity in nineteen of these isolates. The existence of acetyl xylan esterase (EC 3.1.1.6) in fungal cultures was first reported by Biely et al. [449]. The esterases of T. reesei, A. niger, S. commune and A. pullulans efficiently de-acetylated steamed xylan. Partially purified acetyl-esterases and endoxylanases from T. reesei and S. commune exerted a synergistic action in the hydrolysis of acetylated xylan, and xylo-oligomers, xylose, and acetic acid were produced. The acetyl-esterase fi-om T. reesei had a Mr of 45 kDa, and the activity could be separated by chromatofocussing into two isoenzymes with pI 6.8 and 6.0, respectively [-450]. The enzymes exhibited activity toward naphthyl acetate, triacetin, and glucoseand xylose-acetate. Feruloyl and p-coumaroyl esterases have also been detected in the culture supernatants of fungi and bacteria, including S. commune, A. niger, A. phoenicus, A. awamori and F. succinogens [451, 452]. These enzymes are responsible for the selective hydrolysis of ester bonds between L-arabinosyl residues and ferulic (4-hydroxy-3-methoxycinnamic) or p-coumaric (4-hydroxycinnamic) acids. Recently, McCrae et al. [452] isolated and characterized these enzymes from A. awamori. Wheat straw xylan was de-esterified by the esterases without prior degradation of the polysaccharide. However, esterified short-chain xylooligosaccharides, which were generated by a purified fungal xylanase, were better substrates.

4.2.2 Mannan-Degrading Enzymes

1,4-[3-D-Mannanase (1,4-[3-D-mannan mannanohydrolase, EC 3.2.1.78) is capable of hydrolyzing the 1,4-13-D-mannopyranosyl linkages of D-mannans and D-galacto-D-mannans. Highly purified enzyme preparations from B. subtilis and A. niger are also capable of hydrolyzing the D-gluco-D-mannans of konjac and arum root, providing D-glucose, D-mannose, and a series of manno- and glucomanno-oligosaccharides. Both endo and exo types of D-mannanases have been characterized and are produced by various microorganisms including intestinal and rumen bacteria and fungi. Microbial endomannanases have been reported to be both inductive and constitutive, usually being secreted extracellularly [453]. However, in some bacteria such as Sporocytophaga myxococcoides, Aerobacter mannanolyticus, and Xanthomonas campastris, intracellular or membrane-bound endomannanases have been reported [23]. Endomannanases hydrolyze ~-D-mannans to D-mannose and a series of mannose oligosaccharides of DP 2-6. Their action on larch glucomannan (mannose:glucose ratio 3:1) also yield D-glucose in addition to mannose [454]. Galactoglucomannans from spruce and Canadian hemlock were degraded by A. niger endomannanase to mannobiose, mannose, glucose, and

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mannose-oligosaccharides [-455]. Several mannose oligosaccharides containing glucose and galactose have also been identified in enzymatic hydrolysates of Canadian hemlock. Araujo and Ward [456] isolated and characterized four ~-D-mannanases and one ~-mannosidase components from Thielavia terrestris. The enzymes were glycoproteins with carbohydrate contents ranging from 6 to 37%. The enzyme components hydrolyzed coffee bean mannan in a synergistic manner. Probably due to a binding requirement near or at the active site, most fungal endomannanases studied to date need four or more mannosyl residues for efficient hydrolysis [457]. Some mannanases are able to hydrolyse mannotriose but at a slow rate [458]. A multiplicity of mannanases have been observed in several fungal species including Tyromyces palustris [459], Polyporus versicolor F460], and T. terrestris [-456]. 1,4-~-Mannosidases (~-D-1,4-mannoside mannohydrolase, EC 3.2.1.25) hydrolyze 1,4-1inked-[~-D-mannosyl groups from the non-reducing end of their substrates, manno-oligosaccharides and mannose-containing glycopepetides [-461]. Its absence from mannanase preparations results in the accumulation of oligosaccharides during mannan hydrolysis. A purified mannosidase from Polyporus sulphurius liberates ~-D-mannopyranosyl units from various natural substrates [-462]. A [3-mannosidase from A. niger exhibited high activity towards p-nitrophenyl-[3-D-mannoside, but did not hydrolyze the corresponding 0t-D-mannoside or other tested p-nitrophenyl glycosides [463]. The enzyme activity against 1,4-[3-mannotriose was higher than the activity against mannobiose. However, the rate of hydrolysis of 1,4-[3-1inked D-mannooligosaccharides by ~-mannosidase from an edible mushroom, Tremella fuciformis, was found to be markedly reduced with increasing degrees of polymerization [464]. Thus [3-mannotriose was hydrolyzed at a rate which was 30%, and ~-mannotetrose at a rate which was 13% of the rate of hydrolysis of mannobiose. ~-Galactosidase (a-galactoside galactohydrolase, EC 3.2.1.22) occurs widely in microbes, plants, and animals. The enzyme catalyzes the hydrolysis of melibiose, methyl-, ethyl-, phenyl-, and o-nitrophenyl-0t-D-galactosides. Addition of any one of the above 0t-galactosides to an exponentially growing culture of A. aerogenes causes as much as a 100-fold increase in the rate of ~-galactosidase production [465]. The enzyme can remove the 0t-D-galactosyl units found as branched substituents on softwood O-acetylgalactoglucomannans. The purified enzyme of A. niger was found to be highly specific for the anomeric configuration of the glycosidic linkages [-466]. Purification and crystallization of an ~-galactosidase from the fungus Mortierella vinacea have been reported [-467]. This hydrolyzes the usual ~-D-galactopyranosides but does not liberate D-galactose from galactoglucomannan of guar gum. Similarly, two 0t-galactosidases from Aspergillus tamarii catalyzed the hydrolysis of o-nitrophenyl-0t-D-galactoside, melibiose, raffinose, and stachyose but did not attack the galactoglucomannans [468]. However, a T. reesei 0t-galactosidase has been shown to release galactose from polymeric galactomannan [469].


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4.3 Assay of Hemicellulose-Degrading Enzymes The activity of endo-xylanase is generally assayed by measuring the increase of reducing sugars released from xylan substrates (Table 8). However, there are many factors causing variations in the determination of xylanase and xylosidase activities. It has been demonstrated that the reported activities in international units (IU) varied by a factor of 3 to 107, depending on the dilution of the enzyme prior to assay [470]. Xylanase activity is also affected by the type of substrate, varying from 1.5 to 104 IU/ml among the different xylan preparations from larchwood. The units of enzyme activity obtained also vary with the availability of easily degradable xylo-oligosaccharides in the substrate. The measurement of xylanase activity is clearly dependent upon the differences in the mode of action of different xylanases from different sources and also upon the considerable heterogeneities in xylans from different sources. It is therefore necessary to use standardized methods for preparation of xylan substrates and standardization of enzyme dilution to minimize variations. Other methods such as viscometry [471] and nephelometry [472] have also been used to assay xylanase activity. The substrates p-nitrophenyl-[3-D-xylopyranoside and methyl-13-D-xylanopyranoside are commonly used for the assay of 13-xylosidase activity, p-Nitrophenol and xylose, respectively, released by enzymatic action on these

Table 8. Assay of hemicellulose-degradingenzymes Enzyme

Substrate

Measurement

D-Xylanase

Xylan

Reducing sugars Viscometry

[3-Xylosidases

p-Nitrophenyl-[3-D-xylopyranoside Methyl-13-D-xylanopyranoside Xylobiose

p-Nitrophenol Xylose

ct-Arabinosidase

p-Nitrophenyl-~-L-arabinofuranoside Phenyl-ct-L-arabinofuranoside

p-Nitrophenol Reducing sugars

~-Glucuronidase

2-O-(4-O-methyl-ct-D-glucupyrano-xyluronic acid)-xylobiose

Uronic acid

Esterases

p-Nitrophenyl acetate Birch hemicellulose Qt-Napthyl acetate Methyl esters of ferulic and p-Coumaric acid

p-Nitrophenol Acetic acid ct-Napthol Ferulic acid p-Coumaric acid

D-Mannanases

Locust bean galactomannan RBB-dyed carob galactomannan

Reducing sugars

~-Mannosidase

p-Nitrophenyl-[3-D-mannopyranoside 1,4-13-D-mannooligosaccharides

p-Nitrophenol Mannose

~-Galactosidase

p-Nitrophenyl-~-D-galactopyranoside Melibiose Raffinose

p-Nitrophenol Reducing sugars

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substrates, are then measured. ~-Glucuronidase activity is generally determined by using 2-O-(4-O-methyl-ct-D-glucupyranoxyluronic acid)-xylobiose as a standard substrate which is obtained by alkali-extraction of (4-O-methyl-Dglucurono)-D-xylan from birchwood [439]. In assays of a-arabinosidase activity, p-nitrophenyl-0t-c-arabinofuranoside and phenyl-~-L-arabinoside have been used as substrates. However, it is difficult to evaluate how specific these assay techniques might be, since 13-xylosidase of A. niger [473] and Penicillium wortmanni [474] were also reported to show activity toward p-nitrophenyl-0t-Larabinofuranoside. For estimation of acetyl-esterase activity, the rate of release of p-nitrophenol from p-nitrophenyl acetate can be used. Biely et al. [449] used a solution of a non-dialyzable fraction of birch hemicellulose for acetyl-xylan esterase activity measurement. One enzyme unit was defined as the amount of enzyme liberating one gmol acetic acid per min from 10 mg of the substrate dissolved under standardized conditions. Recently, McCrae et al. [452] described the assay methods for feruloyl- and p-coumaroyl-esterase activities, where methyl esters of ferulic and p-coumaric acid, respectively, were used as substrates, and the amounts of released ferulic and p-coumaric acid were determined by HPLC. Locust bean galactomannan and Remazol brilliant blue-dyed carob galactoglucomannan have been routinely used as substrate for the assay of endomannanase activity [475]. The measurement of the rate of cleavage of glycosidic bonds by the rate of release of p-nitrophenol is the most common assay of 13-mannosidase activity. Measurement of the release of D-mannose reducing sugar equivalents from 1,4-13-D-manno-oligosaccharide substrates is another assay technique for this enzyme [23]. Several different substrates have been used for the assay of a-galactosidase activity. Melibiose and raffinose, presumed to be natural substrates, and p-nitrophenyl-0t-D-galactopyranoside are commonly used for the assay of this activity [476].

4. 4 Possibilities f o r Biotechnology Based on Hemicellulolytic Enzymes Important criteria for industrial implementation include the existence of inexpensive and highly active enzyme preparations which can be obtained in bulk quantities. The judicious use (proper enzyme/enzymes and enzyme/substrate ratio) of proper mixes of xylan- and mannan-degrading enzymes could result in cleaner reactions, higher yields, and lower consumption of enzyme and energy [404]. Table 9 shows some industrial applications of hemicellulose-degrading enzymes. For bioconversion of polysaccharides in lignocellulosic materials to fermentation products, maximal utilization of the polymeric sugars is desirable. Complete cellulolytic and hemicellulolytic enzyme systems are required to achieve maximum hydrolysis of complex substrates to yield monomeric sugars. In order

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Table 9. Applications of hemicellulosedegrading enzymes Field

Applications

Bioconversion

Enzymatic hydrolysisof polymericsugars to monomericsugars which can be fermented to ethanol, xylitol, other chemicals,and single cell protein Enzymatic prebleaching,debarking, beating, pulp fiber refining,and production of dissolving pulps Extracting coffeeand plant oils, improving starch recovery,processingcereal flours, producingfood thickeners,providing differenttextures to bakery products, clarifyingjuices and wines, and production of xylo-oligosaccharidesfrom xylans Feed supplementationto improve nutritional properties of agricultural silage Retting of flax

Pulp and paper Food

Feed Textile

to obtain high yields of sugar, the lignocellulosic materials need to be pretreated prior to enzymatic hydrolysis. Steam explosion is one such pretreatment that has proven to be efficient and to require less energy than most other pretreatments. Sugars obtained in saccharification processes can be further utilized for the production of xylitol, organic acids, ethanol, and other solvents via fermentation [477]. Pentoses can also be used as substrates for microorganisms (C. utilis) for the production of protein-rich biomass (single cell protein), and various processes have been developed for this purpose [23]. Xylanolytic enzymes are now being used in the pulp and paper industry for the improvement of existing processes. These enzymes are used in the treatment of cellulosic pulps to remove residual xylans in the production of dissolving pulps. Hemicellulases have also been studied for enzymatic debarking and pulp refining to reduce energy demands in mechanical pulping processes [478]. In an enzymatic beating process, the enzymes are added to bleached pulp fibers to increase external fibrillation and thus improve paper-making properties [479]. Xylanase treatment produces pulp fibers with properties similar to those of slightly beaten pulps. Crude enzyme preparations containing both hemicellulases and cellulases have been used to improve fibrillation and drainage properties of recycled fibers. Mill trials of this concept have been carried out successfully using a commercial T. reesei enzyme called Liftase A40 [-480]. Xylanases probably enhance the process of fibrillation as well as facilitate enzymatic deinking of recycled fibers in combination with cellulases [481]. Dissolving pulps are used to produce cellulose derivatives such as acetates, cellophanes, and rayons. Hemicellulose contaminants lead to color and haze in the product and to poor cellulose derivatization. The use of xylanases to obtain a cellulose free of xylan was first proposed by Paice and Jurasek [482]. Xylanase treatment may reduce the chemical loading required in caustic extraction of xylan from kraft pulps. However, the feasibility of using xylanases for the production of hemicellulose-free cellulose will be dependent on a high selectivity of the enzymes used. The absence of cellulolytic activities is a particular requirement.

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Unbleached kraft pulps have a low brightness due to chromophoric groups in the lignin, the latter being removed in successive bleaching stages. Bleaching is necessary to attain the target brightness with retention of pulp strength properties [483]. The most reactive bleaching chemicals are elemental chlorine (C), ozone (Z), and peroxy acids, which react with the aromatic structures of lignin. Chlorine dioxide (D) and oxygen (O) react with lignin structures containing free hydroxyl groups. Sodium hypochlorite (H) and hydrogen peroxide (P) react with certain functional groups such as double bonds. Effective delignification can thus be achieved by the action of different types of bleaching chemicals active on different sites in lignin. Addition of an enzymatic step, i.e., xylanase, to any conventional chemical bleaching sequence results in a higher final brightness value of the pulp. The incorporation of xylanase prebleaching also permits the use of lower chlorine charges and lower charges of unselective bleaching chemicals such as peroxide and ozone. The dosages of xylanase reported to be used in enzyme-aided bleaching have varied between 30 nkat/g and about 8300 nkat/g. However, an economically realistic dosage seems to be about 100 nkat/g [483]. Yang et al. [484] developed a non-chlorine bleaching process (The EnZone Process) consisting of a xylanase treatment (X) in combination with O, Z, and P bleaching stages for bleaching hardwood (eucalyptus) kraft pulp. Pulp delignifled with an OXZP sequence readily yielded a pulp brightness of 85-90% (ISO) compared with a brightness of 78.0-84.7% for OZP bleached pulp. When evaluated against a reference pulp (ODEDED-bleached to 90.2% brightness), Enzone-bleached hardwood pulp (OXZP) had a higher brightness stability, similar tensile index, and slightly lower tear index. More recently, bleaching of softwood pine kraft pulp by the Enzone Process using a bleaching sequence of OXEpZP was reported [485]. Not only xylanases but also endomannanases and side-group cleaving enzymes have been shown to aid lignin extraction. Treatment of kraft pulps by endomannanase from T. reesei [486], B. subtilis, and A. niger [487] have been shown to improve the bleachability in subsequent bleaching stages. In xylanaseaided delignification side-group cleaving enzymes such as a-arabinosidase and a-glucuronidase provide some beneficial effects, as was shown by a lower kappa number and a higher final brightness for pine kraft pulp [488]. More than 100 mill trials of enzyme-aided bleaching processes have been carried out, about half of them in Europe. The amount of chlorine needed after xylanase prebleaching has been shown to be reduced by 20-30% [483]. The enzymatic treatment is fully compatible with existing industrial equipment, and generally no expensive investments in the mills are necessary [489]. The price of enzymatic treatment is estimated to be 2-5 USD per ton of pulp, which is expected to decrease with adaptation of more efficient enzyme-producing strains and technologies [483]. Xylanases have also been used for clarifying juices and wines, for extracting coffee and plant oils, for improving nutritional properties of silage, for macerating plant cell walls, for producing food thickeners, and for providing different

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textures to bakery products [490]. Pure xylanase is not required in many of these applications. Xylo-oligomers produced by enzymatic hydrolysis of xylan may be valuable for their rheological properties [491]. Xylanases have also been proposed for use in the processing of cereals. Addition of xylanase causes loss in wheat flour dough water absorption and consistency, as well as changes in loaf texture [4923. Xylanases have also been used to enhance the recovery of starch from wheat flours [493]. The major industrial use of ~-galactosidase is in the hydrolysis of raffinose in the sugar beet industry [494]. Limited endo-hydrolysis or removal of side-groups may be used for the modification of polymers, e.g., galactoglucomannans used as thickeners in the food industry.

5 Degradation of Lignin 5.1 Microorganisms Involved in Lignin Degradation Lignin is degraded to different extents by different microorganisms, of which wood-rotting fungi are the most effective, white-rot fungi in particular [23, 52]. These predominantly degrade wood from deciduous trees (angiosperms), but coniferous wood trees (gymnosperms) are also degraded [23, 495]. Most whiterot fungi degrade wood by a simultaneous attack on the lignin, cellulose, and hemicelluloses, but a few are rather specific lignin degraders [23, 98, 99]. The white-rot fungi produce an array of extracellular oxidative enzymes (Table i0), the best characterized of which are lignin peroxidase (LIP), manganese peroxidase (MnP), and laccase [52, 496 5003. The regulation of the production of individual enzymes and lignin degradation is a complex phenomenon. Results obtained in laboratory conditions cannot easily be extrapolated to the decay process in the cell walls of woody plants. However, Lip and MnP have also been isolated from rotting wood [501]. The effects of nutritional and cultivation conditions on lignin degradation and production of ligninolytic enzymes have been studied by various groups throughout the world and discussed in detail in Eriksson et al. [233. The role of regulatory factors such as Mn on lignin degradation and enzyme production is not fully understood. During degradation studies, Mn has been found to be deposited in white-rotted wood [23, 100], and has been reported to play a regulatory role in the degradation of lignin and the expression of laccase, MnP, and LiP [502, 5033. Soft-rot fungi efficiently degrade wood polysaccharides but degrade lignin slowly and incompletely [47]. They have been observed more commonly on hardwoods than on softwoods. Some species of soft-rot fungi studied for their degradation of lignin and lignin model compounds are shown in Table 11. Degradation of lignin in beechwood by Chaetomium globosum and lignin loss along with wood weight loss was correlated with demethoxylation of lignin

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[504]. Chaetomium globosum has been reported to release 20-30% 1 4 C 0 2 from differently labelled corn stalk lignins in seven weeks [80]. Daldinia concentrica was found to cause some lignin degradation, syringyl-propane units being degraded before guaiacyl-propane units [505]. Many soft-rot fungi produce enzymes which demethoxylate vanillic acid in the presence of NADH and oxygen to form protocatechuic acid and formaldehyde [506]. Although soft-rot fungi have been shown to demethoxylate lignin in wood, demethoxylation of isolated lignins has not been reported. Soft-rot fungi have also been reported to possess low levels of protocatechuate 3,4-dioxygenase [507]. Low levels of peroxidase activity in C. cellulolyticum and C. piluliferum have also been reported [508]. Brown-rot fungi (Table 11) prefer coniferous wood. According to a survey of substrate/fungal relationships, 19% of North American Polyporus species are brown-rot fungi, among which 60 out of 71 are found primarily on coniferous wood, i.e., brown-rot fungi are mainly softwood degraders [97]. These fungi cause extensive degradation of the polysaccharides and only a limited degradation of lignin [-100]. In advanced stages of degradation, the residue is a brown mass, which mostly consists of lignin. The limited change in lignin caused by brown-rot fungi involve (1) demethoxylation, and (2) aromatic hydroxylations and limited side-chain oxidations. These fungi do not cleave the aromatic rings in lignin efficiently. Brown-rot fungi have the capability of cleaving and metabolizing methoxy groups of vanillic and ferulic acids [508]. The fungi remove the methoxy groups as methanol, whereas soft-rot fungi appear to release formaldehyde as the initial product. Brown-rot like white-rot fungi have been observed to decarboxylate vanillate to methoxy-hydroquinone, and the enzyme vanillate hydroxylase was detected in brown- and white-rot fungi but not in soft-rot fungi [507]. Most of the brown-rot fungi [509] were found to produce ethylene from 2-keto-4methiolbutyric acid, which can be used as a characteristic for ligninolytic activity [510]. At least four nonenzymatic agents, e.g., oxalic acid [199], siderophores [198], Fenton's reagent (leading to hydroxyl radical production) [511], and glycopeptides [512] have been reported to be implicated in the lignin degradation process by brown-rot fungi. However, the mechanisms by which these agents function with enzymes for in situ degradation of lignin is not understood [513]. White-rot fungi, based on the production pattern of their extracellular enzymes, have been classified in four groups by Hattaka [499] and six groups by Tuor et al. [97]. Both types of classification have some overlaps and exceptions with respect to different strains which vary in their ability to produce one or the other enzyme. An attempt to group the white-rot fungi to avoid the confusion of their belonging to one or the other group has been made by Eggert et al. [514]. Table 10 shows the white-rot fungi categorized on the basis of their enzymeproducing ability. For details, readers are referred to refs. [97, 499]. Much less attention has been given to lignin-degrading bacteria compared to fungi [515]. Most of the studies of bacterial degradation of lignin were based on


99

Table 10. Phenol-oxidase production by various lignin-degrading white-rot

fungi LIP-, MnP-, and laccase-producing Coriolus versieolor Cyathus buUeri Phlebia radiata Pycnoporus sanguineus Pleurotus ostreatus Pleurotus sajo-caju LiP- and MnP-producing

Coriolus pruinosum Phanerochaete chrysosporium

LiP- and laccase-producing

Oudemansiella radicata Pleurotus florida Polyporus pletensis Polyporus brumalis Phlebia tremellosus Phlebia ochraceofulva

LiP-producing Bjerkandera adusta Daedaleopsis confragosa Polyporus varius Laccase-producing Pycnoporus c.innabarinus

MnP- and laccase-producing Ceriporiopsis subvermispora Dichomitus squalens Lentinus edodes Panus tigrinus Rigidoporus lignosus

Novel oxidase*-producing Bjerkandera adusta Pleurotus eryngii Pleurotus ostreatus Trametes versicolor Pluerotus sajor-caju

* Aryl alcohol oxidase (AAO), Veratryl alcohol oxidase (VAO)

species of Pseudomonas and filamentous Actinomycetes [516]. Bacteria generally cause a low percentage of mineralization of lignin in 14C-labelled lignin in lignocellulosic materials or in DHPs as observed in pure cultures [517]. Actinomycetes have been reported to mineralize lignin successfully, although not as fast or comprehensively as white-rot fungi [128]. A number of Gram-negative bacterial species (Table 11) belonging to the genera Pseudomonas, Xanthomonas, Acinetobacter, Achromobacter, Agrobacterium, Aerobacter, and Erwinia have been reported to degrade lignin [-128]. However, much of the present knowledge has been obtained from studies of the species Xanthomonas and Pseudomonas. Xanthomanas sp. was reported to convert 30% of 14C-labelled DHP to 14CO2 [518]. However, lignin polymers were degraded only up to a molecular weight of 1000 Da, suggesting the involvement of intracellular enzymes for lignin degradation [517]. P. fluorescens produces benzaldehyde lyase which cleaves the acyloin linkage of 1,2-diarylethane compounds anisoin and benzoin [519]. Pseudomonas spp. have also been reported to degrade biphenyls via oxidation to 2,3-dihydroxyphenyl and 2hydroxy-6-oxo-phenylhexa-2,4-dienoate to benzoic acid [520]. P. paucimobilis SYK6 produces a protocatechuate-4,5-dioxygenase which catalyzes ring fission of protocatechuate and 3-methylgallic acid, an intermediate of the metabolism of lignin model compounds with biphenyl structure [521]. Actinomycetes (Table 11) degrade grass lignins by release of lignin-rich, water-soluble fragments called acid precipitable polymeric lignin (APPL) [522].

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Table 11. Lignin-degrading brown- and soft-rot fungi and bacteria Brown-rot fungi Fomitopsis pinicola Gleophyllum trabeum Poria placenta Lentinus lepideus Pholiota adiposa Spongiporus sinuosus T yromyces palustris

Actinomycetes Arthrobacter sp. Microbispora sp. Nocardia sp. Streptomyces badius Streptomyces cyaneus Streptomyces setonii Streptomyces viridosporus Thermomonospora mesophila

Soft-rot fungi Chaetomium 91obosum Daldinia concentrica Lecythophora hoffmannii Petrillidium boydii Pialophora mutabilis

Other bacteria Acinetobacter sp. Xanthomonas sp. Pseudomonas sp. Achromobacter sp. Aerobacter sp. Erwinia sp.

APPL is not metabolized further by Actinomycetes, and APPL production seems to be a unique property of these microorganisms, observed mainly in Streptomyces and Thermomonospora [516, 522, 523]. Purified lignin peroxidase (ALip-P3) from S. viridosporus was found to catalyze C-C bond cleavage in the side chains of phenolic and nonphenotic lignin models in the presence of H z O 2 1-524]. The enzyme also oxidizes polymeric lignin, as shown by the rapid H202 consumption upon addition of purified enzyme to lignin. In addition, S. viridosporus also produces aromatic acid esterases [525] and aromatic aldehyde oxidases [526]. Ball et al. [527] screened 20 Actinomycetes for ligninolytic activity and found S. badius, T. mesophile, and S. cyaneus to be the most active lignin degraders. S. badius secretes four extracellular peroxidases similar to those of S. viridosporus [528]. With the discovery of extracellular LiP from Streptomyces spp., it seems likely that these Actinomyces attack lignin by a similar method to that of white-rot fungi [529]. S. viridosporus has also been reported to produce one or more extracellular HzOz-generating oxidases [526], but no evidence of their role in supplying H 2 0 2 to LiP for lignin degradation was demonstrated.

5.2 Ligninolytic Enzymes In the complex ligninolytic enzyme system, peroxidases, laccases, and H 2 0 zproducing oxidases are the most studied (Fig. 3). Peroxidases (LiP and MnP) and laccase are defined as phenol oxidases [23]. The reactions catalyzed by these enzymes are very similar (Fig. 3). They oxidize phenolic compounds, thereby creating phenoxy radicals, while non-phenolic compounds are oxidized to the corresponding cation radicals [530-533]. While all lignin-related phenolic compounds are oxidized by phenol oxidases, the different enzymes have altogether different substrate ranges for the non-phenolic ones [533].


101

LiP and MnP from P. chrysosporium have been crystallized and their tertiary structure examined, while no such report has yet appeared about laccase. The crystallographic structure of LiP from P. chrysosporium has been studied at different resolutions, i.e., at 2.0, 2.5, and 2.6 A [534 536]. The model comprises all 343 amino acids, one heme molecule, and three sugar residues. The crystal structure revealed that the enzyme consists mostly of helical folds with separate domains on either side of the catalytic heme. The enzyme also contains four disulfide bridges formed by eight cysteine residues [535, 536] and two structural calcium ions, which appear to be important for maintaining the integrity of the active site [536]. MnP presents a good sequence homology with LiP. However, except for preliminary information [537], no detailed reports on the structure of MnP have yet been published. Presently, an attempt to crystallize laccase is in progress in this laboratory at the University of Georgia, Athens (UGA).

5.2.1 Lignin Peroxidase Lignin peroxidase (LIP) (ligninase, EC 1.11.1.14), was first discovered in ligninolytic cultures of P. chrysosporium [153, 154], and seems to constitute a major component of the ligninolytic system. LiP catalyzes a large variety of reactions, e.g., cleavage of 13-O-4 ether bonds and C~-C~ bonds in dimeric lignin model compounds - the basis for the depolymerization reactions catalyzed by LiP. The enzyme also catalyzes decarboxylation of phenylacetic acids, oxidation of aromatic C~-alcohols to C~-oxo compounds, hydroxylation, quinone formation, and aromatic ring opening [538]. LiP oxidizes its substrate by two consecutive one-electron oxidation steps, with intermediate cation-radical formation. Due to its high redox potential, LiP can also oxidize non-phenolic methoxy-substituted lignin subunits. Several studies have indicated the importance of LiP in the degradation of lignin and xenobiotics. The enzyme can depolymerize dilute solutions of lignin [539], oxidize and degrade a variety of dimers and oligomers structurally related to lignin in vitro [532], and catalyze the production of activated oxygen species [540]. Studies of LiP oxidation of DHPs [541] confirm the involvement of LiP in the initial degradation of lignin [542]. LiP has a similar catalytic cycle to that of horseradish peroxidase. The five redox states of LiP have been characterized, and its catalytic cycle has been investigated by various groups of scientists [543 545]. In the catalytic cycle of LiP, the native Fe 3 + enzyme is first oxidized by H 2 0 / to compound I, and then a one-electron reduction of compound I with veratryl alcohol takes place [546] or H 2 0 2 oxidation results in compound II. One further electron reduction of compound II with veratryl alcohol brings the enzyme back to its native form, and in this way the catalytic cycle is maintained [543,547]. However, in competition with a reducing substrate, compound II can react with H 2 0 2 and result in the formation of the catalytically less active compound III [548].

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Compound III is stable but becomes inactivated in the presence of H 2 0 2 [544]. It has further been shown that compound III can transform back to the native enzyme either spontaneously or in the presence of H202 and veratryl alcohol [545], and the cycle can get restarted. The generation of phenoxy radicals during oxidation of phenolic substrates, in demethoxylation, and in ether cleavage reactions [549] explains the repolymerization of lignin in reactions with LiP [550]. Oxidation of phenolic compounds causes inhibition of LiP [551]. This inhibition can be overcome by lowering the H202 level relative to phenol and enzyme levels. Moreover, LiP has a preference for phenolic compounds as compared to veratryl alcohol. Thus, in order to achieve maximum enzyme (LIP) turnover and oxidation of nonphenolic compounds, the presence of phenolic compounds in the vicinity of LiP needs to be low [552]. Harvey et al. [552] have shown that veratryl alcohol is oxidized to a cation radical capable of participating in intermolecular charge transfer reactions. Based on this property of the cation radicals, it was proposed that the cation radicals of veratryl alcohol, the product of LiP catalysis, may mediate in the oxidation of lignin. In addition, these radicals may assist in the reaction of LiP compound II with the reductant and thereby maintain the active peroxidase cycle. The reduction of LiP II to native enzyme results in the release of two substrate cation-radicals (Fig. 5a). The mechanism by which substrate cationradicals promote reduction of compound II is not clear. The cation-radicals may form a pathway of charge transfer away from the vicinity of the porphyrin so that the chance of radical generation in the vicinity of the active site of LiP is

HzOz native enzyme FemP

H202

~'.

-

Con'.pound [ Pel~lO)P"

notive er~yme

Compouncl ! FeZglO2P"

FelIIP

\

e'b Compound11 FelglOl P ~ V A +"

~

e"

Compound FeIXIO) p ~ X P" -,--- Fe~IO]P__X ~

,.,,.. ~ O z

,- Compound'mr

F~mtOa'lP

Compound ~r FenttOl-}P

b

Fig. 5. Schematicillustrationof the catalyticcycleof LiP showingthe oxidationof (a) VA veratryl alcohol,(substratemodifiedcompoundII) and (b) X phenoliccompounds(enzymefeedbackcontrol) 1-552]


103

minimized. This is unlikely to happen with phenoxy radicals which are incapable of participating in charge transfer reactions. Their formation would also result in LiP inhibition (Fig. 5b). Results from different laboratories have shown that veratryl alcohol can act both as a stabilizer of LiP and as a charge transfer mediator. An excellent review concerning the physiological importance of aryl alcohols in ligninolytic fungi has recently been published 1-533].

5.2.2 Manganese Peroxidase Manganese peroxidase (MnP), another peroxidase found in ligninolytic cultures of P. chrysosporium, was discovered by Kuwahara et al. 1-155]. The enzyme acts exclusively as a phenoloxidase on phenolic substrates using Mn 2 +/Mn 3§ as an intermediate redox couple. MnP has also been shown to produce H202 in the oxidation of glutathione, NADPH, and dihydroxy-malic acid 1-553]. However, H202 production is not the major function of MnP. Instead, this enzyme is involved in the oxidation of phenols and phenolic lignin structures. It oxidizes Mn(II) to Mn(III) in the presence of a proper chelating agent, and Mn(III) must form a complex with the chelator before it oxidizes phenolic substrates 1,554]. Organic acids are good chelators, and white-rot fungi are producers of oxalic acid, malonic acid, pyruvic acid, and malic acid 1-533]. Mn(III)/oxalate and Mn(III)/malonate form very stable chelators which probably also function in vivo. Malonate facilitates Mn(III) dissociation from the enzyme and has a relatively low Mn(II) binding constant 1,554]. MnP has a catalytic cycle very similar to that of LiP. However, in this case, compound II is readily reduced by Mn 2 § to the native enzyme to complete the catalytic cycle (Fig. 6) 1-156, 555]. A Mn(III) complex can oxidize phenolic lignin substructures by acting as a mediator between the enzyme and the polymer, and leads to the formation of

Fe 3" .~j31'q ~

AH2 )~tHMnlTIT

AH l

~AH" Compound TIT F e 3 + O ~ FeZ+ O2

Compound I Fe'I~T= O [P]t

Fig. 6. The catalytic cycle of manganese peroxidase (MnP) and its five oxidation states. ROOH H202, AH2 phenolic substrate [555]

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phenoxy radicals as intermediates [530]. Subsequently C~-C~ cleavage or alkyl-phenyl cleavage would yield depolymerized fragments, including quinones and hydroxyquinones. The purified MnP has been reported to depolymerize D H P [556] and also to degrade high molecular mass chlorolignins [557]. MnP appears earlier than LiP in cultures of P. chrysosporium and can oxidize phenolic lignin structures, while LiP can also oxidize non-phenolic lignin substructures [531]. Thus, MnP and LiP jointly provide a good system for the degradation of lignin-rich woody and non-woody plant materials under proper reaction conditions. Several white-rot fungi have been found to produce peroxidases which are different from LiP and MnP, and which are generally termed manganeseindependent peroxidase, MIP [558-560]. However, information about the function of MIP in lignin degradation is scanty [533, 561].

5.2.3 Laccase Laccase (benzenediol: 02 oxidoreductase, EC 1.10.3.2) appears to play an important role in the degradation of lignin and is produced by most white-rot fungi. Both constitutive and inductive forms of laccases are known [-102, 562-564]. All laccases are glycoproteins [565], and they generally contain four copper ions [566]. These are distributed among three different binding sites, and each copper ion appears to play an important role in the catalytic mechanism. Laccase is a true phenoloxidase with broad specificity towards aromatic compounds containing hydroxyl and amine groups [500]. The enzyme oxidizes phenols and phenolic substructures by one-electron abstraction with formation of radicals that can either repolymerize or lead to depolymerization [567]. Laccase catalyzes demethoxylation reactions of terminal phenolic units. It can also degrade [3-dimers and [3-0-4 dimers via Ca oxidation alkyl-aryl cleavage, and C~-C~ cleavage [500]. Laccase has also been shown to catalyze the cleavage of aromatic rings in a similar way to LiP [-568]. It was observed that laccase from C. versicolor transformed 4,6-di-tert-butylguaiacol into a ringopening product, the muconolactone derivative 2,4-di-(tert-butyl),4-(methoxycarbonylmethyl)-2-buten-4-olide. This work clearly demonstrated that 1SO incorporated into the muconolactone was from 1802 but not from H2180. It is obvious from the results that laccase, like LiP, can cleave aromatic rings. Studies of side-chain cleavage and ring-opening of lignin model compounds indicated that both laccase and LiP, which catalyze one-electron oxidation of either phenolic or non-phenolic compounds, are involved in the initial degradation of lignin substructure model compounds [542]. Until 1990, laccase had been considered to be able to degrade only phenolic lignin model compounds [567]. However, Bourbonnais and Paice [569] then reported that oxidation of non-phenolic lignin substructures by laccase from T. versicolor took place in the presence of a suitable redox mediator, i.e., the dye


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2,2'-azinobis(3-ethylthiazoline-6-sulfonate) (ABTS). More recently, laccase along with ABTS has been shown to also delignify kraft pulp [570]. The presence of the mediator prevented re-polymerization of kraft lignin by T. versicolor laccase [571]. Other phenolic reagents have also been proposed and found to work as mediators [572]. Currently, brightening and delignification of kraft pulp has been shown with another mediator, 1-hydroxybenzotriazole (HBT) [573]. Moreover, the production of a fungal metabolite which acts as a physiological laccase/redox mediator has recently been shown in our laboratory (UGA) to be produced by the white-rot fungus Pycnoporus cinnabarinus [514]. Laccase and other phenoloxidases act in cooperation with other enzymes. Intracellular NAD(P)H-dependent quinone-reducing enzymes have been reported from P. chrysosporium [574 576]. These enzymes may constitute part of a metabolic system which facilitates fungal utilization of small fragments of lignin to degrade them intracellularly [575]. The reaction of laccase with high molecular mass fractions of lignosulfonates in the presence of glucose oxidase gives rise to more depolymerization than polymerization. Glucose oxidase causes a decrease in the generation of quinone intermediates by laccase [577]. Rapid reduction of phenoxy radicals and quinone compounds by the cooperation oflaccase with oxido-reductase type enzymes could be one possible route to shift the polymerization/depolymerization equilibrium towards depolymerization [23]. Laccase has been reported to act synergistically with MnP to bring about efficient degradation of radiolabelled 1-Ievea lignins in vitro [578]. Recently, laccase from T. versicolor has been demonstrated to generate Mn(III) chelates from Mn(II) [579], which have been shown to be implicated in the degradation of various natural [578, 580] and synthetic lignins [156, 556]. More recently, a FAD-dependent aryl alcohol oxidase (veratryl alcohol oxidase, VAO) has been observed to reduce synthetic DCPIP and guaiacyl quinonoids and phenoxy radicals, generated by laccase, with concomitant oxidation of veratryl alcohol to veratryl aldehyde. The cooperative action of laccase and VAO could prevent the polymerization of phenolic compounds and depolymerize (reduction of Mr) soluble lignosulfonates considerably [581]. Continued investigations are necessary to elucidate the role of laccase and other cooperative enzymes in lignin degradation.

5.2.4 HeO2-Producing Enzymes

White-rot fungi also produce several H202-generating enzymes. Intracellular enzymes are glucose-l-oxidase, glucose-2-oxidase, methanol oxidase, [23] and fatty acyl-CoA oxidase [582]. Extracellular H202-generating enzymes are glyoxal oxidase secreted by P. chrysosporium [161] and aryl-alcohol oxidase found in cultures of T. versicolor, P. sajor-caju, P. ostreatus, and P. adusta [533].

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Both glucose oxidases have been purified and shown to exhibit the following reactions: glucose-1-oxidase [5-D-glucose + 02 , ~-D-gluconolactone + H202 I~-D-glucose + 02

glucose-2-oxidase , D-arabino-2-hexulose + H202

Methanol oxidase oxidizes methanol to formaldehyde and H202 [160]. In addition to methanol, ethanol is also oxidized, probably to acetaldehyde. Glyoxal oxidase (GLOX) is thought to be very important for H202 generation since it oxidizes several aldehydes and ~-hydroxy carbonyl compounds which occur as secondary metabolites [583]. Recently, a role of GLOX activity in the regulation of ligninolysis by peroxidase has been suggested. The inactive GLOX can be reactivated when coupled to LiP with veratryl alcohol as the substrate for LiP [584]. Aryl-alcohol oxidase oxidizes aromatic alcohols to aldehydes and reduces 02 to H202 [533]. Moreover, MnP has also been found to generate HzOz by catalyzing the oxidation of NADP(H), glutathione, dithiothreitol, and dihydroxymaleic acid [585, 586]. 5.2.50xidoreductases Other enzymes of importance in lignin degradation are NAD(P)H: quinone oxidoreductase, aryl alcohol dehydrogenase, and probably also CDH(CBQ). P. chrysosporium produces at least two different intracellular NAD(P)H: quinone oxidoreductases [-23,574, 576, 587]. These enzymes reduce methoxyquinone using either NADH or NADPH as electron donors. Laccase can interact with this enzyme to increase decarboxylation of lignin model compounds. In addition, glucose-l-oxidase from C. versicolor could function as glucose: quinone oxidoreductase and use quinone as co-substrate in place of 0 2 at low O2 tension [588]. CDH or CBQ (the proteolytic breakdown product of CDH[263, 309]) have probably not beyond doubt been shown to play an important role in lignin

Table

12. Electronacceptorsfor CDH (CBO)and CBQ

ABTScation-radical 4-Aminopyridinecation-radical Benzylviologen Cerulignol Cytochromec 2,6-Dichlorophenol-indophenol Ferricyanide Ferriacetate Ferricitrate Guaiacylphenoxyradical Oxygen(slow)

3-Methoxy-5-tert-butyl-benzoquinone 2-Methoxy-benzoquinone Methyleneblue Mn3+ malonate Osmium(II1) TMB radical Triiodine ion Tetramethoxy-azo-p-methylenequinone Vanillicacid phenoxyradical Veratryt alcohol cation-radical


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degradation [257], but, when first discovered, C B Q was suggested to be a link between cellulose and lignin degradation. Both C D H and C B Q have been discussed in depth in Section 3. Both enzymes reduce a number of electron acceptors including quinones, phenoxy- and cation-radicals, complexed ferric ions, c o m p o u n d II of LiP, MnP, and molecular oxygen (Table 12). Quinone reduction is more rapid than oxygen reduction, oxygen being slowly reduced to superoxide and/or H 2 0 2 [589]. The following are suggested to provoke the participation of C D H in lignin degradation: (1) reduction of quinones for detoxification [260], (2) reduction of phenoxy radicals to prevent their polymerization [264], (3) reduction of compound II of LiP and M n P [590] (4) production of - O H in order to degrade lignin and other wood components [162], and (5) support of M n P action/function by means of solubilization of Mn(IV) O2(s) and production of chelating agents for manganese ions [591].

5.3 Assay of Ligninolytic Enzymes Assays for various ligninolytic enzymes are presented in Table 13. Lignin peroxidase (LIP) activity is routinely assayed in a reaction mixture containing veratryl alcohol and H 2 0 2 in a suitable buffer. The reaction is initiated by the addition of H202, and the oxidation of veratryl alcohol to veratraldehyde is monitored at 310 nm [592].

Table 13. Assay of major ligninolytic enzymes Enzyme

Substrate

Measurement

Lignin peroxidase

Veratryl alcohol H202

Monitored at 310 nm

Manganese peroxidase

Vanillylacetone Phenol red MBTH and D M A B H202

Monitored at 336 nm Monitored at 610 nm Monitored at 590 nm

Laccase

Syringaldehyde ABTS MBTH and D M A B

Monitored at 525 nm Monitored at 436 nm Monitored at 590 nm

Cellobiose: quinone oxidoreductase

DCPIP MTBBQ Cellobiose

Monitored at 600 nm (decrease in absorbance)

Cellobiose oxidase (CBO/CDH)

Cytochrome c Cellobiose

Monitored at 550 nm

ABTS, 2,2'-azino-bis(3-ethylbenzothiazoline)-6-sulphonate;MBTH, 3-methyl-2-benzothiazoline hydrazone; DMAB, 3-dimethylaminobenzoicacid; DCPIP, 2,6-dichlorophenol-indophenol; MTBBQ, 3-methyl-5-t-butyl-benzoquinone

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There are several methods for the assay of manganese peroxidase activity. However, most of them have certain disadvantages, including lack of definition of units of activity, low sensitivity, measurement in the UV range, and no determination of extinction coefficients [593]. The most common method is monitoring the disappearance of the substrate (vanillylacetone) at 336 nm [553]. The reaction mixture contains vanillylacetone, H2Oz, and MnSO4 in a suitable buffer. The reaction is initiated by the addition of peroxide, and the linear decrease in absorbance is monitored. Another method uses phenol red in succinate buffer containing H202 MnSO4, gelatin, and lactate. A certain volume of the reaction mixture is removed at one min intervals and added to an N a O H solution. Oxidized phenol red is measured at 610 nm [594]. Recently, Pillar Castillo et al. [593] developed a new method for M n P activity. The assay was based on the oxidative coupling of MBTH and DMAB, which, in the presence of H 2 0 2 , Mn z +, and MnP, gives a deep purple-blue colour with a broad absorption band and a peak at 590 nm. The extinction coefficient is high, which facilitates detection of even low activities. The activity of laccase is usually measured by the oxidation of syringaldehyde to its quinone form at 525 nm [595]. The molar absorbancy of the color formed is high. However, syringaldehyde is not very soluble in an aqueous medium, which necessitates a high alcohol concentration [596]. Laccase activity is also assayed using the dye ABTS by monitoring the progress of oxidation at 436 nm [-569]. However, if peroxidases and other oxidases are present in addition to laccase, it would present a problem, since certain oxidases would generate H 2 0 2 , a substrate for peroxidases [596]. Lonegan and Baker [596] developed a method for laccase assay using the coupling reaction between 3-methyl-2benzothiazoline hydrazone (MBTH) and 3-dimethylaminobenzoic acid (DMAB). Prior to estimating enzyme activity, samples were pretreated with an active catalase to destroy hydrogen peroxide and hence minimize peroxidase activity. CBO and CBQ activities are assayed by measuring their reducing activities for suitable electron acceptors [263, 264]. Both C D H and CBQ can utilize D C P I P as an electron acceptor, while only C D H is able to reduce cytochrome c. D C P I P reducing activity [263] is assayed at 30~ in a reaction mixture containing cellobiose and DCPIP in a suitable buffer. The decrease in absorbance at 600 nm is monitored, and one unit of activity is defined as the amount of enzyme necessary to reduce 1 tamol substrate per min. Cytochrome c reducing activity is determined in a reaction mixture containing cellobiose and cytochrome c in a suitable buffer [264]. The activity is measured at 30 ~ as the increase in absorbance at 550 nm. One unit of cytochrome c reducing activity is defined as the amount of enzyme necessary to reduce 1 ~tmol substrate per min.

5. 4 Possibilities for Biotechnology Based on Ligninolytic Enzymes The ligninolytic enzymes produced by white-rot fungi have broad substrate specificities and should therefore have potential for use in different areas of


109

Table 14. Applications of lignin-degrading microorganisms and enzymes Field

Application

Food and feed

Mushroom production; improvement in digestibility of wood and straw residues for use as ruminant feed

Pulp and paper

Biopulping and biobleaching; modification of pulp fibers; delignification of wood chips to reduce energy input in mechanical and chemical pulping

Chemical

Modification of isolated lignins to produce useful lignins and chemicals

Environment

Treatment of spent kraft bleach liquors and organic pollutants such as PAHs, PCBs, and dioxins

Biosensors

Identification of cellodextrins; measurement of cellulose surfaces of various pulp fibers using CDH

forest industrial processes, pulp and paper manufacture, by-product utilization, and waste management (Table 14). LiP, MnP and laccase, produced by white-rot fungi, have all been studied for use in pulp bleaching [597]. With mixtures of ligninolytic enzymes, a 20-30% reduction in kappa number has been observed [598]. LiP has been implicated in biobleaching with P. chrysosporium [599]. MnP and laccase were detected in cultures of T. versicolor used for bleaching of wood pulp and also in other white-rot fungi [600-602]. Paice et al. [602] reported that partially purified MnP alone demethylated and delignified pulp to almost the same extent as the complete cell-free culture solution from T. versicolor cultures. More recently, pure MnP from the fungus SKB-1152, used for pulp bleaching, has been shown to successfully bleach oxygen-alkali treated hardwood kraft pulp [603]. CBQ has also been observed in culture solutions of T. versieolor used for pulp bleaching [602]. Various studies have shown its involvement in the reduction of oxidation products of laccase or peroxidase such as quinones [260], phenoxy radicals [589], and Mn III [604]. Different approaches, such as the use of suitable redox mediators for enhanced laccase delignification and brightening of kraft pulp have been worked out [570, 572, 573]. Recently, an efficient laccase mediator system for enzyme pulp bleaching has been reported from the company Lignozym [605]. The performance of enzyme mediator systems is now being tried on the pilot plant scale. Effective pulp bleaching by cell-free oxidative enzyme such as laccase or MnP seems to require additional reagents like mediators/activators or other process controls [606]. Laccase requires a high pressure of Oz and a low molecular mass electron carrier, while MnPs require a low and controlled supply of H 2 0 2 along with Mn(II) ions and a suitable chelator such as oxalate or malonate [606]. The ligninolytic system of P. chrysosporium is also capable of degrading a number of organic pollutants and xenobiotics. Halogenated phenols too are

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oxidized to form oligomeric and polymeric products, usually less toxic than the original low molecular mass substrates. Among the polyaromatic hydrocarbons (PAHs) studied for oxidation and degradation by white-rot fungi and their enzymes are anthracene, methylanthracene, fluorenthrene, biphenyl, phenanthrene, pyrene, benzopyrene, fluorene, acenaphthene, some monoaromatic compounds, and the PAH component of the anthracene oil from coal tar distillation 1-607, 608]. Because of the high redox potential of lignin peroxidase from P. chrysosporium, the latter has been suggested for use in bio-remediation of chlorinated hydrocarbons such as DDT, CC14, TCDD, and PCBs [-609]. Use of free enzymes may have some significant drawbacks such as thermal instability, susceptibility to protease attack, inhibition of activity, and the difficulty of separating and reusing them when the reactions are completed. The phenoloxidases have been successfully immobilized on natural and synthetic supports such as soils, clays, and porous glass beads to overcome some of these problems 1-610, 6ll]. Immobilized laccase has been successfully used to remove color from phenolic industrial effluents 1,612]. Recently, immobilized phenoloxidases were found to efficiently remove naturally occurring xenobiotic aromatic compounds from aqueous solutions 1-613]. LiP produced by P. chrysosporium has also been implicated in the decolorization of several dyes 1-614, 615]. The presence of veratryl alcohol stimulates azodye oxidation. The ability of peroxidases and laccases to decolorize dyes indicates the possibility of developing processes to reduce color in pulp mill effluents. Most of the color in chlorine bleach plant effluents emanates from quinone and conjugated lignin structures, particularly in the effluent from the first alkali extraction stage (El). The color is mainly associated with the high molecular mass lignin fractions. Therefore, efficient decolorization and removal of these high molecular mass compounds are necessary. Among the white-rot fungi, P. chrysosporium has been tried for bleaching of E1 effluents from bleach kraft mill processes 1-616]. A continuous process for color removal from bleach plant effluents with P. chrysosporium was developed to the pilot plant scale [-617]. The process (mycelial color reduction, MyCoR) utilizes a rotating biological contactor on the surface of which the fungus is immobilized. In the MyCoR process, chlorolignins are somewhat degraded, and low molecular mass chlorinated derivatives are metabolized. Extracellular LiP and MnP may play an important role in the bleaching process. The potential of the enzymatic pool, LiP and MnP, in particular, from Pycnoporus sanguineus cultures for decolorization of bleach kraft pulp effluents [618] has also been indicated. However, a recent study [619] has demonstrated the partial decolorization of E1 effluent by purified MnP, whereas LiP did not appear to have any role in this process. A CDH/CBO-based biosensor has been developed by Elmgren et al. 1-620]. CDH is immobilized or cross-linked in a redox polymer matrix containing osmium(III) on a rotating disc electrode functioning as a biosensor. A combination biosensor, utilizing glucose oxidase in addition to CDH, has also been developed. The biosensor is coupled to a chromatographic column, and provides for the identification of cellodextrins 1-621].


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R e s e a r c h a n d d e v e l o p m e n t i n v o l v i n g e n z y m e s i n v o l v e d in l i g n i n d e g r a d a t i o n is p r o g r e s s i n g w i t h c o n s i d e r a b l e p a c e , a n d o n e c a n see t h e p o t e n t i a l o f l i g n i n o l y t i c e n z y m e s e i t h e r a l o n e , in m i x t u r e s , o r p e r h a p s p a r t i c u l a r l y in c o m b i n a t i o n w i t h r e a d i l y o x i d i z e d s u b s t r a t e s ( m e d i a t o r s ) for t h e b l e a c h i n g of w o o d pulp.

Acknowledgements. The authors wish to express their appreciation to the Department of Biotechnology (Government of India). Ministry of Science and Technology, New Delhi for the DBT Overseas Associateship award to R.C. Kuhad, and to the University of Delhi for grant of duty leave. We also extend our sincere thanks to D.E. Akin and G. Henriksson for their suggestions during the preparation of this chapter. For the drawing of Fig. 6 we are indebted to Paul Simon.

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Lignin Dimitris S. Argyropoulos and Sam Ben Menachem Department of Chemistry, McGill University, Pulp and Paper Research Centre, 3420 University Street, Montreal, Quebec, Canada, H3A 2A7

1 Occurrence and Role of Lignin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1 W o o d Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Synthesis of Precursors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 The Dehydrogenation Stage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 The Radical Polymerization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Lignin - Carbohydrate Connectivity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Lignin Architecture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 The Gel Degradation Theory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Possibility of Order in Lignin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Solution Properties of Lignin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Lignin Associative Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Lignin Polydispersity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Lignin Preparations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Laboratory Preparations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Commercially Produced Lignins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Methods of Lignin Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1 Degradative Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Non-Degradative Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

128 128 130 130 132 133 137 139 141 142 144 145 145 146 146 147 147 147 149 151

The objective of this chapter is to provide the reader with a general account of various aspects of lignin chemistry with emphasis on issues that have seen rapid growth during the past two decades. This is accomplished by describing the research efforts of over 360 literature citations, embarking from early concepts and concluding with the current views on the subject. After a general introduction that deals with the occurrence and role of lignin in the cell wall and within woody tissue, recent advances of lignin biosynthesis are discussed commencing with a description of the metabolic pathways that determine the synthesis of the various lignin precursors. The main reactions leading to the various bonding patterns in lignin are then discussed including accumulating evidence that pertains to the connectivity of lignin to carbohydrates. The overall architecture of the lignin macromolecule is then dealt with by critically examining various aspects of the early literature and recent scientific evidence that points to the possibility of order in it. After a brief description of the m e t h o d s available to isolate lignin, the chapter concludes with an outline of the various major procedures currently available for its structural determination.

Advancesin BiochemicalEngineering/ Biotechnology,Vol. 57 ManagingEditor: Th. Scheper 9 Springer~VerlagBerlinHeidelberg 1997

128

D.S. Argyropoulosand S.B. Menachem

1 Occurrence and Role of Lignin Second only to cellulose, lignin is amongst the most abundant biopolymers on earth. It is estimated that the planet currently contains 3 x 1011 metric tons of lignin with an annual biosynthetic rate of approximately 2 x 10 l~ tons [1, 2]. Lignin constitutes approximately 30% of the dry weight of softwoods and about 20% of the weight of hardwoods [3]. Lignification is associated with the development of vascular systems in plants, providing resistance to biodegradation and environmental stresses such as changes in the balance of water and humidity [4]. Lignin is absent from primitive plants such as algae, and fungi which lack a vascular system and mechanical reinforcement. The presence of lignin within the cellulosic fibre wall, mixed with hemicelluloses, creates a naturally occurring composite material which imparts strength and rigidity to trees and plants. An additional role for lignin has recently been revealed [5-7] involving complexes of lignin phenolic acids in forage legumes and grasses. The presence of lignin phenolic acids is thought to inhibit the digestion of potentially digestible carbohydrates by ruminants. Industrially one encounters lignin during the process of paper making, which involves the chemical or mechanical separation of the cellulosic fibres from woody or other lignified plant material. The chemical separation of lignin from cellulose has been termed "delignification" and it is one of the complex processes of the pulp and paper industry [8]. The process of delignification results in the production of vast amounts of lignin whose properties may vary depending on which delignification process has been employed and at which stage of delignification the lignin was isolated. In general, the lignin by-products resulting from the process of wood delignification are of a polymeric nature that may serve, after some modification, as additives in various formulations involving adhesives, thermosets or thermoplastics. Although chemical pulping is a long established practice, many controversial questions remain concerning the structure of native lignin (protolignin), its reactions during delignification and bleaching [9]. To expose the problem further requires an understanding of wood anatomy, lignin structure and biosynthesis and the way lignin is linked to carbohydrates in the cell wall. Since wood is the predominant raw material for the pulp and paper industry the focus of this chapter will be toward understanding wood derived lignins as opposed to those from annual plants.

1.1 Wood Structure Figure 1 shows the detailed macroscopic composition of a softwood fibre. The cellulose microfibrils are arranged characteristically within the various layers in the cell wall, namely; the primary wall (P), the secondary wall, and the middle lamella (ML). The secondary wall is further subdivided into three sublayers ($1, $2, and $3) each comprised of cellulose microfibrils in distinct orientations with reference to the main fibre axes. The $1 and $3 layers are both 0.1~).2 gm in

Lignin

129

Fig. 1. D i a g r a m m a t i c r e p r e s e n t a t i o n o f a softwood tracheid (Reprinted with permission from C6te [12])

thickness while the S 2 layer is about 1 10 gm thick containing within it 80-95% of the cell wall material. The primary wall, (0.1 gm), contains a network of microfibrils and significantly higher lignin concentration than the secondary wall, while the middle lamella is composed predominantly of lignin [10]. Despite the fact that the concentration of lignin in the middle lamella is extremely high, it is the secondary wall that contains about 70% of the overall lignin present in wood due to its large volume [11]. Chemically, lignin is built from phenylpropane units linked together by different bonds. A description of how the different bonding patterns emerge will follow. However, at this point it is essential to mention the fundamentals of numbering the various carbon atoms in lignin. More specifically, while regular numbers are used for labelling the aromatic carbons, Greek lettering is used to label the side chain of the phenylpropanoid. The illustration of Scheme 1 may assist the reader in comprehending the meaning of the various bonding patterns described in latter parts of this chapter. "~C H 2 O H

R = another phenyl p r o p a n e unit

I

[3 C H R

R " = H or R

I

ct C H R ....

R"

R " ' = O H or R

R' OR"

guaiacyl :

R' = OCH3, R " = H

syringyl :

R' = R " = O C H 3

para-hydroxyphenyl :

R' = R " = H

S c h e m e 1. T h e e l e m e n t a r y p h e n y l p r o p a n e b u i l d i n g b l o c k s o f v a r i o u s lignins

130


2 Biosynthesis Lignification is believed to occur in the intracellular layers of the cambium, where the sapwood and the bark layers of the tree merge [13]. Tracer experiments [14] and UV-microscopic observations [15] have shown that lignification is initiated within differentiating wood cells and extends to the intracellular areas, i.e. primary and secondary cell wall. Terashima [16] has shown that lignification and cellulose deposition in the plant cell wall proceeds in three distinct phases. Initially lignification occurs at the cell corner and middle lamella, after the deposition of pectins is complete and the formation of the secondary wall $1 has been initiated. During the second phase, an extensive deposition of cellulose microfibrils, mannan and xylan in the $2 layer takes place. Yet, the lignification process proceeds very slowly during this stage. Finally, during the third phase, lignification proceeds extensively. This occurs after the deposition of cellulose microfibrils in the $3 layer of the secondary wall has taken place.

2.1 The Synthesis of Lignin Precursors In 1971, it was demonstrated that lignin is synthesized from/-phenylalanine and cinnamic acids [15]. These acids are derived from carbohydrates through the shikimic and cinnamic acid pathways. Supporting evidence for this route was obtained when radioactive glucose was administered into plants, producing shikimic acid [17] and radioactive lignins [18-20]. Lignification proceeds with the conversion of/-phenylalanine to form trans-cinnamic acid (Scheme 2). This deamination process is catalyzed by l-phenylalanine ammonia lyase (PAL), a key enzyme found only in plants that can synthesize lignin [21,22] and some cinnamic acid derivatives [22]. It is worth mentioning that an additional enzyme, tyrosine ammonia lyase (TAL), which catalyses the formation of pcournaric acid from /-tyrosine, is characteristically found only in grasses [23-25]. The presence of this enzyme may account for the presence of p-coumaryl alcohol as an additional lignin monomer as well as esterified p-coumaric acid present mainly in grasses. As lignification proceeds, cinnamic acid is hydroxylated to p-coumaric and caffeic acids by specific hydroxylase enzymes [26, 27]. The caffeic acid thus formed, is then methylated to ferulic acid by O-methyl transferase (OMT) [28-30]. Up to this point the biosynthetic pathways for softwood and hardwood lignins is believed to be common [15]. However, they seem to diverge beyond it [31, 32]. This is because OMT enzymes, of different functionality, possessing different substrate specificities were found to be present in softwoods and hardwoods [33, 34]. One of the reasons which accounts for the almost exclusive presence of guaiacyl lignin in softwoods, is that the monofunctional OMT enzyme is inhibited competitively by caffeic acid (Scheme 2). For hardwoods, the

Lignin

131

difunctional OMT is inhibited via a feedback mechanism involving 5-hydroxyferulic acid [-34, 35]. This limits the concentration of this intermediate and the production of ferulic acid, thus restricting the formation of syringyl precursors. Another key enzyme which is responsible for the differentiation of the G and G,S pathways, is the ferulic acid-5-hydroxylase. The absence of this enzyme in gymnosperms also accounts for the near exclusive formation of guaiacyl precursors in softwoods. The synthesis of lignin precursors is complete with the reduction of ferulic and sinapic acids to the corresponding coniferyl and sinapyl alcohols. These reactions are catalysed by three specific enzymes which constitute the reductase H o OH H. CH20H H00C_+_NH2 ~OH OH~ Q shikimicacid CH2 CAH ~ ~ [ pathway 0 PAL O H ~ OH COO~/ H~J~-OH OH glucose H-H~n~H 1-phenylalanine cinnamicacid p-hydroxycinnamicacid

shikimicacid

CMS1

oyoH

competitiveinhibition (in gymnosprems)

~feedback

inhibition (in angiosperms)

( J"1

2 3 4 5 6

OMT 1

o.3,-0"

o..ro.

CH2OH

OCH3~OCH 3 q OH sinapylalcohol

OH caffeicacid

"3COocH3 OH sinapic a c i d

FA. OH 5-hydroxyferulic acid

dOCH OH femlic acid

Rs]

1-phenylalanineammonialyase(PAL) cinnamicacid 4-hydroxylase(CAH)* 4-coumarate3-monooxygenase(CMS)* o-methyltransferase(OMT) ferulicacid 5-hydroxylase(FAH)* reductasesystem(RS)*

CH20H .A d

OeH OH coniferylalcohol *These abbreviations have been used for brevity purposes only Scheme 2. A simplifiedmetabolicpathway of 1-phenylalanineto lignin precursors

132


system [32, 35]. The enzymes which were recently isolated from various plants [36, 37] include the hydroxycinnamate-CoA ligase, hydroxycinnamoyl-CoA reductase and hydroxycinnamyl alcohol dehydrogenase. The ligase and reductase enzymes of angiosperms and gymnosperms respectively, were found to show pronounced differences in substrate specificities [30, 38]. In gymnosperms, both enzymes were found to be inactive with sinapic acid and sinapoyl-CoA [39,30]. The lack of activation or reduction of sinapate in gymnosperms contributes toward the exclusive formation of guaiacyl lignins in them. The lignin precursors formed are of low solubility and are readily oxidized; in the cell wall they are stabilized as glucosides. The typical glucoside for softwoods is coniferin and its structure is shown in Scheme 3.

2.2 The Dehydrogenation of the Precursors Early investigations [40, 41] coupled with the more recent results of Harkin and Obst [42] have shown that the dehydrogenative polymerization oflignin monomers in plants is caused by a class of enzymes called peroxidases, or the peroxidase-H202 system. These enzymes, are capable of abstracting a proton from the phenolic hydroxyl of the precursor molecules, creating resonance stabilized free radicals. Examples of such radicals, typical of softwood lignin are depicted in Scheme 4 [42, 43]. More specifically, the dehydrogenation reaction involves hydrogen peroxide as the electron-accepting substrate for the peroxidase [45, 46]. In addition, superoxide radicals were suggested to form via the reduction of oxygen by N A D (nicotinamide adenine dinucleotide) [47]. Recent evidence suggests that hydrogen peroxide is produced by the peroxidase enzyme itself, and may play a key role in the control mechanism [47]. An enzymatically controlled cycle for the CH20H H ~ " r - O. w- O ---~,

,~-----CH

H

H

()H

Scheme 3. Coniferin,a coniferylalcohol glucoside

CH30

CHzOH I CH II CH

CH2OH I CH II CH

CH2OH I CH II CH

CH2OH I CH II CH

CH2OH I 9CH I CH

Peroxidase OH

OCH3

O

H3

Scheme 4. Resonanceforms of softwoodlignin phenoxy radicals (43,44)

3

3

Lignin

133

production of hydrogen peroxide, has also been reported [42]. However, excess peroxide was found to inactivate the peroxidase by the formation of a complex of an unknown structure [-48]. Recent research has also shown that the presence of laccase-like phenoloxidase can be correlated to lignin biosynthesis in Zinnia elegans stem tissues [49]. A review of the recent literature regarding plant laccases and selected fungal laccases and their role during lignin biosynthesis has been compiled by Dean and Eriksson [50]. After the oxidation of the monomeric alcohols to phenoxy radicals, the reaction changes dramatically; the reactions are no longer subjected to enzymatic control, but to a random polymerization process [51, 52].

2.3 The Radical Polymerization A major milestone in lignin chemistry was Freundenberg's success in providing experimental evidence for the enzyme-initiated dehydrogenative polymerization theory. This became possible by polymerizing coniferyl alcohol to a lignin-like dehydrogenation polymer (DHP) using liquid squeezed from the mushroom, Psalliota campestris, known to contain laccase and other oxidative enzymes [50]. Further attempts to synthesize lignin DHP's using the peroxidase/H202 system were successfully accomplished by Freudenderg [53, 54] and Sarkanen [55]. Yet, as Sarkanen pointed out, "these lignin polymer models", despite the fact that structurally were lignin-like, they were not identical to lignins formed in vivo [55]. When the in vitro polymerization was interrupted, structures of dimeric and oligomeric intermediates could be isolated and the lignin bonding patterns identified [56 58]. These experiments and others, involving sinapyl alcohol, support a random mechanism for the polymerization of phenoxy radicals [59 61]. The reactivity of such radicals and the probability of a specific mesomeric form to form a particular linkage depends on /3 electron spin density and steric considerations. Molecular orbital calculations of ~ electron spin densities of lignin model compounds [62] have shown that free electron spin densities are highest at specific sites within the phenylpropane unit. These reactive sites (Scheme 4) are the C1 and C5 positions of the phenylpropane unit, the phenolic hydrogen, and the aliphatic [3-carbon. Of these reactive sites, those with groups attached at C1 and C3 in coniferyl alcohol and C1, C3 and Cs in sinapyl alcohol, are less reactive. Alternatively, the phenoxy oxygen and the [3 carbon are considered as the most reactive species, readily coupling into aryl ether linkages [63, 64]. This may account for the high abundance of the [3-0-4 inter-unit linkages in lignin, estimated to be as high as 50% in softwoods and almost 60% in hardwoods [64]. The polymerization process proceeds with the coupling of the different resonance structures. Amongst the products, highly reactive quinone methides (Scheme 5) are formed which further react by addition to various nucleophiles. Free radical exchange reactions, such as hydrogen abstraction from suitable portions of other molecules, are also possible [65, 66].

134

D.S. Argyropoulos and S.B. Menachem CIH2OH~__xH3 HC - O X-~.ffx- CH= CH - CH2OH

/

O'~-CH=

With coniferyl alcohol OH

CH-CH2OH

OCH 3 OCH 3

Ct, 13ether structures

OCH 3

.c-o% H2COH )='xOCH3 [H20]

HC - O _ _ ' ~,,)- CH= CH - CH2OH HC- OH

O

Quinone Methide

~OCH OH

3

~ - O - 4 structures

H2COH ~OCH3

[glucose]

,

HC -O ~ C H =

CH-CH2OH

Lignin carbohydrate bond Scheme 5. Addition reactions to a quinone methide leading to the formation of the various interunit linkages in lignin

The first dimeric lignol to be isolated and identified was a phenylcoumaran structure analogous to that of dehydrodiconiferyl alcohol [67]. This structure, which comprises about 10% of the lignin units, is formed by the coupling of two free coniferyl alcohol radicals centered at C5 and C 0 positions of the phenylpropanoid (structures 2 and 4 in Scheme 4), giving rise to a dimeric quinonemethide. This coupling followed by aromatization of the ring allows the creation of an a-O-4 linkage (Scheme 6). The C~-C5 bond thus formed is actually quite stable under pulping conditions, as long as its phenolic group is etherified. Other bonding patterns that occur in lignin are the so-called "diaryl propane" units or 13-1 (Scheme 7). They are present in about 5-10% of the total phenylpropane units in lignin and they are thought to be relatively stable under alkaline pulping conditions. Ether and ester linkages are also common in lignin. With respect to ether linkages, several types have been identified to be present, namely: biaryl ethers, non-cyclic benzyl alkyl ethers, 13-O-4 ethers and diphenyl ethers. Biaryl ethers

Lignin

135 H2COH I HC II HC

H2COH HC II HC

HC ~H"~-"i"f OCH3 HC 44.._-.--0

Ioc.3

. c , / --y "oc. HC

~OCH OH

0

0

3

Scheme 6. The phenylcoumaran, (C~-C5)interunit linkage of lignin

R

o.

HCOH R~OCH3 OH

\OCH3 R=H, or OCH3 Scheme 7. The diaryl propane (13-1)interunit linkages of lignin

are generated by the combination of an o-methylquinonoid (structure 2 in Scheme 4) with a phenoxy radical [60, 68, 69]. They comprise only about 6% of the phenylpropanoid units in spruce lignin [60, 68]. Non-cyclic benzyl alkyl ethers, present in relatively small amounts in lignin (2-3%), have been cited for their beneficial effects during pulping [70]. However, recent attempts to detect them in milled wood lignin, using advanced N M R techniques, have failed [-81]. The combination of C~ and phenoxy radicals (Scheme 5) results in the formation of 13-O-4linkages [68, 60, 69], linking 3 ~ 5 0 % of the lignin units [64]. Structures which contain ~, 13ethers are relatively infrequent and have been estimated to account for less than 1% of the interunit linkages in lignin. However, recent efforts using advanced N M R techniques have questioned these levels [81,364]. Another lignin structure, containing two ether groups is the pinoresinol (present in less than 5% of the units) [71]. This structure, is formed by the [3-13 coupling of two coniferyl alcohol radicals followed by double ring closure, as illustrated in Scheme 8. Benzyl alcohol groups are also common structures in lignin. These, which account for more than 30% of the phenylpropane units, are formed by the addition of a water molecule to a quinonemethide. Like aliphatic esters, the benzylic structures are also beneficial during alkaline pulping [72] since they facilitate cleavage of the arylglycerol-[3-aryl ether residues under soda or kraft pulping conditions with the intermediate formation of epoxide or episulfide structures. 5-5'biphenyl and diarylmethane structures which link two aromatic units either through the C5-C5 or Ca-C5 positions,

136

D.S. Argyropoulos and S.B. Menachem

i~ ~ O

1

CH2OH

.2?

CH3Of~[

I

~o~~

cn30

coniferylalcoholradicals

'

H C - - CH I I HC~ CH2

CH2OH

o

OH .fOCH3

~,,~OCH~I

.,o~"~ HzC CH I HC- - CH I

I

C H 3 0 ~ O/CHz OH pinoresinol

Scheme 8. Formation of pinoresinol linkages in lignin CH2OH CH2OH CH CH II II CH CH

CH30~OCH3 OH OH

Scheme 9. The biphenyl 5 5' interunit linkage of lignin

contain carbon-carbon bonds, which are stable under pulping conditions. While 5-5' biphenyl units are present in the original wood the diarylmethane units are formed by the condensation of two aromatic rings in lignins during alkaline and/or kraft pulping (Scheme 9) [73]. As such, their importance becomes of significance in technical lignin preparations. The frequency of occurrence of such structures is obviously higher in pulped softwoods, since their lignin is composed almost exclusively of guaiacyl units which possess aromatic units that contain a free C5 position. Recently Brunow's group announced the discovery of another bonding pattern present in softwoods lignin [74, 75]. This involves the formation of a, [3 ethers on the same 5-5' biphenyl structures. The new octagonal moiety has been identified as the dibenzodioxocin of Scheme 10. Based on these structural details, a number of lignin models have been proposed by a variety of investigators. Amongst them one may cite those of Freudenberg [68], Brauns [76], Erdtman [77], Adler [78], Forss and Fremer [79], and Glasser and Glasser [80]. Amongst the models proposed for softwood lignin, Freudenberg's is still the most widely cited [53, 68]. A very significant analytical effort has been carried out through the past several decades attempting to quantify the various bonding patterns in softwood and hardwood lignins. The results of some of these efforts are now summarized in Table 1.

Lignin

137

O

',

OH

(

O

O.~.

R = CH2OH, CH 3

Scheme 10. Dibenzodioxocin present in softwood lignin

Table 1. The frequency of the major linkages in softwood and hardwood lignins* [8, 55, 64, 81-84]

% of Total Phenylpropane Units Type of Linkage

In Softwoods

In Hardwoods

[3-Aryl ether ([~-O-4) a-Aryl ether (a-O-4) Diphenyl ether (4-0-5) a-Alkyl ether (c~-O-7) Biphenyl (5-5') 13-1 Pinoresinol (13-[3) Phenylcoumaran (f3-5) Lignin-carbohydrate links Dibenzodioxocin

45~48 6-8 3.5 8 small 9.5-17 7 10 3 9 12 not determined >>

60 6-8 6.5 small 4.5 8 6 not determined >>

* The numbering of the carbons used in the nomenclature of this table is shown in Scheme 1.

2.4 The Lignin-Carbohydrate Connectivity Detailed studies of wood and grasses point to the likelihood that lignin is not simply deposited in the cell wall of polysaccharides but is closely linked and associated in a certain architecture with the carbohydrates [85 88]. A considerable research effort has been carried out in studying the nature of the actual chemical bonds between lignin and carbohydrates [89-91]. Advances in the technology of nuclear magnetic resonance spectroscopy coupled with a better understanding of the nuclear spin relaxation mechanism in solids, have provided a powerful tool for probing the molecular motion of carbohydrates and lignin in solid wood and pulp samples [9~95]. By using ferric ions as a probe, coupled to proton spin-lattice (T1) and spin-lattice in a rotating frame (Tlo) relaxation time measurements, Gerasimowitz et al. [94] derived information related to the structural connectivity of carbohydrates and

138


lignin in wood pulp. Similar linkages were also invoked by Kolodziejski et al. who studied the I3C CP/MAS NMR spectra of lodgepole pine wood [96]. By measuring the proton spin-lattice relaxation times of progressively sulphonated and methylated wood and pulp samples, Argyropoulos and Morin evaluated the molecular mobilities of lignin and carbohydrates in the presence of different counterions [95]. Lignin-carbohydrate associative interactions were invoked, once again, to rationalize for their observations. On another front, significant advances in clarifying this issue have been made by carrying out in vitro lignification experiments in the presence of model compounds. Small sugar molecules were found to undergo addition reactions resulting in the formation of ether or ester linkages [97-100]. Early work by Freudenberg demonstrated that sorbitol or sucrose can readily add to lignin, forming benzyl alkyl ether linkages [101]. These bonds were found to be further stabilized when the phenolic hydroxl group of the lignol portion was etherified. Such an etherification step is not uncommon to occur via a dehydrogenative mechanism during lignification. Such an ether linkage, which seems to be considerably more stable than those of glucosides, is considered to be one of the main reactions leading to a stable crosslink between lignin and plant polysaccharides. Another lignolsugar that has been isolated by Harkin and Freudenberg from in vitro lignification experiments, is the guiacylglycerol-[3,y-disucrose ether [98]. The formation of this ether is believed to follow a radical mechanism which could be associated with that of lignin biosynthesis. According to this mechanism, coniferyl phenoxy radicals (catalysed by peroxidases) abstract a hydrogen from a sugar molecule to generate a sugar radical. This, quickly adds to another lignin C~ radical. The nature of the resulting bond, (C-C or ether) depends upon the formation of a carbon radical or an oxygen radical on the sugar molecule. However, the attachment of two sugar units on a phenyl propane unit is an unlikely event when one considers that during lignification a variety of competing nucleophiles are present, including water and other lignols. More recently Leary and coworkers have shown that sugars will readily add to monomeric lignin quinonemethides or benzyl alcohols forming predominantly C~ linked carbohydrate benzyl ethers or esters [102]. Ester bonds are also involved in the attachment of lignin to the plant carbohydrate moiety [103, 113]. Their importance is very significant for plants and grasses and somewhat less for wood. Ferulic acid in grasses is known to be esterified with carbohydrates, and etherified with lignin. However, the topochemistry of its attachment to lignin is not well understood [-104-106]. Similarly, p-coumaric acid is known to be extensively esterified with lignin, but the regiochemistry of lignin acylation is still a matter of debate [107 109]. The following description of events represents the state of our knowledge as far as the biosynthetic pathways leading to such species are concerned. The a-position of quinone methides, formed during the dehydrogenative polymerization process [64], apart from being attacked by water, may also be attacked by free acids and alcohols, leading to a-esters and a-ethers. This is the case of feruloyl esters and phenolic glycosides of p-coumaric acid, where the free phenol or the free

Lignin

139

carboxylic acid groups respectively, may trap quinone methides by an addition to yield a-ethers and a-esters [110, 111]. Alternatively, feruloyl esters can directly participate in the free radical polymerization process giving rise to a number of different structures [112]. Furthermore, enzymatically pre-esterifled p-coumaric acid with p-hydroxycinnamyl alcohol monomers, may enable the formation of p-hydroxycinnamyl p-coumarates which could participate in the formation of the lignin macromolecule by conventional oxidative coupling reactions to yield 7-p-coumaroylated lignin [-108]. While ether-linking ferulic acid to the a position of the lignin side chain, via "opportunistic" quinone methide trapping is still speculative, it has been reported by Ralph et al. [104] that feruloyl esters, if present in the lignifying matrix, are capable of participating in the free radical lignification process. Moreover, the identification of new ether-linked ferulic acid-coniferyl alcohol dimers in grass straws, by Jacquet et al. [106], demonstrates the occurrence of radical coupling reactions between ferulic acid and coniferyl alcohol to yield 13-aryl ether structures. As noted above, the lignin-carbohydrate bonds are possible due to the reaction of quinone methides with various lignols and carbohydrates during lignin biosynthesis. Despite this there is still controversy whether these bonds occur between lignin and cellulose or between lignin and hemicelluloses. This is certainly not an easy issue to resolve but the evidence is more in favour of a predominant lignin-hemicellulose connection. Since in hardwoods the predominant hemicellulose is xylan and considering that about 90% of this sugar can be removed by a mild alkaline treatment, ester links have been proposed to occur between the lignin and the C6 uronic acid groups of the sugar [85, 114].

3 Lignin Architecture Amongst the features of any delignification experiment is the observation that the molecular weights of the solubilized lignin become progressively higher as delignification proceeds [114 125]. Molecules of relatively small size become soluble early in the delignification process, while larger fragments appear in solution in latter stages. This usually occurs when lignin in wood is made soluble by chemical treatments [115, 116, 126, 127]. Szabo and Goring in 1968 recognized this effect and proposed the gel degradation theory to account for these observations. However, prior to their proposition a number of other suggestions were made. These will now be briefly discussed in order to expose some additional salient features of lignin's polymer chemistry. The conjugated phenolic nature of the lignin precursors (Scheme 1) allows for the possibility that condensation reactions occur under delignification conditions. It is thus possible that the molecular weight of soluble lignin may increase if lignin in wood consists of finite macromolecules with reactive sites that

140


condense under acidic or basic [128-131] delignification conditions [132-136]. This is generally known as the "condensation theory". With respect to the condensation theory, a point of major criticism emerges from the results of Adler et al. [137], Felicetta and McCarthy [138], and Yean and Goring [116], where a decrease in the molecular weight of soluble lignosulfonates was observed upon a second exposure to the reaction conditions. This contradicts the condensation theory because one would expect, on further reaction, either an increase or no change in the molecular weight. Nevertheless, it is now accepted that condensation reactions occur during delignification. Model compound studies have suggested that such reactions operate to some degree under both acidic [128] or basic conditions of delignification [129]. During acidolysis, the formaldehyde formed from lignin has been found to be partly consumed in condensation reactions [139]. Quantitative gel filtration studies on acidolysis products [128], however, have revealed that the fraction of "unidentified condensation products" amounts up to 14% by weight of the isolated lignin. It is thus more prudent to conclude that condensation possibly occurs during delignification [140]. Its effects, however, are only secondary contributions to the molecular weight increase phenomenon observed in sulfonic acid delignification. Evidence substantiating this suggestion has been given by Argyropoulos and Bolker [141]. In their communications, Brown et al. [142, 143] concluded that lignin in wood consisted of finite molecules, with two major components of different molecular weights. Similarly, Obiaga et al. [144] postulated that lignin in wood existed as an assembly of finite entities with a degree of polymerization equal to 18. Both conclusions were based on the apparent bimodal distributions observed when lignin samples were chromatographed on Sephadex gels. These interpretations were later proved to be incorrect by Bolker et al. [145]. The peak at the high molecular weight end of the chromatograms was shown to be an artifact, the effect of a high molecular weight tail manifesting itself at the exclusion volume of the Sephadex gel. Later results of Mbachu [146] further strengthened the findings of Bolker et al. A low exclusion volume peak has indeed been reported by Cooper [147] for high molecular weight polystyrene. Therefore, condensation reactions most likely occur during delignification, thus producing a network polymer which must then be degraded during delignification. The effects of condensation, however, are secondary contributors to the molecular weight increase phenomena observed during wood delignification. Another hypothesis attempting to rationalize the molecular weight increase of solubilized lignins, is the pre-sieving concept [163]. According to this hypothesis the lamellar structure of the cell wall or more specifically the pore size, determines the size of the dissolved lignin macromolecules when wood is delignified. This is particularly true during the washing of kraft pulp when high molecular weight lignins diffuse out of the fibres [148, 149]. Lignin fractions, which turned to be of the order of hundreds of thousands [150], would be too large to pass through the porous structure of the cell wall. Similarly it was found that when lignin macromolecules were spread on the surface of a non-solvent [121,152] or deposited on a carbon coated grid for electron microscopy [153], they assumed

Lignin

141

a thickness of 2 nm, which is considerably larger than the pore size. Only after the size of the pores becomes larger, as lignification proceeds [148] could these fractions diffuse out of the cell. This may account for the presence of small molecules early in the delignification, while larger molecules are released later, as delignification proceeds. This theory was criticized by Bolker et al. [,145] on the basis of the work of Bogomolov et al. [123], who found that the molecular weights of milled wood lignins (see Sect. 5.1) increased with increasing yield. This could not have occurred if the cell wall pores were responsible for the sieving action. In milled wood there is no longer need for lignin fragments to diffuse through any pores because the wood has already fragmented to sizes much smaller than the size of individual cells.

3.1 The Gel Degradation Theory The pioneering work of Szabo and Goring [127], which was followed by Bolker and Brenner [155], and Bolker, Rhodes and Lee [145], dealt with the gel degradation concept of delignification. In broad terms, the Szabo and Goring treatment envisages delignification as the reverse of polymerizing a trifunctional network, while the Bolker and Brenner approach is the reverse of polymerizing a network formed by the random crosslinking of monodisperse preformed chains. The difference between the two treatments arises from the assumption of Szabo and Goring that all ether bonds in lignin are of comparable reactivity [-145, 156]. Bolker and Brenner, however, proposed that the sites of crosslinking were the benzyl ether groups, which are expected to be the most reactive of all the ether linkages found in lignin. They further invoked a monodispersity approximation for the primary chains of lignin. They based their argument on Hess' measurements of the molecular weight distribution of Brauns' native lignin [157]. According to the gel degradation theory, Brauns' native lignin represents the final sol fraction remaining from the gelation process of lignin. Accordingly, Bolker and Brenner argued that Brauns' native lignin mainly comprised primary uncrosslinked chains, with a monodispersity ratio (as they calculated from Hess) of Mw/M,, = 1.04. Yan [158], not fully convinced of the validity of the monodispersity approximation, extended the Flory-Stockmayer theory so as to include the crosslinking of primary chains with any initial size distribution. In a series of three papers Argyropoulos and Bolker undertook to experimentally test the reversibility of the Flory-Stockmayer statistics of gelation by using synthetic model networks [159 161]. The results of their efforts have provided support not only for the gel degradation theory but supplied information on the solution properties and the molecular weight species distribution expected from the random degradation of network polymers [162]. Evidence of the validity of the network structure of protolignin arrives from the viscosity, polydispersity, and molecular weight data of soluble lignins discussed previously. The random degradation of a crosslinked network is expected

142


(according to the gel degradation theory) to yield polymeric fragments of spherical configuration whose molecular weights, yields, and polydispersity ratios will increase as bond fragmentation intensifies within the network. The network theory of protolignin finds progressively wider acceptance among researchers in the field as the experimental evidence pointing toward its validity grows. Although there is still lack of agreement on the precise architecture of protolignin, major efforts toward solving this controversial issue are emerging from the application of the gel degradation theory and its various modifications [158, 164].

3.2 Possibility of Order in Lignin Despite the fact that a significant compilation of scientific evidence has been accumulated over the years, pointing to lignin as an amorphous, crosslinked network polymer [165 167], formed via a random free-radical polymerization process [169 172], recently discovered aspects of lignin chemistry and details of its deposition into the cell wall have challenged the universality of this description [168]. Detailed analytical studies of synthetic (DHP's) and milled wood lignins have documented that such preparations are rather distinct as far as their bonding patterns and composition is concerned [171]. However, synthetic lignins prepared in the presence of carbohydrates, under conditions that mimic the micro environment of the cell wall, were found to resemble native lignins [173]. These observations, coupled with studies of the organization of lignin within the woody tissue [85-88], point toward the presence of an organizing influence operating during lignin synthesis and deposition [174, 175]. This dominant organising influence, according to several researchers [-176 181], can be provided by strong associative interactions operating between lignin precursors and the ordered polysaccharidic matrix. Experimental evidence for these interactions has been provided by studying the adsorption of oligolignols onto cellulose [182] and by measuring the photoconductivity of wood tissue

[183]. While the cellulose microfibrils are known to be deposited in the cell wall in complex ordered patterns [,184, 185], the precise deposition patterns of hemicelluloses remain unknown. Evidence pointing toward the possibility of hemicelluloses being aligned to cellulose [186, 187] and linked to lignin [,188] has been provided. These observations coupled with the associative interactions to be discussed later (Section 4.1), and the observation that the cellulose microfibrils are closely packed [-189-192], are factors that may indeed contribute to some organization within lignin. In actual fact, Jurasek in his recent computational efforts, aimed at modelling lignin, has showed that the spatial constraints of the cell wall may indeed impose some degree of regularity in lignin [193]. Gravitis and Erins [194] have also examined lignin with various theoretical

Lignin

143

conformational models and have arrived at the conclusion that "under some conditions quasi-ordered regions of lignin structure can be expected". On a different front, Atalla has provided evidence indicating that the aromatic rings of lignin are aligned tangentially to the secondary wall [195, 196]. Based on these observations, he proposed a new model for the assembly of lignin [174]. The model suggests that variations in the structure of hemicelluloses, may result in corresponding systematic changes in the structure of lignin and cellulose [175, 197-199]. While cellulose provides the primary framework, hemicelluloses furnish various branches which associate themselves with specific lignin precursors. More specifically, it would be anticipated that specific monosaccharide branches are designed to organize the monolignols, while di- and trisaccharidic branches are designed to selectively bind specific di- and trilignols. As such, lignin precursors may be subjected to a certain regulatory mechanism which involves steric factors and sugar binding specificities. Perhaps the most attractive feature of this model is that it introduces a hierarchic and sequential control that occurs at multiple levels, in different phases and separate locations throughout the biosynthetic pathway. This model is different from traditional models which emphasize the compartmentation of the process [200] involving different extracellular or membrane bound enzymes [201-203]. Other evidence pointing toward lignin possessing certain orientation with respect to the cell wall has been provided by its ability to conduct electric charge as witnessed by photoconductivity measurements of wood [183, 204]. Since conductance is highly depended on the coherent orientation of the structures involved in the conduction, the ability of lignins to carry ionic charge has been taken as proof for the presence of a regular array of similar functionalities which may become the vehicle for the excitation transfer of electric charge [-205-207]. A reasonable explanation for the observed photoconductivity in wood is that the regularity of the surfaces in the polysaccharidic matrix probably imparts to the lignin a coherence of spatial organization that is sufficient to facilitate some interactions or coupling between its lowest unoccupied molecular orbitals. In 1994 a report of "visual encounter" of order in lignin appeared [208]. Using a scanning tunnelling microscope (STM) images of building units or modules assembled into larger assemblies were claimed to have been recorded. During the same year Faulon and Hatcher [209] presented their calculations that suggested the biopolymer to have a helical structure, characteristic of many naturally occurring macromolecules. When one considers most of the evidence presented above, it becomes apparent that a better paradigm for lignin needs to be developed. As Goring concluded [210], one should distinguish between lignin in the middle lamella and that of the secondary wall. Until the lignification process has been fully understood, the traditional concept of lignin being a uniform, amorphous, three dimensional network polymer, is probably too simple to reconcile with all the recent scientific data relating to its native structure and properties.

144


4 Solution Properties of Lignin Early attempts to study the properties of lignin in solution commenced with the observation that native lignin is insoluble in all good solvents [-211]. In 1956, Bjorkman [212] discovered that when spruce wood is milled thoroughly, up to 50% of the lignin could be degraded and extracted in aqueous dioxane. During the milling process, covalent bonds were found to rupture [-213], and low molecular weight lignin fractions solubilize. This behaviour was also described by Lindberg [214] and Schuerch [-211]. In 1960, Gupta et al. [215, 217] pointed out that by invoking a three dimensional polymer network architecture for lignin, the molecular weight distributions of isolated solubilized lignins could be accounted for (Section 3). The soluble macromolecules believed to reflect the properties of the network from which they emerged. Amongst the most significant of the early observations were the predominant physicochemical characteristics of lignin molecules in solution. These include the intrinsic viscosity, branching parameter, and the degree of polydispersity. Their determination provided useful structural information in relation to the overall architecture of protolignin. The intrinsic viscosities [-216], at comparable molecular weights, were found to be 1/40 of those of polysaccharides and about 1/40 of other synthetic polymers [-11, 215, 217]. The low intrinsic viscosities of dioxane [218], kraft [-219], lignosulfonate [220], and alkali [215, 217] lignins, in various solvents, led Goring to conclude that these molecules in solution are compact spherical microgel particles [-11,218]. He also reported that the values of the Mark-Houwink exponent (a), ranged between 0 and 0.5. This constituted further confirmation that lignins in solution behave as molecules whose solvated shape is something between an Einstein sphere (a "ball" impermeable to solvent) and a non free-draining random coil. Similar conclusions were derived when other parameters, such as sedimentation coefficients and diffusion constants, were measured [221,222]. These measurements showed that soluble lignins are rather compact molecules in solution, though not as compact as simple solid spheres. In general the chains of the lignin macromolecules in solution are more densely packed than those of linear flexible polymers such as polystyrene. The branching parameter, or contraction factor (g) introduced by Zimm and Stockmayer [-223], when measured on various alkali lignin fractions, was found to decrease with increasing molecular weight [-217], as expected for such molecular configurations. Independently, Alekseev et al. [224] calculated the value of the Mark-Houwink exponent (~) for lignin solutions using viscosity, sedimentation, and diffusion data. His calculations supplied further support to the idea that the lignin macromolecules in solution are of a rigid spherical configuration [221,222]. More recent results of Pla and Robert [225] showed ~ to be 0.5 for dioxane lignin solutions. They, too, interpreted their results in terms of the branched or crosslinked nature of the dissolved lignin macromolecules. Low Mark-Houwink exponents are not unusual for truly branched macromolecules in solution. A good example of this can be found in the work of Argyropoulos

Lignin

145

and Bolker who have shown that the soluble fragments isolated from a series of polyester network polymers beyond the gel point showed an 0~value of only O.15 [226].

4.1 Lignin Associative Interactions Several authors have raised the possibility that lignin in wood is composed of low molecular weight molecules which on becoming soluble tend to associate by non-covalent linkages into complexes of higher molecular weight [227-232]. It was shown by S. Sarkanen [228, 232] that this process seems to be reversible and solvent dependent. Low molecular weight kraft lignin fractions generated higher molecular weight complexes in certain solvents. Such associative interactions among lignin components have been found to occur during gel permeation chromatography in different mobile phases. Non-aqueous solvents exhibited multimodal chromatograms, while alkaline aqueous eluents gave rise to broad chromatographic envelopes with sufficient resolution only in the low molecular weight region [233, 234]. Connors, Sarkanen and McCarthy [228] have discovered that in non-aqueous solvents, association complexes of kraft lignins gave rise to apparent molecular weights (as determined by GPC) as much as three orders of magnitude larger than their constituents. These effects were eliminated when LiC1 was added to the DMF used as the mobile phase. Accounts by Ekman and Lindberg [235] and Lindberg [236] show very major changes occurring in the solubility, hydrogen bond behaviour, and size exclusion chromatograms of lignins after they have been methylated or acetylated. Conclusions on extensive hydrogen bond formation between free hydroxyl groups in dioxane lignins have also been reported by Hatakeyama et al. [237] and Bogomolov et al. [238]. Obviously, any association between lignin molecules in solution would cause an increase in their apparent molecular weight values.

4.2 Lignin Polydispersity A considerable amount of scientific data exists supporting the fact that soluble lignins and derivatives are of high polydispersity [239-246]. The weight to number average molecular weight ratios for different lignins may vary from about 3 to over 10-11 [270, 2713, indicating that lignins are extremely polydisperse materials. Fractionation studies on lignosulfonates [114] and alkali lignins [241,217] have shown that they contain molecular weights ranging from 1000 to greater than a million (g tool- 1) [247]. The molecular weight distribution curves for lignins are expected to be unimodal. However, a number of authors have reported bimodal distributions for soluble lignin [227, 248 249]. Such behaviour was also observed when lignin was isolated from the pulping liquors of wood [227, 250]. It was only after improvements in chromatographic techniques that these bimodal distributions could be to resolved into

146


distributions with several maxima [251-253]. More information on molecular weight distribution within the soluble phase beyond the gel point, which according to the gel degradation theory should resemble those obtained during lignin degradation, can be found in the work of Argyropoulos and Bolker [162]. Recently, Glasser has produced a universal plot of polydispersity versus weight average molecular weight for a series of prototype lignins supplying a linear expression with some predictive power [270, 271].

5 Lignin Preparations Generally, there is a lack of agreement on the ultimate structure of lignin and this may be because, as Pearl wrote [254] "It is practically impossible to isolate two lignin preparations with identical properties, even by the same procedure". Although many methods are available for the isolation and purification of lignin, none provides 100% yield and structural authenticity. To isolate lignin which closely represents the native material, reactive chemicals and elevated temperatures must be avoided.

5.1 Laboratory Lignin Preparations Milled wood lignin (MWL) [255] and cellulase enzyme lignin (CEL) [256] are good examples of preparations closely resembling the native material. Yet, these preparations are never free of even minor chemical modifications. For example several secondary reactions may occur during the milling process, as a result of free radicals produced during the process [257]. Moreover, CEL preparations are known to contain protein impurities introduced during the enzymatic treatment. Despite these shortcomings, MWL and CEL preparations show minimal structural modification with yields ranging between 25%-66% of the total lignin and with carbohydrate contents ranging between 2-10%. The molecular weights (Mw) of these lignin preparations range between 15,000-20,000 and predominantly consist of alkyl-aryl ether linkages [258]. Other techniques for isolating lignin involve treatment of the wood with organic solvents such as dioxane [294] or ethanol [260, 261], sometimes in the presence of catalytic amounts of mineral acids (H/SO4), or inorganic Lewis acids, at elevated temperatures and pressures [262]. The combination of acids and organic solvents causes the hydrolysis of the ether bonds in lignin and those between lignin and carbohydrates. Such products are relatively free of carbohydrates, and of low molecular weight fragments [263]. Another technique involves the degradation of the cellulosic constituents of wood by sodium paraperiodate [259, 260] or their solubilization by complexation with cuprammonium hydroxide [261] (cuoxam lignin). The latter gives lignin in high yields

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which is known to retain the morphological features of wood [264, 265] and is totally insoluble in organic solvents. Amongst various lignin preparations, Fleming and Bolker found cuoxam lignin to be the most suitable for model de!ignification experiments [266]. For additional methods of lignin isolation procedures the reader is referred to more specialized texts [11,255, 263, 267]. Due to the variety of techniques available for isolating lignin, and the structural variations introduced during the isolation, each lignin preparation is usually identified by the wood species and the isolation technique, e.g., sprucedioxane lignin, or birch-cuoxam lignin. It is important to distinguish that all these preparations are distinct from protolignin, which is a term used for the material as it occurs in the plant tissue [268, 269].

5.2 Commercially Produced Lignins Commercial lignins from hardwood and softwood trees are available from the two major pulping processes, sulfite pulping and kraft pulping, which uses sodium sulfide and sodium hydroxide. In addition, pilot plant scale procedures exist which employ either organic solvents (organosolv pulping) or high pressure steam followed by alkaline extraction (steam explosion pulping). While the sulfite pulping process generates water-soluble lignosulfonates as their sodium, magnesium, or calcium salts, the kraft process produces lignins that are soluble only in aqueous alkali. In addition to differences in structure and botanic origin, commercial lignins also vary in molecular weight. Most such lignins are of weight average molecular weights (Mw) ranging between 3000 and 20,000 and polydispersity ratios (Mw/Mn) between 2 and 12 [270,271].

6 Methods of Lignin Analysis 6.1 Degradative Methods The methods described in this section require the sample to be exposed to certain reagents and specified conditions that degrade the lignin in a prescribed manner, in accordance with documented reaction sequences. The products are then quantitatively analysed by chromatographic techniques yielding structural information about the building blocks of the lignin sample [272]. Such techniques suffer from being laborious, involving many steps and complex chemical manipulations, often subjecting the derived quantitative information to large errors and diminished reproducibility. Despite these limitations these methods have offered significant advances to our knowledge of lignin structure and reactivity. Lignins can be selectively oxidized in alkaline nitrobenzene or permanganate solutions. The alkaline nitrobenzene oxidation of lignin was first introduced in

148


1939 by Freudenberg and co-workers [273,274] and was further optimized by Leopold [275 277]. Traditionally, nitrobenzene oxidation is used for the classification of wood or plant tissue and of lignin according to its botanical origin. During this procedure the lignin or the fibrous sample is oxidized in sodium hydroxide (2N) in the presence of nitrobenzene at 170-180 ~ for 3-4 hours. The products, mainly aldehydes, originating from the guaiacyl, syringyl and phydroxyphenyl structures in lignin are then subjected to quantitative determination. The oxidation of lignin in pulps by standard permanganate solutions has been the basis of the permanganate and kappa number tests, extensively used by the pulp and paper industry as a means of rapidly determining the lignin content of pulps. These techniques are based on the fact that potassium permanganate readily and selectively oxidizes lignin in the presence of carbohydrates. The permanganate number is conventionally reported to be the number of millilitres of 0.1 N KMnO4 consumed by 1 g of oven-dry pulp. However, since the size of the pulp sample and the amount of permanganate applied affected the results it was modified by Tasman and Berzins [278,279] who proposed the kappa number. As such, the sample size was adjusted to ensure that approximately half of the applied permanganate is consumed. This test has been of extreme significance to the pulp and paper industry in determining the degree of delignification of a fibrous feedstock during pulping and bleaching operations. Recent research, however, has brought to light that hexeneuronic acids, formed during pulping, consume permanganate during the kappa number test leading to significant errors specially when hardwoods are examined [280]. Analytical protocols involving alkaline permanganate oxidations of fibrous and lignin samples may provide similar information to that obtained by nitrobenzene oxidation. Moreover, these techniques offer structural details about the frequency of the various bonding patterns in lignin [281]. Despite the many steps involved and the fact that the obtained information is confined to units possessing free phenolic hydroxyl groups, alkaline permanganate oxidations have been extremely important in many aspects of lignin chemistry. The significance of the technique coupled with the complex nature of the various chemical manipulations have caused the development of a four step standardized procedure [53]. The procedure commences with an alkaline CuO predegradation step, followed by a methylation step, and ends with two oxidation steps involving permanganate and hydrogen peroxide [282]. The structure of lignins present in neutral sulfite [283-285] and in kraft pulps [286-288] has been elucidated using such techniques. Another procedure which has significantly contributed to our understanding of the lignin structure is that of hydrogenolysis [289-291]. The procedure consists of hydrolytic and thermolytic reactions which cause the partial breakdown and solubilization of lignin. The lignin fragments, diffuse into the liquid phase where they are rapidly stabilised against condensation reactions by in situ generated hydrogen atoms on the surface of a catalyst [292]. Despite the fact that the procedure is not in wide use today the principle may be applicable to converting lignin into valuable low molecular weight products [293].

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Pepper et al. were the first to propose the principle of acidolysis as a procedure for isolating lignin from plant material [294] and recently from kraft pulps [-280]. Later the same principle was used as an analytical tool for determining the occurrence of [3-0-4 and 13-5 structures in spruce lignins [-295]. The procedure calls for refluxing the lignocellulosic material in 0.2 N HC1 in a dioxane: water mixture (9 : 1, v/v). These conditions have been shown to cleave both a and [3 aryl ethers linkages in lignin. Such studies have provided evidence for the existence of additional structural elements in lignin such as 13-13, [3-1, and quinonoid [69, 296 299]. Acidolysis protocols in the presence of catalysts, other than hydrogen chloride, have been proposed by Yashuda [300] and Karlsson [301] and reductive cleavage protocols by Shevchenko et al. [-302]. The main critique of all methods involving acidolysis has been the fact that almost invariably the product yields are rather low as a result of condensation reactions taking place in acidic media [,168]. Consequently efforts have been made to address these limitations. Since the combination of boron trifluoride and thioethanol in anhydrous media is known to convert benzylic cation intermediates to thio-benzyl derivatives, Lapierre et al. have used this principle to develop the technique of thioacidolysis [-303]. This combination has been shown to quantitatively and selectively cleave the arylglycerol-[3-aryl ether linkages in lignin [-303-305]. Recent developments of the technique involving the desulfurization of the products using Raney nickel have been claimed to offer improvements over earlier protocols [-306].

6.2 Non Degradative Methods Non-degradative methods, involve mostly modern spectroscopic and other non evasive techniques, yielding structural information about the sample without the need of subjecting it to harsh chemical environments. Characteristic rc-~z* transitions of the aromatic nuclei in lignin make it a strong absorber of UV light. As such, lignin absorbs much more strongly than carbohydrates with an absorption pattern which has a maximum at approximately 280 nm. Based on detailed measurements of the UV absorption spectra of model compounds involving guaiacyl and syringyl moieties [-307], Fergus and Goring pioneered the use of UV microscopy for studying the distribution of lignin across the cell wall, in different morphological regions within woody tissue [308, 309] and during delignification in kraft and neutral sulphite pulping [,310]. Similar efforts by Boutelje and Eriksson E311] and Yang and Goring [-312] firmly established the use of UV microscopy for such endeavours. The UV absorption spectra of lignins may also be used to estimate their free phenolic hydroxyl content [-313]. The method is based on the fact that ionized phenolic groups in lignin absorb at 300 nm fully obeying the Beer-Lambert law. The use of this technique allowed the determination of the phenolic hydroxyl content in the secondary wall, which was found to be 50% greater to that of the middle lamella [,314]. The characteristic UV absorption spectra of lignin have been used for the quantitative and

150


qualitative determination of native lignins [315, 316], lignosulfonates [317] and the study of delignification of wood during pulping [318, 319]. Despite its numerous advantages the method can give erroneous results in the presence of contaminants that are strong UV absorbers. Its reproducibility has also been questioned as a consequence of an international round robin study involving many lignin standards [-320]. In recent years the application of nuclear magnetic resonance spectroscopy to the characterization, classification and detailed structural elucidation of lignins has seen widespread utility. The fact that the proton nucleus is of 100% natural abundance and of high sensitivity made the 1H NMR experiment rather popular in the early days of the application of this technique to lignin [321 323]. Almost invariably, 1H-NMR is used on acetylated lignins [322-326] since this affords better signal resolution. 1H-NMR signal assignment has been based on model compound [327-329], decoupling [330], and 2D-NMR experiments [331]. Nevertheless, there are some essential limitations to 1H-NMR spectroscopy of lignins. These include the rather limited range of chemical shifts (12 ppm), extensive signal overlapping and proton coupling effects. The technique is only suitable for the qualitative study of the proton distribution in lignins, while carbon containing groups and labile proton functionalities (OH, COOH, SH) remain beyond the capabilities of the technique. Recent developments have claimed that after appropriate derivatization it is possible to quantitatively determine the phenolic hydroxyl groups in lignin [332]. The early work of Ludemann and Nimz [49] has allowed the use of solution 13C-NMR spectroscopy to become an indispensable tool for the structural elucidation of the carbon skeleton of lignin. Compared to proton NMR, the 13C_NMR spectra of lignin offer considerably better resolution, with no coupling effects, over a much wider chemical shift range (200 ppm). Convenience, speed, and a wealth of even quantitative information [333,334,337] have contributed to the widespread use of ~3C-NMR for the structural analysis of lignins. Furthermore, a variety of newly developed pulse sequences such as DEPT [335], INADEQUATE [336], HMQC [81], as well as acquisitions on 13C-enriched lignin samples [337-339] have improved our understanding of lignin structure and reactivity. Despite all these, some drawbacks remain as far as the efficient use of ~3C NMR is concerned. These stem from the fact that the a3C nucleus is not sensitive enough (1/5800 compared to proton) and that it is only 1.1% naturally abundant. Consequently, relatively large sample sizes and lengthy acquisition protocols are essential. When quantitation is sought extreme care is required to allow for complete relaxation of the magnetization which translates to very lengthy acquisition protocols and the need of external methoxyl analyses so that the OCH3 signal to be used as internal standard [333, 340]. Advances in instrumentation have made the use of solid-state NMR spectroscopy a routine operation [341,342]. As such, various solid-state 13C-NMR experiments can now be performed routinely on lignin [343, 344] and solid wood or plant samples [345, 346, 95]. It is thus possible to obtain some information about the lignin within a sample without its prior isolation [87]. Dipolar

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dephasing when used as part of the acquisition protocol of solid-state spectra of wood and pulps has been documented to yield significant information in relation to the degree of chemical modification occurring in lignin [-347]. Efforts to overcome some of the limitations imposed by proton and 13C_NM R spectroscopies have prompted the examination of other NMR-active nuclei which when covalently linked to lignin by appropriate derivatization procedures may provide additional structural information for these heterogeneous biopolymers. Early attempts examined the potential of silylation followed by silicon N M R for the determination of hydroxyl groups in kraft lignin and related model compounds [348, 349]. While the method offers resolved signals for aromatic, phenolic and carboxylic acids, large sample concentrations and long acquisition protocols are essential due to the low natural abundance, low magnetic moments and high relaxation times of the 298i nuclei. 19F-NMR has also been proposed as a means of detecting different functional groups in lignins [350-353]. Finally, 31p_NM R spectroscopy has provided a new analytical tool capable of detecting a variety of functional groups in isolated soluble lignins and within lignin containing papers in the solid-state [-354]. More specifically, solution 3~p N M R has been used to examine soluble lignins [355] and carbohydrate [356] samples after phosphitylation with 1,3,2-dioxaphospholanyl chloride and 2-chloro-4,4,5,5-tetramethyl-l,3,2-dioxaphospholane [357, 358]. F r o m a single quantitative 3aP_NMR experiment it is possible to determine the three principal forms of phenolic hydroxyls present in lignins i.e. p-hydroxyphenyl, guaiacyl, and syringyl structures as well as primary hydroxyls, carboxylic acids, and the two diastereomeric forms of arylglycerol-beta-aryl ether units (13-O-4 structures) [-359]. Most significantly, however, the technique offers additional rapid (1-2 h acquisition time) information in relation to the condensed structures in lignin that can only be obtained by permanganate oxidation [360 364].

Acknowledgments. The

authors would like to acknowledge the following individuals for having critically examined the manuscript prior to dissemination: Dr. R.M. Berry, Professor K.E. Eriksson, Dr. I. Falkehag, Professor R. Francis, Dr. G. Leafy, Dr. L. Jurasek, Professor R.H. Marchessault, Mr. Y. Sun and Dr. S. Shevchenko.

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Fungal Delignification and Biomechanical Pulping of Wood M. Akhtar 1, Robert A. Blanchette2, and T. Kent Kirk 3 1 University of Wisconsin Biotechnology Center and Forest Products Laboratory, Madison, WI, USA 2 Department of Plant Pathology, University of Minnesota, St. Paul, MN, USA 3 Forest Products Laboratory, Madison, WI, USA

1 Fungal Degradation of Wood . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2 Wood Decay by White-Rot Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.1 Microstructural Changes and Delignification . . . . . . . . . . . . . . . . . . . . 1.2.2 Lignin-Degrading Enzyme Systems . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.3 Enzyme Localization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Biopulping . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Past Work . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Biomechanical Pulping . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.1 Screening of Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.2 Evaluation of Selected Fungi in Biopulping Runs . . . . . . . . . . . . . . . . . 2.3.3 Optimization of Biopulping Runs . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.4 Microscopy Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.5 Engineering and Economic Studies . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3.6 Prospects for Biopulping Commercialization . . . . . . . . . . . . . . . . . . . . 3 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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This review article summarizes the results on microstructural changes and delignification, lignin-degrading enzyme systems, and biopulping of wood with lignin-degrading fungi. Biopulping, defined as the treatment of wood chips with lignin-degrading fungi prior to pulping, saves substantial amount of electrical energy during mechanical pulping, results in stronger paper, and lowers the environmental impact of pulping. Optical properties are diminished; however, brightness can be restored readily with peroxide bleaching. The economics of the process look attractive if the process can be performed in a chip-pile based system. Past work on biopulping had been minimal, however a comprehensive evaluation of biopulping at the Forest Products Laboratory suggests that biopulping has a good chance of commercial success.

Advancesin BiochemicalEngineering/ Biotechnology,Vol. 57 Managing Editor: T. Scheper 9 Springer-VerlagBerlin Heidelberg 1997

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1 Fungal Degradation of Wood 1.1 Introduction Although wood can be attacked by an array of microorganisms, fungi are the predominant decomposers in terrestrial ecosystems. Extracellular enzymes produced by decay fungi and subsequent decomposition processes differ among the various groups of fungi, resulting in different types of wood degradation. Brown, soft, and white rot are categories used to separate different forms of decay. These categories are based on the macroscopic characteristics of the advanced stages of decay. Brown rot is caused by fungi taxonomically classified in the Basidiomycotina. These fungi cause rapid and extensive depolymerization of cellulose early in the decay process [ 1 4 ] . Wood polysaccharides are degraded, lignin modification occurs, and relatively small amounts of lignin are lost as decay progresses [-5-7]. In advanced stages of decay, the residue is a brown mass, which mostly consists of lignin. This decayed wood is sponge-like when wet, but often cracks and checks into cubical pieces as the wood dries. Brownrot fungi are common decomposers in conifer forests and also are responsible for most decay found in buildings and wood in service. Soft rot is a term first used to describe decay by fungi attacking wood surfaces in wet environments [8]. Fungi in the Ascomycotina and Deuteromycotina attack wood surfaces resulting in soft, gray to brown decay. Since soft rot was first reported, knowledge of these organisms has expanded--soft rot fungi have been found associated with wood in many different situations, such as in wood treated with preservatives or in wood that receives only intermittent moisture [6, 9-11]. Two forms of soft rot attack, Type I and Type II, have been identified [12]. Type I consists of cavities within the secondary wall. Mycelia in cell lumina produce fine penetrating hyphae that enter the secondary wall, align growth along the microfibrillar axis of the cell wall, and produce chains of cavities. The Type II form of attack consists of an erosion of the entire secondary wall originating from hyphae in cell lumina and progressing toward the middle lamellae. The entire secondary wall may be degraded, but the middle lamella is not attacked. Certain fungi, including Hypoxylon spp. and Xylaria spp., were previously considered to be white-rot fungi, but their mode of attack is more correctly classified a Type II form of soft rot [6, 13]. Fungi that cause white rot belong to the Basidiomycotina and have the capacity to degrade all cell wall components, including lignin. The extent of lignin degradation can vary considerably among species of white-rot fungi [14]. Some species, such as Trametes versieolor, are nonselective in how they degrade the wood, i.e., they simultaneously degrade lignin, cellulose, and hemicelluloses. Other species, such as Phellinus pini, Ceriporiopsis subvermispora, and Phlebia tremellosa, cause preferential degradation of lignin [15]. Some species deplete lignin, cellulose, and hemicellulose in varying ratios; many species attack both nonselectively and selectively in different areas of the same substrate [14].

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Although white-rot fungi have been categorized by whether they cause selective lignin degradation or nonselective decay, it is apparent that the degradation processes of these fungi are extremely variable. Even different strains of one species of white-rot fungus were recently shown to degrade cell wall components differentially [16].

1.2 Wood Decay by White-Rot Fungi 1.2.1 Microstructural Changes and Delignification White-rot fungi enter cell lumina and rapidly colonize ray parenchyma cells that contain free sugars and other nutrients. The radial arrangement of the ray parenchyma facilitates access into the wood and allows widespread distribution of the fungus in the substrate. Access to adjacent cells occurs via pit apertures, or direct penetration may take place directly through the cell wall [2, 4]. Once easily-assimilated substances are depleted, degradation of the cell wall is initiated. White-rot fungi that attack all cell wall components simultaneously cause a localized erosion of all cell wall layers. The attack progresses through the secondary wall layers and middle lamella (Fig. 1A and B). In advanced stages of decay, cell walls are eroded extensively, and holes within adjacent cell walls are frequently observed (Fig. 1B). A different form of cell wall attack occurs in white-rot fungi that selectively degrade lignin. Hyphae in cell lumina degrade lignin progressively from the lumen edge of the secondary wall toward the middle lamella (Fig. 1C and D). Investigations using brominated wood and X-ray microanalysis of the bromine-lignin complex showed that white-rot fungi remove lignin from the secondary wall before the middle lamellae between cells are degraded [17,18]. As the delignification process continues, the middle lamella is degraded and cells separate from adjacent cells (Fig. 1C and D). The delignified, cellulose-rich secondary wall remains relatively unaltered (Fig. 1C and D). The degradation of lignin is extensive throughout the cell walls, originating from only one or two hyphal filaments within each cell lumen (Fig. 2A and B). Wood degradation may be influenced significantly by the lignocellulosic substrate. An important factor that governs the extent and rate of decay is the amount and type of lignin present in the wood. Wood from gymnosperms has greater concentrations of lignin than wood from angiosperms and consists primarily of guaiacylpropyl units. Lignin from angiosperm wood is composed of varying amounts of syringylpropyl and guaiacylpropyl units. In studies evaluating decay by white-rot fungi in different types of wood, angiosperm wood was found to degrade more rapidly and to a greater extent than gymnosperm wood [1, 15, 19]. Synthetic syringyl lignin has also been shown to be depolymerized more rapidly than synthetic guaiacyl lignin in a laboratory investigation using Phaneroehaete chrysosporium [20].

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Fig. 1. Wood decayed by white-rot fungi with simultaneous removal of all cell wall components (A and B) and preferential degradation of lignin (C and D). A nonselective attack erodes all cell wall layers, including the compound middle lamellae. Erosion zones coalesce to form holes through cell walls (arrows).C and D: Lignin in secondary walls and middle lamella regions has been removed by selective delignification. Cells separate from adjacent cells as a result of the lack of middle lamellae. The cellulose-rich secondary wall remains. Transverse sections, scanning electron micrographs. Bar = 10 mm D e l i g n i f i c a t i o n in a n g i o s p e r m s b y w h i t e - r o t fungi also a p p e a r s to be influenced b y the syringyl lignin c o n t e n t of the w o o d . Studies of w o o d d e c a y in Nothofagus dombeyi from the t e m p e r a t e rain forests of s o u t h e r n Chile s h o w e d extensive delignification b y Ganoderma australe [14, 21]. This is one of the few l o c a t i o n s in the w o r l d where h u g e logs m a y be c o m p l e t e l y delignified. In


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Fig. 2A, B. Selective delignificationin cell walls appears as electron-lucid zones after KMnO4 fixation and transmission electronmicroscopy.A Lignin is progressivelyremovedfrom the secondary wall and then from the middle lamella (arrows show extent of delignification).B: Completely delignifiedcells-detachedcells free of middle lamellae. Bar = 5 mm temperate forests of North America, Europe, and Japan, delignification usually occurs in small pockets within the wood or sometimes in large localized zones, but the delignification of entire logs has not been reported. Nothofagus dombeyi has an extremely high syringyl lignin content [21J--three to six times that of Acer, Betula, and other hardwood species--and, worldwide, it appears to be among woods with the highest content of syringyl lignin. Other angiosperms growing in southern Chile, such as Laurelia phillipiana, that have low syringylto-guaiacyl lignin ratios are not delignified when decayed by G. australe. Instead, a nonselective white-rot type of degradation occurs [21]. These results strongly suggest that the concentration of syringyl lignin within angiosperm wood greatly influences the delignification process. Although white-rot fungi preferentially degrade syringyl lignin in angiosperms, some white-rot species extensively degrade guaiacyl lignin in gymnosperms. Fungi such as P. pini, Heterobasidion annosum, and P. weirii are usually found only in coniferous wood and apparently have evolved highly efficient mechanisms of lignin degradation. Other species of white-rot fungi also do not appear to be deterred by the guaiacyl lignin of pines and other conifers. Species such as C. subvermispora can readily delignify the sapwood of loblolly pine, causing considerable amounts of lignin removal [16]. These species are of special interest for use in biological pulping processes because they can degrade lignin in conifers as well as hardwoods.

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The process of preferential delignification of woody cell walls by white-rot fungi has been elucidated using a variety of techniques. Histological stains and electron-dense compounds that react with lignin can be used to visualize lignin removal from the cell wall [22, 23]. KMnO4 can be used to fix wood for transmission electron microscopy and to observe the distribution of lignin in cell walls. Moderate electron density is evident in the secondary wall where lignin is found with cellulose and hemicellulose, but intense electron density can be observed in the highly lignified middle lamellae [17]. As hyphae of white-rot fungi, located in cell lumina, begin to delignify the secondary wall, an electrontransparent zone develops (Figs. 2A). This clear zone progressively moves into the secondary wall. Once lignin is removed from this region, the middle lamella between cells and, in very advanced stages of decay, the cell corner regions are degraded and become less electron dense (Fig. 2B) [14]. A light-based microscopic method that employs bright colored stains (astrablue and safranin) has recently been used to differentiate zones of delignification from nondecayed cells that retain lignin [24]. Colloidal gold cytochemistry using gold-labeled endo-l,4-b-glucanase II, 1-4-b-o-glucan cellobiohydrolase I, and endo-1,-4-b-xylanase revealed that residual delignified wood contains crystalline and amorphous cellulose but little xylan [25, 26]. Other studies also have shown that hemicelluloses are usually depleted from wood as lignin is being degraded [14, 27].

1.2.2 Lignin-Degrading Enzyme Systems Lignin-degrading enzymes were discovered, and subsequently characterized, in P. chrysosporium (28 32). Research on the biochemistry of lignin degradation began in earnest in the 1970s, when 14C-lignins were prepared and used to determine which groups of microbes are able to mineralize lignin (decompose it to 14CO2). The higher basidiomycetous fungi were found to be the most proficient, and P. chrysosporium was chosen for detailed study for a number of reasons related to ease of experimentation. Subsequent research with this fungus described the culture parameters important for lignin mineralization and showed that one or more steps are "secondary metabolic" events triggered by limitation for certain nutrients. At the same time, the chemical changes that occur in the lignin polymer during decay by ligninolytic fungi were described (reviewed by Kirk and Farrell [29]). In the early 1980s, specific degradative reactions accomplished by ligninolytic cultures of P. chrysosporium were described using synthesized "dimeric" lignin model compounds; it was also found that certain polymeric dyes are decolorized. Cell-free enzyme preparations that catalyzed dimer cleavage and dye decolorization were reported in 1983, and subsequently the responsible peroxidase (lignin peroxidase) was described. Evidence to date indicates that three oxidizing enzymes, lignin peroxidase (LIP), manganese peroxidase (MnP), and laccase are responsible for the initial fragmentation of the lignin polymer and production of low molecular mass


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breakdown products in white-rot fungi. Not all white-rot fungi apparently produce all three enzymes, although some, including T. versicolor, do. P. chrysosporium produces only LiP and MnP (however, laccase has recently been reported in P. chrysosporiurn [33]), whereas C. subverrnispora produces only MnP and laccase, and Phlebia ochraceofulva produces only LiP and laccase [32]. The extracellular H202 required as electron acceptor for the peroxidases is supplied by glyoxal oxidase, which, with its substrates, is extracellular, and perhaps also by intracellular sugar and alcohol oxidases. Molecular oxygen is the electron donor for laccase. Low molecular mass products of lignin degradation are taken up by the hyphae and further oxidized. The enzymes responsible for this intracellular oxidation are only beginning to be described [34]. Lignin peroxidase oxidizes aromatic nuclei (phenolic and nonphenolic) by removal of one electron, generating both phenoxy radicals and cation-radicals. The latter react spontaneously with nucleophiles (primarily water) and molecular oxygen. The result is an "enzymatic combustion" in which C-C and C-O linkages are cleaved, depolymerizing the polymer and opening aromatic rings. A plethora of aromatic and aliphatic products are thereby formed. In vitro depolymerization of lignin by pure LiP in the presence of H202 and veratryl alcohol was recently demonstrated [35]. Veratryl alcohol (3,4-dimethoxybenzyl alcohol), found in the extracellular milieu of ligninolytic cultures of examined LiP-producing fungi, is a secondary metabolic product synthesized de novo. It is a substrate for LiP and stimulates its action, probably not as an electron mediator as originally thought (see Kirk and Farrell [-29]), but by donating electrons to LiP so that its catalytic cycle is completed [36]. As mentioned, LiP is apparently not produced by some white-rot fungi, including C. subvermispora [37], suggesting that it is not required in all fungi, i.e., that the white-rot fungi have more than one enzyme system for degrading lignin. Interestingly, C. subvermispora and certain other LiP-negative fungi do have "LiP-like" genes [37]. Manganese peroxidase has been found in nearly all studied white-rot fungi. It catalyzes the oxidation of (complexed) Mn 2 + to Mn 3 +, which in turn oxidizes lignin. Mn / + is a fairly abundant element in wood. The "physiological" complexes that have been studied, such as the lactate chelate, oxidize only phenolic units, which constitute only about 10% of the total in lignin in wood [38]. The phenolic units are oxidized to phenoxy radicals, which can undergo certain degradative reactions [39]. The MnP/HzO2/Mn 2+ has been shown to depolymerize lignin in vitro [40]. The exact function of MnP and the chemistry of its actual oxidation of lignin are not yet clear. In a recent study, Bao et al. [41] discovered a lipid peroxidation system involving MnP. In the presence of Mn(II), MnP promotes the peroxidation of unsaturated lipids, generating transient lipoyl radical intermediates that are known to act as potent oxidants of other molecules. This system, unlike MnP alone, oxidizes and cleaves nonphenolic model compounds via benzylic hydrogen abstraction. It also depolymerizes both nonphenolic and phenolic synthetic lignins, which strongly supports a ligninolytic role for this system in vivo. The

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ligninolytic system of P. chrysosporium is depicted schematically in Fig. 3. We have included the lipid peroxidation system, even though its role is not yet established. Laccase is a blue copper oxidase that is produced (secreted) by most but not all white-rot fungi. Like LiP, laccase is apparently not required for lignin degradation in all fungi. Laccase oxidizes phenolic units in lignin to phenoxy radicals, which is the same process as that brought about by the chelated Mn(III) produced by MnP (Fig. 3). However, in the presence of appropriate "primary" substrates (such as ABTS), the effect of laccase apparently can be enhanced; laccase/primary substrate systems have recently been reported to degrade lignin in kraft pulp [42] and to oxidize nonphenolic compounds that otherwise are unattacked by laccase. However, it is not known whether such "primary" substrates occur and function in vivo, and the actual role of Iaccase, like MnP, remains unclear. (Note that if "primary" substrates augment laccase activity in vivo, they can be expected to augment the MnP/Mn 2+ system as well). The characteristics of LiP, MnP, glyoxal oxidase, and laccase have been described in review articles [28-31, 39, 43]. General properties of LiPs, common to all studied LiP-producing fungi, include the following. Lignin peroxidases consist of acidic isoenzymes encoded by multiple structural genes whose expression

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Fungal Delignificationand BiomechanicalPulpingof Wood

167

is nutrient-regulated. The extracellular proteins have protoporphyrin IX-type heme prosthetic groups, are glycosylated, and have molecular weights around 40 kDa. Lignin peroxidases exhibit the common peroxidase catalytic cycle, being first oxidized by H202 by removal of two electrons to give compound I, which oxidizes its substrates by removing one electron to give compound II, which oxidizes its substrate, returning to resting enzyme. The LiP from P. chrysosporium has been crystallized and its three-dimensional structure resolved by X-ray crystallography [44, 45]. Manganese peroxidases are like LiP in that they consist of multiple acidic isoenzymes encoded by multiple structural genes whose expression is nutrientregulated; in the case of MnP, regulation by Mn 2+ also occurs. Manganese peroxidases are slightly larger than LiPs, but exhibit the same basic peroxidase catalytic cycle. Interestingly, MnP, in the presence of reducing agents such as glutathione, transfers electrons to molecular oxygen, generating H z O 2. Like LiP, MnP has been crystallized and its three-dimensional structure determined [46]. Note that LiP can display MnP activity; in the presence of H202, O2, and the metabolites veratryl alcohol and oxalate, LiP oxidizes Mn 2 + to Mn 3 + [38]. In at least some white-rot fungi, including T. versicolor, laccases also consist of acidic isoenzymes encoded by multiple structural genes. The molecular weights of the laccases of white-rot fungi are in the range 50-65 kDa. Laccases "store" four electrons from four sequential one-electron oxidations before reducing molecular oxygen to water. The molecular genetics of the lignin-biodegrading system have received attention in the last few years. Such studies have now been done with several fungi, although P. chrysosporium has been the most thoroughly studied. With that organism (and with the other studied fungi as well), complex structural gene families have been found. In P. chrysosporium, 10 lip, 4 mnp, and 1 91ox have been cloned and sequenced. All the genes contain introns and multiple glycosylation sites. Further detail is found in recent reviews [31, 43].

1.2.3 Enzyme Localization Immunological cytochemistry in conjunction with electron microscopy can be used to localize enzymes within substrates and observe their spatial relationship within decaying wood. The association of delignified zones with specific enzymes would indicate that the enzymes were able to penetrate the cell wall and accumulate at sites of lignin degradation. Investigations using polyclonal and monoclonal antibodies to lignin peroxidase and manganese peroxidase have shown that these enzymes are present in cell walls of wood decayed for 6 to 12 weeks [25, 4749]. High concentrations of enzymes were found in decayed cell walls that had a loose, modified ultrastructural matrix. Lignin peroxidase and manganese peroxidase were evident at the edge of electron-dense regions in the secondary wall or at sites of middle lamella degradation. Significant alteration of the cell wall was always associated with localization of these enzymes. No

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labeling was evident in nondecayed wood or in wood decayed by a brown-rot fungus [25]. To more accurately identify areas of the cell wall where enzymes could penetrate successfully, several ultrastructural studies were completed after infiltrating decayed wood with extracellular extracts from white-rot fungi or with purified lignin peroxidase and manganese peroxidase [48, 50]. After impregnation, gold-labeled antibodies were used to determine the extent of penetrability in the cells. In nondegraded cells, no enzyme preparations were found to enter the secondary wall. Enzymes were found only on the surface of the cell wall at the lumen. In samples decayed by white-rot fungi, lignin peroxidase and manganese peroxidase were found within altered cell walls. The enzymes diffused into the peripheral areas of the secondary wall and into areas where the middle lamella was becoming less electron-dense. The labeling appeared to be located at sites where lignin was being degraded. In areas with advanced decay, enzymes were located within the cell wall at the edges of undegraded cell corner regions [48].

2 Biopulping 2.1 Introduction The pulp and paper industry is a large and growing portion of the world's economy. In 1991, paper sales were valued at $122 billion [51], and 267 million metric tons of paper and paperboard were consumed worldwide. Worldwide consumption is expected to increase to 300 million metric tons in 1996. A number of pulping processes have been developed to meet industrial and consumer needs. Pulping processes are generally divided into two broad classes, chemical and mechanical, which produce substantially different fiber characteristics. The choice of process depends on the end application of the pulp and the raw material. In many papermaking operations, a combination of chemical and mechanical pulps is used to obtain the desired paper characteristics. Chemical pulping involves the use of chemicals to degrade and dissolve the lignin from the wood cell walls, releasing cellulose fibers. Chemical pulping processes are low yield (about 40-50%) and require significant waste treatment and chemical recycling operations; however, the pulps produced have high strength. Mechanical pulping involves the use of mechanical force to separate the wood fibers. Mechanical processes are high yield (up to 95%) and give paper with high bulk, good opacity, and excellent printability. However, these processes are energy-intensive and produce paper with lower strength and high color reversion (tendency to turn yellow with time). Bleaching of chemical pulps using a combination of chlorination and alkaline extraction has been used in the pulp and paper industry for many years.

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Unfortunately, the effluents from chlorination and alkaline extraction stages cannot be recycled back to the chemical recovery furnace because of their high level of corrosive chloride. In addition, the effluents contain large amounts of chlorinated organic compounds, which are known to have toxic, mutagenic, and carcinogenic effects. In most parts of the world, increased public concern about the environment is having a large impact on the pulp and paper industry. Reductions in allowable air and water discharges from pulp and paper mills are requiring restraint in the use of chlorine and chlorine dioxide for bleaching. Consequently, the industry is rapidly moving towards alternative technologies to alleviate this and other problems related to the environment. Biopulping, defined as the treatment of wood chips with lignin-degrading fungi prior to pulping, is an experimental process that has been researched extensively during the past 8 years. It has been studied mainly as a pretreatment for mechanical pulping. Biopulping reduces electrical energy consumption (which is the major cost in mechanical pulping), improves paper quality, and reduces the environmental impact of pulping [52, 53]. The following sections present a summary of this research and describe key findings.

2.2 Past Work The use of white-rot fungi for the biological delignification of wood was perhaps first studied by Lawson and Still [54] at the West Virginia Pulp and Paper Company research laboratory (non Westvaco Corporation). These researchers published a survey of the literature (covering 72 lignin-degrading fungi), which pointed to the dearth of knowledge about the fungal degradation of lignin. Research was then done at the US Forest Products Laboratory in Madison and the Swedish Products Laboratory (STFI) in Stockholm. The first published report on biopulping per se demonstrated that fungal treatment could result in significant energy savings for mechanical pulping [55]. That research resulted in a US patent [56], which described a "method for producing cellulose pulp." Considerable efforts at STFI were directed toward developing cellulase-less mutants of selected white-rot fungi to improve the selectivity of lignin degradation and thus the specificity of biopulping [57]. However, the mutant strains degraded less lignin than did wild-type strains when grown on wood [58] and did not result in energy savings during subsequent mechanical pulping [59]. Attempts by this group to scale up the biopulping process were not notably successful [60]. However, subsequent work with Cuban scientists on a pilot scale with bagasse using mutant strains gave more promising results 1-61]. Eriksson et al. [62] showed that chip colonization is not the rate-limiting step in biopulping. At the Forest Products Laboratory, Bar-Lev et al. [63] showed that the treatment of primary thermomechanical pulp with a white-rot fungus prior to secondary refining reduced energy requirements and increased paper strength properties. Similar results were obtained in Japan by Akamatsu et al. [64] during thermomechanical pulping of fungus-treated popular chips. Other

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details on biopulping research were described in two review articles and the literature cited therein [65, 66]. A comprehensive evaluation of biomechanical pulping was launched in 1987 at the Forest Products Laboratory after the establishment of a Biopulping Consortium, which involved the Forest Products Laboratory, the Universities of Wisconsin and Minnesota, and pulp and paper and related companies. The overall goal was to establish the technical feasibility of using a fungal pretreatment with mechanical pulping to save energy and/or improve paper strength. In addition, it was assumed that the fungal pretreatment would have less environmental impact than would chemical pretreatments, which turned out to be the case. The consortium research was conducted by seven closely coordinated teams: fungal, pulp and paper, enzyme, molecular genetics, economics, engineering scale-up, and information. However, in this review article, we will focus only on the work conducted by the fungal pulp and paper, and engineering scale-up research teams.

2.3 Biomechanical Pulping 2.3.1 Screening of Fungi There are hundreds of white-rot fungi with varying capacities to degrade lignin, cellulose, and hemicellulose. We assumed at the outset that the fungi that degraded lignin selectively would be the best candidates for biopulping. To ascertain the most appropriate species, a screening program was initiated that selected fast-growing species that could selectively remove lignin from wood.

2.3.1.1 In Vitro Wood Decay Test Several methods have been developed to select fungal species with selective lignin-degrading ability [15, 27, 67-70]. However, one of the most appropriate methods appeared to be an assessment of decay (chemical analyses of lignin and wood sugar content) using wood blocks in accelerated decay chambers [15, 71]. Based on this in vitro screening procedure, we selected several species of fungi; among the best were P. chrysosporium, C. subvermispora, Phlebia brevispora, Phlebia tremellosa, Dichomitus squalens, and Phellinus pini [15, 71]. Different strains of these selected species varied in their selectivity towards lignin. Two fungi were examined in detail: P. chrysosporium and C. subvermispora. As we tested various strains on different species of wood, it became clear that some strains are effective with hardwood only, whereas others are effective on both hardwood and softwood (Tables 1-4). These results clearly showed large differences among the strains in capacity to degrade lignin and in selectivity [16]. The species or strains selected by this method were then evaluated for their biopulping efficacy. No apparent relationship was found between the lignin removal from the wood chips and energy savings or strength improvements during the actual biopulping runs [72, 73]. This suggests that lignin

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Fungal Delignification and Biomechanical Pulping of Wood Table 1. Loss of weight, lignin, and wood sugars in aspen wood blocks decayed by different strains of P. chrysosporium (12-week incubation) Loss (%) Strain

Weight

Lignin

Glucose (glucan)

Xylose (xylan)

Mannose (mannan)

BKM-F-1767 5161 ME-8 FP-104297-sp FP-102169 HHB-6251-sp 5157-A- 1 Gold-9-420-1

61.0 53.6 55.3 50.2 56.2 23.0 61.2

80.7 54.4 47,5 56.7 58.9 30.6 73.4

49.9 53.0 51,3 51.4 55.0 5.2 56.1

77.7 61.0 63,0 52.0 59.4 27.4 71.7

61.0 77.6 81.5 55.3 59.2 60.2 67.9

Table 2. Loss of weight, lignin, and wood sugars in loblolly pine wood blocks decayed by different strains of P. chrysosporium (12-week incubation) Loss (%) Strain

Weight

Lignin

Glucose (glucan)

Xylose (xylan)

Mannose (mannan)

BKM-F-1767 5161ME-8 FP-104297-sp FP-102169 HHB-6251-sp 5157-A- 1 Gold-9-420-1

24.5 8.4 22.7 19.7 11.9 13.6 18.7

20.9 3.6 12.9 16.9 8.3 12.8 18.3

26.1 11.2 30,5 23.0 8.7 10.3 17.5

19.1 9.7 23.8 17.4 6.9 14.8 25.7

31.4 14.5 39.3 24.3 5.3 11.5 20.7

Table 3. Loss of weight, lignin, and wood sugars in aspen wood blocks decayed by different strains of C. subvermispora (12-week incubation) Loss (%) Strain

Weight

Lignin

Glucose (glucan)

Xylose (xylan)

Mannose (mannan)

ME-485 L- 14807-sp L- 15225-sp FP-104027-T L-39292-sp FP- 105752-sp CZ-3 L-6133-sp FP-90331-sp

28.4 24.4 25.4 26.4 25.6 22.7 23.8 24.4 26.5

61.5 57.2 58.8 65.9 63.7 55.7 71.2 70.7 50.1

2.5 6.8 2.9 2.2 2.1 0.6 6.3 3.4 7.3

45.4 36.9 40.4 44.6 47.9 31.0 43.8 38.4 31.5

72.3 39.1 66.3 66.8 66.4 30.2 28.7 29.3 31.3

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Table 4. Loss of weight, lignin, and wood sugars in loblolly pine wood blocks decayed by different strains of C. subvermispora (12-week incubation) Loss (%) Strain

Weight

Lignin

Glucose (glucan)

Xylose (xylan)

Mannose (mannan)

ME-485 L-14807-sp L- 15225-sp FP-104027-T L-39292-sp FP- 105752-sp CZ-3 L-6133-sp FP-90331-sp

22.8 22.5 23.7 28.3 29.0 19.6 21.3 30.1 22.7

31.0 37.0 38.2 40.6 42.2 33.9 31.8 34.1 38.2

20.3 14.7 12.4 18.8 22.4 7.1 14.0 26.2 14.1

0 33.2 27.2 33.8 31.1 27.0 31.0 34.0 30.0

24.2 29.9 28.1 26.9 26.2 10.1 20.3 18.9 15.9

modification rather than removal is involved. Screening by way of the in vitro wood decay test is also time-consuming and labor-intensive. We therefore sought alternative screening methods.

2.3.1.2 P F I Milling P F I milling and freeness measurements have previously been used to assess energy consumption of fungus-treated pulps [74]. We further evaluated this approach and tried to correlate changes in freeness after P F I mill refining of coarse aspen pulp treated with selected white-rot fungi with those of paper strength properties or energy savings obtained during biomechanical pulping of wood chips with the same fungi. We found that P F I milling and freeness measurements of pulp can, in themselves, give a good estimate of paper strength properties [75]. However, follow-up studies suggested that this method can only be used to evaluate the effect of fungal treatments on energy savings compared to the control; they cannot be used to discriminate the effect of different fungal treatments on energy savings [76]. 2.3.1.3 Simons Staining Simons stain [77, 78] had been used previously in various investigations to evaluate the degree of fibrillation in beaten pulp fibers. The stain consists of a 1% aqueous solution of Pontamine Sky Blue 6BX and a 1% aqueous solution of Potamine Fast Orange 6RN mixed in a 1 : 1 ratio. Fibers on microscope slides are first flooded with stain and then heated at 60 ~ to evaporate the water. The fibers are rinsed with distilled water to remove excess stain and then covered with glass. The fibers are then immediately examined under a microscope and photographed. Fibrillated fiber walls stain orange, and nonfibrillated or undamaged fiber walls stain blue. Our earlier electron microscope studies in connection with biopulping had shown that pulp fibers obtained from the fungus-treated wood chips had more uniform fiber length and greater fibrillation, and appeared woolly compared


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with control pulps [79], suggesting that extent of fibrillation might be a good indicator of biopulping efficacy. Therefore, aspen wood chips were treated with the white-rot fungus C. subvermispora for 4 weeks and then refined through a single-disk mechanical refiner. Pulp obtained from the fungus-treated chips had extensively fibrillated fibers that stained a deep orange with Simons stain (Fig. 4). In contrast, pulp obtained from the untreated control chips exhibited little fibrillation and stained a deep blue (Fig. 4). This showed that fibers obtained from the fungus-treated chips could be differentiated from those obtained from control chips based on the yellowing of fiber ends [80]. We

Fig. 4. Simons staining of control (top) and biopulp (bottom). Pulps from untreated control wood chips and Ceriporiopsis subvermispora-treated chips (4-weekincubation) were passed through the refiner only once (Canadian Standard Freeness of pulps at about 700 ml) and then stained [80]

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Table 5. Correlation of yellowing of fiber ends with energy savings using selected Ceriporiopsis spp. on aspen wood chips (2-week incubation) Yellowing of fiber ends (%) Species/strain Control

C. rivulosa L-10602-sp. C. rivulosa PiiRTO-26K225 C. pannocincta FP-101181-sp. C. subversmispora L-3292 C. subvermispora CZ-3 C. subvermispora L-9186-sp.

None + + (0) + a +

Slight

Intermediate

_1_a (3%)b + " (4%) +

+ (7%) + (12%) +

Advanced

+ (16%)

Two plus signs for one treatment indicate that the staining pattern was in-between the two categories b Values in parentheses represent refining energy savings compared to that of the untreated control

further evaluated this approach to determine whether it can be used to monitor differences among fungal pretreatments. We found that the intensity of yellow staining of biopulp fibers obtained under different experimental conditions correlates very well with energy savings from completed biopulping runs (Table 5). These results were summarized in a recent publication [81].

2.3.2 Evaluation of Selected Fungi in Biopulping Runs White-rot fungi screened in the wood decay test or with Simons stain were evaluated for their performance in refiner mechanical pulping. The process involved the treatment of wood chips with the fungi in bioreactors on a bench scale at appropriate temperature and humidity, mechanical pulping of control and fungus-treated chips through a single-disk atmospheric refiner, preparation of paper, and testing of the paper for physical properties. 2.3.2.1 Method Three types of bioreactors were designed and used: a rotating drum bioreactor, a stationary tray bioreactor, and an aerated static bed bioreactor. Details of the configuration of each bioreactor have been published (rotating drum bioreactor [82], stationary tray bioreactor [83], and aerated static bed bioreactor [84]). In recent studies, we have used the simple and inexpensive aerated static bed bioreactor (Fig. 5). Chips (1500 g, dry weight basis) are introduced into each 21-1 bioreactor with water (containing nutrients [85] and additive, if any), and the loaded bioreactors are then usually sterilized by autoclaving. Chip moisture content is adjusted to 55%-60% on a wet weight basis. The chips are then inoculated with the fungus and incubated at an appropriate temperature (39 ~ for P. chrysosporium and 27 ~ for other fungi) for 2-4 weeks (in most cases 2 weeks) with humidified air (0.022711-1min-t). Details are described in previous publications [83, 84, 86].

Fungal Delignification and Biomechanical Pulping of Wood

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Fig. 5. Diagram of a static bed bioreactor [84]. The bioreactor was fabricated from a polypropylene vessel(L). The top of the vesselis sealed with a lid (M), which is vented to the atmosphere through an exit tube (N). Suspended above the bottom of the reactor (L) is a perforated polypropylene floor (.1), supported by a stand (K). Air for the bioreactor comes from a regulated supply and passes through tubing (A) to a fritted glass gas dispenser (B) in a humidification flask (C) containing sterile water. Humidified air passes through tubing (D) to a rotameter (E) and through tubing (F) to a manifold (G). Humidified air from manifold(G) is passed through a filter (H) before passing through tubing (1) to the reactor (L) base

After harvest, the untreated control chips and the fungus-treated chips are fiberized by multiple passes through a S p r o u t - W a l d r o n Model D 2202 single rotating 300-mm-diameter disk atmospheric refiner. Energy consumed during fiberization and subsequent refining is measured using an Ohio Semitronic Model W H 30-11195 integrating watt-meter attached to the power supply side of the 44.8-kW electric motor. Pulps are then refined to Canadian Standard Freeness (CSF) values just above and just below 100 ml. CSF is an arbitrary measure of the rate of water drainage from a pulp slurry. Handsheets (60 g m - z) are made with these two pulp samples and tested for physical properties using Standard T A P P I methods. Energy values and physical properties are regressed to 100 ml CSF to facilitate comparison. Details of energy measurements, handsheet preparation, and testing methods have been described [72, 82, 87, 88].

2.3.2.2 Energy Savings and Physical Properties In early experiments, several white-rot fungi screened initially by the wood decay test were further screened for their biomechanical pulping performance using aspen (hardwood) wood chips [72, 73, 82, 88]. Based on energy savings and paper strength improvements, six fungi were initially selected: P. chrysosporium, P. tremellosa, P. subserialis, P. brevispora, D. squalens, and Poria medulla-panis. Of these, more emphasis was given to P. chrysosporium because it is by far the most studied white-rot fungus, grows rapidly, and competes well with indigenous microorganisms of wood chips. Periodically, additional fungi

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Table 6. Energy savings and changes in physical properties from biomechanical pulping of loblolly pine chips with different white-rot fungi (4-week incubation) Strength properties

Optical properties

Fungus

Energy savings (%)

Burst index (kN g - i )

Tear index (mN m2g - i )

Tensile index (N mg-1)

Brightness (%)

Opacity (%)

Scattering coefficient (m 2 kg- l)

Pc Hs Pb Ps Pt Cs

14 26 28 32 36 42

14 12 - 4 -29 - 18 32

1 32 21 9 12 67

19 - 12 - 9 -36 - 17 -1

-27 - 21 - 24 -28 -24 -22

-2.3 - 3.6 - 2.9 -3.7 -4.5 -4.4

-29.8 - 27.6 - 28.1 -34.6 - 32.6 -30.9

Pc = Phanerochaete chrysosporium; Hs = Hyphodontia setulosa; Pb = Phlebia brevispora; Ps = Phlebia subserialis; Pt = Phlebia tremellosa; Cs = Ceriporiopsis subvermispora Table 7. Energy savings and changes in physical properties from biomechanical pulping of aspen wood chips with three strains of C. subvermispora (4-week incubation) Strength properties

Optical properties

Strain

Energy Burst savings index (%) (kN g - i )

Tear index (ran m2g -1)

Tensile index Brightness (N mg 1) (%)

Opacity (%)

Scattering coefficient (m 2 k g - i )

FP-90031-sp. L-6133-sp. CZ-3

40 44 48

131 137 162

17 20 27

- 2.0 - 1.3 -- 1.3

-- 33.8 -- 34.0 - 37.1

23 27 40

- 18 - 21 -- 21

Table 8. Energy savings and changes in physical properties from biomechanical pulping of loblolly

pine chips with three strains of C. subvermispora (4-week incubation) Strength properties

Optical properties

Strain

Energy savings (%)

Burst index (kN g - i )

Tear Tensile index index (raN mZg l) (N mg-1)

Brightness (%)

FP-90031-sp. L-14807-sp. FP-104027-T

37 30 21

41 44 45

54 59 47

4 3 11

- 21 -- 19 - 21

Opacity (%) - 0.94 -- 0.63 0.0

Scattering coefficient (m 2 k g - l ) -- 27.2 - 24.9 - 23.1

identified for lignin selectivity on the basis of the wood decay test were also e v a l u a t e d f o r t h e i r b i o m e c h a n i c a l p u l p i n g efficacy. As the research progressed, emphasis was given to screening the fungi on loblolly pine (softwood) chips because this wood, together with other southern p i n e s , is a m a j o r w o o d s o u r c e f o r m e c h a n i c a l p u l p m i l l s i n t h e U n i t e d S t a t e s . Some of the fungi selected as being best on aspen wood chips were evaluated on l o b l o l l y p i n e c h i p s . S o m e o f t h e s e r e s u l t s a r e p r e s e n t e d i n T a b l e 6. All t h e f u n g i s a v e d e n e r g y a n d s o m e i m p r o v e d p a p e r s t r e n g t h , b u t all a d v e r s e l y a f f e c t e d


177

paper optical properties. Based on energy savings and strength improvements, C. subvermispora was identified as the best fungus [86]. Different strains of this fungus were found to be effective on both aspen (Table 7) and loblolly pine (Table 8) [84]. A US patent was issued on the use of C. subvermispora for biomechanical pulping [89].

2.3.3 Optimization of Biopulping Runs For biopulping, like any industrial microbial process, there is great opportunity to increase the effectiveness and efficiency and decrease the cost of the process through optimizing the variables. Our initial work was governed by "best guesses" for optimal biopulping conditions based on the literature, knowledge of fungal growth, and past experience. Initial research focused on the fungus P. chrysosporium and aspen chips; later, C. subvermispora became the focus of our study. Some of the initial results are in the following text.

2.3.3.1 Wood Batch, Chip Storage, and Chip Movement During Incubation Some parameters, including wood batch and chip storage conditions (frozen, fresh, or dried at room temperature) did not seem to affect biopulping performance. In early experiments, fungus sensitivity to chip movement was observed when stationary and rotating drum bioreactors were used. Chip movement affected the extent of chip degradation, energy consumption during refining, and paper strength properties [72, 88]. Later studies, however, showed that shaking the aerated static bed bioreactors once a week during the 4-week incubation period had no appreciable effect on energy savings or paper strength properties. 2.3.3.2 Inoculum In any industrial fermentation, the inoculum is of key importance. A number of variables affect the fermentation, including level, physical form, age, and viability. We examined some of these variables in a series of experiments. The effect of different inoculum levels (2.5%, 5%, 10% and 20%, dry weight basis), using precolonized chips as inoculum, was studied with P. chrysosporium on aspen wood chips. The lowest inoculum level (2.5%) gave slightly lower energy savings than the other three levels. We postulated that the addition of nutrient nitrogen to the inoculum would help build fungal biomass and vigor, which in turn would improve biopulping performance of P. chrysosporium. To test this hypothesis, two concentrations of glutamic acid (500 and 5000 ppmN) were added to the 5% wood chips inoculum prior to introducing the fungus. The results suggested that increased inoculum nitrogen had beneficial effects in terms of energy savings. However, the weight loss stimulated by the high nitrogen offsets these benefits. Another approach that was tried to increase the inoculum vigor of P. chrysosporium was by increasing the age of the inoculum (2, 4, and 6 weeks). We concluded that inoculum age has little influence under the conditions used.

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In all of these experiments, precolonized wood chips were used as an inoculum. In other experiments, we evaluated a liquid inoculum (mycelial suspension) of C. subvermispora on aspen wood chips (0.1% on a dry wood basis). Results with the liquid inoculum were acceptable but not as good as the results with the chip inoculum. Because we did not measure the amount of fungus added with the colonized chips, we do not know whether the increased biopulping efficacy with chip inoculum was due to increased fungus content or an inherent advantage of delivering inoculum with wood chips.

2.3.3.3 Nutrients Requirement The effect of nitrogen and carbon was studied with P. chrysosporium on aspen chips. Two levels each of nitrogen (25 or 250 ppm N as glutamic acid) and glucose (4000 or 40 000 ppm) were added to bioreactors. The combination of 25 pprn nitrogen and 40 000 ppm glucose produced the best results in terms of energy savings. Subsequently, the effect of another nitrogen source (ammonium tartrate) (0, 36, 108, and 324 ppm N) was examined. We found that 324 ppm N gave optimum results. The use of a nonchemically defined nitrogen source, i.e., yeast extract, was also investigated with both P. chrysosporium and C. subvermispora on aspen wood chips. With P. chrysosporium, two levels of nitrogen (108 and 324 ppm) on a dry weight basis were tested with a uniform level of 40000 ppm lactose (dry weight basis). With C. subvermispora, three levels of nitrogen (108, 324 and 976 ppm) with a uniform level of 4 000 ppm lactose (dry weight basis) were used. An energy saving of 32% with significant strength improvements was obtained after 2 weeks, one of the best results ever achieved with P. chrysosporium. With C. subvermispora, an energy saving of 52% with strength improvements was obtained for the highest nitrogen level (976 ppm); this result was better than average for this fungus on aspen chips. In a few runs made without the addition of nutrients, we noted significant energy savings and paper strength improvements, although addition of nutrient nitrogen seemed to enhance the biopulping efficacy of these fungi. 2.3.3.4 Aeration Solid-substrate fermentations are known to be affected markedly by aeration. In addition, the ligninolytic activity of fungi depends on oxygen availability. Consequently, we evaluated the influence of three air flow rates on the biopulping efficacy of P. chrysosporium on aspen chips: low (0.001 11-1 min 1), medium (0.02211-1 rain- ~), and high (0.10011- ~min 1). The low flow rate was achieved using intermittent aeration. The medium and high flow rates gave comparable energy savings and had similar effects on strength properties. The low flow rate was suboptimal. 2.3.3.5 Wood Chip Sterilization Our studies showed that P. chrysosporium could colonize unsterilized wood chips and perform biopulping when incubated in a bioreactor at its optimum


179

growth temperature (39 ~ but not at suboptimal growth temperatures; C. subvermispora was found to be ineffective on unsterilized wood chips even at its optimum temperature (27-32 ~

2.3.3.6 Bleaching Studies Fungal pretreatment reduces the brightness of the resulting mechanical pulps by as much as 15 20 Elrepho brightness points in 4 weeks and 8-10 Elrepho brightness points in 2 weeks. Consequently, the bleachability and brightness stability of aspen biorefiner mechanical pulps (BRMPs) were investigated. Comparisons were made with untreated aspen refiner mechanical pulp (RMP) and commercial mechanical pulps: chemithermomechanical pulp (CTMP), thermomechanical pulp (TMP), and groundwood (GW) pulp [90]. P. chrysosporium was used for the pretreatments reported here; subsequent experiments demonstrated that aspen chips treated with C. subvermispora respond similarly to bleaching. Either alkaline hydrogen peroxide or sodium hydrosulfite readily increased the brightness of aspen BRMP (Table 9). Fungal pretreatment enhanced the bleachability of the aspen pulps examined. The BRMPs increased by more brightness points than did corresponding untreated pulps. However, because the initial brightness values of the BRMPs were lower than those of the corresponding untreated pulps, the bleached brightness values were not as high at a given chemical charge as those of untreated mechanical pulps. Aspen BRMP was readily bleached to 60% Elrepho brightness with 1% sodium h y d r o s u l f i t ~ a brightness suitable for newsprint; brightness values approaching 80% were achieved with a two-step bleach sequence. Thus, bleachability of biomechanical pulps appears not to be a serious problem. Table 9. Bleaching results with aspen pulps (90) Pulp

Chemical charge

GW

Unbleached 1.5% H202 1% Na2S204 Unbleached 3% H2Oz 1% NazS2O 4 Unbleached 3% HzO2 1% Na2SzO4 Unbleached 3% H2O 2 1% Na28204 Unbleached 3% H202 1% Na2S204 Two-step bleach 3% H202 1% NazSzO 4

CTMP

TMP

RMP (control)

BRMP

Brightness (%) 63.1 80.8 71.9 62.0 78.3 66.3 60.2 78.6 66.9 62.2 80.0 77.2 51.8 76.0 59.3 78.0

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M. Akhtar et al.

Table 10. Effluents from first refiner pass of control and fungus-treated aspen chips (91) Treatment

BOD (g kg-1 pulp)

COD (g kg 1 pulp)

Toxicity (100 EC50-1)

Control C. subvermispora

40 36

74 100

17 4

Brightness stability was evaluated by subjecting handsheets to accelerated thermal and photo-aging tests [90]. The stability of BRMP was slightly lower than that of RMP but slightly higher than that of CTMP.

2.3.3.7 Analysis of Effluents Samples of the wastewater from the first refiner passes of aspen chips pretreated with either C. subvermispora or P. chrysosporium were analyzed for biochemical oxygen demand (BOD), chemical oxygen demand (COD), and Microtox toxicity. Fungal pretreatment of aspen chips with either P. chrysosporium or C. subvermispora during biopulping substantially decreased effluent toxicity (Table 10). The BOD values for effluents from fungus-treated pulps were either slightly lower or higher than those of RMPs, depending upon the fungal species and whether nutrients were used. The COD values for effluents from fungustreated pulps were considerably higher than those from RMP runs, probably because of the release of lignin-related products. Based on these data, we conclude that the effluent load from the biopulping runs should be lower and more benign than that of commercial CTMP mills [91].

2.3.3.8 Global Analysis of Data The data from all runs, regardless of experimental conditions, were analyzed statistically in a global manner. The data were plotted as notched box plots [92] so that any statistically significant differences ( _+ 0.95 level) between the untreated control and biopulping runs could be observed. The global analysis greatly simplified the overall interpretation of a large amount of data. To illustrate the analysis, a comparison of the efficacy of fungal species in terms of refiner energy consumption in the biopulping of aspen chips is shown in Fig. 6. In subsequent analysis, P chrysosporium was found to be less effective than C. subvermispora on aspen wood chips and relatively ineffective on loblolly pine chips. The energy required for refining was reduced, in many cases by 40% or more (e.g., Fig. 6) with little loss of wood substance ( < 10%), except when chemically defined, high nutrient nitrogen supplements were used. Handsheet density was decreased somewhat by treatment of aspen with P. chrysosporium, and more with C. subvermispora; with pine, density was unaffected (Fig. 7). We treated P. chrysosporium/aspen, C. subvermispora/aspen, and C. subvermispora/pine as three separate sets of data, with all runs of these combinations


181

Wc Pt Ps

Pm Pc

9 "'F~'"

Pb ..... ~

None

Ds Cv

LL

........

9

.

Cs - ~

..... I

I

I

I

1000

1500

2000

2500

3000

Wc Pt

I

Ps

Pm Pc Pb

I

Fig. 6. Refiner energy consumption of aspen chips treated with different fungi (top dot-plot, bottom box plot) [52]. Wc Wolfiporia cocos (a brown-rot fungus), Pt Phlebia tremellosa, Ps

I

None Ds Cv Cs - C ~ f

1000

1500

I 2000

2;00

Phlebia subserialis, Pm Pholiota mutabilis, Pc Phanerochaete chrysosporium, Pb Phlebia brevispora, Ds Dichomitus squalens, Cv Coriolus (Trametes) versicolor, Cs Ceriporiopsis subvermispora (all white-rot fungi)

3000

Energy

included. Chip pretreatment with either fungus significantly increased burst index of handsheets from aspen but not that of handsheets from pine (Fig. 8). Chip pretreatment with either fungus also significantly increased tear index (single-ply test). Only C. subvermispora was examined with pine; tear index increased significantly (Fig. 9). Tensile index was increased significantly by chip pretreatment with either fungus in the case of aspen, but not pine (Fig. 10). Handsheet brightness was significantly decreased by pretreating chips of either wood with either fungus (Fig. 11), as was scattering coefficient with C. subvermispora on both woods and P. chrysosporium on aspen (data not shown); opacity was not significantly affected by either fungus (data not shown).

2.3.4 Microscopy Studies To gain insight into the mechanism of biopulping, we examined at the microscope level the growth patterns of the fungi in wood, the effects of fungi on wood cell walls, and the appearance of handsheets made from biopulps.

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M. Akhtar et al.

Pc

None

Cs

.@

m Q n Itl

250

I

I

I

350

450

550

Pc

None

Cs

250

!

I

I

350

450

550

Fig, 7. Density of handsheets made from aspen (top) and loblolly (bottom) pine chips treated with Phanerochaete chrysosporium (Pc) or Ceriporiopsissubvermispora (Cs) [52]

Density

2.3.4.1 Fungal Growth Patterns Scanning electron microscopy was used to observe fungal growth in and degradation of nutrient-supplemented aspen chips after a 3-week treatment with P. chrysosporium. The fungus grew well both across the chip surfaces (Fig. 12) and throughout the cell walls. The hyphae penetrated the chips through the lumina of wood vessels and fiber cells as well as through natural wood cell pits and fungal boreholes. Partial degradation of the cell lumen walls was evident. Erosion troughs and localized wall fragmentation or thinning were clearly visible as was a generalized swelling and relaxing of the normally rigid wood cell wall structure (Fig. 13). Aspen wood chips treated with C. subvermispora showed packed hyphae within the ray cells. Many crystals of calcium oxalate were found on the hyphae (Fig. 14) during both the incipient and the advanced stages of growth. Our observations suggest that the physical basis for the biopulping efficacy of the fungal treatment is likely to involve an overall softening and swelling of the wood cell walls as welt as thinning and fragmentation in localized areas [93].


I

Pc

~

~o

183

o

None

Cs

0

CL r

I

I

!

1

2

3

tLL

pc ~

@

None

Cs

0

I

I

I

1

2

3

Fig. 8. Burst index for handsheets made from aspen (top) and loblolly pine (bottom) chips treated with Phanerochaete chrysosporium (Pc) or Ceriporiopsis subvermispora (Cs) [52]

Burst index

2.3.4.2 Fibers In another investigation, we compared the microscopic appearance of B R M P with that of RMP, G W pulp, TMP, C T M P , neutral sulfite semichemical pulp (NSSC), and kraft pulp. When fiberized, B R M P emerged wool-like, looser, and bulkier, with fibers rather uniform in length; the pulp also exhibited abundant fibrillation. In contrast to BRMP, R M P fibers were not as wide, appeared to be stiffer, were of different lengths, and had only moderate fibrillation. The G W pulp fibers were stiff and of various lengths, showed reduced fibrillation, and were accompanied by debris. The T M P and C T M P fibers appeared stiff and were of various lengths, although longer than R M P fibers, and had moderate fibrillation. The T M P and C T M P fibers were more twisted than the B R M P fibers. The NSSC pulp exhibited few stiff fibers and the fibers were of various lengths; they appeared to be more compressible and conformable when compared to the mechanical pulp fibers. Compared to BRMP, NSSC pulp fibers were not as compressed and were more variable in length. Kraft pulping

184

M. Akhtar et al.

Pc

None

Cs

O~

0

O.

I

I

I

I

I

1

2

3

4

5

g LL

None

Cs

0

I

I

I

I

I

1

2

3

4

5

Fig. 9. Tear index (single-ply tear) for handsheets made from aspen (top) and loblollypine (bottom) chips treated with Phanerochaete chrysosporium (Pc) or Ceriporiopsis subvermispora (Cs) 1-52]

Tear index

produced more uniform, separated, and collapsed fibers, with abundant fibrillation. The BRMP appeared to be similar to the kraft pulp. These results were summarized by Sachs et al. [79, 94]. 2.3.4.3 H a n d s h e e t s

Aspen BRMP produced a stronger handsheet than did aspen T M P and G W pulp. Aspen NSSC pulps appeared to be superior to aspen BRMP in handsheet properties. Of all the pulps, aspen kraft pulp had the highest sheet strength properties. To gain insight and visually assess how the fiber morphology in these pulps may have contributed to sheet strength properties, we examined cross sections of handsheets. Handsheets made from mechanically processed pulps showed uncollapsed fibers, leading to poor conformability and reduced bonding. The NSSC and kraft pulps gave handsheets that exhibited fibers of enhanced compressibility and conformability. Handsheets prepared from B R M P visually resembled the kraft handsheets, exhibiting good compressibility and conformability of the fibers [79, 94].

Fungal Deligniflcation and Biomechanical Pulping of Wood

Pc

9

185

,

None

Cs

10

2'o 3'0 ,'o

I

50

6~0

== I.L

I

Pc

None

Cs

10

20 '

3'0

40 '

50 '

'0

Fig. 10. Tensile index for handsheets made from aspen (top) and loblolly pine (bottom) chips treated with Phanerochaete chrysosporium (Pc) or Ceriporiopsis subvermispora ( Cs) [52]

6

Tensile index

2.3.5 Engineering and Economic Studies The engineering and economics studies related to biopulping scale-up are summarized in the PhD thesis of Wall [95]. The goal of these studies was to develop the basic scientific and engineering knowledge needed for the commercial utilization of fungi in chip piles. We investigated process scale-up by obtaining kinetic data, developing process models, and implementing the processes on a pilot scale [96]. Initial studies were performed using P. chrysosporium on aspen wood chips. Mathematical modeling of biopulping was done to assess the factors that might kinetically limit fungal growth. Assumptions concerning the stoichiometry of biological wood degradation were used to predict important parameters needed to model the growth of P. chrysosporium in wood chips. Next, the effects of mass transfer on the growth of P. chrysosporium under typical biopulping conditions were considered. Finally, mass and energy balances were modeled for an isolated plug of wood chips, and the resulting system of differential equations was integrated numerically.

186

M. Akhtar et al.

Pc

o

..@

None

Cs

5'0

30

LL

o'o

!

7O

Pc None

~ I

;,o

30

I

Fig. 11. Brightness of handsheets made from aspen (top) and loblolly pine (bottom) chips treated with Phanerochaete chrysosporium (Pc) or Ceriporiopsis subvermispora (Cs) [52]

Brightness

Based on the kinetic and modeling studies, two bioreactor designs were considered: a packed-bed reactor and a chip-pile-based system (Fig. 15). The packed-bed bioreactor envisaged for the biopulping process is a bed of wood chips with controlled air flow. Packed-bed reactors allow better control of process conditions. The drawbacks to these reactors are that they require much greater capital expenditure and entail higher operating costs than chip-pile based systems. A chip-pile based system is defined here as an industrial-size chip pile modified to increase temperature and moisture control. The advantage of the chip pile is reduced cost as compared to that of a packed-bed reactor; the disadvantage is reduced process control. Early in the biopulping research, Harpole et al. [97] conducted an economic evaluation based on a thermomechanical process model. Results indicated that a 25% reduction in pulping energy by fungal treatment would save $21 (US) per air dry ton (adt) of pulp ($33 with 40% energy savings). The capitalized value of the energy savings was estimated to be about $250000 for each percentage of energy saved, at an electricity cost of $0.035/kWh. Thus, a sizeable capital


187

Fig. 12. Web-like hyphal network on surface of nutrient-supplemented aspen wood chip during 3-week treatment by Phanerochaetechrysosporium[93]. Bar = 1 mm

expenditure for the biotreatment could be accommodated. More recently, an economic model based on mass and energy balances was made for a controlled packed-bed reactor process and a chip-pile-based system. The controlled reactor process yields a pretax return on investment (ROI) of 21%, whereas the chip-pile-based system shows a pretax R O I of 1 0 6 % - 2 1 7 % (depending on the inoculum costs). The details are summarized in the following text.

2.3.5.1 Economic Analysis of Packed-Bed Reactor The capital and operating costs were estimated for a 300 metric ton/day biological pretreatment using P. chrysosporium on aspen and assuming a treatment time of 2 weeks and a dry weight loss of 5%. The air flow rate was assumed to be 0.59 VVM. The operating costs considered were steam, inoculum, and electricity for aeration. A reasonable figure for inoculum costs was chosen based on the information concerning a similar commercial process (see Section 2.3.6). The most optimistic scenario yields a pretax R O I of 21% (Table 11, Case 1).

2.3.5.2 Economic Analysis of Chip-Pile-Based System A chip-pile-based system with ductwork to provide aeration, piping to provide steam, and a sprayer to apply inoculum to chips on a conveyor was chosen for this analysis. The aeration rate and inoculum level were the same as those used for the packed-bed reactor; therefore, the operating costs for a chip-pile-based system are the same as those given for the packed-bed reactor. The capital costs

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M. Akhtar et al.

Fig. 13. The normally rigid cell wall structure within aspen chips (top) was modified (bottom) by 3-week treatment with

Phanerochaete chrysosporium [93]. Modifications included cell wall swelling (a), enzymatic softening or relaxing resulting in the partial collapse of the tube-like structure (b), and localized areas of wall thinning (c) or fragmentation (d). Bar = 10 gm

considered were those for fans, ductwork, i n o c u l u m tanks, and humidification. The analysis showed a pretax R O I of 106% 217%, depending on the i n o c u l u m costs (Table 12). O u r recent w o r k has focused on C. subvermispora and loblolly pine, with special attention to those factors m o s t likely to affect the economics of a chippile-based system. These factors are achieving the necessary degree of asepsis, lowering the cost of fungal inoculum, and maintaining a hospitable e n v i r o n m e n t in the chip pile.

2.3.5.2.1 Asepsis. In early work, the beneficial effects of biopulping on a laboratory scale were seen when w o o d chips were sterilized by autoclaving. C. subvermispora was not found to be aggressive e n o u g h to compete with indigenous microorganisms in unsterilized w o o d chips. Recently, we discovered that


189

Fig. 14. Calcium crystals on the surface of hyphae of Ceriporiopsissubvermisporaon aspen wood chips (4-weektreatment). 1 cm = 5 gm

a brief atmospheric steaming of the chips allowed C. subvermispora to colonize and be effective [98]. This condition is now being studied from an engineering standpoint.

2.3.5.2.2 Fungal Inoculum. One of the major costs foreseen during the scale-up of biopulping is inoculum production. Therefore, several experiments were aimed at determining the best inoculum level of C. subvermispora for saving energy and improving paper strength properties in a 2-week incubation. We found that 0.3% inoculum (dry weight basis) saved 19% energy and improved paper strength properties, such as tear index, significantly compared to the control (Table 13). This amount of inoculum is quite high. However, we discovered that the amount of inoculum can be lowered to 0.0005% (dry weight basis) or less by adding a cheap and commercially available nutrient source, corn steep liquor, to the mycelial suspension. This low amount of inoculum is now well within a commercial range. Subsequent studies have also identified better strains of C. subvermispora that yield up to 38% energy savings and improve tear index by 51% [98, 99]. Other nonchemically defined additives, including yeast extract and molasses, have shown promise in biopulping, but they have not been found to be as effective as corn steep liquor. 2.3.5.2.3 Chip Pile Environment. Instrumented bioreactors were constructed and are now in use to mimic chip piles. Current research is aimed at methods for controlling temperature, aeration, and moisture in chip piles so that the fungal treatment is effective. These data would support the design of a chip-pile-based system.

190

M. Akhtar et al.

Wood z= Harvesting Debarking Chipping

Chips ]=i..................... i

I

i (Asep~s) "--~ (Nutrient i components) .e----::-----~ (Nutrients) i ..................... -]~ IMIxer) :................... ]~,

Air

! i ~ : : i

>i'~'l'~"-~i;~r--~.._..T.....~. [ Humidifier i

i Air

I

.[

Recycle air

Inoculum train

I

I Ir~culum Softened chips). Mechanical pulping Biopolping reactor

Waste

Bleaching )

Bleed air

i Fan2 ! L.......... 1 =

i F~3i treaV'~nt

Fig. 15. Flowsheets for packed-bed reactor parentheses are optional [95]

(top) and

chip-pile-basedsystem (bottom). Operations in

2.3.6 Prospects for Biopulping Commercialization During the course of the biopulping research, one of the sponsoring companies developed and commercialized a biotechnological process that is similar to biopulping in many respects, and it deserves mention here. Cartapip is an industrial fungal inoculum of Ophiostoma piliferum which was developed by Sandoz Chemicals Biotech Research Corporation (now Clariant Biotech Research Corporation) and is marketed by the US Sandoz Chemicals Corporation (now Clariant Corporation). The fungus is a naturally occurring and ubiquitous "blue stain" organism. It is nonpathogenic. A dilute slurry of the product (a powder), consisting of fungal spores, is sprayed onto wood chips as they are piled for storage prior to pulping. The spores germinate and the fungus grows aggressively in the chip piles. Within the wood chips, the fungus grows mainly in ray

191

Fungal Delignification and Biomechanical Pulping of Wood Table 11. Economic feasibility of a packed-bed reactor [95] Case 1

Case 2

Case 3

Installed equipment costs Working capital

$5 000 000 $206 750

$10 000 000 $206 750

$17 000 000 $206 750

Total capital investment Utility costs Inoculum costs Labor Yield losses Depreciation

$5 206 750 $2.46 $3.00 $0.76 $2.46 $4.96

$10 206 750 $2.46 $3.00 $0.76 $2.46 $9.72

$17 206 750 $2.46 $3.00 $0.76 $2.46 $16.39

Total operating cost Pretreatment value Gross profit Pretax ROI

$13.64 $23.49 $9.85 21%

$18.40 $23.49 $5.09 5%

$25.07 $23.49 - $1.57 - 1%

Table 12. Economic feasibility of a chip-pile-based reactor 1-95] Case 1

Case 2

Case 3

Installed equipment costs Working capital Total capital investment

$500 000 $206 750 $706 750

$500 000 $206 750 $706 750

$500 000 $206 750 $706 750

Utility costs Inoculum costs Labor Yield losses Depreciation Total operating cost Pretreatment value Gross profit Pretax ROI

$2.46 $3,00 $0.76 $2.46 $0.76 $9.35 $23.49 $14.14 217%

$2.46 $5.00 $0.76 $2.46 $0.76 $11.35 $23.49 $12.14 180%

$2.46 $10.00 $0.76 $2.46 $0.76 $16.35 $23.49 $7.14 106%

Table 13. Energy savings and strength properties during biomechanical pulping of loblolly pine chips with C. subvermispora (2-week incubation) Strength properties Treatment (% inoculum on dry weight basis)

Energy savings~ (%)

Burst index (kNg 1)

Tear index (mN m 2 g 1)

Control 0.01 0.05 0.10 0.15 0.30

4 11 12 12 19

0.62 0.63 0.71 0.74 0.70 0.70

1.67 _+ 0.13 1.89 • 0.09 2.16 _+ 0.20 2.13 _+0.14 2.04 +_ 0.15 2.14 _+ 0.15

_+ 0.05 b _+ 0.04 _+ 0.04 +_ 0.03 + 0.06 + 0.05

a Energy savings are calculated on the basis of untreated control values b Standard deviation

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M. Akhtar et al.

parenchyma cells and resin ducts [100], where its distinctive activity is to degrade extractives within about 2 weeks during storage [101]. It is not capable of degrading lignin or cellulose, but it degrades hemicelluloses to a very small extent. Partial removal of extractives helps solve several problems associated with pitch, including downtime for cleaning equipment, breakage of paper on the paper machine, decreased paper strength, and holes in the paper caused by sticky spots on rolls [102]. The Cartapip process has shown other beneficial effects as well. It improves incoming chip quality by inhibiting the growth of dark-colored fungi [103, 104]. This in turn increases chip and pulp brightness, and reduces the need for bleaching chemicals. The Cartapip process also prevents wood losses caused by wood-degrading microorganisms. During chemical pulping, the process increases yields and reduces rejects because of the improved penetration of pulping liquors through empty resin canals and ducts of wood chips [105]. The development of the Cartapip process shows that biological treatment of wood chips can be successfully implemented on an industrial scale. At this point in the investigations, it would appear that biopulping has a good chance of commercial success. Four recent developments have led to this optimism: (1) the discovery of C. subvermispora, which is effective on both hardwood and softwood species, (2) the finding that brief steaming can decontaminate the surfaces of wood chips so that C. subverrnispora can take over, (3) the use of unsterilized corn steep liquor to dramatically reduce the inoculum quantity (from 3 kg to 0.25 g fungus per ton of wood [dry wood basis]), and (4) the demonstration of a successful 1-ton chip pile (green wood) where the fungal pretreatment saved 32% electrical energy. A recently conducted 100-ton (green wood) outdoor chip pile experiment produced results similar to those obtained using laboratory scale bioreactors. Our efforts so far have focused on the use of fungal treatments prior to refiner mechanical pulping. Recent studies in our laboratory and in others suggest that the fungal pretreatment is also effective for depitching [106] and that it gives benefits with thermomechanical pulping, chemithermomechanical pulping [107], sulfite pulping [106, 108, 109], and kraft pulping [110, 111].

Acknowledgements. We thank the following for their significant contributions to this research: Michael Attridge, Todd Burnes, Dough Cameron, Kory Cease, Leatha Dameron, Marilyn ElItand, Mary Greenheck, George Harpole, Eric Horn, Jane Kohlman, John Koning Jr., Gary Leatham, Michael Lentz, Ed Lightfoot, Efrat Livney, Lou Lunte, Gary Myers, Lewis Otjen, Jean Pierick, Sanya Reyes-Chapman,Irv Sachs, Gary Scott, Jane Sherwood,Marguerite Sykes,Freya Tan, Ted Wegner, and Mary Beth Wall. This research was partially funded by the following companies: Andritz Sprout-Bauer,Boise-CascadeCorporation, Cellulosa Puerto Piray S.A., Champion International Corporation, Chimica del Friuli, Consolidated Papers Incorporated, The Dow Chemical Company, Great Northern Nekoosa Corporation (Georgia Pacific), James River Corporation, Leykam Miirzataler AG, Mead Corporation, Mets/i Serla Oy, Nelco Chemical Company, Novo Nordisk A/S, Potlatch Corporation, Procter and Gamble Cellulose Company, Clariant Corporation, Scott Worldwide, Union Camp Corporation, and WeyerhaeuserPaper Company.


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3 References 1. Cowling EB (1961) US Dept Agric Tech Bull 1258 2. Wilcox WW (1968) Changes in wood microstructure through progressive stages of decay. US For Serv Res Pap FPL-70 3. Kirk TK, Cowling EB (1984) Biological decomposition of wood. In: Rowell RM (ed) The chemistry of solid wood. Adv Chem Series 207. American Chemical Society, Washington DC, pp 455-487 4. Eriksson KE, Blanchette RA, Ander P (1990) Microbial and enzymatic degradation of wood and wood components. Springer, Berlin, Heidelberg, New York 5. Kirk TK (1975) Holzforschung 29:99 6. Blanchette RA, Nilsson T, Daniel G, Abad A (1990) Biological degradation of wood. In: Rowell RM, Barbour RJ (eds) Archaeological wood: properties, chemistry and preservation, Adv Chem Set 225. Am Chem Soc, Washington, DC, p 141 7. Goni MA, Nelson B, Blanchette RA, Hedges JI (1993) Geochim. Cosmochim. Acta 57:3985 8. Savory JG (1954) Ann Appl Biol 41:336 9. Daniel G, Nilsson T (1988) Int Biodeterior 24:327 10. Daniel G, Nilsson T (1989) Inst Wood Sci 11:162 11. Blanchette RA, Simpson E (1992) IAWA Bull 13:201 12. Eaton RA, Hale MDC (1993) Wood: decay, pests, and protection. Chapman and Hall, London 13. Nilsson T, Daniel G, Kirk TK, Obst JR. (1989) Holzforschung 43:11 14. Blanchette RA (1991) Annu Rev Phytopathol 29:381 15. Otjen L, Blanchette RA, Effland M, Leatham G F (1987) Holzforschung 41:343 16. Blanchette RA, Burnes TA, Eerdmans ME, Akhtar M (1992) Holzforschung 46:109 17. Blanchette RA, Otjen L, Carlson MC (1987) Phytopathology 77:684 18. Otjen L, Blanchette RA, Leatham G F (1988) Holzforschung 42:281 19. Highley TL (1982) Can J For Res 12:435 20. Faix O, Mozuch MD, Kirk TK (1985) Holzforschung 39:203 21. Agosin E, Blanchette RA, Silva H, Lapierre C, Cease KR, Ibach RE, Abad AR, Muga P (1990) Appl Environ Microbiol 56:65 22. Liese W (1970) Annu Rev Phytopathol 8:231 23. Yoshizawa N, Watanabe J, Kobayashi S, Idei T (1989) Bull Utsunomiya Univ For 25:23-28 24. Srebotnik E, Messner K (1994) Appl Environ Microbiol 60:1383 25. Blanchette RA, Abad AR, Cease KR, Lovrien RE, Leathers TD (1989) Appl Environ Microbiol 55:2293 26. Blanchette RA, Abad AR, Farrell RL, Leathers TD (1989) Appl Environ Microbiol 55:1457 27. Ander P, Eriksson K-E (1977) Physiol Plant 41:239 28. Buswell JA, Odier E (1987) Crit Rev Biotechnol 6:1 29. Kirk TK, Farrell RL (1987) Annu Rev Microbiol 117:277 30. Tien M (1987) Properties of ligninase from Phanerochaete chrysosporium and their possible applications. Critical Reviews Journals in Microbiology 15(2): 141 168, CRC Press, Boca Raton, FL 31. Gold MH, Alic M (1993) Microbiological Reviews 57:605 32. Hatakka A (1994) FEMS Microbiology Reviews 13:125 33. Srinivasan C, D'Souza TM, Boominathan K, Reddy CA (1995) Appl Environ Microbiol 61: 4274 34. Rieble S, Joshi D, Gold MH (1994) J Bacteriol 176:4838 35. Hammel KE, Jensen KA Jr, Mozuch MD, Landucci L, Tien M, Pease EA (1993) J Biol Chem 268:12274 36. Koduri RS, Tien M (1994) Biochemistry 33:4225 37. Ruttimann C, Schwember E, Salas L, Cullen D, Vicuna R (1992) Biotechnol Appl Biochem 16:64 38. Popp JL Kalyanaraman B, Kirk TK (1990) Biochemistry 29:10475 39. Higuchi T (1993) J Biotechnol 30:1 40. Wariishi H, Valli K, Gold MH (1991) Biochem Biophys Res Commun 176:269 41. Bao W, Fukushima Y, Jensen Jr. KA, Moen MA, Hammel KE (1994) FEBS Lett 354:297 42. Call HP, Mucke I (1994) State of the art of enzyme bleaching and disclosure of a breakthrough process. Paper presented at the 1994 International Nonchlorine Bleaching Conference, Amelia Island, FL

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43. Cullen D, Kersten P (1992) Fungal enzymes for lignocellulosie degradation. In: Kinghorn JR, Turner G (eds) Applied molecular genetics of filamentous fungi. Blackie, Glasgow, UK, p 100 44. Edwards SL, Raag R, Wariishi H, Gold MH, Poulos TL (1993) Proc Natl Acad Sei USA 90: 750 45. Pointek K, Alumoff P, Winterhalter K (1993) FEBS Lett 315:119 46. Sundaramoorthy M, Kishi K, Gold MH, Poulos TL (1994) J Biol Chem 269:32759 47. Daniel G, Nilsson T, Petterson B (1989) Appl Environ Microbiol 55:871 48. Daniel G, Petterson P, Nilsson T, Volc J (1990) Can J Bot 68:920 49. Daniel G, Jellison J, Goodell B, Paszczynski A, Crawford R (1991) Appl Mierobiol Biotechnol 35:674 50. Srebotnik E, Messner K, Foisner R (1988) Appl Environ Microbiol 54:2608 51. Slatin B (ed) (1992) 1992 Statistics of paper, paperboard and wood pulp. American Paper Institute, New York 52. Kirk TK, Koning JW Jr, Burgess RR, Akhtar M, Blanchette RA, Cameron DC, Cullen D, Kersten P J, Lightfoot EN, Myers GC, Sykes M, Wall MB (1993) Biopulping: A glimpse of the future? Res Rep FPL-RP-523, Madison, WI 53. Kirk TK, Akhtar M, Blanchette RA (1994) Biopulping: Seven years of consortia research. Tappi Biological Sciences Symposium, pp. 57-66, Tappi Press, Atlanta, GA 54. Lawson LR, Still CN (1957) Tappi J 40: 56A 55. Ander P, Eriksson K-E (1975) Svensk Papperstidning 18:641 56. Eriksson K-E, Ander P, Henningsson B, Nilsson T, Goodell B (1976) Method for Producing Cellulose Pulp. US Patent 3 962 033 57. Johnsrud SC, Eriksson K-E (1985) Appl Microbiol Biotechnol 21:320 58. Eriksson K-E, Johnsrud SC, Vallander L (1983) Arch Microbiol 135 161 59. Eriksson K-E (1990) Wood Sci Technol 24:79 60. Samuelsson L, Mjober P J, Hartler N, Vallander L, Eriksson K-E (1980) Svensk Papperstidning 8:221 61. Johnsrud SC, Fernandez N, Lopez P, Guitierrez I, Saez A, Eriksson K-E (1987) Nordic Pulp & Paper Research Journal, Special Issue 2:47 62. Eriksson K-E, Grunewald A, Vallander L (1980) Biotechnol Bioeng 22:363 63. Bar-Lev SS, Kirk TK, Chang H-M (1982) Tappi J 65:111 64. Akamatsu I, Yoshihara K, Kamishima H, Fujii T (1984) Mokuzai Gakkaishi 30:697 65. Eriksson K-E, Kirk TK (1985) Biopulping, biobleaching and treatment of kraft bleaching effluents with white-rot fungi. In: Cooney CL, Humphery AE (eds) The Principles of Biotechnology: Engineering Considerations. In: Moo-Young M (ed). Comprehensive Biothechnology: The Principles, Applications and Regulations of Biotechnology in Industry, Agricultural and Medicine, Pergamon, New York, p 271 66. Kirk TK, Burgess RR, Koning JW Jr (1992) Use of fungi in pulping wood: An overview of biopulping research. In: Leatham GF (ed) Frontiers in industrial mycology. Proceedings of Industrial Mycology Symposium, June 25-26, 1990, Madison, WI. Routledge, Chapman and Hall, New York, Chapter 7, p 99 67. Setliff ED, Eudy WW (1980) Screening white-rot fungi for their capacity to delignify wood. In: Kirk TK, Chang H-M, Higuchi T (eds) Lignin biodegradation: microbiology, chemistry, and protein applications (vol 1). CRC, Boca Raton, FL, p 135 68. Blanchette RA (1984) Appl Environ Microbiol 56:210 69. Nashida T (1989) Mokuzai Gakkaishi 35:675 70. Kimura Y, Asada Y, Kuwahara M (1990) Appl Microbiol Biotechnol 32:436M42 71. Blanchette RA, Burnes TA, Leatham GF, Efftand MJ (1988) Biomass 15:93 72. Leatham GF, Myers GC, Wegner TH, Blanchette RA (1990) Energy savings in biomechanical pulping. In: Kirk TK, Chang H-M (eds) Biotechnology in pulp and paper manufacture Applications and fundamental investigations. Butterworths-Heinemann, Boston, p 17 73. Leatham GF, Myers GC, Wegner TH, Blanchette RA (1990) Tappi J. 72:249 74. Eriksson K-E, Vallander L (1982) Svensk Papperstid. 85:R33 75. Leatham GF, Myers GC (1990) Tappi J 72:192 76. Akhtar M, Leatham GF, Myers GC, Attridge MC (1989) PFI milling: A possible method to assess both energy savings and paper strength properties in biomeehanical pulping. Conf Abs Fourth International Conference on Biotechnology in the Pulp and Paper Industry, Raleigh, NC 77. Simons FL (1950) Tappi J 33:312 78. Wurz O (1969) The Paper Maker 38:59

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Sachs IB, Leatham GF, Myers GC, Wegner TH (1990) Tappi J 73:249 Blanchetter RA, Akhtar M, Attridge MC (1992) Tappi J 75:121 Akhtar M, Blanchette RA, Burnes T (1995) Wood Fiber Sci 27:258 Myers GC, Leatham GF, Wegner TH, Blanchette RA (1988) Tappi J 73:105 Akhtar M, Attridge MC, Myers GC, Kirk TK, Blanchette BA (1992) Tappi J 75:105 Akhtar M, Attridge MC, Blanchette RA, Myers GC, Wall MB, Sykes MS, Koning Jr JW, Burgess RR, Wegner TH, Kirk TK (1992) The white-rot fungus Ceriporiopsis subvermispora saves electrical energy and improves strength properties during biomechanical pulping of wood. In: Kuwahara M, Shimada M (eds) Biotechnology in pulp and paper industry. UNI Publishers, Kyoto, Japan, p 3 Leatham GF (1983) Mycologia 75:905 Akhtar M, Attridge MC, Myers GC, Blanchette RA (1993) Holzforschung 47:36 Akhtar M (1994) Holzforschung 48:199 Leatham GF, Myers GC, Wegner TH (1990) Tappi J 73:197 Blanchette RA, Leatham GF, Attridge MC, Akhtar M, Myers GC (1991) Biomechanical pulping with C. subvermispora. US Patent no. 5 055 159 Sykes MS (1993) Tappi J 76:121 Sykes M (1994) Tappi J 77:160 McGill R, Tukey JW, Larsen WA (1978) The American Statistician 32:12 Sachs IB, Leatham GF, Myers GC (1989) Wood Fiber Sci 21:331 Sachs IB, Leatham GF, Myers GC, Wegner TH (1990) Biomechanical pulping of aspen chips: Fungal growth pattern and effects on cell wall, fiber, and pulp morphology. In: Kirk TK, Chang H-M (eds) Biotechnology in pulp and paper Manufacture-Applications and fundamental investigations. Butterworths-Heinemann, Boston, p 27 Wall MB (1993) Biopulping process design and analysis, PhD Thesis, University of Wisconsin Chemical Engineering Department Wall MB, Cameron DC, Lightfoot EN (1993) Biotechnology Advances 11:645 Harpole GB, Leatham GF, Myers GC (1989) Economic assessment of biomechanical pulping. In: Proceedings of the international mechanical pulping conference 1989 Mechanical pulp responding to end product demands (vol 2). Multiprint, Helsinki, p 398 Akhtar M, Kirk TK, Blanchette RA (1996). Biopulping: An overview of consortia research. In: Srebotnik E, Messner K (eds) Biotechnology in the pulp and paper industry: Recent advances in applied and fundamental research, Facultas-Universit~tsverlag, Berggasse 5, A-1090 Wien, Austria, p 187 Akhtar M, Lentz M J, Blanchette RA, Kirk TK. Corn steep liquor lowers the amount of inocutum for biopulping. Tappi J 1997, Vol 80, Nov 2 Blanchette RA, Farrell RA, Burnes TA, Wendler PA, Zimmerman W, Brush TS, Snyder RA (1992) Tappi J 75:102 Brush TS, Farrell RL, Ho C (1994) Tappi J 77:155 Farrell RA, Blanchette RA, Brush TH, Gysin B, Hader Y, Perollaz J-J, Wendler PA, Zimmermann W (1992) Cartapip: A biopulping product for control of pitch and resin acid problems in pulp mills. In: Kuwahara M, Shimada M (eds) Biotechnology in pulp and paper industry. UNI, Kyoto, Japan, p 163 Blanchette RA, Behrendt CJ, Farrell RL (1994) Biological protection of sapstain for the forest products industry. Tappi Biological Sciences Symposium, Tappi, Atlanta, GA, pp 77-80 Behrendt CJ, Blanchette RA, Farrell RL (1995) Phytopathology 85:92 Wall MB, Brecker J, Fritz A, Iverson S, Noel Y (1994) Cartapip treatment of wood chips to improve chemical pulping efficiency. Tappi Biological Sciences Symposium, Tappi, Atlanta, GA, pp 67 76 Fischer K, Akhtar M, Blanchette RA, Burnes TA, Messner M, Kirk TK (1994) Holzforschung 48:285 Myers GC, Akhtar M, Lentz M, Scott GM, Sykes MS (1996) Biological pretreatment for thermomechanical (TMP) and chemithermomechanical (CTMP) pulping processes. Conf Abs 21 lth American Chemical Society National Meeting, New Orleans, LA Messner K, Srebotnik E (1994) FEMS Microbiology Reviews 13:351 Scott GM, Akhtar M, Lentz M, Sykes M, Abubakr S (1995) Environmental aspects of biosulfite pulping. Tappi Environmental Conference (Book 2). Tappi Press, Atlanta, GA, p 1155 Oriaran TP, Labosky P Jr, Blankenhorn PR (1990) Tappi J 73:147 Oriaran TP, Labosky P Jr, Blankenhorn PR (1991) Wood Fiber Sci 23:316

Solving Pitch Problems in Pulp and Paper Processes by the Use of Enzymes or Fungi Roberta L. Farrell 1, Kunio Hata 2 and Mary Beth Wall 3 1 Dept. of Biological Sciences, University of Waikato, Hamilton, New Zealand 2 Nippon Paper Industries Co., Ltd. 5-21-10ji, Kita-ku, Tokyo 114, Japan 3 Clariant Corp, 4000 Monroe Rd, Charlotte, NC 28205, USA

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Extractive Degradation During "Natural" Storage . . . . . . . . . . . . . . . . . . . . . . 3 Degradation of W o o d Extractives by Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Molds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Basidiomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Sap-Stain Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Industrial Use of Fungi to Solve Pitch Problems . . . . . . . . . . . . . . . . . . . . . 4 Enzymatic Pitch Control in the Papermaking Process . . . . . . . . . . . . . . . . . . . . 4.1 F u n d a m e n t a l Research and Theory of Lipase Application . . . . . . . . . . . . . . . 4.1.1 Identification of C o m p o u n d Causing Pitch Trouble . . . . . . . . . . . . . . . . 4.1.2 Application of Lipase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Effect of Lipase Treatment on Prevention of Pitch Deposition . . . . . . . . . . . . . 4.3 Application to Papermaking Process ........................... 4.4 Mill Trial I . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Mill Trial II . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

198 199 200 200 201 202 204 206 206 206 207 207 208 208 210 211 211

Pitch problems in pulp mills are often caused by the resinous materials (pitch) in wood which comprise approximately 2 - 8 % of total composition depending upon the species and time of year. Traditional methods to control pitch problems include natural seasoning of wood before pulping and/or adsorption and dispersion of the pitch particles with chemicals in the pulping and papermaking processes, accomplished by adding fine talc, dispersants and other kinds of chemicals. Within the last five years, two new and different methods of combatting pitch, both of which are biotechnological in basis, have been developed independently and are now used industrially. H a t a and colleagues of Nippon Paper Industries developed a pitch control m e t h o d using the enzyme lipase, which catalyzes the hydrolysis of triglycerides. Farrell and colleagues of Sandoz Chemicals Biotech (now known as Clariant) developed a method of pitch control and biocontrol using a fungus developed in the laboratory from the same type of organisms which cause natural aging, the Ascomycetes. This fungus is non-colored and prevents the staining and decrease of brightness normally associated with aged wood.

Advancesin BiochemicalEngineering/ Biotechnology,Vol. 57 ManagingEditor: T. Scheper 9 Springer-VerlagBerlin Heidelberg 1997

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1 Introduction In today's pulp and paper industry, pitch trouble is often caused by the resinous materials (pitch) in wood. These are some of the materials ("extractives") that are extracted from wood during the pulping process, and comprise about 2-8% of its total composition, depending upon the species and time of year. Not all of the extractives are troublesome, most problems occurring in pulping and papermaking when there are shifts in pH and/or temperature. During the pulping process, these resinous materials are released from wood and later stick to the tile and metal parts including the rolls and wires of the papermaking machines. The pitch also stains the felts and canvas, and eventually reaches the dryer section. This pitch accumulation can cause paper spotting and web breaks on the machine, which are severe problems in production. Pitch content arid severity of problems from pitch vary with wood species. The pitch of pines, including loblolly, slash, and red pines is known to cause serious problems. Hardwood pitch, particularly from tropical hardwood species, and eucalyptus can also be detrimental. Traditional methods of controlling pitch problems include seasoning of wood before pulping. Seasoning requires raw wood logs (roundwood) to be left outdoors for several months or chips to be piled and left for weeks. It is the most commonly used method around the world because wood extractives are decomposed during the seasoning process. However, accompanying seasoning are potential losses due to biological deterioration, such as decreased pulp brightness and pulp yield. Moreover, seasoning increases working capital costs due to high wood inventory and land use. Thus, this method is often unacceptable, especially in areas where space is limited. Another method used to reduce the accumulation of pitch is the adsorption and dispersion of the pitch particles with chemicals in pulping and papermaking processes. This is accomplished by adding fine talc, dispersants, and other kinds of chemicals. In Japan, red pine is the most important wood for groundwood pulp. Red pine groundwood pulp has high opacity and printability. Therefore, it is an indispensable pulp for the manufacture of newsprint and light weight paper. However, the red pine groundwood pulp contains a large amount of pitch. Hata et al. conducted fundamental research and determined that pitch trouble was caused by triglycerides within the resinous materials in wood [1-1. These triglycerides form a nucleus upon which other resinous materials tend to accumulate, causing pitch troubles. Hata et al. developed a new pitch control method using the enzyme lipase [1 3]. This method was put into practice in a large scale papermaking process as a routine operation in the early 1990s, and was the first case in the world in which enzyme was successfully applied in the actual papermaking process. In the USA, Farrell and coworkers independently and concurrently were also studying biotechnological solutions to pitch problems [4, 5]. These studies were initiated with the goal of solving pitch problems in loblolly pine by the

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application of a fungus developed in the laboratory from the same Class of organism which causes natural aging, the Ascomycetes [6]. This organism, Ophiostoma piliferum, belongs to the sap-staining type of organisms, but was bred in the laboratory to be colorless and non-staining. Thus, the positive benefits of aging were achieved with increases in pulp brightness. Also, by directly applying the fungus to the wood chips or logs, the effect of aging was accelerated. This product, marketed as Cartapip, was the first case in the world where an organism was commercially successfully applied prior to pulping to achieve beneficial effect [-7, 8].

2 Extractive Degradation during "Natural" Storage Living cells are contained in the bark, foliage, and sapwood when the tree is cut. These cells remain viable for periods of up to six months when the wood is stored. The living cells in the wood rays (ray parenchyma) respire and release heat. Bacteria and fungi are provided with good growth conditions during this heat generation, and the starches and simple sugars of the rays and subsequently by the extractives of the wood can be metabolized as a source of carbon and energy. This metabolism results in an overall decrease of pitch with storage of wood. The outside storage of pulpwood was introduced in the 1920s as whole logs (roundwood), and in the early 1950s as chips [9]. This method was the direct result of the need to stockpile wood as inventory to mills, to handle intermittent flow of chips to the mill, and to season wood, which resulted in decreased resin deposition. The reduction in pulp brightness and yield during storage was shown to be of the same order whether the wood had been stored as chips or as roundwood [10]. Conditions which affect the wood were shown to be the following: species of wood, time of cutting, removal of bark, presence of insects, methods of piling, length of storage time, general housekeeping conditions in the woodyard, and climatic conditions, especially temperature and moisture. Temperature appeared to be the single most important factor affecting distribution and prevalence of microorganisms in various sections of the chip pile in one study on microbiological effects of seasoning on hardwoods 1-11]. Outside storage of white spruce (Picea glauca) and lodgepole pine (Pinus contorta) showed decreased wood substances by 3.8% and 4.5% respectively after 6 months. Most of this decrease was attributed to pitch components [12]. This study also showed that pine kraft pulp yield increased based on seasoned chips, though spruce kraft pulp yield decreased slightly with time of storage. Seasoning has been recommended for pine unless the recovery of maximum tall oil and turpentine yields was desired [13]. Eighty percent losses in tall oil and turpentine yields resulted after 30 weeks storage. Pulp strength in this and other studies was shown not to be affected by seasoning, with the possible

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exception of tear strength. Although seasoning chips reduces pitch troubles, the negative effects of seasoning on various wood species were also thoroughly studied in North America and Europe [14, 15]. The single most detrimental effect was loss of brightness of mechanical and sulphite pulps after storage, particularly with softwoods [10].

3 Degradation of Wood Extractives by Fungi A variety of wood-inhabiting fungi including molds, sap-stain, brown-rots, and white-rots are capable of degrading wood extractives.

3.1 M o l d s Although molds are capable of degrading wood extractives, the contribution of molds to extractive degradation is expected to be minimal because molds grow less prolifically in wood than do other wood-inhabiting fungi. Molds grow best on wood that is very wet or that has been exposed to very high humidity for a long time. On softwoods, molds grow mainly on wood surfaces. On hardwoods, molds can enter the wood at exposed parenchyma, vessels, and ruptured cells and can move throughout the wood by rupturing pit membranes [16]. Nilsson and Asserson have shown that the following molds degrade wood waxes in liquid culture: Penicillium roqueforti, Penicillium funicolosum, Rhizopus arrhizus, and Trichoderma lignorum [17]. In addition, they showed that the following molds could reduce the ethanol/benzene (1:2) extractive content of wood chips: R. arrhizus, Gliocladium viride, Penicillium rubrium, T. lignorum, and Aspergillus fumigatus. The wood chips were

Table 1. DCM extractive content of nonsterile southern yellow pine treated with various molds

Fungal Species

Phlebia roqueforti Leptographium terrebrantis Verticicladiella truncata Diplodia pinea Codinaea sp. Aureobasidium pullulans

Extractives (%) Control 1

Extractives (%) Treatment

Reduction %

3.34 2.27

2.16 1.92

35 15

2.27

1.96

14

2.27 2.27 3.34

2.01 2.07 3.26

11 9 2

1 The control was chips that had been frozen at -- 20 ~ since the start of the experiment

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stored at 35~ and sampled at 30 days. The type of wood chips tested was not given. Iverson et al. screened various molds for their ability to degrade wood extractives 1-18]. Nonsterile southern yellow pine chips were inoculated with 104 to 108 colony-forming units/g wet weight wood and incubated at room temperature for 2 weeks. As shown in Table 1, the best fungus tested was P. roqueforti, which reduced the dichloromethane (DCM) extractive content by 35%.

3.2 Basidiomycetes Basidiomycetes, including white-rot and brown-rot fungi, extensively colonize wood. Brown-rot fungi preferentially degrade wood polysaccharides including cellulose and cause rapid decreases in its degree of polymerization. Brown-rotted wood usually shows virtually no decrease in total lignin content. Rather than degrading lignin, brown-rot fungi modify it by oxidation and demethylation of methoxy groups. White-rot fungi are the predominant degraders of lignin in nature. Some species of white-rot fungi preferentially degrade lignin to wood polysaccharides, and other species degrade all wood components simultaneously. The Basidiomycetes have been shown to degrade pitch extractives. Several fungi observed to have "biopulping" activity have also been shown to degrade wood extractives. Biopulping involves the use of fungally treated chips to obtain pulping benefits such as reduced energy used during mechanical pulping or improved chemical pulping efficiency. Lim and Cho have shown that the ethanol/benzene (1:2) extractive content of oak treated with Phanerochaete chrysosporium decreased by 46% after twelve weeks [19]. Treatment of sterile southern yellow pine wood chips with P. chrysosporium for 2 weeks resulted in a 21% reduction in dichloromethane (DCM) extractives [20]. Fischer et al. have shown that the DCM extractive content of sterile loblolly pine chips treated with C. subvermispora decreased after four weeks by 32% [21]. In addition, they showed that both C. subvermispora and P. chrysosporium decrease the DCM extractive content of sterile spruce chips [21]. Various Basidiomycetes were screened for their ability to degrade wood extractives [-18]. Sterile southern yellow pine was inoculated with 104 to l0 s colony-forming units/100 g wet weight wood and incubated for 2 weeks at room temperature. As shown in Table 2, P. chrysosporium performed best, reducing DCM extractives by 51%. Hyphodontia setulosa, Perenniporia subacida, P. gigantea, and Phlebia tremellosa also performed well, reducing extractives by about 40%. Poller and Schultze-Dewitz investigated the effect of two brown-rot fungi, Coniophora puteana and Gloeophyllum saepiarium, and one white-rot fungus, Phellinus igniarius, on the extractive content of pine [22]. All three fungal treatments resulted in large reductions in extractive content.

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Table 2. Extractive content of sterile southern yellow pine treated with various Basidiomycetes

Fungal species

Control Extractives (%)

Treatment Extractives (%)

Reduction (%)

Phanerochaete chrysosporium P. subacida P. gigantea P. tremetIosa H. setulosa Coriolus versicolor Inonotus rheades Trichaptum abietinium C. subvermispora Trichaptum biforme Schizophylum commune Sistotrerma brinkmanii Pleurotus ostreatus Alurodiscus sp. Ganoderma collosum PheUinus igniarius

2.19 3.34 3.34 1.98 1.98 1.98 3.34 4.70 3.34 4.70 2.50 2.44 2.44 2.44 2.29 2.44

1.30 2,01 2.03 1.21 1.20 1,28 2,18 3.13 2.18 3.13 2.03 2.17 2.30 3.70 1.75 2.48

41 40 39 39 39 36 34 33 29 24 17 11 6 0 0 0

3.3 Sap-Stain Fungi Sap-stain fungi rapidly colonize the sapwood of logs and wood chips. These fungi grow mainly in ray parenchyma cells and are capable of deeply penetrating logs and wood chips. In addition, these fungi can grow within resin canals, tracheids, and fiber cells, and penetrate simple and bordered pits, occasionally forming boreholes through wood cell walls. Sap-stain fungi are not capable of degrading the major components of the wood cell wall: cellulose and lignin. Hemicellulose is degraded to a very slight degree. Extractives and simple sugars found in the parenchymal cells are the major nutrient source for these fungi. Sap-stain fungi cause a characteristic staining of sapwood, resulting in a blue, black, grey, or brown discoloration of the wood. Sap-stain causes major economic losses in the lumber and mechanical pulping industries. Problems with sap-stain are most prevalent in warm, humid climates and when wood with a high sapwood content is used. Common species of sap-stain on softwoods include: Ophiostoma ips,

O. piliferum, O. piceae, Aureobasidium pullulans, Leptographium lundbergii, Alternaria alternata, Cephaloascusfragrans, Cladosporium spp., Lasiodiplodia theobromae, and Phiolophora spp. [-23]. Common species of hardwood sap-stain include: Ophiostoma pluriannulatum, Ceratocystis moniliformis, L. theobromae, Ceratocystis rigidum [23]. Many of these species are capable of degrading wood extractives. Extractive degradation by Ophiostoma spp., particularly O. piliferum and O. piceae, has been most widely studied. Iverson et al. screened a variety of sap-stain fungi for the ability to degrade wood extractives [18]. Sterile southern yellow pine was inoculated with the fungi listed in Table 3 and incubated at room temperature for 2 weeks. The best


203

Table 3. Extractive degradation by sap-stain fungi on nonsterile southern yellow pine Fungal Species

Control Extractives (%)

Treated Extractives (%)

Reduction (%)

c. adiposa O. piliferum C. adjuncti C. minor O. piceae O. populina O. abiocarpa C. tremuloaurea O. fraxinopennsylvanica O. plurianulatum E. aereum C. ponderosa C. penicullata O. olivaceum E. clavigerum C. hunti C. ambrosia O. distortum E. robustum C. virescens O. 9aleiformis C. coerulescens O. dryocetidis O. stenorcerns X. conudamae X. hypoxylon

2.13 3.34 1.98 2.13 2.13 2.13 2.13 2.13 2.13 2.13 1.98 2.19 1.98 2.27 2.13 2.13 2.13 2.27 2.27 2.27 2.13 2.13 2.24 2.13 2.44 2.44

1.26 2.27 1.44 1.57 1.57 1.62 1.61 1.65 1.71 1.73 1.61 1.86 1.58 1.93 1.84 1.89 1.92 2.07 2.06 2.11 1.98 2.13 2.27 2.21 3.16 2.88

41 32 27 26 26 24 24 23 20 19 19 18 15 15 14 11 I0 9 9 7 7 0 0 0 0 0

species for extractive reduction were C e r a t o c y s t i s adiposa, O. piceae, and O. piliferum. Nine different isolates of O. piliferum were also screened on sterile southern yellow pine using the same procedure; 22% of the isolates did not reduce D C M extractives and 55% of the isolates reduced D C M extractives by 25 35%. In addition, 45 different strains of O. piceae were screened on sterile aspen chips. These strains can be divided into four groups based on their ability to reduce ethanol/toluene extractives: 24% of the isolates did not degrade extractives, 46% reduced extractives by 1-15%, 28% reduced extractives by 16 35%, and one isolate reduced extractives by 60%. Chen et al. studied the effect of five sap-stain fungi on the composition of aspen and lodgepole pine extractives [24]. Four fungi were selected on the basis of high lipolytic activity from 100 fungal strains isolated from Canadian lumber mills and compared with the commercial strain, Cartapip | 97. Analysis of untreated wood showed that triglycerides were the most abundant component of both aspen and lodgepole pine sapwood extractives. The wax and steryl ester content of aspen was about 3 times that of lodgepole pine, and fatty and resin acids were the second most c o m m o n component of lodgepole pine extractives but were present in very small amounts in aspen. All five fungi decreased the total acetone extractive content of aspen and lodgepole pine sapwood to

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Table 4. Extractive content of sterile lodgepole pine and aspen treated with sap-stain fungi Treatment

Aspen Extractives (%)

Lodgepole pine Extractives (%)

Control Aged Control Cartapip 97 Strain A Strain B Strain C Strain D

3.09 _+ 0.07 2.88 _+ 0.04 2.15 _+ 0.01 2.13 _+ 0.02 2.22 _+ 0.01 2.07 _+ 0.04 2.08 + 0.05

2.31 _+ 0.03 2.26 _+ 0.02 1.94 _+ 0.03 2.08 _+ 0.01 NA 1.92 _+ 0.11 1.92 + 0.01

a similar degree - 28-33% for aspen and 1 ~ 1 7 % for lodgepole pine (Table 4). Extractive component analysis showed that all five fungi decreased triglyceride content and steryl esters/waxes content, and that four of the five fungi increased free fatty acid content.

3.4 Industrial Use o f Fungi to Solve Pitch Problems Several studies have shown that wood extractive components such as triglycerides, resin acids, and steryl esters are major components of paper machine pitch deposits [1, 2, 6]. In addition, pitch outbreaks are more common when resinous wood species are used and during seasons when wood resin content is particularly high. There is a living fungus, marketed to the pulp and paper industry, which metabolizes and thus removes pitch. This fungus is a colorless strain of O. piliferum, an Ascomycete of the same species of that often dominates in naturally seasoned piles. Marketed as Cartapip, with different numbers denoting different strains such as 97 and 58, it is commercialized as a powder inoculum. Moreover, Cartapip use results in a biocontrol effect, i.e., the presence of Cartapip reduces growth of other, undesired organisms. One kilogram of the powder can treat about 1200 tons of wood chips. Industrial use involves dispersing the powder in mill water and spraying it onto chips as they are conveyed to a chip pile. Selective breeding was used to obtain this isolate, which rapidly colonizes nonsterile wood chips, rapidly degrades extractives, and is colorless and nonstaining. Most O. piliferum strains are a bluish-black color. Growth of pigmented fungi on wood chips reduces chip brightness and increases bleach usage when these chips are used to produce T M P or sulfite pulp. Because Cartapip outcompetes indigenous microorganisms and maintains chip brightness, use of this product reduces bleach chemical usage during T M P production, in addition to reducing the extractive content of chips and pulp, and alleviating pitch problems. Use of treated chips has also been shown to increase paper strength [25]. Moreover, treatment of wood chips with Cartapip also results in improved


205

chemical pulping efficiency [26]. Reductions in kappa number were observed during laboratory-scale kraft and sulfite pulping. Wall et al. hypothesize that the improved pulping efficiency observed experimentally is caused by more rapid and more uniform penetration of steam and cooking chemicals in the fungally treated chips [27]. Two commercially available strains of O. piliferum, Cartapip 28 and Cartapip 58, have been shown to degrade the extractives of both hardwoods and softwoods including aspen, southern yellow pine, red pine, and spruce [27], Both fungi in two weeks reduced the diethyl ether extractive content of fresh nonsterile southern yellow pine wood chips by 40%, or 22% if the chips were aged but not inoculated. In addition, the white strain, Cartapip 58, maintained chip brightness. This study also showed that Cartapip 28 decreased the D C M extractive content of sterile southern yellow pine chips by 30% for a 2 week treatment. O. piliferum has also been shown to reduce more DCM extractives of nonsterile red pine in 31 days (48% reduction) than in 21 days (32% reduction) [18]. O. piliferum, strain Cartapip 97, also reduces the DCM extractives of spruce by 25% in a 2 week treatment, and the DCM extractive content of sterile loblolly pine chips by up to 35% in a 4 week treatment [21]. Cartapip 97 reduced the acetone extractive content of fresh nonsterile aspen chips by 36% after a 3 week incubation, or 13% for uninoculated aged chips [27]. Cartapip 28 and Cartapip 58 significantly decreased the fatty acid content and the unidentified compound content, which includes waxes, alcohols, and sterols, of southern yellow pine extractives l-6]. In particular, Cartapip 58 decreased esterified fatty acids by 60%. Both esterified fatty acids and nonsaponifiable compounds such as waxes and steryl esters have been shown to be major components of industrial pitch deposits [6]. Both fungal treatment and natural microbial activity increased the free fatty acid content of the extractives. The increase in free fatty acid content results from initial hydrolysis of esterified fatty acids to free fatty acids. The free fatty acid content of the Cartapip-treated chips is lower than that of naturally aged chips, indicating further metabolism and removal of these components by the fungus. In addition, analysis of the individual fatty acids showed that fungal treatment significantly decreased the content of the three fatty acids found in highest concentration in the untreated southern yellow pine chips oleic acid by 44%, linoleic acid by 64%, and palmitic acid by 45%. The results of Cartapip treatment such as pitch removal and maintence of chip brightness and improved paper machine runnability have been documented by use in mills. In a T M P mill using southern yellow pine, a trial was performed comparing a two week period using the Cartapip product on their wood chips to a two week period without product use, and the results are shown in Table 5 [24]. Reductions in the DCM extractive content of secondary refiner pulp caused expected reductions in alum, a pitch control chemical. Because Cartapip 97 is a colorless strain that outcompetes indigenous microorganisms, its use results in brighter chips. This effect was observed as a 36.9% reduction in bleach usage along with increased paper brightness of 0.9%. In

206 Table

R.L. Farrell et al. 5. Use of a depitching organism in a TMP mill

DCM extractive content of secondary refiner pulp Alum usage Bleach usage Brightness Tensile index Tear index Burst index

- 37.5% - 31.7% -- 36.9% + 0.9% + 5.4% + 3.4% + 3.3%

addition, strength properties were increased, probably due to the lower extractive content of the paper. Brandal and Lindheim have shown an inverse relationship between paper strength and pitch content [28]. A two m o n t h Cartapip 97 trial at a US T M P mill using southern yellow pine showed significant reductions in the D C M extractives of wood chips and an increase in burst index [24]. A one week trial was performed at a mill in Northwestern USA using a blend of 60% lodgepole pine and spruce and 40% fir and hemlock. Only the pine/spruce mixture was treated because this mixture caused the most serious pitch problems. Cartapip 97 treatment reduced the averaged D C M extractive content of the reclaim chips by 25% [25].

4 Enzymatic Pitch Control in the Papermaking Process 4.1 Fundamental Research and Theory of Lipase Application Initial studies to demonstrate the effect of lipases on pitch were conducted on Japanese red pine groundwood by the group of H a t a and colleagues at N i p p o n Paper [1-3]. Subsequent studies have also been carried out on softwood sulfite pulp and birch sulfate pulp [29-32].

4.1.1 Identificationof Compound CausingPitch Trouble A new method using an adsorption resin was established, instead of the previous solvent fractionation method, in order to fractionate red pine pitch and to determine what were the components that were sticky and causing pitch troubles [33]. Pitch compounds in red pine as well as deposited pitch were fractionated using the method and analyzed by gas chromatography. The changes in pitch compounds during the seasoning period and the contents of pitch in fresh wood were also investigated in great detail to understand the seasoning mechanism. These investigations produced the following results:


207

1. Pitch compounds could be fractionated into polar and nonpolar fractions [2]. 2. Fresh wood contained more nonpolar compounds, especially in winter. The main component consisted of triglycerides (TG) [2]. 3. 96% of the fatty acids that composed TG were oleic and linoleic acids [2]. 4. TG was rapidly decomposed and reduced during seasoning [1]. 5. Deposited pitch in the papermaking process always contained much TG [2]. Based on these results, TG was estimated to be the key to pitch troubles. In general, nonpolar compounds such as TG may easily adhere to hydrophobic surfaces, such as rolls, by Van der Waals forces and build to become pitch deposits. It was hypothesized by Hata and his coworkers that if TG in pulp slurry could be converted to less adhesive components, pitch deposits would decrease. The conversion of TG would enable the use of flesh wood with less probability of pitch trouble.

4.1.2 Application of Lipase Lipase specifically hydrolyzes TG, and thus was not expected to affect the environment or the paper quality. Three kinds of lipase, each produced by a different microorganism, were used in the original work by Hata and colleagues, and their properties are given in Table 6.

4.2 Effect of Lipase Treatment on Prevention of Pitch Deposition Resinuous materials extracted from red pine wood and groundwood pulp (GP) were treated with lipase, and their adhesiveness to the hydrophobic surface was determined [1, 23. As shown in Table 7, the pitch deposits increased when the ratio of nonpolar compounds to polar compounds increased. Thus, evidently the nonpolar compounds of the pitch materials had higher adhesiveness to hydrophobic material and seemed to play an important role in pitch deposition. TG was shown to be a key material in pitch deposition because the enzymatic hydrolysis of TG reduced pitch deposition significantly [2]. TG was hydrolyzed to glycerol and fatty acids with the lipase, and the resulting glycerol dissolved into water. Fatty acids existed in the form of aluminum salt in the presence of alum, and were dispersed into the pulp slurry and fixed on the surface of fibers. CHzOCORi ~HOCOR2

Lipase

CH2OH ~HOH

+

R1COOH R2COOH

~H2OCOR3

~H2OH

R3COOH

Triglycerides

Glycerol

Fatty acids

(Water-soluble)

(Fixed on fibers as A1 salt)

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4.3 Application to Papermaking Process Since the effect of lipase on reducing pitch deposits was confirmed, the technology was applied to the actual papermaking process [-2,3]. To select optimum conditions for the lipase treatment in mills, the following factors were investigated: the effects of enzyme concentration, reaction temperature, reaction time, and agitating mode on the hydrolysis of TG. The following results were obtained from the investigation: 1. It was necessary to have a strong mixing system to keep contact between enzyme and TG for the effective reaction by enzyme. 2. Under sufficient mixing conditions, lipase 5 000 U/kgGP (300 ppm Lipase B) could hydrolyze more than 80% of TG in the surface pitch (n-hexane extract from GP slurry) within two hours. 3. No effect of the lipase treatment on the brightness and strength of pulp was observed.

4.4 Mill Trial I Based on these results, the first long run mill trial was conducted using a large paper machine [2]. In this mill, the paper machine using red pine GP always had serious pitch problems because large amounts of red pine were used as raw materials for GP. Normally, 50% of unseasoned wood and 50% of wood seasoned for six months were consumed. Therefore, GP had a high content of pitch and a 30% TG content in the pitch. In order to reduce the pitch problem, it was in the past necessary to extend the seasoning period of wood to supply the mill, and the use of fine talc and dispersant was also increased. As an attempt to solve this problem, lipase was added to the groundwood pulping line just before the post refiner (Fig. 1). The operating conditions were as follows: Paper machine: Paper product: Pulp: Machine speed: Production rate: Enzyme: Reaction time:

Bel Baie II, wire width 5,080 mm Yellow Telephone Directory paper (YTD) (34 g/m2) Newsprint (46 g/m2) Red pine groundwood pulp (15-40%), de-inked pulp, softwood semi-bleached kraft pulp 830 m/rain 220-270 t/d Lipase A 75-125 ppm GP Lipase B 50~750 ppm GP 40-60 rain

An initial test was done in order to understand the proper dosage for a long run mill trial and to show the hydrolysis rate of TG by lipase in the actual papermaking process. Lipase A (125-ppm addition) on the first day of the mill trial reduced the content of TG by 74%.


209

For a one month lipase trial in the mill, the following parameters were compared between the usual operation and the lipase treatment operation in major products, such as newsprint and YTD: Content of the surface pitch and TG, first pass retention (FPR) of pulp and pitch, pitch deposits on the wall of the machine chest, amount of wet pitch deposits, number of defects in paper web, and dynamic friction coefficient (DFC) of paper. For 1.5-2.0% of oven-dried GP, the content of TG was 16-26% of the surface pitch [1]. Apparently the lipase hydrolyzed 70% of TG until reaching the mixing chest inlet. Furthermore, the accumulation of the pitch in the recycled white water (stock inlet, saveall) decreased to a lower level after the lipase treatment. As shown in Fig. 2, the first pass retention (FPR) of pulp did not change with lipase addition [2]. However, the F P R of the pitch increased from 5-9% to 12-19% in YTD, and from 9 14% to 13-24% in newsprint. As the lipase hydrolyzed TG, the pitch was dispersed into the pulp slurry and distributed onto the fibers without unevenness to the surface. Lipase also prevented the accumulation of pitch in the recycled white water system. As shown in Fig. 3, pitch deposit was observed as a black piling during the usual operation. However, pitch deposits could rarely be observed after a 1-month trial with the addition of lipase. Fig. 4 clearly shows that the lipase prevented the pitch deposition on the chest wall. In order to evaluate the pitch deposits in the wire and press sections, pitch deposit was collected from each section and measured every day. Results showed a dramatic decrease in the weight of pitch deposit with the lipase treatment compared to that of pitch in normal operation I-2]. The above results strongly proved that TG in the pitch was hydrolyzed and then converted to less sticky compounds. Long term data collected by spot detectors showed that the number of defects, holes, and spots larger than 1.5 mm was reduced from 61 to 19 as a daily average by the addition of lipase. When comparing long term data between the normal and the lipase operations, it was clear that the quality of products improved with lipase use. When a paper roll is printed on a web offset press, it is very important to prevent runnability problems such as wrinkling and uneven movement of the paper roll. This runnability performance is especially a concern in newsprint rolls. The dynamic friction of paper is thought to be related to these problems, and dynamic friction coefficient (DFC) is regarded as a quality control parameter at some Japanese paper companies. When D F C is low, the web tends to snake on the printing press. Therefore, when D F C of newsprint drops to a low level, white carbon (amorphous silica gel SiO2 x H20) is usually added to the paper furnish to increase DFC. As lipase treatment was incorporated into the production process, there was an increase in the newsprint D F C and a decrease in the amount of white carbon dosage. In order to reach a certain D F C level during newsprint production, about 2% of white carbon is added in the production process. However, by incorporating the lipase treatment of GP, there was a decrease in white carbon

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dosage to 1%. With the lipase treatment of GP, an increase of DFC was also confirmed [3].

4.5 Mill Trial H Another long term mill trial was conducted in March of 1990 [3]. Newsprint consisting of 15-35% GP was produced from red pine. This wood was normally seasoned in the mill yard for at least three months after cutting and collecting. This treatment reduced the TG content in wood and prevented severe pitch problems in the newsprint production process. However, there were still pitch deposits on the center roll of the paper machines, especially during winter, making it necessary to clean the center roll frequently. Lipase was added to make the operation smooth and to increase the unseasoned wood ratio in its furnish. The Machine conditions during the trial were as follows: Paper machine: Product: Pulp: Machine speed: Production rate: Enzyme:

Bel Baie II, wire width 3,800 mm Newsprint (46 g/m2) GP (15 35%), KP, TMP, de-inked pulp 1,000m/min 200t/d Lipase A 80-100 ppm GP Lipase C 400-500 ppm GP

The long term mill trial yielded the following results: 1. The frequency of cleaning was increased during winter (from January to April). However, the addition of lipase decreased the pitch deposits on the center roll, and the frequency of cleaning decreased to the level of summer (Table 8). 2. Increase of pitch deposit was not apparent on using 50% unseasoned wood. 3. Based on the two mill trials, lipase should be added to the pulp slurry before the addition of alum. Therefore, with the use of lipase, there was a decrease in TG content in fresh wood and reduction in the cost of seasoning and bleaching chemicals. Lipases for hydrolysis of pitch components have subsequently been applied to other wood species and to chemical pulps with successful results. Fischer and Messner applied the commercial lipase product Resinase A 2X (Novo Nordisk A/S) to unbleached softwood sulfite pulp [29-31]. In these studies, they drew the following conclusions concerning the activity of lipase on sulfite pulps: 1. Lipase is rapidly absorbed onto pulp fiber within a minute of addition [29]. 2. In a pilot mill scale treating 12 tons pulp day, with pulp at 4% consistency, 85 90% of the triglycerides were hydrolyzed, as determined by reduction of one triglyceride fraction from gas chromotography analysis [30], and the resin content of the pulp was reduced by 60% [31].


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Lipases from Candida cylindracea have been shown to be effective in hydrolyzing triglycerides in extractives of fresh birch and birch sulfate pulp [32]. The total amount of esterified compounds in fresh birch was decreased by 34%; 50% and 65% respectively of the esterified fatty acids and saturated fatty acids were also hydrolyzed. For treatment of birch sulfate pulp, the lipases of C. cylindracea hydrolyzed 30% of the esterified lipids as compared to 40% hydrolyzed by Resinase A (source organism Aspergillus from Novo Nordisk A/S). Esters of saturated fatty acids, alcohols and sterols were hydrolyzed by both lipases, though the C. cylindracea lipase was incapable of degrading esters of betulaprenols and triterpenoids, whereas the Aspergillus enzyme hydrolyzed them to some extent.

5 Conclusions The enzymatic pitch control method using lipase was the first successful example of the use of an enzyme as a solution to pitch problems and in the papermaking process. The fungal pitch control method, using the colorless isolant of Ophiostoma piliferum, was the first successful case of using a live organism as a solution to pitch problems and in the pulping process. Both technologies use biotechnology as their basis and have been successfully used in full scale industrial mills in various parts of the world.

6 References 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16.

Irie Y, Hata K (1990) Proceedings of 1990 Papermakers Conference, Atlanta Fujita Y, Hata K (1991) Japan Tappi J 45:55 Fujita Y, Hata K (1992) Tappi Journal 74:112 Blanchette RA, Farrell RL, Burnes TA, Wendler PA, Zimmerman W, Brush TS, Snyder RA (1992) Tappi Journal 75:102 Farrell RL, Hadar Y, Wendler PA, Zimmerman W (1992) European Patent 470 929 Brush TS, Farrell RL, Ho C (1994) Tappi Journal 77:155 Hoffmann GC, Brush TS, Farrell RL (1992) Naval Stores Review 102:10 Wendler PA, Brush TS, Iverson S, Krisa K, Zimmerman W, Farrell RL (1992) Kemia Kemi 19: 262 Backerby LH (1958) Pulp Paper 32:129 Hajny GJ (1966) Tappi Journal 49:97 Eslyn WE (1967) Tappi Journal 50:297 Hatton JV (1970) Tappi Journal 53:627 Somsen RA (1962) Tappi Journal 45:623 Dahm HP (1966) Norsk Skogindustri 1:10 Assarsson A, Croon I, Irisk E (1972) Svensk Papperstid 73:493 Messner K, Masek S, Srebotnik E, Techt G (1992) Proceedings of the 5th International Conference on Biotechnology in the Pulp and Paper Industry, Kyoto, Japan, 9

212 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33.

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Nilsson T, Asserson A (1969) US Patent 3 486 969 Iverson S, Blanchette RA, Farrell RL, Holzforschung, submitted. Lim CS, Cho NS (1990) Journal of Tappi-Korea 22:32 Farrell RL, Blanchette RA, Brush TS, Hadar Y, Iverson S, Krisa K, Wendler PA, Zimmerman W (1993) J Biotechnol 30:115 Fischer K, Akhtar M, Blanchette RA, Burnes TA, Messner K, Kirk TK (1994) Holzforschung 48:285 Poller VS, Schultze-Dewitz G (1982) Holzforschung 23:195 Zabel RA, Morrell JJ (1992) Wood Microbiology: Decay and Its Prevention. Academic Press, New York Chen T, Wang Z, Gao Y, Breuil C, Hatton JV (1994) Appita J 47:463 Haller T, Kile G Proceedings of the 1992 TAPPI Pulping Conference, p 1243 Wall MB, Stafford G, Noel Y, Fritz A, Iverson S, Farrell RL Proceedings of the 6th International Conference on Biotechnology in the Pulp and Paper Industry (in press) Farrell RL, Fritz A, Iverson S, Kile G, Brush TS (1994) 80th Annual Meeting, Technical Section of Canadian Pulp and Paper Association Preprints "B", Montreal Brandal J, Lindheim A (1966) Pulp Paper Mag Can T431 Fischer K, Messner K (1992) Proceedings of the 5th International Conference on Biotechnology in the Pulp and Paper Industry, p 169 Fischer K, Messner K (1992) Enzyme Microb Technol 14:470 Fischer K, Puchinger, Schloffer K, Kreiner W, Messner K (1993) J Biotechnol 27:341 Mustranta A, Fagernas L, Viikari L, Tappi Journal 78:140 Assarssopn A, Akerlund G (1966) Svenks Papperstidn, 69:291

Reduction of Organochlorine Compounds in Bleach Plant Effluents Pratima Bajpai and Pramod K. Bajpai Chemical Engineering Division, Thapar Corporate Research and Development Centre, Patiala - 147 001, India

List of Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Environmental Impact of Organochlorines . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Reducing the Generation of Organochlorine C o m p o u n d s . . . . . . . . . . . . . . . . . . 3.1 Enzymatic Prebleaching . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.1 Hemicellulase Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.2 Ligninolytic Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Fungal Prebleaching . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Treatment of Bleach Plant Effluents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4,1 Biological Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1.1 Using Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1.1.1 Aerobic Processes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1.1.2 Anaerobic Processes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1.2 Using Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Enzymatic Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

214 216 220 226 227 227 227 230 235 236 236 236 237 238 247 251 251

Organochlorines have been a matter of concern in the pulp and paper industry for the last two decades. These c o m p o u n d s are produced mainly by the reactions between residual lignin present in wood fibres and the chlorine used for bleaching. Some of the organochlorine c o m p o u n d s are found to be toxic, mutagenic, persistent, bioaccumulating and cause h a r m in biological systems. Earlier measures taken by the pulp a n d pape r industry to solve the chlorine problem have focussed on improving effluent treatment methods. Today, the emphasis of research and development work in this area has shifted more towards improving the processes. In the search to produce pulp with non-polluting chemicals, more efficient pulping m e t h o d s reducing the a m o u n t of residual lignin passing to the bleaching process and alternative bleaching methods, are being developed. There are also possibilities for treating effluent with microorganisms and enzymes to remove the dechlorinated organic material. Each option has inherent advantages and disadvantages with regard to capital costs, operating costs, ease of retrofit, fabrication a n d installation time. Impact on other mill unit operations is also considered in choosing the best options. M a n y factors have to be considered in choosing an effective and economical bleaching/treatment process that meets all the environmental guidelines. This review describes the environmental impact of organochlorines, use of enzymatic and biological bleaching for reducing the generation of organochlorine compounds, and treatment of bleach plant effluents by biological and enzymatic methods. Advantages and limitations of various biotechnological methods are discussed.

Advancesin BiochemicalEngineering/ Biotechnology,Vol. 57 ManagingEditor: T. Scheper 9 Springer-VerlagBerlinHeidelberg 1997

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P. Bajpai and P.K. Bajpai

List of Abbreviations ADI AOX BOD COD CTMP DDT ECF EOC1 EOX EPA HC HRT ISO LMS MBC MCC MW OKP PCB PCDD PCP Pow

ppb ppm ppt RBC t TCDD TCF TCP TEQ TMP TNT TOC1

Acceptable daily intake Adsorbable organic halogens Biological oxygen demand Chemical oxygen demand Chemi-thermomechanical pulp Dichlorodiphenyltrichloroethane Elemental chlorine free Extractable organic chlorine Extractable organic halogen Environmental protection agency High concentration (consistency) Hydraulic residence time International standards organization Laccase mediator substrate Modified batch cooking Modified continuous cooking Molecular weight Oxygen bleached kraft pulp Polychlorinated biphenyl Polychlorinated dibenzo dioxins Polychlorophenol Octanol/water partition coefficient Parts per billion (1/109) Parts per million (1/106) Parts per trillion (1/1012) Rotating biological contactor metric ton Tetrachlorinated dibenzodioxins Totally chlorine free Trichlorophenol Toxicity equivalent Thermomechanical pulp 2,4,6-Trinitrotoluene Total organochlorine

Bleachin9 stages C (CD)

Chlorination Treatment by mixing chlorine and chlorine dioxide simultaneously (C1 proportion is higher than D) D Chlorine dioxide treatment D1 & D2 First and second treatment, respectively with chlorine dioxide

Reductionof OrganochlorineCompoundsin Bleach Plant Effluents (DC) E E1 & E2 Eo Eov Ev F H L O P

Q X Z

215

Treatment by mixing chlorine dioxide and chlorine simultaneously (D proportion is higher than C1) Alkaline extraction First and second alkaline extraction respectively Alkaline extraction in presence of oxygen Alkaline extraction in presence of oxygen and hydrogen peroxide Alkaline extraction in presence of hydrogen peroxide Fungal treatment Hypochlorite treatment Laccase enzyme treatment Oxygen delignification/bleaching Hydrogen peroxide treatment Chelation of metal Xylanase enzyme treatment Ozone treatment

216


1 Introduction The bleaching of pulp became an issue of great concern during the 1980s primarily because of growing alarm over chlorinated organic compounds, referred to as adsorbable organic halogen (AOX), in bleach plant effluents. In particular, the detection of chlorinated dioxins and furans led to strong reactions. The paper industry recognized chlorine bleaching to be a potential problem. As the analytical instruments and methods became available, especially for chlorinated organic substances, studies were made on the chemical composition of bleach plant effluents. The new analytical methods and studies also created a basis for governmental regulations. The focus has been on setting upper limits for AOX/TOC1. At present, the strictest general emission limit value for AOX is 1 kg/t of pulp. This limit applies to sulphite pulp e.g. in Austria, Germany and Norway. This is because AOX emissions are easier to reduce with sulphite pulp than with softwood kraft pulp. However, in Sweden, kraft pulp mills have individual limits as low as 0.3 kg/t pulp. Some decided or planned regulations for AOX are shown in Table 1 [1]. The permitted levels are likely to decrease to about 1 kg/t in the next few years in many countries. In several countries or provinces, emission limits are decided mill by mill taking into account the local

Table 1. Regulations for discharge of chlorinated organic compounds measured as AOX (kg/t pulp) from bleaching of chemical pulps Country

1994

1995-2000

Australia Austria Belgium Canada - Alberta British Columbia Ontario Quebecb Finlandb Germanyb India~ Norwayb Japanc USAa Sweden

1.0a 0.75 1.5 1.5

0.5 1.0

2000-2005

0.29a/1.5 1.5 1.5 (HW)

1.5 1.0-2.0 1.0-2.0

0 0.8

1.0 2.0 1.0-2.0 1.5 1.2-1.5

0.156 0.3-1.0

0.3-0.5

"Limits for new mills bLower limits for hardwood pulps cGuidelines of Japan Pulp, Paper and Paperboard Association a Proposed regulation on "Cluster Rules" by EPA eProposed by Ministry of Environment and Forest, Government of India Based on data from Ref. [1]

Reduction of OrganochlorineCompoundsin Bleach Plant Effluents

217

factors. Thus, for individual mills stricter emission limits may apply than those given in the table. According to a decision in 1992 by PARCOM (Paris Convention for Prevention of Marine Pollution from Land Based Sources and Rivers) signed by twelve European countries, a general AOX emission limit would be 1 kg/t from 1995. The limit applies for all types of bleached chemical pulp and has been accepted by Belgium, Denmark, France, Germany, Great Britain, Ireland, Luxembourg, the Netherlands, Norway, Portugal, Spain and Sweden. In the USA, the Environmental Protection Agency (EPA) has proposed new emission limits for several categories of pulp, the so called cluster rules. The proposed emission limit for AOX for bleached kraft pulp is 0.156 kg/t. The new regulations will affect every existing and new facility in the pulp and paper industry and are expected to be promulgated soon. An AOX limit of 0.156 kg/t as proposed by the US EPA is very stringent when compared to current emissions even from Scandinavian mills (Fig. 1). Germany is also discussing legislation which will not only ban production of pulp using chlorine-containing chemicals but also consumption of other pulps than totally chlorine free (TCF). The Ministry of Environment and Forest, Government of India, has categorized the pulp and paper industry as one of the twenty most polluting industries and advised the paper industry to self-impose the AOX discharge limit of 2 kg/t of paper. Some of the state pollution control boards in India have already introduced the AOX limit of 2 kg/t of paper as a controlling parameter.

1.4

1.2

]

Scandinavian mills

II

Canadian rnitis

1.0

0.8

v

0.6

x

M

o 0.4

0.2

Cluster rulesl

II

I

R 3456

10 11 1 2 1 3 1 4 1 5

I I

161718192021

22,23:425

26 27 28 29 30 31

Mill

Fig. 1. AOX emissionsfrom Scandinavianand someCanadianbleachingkraft pulp millsin 1994

218


Despite some shortcomings, the kraft process is the most cost effective, versatile and efficient wood delignification method available. Kraft pulp is generally more difficult to bleach than sulfite pulp. With softwood pulp, it is harder to achieve a high brightness product than with hardwood. Chlorine is one of the popular bleaching chemicals, reacting with most of the lignin remaining in pulp, and degrading and dissolving the lignin as chlorinated organics. The acidic chlorination is usually followed by alkaline extraction to increase the solubility of chlorinated lignins. The substitution of organically bound chlorine by hydroxyl ions takes place under alkaline conditions, eliminating 60-90% of organically bound chlorine in the pulp I-2]. The stages following the chlorination and alkaline extraction are known as bleaching stages, and use more powerful oxidizing chemicals such as chlorine dioxide, hydrogen peroxide and hypochlorite. The main reaction is to oxidize the chromophoric structures in the pulp and to increase the pulp brightness. In the production of softwood kraft bleached pulp by the conventional CE1DIE2D2 sequence, approximately 75% of the dissolved material (COD and colour) and 95% of the organically bound chlorine were contained in the C and E1 prebleaching effluents. The major source was the E1 stage effluent. Lindberg 1-3] reported that the E1 effluent, which constituted about 12% of the total bleach effluent, accounted for 96% of the colour, 70% of the COD, and 50% of the BOD in whole bleach plant effluent. The amount of TOC1 produced during pulp bleaching varies with wood species, kappa no. of pulp, bleaching sequence and conditions employed. Typically, TOC1 in the effluent from a bleached softwood kraft pulp by a conventional sequence is 5-8 kg/t of pulp bleached, representing about 10%o of the total chlorine charged in the chlorination stage [4-6]. A physical-chemical classification of this chlorinated organic material, present in spent liquors from conventionally pulped and bleached softwood kraft pulp, is shown in Fig. 2 [7 9]. About 20% of the organically-bound chlorine found in the bleaching effluent corresponds to relatively low-molecular-mass (M r < 1000) material. In recent years, considerable research effort has been directed towards characterizing this fraction with respect to its individual chlorinated compounds [10-12], as this fraction is expected to contain those compounds which are potentially toxic to aquatic organisms because of their ability to penetrate cell membranes or their propensity to bioaccumulate in the fatty tissues of higher organisms. Some of the major components of this low-molecular-mass fraction were found to consist of relatively water-soluble substances such as chlorinated acetic acids or chlorinated acetones, which are easily broken down [8, 9] before or during biotreatment and are thus of minimal environmental significance. The fraction of AOX which is extractable by a non-polar organic solvent and is referred to as EOX (or extractable organically-bound halogen) accounts for about 1-3 %0 of the total organically-bound chlorine. This fraction contains relatively lipophilic (i.e. fatsoluble) neutral organic compounds, primarily of low molecular weight, and is therefore of greater environmental significance than the remaining 99 %0 or so of

Reduction of OrganochlorineCompoundsin BleachPlant Effluents

8o%

219

I

High M r material I

I

AOX 100%

Relatively hydrophilic (Water soluble) mainly non-aromatic Does not permeate cell walls < 10% chlorine by weight

Low M20m~

~ i R ~19% I elatively hydrophilic ncludes compounds which can easily be hydrolyzed or metabolized e.g. trichloroacetic acid)

Relatively lipophilic (fat soluble) Potentially toxic potentially bioaccumulable

Log Pow > 3 Highly lipophilic bioaccumulable (e.g. Dioxin - 44% chlorine by weight)

Fig. 2. The character of AOX in the effluentfrom conventionallypulped and bleached kraft pulp the AOX material. The EOX material can be further fractionated according to its octanol/water partition coefficient (Pow). The fraction having a partition coefficient greater than 1000 (i.e. log Pow > 3) makes up only 0.1% (or less) of the AOX and contains those compounds which are most readily bioaccumulable and considered to be potentially the most toxic and persistent. A component of particular concern in this fraction is the polychlorinated aromatic material that has a relatively high level of chlorine substitution, typically three or more chlorine atoms per aromatic ring. The major bleaching parameters such as incoming kappa number, C12 dosage, and chlorination and extraction pH and temperature have a significant effect on the effluent BOD, COD, color loadings and the formation of chlorinated compounds. It has been reported that the pollution load and amount of chlorinated material produced in the chlorination and extraction stage are a function of the amount of chlorine applied to the pulp, which is determined by residual lignin in the pulp. The amount of TOC1 in the C stage spent liquor was strongly dependent on the C12 dosage and lignin content in the pulp [13-15]. The lipophilic chlorinated organic substances were found to increase with an increase in the pulp kappa number and C12 dosage [15]. An increase in the chlorine dosage results in increasing BOD and COD in the chlorination and extraction effluents [16, 17]. Voss et al. [4, 17] have reported that significant reductions in the loadings of toxicity and chlorinated phenolics of combined C and E effluents can be achieved by a higher final pH of chlorination. The

220


formation of TOC1 and chlorinated phenolics increased with increasing temperature and decreasing pH of chlorination. Alfthan et al. [16] reported that increase in temperature of the chlorination resulted in increase in pollution loadings in chlorination and extraction effluents. Crawford et al. [18] reported that the increase in the end pH and temperature of chlorination and chlorine charge increases the formation of chloroform, potential mutagen and carcinogen in the chlorination liquor. The hypochlorite stage is the major source of chloroform formation. In the C stage, chloroform production is normally much less, but it increases with increasing pH, temperature and kappa factor. The higher kappa number also results in an increase in chloroform concentration in the E stage. Chan and McDonald [19] reported that the use of high C102 substitution in the C stage reduced the chloroform concentration. While C102 substitution reduces the effluent TOC1, color and chloroform, it had little impact on the production of chlorinated dioxins [20]. The major source of chlorinated dioxins was found to be the C stage followed by the E stage. The formation of dioxins is mainly dependent on the brown-stock kappa number, kappa factor, mixing condition in the chlorination, carry-over and wash water quality. This article presents a state-of-the-art review of the literature relating to the environmental impact of the chlorinated organic compounds and the measures which are used for their reduction. Emphasis is laid upon biotechnical methods for reducing the discharge of chlorinated organics.

2 Environmental Impact of Chlorinated Organic Compounds About 300 different chlorinated organic compounds in bleached pulp mill effluents have been identified to date. About 200 of these are compounds which include chlorinated resin acids, chlorinated phenolics and dioxins [20-22]. The main compounds of the general type are listed in Table 2. These compounds, classified as acidic, phenolic and neutral compounds, are at least partly responsible for the oxygen demand (BOD and COD), effluent color, toxicity, mutagenicity and carcinogenicity [23-29]. Various chlorinated phenolic, acidic

Table2. Chlorinated organic compounds in bleached pulp mill effluents Type

Number of varieties

Amounts(g/t pulp)

Chlorinated phenolics Chlorinated aldehydes,ketones and lactones Chlorinated acids Chlorinated hydrocarbons Chlorinated ethers High molecular mass

40 45 40 45 20

Up to 100 500 Up to 500

Based on data from Refs. [2~22]

Up to 4 kg C1

Reduction of Organochlorine Compounds in Bleach Plant Effluents

221

and neutral compounds and chlorinated dioxins have been found to be bioaccumulative [20, 30]. Up to 2000 ppm organic chlorine has been detected in the fat of fish from waters receiving bleaching effluent [31]. Untreated pulp and paper mill effluents can be acutely toxic to fish at concentrations as low as 2% by volume [26]. Chlorinated phenolics, resin and fatty acids are the principal contributors to effluent acute toxicity [23, 32-34]. Table 3 lists 12 polychlorinated phenolics selected by the US EPA for possible regulation [35]. The toxicity of a chlorinated compound increases with increasing number of chlorine atoms on the organic compounds. Polychlorinated dibenzodioxins (PCDDs) and 2,3,7,8-tetrachlorinated dibenzodioxin (TCDD) are toxic. It is generally believed that they are of precisely the right size to fit as the key in the lock in some vital molecules in living cells and to be firmly attached to this site by the chlorine atoms at both ends, thus preventing normal functioning of the cells. T C D D seems to fit perfectly. If, however, more hydrogen atoms in the benzene rings are substituted with chlorine, the toxicity is reduced, e.g. if all eight available hydrogen atoms are replaced with chlorine atoms, the toxicity drops to one per thousandth of that of TCDD. The key does not fit in the lock any more. Dioxins have received extensive media attention. 2,3,7,8-TCDD is extremely toxic and bioaccumulative. The toxicity of individual members of the abovementioned family of 210 isomers (varieties) of polychlorinated dioxins and furans differ substantially. The most toxic of them is at least 100000 times as toxic as the least toxic, so that it is meaningless to judge the quality of an effluent or pulp by quoting the total dioxin concentration. Dioxins are frequently reported as toxicity equivalents (TEQ), which is the sum of total chlorinated dioxins and furans corrected for the toxicity of each relative to 2,3,7,8-TCDD. For practical purposes, in the pulp industry, the T E Q generally corresponds to somewhat more than the concentration of 2,3,7,8-TCDD [36, 37]. Table 3. Polychlorinatedphenoliccompoundsproposed for regulation by the US EPA Polychlorinatedphenols

Minimum level,ppb (gg/1)"

Pentachlorophenol 2,3,4,6-Tetrachlorophenol 2,4,5-Trichlorophenol 2,4,6-Trichlorophenol 3,4,5-Trichloroguaiacol 3,4,6-Trichloroguaiacol 4,5,6-Trichloroguaiacol 3,4,5-Trichlorosyringol 3,4,5,6-Tetrachlorocatechol 3,4,6-Trichlorocatechol 3,4,5-Trichlorocatechol 3,4,5,6-Tetrachloroguaiacol

5.0 2.5 2.5 2.5 2.5 2.5 2.5 2.5 5.0 5.0 5.0 5.0

"The minimum level is defined as the concentration at which the analytical system gives recognizablemass spectra (correctedfor background) and acceptable calibration points, using EPA method 1653 Based on data from Ref. [35]

222


Greenpeace claims that there is no safe level of dioxin [36]. On the other hand, over a billion dollars worth of research has failed to prove any serious health damage to humans due to dioxin exposure [36]. It is fairly well accepted by scientists in the field that 10 picograms of dioxin per kg body weight per day is an acceptable daily intake (ADI) for a one in a million risk of mortality, over and above the other risks of living. However, a concern about dioxins in effluents is that they bioaccumulate in fish. This means that they concentrate as they move up the food chain through predator fish to humans. In Canada, the Federal Government recommends that fish with over 20 ppt dioxins in the flesh should not be eaten. Occasional fish caught downstream of mills have concentrations of up to about 100 ppt, but these incidents can be expected to become things of the past if mills adopt the dioxin control measures already announced. To help put this all into perspective, it is worth noting that collection of the ADI includes several safety factors. Extensive studies of 1200 US soldiers who worked with dioxin-contaminated herbicides in Vietnam failed to demonstrate any health effect, but there was an out-of-court settlement of almost $200 million. One might say that legal dangers have been demonstrated to be associated with dioxin but no serious biochemical danger to humans. Recently, the EPA released its 2000-page draft reassessment of the environmental and health effects of dioxins [38]. The report reaffirms the EPA's 1985 conclusion that dioxins and related chemicals are a probable cause of cancer in humans. It also presents new evidence that dioxins, even in trace amounts, may cause a wide range of other adverse human health effects including disruption of regulatory hormones, reproductive and immune system disorders and abnormal fetal development. The EPA defines dioxins and related compounds as tetra- to octa-chlorodibenzo-dioxins and furans with chlorine atoms in at least the 2,3,7 and 8 positions, as well as coplanar polychlorinated biphenyls (PCBs). The EPA expresses the mass of these compounds in terms of their toxicity compared with the most toxic dioxin congener 2,3,7,8-tetrachlorodibenzo p-dioxin, denoted as 2,3,7,8-TCDD toxicity equivalents (TEQs). In other words, if the mass of a particular dioxin congener is 100g and it has one tenth the toxicity of 2,3,7,8-TCDD, its mass in TEQs is 10 g. The reassessment emphasizes dioxin's effects on fetal development, because these effects have been seen in the offspring of laboratory animals exposed to very low concentration, and are now being observed in children accidentally exposed to dioxin-like compounds in the womb [38]. For example, the offspring of women who in 1979 ate rice oil contaminated with furans and PCBs in the Yu-Cheng Province of Taiwan exhibit changes in skin pigmentation and hair growth and are showing signs of abnormal sexual development, says Aronold J. Schecter, Professor of Preventive Medicine at the State University of New York, Clinical Campus, Binghamton. Dioxins and dioxin-like compounds apparently target many sets of genes which encode a variety of proteins, including hormones, enzymes and growth factors. As a consequence, the cell produces an inappropriate amount of protein. Depending upon the dose, timing and age of


223

the individual, this cell disruption can lead to diverse biological outcomes, effects on the reproductive system of the developing fetus, effects on the brain, disruption of the immune system and cancer. Dioxin's effect has been seen in all species studied, although they occur at different doses. Humans lie somewhere in the middle of the sensitivity range, neither extremely responsive nor extremely resistant, according to the EPA report. Unlike natural hormones, like the estrogen estradiol, dioxins can be deactivated by binding to sex-hormonebinding globulin and are not easily metabolized. Dioxins disrupt multiple endocrine systems, at times behaving as antiandrogens, at others like antiestrogens or even estrogens. The half-life of the most studied dioxin 2,3,7,8-TCDD is 10 years in humans [39]. Low molecular mass chlorinated neutral compounds are major contributors to mutagenicity [31]. The dominant chlorophenolics in C stage effluent are chlorocatechols, whereas chloroguaiacols are the major species in E stage effluent. In hardwood bleaching effluent, additional chlorinated compounds such as chlorinated vanillins, syringealdehydes and syringals were found. But the concentrations of chlorinated phenolics in the hardwood bleaching effluents are generally lower than those in softwood bleaching effluent [40]. At least 14 different chlorinated phenolics in conventional C and E1 bleaching effluents were detected [41]. Sublethal effects of pulp and paper mill effluents are varied. The threshold concentration for sublethal effects appears to be near 1/10 of the 96 h LCso concentration [33]. Davis [42] reported the lethal effect at 5 % of the 96 h LCso concentration of the spent bleaching liquor. The Swedish pulp and paper industry sponsored studies to determine the environmental effect of spent bleach liquors and to develop environmentally compatible bleaching processes [15, 43, 44]. It was concluded that the effects of bleach plant effluents actually observed in receiving waters were few in number and limited to receiving waters with poor water exchange or to areas close to the outlet. Laboratory tests suggested that all sublethal effects on fish of effluents from the conventional bleaching of softwood kraft pulp disappear after a total dilution of 70-1000 times [15]. Based on the results of chemical and biological comparisons of the effluents, various alternative bleaching sequences and treatments were ranked as shown in Table 4. The Swedish project SSVL-85 was designed to assess long-term large-scale effects of bleach plant effluents and used model ecosystems to characterize bleach plant effluents [15, 44]. Two concentration levels were used corresponding to dilutions of 400 and 2000 times for a total effluent volume of 50 m3/t pulp. Five different effluents were tested for fish and invertebrate toxicity, reproductive disturbances, physiological changes and disease, parasitic infestation, bioaccumulation of chemicals and genotoxic effects (mutagenicity and/or carcinogenicity). The ranked sequences did not change significantly from that in Table 4. However, biological effects were found at dilutions which were previously thought to render the effluents harmless. The effluent from a conventional (C95D0s)E1DE2D bleach plant showed strong biological effects after 166-fold dilutions, with low survival of fish, decreased

224


Table 4. Bleaching processes in order of decreasing environmental effect

S.no. a

Bleaching process

Comments

1 2

(C90DIo) E1HD1E2D2 (CgoDIo) E1HD1E2D2

3 4

(C9oDlo)E1HD1E2D2 + UfE1 0(C85D15) E1D1E2Dz

5

(C9oDao)EaHD1E2D2 + UfE1 + Z

6

0(C85Dxs)E1D1E2D2

Traditional technique Improved washing compared to traditional technique Ultrafiltration of Ex effluent Oxygen bleaching to kappa no. 20 and washing to 1~12 kg COD/t Ozone treatment of permeate and C-stage effluent Oxygen bleaching to kappa no. 20 and washing to 4-6 kg COD/t Total effluent through aerated lagoon Ion exchange of total effluent High substitution of chlorine dioxide Oxygen bleaching to kappa no. 20 Oxygen bleaching to kappa no. 17

7 8 9 10 11

(C90Dlo)E1HDaE2D2 + aerated lagoon (CgoDlo) + Ion exchange (D85C15)EID1E2D 2 0(C85D15)E1D1E2D2 + aerated lagoon 0(CssD15)E1D1EzD2

Softwood Kraft Pulp UfE~ = Ultrafiltration of E1 effluent Worst 1 Best 11 Based on data from Ref. 1-15] a

invertebrate density and parasitic infestation of stationary fish species. Even at dilutions over 5000 times, all these effects could be found, although to a substantially decreased degree. In 1983, the Swedish Government sponsored Project Environment/Cellulose, a three year study of the Gulf of Bothnia, that portion of the Baltic sea which lies between Sweden and Finland [21]. The study focussed on the area around the Norrsundet pulp mill, which discharges untreated effluent into the Gulf and which was regarded as representative of Swedish pulp mills. Near the discharge point, fish biomass was low, species composition was changed, reproduction was reduced and physiological disturbances were seen. The effluent also affected the diversity, biomass and distribution of invertebrates and plants in the region [21]. The toxic effects were more pronounced within 4-5 km of the outlet of the mill, but biological effects were observed in the fish caught 8 to 10 km from the outlet, where the dilution was estimated to be 5000 times [45]. Bioaccumulating compounds in the fish declined in concentration from the discharge point to about 5 km out from the shore [21]. The concentration of AOX in the water, however, showed a larger area of influence. High concentrations of solvent-extractable organically bound chlorine (EOC1) were found in the sediment 15 km out from the Norrsundet mill and 29 km out from another mill 75 km north of Norrsundet. Large areas of the Gulf of Bothnia were found to have higher levels of EOC1 in the sediment. Compared with areas further from industry, EOC1 levels in sediment 20 to 50 km off the coast of bleached kraft pulp mills were 10 times the background levels [21]. Between 1979 and 1984, the Norwegian Centre for Industrial Research studied the content of EOC1 in fish in


225

relation to the effluent of a CEHD bleached sulphite pulp mill [46]. While the mill was in operation, the concentration of EOC1 in fish near the outlet was up to 30 times that in fish from a nearby reference area. EOC1 levels in fish did not drop to the background concentration until 3.5 years after the mill had shut down. In 1987, Swedish scientists stated that in spite of extensive investigations, a clear-cut relationship between release of chlorinated organic material and bioecological effects in the receiving waters has not yet been established. Thus, the TOCI regulations give a general feeling that the release of chlorinated organic material from bleach plant is critical to the environment [47]. However, concern has been voiced that the effects found in Sweden are specific to that area, because the Baltic sea may be a unique ecosystem. It contains a relatively small number of species and the media is brackish and has a long residence time, of the order of several decades [483. Extensive long-term environmental impact studies have also been carried out in the United States. Since virtually all the mills in the USA have secondary biological treatment, treated bleach plant effluents were tested. Test work, using experimental stream channels, has been conducted by National Council of the Paper Industry for Air and Stream Improvement (NCASI) as part of an aquatic biology investigation going on since the 1970s. In the latest report, the results of year-long exposure to treated bleached plant effluent of 20:1 dilution were reported [49]. Productivity of the rainbow trout population and aquatic flora was increased compared to that in the control stream, while productivity of the benthic macroinvertebrate population was unchanged. There was no effect on the diversity index or histopathology of 20 different fish-tissue types. These results are clearly contradictory to the general conclusion of the Swedish investigations. The reason for the differences is not apparent; it may be the effect of effluent treatment or the sensitivity of the Baltic eco-system. Another category of compounds which has aroused environmental concern comprises the high-molecular-mass chlorinated compounds (Mr > 1000). In bleaching softwood kraft pulp, this fraction of chlorinated compounds accounts for 50% of dissolved organic material [50] and 70% of TOC1 [20]. Although these compounds contribute little to BOD5 and acute toxicity due to their inability to pass the bacterial cell membrane, they are the major contributors to effluent colour, COD and chronic toxicity. Eriksson and Kolar [51] and Eriksson et al. [52] found that the high-molecular-mass material was not as stable as previously thought. High-molecular-mass chlorolignins from prebleaching stages were chemically unstable under conditions that may prevail in receiving waters. The material was slowly decomposed to various chlorinated catechols and guaiacols which were methylated in cases where a complete mixture of bacteria or the white rot fungus alone was used, a condition appearing to be common in nature [15, 53]. The low-molecular-mass phenolics and their methylated counterparts (more lipophilic) may cause toxicity and bioaccumulation in fish. Chlorate is yet another pollutant recently arousing environmental concern. The formation of chlorate is largely dependent on the amount of C102 used in

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the bleaching. It is a well-known herbicide and biocide, especially for brown algae [54]. A conventional bleaching sequence produces about 3-6 kg/t of pulp bleached depending on the level of C102 substitution.

3 Reducing the Generation of Organochlorine Compounds The emission of chlorinated organic compounds from bleach plants can be reduced by modifying the pulping and bleaching processes using one or more of the following strategies: (1) removing more lignin before starting the chlorination, i.e., reducing the kappa number of unbleached pulp, (2) modifying the conventional bleaching process, (3) eliminating all chlorine compounds in bleaching, or (4) recovering bleach plant effluents. These methods may be physicochemical or biochemical in nature or a combination thereof. Lignin from the pulp before chlorination can be removed by extended delignification of the brown stock pulp using modified continuous cooking (MCC) [55-58] or modified batch cooking (MBC) [59-61]. The lignin content can also be removed by oxygen delignification [20, 58, 6~69]. Ozone is also capable of delignifying an oxygen-treated pulp to a very low kappa number of 5-6 [70, 71]. A lower lignin content in the pulp requires lower chlorine usage resulting in reduced generation of chlorinated organics. The conventional bleaching process can be modified by (a) reducing the kappa factor or active chlorine multiple, i.e., the total active chlorine charge derived by the kappa number of the pulp entering the chlorination stage, (b) slow and multiple addition of chlorine [72], (c) chlorine dioxide substitution in the chlorination stage [4, 13, 17, 72-77], (d) oxidative extraction using oxygen [78-80] and/or hydrogen peroxide [81-91], or (e) converting to ECF bleaching sequence [89, 92]. Further decrease in AOX can be achieved by a combination of extended delignification in the digestor, oxygen delignification, and high chlorine dioxide substitution in the chlorination stage [72]. The trend now is to adopt a totally chlorine-free (TCF) bleaching sequence in some European countries or ECF bleaching in North America, which can involve the use of oxygen, enzyme, hydrosulfite, peroxide, peracetic acid, thiourea dioxide, chelating agents [72, 93], ozone [70-72, 94-97] etc. Once chlorine-free bleaching is introduced, the bleaching effluent can be taken to the black liquor recovery section [98]. A non-polluting bleach plant has been reported where counter-current washing decolorization and reuse of the extraction stage effluent as wash water in the chlorination stage is practiced [99, 100]. Ultrafiltration and reverse osmosis membrane techniques can also be applied for complete recycling of the effluent [101, 102]. The best-known closed cycle effluent-free bleaching process was reported in the mid-1960s by Rapson and Reeve [103-105].

Reduction of Organochlorine Compoundsin Bleach Plant Effluents

227

Apart from the above methods, several biochemical and biological methods can be used to reduce the generation of organochlorine compounds, as discussed in the following sections.

3.1 Enzymatic Prebleaching Pretreatment of pulp with enzymes prior to bleaching helps in substantially reducing or eliminating the use of chlorine and chlorine compounds. Hemicellulase enzymes and ligninolytic enzymes have been investigated. Hemicellulase enzymes have no immediate effect on the kappa number and lignin removal of the treated pulp. However, the consumption of the bleaching chemicals is reduced and/or brightness ceiling is increased. Ligninolytic enzymes selectively remove the lignin from the pulp fibre, hence are more useful. The role of different enzymes on pulp components has been reviewed by Eriksson [106].

3. I. 1 Hemicellulase Enzymes These enzymes are used commercially in pulp bleaching. The main enzyme needed to enhance delignification of kraft pulp is reported to be endo-~xylanase, but enrichment of xylanase with other hemicellulolytic enzymes has been shown to improve the effect of enzymatic treatment [107-110]. Xylanases act mainly on the relocated, reprecipitated xylan on the surface of the pulp fibres. Enzymatic hydrolysis of this specific type of xylan renders the structure of the fibre more permeable, allowing lignin and lignin carbohydrates to diffuse more easily into the bleaching liquor. An alternative explanation is that xylanases attack the bonds that exist between xylan and lignins, releasing lignins which can then diffuse more easily into the bleaching liquor [109, 111, 112]. Xylanase treatment has been shown to reduce the requirement of chlorine for bleaching, while still achieving high brightness and good pulp properties [113-115]. Results from laboratory study and mill trials show about 3 5 4 1 % reduction in active chlorine at the chlorination stage for hardwoods and 10-20% for softwoods, whereas the saving in total active chlorine is found to be 20-25% for hardwoods and 10-15% for softwoods [116-123]. Xylanase treatment represents a successful new technology for reducing chlorine use. More details on xylanase bleaching are available in a recent review by Bajpai and Bajpai [124] and in another chapter in this volume written by Viikari et al. [125].

3.1.2 Ligninolytic enzymes These enzymes, unlike xylanases, attack lignin directly, and hence are more effective. White-rot fungi are the main producers of ligninolytic enzymes. These

228


fungi secrete a number of oxidative enzymes and some hitherto unknown substances (mediators) into their environment, together effecting a slow but continuous degradation. The most important lignin-degrading enzymes are lignin peroxidases, manganese peroxidases and laccases. The action of both lignin peroxidases and manganese peroxidases needs Mn z +, which is oxidized to Mn 3 +. Mn a + is the real oxidizing agent, attacking the lignin molecule. Laccase uses molecular oxygen as a cosubstrate. In the absence of the living organism, the various peroxidases and laccases perform only negligible kappa number reduction [126, 127]. Egan [128] has reported kappa number reductions of 24 and 26% after treatment with ligninase 118 from P. chrysosporium followed by alkaline reduction. Viikari et aI. [129] were unable to demonstrate any bleaching effect on pine kraft pulp with ligninases and oxidase from the white-rot fungus Phlebia radiata or purified ligninase from P. chrysosporium. When the ligninases were applied after treatment with hemicellulases, there were slight reductions in kappa number. Arbeloa et al. used lignin peroxidase enzyme from P. chrysosporium to improve the bleachability of hardwood and softwood kraft pulps [130]. Enzyme treatment prior to chemical bleaching increased brightness and decreased lignin content in the pulp. The final brightness of pulp was found to be higher by about 0.84).9 points than that of the control (Table 5). Although much of the research has focussed on lignin peroxidases, these enzymes are not necessarily involved in lignin degradation and may not be secreted by all lignin-degrading fungi. C. versicolor produces laccases as well as lignin and manganese-dependent peroxidases [131-134]. However, Archibald [135] recently found that lignin peroxidases secreted by C. versicolor did not appear to play an important role in lignin degradation. Dichomitus squalens [136] and Rigidoporus lignosus [137] produce both laccase and a manganesedependent peroxidase but do not produce lignin peroxidase. It is now known that although lignin peroxidases and laccases play an important role in degrading the lignin in vivo, in vitro the oxidation reactions catalyzed by the enzyme result in further polymerization of the lignin [138]. Table 5. Effects of lignin peroxidase on kappa number and bright-

ness in softwood and hardwood kraft pulp After treatment

After bleaching

Kappa number

Brightness (%)

Brightness (%)

29.7 26.9

29.1 24.1

88.8 90.7

14.6 11.8

31.0 39.0

88.6 89.4

Softwood Control Treated

Hardwood Control Treated

Conditions of enzyme treatment: pH 3, temperature 30 ~ alcohal 2 mM/l, H202 addition at 100 ppm/h for 3 h Based on data from Ref. [130]

veratryl


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Recently, Hammel and Moen [139] reported depolymerization of a synthetic lignin by a lignin peroxidase in the presence of H202 and veratryl alcohol, but this effect has not been demonstrated with lignin in wood or pulp. It is likely that fungi possess enzyme systems that prevent polymerization [138, 140]. These results imply that single enzymes are not able to mimic the complete biological system. Small improvements can be achieved by the addition of low-molecular-mass aromatic compounds like veratryl alcohol or other substances such as ABTS [2,2'-azinobis(3-ethylbenzthiazoline-6-sulphonate)] and Remazol blue [126, 127]. In consequence, most scientific and industrial groups have meanwhile withdrawn from research programs focussing on direct lignin degradation. Lignozyme GmbH (Germany) continued work with enzymes plus chemical mediators which create a redox system throughout the pulp treatment period [-141-143]. Their idea was to find a system which is a good mimic of the natural situation. Starting in 1987 with the enzyme mediator concept, Lignozyme has very recently improved the performance of the mediator system for the laccase of Coriolus versicolor by changing and further fine-tuning the chemical nature of the component [144-146]. The treatment of pulp with laccase alone does not result in any degradation of lignin but merely produces in a structural change or repolymerization, whereas the laccase mediator system causes a significant kappa number reduction at reasonable treatment times even if the enzyme mediator system is applied in several consecutive treatment steps to the same pulp. In contrast to commonly used pulp-bleaching chemicals (oxidizing and reducing), no passivation can be observed in enzymatic delignification, i.e., efficiency does not drop significantly during consecutive treatment steps. Laccase enzyme contains four atoms of copper per molecule and requires oxygen as a cosubstrate for the oxidation reaction which has to be provided. According to the present understanding, the laccase, while oxidizing the chemical mediator, is generating a strongly oxidizing co-mediator which is the real bleaching agent. Table 6 illustrates the conceptual difference between the indirect and the direct enzymatic lignin attack. The general technical conditions for enzymatic bleaching with the laccase mediator system are: temperature 40-65~ pH 4-7, consistency 1-20%, pressure 1-14 bar, duration 1~4 h. The system provides a broad flexibility with respect to the pulp substrate, the technical requirements for application, and the final quality of the pulp. The principal applicability has been demonstrated for softwood and hardwood pulps as well as for annual plant

Table 6. Differencesbetweenxylanaseand laccase/mediatortreatment Xylanase

Laccase/mediator

No or verypoor kappa reduction Moderate bleachingeffect Saving of bleachingchemicals

Very good kappa reduction Good bleachingeffect Saving of bleachingchemicals TCF pulp productionpossible

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fibres. A repeated enzymatic treatment is possible and results in a 50-70% kappa number reduction per treatment step. Laccase mediator system (LMS) is compatible with all other bleaching sequences. The performance of the LMS (1-hydroxybenzotriazole) has been proven in several pilot trials and is now ready for large-scale application. Recently, Kondo et al. [147] examined the bleaching of kraft pulp with Mn peroxidase (MnP) enzyme, which was isolated from cultures of P. sordid YK624. Pulp brightness was increased by about 15 points and kappa number decreased by about 8 points when unbleached kraft pulp was treated with MnP in the presence of MnSO4, Tween 80 and sodium malonate with addition of H 2 O 2 at a rate of 3 ml/h at 45 ~ for 12 h. To establish an absolutely chlorinefree bleaching process, oxygen-bleached kraft pulp (OKP) was treated with a four-stage biobleaching process consisting of sequential MnP treatment, alkaline extraction, MnP treatment and hydrogen peroxide treatment. Fully bleached kraft pulp (brightness 91%, yield 97%) could be obtained from OKP by combination of enzyme treatment and hydrogen peroxide bleaching. MnP from Trametes versicolor was found to delignify softwood kraft pulps over a wide range of initial lignin contents [148]. Demethylation of residual lignin in the pulp also occurred. Subsequent treatment of pulp with TCF chemicals resulted in brighter pulp than predicted from the initial kappa number. The MnP stage required small doses of hydrogen supplied by the action of glucose oxidase on glucose. Mn ions already present in the pulp were chelated by malonate or gluconate during the catalytic cycle. Much of the bleaching effect was evident after 4 h of treatment with 1 U/ml of MnP, although bleaching was improved further after 24 h under the conditions used. Laccase, especially in the presence of the mediator, ABTS, produced a similar effect to MnP when combined with subsequent TCF bleaching. MnP generates oquinones by demethylation of residual lignin, and these structures are susceptible to oxidative ring opening during TCF bleaching, resulting in hydrophilic soluble lignin. Ducka and Pekarovicova [149] used crude ligninases from P. chrysosporium for bleaching of softwood kraft pulp. The pulp, after oxygen bleaching with kappa number 19.7 and 36.5% MgO brightness, was pretreated with ligninases (L) or xylanases (X) and bleached in QEopDP sequence. The brightness of pulp bleached in this sequence was 87.8%, which is about 3.2 points higher than the control and is approximately the same as that obtained with the use of commercial xylanase.

3.2 Fungal Prebleaching Pretreatment with fungi has been shown to replace up to 70% of the chemicals needed to bleach kraft pulp [150]. The usual specificity of biological reactions and their mild reaction conditions make biological delignification an interesting alternative to bleaching with chemicals such as pressurized oxygen or ozone.


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Only a few white-rot fungi have been tested for their ability to delignify kraft pulps. Phanerochaete chrysosporium reduced kappa number by about 33% in hardwood kraft pulp during a 10 day incubation period, whereas Coriolus versicolor reduced kappa number by about 20 and 33% in a 5 day period [151-153]. Table 7 shows the performance of various white rot fungi used in treating hardwood kraft pulps. The largest bleaching effect was noted with white-rot species IZU-154 [154]. Treatment of hardwood pulps with alkaline extraction following fungal bleaching did not significantly improve brightness. Softwood kraft pulps were found to be more resistant to attack by P. chrysosporium and C. versicolor, possibly because both P. chrysosporium and C. versicolor tend to attack hardwoods more often than softwoods in nature [155-157]. Softwood lignin has a different character from hardwood lignin and is susceptible to blocking reactions that restrict delignification [158, 159]. On delignification of softwood pulps by P. chrysosporium, kappa number reductions of 50-70% were achieved [160, 161]. Kirk and Yang [160] reported a considerable loss in cellulose content during fungal bleaching, which was attributed to cellulase production in the strain being used. A strain of P. chrysosporium without cellulase activity has since been developed [162].

Table 7, Change in brightness of hardwood kraft pulp after exposure to various white-rot fungi Brightness (%ISO) Fungus

Increase

Final

Incubation Period (d)

Phanerochaete Chrysosporium

11 0 2.2

39 37.7 35.7

13 14 5

2.0

35.5

5

P. chrysosporium (cellulase-free mutant # 431) P. chrysosporium (cellulase-free mutant # 432) Coriolus versicolor

IZU-154 Coriolus hirsutus Phellinus pini Pleurotus eryngii Pleurotus sajor-caju Lentinus edodes Aureobasidium puUulans

1.9 8 25 7.1-17.8 14.5 13.6 13.4a 6 35 23.7 12 2.3 1.9 2.3 2.4 2.5

"Fungus was immobilized Based on data from Refs. [152 154,163,195,199]

35.4 50-67 38.5 50.3 48.0 50.4 50.2 34.0 63 52 40 35.8 34.4 35.8 35.9 36.0

5 10 5 5 5 5 15 11 5 11 5 5 5 5 5

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C. versicolor was found to be ineffective in brightening softwood kraft pulps despite reductions in kappa number of 8% [163] and 47% [154, 157] over 14 days. However Reid et al. [157] reported that when the pulp was subsequently subjected to alkaline extraction, brightness was increased by 15 points and kappa number was further reduced. The softwood kraft pulp exhibited a more pronounced darkening than the hardwood kraft pulp during the first 6 days of treatment, and a longer time lag occurred before kappa number began to decrease. Direct brightening of softwood kraft pulp was noted when the C. versicolor was immobilized on polyurethane foam, possibly because of a higher fungus/pulp ratio [157]. In Japan, a 5-day fungal (F) treatment of hardwood kraft pulp with IZU-154 replaced a CE1DEzD sequence with an FCED sequence, yielding respective brightness of 88.8 and 88.1% ISO (84.2 and 85.3% ISO after aging) [154]. The resulting chlorine saving was 72%, despite a kappa number reduction of less than 60% in the fungal stage. In another study, a 5-day fungal stage with C. versicolor essentially replaced the chlorination stage in a conventional bleaching sequence, achieving a brightness of 82% ISO with a FDED sequence compared to a brightness of 88% ISO with a CE1DEzD sequence [154]. Recently, Kondo et al. have reported biobleaching of hardwood kraft pulp with lignin-degrading fungi P. sordia YK-624 [147]. A three-stage bleaching process (5 days fungal treatment, alkaline extraction and again 5 days fungal treatment) bleached the kraft pulp to about 80% brightness in 10 days, although brightness increase of the pulp with a one-stage continuous fungal treatment for 10 days leveled off after 5 days incubation. Wroblewska and Zielinsk [164] examined biodelignification of beech and birch pulp wood by selected white-rot fungi. One of the strains, designated as DL-Sth-4, was found to be the best for selective delignifiction of beech wood. About 25% lignin was lost with very little loss in cellulose content. Pazukhina et al. used the culture filtrate of several white-rot fungi P. sanguinea, C. versicolor, Ganoderma applanatum and Trichoptum biforma for bleaching hardwood kraft pulp [165]. P. sanguinea showed the highest selectivity in lignin degradation. Nishida et al. [166] investigated the biobleaching of hardwood unbleached kraft pulp by P. chrysosporium and T. versicolor in the solid state and liquid state fermentation systems with four different culture media (low nitrogen-high carbon, low nitrogenqow carbon, high nitrogen-high carbon and high nitrogen-low carbon). In the solid state fermentation system with L N - H C culture medium, pulp brightness increased by 15 and 30 points after 5 days of treatment with T. versicolor and P. chrysosporium respectively. The pulp kappa number decreased with the increasing brightness, and a positive correlation between the kappa number decrease and brightness increase of the fungus-treated pulp was observed. Delignification by white-rot fungi is strongly influenced by culture conditions. The major culture parameters affecting lignin degradation are: growth substrate, nitrogen availability, other culture conditions, oxygen concentration and mode of cultivation. P. chrysosporium and C. versicolor require a carbon


233

source such as glucose or cellulose to metabolize lignin to form carbon dioxide [167], although the role played by the carbon source is not clear. Glucose supplementation at low concentration (0.47%) was found to stimulate delignification in kraft pulps [15!] and lignin in aspen wood [168]. However, there was no difference in the degree of delignification for thermomechanical pulp (TMP) at glucose concentrations ranging from 0 to 35% [169]. Glucose was added in most studies to maintain pulp yield because it is a repressor of endo-l,4,flgluconase, mannanase, xylanase, aryl-/%glucosidase, pectinase and cellulase, which attack the carbohydrate fraction of the pulp. The white-rot fungus IZU-154 was able to degrade lignin in hardwood kraft pulp without an exogenous energy source and without significantly affecting pulp yield. Fusarium solani and Aspergillus japonicus were also able to degrade lignin without additional carbon source [154, 170-174]. Nutrient depletion, particularly nitrogen depletion, appears to trigger the development of the ligninolytic system in P. chrysosproium, C. versicolor and Phlebia brevispora, Phanerochaete sordida and Phlebia radiata with a few exceptions [152, 168, 175-179]. The effect of nitrogen depletion does not necessarily hold true for all white-rot fungi, as the lignin-degrading abilities of P. sajor-caju and L. edodes do not show strong stimulation by nitrogen depletion [180]. The ligninolytic system of P. chrysosporium is also triggered by limitations in carbon, sulphur, Zn z+, Fe 2+ and Mo 4+, but not phosphorus [30, 181, 182]. Other culture conditions such as temperature and pH have not been studied extensively [181]. Tran and Chambers [151] reported that the optimal temperature for delignification by P. chrysosporium was 38 ~ but Drew and Kadan [183] found that the extent of degradation of 14C-kraft lignin in 15 days at 28 ~ was about 4 times that at 38 ~ Optimum pH values reported for lignin degradation are different for different fungal strains. Tran and Chambers reported that the optimal pH of P. chrysosporium was 3.5 for delignification and 4.5 for growth [151]. Kirk et al. [184] reported an optimum pH of 4 4 . 5 for their strain. Suppression of delignification was noted at pH less than 3.5 and greater than 5.5 [ 184]. Recently, Royle et al. reported increased lignin degradation by P. chrysosporium and L. edodes at pH 3 [180]. o-Phthalate was found to inhibit delignification [185]; in one study, lignin degradation rates were doubled by changing the buffer from o-phthalate to 2,2-dimethylsuccinate (DMS) [176]. Lignin degradation in white-rot fungi is primarily oxidative. It occurred faster in cultures of P. chrysosporium which had been flushed with pure oxygen than in those flushed with air [-135, 151,169, 186-188]. Enhanced lignin degradation due to elevated oxygen partial pressures has also been reported for other white-rot fungi including C. versicolor, Pycnoporus cinnabarinus, Lentinus edodes, Giifola frondosa, Polyporus burmalus and Merulius tremellosus [189]. However, other species such as Gleoporus dichrous, Pleurotus ostreatus, Bondarzewia berkeleyi and IZU-154 were less responsive to increased oxygen concentrations [154, 189]. The addition of polydimethylsiloxane (PDMS) oxygen carriers enhanced lignin degradation by C. versicolor [190]. However, they alone were not found to appreciably increase the brightness of hardwood kraft

234


pulp [191]. Together with C. versicolor they acted synergistically, resulting in an overall increase of approximately 10 brightness points [190]. Lignin degradation is also strongly influenced by the mode of cultivation. Most researchers have studied stationary cultures, although dependence on stationary cultures causes difficulties in scale-up. Also, agitation is an important consideration because of the role of oxygen mass transfer. In a number of studies, agitation in reciprocating and gyratory shakers caused the formation of pellets consisting of mycelia entangled with pulp. The formation of mycelial pellets resulted in suppression of degradation of kraft lignin and lignin model compounds by P. chrysosporium [-151,156, 169, 192, 193]. Similar delignification rates were noted in agitated and stationary cultures and also with the addition of detergent to the culture [187, 194]. Effective delignification has also been reported in agitated cultures of C. versicolor in which the formation of mycelial pellets was prevented and in cultures of P. chrysosporium in the presence of veratryl alcohol [195, 196, 199]. P. chrysosporium and C. versicolor have been successfully immobilized on polyurethane foam [153, 157, 190, 197]. Immobilized and free cultures of C. versicolor have been found to bleach hardwood and softwood kraft pulp at a comparable rate and to a similar extent [153, 157]. The results showed that intimate contact between the fungal hyphae and pulp fibres was not required as long as the media was renewed through contact with the fungus [132, 153]. Immobilization enabled the pulp to be separated from the mycelia. Another advantage of immobilization was that the same fungal biomass could be reused to treat other batches of pulp either immediately or after storage at 4~ [153]. A serious shortcoming of the fungal bleaching process is the long incubation time required, ranging from 5 to 14 days for both hardwood and softwood pulps. Softwood incubation periods are likely to be longer than those for hardwood pulps because softwood pulps require a longer time lag (6 days) before kappa number decrease or brightness increase can occur [157]. Hardwood pulps generally have a lag time of 1-2 days [152-154, 160], followed by rapid and then slower delignification. Unfortunately, the size of the fungal bioreactor would have to be very large, considering that daily production could range from 200 to over 1000 air-dried tonnes of pulps. Most researchers have performed fungal bleaching experiments at low pulp consistencies of 0.5-2% (w/v) [151 154, 190]. Only Fujita et al. [154] have investigated fungal bleaching at high consistency (16-24%), which would allow for a smaller reactor. Both reactor size and incubation period will have to be reduced in fungal bleaching if it is to become an economical alternative. Jurasek and Paice [198] suggested that the lignin may become more flexible and hydrophilic as a result of fungal enzyme action, resulting in a sorer pulp with improved bonding and stronger paper characteristics. Reduced colour reversion was another benefit noted with the fungus IZU-154 [154]. Some viscosity loss, indicating limited cellulose depolymerization, has been reported

Reductionof OrganochlorineCompoundsin BleachPlant Effluents

235

as a result of fungal bleaching [153, 154, 157, 199]. However, based upon experiments done with free and immobilized cultures, Kirkpatrick et al. [ 153] reported that up to 25% of the reduction in the pulp viscosity may be due to the presence of fungal mycelia rather than cellulose cleavage. Although fungal bleaching is primarily an oxidative process, it appears to be more selective than oxygen bleaching at high pH and at kappa number less than 17 because there is a better retention of pulp viscosity. Only a few researchers have measured the impact of fungal bleaching on effluent quality. In a Japanese study with FCED bleaching sequence, the COD and color in the bleach plant waste water were reduced by 50% and 80% respectively, assuming that the filtrate from the fungal bleaching stage was sent to a kraft chemical recovery system [1541. Whether this could occur in practice would depend on the capacity available in the recovery furnace. The authors suggested that higher reductions could be obtained with an FED or FE1D1EzDz sequence, although there may be slight loss in pulp yield [1541. Despite the emphasis on fungal bleaching as a means to reduce the use of chlorine and the associated formation of chlorinated organics, the effect upon chlorinated organic discharges has not been reported. As this is an important factor in the choice of any alternative bleaching sequence, quantitative information in this area is needed.

4 Treatment of Bleach Plant Effluents Numerous physicochemical methods have been used for the treatment of bleach plant effluents. These treatments include precipitation with lime [200-203], alum and metal ions [204, 205] and synthetic polymeric coagulants [2061, adsorption on activated carbon [207], natural clays [208-210] and polymeric adsorbents [211, 212], membrane techniques [2131, rapid filtration on soil [214-2161, UV-irradiation [217-2191, and oxidation using oxygen [2201, sulfurdioxide [2211, hydrogen peroxide and sodium hypochlorite [222-224]. The problems underlying the physicochemical treatments are those associated with cost and reliability. Coagulation and precipitation produce a voluminous sludge which is very difficult to dewater. Usually, an extreme pH range is used for optimum treatment, and the pH needs to be readjusted to neutral before discharge. Oxidation using ozone and hydrogen peroxide are costly, and oxidation using chlorine species generates secondary pollutants such as chlorinated organics. The membrane techniques require pretreatment and a large capital investment. Membrane fouling is also a problem with the membrane technique. Biotechnological methods have the potential to eliminate/reduce the problems associated with physicochemical methods, and are described in the following section.

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4.1 Biological Treatment Biological treatment is known to be effective in reducing the BOD and simultaneously the toxicity of kraft mill effluents [225, 226]. Some aspects of biological treatment have been reviewed by Boman et al. [227].

4.1.1 Using Bacteria 4.1.1.1 Aerobic Processes. Biological oxidation is the most widely used technique to remove BOD and chlorinated organics because of its effectiveness and low cost. Rogers et al. [228] treated the bleached kraft mill effluent in a benchscale aerated lagoon for 29, 58 and 99 h and showed that toxicity, BOD and resin acids were most consistently reduced during the 99 h treatment. Leach et al. [229] reported the biodegradation of seven compounds representing the major categories of toxicants in a laboratory batch aerated lagoon. Resin acids (major source of acute toxicity) were readily biodegradable, but only part (less than 30%) of the load of chlorophenolic compounds was removed [34, 77, 229, 230]. However, Gergov et al. [230] reported that biological treatment, especially the activated sludge process, removed 75-95% of chlorophenolics. Chlorinated neutral organic compounds (mutagenicity of the spent liquor) are removed effectively [231]. Chloroform is stripped off by the air during biological treatment. COD, TOC1 and high molecular weight material are reduced to a lesser extent [44]. Eriksson and Kolar [51] have shown that the high-molecular-mass fraction in bleach effluent cannot be degraded in an aerated lagoon. While the Swedish researchers found that the biological treatment is ineffective in removing TOC1, American mills reported, an average of 5 0 4 0 % TOC1 removal by an aerated lagoon or activated sludge process. Gergov et al. [230] investigated pollutant removal efficiencies in mill scale biological treatment systems. They found that 48-65% of AOX was removed in the activated sludge process. The aerated lagoon was found to be less efficient than the activated sludge. Recently, Deardorff et al. [232] reported that the efficiency of AOX removal through biotreatment of combined bleach plant effluent increases with increasing chlorine dioxide substitution. Biological treatment in an aerated lagoon reduced the concentration of polychlorinated phenolic compounds by 97%. Jokela et al. [233] reported that aerobic lagoon system removed 58-60% of the organochlorine compounds from the water phase, and the full scale activated sludge plants removed 19-55%. Both biotreatments removed all sizes and classes of organochlorine molecules and slightly shifted the relative size distribution of the compounds remaining in the water phase towards the higher molecular masses. Graves et al. [234] examined the combined effects of oxygen delignification, C102 substitution and biological treatment on pollutants levels in bleach plant


237

effluents. Biological treatment reduced COD, BOD, AOX and toxicity, but did not reduce colour. Lafond and Ferguson [-235] reported that aerobic treatment in activated sludge reactors removed 14-25% of AOX. To enhance the efficiency of the biological treatment, Ek and Eriksson [236] combined ultrafiltration and biological treatment. The result of the process was better than the sum of the two processes. The detoxification of the effluent during ultrafiltration was believed to be responsible for the improvement. The mechanisms leading to the removal of chlorinated organics include stripping of voltatiles, hydrolysis, chemical oxidation, biological oxidation, foam separation, adsorption, and precipitation. 4.1.1.2 Anaerobic Processes. Anaerobic biological treatment can also efficiently

destroy chlorophenolic compounds, mutagenicity and acute toxicity [77, 237]. The Enso-Fenox Process was capable of removing 64-94% of the chlorophenol load and toxicity, mutagenicity and chloroform in the bleaching effluent. McFarlane and Greenwood [-238] removed 92% of color in kraft bleaching effluent in a granular activated carbon anaerobic system in a 2-day hydraulic residence time. The decolorization was decreased to 75% when the retention time was shortened to 0.85 day. Gunnarsson et al. [239] treated 40% E stage effluent combined with sulphite evaporator condensate in an anaerobic reactor. Approximately 60% COD and 90% BOD reductions were achieved. The methane yield was high, but the reduction of TOC1 was limited, which was still better than the parallel aerobic treatment of the same effluent. Many compounds in the bleaching effluent were reported to be toxic to anaerobic microorganisms. These compounds include residual H202 in CTMP bleaching effluent, chelating agents, extractives and chlorinated compounds [240-243]. Adoption of microorganisms to the effluent containing these compounds is necessary to achieve a successful treatment [-240]. Raizer-Neto et al. [243] studied the efficiency of anaerobic treatment in reducing chlorinated organic compounds in a fixed bed reactor. AOX removal efficiency was primarily affected by increasing AOX concentration and was only about 50% for 50 mg/1 COD. BOD 5 removal efficiencies were affected at higher AOX concentrations of 100 mg/1. AOX loading rate or hydraulic residence time were found to be more important limiting factors than bleaching effluent AOX concentration. For AOX loading rates under 40 mg/1 and an AOX concentration as high as 130 mg/1 in the effluent feed, COD and BOD5 removal were about 80%. Fitzsimons et al. [244] reported 35-40% COD and 42-45% AOX reductions in a 1.5-day anaerobic treatment. They found that at least part of the higher molecular mass was dechlorinated. Since anaerobic microorganisms are believed to be unable to degrade the highmolecular-mass chlorolignin, the dechlorination of this could be mainly due to physical and/or chemical action in the anaerobic process. The anaerobic dechlorination of chlorolignins is due to a combination of energy metabolism growth, chemical hydrolysis and probably adsorption and/or insolubilization [244, 245].

238


E K and Eriksson proposed a process based on U F and anaerobic and aerobic biological treatments [236]. The U F was used to separate the highmolecular-mass material, which is relatively resistant to biological degradation. Anaerobic microorganisms were believed to be able to remove highly chlorinated substances more efficiently than aerobic microorganisms. The last remaining chlorine atom was removed by aerobic microorganisms. The combined treatments typically removed 80% of the AOX, C O D and chlorinated phenolics and completely eliminated chlorate. Lafound and Ferguson [235] reported that anaerobic treatment in an upflow hybrid reactor removed 17-40% of AOX. Armentate et al. [246] investigated an integrated anaerobic/aerobic process for the biodegradation of chlorinated aromatic compounds. The sludge obtained from the anaerobic digester of a commercial treatment plant was used to obtain an anaerobic consortium capable of partially dechlorinating 2,4,6-trichiorophenol. The clarified and sterilized effluent from the same anaerobic digester was used as the medium for the anaerobic consortium. During the anaerobic process, 2,4,6-TCP was first dechlorinated to 2,4-dichlorophenol and then to 4-chlorophenol. The stoichiometric amount of 4-CP was recovered. Similar results were obtained when the anaerobic microorganisms were immobilized. After immobilization, the consortium was able to dechlorinate 150 laM of 2,4,6-TCP in four days. Pseudomonas 91athei and an indigenous culture obtained from same sludge used to produce the anaerobic enrichment culture were shown to be able to degrade the 4-CP produced from the anaerobic dechlorination of 2,4,6-TCP. Strehler and Welander [247] investigated the removal of AOX and C O D from bleached kraft mill effluent in laboratory and pilot scale aerobic suspended carrier reactors and abiotic thermoalkaline reactors. At pH 7.0, 37 ~ and HRTs longer than 3.5 hours, a maximum C O D removal of 55% was achieved in the suspended carrier process. The C O D conversion rate at the minimum H R T was 2.6 kg C O D m 3 d-1. The suspended carrier treatment was operated successfully at pH 9.0 and 45 ~ and at pH 7.0 and 50 ~ giving > 50% C O D removal with an H R T of four hours. The AOX removal achieved at p H 9.0 and 45 ~ (50%) was significantly higher than that at pH 7.0 and 37 ~ (39%) because of an increased abiotic dechlorination at the higher p H and temperature levels. These researchers further studied sequential thermoalkaline and biological treatment on a pilot scale. Thermoalkaline treatment at pH 10.0, 54 ~ and an H R T of two hours followed by biological treatment at pH 8.0, 35 ~ and an H R T of four hours removed almost 80% of the AOX and 50% of the C O D from the kraft mill effluents.

4.1.2 Using Fungi The reason why bacteria show low efficiency in removing COD, TOC1 and high-molecular-mass chlorolignins from bleaching effluent is that bacteria degrade the substrate by an intracellular enzyme system. The substrate must be

Reduction of Organochlorine Compoundsin Bleach Plant Effluents

239

able to pass the bacterial cell membrane in order to be degraded. In the light of this, the inefficiency of bacteria in degrading high-molecular-mass chlorolignins is understandable. Since most TOC1 and COD is due to the high-molecularmass chlorolignins, low COD and TOC1 removal efflciencies are also to be expected. An approach to degrading high molecular weight chlorolignins is to use white-rot fungi, the only known microorganisms to efficiently degrade lignin. They generate a lignolytic activity to degrade lignin in a so-called secondary metabolism stage when one of several nutrients, nitrogen, phosphorous or carbon is depleted (unfavorable natural conditions). White-rot fungi excrete an operating system including extracellular enzymes which can degrade highmolecular-mass chlorolignins effectively. So far, the widest applications of the white-rot fungi in waste-water purification have been concentrated on the decolorization of bleaching or pulp mill effluents [248-250]. The reaction mechanism by which white-rot organisms degrade lignin is in four steps. Ligninase enzymes catalyze the first two steps. This is followed by an aromatic hydroxylation step that produces catechol structures in the resulting fragments [251, 252]. The fourth step is an oxidative ring cleavage catalyzed by a dioxygenase enzyme. The products of this reaction are converted by the organism into CO2 and water. Chlorinated aromatics are dealt with by the organism in a like manner by converting them into catechol structures whose rings can be cleaved via the same dioxygenase enzyme, the products being CO2, water and inorganic chlorides. Many other organisms can grow directly on chlorinated organics, degrading them via the same or similar dioxygenase pathways or in some cases monooxygenase pathways. The best known white-rot fungus is Phanerochaete chrysosporium. This fungus is known to secrete a family of enzymes which degrade both lignin and modified lignin/-249]. Both high- and low-molecular-mass chlorolignins generated during the pulp bleaching are significantly degraded [253, 254]. The fungus reduces the COD by degrading the chlorolignins to CO2 and chloride, decolorizes the bleaching effluent by destroying both the color bodies and chromophoric structure, and removes TOC1 by converting it to inorganic chloride [253-255]. Most of the low-molecular-weight chlorophenolics disappear after 1 day's fungal treatment [256, 257]. This particular fungus is also known to be able to degrade refractory compounds such as TNT, PCB and Lindane, DDT, chlorinated dioxin and other difficult biodegradable compounds [258-260]. Decompositions of lignin to CO2 by the lignin-decomposing fungus P. chrysosporium requires a growth substrate such as cellulose or glucose. Growth on lignin as a sole carbon source is negligible. In addition to its requirement for a co-substrate, the ligninolytic system (a) is produced even in the absence of lignin, (b) is expressed only during secondary metabolism, (c) is triggered by carbon, sulfur, or nitrogen limitation, (d) is strongly repressed by glutamate and certain other amino acids, (e) is sensitive to the balance of trace metals supplied, (f) has a relatively narrow pH optimum (4 5), and (g) is

240


markedly affected by oxygen concentration [-261-265]. The fungal decolorization, like lignin metabolism, is a secondary metabolic event and is probably attributable to lignin metabolism since lignin and its degradation products are the main color source in E1 effluent. The optimum conditions favoring fungal growth are quite different from those favoring decolorization. The pH range for optimum growth is 4.3~4.8 and decolorization is greatly retarded below pH 4.0 or above 5.0 because of poor growth [261]. The optimum temperature for the growth of fungus is 40 ~ whereas decolorization is not limited to the same narrow range of temperature, but takes place with little decrease in rate at temperatures as low as 25 ~ [261]. The fungal decolorization requires oxygen and a co-substrate, but, unlike fungal growth, the addition of a nitrogen source is not necessary. Eaton et al. [266] have outlined a process based on laboratory and bench scale experiments for decolorization by P. chrysosporium. The fungi require a growth substrate for decolorization just as they do for lignin degradation. Investigations have demonstrated that the cellulose-rich primary sludge serves this purpose well. The fungus requires a fixed surface for pregrowth without agitation in shallow medium for efficient color reduction. Fixed film reactors of the rotating biological contactor (RBC) design, which are commercially available and are presently used in waste treatment have been proved to be effective. Color removal is greatly stimulated by an oxygen-enriched atmosphere. The RBC provides good aeration and can be enclosed for the addition of supplementary oxygen. A growth stage is necessary before decolorization begins. During this stage, nutrient nitrogen is depleted, and the fungus becomes ligninolytic and able to decolorize. Based on the above considerations, the process shown in Fig. 3, which has been termed the FPL/NCSU/MyCoR process [267], can be outlined. This mycelial color removal process resulted from the cooperative research between the US Forest Products Laboratory and North Carolina State University. A fixed film MyCoR reactor is charged with growth nutrients, which can include primary sludge as the carbon source, and is inoculated with a suitable fungus. Depending on the mill, the sludge will provide

Influent

uent

Fig. 3. Rotatingbiologicalcontactorfor fungaldecolorizationof pulp mill effluent


241

some of the required mineral nutrients and trace elements as well as carbon. Nitrogen-rich secondary sludge can also be used to supply the nitrogen required for growth. After the mycelium has grown over the reactor surface, it depletes the available nitrogen and becomes ligninolytic (pregrowth stage 2 to 4 d). The reactor is then ready for use. Operation for over 60 days has been achieved in bench reactors in a batch mode. The process converts 70% of the organic chlorides to inorganic chloride in 48 h while decolorizing the effluent and reducing both COD and BOD by about half. Huynh et al. [268] used the MyCoR process for the treatment of the chlorinated low-molecular-mass phenols of the E1 effluent. Their results showed that most of the chlorinated phenols and other low-molecular-mass components of the effluent were removed during fungal treatment. Pellinen et al. [269] have reported that the MyCoR process can be considerably improved in terms of COD removal by simply using less glucose as the carbon source for the fungi, P. chrysosporium. However, the dechlorination was reported to be faster at high glucose concentration. The kinetics of decolorization of E1 effluent with P. chrysosporium in an RBC under improved conditions have been studied by Yin et al. [270]. The kinetic model developed for 1 and 2-d retention times showed a characteristic pattern. The overall decolorization process can be divided into three stages, viz. a rapid color reduction in the Ist hour of contact between the effluent and the fungus followed by a zero order reaction and then a 1st order reaction. The color removal rate on the second day of the 2-d batch treatment was less than that on the first day. The decolorization in a continuous flow reactor achieved approximately the same daily color removal rate, but the fungus had a longer working life than when in the batch reactor, thereby removing more color over the fungal lifetime. Pellinen et al. [271] studied dechlorination of high-molecular-mass chlorolignin in first extraction stage effluent with white-rot fungus P. chrysosporium immobilized on an RBC. The total organic chlorine content of chlorolignin decreased almost by 50% during one day of treatment. Correlation studies suggested that dechlorination, decolorization and degradation of chlorolignin (as COD decrease) are metabolically connected, although these processes have different rates. Size exclusion chromatography showed that polymerization took place in the early stage of the treatment. Low-molecular-mass degradation products were not observed. A sequential biological treatment using this fungus and these bacteria was studied to decrease TOC1, color and COD in conventional softwood kraft pulp bleaching effluent by Yin et al. [272]. In six variations of the white-rot fungus/bacteria system studied (Table 8), only the degree of fungal treatment was varied. In three of the six variations, ultrafiltration was also used to concentrate high-molecular-mass chlorolignins and to reduce effluent volume (and thus cost) prior to fungal treatment. The best sequence, using ultrafiltration/white-rot fungus/bacteria, removed 71% TOC1, 50% COD and 65% color in the effluent (Table 8). Fungal treatment enhances the ability of bacteria to degrade and dechlorinate chlorinated organics in the effluent. The fungus is able to remove organic chlorine from chlorolignins and to attack the high- and low-molecularmass chlorolignins.

242


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243

Guo et al. [257] investigated degradation of model compounds - chlorophenols (pentachlorophenols, 2,4,6-trichlorophenol, and 2,4-dichlorophenol) and chloroguaiacols (4,5-dichloroguaiacol and 4,5,6-trichloroguaiacol) in pure water solution by fungal treatment using an RBC. They found that at concentration of 30 mg/1, 80-85% of chlorophenols and chloroguaiacols could be degraded after 3-4 h of treatment. Meanwhile, some of these compounds were methylated to chloroanisoles and chloroveratrols. Messner et al. [273] developed a similar process, i.e. the MyCoPOR. In this process, porous carrier material is inoculated with spores of Phanerochaete chrysosporium. Shaking-flask tests with and without foam as porus carrier material showed that the decolorization of an E1 effluent from a sulfite plant was improved from 50% without foam as supporting material to 70-80% with foam. Decolorization was accompanied by about 70% AOX reduction when using foam. If nutrients were replaced and effluent removed daily, the active lifetime of the mycelium was 6-8 weeks, depending on the fungal strain. A trickling filter system was found to be the most effective for the degradation of effluents. With daily replacement of the medium/effluent solution, the decolorization rate of the E1 effluent was 70% when aerated by air, and AOX reduction was 60%. The fastest degradation of the effluent took place during the first 3-6 hours, when about 50% of the color and AOX were reduced. Another white-rot fungus, which has shown good performance, is Coriolus versicolor. It requires a growth substrate such as cellulose or glucose for the decomposition of lignin to carbon dioxide [274]. The culture conditions favoring lignin degradation are similar to those favoring fungal decolorization. Livernoche et al. [274] showed that C. versicolor in liquid culture removed over 60% of the color of the combined bleach kraft effluent within 6 d in the presence of sucrose. Decolorization of effluent was more efficient when the concentration of sucrose (growth substrate) and inoculum was high. Treatment of the same effluent with this fungus, immobilized in beads of calcium alginate gel, resulted in 80% decolorization after 3 d in the presence of sucrose. The pH of the effluent during decolorization decreased from 5.7 to 3.6, probably due to formation of organic acids resulting from the metabolism of the immobilized fungus. However, decolorization was not due to lowering of pH. The decolorization process affected not only the dissolved chromophores but also the suspended solids. The solids after centrifugation of the zero time samples were dark brown, while the solids after 4-d incubation were light brown. The beads with the immobilized mycelium remained light-coloured throughout the experiments with no indication of accumulation of the effluent chromophores. Decolorization was achieved more rapidly at pH 5.0 than at pH 7.0. Recycled beads could remove color efficiently and repeatedly in the presence of air but not under anaerobic conditions. Biological reactors of the airlift type using calcium alginate beads to immobilize the fungus C. versicolor have been used to study the continuous decolorization of kraft mill effluents [275]. The effluent used contained only sucrose and no other nutrient source. An empirical kinetic model was proposed to describe

244


the decolorization process caused by this fungus, but it did not shed any light on the chemical mechanism involved in the decolorization. Direct use of suspended mycelium of the fungus, C. versicolor, may not be feasible because of the problem of viscosity, oxygen transfer and recycling of the fungus. The fungus was, therefore, grown in the form of pellets, thus eliminating the problems with biomass recycling and making it possible to use a larger amount [276]. Rate of decolorization with fungal pellets was almost ten times as high in batch culture as in continuous culture under similar conditions. The capacity for decolorization decreased markedly with increase of lignin loading [-276]. Bergbauer and Kraepelin [277] showed that C. versicolor efficiently degraded chlorolignins from bleaching effluent. More than 50% of the chlorolignins were degraded in a 9-d incubation period, resulting in a 39% reduction in AOX and an 84% decrease in effluent color. In a 3-1 laboratory fermenter with 0.8% glucose and 12 m M ammonium sulphate, about 88% color reduction was achieved within 3 d. Simultaneously, the concentration of AOX dropped from initially 40 mg/1 to 21.9 mg/1, a 45% reduction within 2 d. With malt extract instead of glucose, reduction of color unit as well as AOX values were nearly the same [278]. Bajpai et al. [279] used pellets of T. versicolor strain B-7 for decolorization of E1 effluent. The mycelial pellets oxidized the chromophores of the effluent in the presence of any of the cosubstrates sucrose, glucose, starch, ethanol, carboxymethylcellulose, microcrystalline cellulose, pulp and malt extract. The highest decolorization was obtained in the case of glucose. Optimum pH and temperature were 4.5-5.5 and 30 ~ respectively. In the batch reactor with an effluent of 7000 color units per litre, the maximum color reduction of 93% was obtained in 48 h with a C O D reduction of 35%, whereas, in a continuous reactor, the same level of color and C O D reduction was obtained in 38 h residence time. No loss in decolorization ability of mycelial pellets was obtained when the reactor was operated continuously for more than 30 d. They also used T. versicolor for decolorization of effluent from a pulp mill utilizing agriresidues [280]. With an effluent of 18 500 color units, a maximum color reduction of 92 % with a C O D reduction of 69% was obtained. Royer et al. [281] described the use of the pellets of C. versicolor to decolorize ultrafiltered kraft liquor in nonsterile conditions with a negligible loss of activity. The rate of decolorization was observed to be linearly related to the liquor concentration and was lower than that obtained in the M y C o R process. This could be due to the lower temperature (22 ~ used in this work and to the use of pellets with relatively large diameters which could limit the microbial activity as compared to the free mycelium used in the M y C o R process. An effective decolorization of effluent having 400-500 color units/1 can be obtained in presence of a simple carbon source such as glucose. In the repeated batch culture, the pellets exhibited a loss of activity dependent on the initial color concentration. The production of the extra cellular enzyme, laccase, was followed but could not be used as an indicator of the ligninolytic activity.


245

Archibald et al. [282] studied the fungal requirements for optimal growth and decolorization and the mechanism of chromophore degradation by C. versicolor. Simple carbohydrates were essential for effective decolorization, and a medium composed of inexpensive industrial by-products provided excellent growth and decolorization. The regulation or control of E1 effluent decolorization appears to differ substantially from that of lignin peroxidase production and the induction of ligninolytic activity. In particular, modulation of decolorization by trace metals, nitrogen and carbon limitation, culture age and veratryl alcohol was not observed. Marton et al. [283] reported that C. versicolor reduced the characteristic dark brown color of diluted alkaline lignin solutions. They found, however, about half of the color and one fourth of the aromatically absorbing substances were recovered from the fungal cells by alkaline extraction. Therefore, it was concluded that adsorption played a major role in lignin degradation. Decolorization also proceeded anaerobically but to a lesser extent. Treatment of E1 stage eff‚ with ozone and the fungus C. versicolor has also been tried [284]. Both ozone treatment and biological treatment effectively destroyed effluent chromophores, but the fungal process resulted in greater degradation as expressed by COD removal. Monoaromatic chlorophenolics and toxicity were removed partially by ozone and completely by C. versicolor. Molecular weight distributions showed roughly equal degradation of all sizes of molecules in both treatments. The combination of a brief ozone treatment with a subsequent fungal treatment revealed a synergism between the two decolorization mechanisms on E1 stage effluent. Effluent was pretreated with ozone (ll0-160mg/1) or C. versicolor (24h with 2-5g/1 wet weight fungal biomass). The pretreatment was followed by a 5-d incubation with C. versicolor. It was noted that partial color removal by ozone pretreatment allowed more effective removal by the fungus than that by fungal pretreatment. After 20h, 46-53% decolorization was observed for ozonepretreated effluents, compared to 29% for fungal treatment alone. The contribution of ozone seemed to be most important in the first 24 h following the pretreatment. Ozone pretreatment also produced a small improvement in the bioavailability of effluent organics to the fungus. A partial replacement of chlorine by ozone in the bleach plant (the addition of an ozone bleaching step) or a brief ozone pretreatment of E1 stage effluent should considerably reduce the low-molecular-mass toxic chlorophenolics. In addition, the use of ozone should also improve decolorization by subsequent fungal and possibly bacterial treatments. Another white-rot fungus, Schizophyllum commune, has also been found to decolorize and degrade lignin in pulp and paper mill effluent [-285]. The fungus was able to degrade lignin in the presence of an easily metabolizable carbon source. The addition of carbon and nitrogen not only improved the decolorizing efficiency of the fungus but also resulted in reduction of the BOD and COD of the effluent. Sucrose was found to be the best carbon source for the breakdown of lignin. A 2 d incubation period was sufficient for lignin degradation by

246


S. commune. The efficiency of treatment of effluent with this fungus was highest at pH 4 5 and was further improved by intermittent aeration. Under optimum conditions, S. commune reduced the colour of the effluent by 90% and also reduced BOD and COD by 70% and 72% during a 2 d incubation. A white-rot fungus, Tinctoporia borbonica, has been reported to decolorize the kraft waste liquor to a light yellow color [286]. About 99% colour reduction was achieved after 4 d of cultivation. Measurement of the culture filtrate by ultraviolet spectroscopy showed that the chlorine-oxylignin content also decreased with time, and measurement of the culture filtrate plus mycelial extract after 14 d of cultivation showed the total removal of the chlorine-oxylignin content. Addition of a carbon and nitrogen source was found to improve decolorization of pulp and paper mill waste water by the fungus Aspergillus niger, leaving 19% of the original color, and reduced about 43% BOD and 41% COD after 2 d of incubation [287]. Tono et al. [288] reported that Aspergillus sp and Penicillium sp achieved 90% decolorization in one week's treatment at 30 ~ and at pH 6.8. Later, Milstein et al. [289] reported that these microorganisms removed appreciable levels of chlorophenols as well as chloroorganics from the bleach effluent. Prasad and Joyce [290] used Trichoderma sp., one of the fungi imperfectii, to decolorize the hardwood E1 stage effluent. Glucose was found to be the most effective carbohydrate utilized by the fungus, as it stimulated substantial color reduction without any increase in COD. Addition of nitrogen did not stimulate the decolorization process, indicating that it is not a rate-limiting factor. The optimum pH for decolorization and growth was 4.0. Under optimal conditions, total colour and COD decreased by almost 85% and 25% respectively after cultivation for 3 d. Since the majority of TOC1 and color is due to higher molecular weight chlorolignins, future research should concentrate on the fate of highmolecular-mass chlorolignins in biological treatment or in the natural environment. Since bacteria significantly degrade only those chloroorganics of molecular mass less than 800-1000, research is needed to decrease the chlorolignin molecular mass or to remove high molecular mass chlorolignins before biological treatment is applied, in order to enhance the biotreatability of bleaching effluent. Three approaches to decreasing the molecular weight of chlorolignin should be studied: (1) a modified bleaching process such as oxygen prebleaching and a high level of chlorine dioxide substitution, (2) a two-stage biological treatment such as white-rot fungus followed by biological treatment, and (3) physicochemical treatment followed by biological treatment. Ultrafiltration can separate high molecular weight material, and biological treatability of permeate can be enhanced. Another method is precipitation, which preferentially removes high-molecular-mass chlorolignins. The treated effluent should be more amenable to biological treatment.


247

4.2 Enzymatic Treatment Some enzymes also seem to have the potential to remove color and AOX from pulp and paper mill effluents. Peroxidase, laccase etc. are the most important of these. The use of microbial or enzyme-based treatment offers some distinct advantages over physical and chemical AOX precipitation methods, in that only catalytic and not stoichiometric amounts of the reagents are needed, and the low organic concentrations and large volumes typical of bleaching effluents are, therefore, less of a problem. Also, both complete microbial systems and isolated enzymes such as peroxidases and laccases have been shown to reduce the acute toxicity by polymer• and thereby rendering less soluble many of the lowmolecular-mass nonchlorinated and polychlorinated phenolics [291 295]. It has been reported that lignin-type peroxidases secreted by white-rot fungi are involved in the degradation of a whole range of organic pollutants including PCB, D D T and Indane, chlorinated anilines, polychlorinated phenolics including P C P and chlorolignins, and many more [296-300]. Lyr [301] reported that laccase of T. versicolor partially dechlorinates PCP, and Hammel and Tardone [302] reported that peroxidase from P. chrysosporium can partially dechlorinate P C P and 2,4,6-trichlorophenol. Arcand and Archibald [303] carried out a systematic study on direct dechlorination of chlorophenolic compounds in pulp and paper mill effluents by laccase from T. versicolor. It was found that most of the laccase secreted by T. versicolor could partially dechlorinate a variety of chlorophenolics. The rate and extent of C1- release were found to be substantially affected by substrate and enzyme concentration and the presence of multiple laccase substrates. The dechlorination was found to be accompanied by extensive polymerization of the substrate. Table 9 shows a comparison of the removal of chlorophenolics from a mixture in E1 effluent by crude laccase with Table 9. Removalof chlorophenoliccompoundsfrom a mixture in high-molecular-mass E1 effluent by crude laccase in 30 mina and T. versicolor myceliumin 3 h Reduction (%) Chlorophenolic Compounds

Concentration (gM)

Laccase

Mycelium

2-Chlorophenol 3-Chlorophenol 4-Chlorophenol 2,4-Dichlorophenol 2,4,6-Trichlorophenol 2,3,4,6-Tetrachlorophenol Pentachlorophenol 4,5-Dichlorognaiacol 4,5,6-Trichloroguaiacol Tetrachloroguaiacol

137 18 31 21 26 18 24 52 32 17

35.5 _+1.4 51.9 + 4.3 48.1 • 2.9 45.0 • 2.3 72.4 _+5.2 40.7 • 8.2 34.0 • 8.9 100 100 76.5 • 21.5

61.5 • 2.0 • 41.6 • 64.9 • 83.8 • 71.2 • 82.1 • 96.8 + 100 95.0 •

2.7 0.1 1.3 1.8 6.2 2.7 3.2 6.1 7.6

aReactions were conducted in 20 ml of E1 effluent containing 20 mM D-glucose,pH 4.6, temp. 25 ~ shaken at 200 rpm in triplicate using either 0.1 U/ml crude T. versicolor laccase or 0.5 g wet weight (25 mgd.w.) washed T. versicolor mycelium Based on data from Ref. [3033

248


that by T. versicolor mycelium, and Table 10 shows the effects of crude laccase on various chlorophenolics. Arcand and Archibald [303] also studied the effects of horseradish and P. chrysosporium peroxidases on the mixture of five chlorophenolics. Both peroxidase enzymes were found to degrade the majority of all substrates except PCP, whereas the P. chrysosporium peroxidases was superior to both horseradish peroxidase and laccase in degrading PCP, it was inferior to horseradish peroxidase in degrading the other four phenolics (Table 11). Paice and Jurasek studied the ability of horseradish peroxidase to catalyze color removal from bleach plant effluents [304]. The color removal from effluents at neutral pH by low levels of hydrogen peroxide was enhanced by the addition of peroxidase. No precipitation occurred during the decolorization process. The catalysis with peroxidase (20 mg/1) was observed over a wide range of peroxide concentrations (0.1-800 mM) but the largest effect was between 1 mM and 100 mM. The pH optimum for catalysis was around 5.0. Compared with mycelial color removal by C. versicolor, the rate of color removal by peroxide plus peroxidase was initially faster (for the first 4 h), but the extent of color removal after 45 h was higher with the fungal treatment. Further addition of peroxidase to the enzyme-treated effluents did not produce additional catalysis. Thus, the peroxide/peroxidase system did not fully represent the metabolic route used by the fungus. One working hypothesis has been proposed to explain the behaviour of enzymes in the decolorization process [304]. Glucose is used by the cell to produce peroxidase. One of the extracellular enzymes often found in white-rot fungi is peroxidase. It seems that this enzyme oxidizes the chromophores and so removes the color from bleaching waste water. Forss et al. [305] have shown that aeration in the presence oflaccase leads to a reduction by over 90% in the amount of phenolic compounds in waste water. They have shown that the amount of chlorophenols is also reduced by 75-99%, depending on the group of chlorophenols (Table 12). The total reduction was 80%. The sample was aerated in the presence of laccase for 1 h and was flocculated with aluminium sulfate. Since polymerization by laccase can be avoided and the degradation of the substrate enhanced by the presence of appropriate reductant, it may be possible to substantially increase the rate and extent of laccase-driven dechlorination. Call (of Lignozym, Germany) has patented a process for decolorization and decontamination of waste water from pulp and paper mill effluents using enzyme [306]. In this process, laccase enzyme from C. versicolor is used along with phenol or nonphenolic aromatic compounds with continuous passing of oxidation materials (H202, 02 or oxygenated air). Almost complete polymerization of the lignin contained in the waste water is achieved, which is 20-50% above the values attainable with the addition of laccase alone. About 70-90% lignin is developed into insoluble form, which is removed by flocculation and filtration. Ferrer et al. reported that immobilized lignin peroxidase decolorized kraft effluent [307]. Novel lignin peroxidases called Pulpases produced by


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Table 12. Concentration of chlorinated phenols in bleach plant effluent from a softwood kraft mill before and after aeration in the presence of laccase and flocculation with alum Phenolic compound

Concentration (rag/l) Initial

Chlorinated phenols Chlorinated vanillins Chlorinated guaiacols Chlorinated catechols Total

Reduction (%)

After 1 h aeration

141 813 516 251

9 176 30 19

93.6 78.3 94.2 92.4

1721

234

86

Based on data from ref. [305]

P. chrysosporium m u t a n t strain SC 26 a n d d e s c r i b e d in 2 p a t e n t s assigned to the Repligen C o r p o r a t i o n [308, 309] have been c l a i m e d to d e c o l o r i z e b l e a c h i n g effluents.

5 Conclusions M a n y process c h a n g e s have been i m p l e m e n t e d o r are being c o n s i d e r e d to reduce the f o r m a t i o n of A O X a n d c h l o r i n a t e d d i o x i n s from c h e m i c a l p u l p b l e a c h i n g o p e r a t i o n s . T h e r e are also possibilities for t r e a t m e n t of effluents with m i c r o o r g a n i s m s a n d enzymes to r e m o v e o r d e c h l o r i n a t e o r g a n i c material. E a c h o p t i o n discussed has i n h e r e n t a d v a n t a g e s a n d d i s a d v a n t a g e s with r e g a r d to c a p i t a l costs, o p e r a t i n g costs, ease of retrofit, f a b r i c a t i o n a n d i n s t a l l a t i o n time. I m p a c t on o t h e r mill unit o p e r a t i o n s is also c o n s i d e r e d in c h o o s i n g the best options. M a n y factors h a v e to be c o n s i d e r e d in c h o o s i n g an effective a n d e c o n o m i c a l b l e a c h i n g / t r e a t m e n t process t h a t meets all the e n v i r o n m e n t a l guidelines. It a p p e a r s t h a t the o n l y l o n g - t e r m s o l u t i o n will p r o b a b l y be to d e v e l o p the t e c h n o l o g y which will allow mills to o p e r a t e with zero effluent.

Acknowledgements We dedicate this article to Shri L.M. Thapar (Chairman, Thapar Group of Industries) on his 65th Birthday, thanking him for his never-failing encouragement. We also thank Dr. M.P. Kapoor, Director, Thapar Corporate R&D Centre, for his encouragement, Dr. M.B. Jauhari, General Manager (R&D), Ballarpur Industries Limited, for helpful discussions and Shri S.S. Gill for excellent typing of the manuscript.

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Hemicellulases in the Bleaching of Chemical Pulps A. Suurn/ikki, M. Tenkanen, J. Buchert, and L. Viikari VTT Biotechnology and Food Research P.O. Box 1501 FIN-02044 VTT Finland

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Pulping and Bleaching . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Wood Hemicelluloses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Chemical Pulping Processes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Bleaching Processes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Modification of Carbohydrates During Pulping . . . . . . . . . . . . . . . . . . . . . 3 Hemicellulases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Xylanases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Mannanases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Other Hemicellulases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Enzymatic Accessibility of Pulp Hemicelluloses . . . . . . . . . . . . . . . . . . . . . . 4 Hemicellulases in the Bleaching of Kraft Pulps . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Proposed Mechanisms of Hemicellulase-Aided Bleaching . . . . . . . . . . . . . . . . 4.2 Analysis of the Enzymatic Action in Pulp . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Factors Affecting the Enzymatic Action . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Effects of Different Hemicellulases on Pulp Bleachability . . . . . . . . . . . . . . . . 4.5 Action of Enzymes in Pulps Produced by Sulfate Cooking Methods . . . . . . . . . 4.6 Action of Enzymes in Sulphite Pulps . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7 Enzymes in Different Bleaching Sequences . . . . . . . . . . . . . . . . . . . . . . . . . 4.8 Impact of Enzymes on Pulp Yield and Quality . . . . . . . . . . . . . . . . . . . . . . 5 Industrial Enzyme-aided Bleaching . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Industrial Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Installation of the Enzymatic Step . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3 Environmental Impacts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4 Economics of the Enzymatic Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

262 263 263 264 265 266 267 267 269 270 272 272 272 274 274 276 277 277 278 279 280 280 280 282 282 283 284

Hemicellulase-aided bleaching is the first full-scale biotechnical application in the pulp and paper industry which truly exploits the unique specificity and safety of biocatalysts. Hemicellulases are used to modify the structure of xylan and glucomannan in pulp fibers in order to enhance the chemical delignification. This technology can be combined with various types of kraft pulping processes and bleaching sequences. The aims of the enzymatic treatment depend on the actual mill conditions, and may be related to environmental demands, reduction of chemical costs, or maintenance or even improvement of product quality. The technology is applied on the mill scale in several countries. This review describes the principles of the enzyme-aided bleaching, the composition of the fiber substrates, the basic enzymology involved, and the present knowledge of the mechanisms of the action of enzymes, as well as the practical results and advantages obtained on the laboratory and industrial scale.

Advances in BiochemicalEnNneering/ Biotechnology,Vol. 57 Managing FAitor:T. Scheper 9 Springer-VerlagBerlin Heidelberg 1997

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1 Introduction

During the past five years, the technology of chemical pulp bleaching has entered a period of extremely rapid change driven by growing concern for the environment. Environmental consciousness is evident not only in the market place, but also in increasingly stringent government regulation of the industry's waste streams, processes and products. Stringent regulation of chlorinated and poorly biodegradable unchlorinated organic compounds in mill effluents is rapidly becoming a universal rule. The industry has assessed various technologies in order to identify means for achieving an effluent level lower than 0.2 kg of AOX per ton of pulps. This target has already been achieved and even exceeded in several mills, e.g. in Scandinavia, using extended cooking, oxygen delignification and various ECF (Elemental Chlorine Free) or TCF (Totally Chlorine Free) bleaching chemicals. Among the new bleaching agents, biotechnical methods have also been developed and used on the mill scale. In Europe, the first signs of growing consumer awareness appeared in late 1980s, leading to an almost desperate search for new methods to reduce environmental impacts. The resulting changes in bleaching technologies occurred so rapidly that they may be considered revolutionary. In this transitional period, the hemicellulase-aided bleaching process was adopted on the industrial scale. Enzymes are readily biodegradable, and offer an environmentally safe method for improving pulp bleaching. The idea of using hemicellulolytic enzymes for increasing the bleachability of chemical pulps was introduced in the 1980s [1, 2]. Previous reports of applications of hemicellulases included total hydrolysis of hemicellulosic raw materials for production of monomeric sugars [3] and the removal of hemicelluloses from chemical pulps to produce dissolving pulps [4]. The concept of enzyme-aided bleaching was based on the realization that limited hydrolysis of hemicellulose in pulps by hemicellulases, mainly xylanases, increases the extractability of lignin from the kraft pulps in the subsequent bleaching sequences. The xylanase pretreatment permits the use of lower chlorine charges during the bleaching of kraft pulps, the bleach-boosting effect being associated with reduced chloroorganic discharges. The first mill trials were conducted in Finland in 1988 [5], a remarkably short time after the first reports. As effective and easily applicable process aids, xylanases are nowadays used in several mills prior to chlorine and non-chlorine bleaching sequences of kraft pulps (reviewed in [6]). The hemicellulase treatment, together with a chemical extraction, leads to a significant reduction of the residual lignin content in the pulps. However, due to the indirect mode of action, the effect of hemicellulase-aided bleaching is limited. The development of a directly delignifying biotechnical method is therefore still the major target for several research groups. The reaction mechanisms of lignin-degrading enzymes on fiber-bound lignin are, however, more difficult to control, and delignification results with these enzymes have been rather poor. Recently, however, very promising results have been obtained using

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a new concept based on the ability of mediators, oxidized by laccase, to degrade lignin specifically [7, 8]. The hemicellulase-aided bleaching method represents a new type of generic technology, already adopted by the pulp and paper industry. Hemicellulases are the first group of specific enzymes to be used in a large-scale application in this industrial area. The approach also represents a truly sustainable technology. Moreover, the technology was shown to be economically competitive a prerequisite not always fulfilled by biotechnical applications. The interest in this type of new technology is also shown by the increasing number of papers published during recent years describing numerous xylanases from new sources, as well as bleaching results obtained using various hemicellulases, pulps and bleaching sequences. Several reviews have also been published [6, 9, 10, 11, 12]. The present review summarizes the major achievements in the field.

2 Pulping and Bleaching 2.1 Wood Hemicelluloses Wood is mainly composed of cellulose, hemicellulose and lignin. Cellulose is the most abundant of the components, generally representing 4 ~ 4 5 % of the wood dry weight [13]. Depending on the wood species, about 20-30% of wood dry weight is hemicellulose and about 15 25% lignin. The two main hemicelluloses in wood are xylans and glucomannans (Table 1). The softwood xylan has a backbone of arabino-4-O-methylglucuronoxylan, which is composed of D-xylopyranose units connected via [3-1,4-glucosidic linkages. The average molar ratio of arabinose: 4-O-methylglucuronic acid: xylose sugar units in softwood xylan is 1.3:2:10 [13]. Hardwood xylan contains 4-O-methylglucuronic acid and acetyl side groups. Methylglucuronic acids are linked to the xylan backbone by 13-1,2-glucosidic bonds, and the acetic acids are esterified at the carbon 2 and/or 3 hydroxyl group. The backbone of softwood glucomannan is composed of [3-1,4-1inked D-glucopyranose and D-mannopyranose units and is partially substituted by ~-galactose and acetyl units [13]. The two fractions of softwood glucomannan differ in their average molar ratio of galactose: glucose: mannose. The low galactose content fraction has a ratio of 0.1:1:4 and is generally called glucomannan [13]. This is the main fraction of glucomannan in softwoods. The corresponding ratio in the high-galactose content fraction, galactoglucomannan, is 1:1:3. Hardwoods also contain a small amount of glucomannan. The ratio of glucose: mannose in this glucomannan varies between 1 : 1 and 1:2, depending on the wood species. In addition to xylan and glucomannan, both softwoods and hardwoods contain minor amounts of other hemicelluloses such as pectin, galactan and arabinan [-13].

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Table 1. Hemicellulose in softwood and hardwood [13]

Wood species

Component

Content % of dry weight

Pine

Xylan Glucomannan Xylan Glucomannan

5-11 14 20 2~30 1~4

Birch

T h e d i s t r i b u t i o n of hemicelluloses in w o o d fibers varies d e p e n d i n g o n the w o o d species a n d g r o w i n g c o n d i t i o n s [14]. H o w e v e r , in softwoods, the o u t e r layer of s e c o n d a r y wall ($2) a n d the i n n e r m o s t fiber wall layer, the w a r t y l a y e r (W), are generally richer in xylan t h a n the m i d d l e of the s e c o n d a r y wall. T h e g l u c o m a n n a n c o n t e n t of s o f t w o o d fibers increases steadily f r o m the o u t e r p a r t s to the inner p a r t s of cell walls. In h a r d w o o d s , the o u t e r m o s t layers of s e c o n d a r y fiber wall are rich in xylan, whereas the c o n t e n t of g l u c o m a n n a n r e m a i n s low a n d c o n s t a n t t h r o u g h the fiber walls.

2.2 Chemical Pulping Processes The m a i n a i m in chemical p u l p i n g is to s e p a r a t e the w o o d fibers from each o t h e r in o r d e r to r e n d e r t h e m suitable for further i n d u s t r i a l processing. T h e delignific a t i o n of w o o d y m a t e r i a l can be c a r r i e d out, for e x a m p l e , b y sulfate (kraft) o r

Table 2. Annual production of wood-based kraft and sulfite pulp in the

world. Estimate for the year 1996 (FAO survey, 1995) Region

Pulping method

Pulp productiona 10 6 tb/a

Northern America

Kraft Sulfite Kraft Sulfite Kraft Sulfite Kraft Sulfite Kraft Sulfite Kraft Sulfite Kraft Sulfite

62.7 8.2 13.3 0.9 8.1 2.4 9.1 0.9 6.1 0.2 1.2 0.1 0.9 0

Scandinavia Europe Latin America Asia, developing Oceania Africa, developing

a Total production (unbleached, bleached) b Metric tons of air dry pulp

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sulfite processes [13]. Today, the predominant pulping method is the kraft process (Table 2). However, for environmental reasons, sulfite pulping is preferred in Germany and also in several other countries. In sulfate, i.e. kraft pulping, the lignified middle lamella located between the wood fibers is removed in highly alkaline conditions and at a high temperature [13]. Recently, modified and totally new sulfate pulping processes using altered cooking conditions such as a relatively even alkali profile and decreased cooking temperatures have been introduced [15]. The main aim of the pulping methods such as modified continuous cooking (MCC), superbatch cooking and isothermal cooking (ITC) is to produce more easily bleachable pulps having both relatively low lignin content and acceptable strength properties.

2.3 Bleaching Processes In kraft pulping, about 90% of wood lignin is solubilized during the cooking process [13]. The remaining 10% of lignin is mainly responsible for the brown colour of the kraft pulp and unbleached paper. The primary goal of bleaching is to remove the residual lignin from the pulp as selectively as possible without degrading the pulp carbohydrates, especially cellulose, which would lead to a decrease in viscosity. Traditionally, the bleaching of chemical pulps has been carried out with elemental chlorine and chlorine dioxide. However, the chlorinated organic compounds formed during chlorine bleaching have attracted negative attention during recent years. Public concern, together with tightened environmental regulations, have driven the pulp and paper industry to seek out and utilize alternative bleaching processes. Reduction in bleach plant effluents can be achieved by reducing the lignin content of pulp prior to bleaching by modified cooking procedures or oxygen delignification, or by replacing the elemental chlorine used in the bleaching process with other chemicals such as chlorine dioxide, ozone, oxygen, peroxide and/or peroxyacids [15, 16]. Worldwide, chlorine gas has traditionally been the main chemical used in pulp bleaching, but the alternative bleaching sequences, e.g. oxidizing agentbased totally chlorine-free (TCF) and especially the chlorine dioxide-based elemental chlorine-free (ECF) sequences, are increasingly used in industrial pulp bleaching [15]. Compared with elemental chlorine or even with chlorine dioxide, the oxygen-based chemicals are less effective or less selective in reacting with pulp lignin [13]. To obtain a fully bleached pulp without elemental chlorine, the lignin content of the pulp entering the bleaching process should be as low as possible. Oxygen delignification is commonly used as a prebleaching stage prior to the ECF and TCF bleaching sequences. A reduction of lignin content by about 50% can be achieved using oxygen, with relatively low loss of carbohydrate yield and without impairing the strength properties of pulp [13, 17].

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2.4 Modification of Carbohydrates During Pulping Wood cellulose is rather resistant to the harsh conditions used in chemical pulp production, because of its crystallinity and linearity. However, hemicellulose components of the wood are heavily modified during pulping processes. At the beginning of the conventional sulfate process, i.e. kraft cooking, xylan in wood is partly solubilized by the alkaline cooking liquid, and many of the side groups and acetic acid residues are cleaved off [18, 19]. Already in the early phases of the kraft cook, the methylglucuronic acid side groups in both residual and solubilized xylan are almost completely converted to hexenuronic acid [20, 213. As the alkali concentration decreases towards the end of the kraft cook, dissolved xylan tends to readsorb on the surface of cellulose microfibrils [22-25]. In addition to free xylan chains, dissolved lignin and covalently bound xylan and lignin are also believed to relocate to fiber surfaces [26, 27], resulting in relatively high amounts of lignin on these surfaces [28-30]. The amount of xylan readsorbed during the cooking depends on the wood species used in pulping. High amounts of xylan have been found to locate on the surface of birch kraft fibers, probably partly due to readsorption, whereas in pine kraft fibers the concentration of xylan on the fiber surfaces has not been observed to be higher than in the whole fibers [31]. A large part of wood glucomannan is also dissolved at the beginning of the kraft cook, but owing to their instability in alkali, the solubilized polymers are completely degraded in the pulping liquor [23, 32, 33]. As a result of the solubilization of hemicelluloses during cooking, the distribution and content of xylan and glucomannan in kraft pulp fibers differs from that in the native wood fibers [14]. In softwood kraft fibers, the xylan concentration is generally higher in outer layers, and glucomannan is more concentrated in the middle layers of the fiber. However, due to different analysis methods, variations in the distribution of polysaccharides in softwood kraft fibers have been reported [34-38]. There is, however, general agreement that the outer surface layer of hardwood kraft fibers is rich in xylan. Recently, several modified kraft pulping methods as well as totally new sulfate pulping methods have been introduced [15]. In the pulps produced by these methods, little or no reprecipitation of xylan and lignin is expected to occur owing to the relatively constant alkali concentration throughout the cooking process. Consequently, the composition of the outer surfaces of the pulp fibers is probably different from that of the conventional kraft pulp fibers. In sulfite cooking, hemicellulose is extensively solubilized to monomeric and oligomeric compounds, and no reprecipitation occurs [39]. Thus, the distribution of hemicellulose is relatively constant across the pulp fibers [40]. The arabinose side-groups of softwood xylan are completely removed in acidic sulfite cooking [41], although some acetyl groups are present in sulfite pulps.

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3 Hemicellulases Owing to the complex structure of hemicelluloses, several different enzymes are needed for their enzymatic degradation or modification. The two main glycanases depolymerizing the hemicellulose backbone are endo-l,4-~-Dxylanase and endo-l,4-~-D-mannanase. Xylanases and mannanases are produced by many species of bacteria and fungi (reviewed in [42 46]). Small oligosaccharides are further hydrolyzed by 1,4-13-D-xylosidase, 1,4-13-D-mannosidase and 1,4-[3-D-glucosidase. The side-groups are removed by 1,3-~-c-arabinosidase, 1,2-a-o-glucuronidase and 1,6-a-D-galactosidase. Esterified side-groups are liberated by acetylxylan esterase and acetylgalactoglucomannan esterase.

3.1 Xylanases Since xylan is the most abundant of the hemicelluloses in pulps, a major part of the published work on hemicellulases deals with the properties, mode of action and applications of xylanases (reviewed in [42,45-48]). EndoXylanases (1,4-13-D-xylan xylanohydrolases, EC 3.2.1.8) catalyze the random hydrolysis of 1,4-13-D-xylosidic linkages in xylans (Fig. 1). Most xylanases are rather small proteins (molecular mass around 20 kDa) with a basic isoelectric point (pI 8 10). Another group of xylanases has also been identified. These are somewhat larger (molecular mass > 40 kDa), with acidic isoelectric point (pI 3-5) [47]. The grouping based on the physico-chemical properties correlates well with the classification of xylanases into glycosyl hydrolase families 10 and 11 (previously F and G), based on amino acid similarities [49]. The xylanases belonging to the two xylanase groups so far identified also differ from each other with respect to their catalytic properties [50]. The family 10 xylanases, with high Mr and low pI, seem to exhibit greater catalytic versatility than those of family 11, with low Mr and high pI. Thus, they are typically able to hydrolyze highly substituted xylans more efficiently. Most of the xylanases characterized are able to hydrolyze different types of xylans, showing differences only in the spectrum of end products, The main products formed from the hydrolysis of xylans are xylobiose, xylotriose and substituted oligomers of two to five xylosyl residues. The chain length and the structure of the substituted products depend on the mode of action of the individual xylanase. Some xylanases, however, show rather strict substrate specificity. A unique xylanase which requires a glucuronic acid substituent in the xylan backbone is produced by Bacillus subtilis [51]. At the other extreme, xylanase produced by Talaromyces emersonii requires at least 24 unsubstituted xylose residues for its action [52]. The three-dimensional structures of several low-molecular-mass xylanases have recently been determined [53-56]. The structure of the Trichoderma reesei pI 9 xylanase

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MeGIcA Ac Ac Ac I~-I

Biotechnology in the Pulp and Paper Industry

Biotechnology in the Pulp and Paper Industry (Progress in Biotechnology)

Biotechnology for Pulp and Paper Processing