Microbial Pentose Utilization: Current Applications in Biotechnology (Progress in Industrial Microbiology)

Microbial Pentose Utilization Current Applications in Biotechnology

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edited by M.J. Bull (1st reprint 1983) edited by M.J. Bull edited by M.J. Bull edited by M.E. Bushell Microbial Polysaccharides, edited by M.E. Bushell Modem Applications of Traditional Biotechnologies, edited by M.E Bushell Innovations in Biotechnologie, edited by E.H. Houwink and R.R. van der Meer Statistical Aspects of the Microbiological Analysis of Foods, by B. Jarvis Moulds and Filamentous Fungi in Technical Microbiology, by O. Fassatiovfi Micro-organisms in the Production of Food, edited by M.R. Adams Biotechnology of Animo Acid Production; edited by K. Aida, I. Chibata, K. Nakayama, K. Takinama and H. Yamada Computers in Fermentation Technology, edited by M.E. Bushell Rapid Methods in Food Microbiology, edited by M.R. Adams and C.F.A. Hope Bioactive Metabolites from Microorganisms, edited by M.E. Bushell and U. Grfife Micromycetes in Foodstuffs and Feedstuffs; edited by Z. Jesenskfi Aspergillus: 50 years on; edited by S.D. Martinelli and J.R. Kinghorn Bioactive Secondary Metabolites of Microorganisms, edited by V. Betina Techniques in Applied Microbiology, edited by B. Sikyta Biotransformations: Microbial Degradation of Health Risk Compounds, edited by V.P. Singh Microbial Pentose Utilization. Current Applications in Biotechnology, by A. Singh and P. Mishra

Microbial Pentose Ut'~ilizatJon Current Applications in Biotechnology

AJAY SINGH Microbial Biotechnology Laboratory, University of Waterloo, Waterloo, Ontario N2L 3G1, Canada

PRASHANT MISHRA Department of Biochemical Engineering and Biotechnology, Indian Institute of Technology, Delhi, Hauz Khas, New Delhi 110016, India

progress in industrial microbiology

ELSEVIER

Amsterdam

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York - Oxford

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Tokyo

1995

ELSEVIER SCIENCE B.V. Sara Burgerhartstraat 25 P.O. Box 211, 1000 AE Amsterdam, The Netherlands

ISBN 0-444-82039-6 (Vol. 33) ISBN 0-444-41668-8 (Series) 9 1995 Elsevier Science B.V. All rights reserved

No part of this publication may be reproduced, stored in a retrieval system or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, without the prior written permission of the publisher, Elsevier Science B.V., Copyright & Permissions Department, P.O. Box 521, 1000 AM Amsterdam, The Netherlands. Special regulations for readers in the U.S.A. - This publication has been registered with the Copyright Clearance Center Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923. Information can be obtained from the CCC about conditions under which photocopies of parts of this publication may be made in the U.S.A. All other copyright questions, including photocopying outside of the USA, should be referred to the copyright owner, Elsevier Science B.V., unless otherwise specified. No responsibility is assumed by the publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. This book is printed on acid-free paper. Printed in The Netherlands

Preface Microbial utilization of the inexhaustible lignocellulosic biomass for production of liquid fuels and protein rich food and fodder offers an attractive approach to meet energy and food demands. Whilst hemicellulose derived sugars consist of appreciable amounts of pentose sugars, their microbial utilization is a major limiting factor in the development of an economically viable process. In the past decade considerable progress has been made in our understanding of the metabolic pathways, genetics and molecular biology of the pentose fermenting microorganisms. In addition, recent developments in fermentation technology have led to advanced processes for bioconversion of lignocellulosic biomass to various industrially important products. Our objective of putting together fundamental aspects of pentose utilization and its biotechnological implications was to bring together, in one place, biological and engineering aspects of pentose utilization tO develop a clear understanding of pentose fermentation technology needed for industrial processes. In this book, chapter 1 is an introduction to the availability of pentose sugars from agricultural and forestry residues and their potential uses. Chapters 2 and 3 are concerned with the biosynthesis and biodegradation of hemicelluloses and extraction of pentose sugars. Since uptake of these sugars and subsequent metabolism is the initial step in their utilization, chapters 4 and 5 deal with the pentose uptake, their metabolism in various organisms and regulation of uptake and metabolic network. Chapters 6,7 and 8 are concerned with kinetics of growth and product formation and fermentation technologies of ethanol, acetone-butanol and butanediol production. Besides these solvents, organic acids, xylitol and SCP/SCO are also attractive end products of pentose metabolism hence chapters 9, 10 and 11 have been devoted to production of respective end products. Since many solvents and organic acids produced during microbial fermentation are known to exert inhibitory effects, chapter 12 describes microbial tolerance to solvents and organic acids. It is well understood that for the development of any successful fermentation technology selection of microbial strain and genetic engineering plays a crucial role, hence chapter 13 has been devoted to current approaches employed for the improvement of pentose fermenting microbial strains. Finally, chapter 14 describes process evaluation and bioengineering aspects of pentose fermentation. We feel that this book should prove to be useful to graduate and postgraduate students of microbiology, biochemistry and biotechnology, scientists and engineers both in academia and industry. We thank our colleagues Drs K. Hayashi, O.P. Ward, P. Ghose, S. Chand, V.S. Bisaria, K. Tokuyasu, M.A. Tariq, N. Izawa, R.C. Kuhad and P.K.R. Kumar for their help and encouragement. We are also thankful to the publishers for their patience, willing help and cooperation. We would like to acknowledge our sincere thanks to our wives Pratima and Simi for their endurance and constant support without which it would have been a distant dream. Ajay Singh Prashant Mishra

This Page Intentionally Left Blank

vii

CONTENTS

I

Overview of Problems and Potential

1 2 3 3.1 3.2 4 4.1 4.2 4.3 4.4 4.5 5 6

II

Energy Demand Lignocellulosic Resources Global Availability of Lignocellulosic Materials Agricultural Residues Industrial Residues Nature of Lignocellulosic Materials Structure of Plant Cell Walls Cellulose Hemicellulose Lignin Extraneous Materials Applications in Biotechnology References

1 3 6 6 8 9 12 13 16 20 22 23 26

Biosynthesis and Biodegradation of Hemicelluloses

1 2 2.1 2.2 2.3 3 3.1 3.2 3.3 4 5 5.1 5.2 5.3 5.4 6

Introduction Biosynthesis of Hemicellulosic Substances Synthesis of Sugar Nucleotides Interconversion of Sugar Nucleotides Polymerization Chemistry of Hemicelluloses Xylans Mannans and Glucomannans Galactans and Arabinogalactans Enzymatic Analysis of the Structure Biodegradation of Hemicelluloses D-Xylanases L-Arabinnanase D-Galatanase D-Mannanase References

33 33 34 38 40 43 46 50 51 52 54 56 60 61 63 64

viii III

Extraction of Pentosans from Lignocellulosic Materials

1 2 3 3.1 3.2 4 4.1 4.2 4.3 4.4 5 6

IV

71 71 76 77 78 79 79 84 90 90 91 92

Microbial Uptake of Pentoses

1 2 3 4 5 5.1 5.2 6 7

V

Introduction Enzymatic Treatment Physical Treatment Milling Irradiation Chemical Treatment Alkali Acids Gases Oxidizing Agents Thermal Treatment References

Introduction Mode of Sugar Uptake Pentose Uptake in Yeast Pentose Uptake in Bacteria Regulation of Pentose Uptake Yeasts Bacteria Genetic Studies on Pentose Uptake References

99 100 101 105 109 109 110 113 115

Microbial Metabolism of Pentoses

1 2 2.1 2.1.1 2.1.2 2.1.3 2.2

Introduction Metabolism of D-Xylose Conversion of D-Xylose to D-Xylulose-5-Phosphate Oxidative Reductive Pathway Xylose Isomerase Pathway Phosophorylation Conversion of D-Xylulose-5-Phosphate to Various End Products

119 119 120 120 124 126 127

3 3.1 3.2 4 5

VI

130 130 133 136 139

Microbial Production of Ethanol

1 2 2.1 2.2 2.3 2.4 3 3.1 3.2 3.3 3.4 4 5 6 7 8 8.1 8.2 8.3 8.4 8.5 8.6 8.7 9

VII

Regulation of D-Xylose Metabolism Genetic Regulation Oxygenation and Cofactor Regulation Metabolism of L-Arabinose References

Introduction Microorganisms used for Ethanol Production Yeast Filamentous Fungi Thermophilic Bacteria Mesophilic Bacteria Kinetics of Growth and Product Formation Yeast Filamentous Fungi Thermophilic Bacteria Mesophilic Bacteria Simultaneous Pentose Isomerization and Fermentation Whole Cell Immobilization Coculture Performance on Natural Substrates Factors Affecting Ethanol Production pH Temperature Nutrition Oxygenation Lipids Metabolic Inhibitors Inhibitors Present in Lignocellulosic Hydrolysate References

147 149 151 153 154 155 156 156 161 163 167 168 170 173 174 178 178 179 180 182 185 186 187 188

Microbial Production of Acetone and Butanol

Introduction Microorganisms used for Acetone-Butanol Production

197 198

3 4 5 5.1 5.2 5.3 5.4 5.5 5.6 5.7 6

VIII

201 206 209 209 210 211 212 213 215 216 218

Microbial Production of 2,3-Butanediol 1 2 3 4 5 6 6.1 6.2 6.3 6.4 6.5 6.6 6.7 7

IX

Kinetics of Growth and Product Formation Performance on Natural Substrates Factors affecting Acetone and Butanol Production pH and Temperature Repeated Subculturing Production of Bacteriocin Nutrition Oxygenation Continuous Culture Whole Cell Immobilization References

Introduction Microorganisms used for 2,3-Butanediol Production Kinetics of Growth and Product Formation Reactor Systems Performance on Natural Substrates Factors affecting 2,3-Butanediol Production pH Temperature Aeration Water Activity Inoculum Nutrient Supplementation Inhibitors References

221 223 226 231 232 234 235 236 237 239 240 240 242 244

Microbial Production of Organic Acids

1 2 2.1 2.2 3 3.1 3.2 4 4.1

Introduction Acetic Acid Microorganisms used for Acetic Acid Production Kinetics of Product Formation Lactic Acid Microorganism used for Lactic acid Production Kinetics of Product Formation Citric Acid Microorganisms used for Citric Acid Production

249 249 250 251 255 256 257 258 258

4.2 5 5.1 5.2 6 6.1 6.2 7 7.1 7.2 8 9

X

259 261 261 262 262 263 263 266 266 267 268 269

Microbial Production of Xylitol 1 2 3 3.1 3.2 4 4.1 4.2 4.3 4.4 4.5 5

Xl

Kinetics of Product Formation Propionic Acid Microorganisms used for Propionic Acid Production Kinetics of Product Formation Itaconic Acid Microorganisms used for Itaconic Acid Production Kinetics of Product Formation Fumaric Acid Microorganisms used for Fumaric Acid Production Kinetics of Product Formation Mixed Acid Fermentation References

Introduction Microorganisms used for Xylitol Production Kinetics of Growth and Product Formation Yeast Bacteria Factors Affecting Xylitol Production pH and Temperature Oxygenation Magnesium Supplementation Nitrogen Sources and Organic Nutrients Methanol Supplementation References

273 274 276 276 287 289 289 291 293 294 296 297

Microbial Production of Single Cell Protein (SCP) and Single Cell Oil (SCO) 1 2 3 4

Xll

Introduction Microorganisms used for SCP Production Microorganisms used for SCO Production References

301 303 310 314

Microbial Tolerance to Solvents and Organic Acids 1 2

Introduction Effect of Solvents and Organic Acids on Cellular Physiology of Microorganisms

317 318

xii

3.1 3.2 4

6

Xlll

332 336 342

Introduction Screening and Mutagenesis Genetic Recombination Hybridization Protoplast Fusion Gene Cloning, Expression and Characterization Yeasts Bacteria References

351 352 356 356 357 359 359 361 366

Process Evaluation and Bioengineering 1 2 3 4 5 6 7

Index

327 327 331

Genetic Improvement of Pentose Fermenting Microorganisms 1 2 3 3.1 3.2 4 4.1 4.2 5

XlV

Adaptive Modifications in Microorganisms Leading to Solvent Tolerance Modification of lipid Composition Induction of Stress Proteins Manipulation of Membrane Lipid Composition and Tolerance to Solvents Genetic Basis of Tolerance to Solvents and Organic acids References

Introduction Pretreatment of Substrate Fermentation Design Downstream Processing Economic Evaluation Future Prospects References

371 372 374 382 387 390 392

397

Overview of Problems and Potential

1

ENERGY D E M A N D

The oil crisis of the mid 1970s and widespread recognition of the finite nature of the world's petroleum resources has led to the examination of alternative sources of materials and energy. The demand for energy is increasing because of more productive economics and growing population. A significant part of this demand will be met by increased consumption of petroleum. However, oil is becoming increasingly difficult to find and recover, and its demand will shortly catch up with supply, which will inevitably result in the increased oil prices. Oil is not only the major source of energy, it also represents the major source of raw material for chemical industries. Therefore, fuel prices will exert a great influence on future feedstock cost. Recently, the price of oil has varied considerably and is still unstable. Thus considerable pressure is building up to change the feedstocks for chemical industries and to stabilize the sources and costs of raw materials [1]. A long term practical solution to this problem is to direct technologies toward the conversion of a major source of continuously renewable, nonfossil carbon, such as organic wastes and biomass - which consists of all growing organic matter, such as plants, trees, grasses, and algae - to produce chemicals that were an attractive alternative to oil. Although in mid 1980s, this approach has dwindled with the drop in oil prices, the need to intensify the biomass utilization activities in future remains because of the finite nature of oil resources. On a worldwide basis, it has been estimated that about 146X109 tons of carbon are fixed annually [2]. Assuming lignocellulosic materials to be 50% carbon and to have a heat of combustion of 8500 Btu/Ib on an ash-free ovendried basis, 5X10 ~8 Btu are stored annually by photosynthesis [3]. On the basis of these data, it can be assumed that only 2 years are required to photosynthetically produce an equivalent in carbon to provide current available categories of natural gas, crude oil and syncrude from oil, or 8-20 years for an equivalent in fixed carbon to the estimated total remaining recoverable amounts of the four fuel categories [2]. The main feedstock or primary precursor for the chemical industry is crude oil. However, a large number of precursors can be produced from other raw materials such 1

as coal, natural gas, and pyrolysis and distillation of wood. A large number of these precursors can also be synthesized from solvents and chemicals produced by fermentation of biomass using microorganisms. Potential substrates include sugar containing materials, starchy crops and lignocellulosic materials (Table 1). Biomass fermentation route for the production of chemical feedstock is becoming increasingly attractive as biomass production costs are not as tightly bound to the energy costs. This trend will continue in the future. However, in their present state, most of these processes require subsidy or special circumstances to compete with more efficient chemical processes. Nevertheless, the microbial conversion of biomass to chemicals and solvents is a versatile process which can be used in various applications for replacing or improving petroleum products, treating wastes, and reducing pollution. Petroleum replacement can be in relation to neat fuel, fuel additives, or raw materials.

Table 1 Potential raw materials for bioconversion to chemicals, solvents, and animal feed Sugar containing

Starch containing

Lignocellulosic

Molasses

Cereal grains

Agricultural residues

Whey

corn

Forest residues

Fruit juices

sorghum

Wood sulfite waste

Sweet sorghum

barley

Fruit/vegetable waste

Sugarbeet

Wheat bran

Waste paper

Sugarcane

Root tubers

Municipal solid waste

A person require about 500 g of food (70 g protein, 80 g fat, 350 g carbohydrates) per day to maintain himself at his energy output of 130 watts [4]. Therefore, the world population of approximately 3.5Xl 08 requires more than 5X10 e tons of food per year and the demand is steadily increasing. Since much of the plant is inedible, unavailable, or eaten by animals, food shortages are developing. In addition to the source of chemicals,

we can look to the lignocellulosic biomass as a source of food and feed, and as a substrate for single cell protein and lipid production using appropriate microorganisms.

2

LIGNOCELLULOSIC RESOURCES

Lignocelluloses are the most abundant renewable natural materials present on the earth. Wood and agricultural by-products are virtually inexhaustive based on the photosynthetic processes. They account for more than 60% of the total biomass. The net productivity of the dry biomass due to photosynthesis by plants on earth has been estimated to be 155 billion tons per year [5]. About two-thirds of the biomass production occurs on land, and about one third occurs in the oceans. Most terrestrial plant materials occur in forest (65%), with a bit more than 15% generated in grasslands and cultivated lands. About three-quarters of the total biomass generated on the cultivated lands and grasslands is unutilized and hence is residue or waste [6]. Crops, trees, and other plants that are grown for food and other economic purposes also generate millions of tons of lignocellulosic waste. One great disadvantage lies in the fact that since these wastes are generated in a solid form and are spread thinly over the land surface, the cost of transportation is high, making this a prohibitory factor [7]. About 1.25% of the total land biomass is projected to be eventually for human food, with about 9% being lost during the processing operations and the rest accounting for the magnitude of availabilty of lignocellulosic wastes. About 40% of the various cultivated crops consists of marginal foods and feed. A large proportion of these materials are carbohydrate wastes produced mostly in the agriculture, forest and food industries. There are several convenient carbohydrate waste streams generated from manufacturing processes such as those from pulp and paper processing units. Currently the pulping and paper manufacturing industries produce a large amount of carbohydrate wastes which are estimated to be over 200 million tons [8]. About 1400 million tons of straw and cornstalks are produced annually worldwide. The potential utilization of the huge quantities of such wastes as a renewable carbon source is of great importance. A wide range and variety of inedible agricultural, forestry and industrial wastes are available (Table 2). The type and availability of the lignocellulosic wastes in any particular geographic region depends on the climatic and environmental factors, use and disuse,

culture, and type and nature of the regional technology. Thus, while rice straw is more prevalent in the far east and southern Asia, wheat straw and maize by-products are abundant in North America and Europe. In United Kingdom considerable amounts of wheat straw are disposed by burning [9].

Table 2 Total carbohydrate content of various waste streams Waste

Total carbohydrates (% dry weight)

Agricultural Stems

50-80

Leaves

80-95

Fibres

90-98

Forestry

60-70

Urban

50-60

Spent sulfite liquor

30-35

Manure

20-25

Waste paper

80-95

Waste fibres

70-90

Estimates for the availability of biomass in United States from waste materials, forestry and agricultural crops have been projected to be 8.56 to 10.8X108 tons per year [10]. Forest and agricultural residues necessary to maintain soil fertility are not included in estimates for total or collectable wastes. Wheat and soybean straw, and corn stover are each produced in excess of 100 million tons per year, while sorghum stover and oat straw are produced in amounts ranging from 15 to 56 million tons per year in the United States [11]. Approximately 200 million tons of agricultural cellulosic wastes are produced

each year in the United States [12-18]. Much of these unutilized resources is disposed by burning, a method which has been increasingly the subject of criticism because of the resultant air pollution. In many of the southeast Asian countries, large fractions of the available straw are alternatively used for thatching of the roofs, mulching and even cattle feed. About half of the total production of plant residues from agricultural and industrial processes remains unused, but much of these materials, if not burnt, is shredded and/or composted for landfills or improvement of soil types. The forest production (tons per year) available for energy production in United States is estimated at 182 to 245 in the late 1970s, and 280 to 560 in the year 2000 [10]. The higher estimates for forest production in the year 2000 is based on a greater role for high-productivity-energy devoted silviculture. The availabilty of agricultural crops in United States is estimated at 69X106 tons per year for corn with process by-products used for animal feed, plus either an additional 0-88X106 tons of corn, or 0-414X106 tons of forage grasses. Utilization of sugar- and starch-rich agricultural crops for fuels and chemical production may be considered to compete with food production. It is generally considered to have a less favourable energy balance compared to lignocellulosic substrates, and has the greatest potential for unfavorable environmental impact [19]. The environmental impacts of solvent and chemical production could be: those resulting from process waste streams; those from the interplay between substrate production and land resource considerations such as soil fertility and erosion, and maintaining wildlife habitat and those relating to product utilization per se [10].The principal streams are airborne emissions, suspended solids and BOD in wastewater, and solids in the form of ash and insoluble salts, particularly arising from neutralization of acid. Land resource issues have sharply differing potential to be environmental problem depending on the chemical feedstock considered. Wood and food processing wastes, animal wastes and collected logging wastes have no significant potential. Crops and logging residues have some potentials if mismanaged, whereas grasses should have few significant adverse impacts for most applications. Other wood sources have high potential but theoretically can be managed. On the other hand, grain and sugar crops have the highest potential. Biomass has the potential to be an energy source that has few significant environmental problems. However, the expansion of bioenergy may still cause serious environmental damage because of poorly managed feedstock supplies and inadequately controlled conversion technologies. Further, some uncertainties remain about the long term effects of intensive biomass harvest on the soil fertility. Some agricultural crops are primarily used for animal feed in a few countries. For

example, over 80% of the total United States corn crop is used for feed, with 55-60% used domestically, whereas only 10% is used for human consumption [10]. The starch fraction of the corn crop can be used for ethanol production while residues and/or processing by-products still retain considerable feed value. Thus ethanol can be produced from corn ultimately used for animal feed, with relatively small incremental resource demands. Forage grasses represent agricultural crops with a more favourable energy balance and lower potential for unfavourable environmental impact in comparison to corn. Grass utilization for ethanol production makes use of the entire plant, and does not depend on by-product utilization. However, there is considerable uncertainty regarding the availability of agricultural land for energy crops in United States with between 0 and 26X 106 hactare available in the year 2000 [16]. The quantity of fermentable carbohydrates for ethanol production from lignocellulosic substrates appears to be large. Biomass supply is on average larger relative to oil consumption. Lignocellulosic substrates can be distinguished from starch- and sugar-rich substrates by their relatively high content of insoluble polymers containing 13-1inked glucose (cellulose) and pentoses (found in hemicelluloses). The difficulty in economically converting these components has been primarily responsible for the incomplete success in efforts to develop practical biological processes.

3

GLOBAL AVAILABILITY OF LIGNOCELLULOSIC MATERIALS

3.1

Agricultural residues

According to an estimate [20], about 2946 million tons cereal straw are produced per year in the world (Table 3). Significant quantities are disposed of by burning in most of the parts of the world, which clearly is a great loss of energy conserved through the process of biosynthesis by the green plants. Air pollution laws and the restriction of burning of straw, and increasing cost of animal feeds have revived interest in the utilization of low quality crop residues as a ruminant feed. Biodegradation and bioconversion of these straws using microorganisms would contribute to the production of directly palatable food and feed material. A number of pulse crops are cultivated in

various parts of the world and for every ton of the pulse that is harvested, two to three times of the inedible plant residue is available. These plant residues contain more nitrogen than those of cereal residues. Annually about 166 million tons of this residue are available worldwide. Maximum amount of pulse crop residues is produced in Asia followed by Central America and United States. Edible oil is one of the basic components of food consumed. After the oilseeds are harvested, significant quantities of the plant residues remain unutilized. For example, in the case of sunflower, the plant residues does not have any value other than serving as a fuel after burning. Annually, approximately 142 million tons of oilseed crop residues are available all over the world and would merit consideration for a number of diversified applications.

Table 3 Estimated global production of lignocellulosic wastes Wastes (million tons) Continent/ Country Africa

Cereal

Pulse

Oilseed

Plantation

165

9

11

34

1135

51

61

174

35

1

2

12

Central America

500

49

21

84

Europe

550

10

8

1

India

240

16

14

88

South America

153

37

10

147

United States

440

44

19

15

2946

166

142

548

Asia Australia

World

The fibrous residue, bagasse remains after extracting juice from the sugarcane

stalks is another potential substrate for bioconversion. In sugar mills, most bagasse is used as fuel. Other substrates like corn stover, stems of castor oil plant, leaves of mulberry, saw dust of Lantinus, milled dry cassava roots, corn stalks and cottonseed husk are available abundantly. About 1.5 tons of straw is produced per acre of cotton cultivated. Its high lignin content (about 25%) limits the value of cotton straw as a direct ruminant feed. Cotton ball Iocules, plentiful in cotton growing areas, is a promising substrate for bioconversion [21-24].

3.2

Industrial residues

Coffee pulp is a major by-product of the coffee industry, representing 28.7% of the coffee beans on a dry weight basis during the wet coffee processing method [25]. This is a potential substrate for mushroom cultivation [26]. A semi-industrial scale mushroomproducing plant has been set up in Mexico, designed to work on one ton of coffee pulp every day. This substrate can also be utilized for the production of fuel and chemicals using microorganisms. Citronella bagasse and lemon grass are the residues of steam distillation of freshly harvested citronell leaves and lemon grass to recover their essential oils [27]. After distillation, the bagasse is partially dried in the field and a fraction is burnt to generate steam for the stripping, and rest is left in the fields where natural degradation takes place. Because of the residual aroma and flavour and animal rejection its use as a ruminant feed is limited. There is an estimated worldwide availability of about 2X10 s tons of dry bagasse per year [28] that could be used as a source of lignoceilulosic material for bioconversion. Shive, a bulky by-product of flax, is left after scutching and has little value [29]. For every ton of fibre produced, 2.5 tons of shive will be left after scutching. However, ruminants can not utilize cellulose because of high lignin content [30]. Apple pomace and orange peels are inexpensive and plentiful available from fruit processing industries. They cause a severe disposal and environmental problems. In Italy, the orange juice industries process about 6X10 s tons of citrus fruits with a residual waste that constitute 60% of the weight of treated fruits [31]. The waste contains a considerable amount of residual sucrose and other carbohydrates but has a low protein content, it has a good digestibilty level, but is of low nutritional value.

Distillery grape stalks have a low carbohydrate and protein contents, but have a high level of cellulosic materials, therefore, they can be good substrate for bioconversion. The residual pulp waste from shochu (alcoholic beverage) contains about 32% carbohydrate and 29% proteins (on a dry weight basis). The other industrial wastes such as corrugated paper, tobacco waste (mid rib) and sulfite pulp waste are produced abundantly in different parts of the world and can be utilized through biological routes [32-35]. A number of forest tree residues from willow, poplar and alder can also be potential lignocellulosic substrates for bioconversion. Most wood and agricultural residues are not generally collected at the time of harvest. Branches, leaf tops and roots of trees are left in the forest. Only 50-75% of the trees are removed during harvest. Most residues are generated during pulping and milling operations. A certain proportion of agricultural residues (about one ton per acre) are left in the soil to maintain the tilth and prevent erosion. Of all biomass resources, low grade hardwoods and agricultural residues are the two largest available components. Each possess characteristics that favours its utilization. Low grade hardwoods are abundant in the southeastern United States and, aside from direct combustion, have few commercial uses. They can be harvested on a year round basis. Some hemicellulose sugars are available as a by-product of hardboard and insulation board manufacture. Others are available as a by-product formed during the manufacture of sulfite and dissolving pulp. As industrial waste of considerable potential as a biomass resource, about 100X106 metric tons of spent sulfite liquor are produced annually as a by-product of the world's pulp and paper operations. The magnitude of available lignocellulosic waste is quite high and their availabilty is varied depending on geographic location and season. Currently their exploitation for value added and economic use is underdeveloped and obviously merits study to create better utilization.

4

NATURE OF LIGNOCELLULOSIC MATERIALS

Regardless of source, lignocellulosic materials contain cellulose, hemicellulose, and lignin as major components. An analysis of the composition of several hardwood and softwood species is presented in Table 4.

10 Table 4 Chemical composition of some forest residues % Dry weight Residues Hexosans

Pentosans

Lignin

Ash

Aspen

50

28

15

0.3

American beech

47

20

23

0.2

Paper birch

41

26

25

1.0

Yellow birch

40

33

21

0.8

Cottonwood

46

19

24

0.6

Sugar maple

42

21

23

0.2

Silver maple

47

18

21

0.2

Red maple

39

33

23

1.0

Poplar

45

19

20

0.1

Black cherry

45

20

21

0.1

White oak

48

18

28

0.4

Sweet gum

40

24

19

1.0

Balsam fir

42

11

29

0.5

Douglas fir

57

8

24

0.4

White fir

56

12

24

0.7

Eastern hemlock

43

10

32

0.4

Jack pine

41

10

27

0.1

White pine

44

11

28

0.1

Red pine

46

12

24

0.2

Black spruce

44

11

27

0.3

Red spruce

43

12

27

0.2

White spruce

44

10

27

0.3

Hardwoods

Softwoods

11 Cellulose and hemicellulose are found in the secondary wall of the cell wall. Crystalline microfibrils of cellulose are surrounded by amorphous hemicellulose, and the whole is embedded in a matrix of lignin [36]. The composition 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. Table 5 presents the composition of various types of agricultural residues. Straw species are more uniform in composition than various wood species.

Table 5 Chemical composition of some agricultural lignocellulosic residues % Dry weight Residues Hexosans

Pentosans

Lignin

Ash

Bagasse

33

30

29

4

Barley straw

40

20

15

11

Corn stover

42

39

14

2

Corn stalks

35

15

19

5

Cotton stalks

42

12

15

6

Groundnut shells

38

36

16

5

Oat straw

41

16

11

12

Rice straw

32

24

13

18

Rice husk

36

15

19

20

Sorghum straw

33

18

15

10

Wheat straw

30

24

18

10

Rye straw

37

30

19

4

Flax shives

35

24

22

3

Soybean stalks

34

25

20

2

12 A wide variations in the chemical composition occurs not only between different species but also within a single species. Generally straws have lower cellulose content than wood but in spite of this, it has a total carbohydrate fraction (holocellulose = cellulose + hemicellulose) approximately equal to that of wood. This is possibly due to its high hemicellulose and low lignin contents compared to wood. The ash (total minerals) content is greater in straw than in wood [37,38]. In natural substrates, cellulose is present in association with hemicellulose, lignin, and extractives. The characteristics such as crystallinity, lignin content, specific surface area are related to the saccharification of the complex polysaccharides. Cellulase enzymes readily degrade easily accessible, amorphous cellulose as compared to crystalline cellulose due to enzyme transport limitation imposed by the closely ordered lattice of the cellulose molecules [39]. Lignin which plays a cementing role in cell wall architecture, creates a hindrance in cellulose hydrolysis. Although lignin is inert in hydrolysis, it can adsorb a part of the active enzyme [40]. Hemicellulose present in lignocellulosic biomass appears to shield cellulose from enzymatic attack. The extractives (resins, waxes etc.) also interfere with the hydrolysis of polysaccharides because of their hydrophobic nature.

4.1

Structure of plant cell walls

The wood cell is a multilayered structure consisting of an external microfibril primary layer and a secondary wall containing three sublayers $1, $2 and $3 [41]. Within each layer of the secondary wall, the cellulose and other cell wall constituents are aggregated into long slender bundles called microfibrils. The microfibrils are distinct entities in that few cellulose molecules cross over from one microfibril to another. In the $1 layer, the microfibrillar groups are in helixes alternately crossed, while in the middle layer ($2), the microfibrillar groups are oriented in bands (lamellae) nearly parallel to the cell axis. In the inner layer ($3) the direction is nearly perpendicular to that in $2. The primary wall has an irregular helical arrangement around the cell axis. Surrounding the fibre is the heavily lignified middle lamellae, shared by adjacent fibers [42]. The concentration of cellulose is highest in sublayer $2 and diminishes towards the middle lamella. The concentration of hemicelluiose is maximum in the middle lamella and

13 decreases toward the lumen. The concentration of lignin in the middle lamella is about 90% in hardwood and 70% in softwood, with the major part (70-80%) of the lignin being distributed within the secondary wall [43,44]. Wood cells may contain upto 90% fibre whereas only about 35-39% of the cells in straw are fibre [45]. The hemicellulose and lignin form a matrix surrounding the cellulose. Within a given microfibril, lignin and hemicellulose may penetrate the space between cellulose molecules in the amorphous region. Most of the cell wall capillaries are closed when iignocellulosics are free of water, but open after the absorption of water. The total surface area exposed in gross capillaries (2Xl 03cm2.g1) is several orders of magnitude smaller than the total surface area exposed within the cell wall capillaries (3x103cm2.gl). The penetration of cellulase enzyme to cell wall capillaries substantially increases the saccharification rate [46]. Cellulose contains the regions of high and low crystallinity so that different regions exhibit different susceptibilities during the progress of enzymatic reactions [47-48]. Together with hemicellulose and pectins, lignin fills the spaces between cellulose fibrils in woody cell tissues and functions as a binding material. Both physical association and chemical bonding have been postulated [49, 50]. The cell wall structure of straws has been less studied than that of wood. Straw is much more heterogenous raw material than wood. Straw fibres, principally derived from cells and internodes, are fairly long and slender with sharply pointed ends [38]. In addition, straw also contain short nonfibrous cells consisting of epidermal cells, platelets, serrated cells, and spirals which are derived from the pitch, nodes, chaffs and rachises.

4.2

Cellulose

As the most abundant organic substance on the earth, cellulose is one of the thoroughly studied chemical compounds. Cellulose is a linear homopolymer of anhydroglucose units linked together with 13-1,4-glycosidic bonds. Some natural materials are practically pure cellulose, e.g. cotton. Cotton is (z-cellulose, a form insoluble in 17.5% NaOH. Plant and wood celluloses generally contain 13-cellulose as well, a material soluble in the above solution. Although cellulose is formed from D-glucose building blocks joined by 13-1,4-glycosidic bonds, there are differences in the degree and types of association within cellulose molecule. The cellulose molecule is a polymer with molecular weight

14 generally in the range of 3-5X105. Table 6 shows typical materials and their degree of polymerization. Glucose, as well as cellobiose, cellotriose, and cellotetraose can be isolated when cellulose is hydrolysed. Complete hydrolysis by acid yields D-(+)-glucose as the only monosaccharide. The cellulose molecule is thread-like, existing as fibril-long bundles of molecules stabilized laterally by hydrogen bonding between hydroxyl groups of adjacent molecules. Molecule arrangement in the fibrillar bundles is so regular that cellulose has a crystalline X-ray diffraction pattern. The consequence of the high degree of order in native cellulose is that not even water molecuels, let alone enzyme, can enter the structure. The structure of acid or alkali swollen cellulose is open and this material is readily split by cellulase enzyme. In nature cellulose exists in a highly organized state known as fibrous crystal. The basic repeating units of the crystal is the unit cell, which was defined as a monoclinic lattice with cellulose chains packed at the corners and the center of the cell [51]. Later the unit cell was redefined and named 'Meyer and Misch unit cell' of cellulose [52]. This structure has since been well received except for some minor corrections and some occasional disputes on the chain orientation [53-55]. The confusion arises from the fact that cellulose is a paracrystalline substance but never is a perfect crystal. The X-ray diffractograph does not show sufficient data points to differentiate one formation from other. However, the evidences strongly support the antiparallel chain orientation in all cellulosic crystals except Valonia cellulose [56,57].

Table 6 Chracteristics of cellulose Source

Native cellulose

Degree of

Molecular

polymerization

weight

3,500-10,000

600,000-1,500,000

Wood pulp

500- 2,100

80,000- 340,000

Chemical cotton

500- 3,000

80,000- 500,000

15 The crystallite structure of cellulose is another interesting feature. Several representative models of the molecular orientation in the crystallite have been developed. According to the fringed fibrillar model (a fibrillar version of the fringed micellar model), cellulose molecules in the elementary fibril are fully extended with the molecular direction in the line with the fibril axis [58]. Among the fibril, however, there are intermittent highly ordered areas, the so-called crystalline regions, separated by the less ordered amorphous regions. Folding chain model of the crystallite structure reveals that the cellulose molecules are being folded back and forth along the fibrillar axis within the 101 plane of the crystalline lattices. Thus, the folding molecule forms a sheet-like 'platellite unit'at the fold length of crystal line and makes up the basic molecular unit of the cellulose fiber [59]. A total of as much as 1,000 degree of polymerization (DP) can be accomodated within this platellite unit. If the whole molecule is very much longer than 1,000 DP, the rest of the chain will enter into the neighbouring platellite above or below in series along the elementary fibril. In this way, the corresponding portions of molecules connecting two platellites are single stranded chains and hang loose from the crystalline structure. These are the weak spots in the molecule vulnerable to relatively mild degradation, such as by exposure to light or by mechanical impact. But the breaking of these portions does not affect the physical and chemical properties of the cellulose fiber. The glycosidic bonds at the folds are different and much weaker than the linear bonds, but structurally very important to the integrity of the crystal. One bond breakage per basic molecular unit (1,000 DP for native and 300 DP for regenerated cellulose) will cause the disintegration of the crystal and severe loss in the mechanical strength of the fiber [59]. The multiple passages of the molecules through the amorphous and crystallite regions have also been suggested. The crystalline structure apparantly plays a very important role in the hydrolytic degradation of cellulose. It has been demonstrated that cellulose of high crystallinity reacts much slower than that of low crystallinity in enzymatic hydrolysis [60]. Native cellulose is water insoluble and its susceptibilty to hydrolytic enzyme attack depends significantly on its structural features i.e. surface area and crystallinity [61,62]. The importance of the former stems from the fact that contact between the enzyme molecules and the surface of cellulose particles is a prerequisite for hydrolysis to proceed, and that of the latter from the fact that the cellulolytic enzymes degrade the more accessible amorphous region of cellulose more readily than the less accessible crystalline region. As the crystallinity increases, cellulose become increasingly resistant to further hydrolysis. The increased initial specific surface area enhances the extent of initial soluble protein adsorption, which in turn, increases the initial hydrolysis rate [63].

16

4.3

Hemicellulose

Hemicellulose by definition are the short branched chain heteropolysaccharides of mixed hexosansand pentosansthat are easily hydrolysed [64]. D-Xyloseand L-arabinose are the major constituents of pentosans while D-glucose, D-galactose and D-mannose are the constituents of hexosans. Chemical structure of some naturally abundant pentoses and pentitols are presented in Figure 1.

CHO H-(~-OH HO-(~-H

H- I-OH

CHzOH

D-Xylose

CHzOH H-(~-OH HO-(~-H H-(~-OH I CHzOH Xylit ol

CliO H-(~-OH HO_f_ H

IHO H- -OH H- IOH

HO- I .

H- I-OH

CHzOH

L-Arabinose

CHzOH

D-Ribose

CHzOH HO-(~-H HO-f- H HO- I -H CHzOH

CHzOH H-(~-OH H" ! -'OH H- -OH CHzOH

Ara bitol

Ri bit ol

Figure 1. Chemical structure of some naturally abundant pentoses and pentitols

The close association of hemicellulose with cellulose and lignin contributes to cell wall rigidity and flexibilty. The majority of the hemicellulosic polysaccharides are derived

17 from cell wall middle lamella. Some of the non-starch, non-cellulose polysaccharides, excluding pectic materials, known as cereal pentosans, are sometimes also considered hemicellulose [65]. Hemicelluloses are composed of neutral sugars, uronic acids and acetyl groups, all present as their respective anhydrides, i.e. xylan, araban, glucan, galactan and mannan (Table 7). As anhydrides, hemicellulose averages about 26% of hardwood, 22% of softwood and about 30% of various agricultural residues [66-69]. The hemicellulose sugar content varies greatly with the plant species. In addition, the individual sugars may be methylated or acetylated.

Table 7 Distribution of hemicellulosics in wood and straw % Dry weight Substrate Xylan

Araban

Galactan Mannan

Glucan

Hardwood

17.4

0.5

0.8

2.5

50.1

Softwood

5.7

1.0

1.4

11.2

46.3

Straw

16.2

2.5

1.2

1.1

36.5

Unlike the orderly crystalline structure of cellulose, hemicellulose exhibit variability in both structure and sugar constituents. Hemicelluloses are commonly composed of two to six different sugar residues with a degree of polmerization of approximately 200. Thus hemicellulose chains are simple or mixed polysaccharides of smaller dimensions than cellulose. The interior chain of hemicellulose consists of polysaccharides that are attached to a variety of sugar residues that are same or different from the sugars that form the side chains.

Except for galactose based hemicellulose with a ~-l,3-1inkage,

most

hemicelluloses are based on ~-l,4-1inkage of their constituent sugars. A D-xylose backbone with L-arabinose side chains is the most abundant form. Table 8 shows the composition of pentose sugars in different lignocellulosic residues.

18 Table 8 Pentose sugar composition of various lignocellulosic residues % of total hemicellulose sugars Residue Xylose

Arabinose

Corn cobs

65

10

Corn stalks

71

9

Corn husk

54

13

Wheat straw

58

9

Soybean stalks

60

7

Soybean hull

27

13

Sunflower

61

2

Flax straw

65

13

Peanut hull

46

5

Sugarcane bagasse

60

15

Maple

33

1

Alder

20

1

Birch

39

3

Beech

28

2

Poplar

24

3

English oak

26

1

Pine

9

2

Tamarack

7

2

Spruce

7

2

Balsam fir

5

1

Agricultural

Wood

19 The types of hemicelluloses are often classified according to the sugar residues present [70,71]. Commonly occurring hemicelluloses are D-xylan, L-arabino-D-xylan, Larabino-D-galactan, L-arabino-D-glucurono-D-xylan, L-O-methyI-D-glucurono-D-xylan, Larabino-(4-O-methyI-D-glucurono)-D-xylan, D-gluco-D-mannan, and D-galacto-D-gluco-Dmannan. The type and amount of hemicellulose varies widely, depending on plant materials, type of tissues, stage of growth, growth environment, physiological conditions, storage and method of extraction [72-74]. The major class of hemicellulose is xylans, which are found in large quantities in annual plants and deciduous trees and smaller quantities in conifers. Xylans of grasses and cereals are generally characterized by the presence of L-arabinose linked as a single unit side-chain to a D-xylose backbone. Substantial differences in sugar constituents are found in wood xylans. Wood xylans are characterized by the presence of 4-O-methyI-D-glucuronic acid linked to a D-xylose backbone. In general, the proportion of 4-O-methyI-D-glucuronic acid is higher in softwood than in hardwood. D-Xylan comprises 15-30% of annual plants, 20-25% of hardwoods and 7-12% of softwoods. True xylans composed exclusively of D-xylose subunits are rare. Xylan appears to be a major interface between lignin and other carbohydrate component in many isolated phenolic-carbohydrate complexes [75-79], likely covalently linked to phenolic residues via its arabinosyl [80] and glucuronosyl [81] residues. The high phenolic content in some of these complexes suggests that the carbohydrates are linked to lignin fragments. However, the phenolic substituents may also cross-link xylan to other carbohydrates [82]. Linkages between xylan and pectic substances have also been suggested, as have xylan-glucan-protein complexes [83]. Association of xylan to other carbohydrates can be covalent or noncovalent in nature. The alkali-labile acetyl substituents and reducing end groups have been recognized in xylans [84], but as much as 50% of the alkaline-labile xylosidic linkages remain to be identified in some plant materials. Xylan tends to adsorb onto cellulose and to aggregate with other hemicellulosic components, likely the result of hydrogen-bonding interactions [85,86]. Xylan may play a major role in cell wall cohesion since its selective removal from delignified wood fiber results in a substantial increase in fiber porosity [87]. There have been observations which suggests that cellulose is protected from enzymatic attack by xylan and mannan [88,89]. When xylan or mannan was selectively removed from delignified fiber using enzymes, the residual cellulose was more accessible to hydrolysis by cellulolytic enzymes. However, a similar prehydrolysis of cellulose or mannan did not improve the accessibility of xylan to xylanases. It shows that xylan may be relatively more important to fiber cohesion so that its selective removal increases accessibility of other

20

polysaccharides by increasing fiber porosity [90]. Only one hemicellulose can be isolated in reasonable yields by direct extraction of fully lignified plant material with water, namely arabinogalactan, which is present in the heartwood of larch in amounts between 10 and 25%. Two other hemicelluloses can be isolated in reasonable representative yields by direct extraction with aqueous alkali, namely hardwood xylan and arabinoxylan from grasses [91]. For quantitative isolation of xylans from hardwoods and straw as for isolation of softwood hemicelluloses, the material has first to be delignified, after which the resulting holocellulose is treated with alkali. Naturally occuring hemicellulose differ from isolated hemicelluloses. Besides impurities of other cell wall materials, isolated hemicellulose suffer from the oxidative delignification procedure, which may lead to reduction in the chain length of the polysaccharides. During pulping, the nature of of wood hemicelluloses are changed. The nature of xylan in wood is dependent upon the type of polymer originally present in the wood and also on the pH of the cooking liquor used to prepare a pulp. If a conventional kraft cook with its high pH is employed to prepare a pulp, xylan or arabinoxylan is found in a good yield dependent upon whether the wood contains 4-O-methylglucuronoxylan or arabino-4-Omethylglucuronoxylan. The kraft process starts under extreme alkaline conditions which causes hemicellulose losses. Two-third of glucomannans are dissolved very quickly and degraded by the alkaline peeling reactions [92]. The peeling reaction in xylan is much more slow compared to the degradation of cellulose or glucomannan due to the unique sequence of sugar units at the reducing end of xylan. The reducing xylose unit is isomerized into a 2-keto sugar unit and cleaved off by 13-elimination, so that the next sugar unit forms the reducing end group, which is galacturonic acid, bound to rhamnose in 2-position. Arabinose, which occurs in the furanose form as a xylan appendix, is much quickly degraded than pyranose substituents.

4.4 Lignin

Lignin is the characteristic cementing constituent between the cell walls of woody tissues. It does not represent a definite, uniform compound, but is a collective form for substances that have very similar chemical properties but very different molecular weights. The molecular weight of lignins may reach the range of 100,000 daltons or

21

greater. A considerable part of the photosynthetic activity in plant is devoted to the conversion of atmospheric carbon dioxide to lignin. The photosynthetic assimilation of atmospheric carbon dioxide by plants leads to the formation of carbohydrates. Carbohydrates are metabolized via a shikimic acid pathway and converted to phenyl propane amino acids. These amino acids supply precursors for the synthesis of plant proteins, flavonoids and lignin. Lignin is most concentrated in and between the primary walls (middle lamella) of vascular plants [93]. It performs a number of functions in the life of plants: Permanent bonding agent between cells, making composite structure of wood. Energy storage system. Prevent enzymatic degradation of other plant components. UV light stabilizer and antioxidant. Water proofing agent. The water-permeation-reducing property of lignin plays an important role in the internal transport of water, nutrients, and metabolites in the plant. The healing process of injured plants involves lignification and the suberization of surface cells, which is associated with an increased resistance to infection [94,95]. The present concept indicates that lignin and cellulose form a mutually interpenetrating system that is largely physical in nature [50]. The nature of ligninhemicellulose bond is suggested to be ether and 4-O-methylglucuronic acid ester to the o~-carbon of lignin unit [96]. Lignin is composed of 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 varies not only with the plant species but also with the plant tissues and location of lignin within the plant cell wall. Ecological factors such as the age of the wood, climate, plant sustenance and amount of sunlight also affect the chemical structure of lignin. A major problem in studying the chemistry of lignin has been the difficulty in isolating lignin from plant materials without secondary reactions. There are several reasons for this complication: Lignins, unlike polysaccharides, proteins and nucleic acids, do not have identical readily hydrolyzable linkages repeating at regular intervals.

22 They are irregular and have no precise chemical structures, but have a series of chemical groupings. Lignins are difficult to extract in an unmodified state. Lignins can not be degraded efficiently into monomeric units because of C-C and diaryl ether type bonds which are difficult to cleave. The hydrolysable linkages in lignin are suggested to be of two types: ~-aryl ether and o~-aryl ether [97]. The predominant ~-aryl ether type bond is more resistant to cleavage [98]. Under mild hydrolytic conditions, the cleavage of the ether bond is exclusively restricted to those of the o~-aryl ether type [99,100]. In terms of physical properties, lignin is an amorphous polymer that has no crystallinity. The mode of polymerization during lignin biosynthesis makes it optically inactive. 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 [101]. Lignin may be oxidized in air. It is insoluble in water, and difficult for microoganisms to penetrate. Lignin is generally acid stable but can be solubilized under alkaline conditions, it is closely bound to cellulose and hemicellulose in plant cell walls and its separation from lignin-carbohydrate complex cannot be achieved by using conventional methods. Sequential enzymatic and chemical hydrolysis of lignin-carbohydrate complex from aspen wood gave evidence for arabinan type polysaccharides and lignin-saccharide ether bonds in the complex [102,103]. Lignin preparations of woody materials contain, beside carbohydrates, significant amounts of protein [104].

4.5

Extraneous materials

In addition to cellulose, hemicellulose and lignin, plant cell walls contain extraneous materials, including extractives and non-extractives. The extraneous component consists of astonishingly wide variety of chemicals. Wood contains 0.4-8.3% extractives on a dry weight basis, whereas agricultural residues contain even greater amounts [105,106]. The extractives can be broadly devided into three groups, namely, terpenes, resins and phenols. The terpenes are regarded as isoprene polymers and are terpene alcohols

23 and ketones. The resins include a wide variety of non-volatile compounds, including fats, fatty acids, alcohols, resin acids, phytosterols, and some less known neutral compounds. The phenols consist of a large number of compounds which are yet to be studied thoroughly. However, the most important among them are tannins, heartwood phenols, and related substances. In addition, low molecular weight carbohydrates, alkaloids, gums and various other cytoplasmic constituents are present [107,108]. Extractives can be removed by treating the substrate with water or neutral solvents like ethyl ether, acetone, ethanol, and benzene [109]. The non-extractives make up 0.2-0.8% of the dry weight and include inorganic components such as silica, carbonates, oxalates, and non-cellular substances [106]. In agricultural residues, the non-extractives make up about 10% of the dry weight. Silica, deposited as crystals is especially abundant in straw. Small amounts of non-cell wall substances, such as starch, pectins, and proteins are not extractable. In spite of their small quantity, the role of extraneous materials is significant in that they render cellulosic materials not only resistant to decay and insect attack but also inhibitory to pulping and bleaching.

5

APPLICATIONS IN BIOTECHNOLOGY

Microbial utilization of the inexhaustible lignocellulosic biomass for the production of value-added products such as industrial chemicals, liquid fuels, protein-rich food and feed, and preparation of cellulose polymers is an attractive approach to help meet energy and food demand. Depending on their availability, the feasibility of several lignocellulosic materials for such purposes has been studied around the world. Another issue that arises in this context is that estimates of conventional feedstock costs are in a conversion range from ca. 30-70% of the product selling price [110,111]. Thus much of the current interest has been focussed on the processes that are based on cheaper cellulosic and hemicellulosic feedstocks. Biomass can be converted to valuable chemicals by either thermochemical or biological means. A thermochemical process requires high temperature and pressure and produces a complex mixture of products. A biological conversion process using microorganisms, on the other hand, operates at a lower temperature and produces specific products in high yields with fewer by-products [112].

24 While biomass has served as a substrate in microbial processes for the production of alcoholic beverages for thousands of years, it has only been recently that broader applications of this material have been investigated. It is possible to generate useful products form wastes by effective recycling of materials with the consequent reduction of overall cost using fermentation technology. Any waste stream containing carbohydrates can be utilized for the generation of useful chemicals [113]. When the alternative technologies of oil-based feedstocks are compared biologically produced chemicals compare favourably with the chemical industry [114]. Fermentation involves simpler technology, and the by-products are mostly nontoxic, unlike those from chemical processes [115]. A fermentation plant can be smaller and dispersed to meet social needs. Biomass-derived alcohols, ketones, and acids produced by fermentation can enter into the current petrochemical synthetic pathways through a number of reactions. The most important being dehydration of alkanols to alkenes to synthesize ethylene, polypropylene, butylene, and butadiene. Industrial processes using the lignocellulosic materials have traditionally made use of only the hexose component of the holocellulose. Therefore, the pentose sugars, which may comprise as much as 40% of the plant materials, have in most cases been wasted [116]. The economics of bioconversion, would be more feasible if both hexose and pentose sugars could be utilized [117]. While the technology employing yeasts and bacteria to produce chemicals and solvents from hexoses is well known, the ability of these organisms to ferment pentoses has been considered problematical [118-121]. The pentose sugars in question being D-xylose and L-arabinose. These sugars alongwith Dglucose, D-mannose and D-galactose (hemicelluloses) form and attractive fermentation feedstock. Hemicellulose derived carbohydrates have many potential uses. Bioconversion of hemicellulose often requires prior hydrolysis of the polysaccharides to their sugar constituents. These sugars can be converted by microorganisms to various products such as ethanol, sugar alcohols, solvents, organic acids, and single cell protein and lipid (Figure 2). The yield, rate of hydrolysis, and type of sugar recovered depend on the source of substrate and its composition. The nature and amount of conversion products depend on the type of sugar, metabolic efficiency of the organism, and culture conditions employed [113,122]. Of the many products available from hemicellulose-derived carbohydrates, ethanol has received the most attention, because of its potential use for blending with petroleum. In addition, it is also a versatile chemical feedstock, and a variety of chemical products can be derived from ethanol.

25

IDEHYDP,AT~

I

I HYDROGENATION,,I

IFERMENTAT~)N I GLUCOSE

L,NO L'ULOS,, FELLULOSEI

-I~ASH ~

BIOMASS

--~ Lr.~IN

IHEMICELLULOSE I

I FERMENTATIONI

Figure 2. Potential utilization of lignocellulosic materials

I

DEHYDRA~ON I

26 Many research groups have been involved in the selection of better microbial strains for the production of industrial chemicals from pentose sugars [122-129]. Noteworthy is the development of yeast strains and recombinant bacterial strains that tolerate high substrate and ethanol concentrations giving high final product yields. Thermophiles and high temperature tolerant strains could be useful, since cooling problems are simplified. Furthermore, high temperature facilitates the hydrolysis of biomass and results in higher rates of product formation. Selection of high ethanol-tolerant flocculating strains is also important because these could be used in a continuous process. Following chapters in this book review aspects of biosynthesis, biodegradation and extraction of hemicelluloses, as well as recent microbiological, biochemical, and biotechnological findings in relation to our previous understanding of pentose fermentation, metabolism, and utilization. The regulatory aspects of microbial pentose metabolism and genetic improvements of pentose fermenting microorganisms are discussed. The state of the art for the production of ethanol, xylitol, acetone-butanol, 2,3-butanediol, organic acids, and single cell protein and lipid is presented.

6

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55

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56

Chang MM, Chou TYC, Tsao GT. Adv Biochem Eng 1981; 20: 16.

57

Ellefson O, Tonnesen BA. Cellulose and Cellulose Derivatives, New York: John Wiley, 1971; 151.

58

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59

Chang M. J Polym Sci 1971; 36: 343.

60

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62

Ghose TK. Adv Biochem Eng 1977; 6: 39.

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Lee Y-H. Biotechnol Bioeng Symp 1978; 8: 75.

64

Whistler RL, Richard EL. The Carbohydrates, New York: Academic Press, 1970; 447.

65

Timell TE. Adv Carbohydr Chem 1965; 20: 409.

66

Brasch DJ, Wise LE. Tappi 1956; 39: 581.

29 67

Rydholm SA. Pulping Processes, New York: Wiley ,1965; 95.

68

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69

Gordon AH, Lomax JA, Chesson A. J Sci Food Agri 1983; 34: 1341.

70

Polyglase WJ. Adv Carbohydr Chem 1955; 10: 283.

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73

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74

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77

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79

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8O

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84

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86

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88

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90

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91

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30 96

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97

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99

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100 Kirk TK, Farrel RL. Annu Rev Microbiol 1987; 4: 465. 101 Hatakeyama T, Hatekayama H. Polymer 1982; 23: 475. 102 Das N. Carbohydr Res 1981;94: 73. 103 Joselen J-P, Gancet C. Svensk Paperstidn 1981; 84: 123. 104 Whitemore F. Phytochemistry 1982; 21:315. 105 Ladisch MR, Lin KW, Valoch M, Tsao GT. Enzyme Microb Technol 1983; 35: 156. 106 McDonald RG. The Pulping of Wood, New York: McGraw Hill, 1969; 34. 107 Cowling EB, Merill W. Can J Bot 1954; 44: 1539. 108 Hillis WE. Wood Extractives and Their Significance to the Pulp and Paper Industries, New York: Academic Press, 1962. 109 Ghosh P, Singh A. Adv Appl Microbiol 1993; 39: 295. 110 Scneider H. Crit Rev Biotechnol 1989; 9: 1. 111 Chahal DS, ed. Food, Feed and Fuel from Biomass, New Delhi: IBH, 1989. 112 Stewart GG, Panchal CJ, Russel I, Sillis AM. Crit Rev Biotechnol 1984; 1: 161. 113 Mishra P, Singh A. Adv Appl Microbiol 1993; 39: 91. 114 Wiegel J. Experientia 1980; 36: 1434. 115 Palson BO, Afsar SF, Rudd DF, Lightfoot EN. Science 1981; 213: 513. 116 Timell TE. Wood Sci Technol 1967; 1: 45. 117 Skoog K, Hahn-Hagerdal B. Enzyme Microb Technol 1988; 10: 66. 118 McCracken LD, Gong C-S. Adv Biochem Eng/Biotechnol 1983; 27: 33. 119 Kurtzman CP. Adv Biochem Eng/Biotechnol 1983; 27: 73. 120 Gong C-S. Annu Rep Ferment Proc 1983; 6: 253. 121 Magee RJ, Kosaric N. Adv Biochem Eng/Biotechnol 1985; 32: 61. 123 Wang PY, Shopsis C, Schneider H. Biochem Biophys Res Commun 1980; 94: 248. 124 Gong C-S, Chen LF, Flickinger MC, Chiang LC, Tsao GT. Appl Environ Microbiol 1981 ; 41: 430. 125 Kumar PKR, Singh A, Schugerl K. Proc Biochem 1991;26: 209. 126 Singh A, Kumar PKR. Crit Rev Biotechnol 1991; 11:1129.

31 127 Singh A, Kumar PKR, Schugerl K. Biotechnol Appl Biochem 1992; 16 296. 128 Lawford HG, Rousseau JD. Biotechnol Lett 1991; 13" 191. 129 Singh A, Kumar PKR, Schugerl K. Adv Biochem Eng/Biotechnol 1992; 45" 29.

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2 Biosynthesis and Biodegradation of Hemicelluloses

1

INTRODUCTION

Hemicelluloses are those plant cell wall polysaccharides which occur in close association with cellulose, especially in the lignified tissues. Hemicelluloses are operationally defined as alkali-soluble polysaccharides, and the term often being restricted to substances extracted with alkaline reagents but not with water [1]. There is, depending on the extraction methods an overlap between hemicelluloses and pectic fractions. The hemicelluloses rank next to cellulose in abundance as naturally occurring compounds. They are linear and branched heteropolymers of D-xylose, L-arabinose, Dmannose, D-galactose, D-glucose and D-galacturonic acid. In addition, homopolymers of xylose, galactose and mannose are known to occur. Individual sugars may be methylated or acylated. Of this group only three of the pentoses (D-xylose, L-arabinose and D-ribose) and two pentitols (D-arabitol and ribitol) are found in significant quantities in nature. The other pentoses are known as unnatural carbohydrates due to their rarity in or apparent absence from the natural environment [2].

2

BIOSYNTHESIS OF HEMICELLULOSIC SUBSTANCES

The discovery of sugar nucleotides in the early 1950s was an important landmark for future studies on the biosynthesis of polysaccharides and complex carbohydrates [35]. There has been considerable increase in our understanding of the mechanism of polymerization of sugars. Involvement of the nucleotide sugars as precursor of cell wall polysaccharides is now well established [6]. While the broad scheme is clear, important details are regularly emerging. It is quite clear by now that the individual sugars are first converted to their sugar nucleotide derivatives, and these high-energy intermediates are then utilized in the biosynthetic and polymerization reactions. 33

34

2.1 Synthesisof sugar nucleotides Sugars must first be phosphorylated in order to form their sugar nucleotide derivatives. The major source of many of the phosphorylated sugars of plants is Calvin cycle, which give rise to D-fructose-6-phosphate (Figure 1). D-Fructose-6-phosphate can then be transformed to D-mannose-6-phosphate, D-glucose-6-phosphate, N-acetyI-Dglucosamine-6-phosphate, and so on [7,8]. These sugar phosphates are then transformed via mutase reactions to sugar nucleotides.

PHOTOSYNTHESIS

Ma~~6-P Man-1 -P

:~_.,

im~

,v

Fru-6-P-.

+

.....

-

'

Glucose

v

Glc-1 -P

GIcN-6-P

+ 6DP-Man

GIc-6-P

UDP-GIc

61cNAc-6-P

+f

GIcNAc-1 -P

UDP-GIcA

UDP-GIcNAc~,~

UD~~yl

#

UDP-GaiNAc

+

UDP-Gal

UDP-L-Ara

Figure 1. Metabolism of hexoses and pentoses, and their conversion to sugar nucleotides

35 In addition, a number of specific kinases are present in plants which can phosphorylate monosacchrides to provide sugar phosphates as substrates for the sugar nucleotide phosphorylases [9]. Kinases have been demonstrated in a number of different plants that can phosphorylate D-glucose, D-mannose, D-fructose and D-glucosamine at the C-6 position, and L-arabinose, D-galactose, and D-galacturonate at the C-1 position. These kinases include such enzymes as hexokinase, D-galactokinase, L-arabinokinase, D-glucuronokinase, and D-galacturonokinase [10-14]. Sugar-6-phosphates are required to be converted to sugar-1-phosphate before they can be utilized as substrate for sugar nucleotide synthesis. Mutases are capable of catalyzing such reactions. For instance, phosphoglucomutase catalyzes the interconversion of glucose-6-phosphate and glucose1-phosphate and phosphomannomutase catalyzes the interconversion of mannose-6phosphate and mannose- 1-phosphate [ 15-17]. Sugar nucleotides are synthesized de novo by the action of sugar nucleotide pyrophosphorylase. A specific example of pyrophosphorylase reaction is that of UDPglucose pyrophosphorylase (UDP:o~-D-glucose-l-phosphate uridylyl transferase, EC 2.7.7.9), which catalyzes the formation of UDP-glucose via the reaction of UTP and Dglucose- 1-phosphate [18]. Although many pyrophosphorylases have been demonstrated in the plant tissues that catalyze the synthesis of different sugar nucleotides, those catalyzing the formation of D-glucose containing nucleotides predominate. In addition to UDP-glucose, plants also contain ADP-D-glucose, GDP-D-glucose, TDP-D-glucose, and CDP-D-glucose which are formed from specific pyrophosphorylases [19-23]. Apart from glucose nucleotides, a large number of other sugar nucleotides have also been identified. Most of these are formed by specific nucleotide pyrophosphorylases. These sugar nucleotides include UDP-D-xylose [24], UDP-L-arabinose [25], UDP-D-galacturonic acid [26], UDP-D-glucuronic acid [27], GDP-L-fucose [28], UDP-D-galactose [29], GDPD-mannose [30], UDP-N-acetyI-D-glucosamine [31], and ADP-N-acetyI-D-glucosamine [32]. Some of the sugar nucleotides may also be formed by transformation of other nucleotides. A number of enzyme preparations [33,34] capable of synthesizing hemicellulosic polysaccharides have been examined for their nucleotide sugar specificity and glycosidic linkage of product (Table 1). They may be associated with epimerases and other enzymes capable of interconverting sugar nucleotides. The incorporation of labeled sugars into polymeric materials has been interpreted as polysaccharide biosynthesis, but in many cases this conclusion has not been warranted on the basis of the experimental data. Polysaccharide synthesis actually involves incorporation of a large number of monomers to synthsize new polymeric chain of high degree of polymerization [35-39].

36 Elongation of the polysaccharide chain, whether by the addition of monosaccharides or oligosaccharides, could occur at the nonreducing end or at the reducing end [40,41].

Table 1 Sugar nucleotide specificity of hemicellulose polysaccharide synthetases Polymer

Linkage

Sugar nucleotide

Xylan Xyloglucan

1,4-13-xylosyl

U DP-D-Xyl UDP-D-GIc UDP-D-Xyl UDP-L-Ara GDP-D-Man

Arabinan Mannan G lucom annan Galactan Galacturonan

1,4-13-mannosyl 1,4-i~-man nosyi 1,4-13-gIucosyl 1,4-13-galactosyl 1,4-o~-galacturonyl

G DP-D- Man G D P-D-G Ic UDP-D-Gal UDP-D-GalU TDP-D-GalU

Hemicelluloses are characteristically heteropolymers. Unlike animal connective polysaccharides [42] and certain bacterial polysaccharides [43], they do not have repeating units in their backbones or regularly inserted side branches [44]. The partially ordered sequence of linkages or monomers in the main chain is consonant with the view that their assembly is not a template-directed process, but is directed by the specificity of the transferases involved. However, it is not known whether the addition of substituents to the main chains of heteroxylans, heteroglucans or heteromannans is achieved concurrently with the main chain synthesis or occurs later. Ester or ether derivatives of monosaccharide residues are frequently found in hemicellulosic polysaccharides. It is not known whether these modifications occur before polymerization, concurrently with polymerization, or after the polysaccharide has been assembled [45,46]. The polysaccharide composition of the primary wall, middle lamella

37 and secondary wall formed during the development show characteristic qualitative and quantitative differences [47-49]. The control of polysaccharide synthesis which leads to these differences may be either through the supply of precursors or the intracellular sugar and sugar phosphate pool or through the modulation of the activity of pre-existing enzymes which establish the pool of nucleotide sugars and the enzymes which utilize these as substrates in polymerized reactions. The activation of monosaccharides to the nucleotide form and the reaction involved in the subsequent interconversion of the activated monosaccharides are possible control points in the biosynthesis of plant polysaccharides (Figure 2).

~ POOLOF SOLUBLE ' ~ Isomerases SUGARS AND | Kinases ,, ~) Mutases SUGAR PHOSPHATES

Sugar nucleotide pyrophosphorylase

sPOOLOF SOLUBLE~

Epimerases UGAR NUCLEOTIDES ) Decarboxylases Dehydrogenases Polysaccharide synthetase

~OLYSACCHARiDES) l

Deposition

p' CELLWALL S~ OLYSACCHARIDE

Figure 2. Biosynthesis of cell wall polysaccharides

38 The activities of various enzymes (dehydrogenases and decarboxylases), responsible for the maintenance of the nucleotide sugar pool, varies with the stage of differentiation or growth of the tissue. These changes also correlate well with changes in the chemical composition of the wall during development. In gymnospermic plants, the specific activities of these enzymes change during differentiation in a way which reflects the smaller amounts of xylan and larger amounts of arabinogalactans and galactoglucomannans.

2.2

Interconversion of sugar nucleotides

Many sugar nucleotides are formed directly from their sugar-l-phosphate and the appropriate nucleotide triphosphate via a pyrophosphorylase reaction. A number of other sugar nucleotides may also be synthesized by transformation of already existing sugar nucleotides. Most widely studied enzymes responsible for the sugar nucleotide interconversions are the 4-epimerases. These enzymes have been known since 1951, when the interconversion of UDP-D-glucose and UDP-D-galactose was demonstrated [4]. The enzyme, UDP-D-glucose-4-epimerase, may represent the only means to synthesize UDP-galactose in plants [50]. It requires NAD as a cofactor, and the reaction mechanism involves UDP-4-ketohexose as an enzyme-bound intermediate [51-53]. In addition to the D-glucose:D-galactose pair, other 4-epimeric pairs are also found in nature, such as L-arabinose:D-xylose and D-glucuronic acid:D-galacturonic acid. Mung bean plants were shown to contain UDP-derivatives of all of the 4-epimer pairs [48,49]. In addition, the tissue extracts catalyze the interconversion of UDP-glucuronic acid and UDP-galacturonic acid [20]. Particulate preparations could also cause the interconversion of TDP-galactose and TDP-glucose [54]. The presence of 2-epimerases which can interconvert D-glucose and D-mannose as the sugar nucleotide derivatives has also been demonstrated in in some tissues. In particulate enzyme preparations of mung bean seedlings, a glucomannan synthesized from GDP-[~4C]mannose was found to contain radioactivity in both the D-glucose and D-mannose moieties [36]. Interconversions of some sugar nucleotides are shown in Figure 3.

39

CH20H

HO/~'---...~0 H~

O-UDP

O-UDP

HO

II

4-Epimerase

OH HO

H0~ O N O O-UDP

O--UDP (b)

(a)

CHzOH

COO-

2 NAD§ HO

HO HO

O--UDP

O-UDP

(c)

Figure 3. Interconversion of sugar nucleotides. (a) UDP-glucose and galactose, (b) UDPxylose and UDP-arabinose, (c) conversion of UDP-glucose to UDP-glucuronic acid

Structural similarities amongst D-glucose, D-glucuronic acid and D-xylose have led to the postulation, that oxidation of D-glucose would give D-glucuronic acid and D-xylose could arise by decarboxylation of D-glucuronic acid. These reactions have now been

40 confirmed at the sugar nucleotide level in plants. UDP-D-Glucose:NAD oxidoreductase has been demonstrated in a number of plants [55]. The enzyme forms a Schiff's base between C-6 of the glucose and an amino group of the appropriately placed lysine on the enzyme. Hemithioacetal thus formed is further oxidized to the carboxylic acids. In many plant tissues, UDP-D-glucose oxidoreductase displays complex inhibition by UDPD-xylose [56], which may represent a control in the synthesis of pentose nucleotides. Conversion of UDP-D-glucuronic acid to a mixture of UDP-D-galacturonic acid, UDP-Dxylose and UDP-D-arabinose has been demonstrated in various plant tissues [57]. Presence of 4-epimerase for the uronic acid as well as UDP-uronic acid decarboxylase has been indicated. However, the origin of pentose nucleotides is not clear. In partially purified extracts from which 4-epimerase had been removed, the first detectable product arising from UDP-D-glucuronic acid was UDP-D-xylose. UDP-L-Arabinose accumulated as the reaction progressed [58]. This suggests that UDP-D-xylose results from decarboxylation of UDP-D-glucuronic acid which is subsequently epimerized to UDP-Larabinose. UDP-D-Glucuronate decarboxylase is involved in the decarboxylation reaction [59].

2.3

Polymerization

"One gene-one glycosidic linkage" hypothesis for glycoprotein assembly states that each glycosyl transferase is specific not only for the transfer of a particular sugar but also for the anomeric configuration and position of the glycosidic linkage formed. This also applies to polysaccharide polymerases [60]. The short oligosaccharide substituents on heteroxylans and heteroglucans are synthesized by successive and specific action of individual synthetases [61]. The existence of specific polymerases for polysaccharide assembly also offers a means for independent control of the process. This is probably achieved by modulating the activity of preexisting enzymes. The clearcut changes in polysaccharide types occurring in the primary to secondary cell wall transition suggests the control of events by alterations in the steady-state level of specific polymerases, which in turn would be controlled by the rate of enzyme synthesis and degradation [62,63]. Figure 4 shows the polymerization reactions from sugar nucleotides.

41

C NUCLEOTIDE-MO'NosACCHARIDE)

(NUdLEOTIDE-OLIGOSACCHARIDE)

( L~P~D-OUGOSACCHAR~DE )

(ACCEPTOR)

J CPROTEIN ~)

(OL.IGOSACCHARIDE)

(~ROTEIN-POLYS~,cCHARi DE)

PR0 TE~N

CPOLYSAcc HA RI6~=)

Figure 4. Polymerization reactions from sugar nucleotides

The major hemicellulosic polysaccharides in the cell wall of dicotyledonous plants are xyloglucans. Xyloglucans have the ability to hydrogen bond to cellulose

42 chains, because they bear a structural relationship to cellulose. The polymer is a 13-1,3glucan core to which single o~-l,6-1inked xylose units are attached. The sugars like Dgalactose and/or D-galactose/L-fucose may be linked to the xylose residues. Cell-free extracts of the elongating pea stems catalyze the incorporation of D-glucose from UDP[3H]glucose and xylose from UDP-[14C]xylose into a water soluble xyloglucan [34,38]. Xyloglucan gave rise to the disaccharide xylosyl glucose upon partial acid hydrolysis. Both UDP-xylose:xyloglucan xylosyl transferase and UDP-glucose:~-1,4-glucan glucosyl transferases have been detected. The addition of unlabelled UDP-glucose greatly stimulated the incorporation of xylose from UDP-xylose into the polymer. UDP-Glucose forms the glucan backbone, and then serve as an acceptor for xylose. In the presence of xylose, the glucan chains become modified to yield a heteropolysaccharide of Dxylose and D-glucose units in which D-xylose residues appear to be attached mainly as non-reducing termini onto the glucan core. Although xylans are mainly secondary wall components, they are found in the primary cell wall in monocots. They are the most abundant hemicellulose polysaccharides in many angiosperms. Biosynthesis of arabinoxylan involves the participation of the sugar nucleotides UDP-D-xylose and UDP-L-arabinose. When the particulate enzyme preparations from immature corn cobs were incubated with UDP[14C]xylose, the radioactive product resembled native arabinoxylan and contained radioactivity in both xylose and arabinose [64]. This also indicate that the particulate enzyme contained the UDP-xylose:UDP-arabinose 4-epimerase. The incorporation of 4O-methyI-D-glucuronic acid into xylan has also been demonstrated in extracts of corn cobs [65,66]. Glucuronic acid from UDP-D-glucuronic acid was found to be incorporated into polymer, and then the glucuronic acid residues became methylated by the transfer of methyl groups from S-adenosyI-L-methionine to the 4-position of glucuronic acid [67]. Glucomannan, another hemicellulosic polysaccharide, is found in cell walls of various higher plants along with galactoglucomannans. However, they are the major cell wall components in gymnosperms. It has been shown in mungbean particulate enzyme preparations that glucose from GDP-[~4C]glucose was incorporated into a j3-1,4-glucan and that this was greatly stimulated by the addition of unlabelled GDP-mannose to the reaction mixture [68]. Mannose from GDP-[~4C]mannose was also incorporated into an alkali-insoluble polymer. The radioactive glucomannan formed from GDP-[14C]mannose was also compared to that synthesized from GDP[14C]glucose in the presence of unlabelled GDP-mannose. Partial acid hydrolysis or enzymatic hydrolysis with a 13mannanase, liberated a similar series of radioactive oligosaccharides from both of the biosynthetic products. These labelled oligosaccharides contained both D-mannose and

43 D-glucose, indicating that both sugars were incorporated into the same polymer. Since both glucose and mannose labelled when only GDP-[14C]mannose was used, it seemed likely that the particulate enzyme contained a 2-epimerase capable of interconverting GDP-glucose and GDP-mannose. Methylation analysis and periodate oxidation of oligosaccharides exhibited two major disaccharides, 4-O-~-D-mannopyranosyl-~-Dglucopyranoside and 4-O-~-D-glucopyranosyl-~-D-mannopyranose [68]. A number of problems arise during in vitro studies on the hemicellulose biosynthetic reactions. The radioactivity from the sugar nucleotide usually ends up in a number of neutral polysaccharides and it is often difficult to determine whether a hemicellulose or pectin, or both have been formed [69]. Since the incorporation of radioactivity is usually low therefore the characterization of product is difficult. In spite of these difficulties, incorporation of radioactive sugars into various polymers has been examined in a number of in vitro systems. Villemez and Hinman [70] have demonstrated the incorporation of D-xylose from UDP-xylose into xyloglucan in particulate enzyme from mungbean seedlings. Incorporation of D-galactose from UDP-galactose into galactan has also been demonstrated [71]. The mucilage or slime polysaccharides from wheat and corn preparations have also been characterized and found to contain the neutral sugar L-fucose (39%) and D-galactose (30%) as the major components and smaller amounts of D-xylose, L-arabinose, D-mannose, D-glucose, and D-galacturonic acid [72].

3

CHEMISTRY OF HEMICELLULOSES

Hemicelluloses include several complexes such as those enriched in xylose (arabinoglucurono- and glucuronoxylans), galactose (arabinogalactans), and mannose (galactogluco- and glucomannans). The hemicelluloses as such represent a distinct group of polysaccharides and are classified according to their chemical composition and structure [73,74]. Three predominant types have been recognized including 1,4-J~-Dxylans, 1,3- and 1,4-J~-D-galactans, and 1,4-~-D-mannans. They usually occur as heteroxylans containing different kinds of sugar residues [75-78]. Classification of plant cell wall polysaccharides by structural family is shown in Table 2

44 Table 2 Classification of plant cell wall polysaccharides Group

Polysaccharide

Glucans

Cellulose Callose 13-D-Glucans Xyloglucans

Arabinans and galactans

Rham nogalacturonans Arabinogalactan I Arabinogalactan II

Mannans

Glucomannans Galactoglucomannans Glucuronomannans

Xylans

Arabino-4-O-methylglucuronoxylan O-Acetyl-4-O-methylglucuronoxylan

Most cell wall polysaccharides, with the exception of cellulose and other J3-glucans, are heteropolysaccharides which fall into a limited number of structural families. Each family contains species with more or less regularly repeating features, usually in the interior chains, but the individual members may exhibit considerable variations in the nature and proportions of other sugar units in side-chains, and of other features such as ether or ester functions.

In addition to that some families

(xylan and

rhamnogalacturonan show discrete variations in the nature of the sugar units in side chains No complete picture of biological materials can be given without reference to their original condition in their natural environment. However, complete extraction of

45 hemicellulose by alkali is only possible after removal of lignin. In case of coniferous wood (e.g. spruce), extraction of hemicellulose is often impossible unless lignin has been previously removed [79]. Of the two common delignification methods, using acidified sodium chlorite [80] and chlorine [81], the previous method probably has the greater degradative effect on both the residual cellulose [82] and the isolated hemicellulose [83]. It has been shown that xylans isolated by alkaline extraction of white spruce wood and the corresponding chlorine holocellulose have similar average molecular weight. Alkaline degradation of polysaccharides results in four types of reactions which may cause modification of the hemicellulose as they occur in plants [84,85]. Deesterification of partially acylated polysaccharides. Chemical degradation initiated at reducing groups. Alkaline hydrolysis of glycosidic linkages. The breaking of chemical bonds between hemicelluloses and other cell wall components. A number of acyl groups (mainly acetyl) are present in wood, which are associated with the xylan components of the hemicellulose fraction. Hemicelluloses, containing acyl group, may be extracted from wood hemicelluloses by dimethyl sulphoxide, and further quantities of acylated polysaccharides may be removed by the subsequent extraction with water. The majority of acyl groups in wood arise from acetyl ester. The acetyl groups are substituents of D-xylose rather than D-glucuronic acid residues, the majority probably being attached to position 3 of D-xylose [86,87]. The types of reactions likely to be initiated at the reducing groups of xylans have been exemplified in studies of the alkaline degradation of xylobiose and xylotriose [88]. The corn cob xylan can not be degraded by alkali because of the oxidation of the reducing end groups during the preparation of the chlorite holocellulose [89]. Chlorite delignification does not necessarily oxidize all reducing endgroups, since oat straw xylans prepared from the chlorite holocellulose are degraded by alkali with the formation of acid products. Rye flour arabinoxylan, isolated by direct aqueous extraction is similarly degraded [90,91]. Treatment of isolated fractions under kraft pulping conditions results in the cleavage of glycosiduronic acid linkages, with the formation of neutral xylans.

46

3.1

Xylans

Angiospermic wood of temperate zone is chemically distinguished from that of the gymnosperms by its lower content of lignin and glucomannans, and high content of xylans. On the other hand, tropical angiospermic wood contains as much lignin as do most coniferous woods and a correspondingly low proportion of xylan [92]. The xylose content varies from 19-39%. The xylans from both hard and softwoods, and also from dicotyledonous plants, are characterized by side-chains of 4-O-methyI-D-glucuronic acid, but in some cases small proportions of L-arabinofuranose side-chains are also present. The xylans of cereals and grasses, on the other hand, are characterized by Larabinofuranose side-chains, but in some cases side-chains of D-glucuronic acid are also present. Pentosan composition of different hardwood species is shown in Table 3.

Table 3 Pentosan composition of various hardwood species % of extractive free wood Species Total Xylose pentosans

4-O-Methylglucuronoxylan

4-AcetyI-O-methylglucuronoxylan

Maple Birch Beech Aspen

20.1 25.5 21.0 18.9

18.6 26.4 18.8 17.2

21.3 30.2 21.5 19.7

25.1 34.6 25.4 23.5

Elm

15.3

12.3

14.9

18.3

The xylose residues in hardwoods probably originate from an O-acetyl-4-Omethylglucuronoxylan. The relative proportion of O-acetyl-4-O-methylglucuronoxylan varies considerably from species to species. More than one third of the white birch wood consists of this polysaccharide, whereas the corresponding value for white elm is less

47

than a quarter. Unlike softwood xylans, hardwood xylans are entirely devoid of arabinose. Although small proportions of such residues (<0.5%) occur in all hardwoods, they are probably derived from other polysaccharides such as arabinogalactan or pectic materials. Hardwood xylans are quite stable in alkaline solutions at room temperature. For quantitative isolation of hardwood xylans, the wood has to be first delignified, after which the holocellulose is extracted with alkali. All hardwoods contain the same type of xylan. About 70% of the xylose residues contain an O-acetyl group at C-2, or more frequently, at C-3 [93]. Earlier, it was believed that hardwood xylans contained a linear backbone. Dutton and Unrau [94] were first to demonstrate that hardwood xylans can be slightly branched. Aspen wood 4-O-methylglucuronoxylan contains two branching points at C-3 per average macromolecule [95,96]. Undegraded hardwood xylans contain approximately 200 xylose residues per average molecule. Light scattering and sedimentation equilibrium measurements with a xylan from white birch indicated that the weight average degree of polymerization of this xylan was only slightly higher than the number average value [95]. The xylose residues, like the glucose residue in cellulose, are present in C-1 chain conformation. Using X-ray and polarized infrared techniques, it has been demonstrated that hardwood xylans possess a three fold screw axis with rotation of 120~ and a repeating length of 15 Angstrom. Hardwood xylans can be induced to form single crystals after prior reduction of the frequency of the side chains and lowering the molecular weight [97-100]. The occurence of xylose residues in the ~-D-configuration is evident from the nature of xylooligosaccharides. The isolation of 2,3-di-O-methyI-D-xylose on the hydrolysis of O-methylxylans, and the consumption of approximately one mole of periodate per equivalent of pentose residues show that hardwood xylans, like all xylans from the land plants, contain a backbone of 1,4-1inked D-xylopyranose units. Isolation of 4-O-~-D-xylopyranosyI-D-xylopyranose (xylobiose) and its higher homologs on partial hydrolysis of hardwood xylan provided further evidence [101,102]. Angiospermic woods usually contain 3-5% O-acetyi groups by weight. Woods of silver birch and eucalyptus may contain 5-12% of O-acetyl groups. On methylation of Oacetylxylan, 30% of the acetate groups were found to be lost. Most of the O-acetyl groups have been found to be attached to C-3, and the remainder to C-2, of the xylose. The role of acetyl group is probably to serve the function of decreasing the hydration capacity of the xylan, which could, otherwise, concievably lower the tensile strength of the woody tissue. O-Acetyl groups are slowly hydrolyzed in living tissue to produce acetic acid which effects on acid hydrolysis of the wood leading to a greater ease of fiber

48 separation. Softwoods can not be directly extracted with alkali for the isolation of hemicelluloses. The reason for this is the relatively higher lignin content of the secondary wall in softwood. Composition of hemicelluloses from various softwoods is shown in Table 4.

Table 4 Pentosan composition of various softwoods % of extractive free wood Species

Arabino-4-O-methylglucuronoxylan

O-Acetyl-galactoglucomannan

Anabilis fir Spruce

9.6 13.4

18.5 18.7

Pine Western hemlock Thuja

9.1 7.3 14.7

18.3 16.2 12.1

Softwood xylans have been found to consist of a framework of 1,4-1inked-~-Dxylopyranose residues. Seven out of ten units carry terminal 4-O-methylglucuronic acid residues attached to C-2, rendering this type of xylan similar to that present in hardwood. However, instead of acetyl-substituents, softwood xylans contain o~-Larabinofuranose residues, directly linked to C-3 of the xylose. One arabinose occurs per eight or nine xylose units. Softwood xylans are more acidic than those usually found in hardwoods having five to six xylose residues per acid side chains. Some properties of wood hemicelluloses are shown in Table 5.

49

Table 5 Properties of wood hemicelluloses Hemicellulose

Amount

Composition

Linkage

(%) Hardwood O-Acetyl-4-O-methyl glucuronoxylan

10-35

1~4I3-D-Xylp 4-O-Me-o~-D-GlupA 1,2O-Acetyl

Glucomannan

3-5

13-D-Manp

1,4-

13-D-Glup

1,4-

Softwood Arabino-4-O-methyl glucuronoxylan

10-15

13-D-Xylp 1,44-O-Me-o~-D-GlupA 1,2L-Araf 1,3-

Galactoglucomannan

5-15

13-D-Manp 13-D-Glup o~-D-Galp O-Acetyl

1,41,41,6-

Larchwood Arabinogalactan

10-20

I3-D-Galp L-Araf 13-D-Arap 13-D-Glup

1,3-; 1,61,61,31,6-

Xylans from annual plants have the same backbone as the wood xylans, but they contain lesser proportions of uronic acid. They are highly branched and contain large proportions of L-arabinofuranosyl units mainly linked to the C-2 position [103]. Every

50 seventh xylose unit of a xylan from the internode of wheat was found to be substituted by a L-arabinofuranose side chain [104]. The content of arabinose in xylan from leaves of wheat and barley has been found with a further increase in xylose:arabinose ratio [105]. Xylans from gramineous plants also contain O-acetyl groups [106]. Acetyl groups account for 1-2% of the cell walls of gramineous plants. Cell walls of grasses also contain 1-2% phenolic acid substituents. One in every thirty one arbinose residues of barley straw xylan was found to be esterified with p-coumaric acid, and one in every fifteen with ferulic acid [107]. The xylans from non-endospermic tissues of wheat are acidic arabinoxylans containing low proportions of non-xylose sugar units. Non-endospermic xylans from corn cobs, fibers and hulls display apparently unusual structural features [108-109] and are more complex than those normally isolated from wheat and other grasses. Some of the structural features account for galactose units in the parent hemicellulosic materials [110]. Both D and L enantiomers are present, but it is often assumed that the galactose in plants is the D enantiomer. Non-endospermic xylans from grasses contain low proportions of non-xylosidic sugar units, whereas endospermic xylans are highly substituted by L-arabinofuranosyl groups. Non-terminal, arbinosyl residues in xylans are indicated by the occasional presence of 2,5-di-O-methylarabinose, 2,3-di-O-methylarabinose, and 3,5-di-O-methylarabinose [111,112] in hydrolysates of fully methylated xylans. In non-endospermic xylans,the various non-acidic side-chains and sugar units are mainly attached to 0-3 atoms of xylosyl residues. Enzymatic hydrolysis of xylans from wheat straw and cocksfoot grass all yield O-L-arabinofuranosyl- 1,3-O-13-D-xylopyranosyl- 1,4-D-xylopyranose [113].

3.2

Mannans and glucomannans

Hemicelluloses based on D-mannose as the major structural unit occur in woods and seeds of many plants. Mannans and glucomannans contain linear chains of 1,4-13-Dmannopyranose and 13-D-glucopyranose residues as their main structural features. True mannans have been examined from vegetable ivory [114,115]. Mannans have also been isolated from spruce sulfite pulp and spruce holocellulose [116-118]. Several mannans and glucomannans contain, in addition, a small proportion of D-galactose residues. Two types of mannans may be isolated from ivory nut. Mannan A, extracted with alkali,

51 occurs in granular form [115], and the X-ray diffraction analysis of both native and extracted polysaccharide show distinct crystalline patterns. Mannan B cannot be extracted directly, but it is separated from cellulose by precipitation with cupramonium solution. Xylans from annual plants contain 3-5% of glucomannan. It contains a glucose:mannose ratio of 1:2, although in some species a ratio of 1:1 seems to prevail. Four of the glucomannans so far studied have been found to be linear. It is not yet clear whether the native hemicellulose is partly acylated or not. The glucomannan is degraded during isolation, both by the oxidizing agent used for delignification and by the alkaline solutions used for extraction. When finely divided hardwood is directly extracted with water, approximately 1% of the wood can be recovered in the form of a mixture of watersoluble polysaccharides. Three such polymers, a glucomannan, a 4-0methylglucuronoxylan, and an arabinogalactan have been isolated [119,120].

3.3

Galactans and arabinogalactans

Water soluble arabinogalactans occur in many coniferous wood and are present in largest amount in larches. They are highly branched with, 1,6- and 1,3-1inked Dgalactopyranose residues predominantly. The L-arabinose residues occur as integral parts of arabinogalactans. D-Galactose and L-arabinose are also present in polymeric forms as galctans and arabinans or as arabinogalactans of Type I (4-1inked arabinogalactans). The polysaccharides of these structural types may be isolated as discrete neutral polymers. A second polysaccharide of similar composition to arabinogalactans of Type I, but quite different linkage type is found in the coniferous woods, especially in larches, where it probably does not function as a cell wall component [121]. Arabinogalactan Type II (3,6-1inked arabinogalactans) are now recognized as of more widespread occurence since they are found in association with pectins in dicot cell walls [122], apparently linked to proteins in cereals [123], and further ramifications of outer chains as the major components of many exudate gums [124,125]. The softwood arabinogalactans with higher degree of branching are the most complicated of all wood polysaccharides [126,127]. The backbone of the macromolecule is composed of 1,3-1inked ~-D-galactopyranose residues, all of which carry a side chain attached to its 6-position. The majority of these side chains are composed of 1,6-1inked

52 J~-D-galactopyranose residues, with two such units present per average chain. The ratio of galactose:arabinose was found to be 6:1 for most of the species. The molecular weight of arabinogalactan decreases linearly with the age of the wood from 46,300 to 70,000. Arabinogalactans from western larch is the first wood hemicellulose to find a commercial use [128]. Although galactoglucomannans are the predominant hemicellulose polysaccharides in all softwoods, they were the last to be discovered. The polymer contains galactose, glucose and mannose in a ratio of 1:1:3. Evidently, softwoods contain a whole family of related galactoglucomannans which differ mainly in their hexoses, and especially galactose composition. Some of the hexose units carry a terminal residue of o~-Dgalactopyranose attached to C-6. All galactoglucomannans have been found acylated in their native state [129]. The acetyl content of softwood mannan has been reported to range between 5.9% and 8.8% [130]. Acetyl group substituents have been found at both glucose and and mannose residues [131].

4

ENZYMATIC ANALYSIS OF THE STRUCTURE

Enzymes, depolymerizing polysaccharides, may have an endo or an exo action pattern, and may hydrolyze, or cleave by elimination. Enzymes may be used to detect major linkage types in polysaccharides, and to purify specific polysaccharides from mixtures by selectively depolymerizing contaminants. Modern methods of protein purification allow the preparation of pure enzymes, free of contaminating activities. The availability of size exclusion gels, on which oligosaccharides can be separated, has allowed the isolation of fractions in reasonable quantities. These fractions can then be examined further using enzymatic degradation, methylation analysis, NMR spectroscopy, and mass spectroscopy. Xyloglucans are susceptible to hydrolysis by cellulase, and they yield hepta-, octa-, and nona-saccharide fractions as major components with a 1,4-~-D-glucan backbone [132]. Treatment of xyloglucan with an enzyme preparation containing cellulase, ~-Dgalactosidase and ~-D-glucosidase, yielded 6-O-e~-D-xylosyI-D-glucose, D-glucose and D-galactose. The polymer consists of a main chain of 13-1,4-1inked D-glucosyl residues to which D-xylosyl groups are attached to three out of every four main chain residues (Figure 5).

53

C,~I--0

~ -0

0 0 ~o O~ H~ 00~~

Figure 5. Structure of a portion of hemicellulosic xyloglucan of primary cell wall of dicots. A, arabinose; F, L-fucose; G, D-glucose; Gal, D-galactose; X, D-xylose

Treatment of a hemicellulosic arabinoxyioglucan from the midrib of tobacco leaves with cellulase gives a complex mixture of oligosaccharides [133]. The characterization of these oligosaccharides provided the first convincing evidence for the attachment of o~-L-arabinofuranosyl groups to D-xylose residues. The arabinoxyloglucan was pretreated with mild acid to obtain oligosaccharides having simpler structures, before treatment with cellulase [134-136]. Nine components were separated by gel chromatography of the cellulase digest of intact arabinoxyloglucan. The major fraction contained L-arabinose, D-xylose and D-glucose in the ratio of 1:2:3. Digestion of xyloglucans from soybean and mungbean hypocotyls released oligosaccharides of similar structures, based on two oligosaccharide units, one of which consists of D-glucose and D-xylose, and the other, of D-glucose, D-xylose, D-galactose,

54 and L-fucose [137]. The structure of a disaccharide from a mungbean hydrolysate was later established with Aspergillus oryzae exo-enzyme, with a D-galactosyl group attached to the xylose unit on the penultimate D-glycosyl residue at the non-reducing end [138]. Treatment with oc-D-xylosidase followed by methylation analysis, confirmed that the Dxylosyl group on the D-glucosyl residue at the non-reducing terminus was unsubstituted. Similar oligosaccharides have also been obtained in a digest of soybean xyloglucan [139]. Xyloglucan from cell walls of barley [140], rice [141], mungbean [142,143], and pea [144] have also been characterized using cellulase preparations. Rice cell wall xyloglucan gave an octa- and a penta-saccharide. The anomeric configuration of the terminal D-galactosyl group was established with ~-D-galactosidase. The xyloglucan of rice cells suspension cultures releases cellobiose with a 1,2-13-D-galactosyl group attached to a D-xylosyl residue [145].

5

BIODEGRADATION OF HEMICELLULOSES

Hydrolytic enzymes attacking the hemicelluloses are referred to as hemicellulases (glycan hydrolases, EC 3.2.1) or hemicellulolytic enzymes [146-148]. Typical hemicellulases are D-xylanases, D-galactanases, D-mannanases and L-arabinases. Other related glycosidases such as oc- and ~-D-galactosidases, ~-D-mannosidases, ~-Dxylosidases and ec-L-arabinosidases are excluded from this group because these enzymes are capable of hydrolyzing not only low molecular weight glycosides but also the short chain or monosaccharide appendages from the main hemicellulosic backbone chain [149-151]. Nevertheless, the action of glycosidases is necessary to achieve total hydrolysis of the hemicelluloses, because they act synergistically. The occurrence of hemicellulases is widespread. Microbial hemicellulases have been studied in considerable details (Table 6). They have been reported to be produced by bacteria from terrestrial and marine environments, fungi, rumen microorganisms (protozoa and bacteria), yeasts, xylophagous insects, certain molluscs, crustaceans, marine algae, and in germinating seeds of land plants.

55 Table 6 Important microbial sources of hemicellulolytic enzymes Enzyme

Bacteria

Fungi

D-Xylanases

Bacillus subtilis Clostridium spp. Streptomyces xylophagus

Aspergillus niger Fusarium oxysporum Aspergillus wentii Chaetomium globosum Neurospora crassa Trichoderma koningi Sclerotium rolfsfi

L-Arabinases

Clostridium felsinium Bacillus subtilis

Aspergillus niger Corticium rolfsfi Penicilfium expansum Rhizopus solani Fusarium oxysporum

D-Galactanses

Bacillus subtilis Butyrivibrio sp. Cellulomonas sp.

Bysochlamus fulva Rhizopus niveus Sclerotium rolfsfi Penicilfium expansum Sclerotinia sp. Trametes gibbosa

D-Mannanases

Aerobacter sp. Bacillus subtilis Streptococcus sp. Cellulomonas sp.

Aspergillus niger Aspergillus awamori Botrytis cineria Paecilomyces varioti Trichoderma viride Trametes versicolor

56 Most of the bacteria, moulds and yeasts secrete hemicellulases extracellularly, although cell wall bound and intracellular hemicellulolytic enzymes have also been reported [152-156]. They are produced both constitutively and inducively. They can attack hemicelluloses in two ways: (i) the attack by exoglycosidases precedes that of the hemicellulases, removing the side chain substituents, and exposing the glycan chain to hemicellulase action, (ii) the endohemicellulases may attack the regions of the glycan chain that are unbranched or relatively moderately branched.

5.1

D-Xylanases

D-Xylanases (1,4-~-D-xylan xylanohydrolase, EC 3.2.1.8) hydrolyze the 1,4-~-Dxylopyranosyl linkages of xylans such as L-arabino-D-xylan, L-arabino-D-glucurono-Dxylans and D-glucurono-D-xylans, are well characterized endo-enzyme type. However, the existence of exo-enzyme (1,4-~-D xylan xylohydrolase, EC 3.2.1.37) has also been reported [148]. It appears that the xylosidic likages in lignocellulosics are not all equivalent and equally accessible to xylanolytic enzymes. The accessibility of some linkages also changes during the course of hydrolysis [157]. Multiple xylanases have been found in the enzymes of microbial origin. Xylanases of bacterial origin are least studied because eukaryotic microbes such as filamentous fungi are better xylanase producers. Most of the studies are confined to Bacillus and Streptomycesxylanases. Alkalophilic Bacillus sp. C-59-2 produces xylanase which degrade rice straw arbinoxylan to xylobioses and xylotriose as major end products with smaller amounts of higher xylooligosaccharides [159]. D-Xylose was not detected within short digestion period (lh). 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 hydrolyses. An arabinose containing xyloside was also detected in the hydrolyzate [160]. A purified xylanase preparation from Streptomyces sp. hydrolyzed corn hull and cob arabinoxylans and several different xylans to xylotriose, D-xylose and L-arabinose [161,162]. The Streptomyces xylanases preferentially attack the xylan chain where the degree of branching by arabinose is less. Streptomycesxylanases yield mainly xylotriose in the early stages of the hydrolysis. As the hydrolysis proceeded, xylose oligosaccharides of degree of polymerization 3-5 are further degraded into xylobiose and

57

some xylose. Prolonged hydrolysis (120 h) showed that xylobiose was further degraded with xylose being major end product of the hydrolysis of xylan. The mechanism of enzymatic degradation of heteroxylans is shown in Figure 6.

~A

ARABINOXYLANS GLUCURONOXYLANS RABINOGLUCURONOXYLANS J

I

Endo-13-xylanase o~-Arabinosidase

~Xy

XYLAN OF LOW DS LOSE-OLIGOSACCHARIDES Endo-13-xylanase 13-Xyiosidase o~-Arabinosidase ~-Giucuronidase

XYLOSE 1 ARABINOSE GLUCURONIC ACID Figure 6. Enzymatic hydrolysis of heteroxylans

Yeasts belonging to the genera Aureobasidium, Cryptococcus, and Trichosporon have been recognized to produce xylanases [163]. T. cutaneum produces a single xylanase (MW 45 kDa) which degrades several xylans but not cellulose or xylobiose [164]. Degradation of oat husk arabinoxylan yielded an array of xylose oligosaccharides, and L-arabinose after short (15 min) hydrolysis time. The major end products were found to be xylose, xylobiose and xylotriose. C. albidus produces an extracellular endotype xylanase which exhibts a low affinity for p-nitrophenyl-13-D-xyloside[165]. Xylobiose was

58 attacked at a very slow rate by this xylanase and cleavage was detected only when radioactivity labelled [1-3H]xyl2 was used [166]. Existence of two types of endoxylanases has been demonstrated in fungal xylanases [167]: debranching or arabinose- releasing xylanases and nondebranching or xylotriose-cleaving xylanases. Both types of xylanases are capable of attacking glucuronoxylans and unsubstituted 1,4-~-D-xylans. Takenishi and Tsujisaka [168] demonstrated that one of the three xylanases purified from Aspergillus niger, was responsible for the hydrolysis of xylotriose, while another was responsible for the hydrolysis of arabinosyl substituents. Hydrolysis of arabinoxylan by xylanase I resulted in the accumulation of arabinoxylotriose and arabinoxylobiose which could be removed by the addition of xylanase I1. Hydrolysis by xylanase II, however, resulted in the accumulation of xylotriose and larger oligosaccharides, which were easily hydrolyzed by xylanase I. The nondebranching xylanases degrade heteroxylans randomly. Five xylanases of different specificities were isolated from a commercial enzyme preparation (Rhozyme HP-150) that did not debranch arabinoglucurono xylans [169]. One of these (20.8 kDa, pl 6.7) degraded heteroxylans and xylose oligosaccharides to mainly xylobiose and xylose. Four other xylanases (12-28 kDa) degraded heteroxylan to xylose oligosaccharides but without liberation of xylose. Substrate binding sites of these enzymes were relatively large (>5 subunits). Vrsanska et al. [170] studied the quantitative binding and hydrolysis of xylose oligosaccharides by A. niger xylanase. It did not degrade xylobiose or aryl-~-D-xylopyranosides. The substrate binding site was found to be composed of seven subunits. Bond cleavage frequencies of oligosaccharides by this enzyme were dependent on the substrate concentration. Hydrolysis of xylooligosaccharides occurred via a unimolecular mechanism, with the result xylotriose yielded xylose, xylotetraose yielded xylobiose, and xylopentaose yielded xylobiose and xylotriose as major end products. Sporotrichum dimorphosporum degrades redwood arabinoglucuronoxylan (Figure 7) to mainly xylose, xylobiose, arabinoxylobiose, arabinoxylotriose, glucoarabinoxylotriose, and glucoarabinoxylotetraose [171]. Although L-arabinose and D-glucuronic acid were identified at the branch point on the reducing end of the D-xylosyl chain of the oligosaccharides, they were not released on hydrolysis. Wood-rot fungus Trametes hirsuta produces a xylanase (23 kDa) which degrade willow 4-O-methyl glucuronoxylan to mainly xylotetraose, xylopentaose and 4-O-methyl derivatives [172].

59

Figure 7. Xylose oligosaccharides released during enzymatic hydrolysis of heteroxylans by endo-l,4-13-D-xylanase from Aspergillus niger, Oxiporous sp. and Sporotrichum dimorphosporum [50]. X, 1,4-1inked ~-D-xylosyl residue; and U, 1,2-1inked 4-O-methyl-e~D-glucopyranosyl uronic acid residue; arrow, site of cleavage

Cooperative interactions among multiple xylanases from N. crassa, S. exofoliatus, T. byssochlamydoides and T. harzianum have been demonstrated [173]. They can increase the extent of hydrolysis of xylan. Cooperative interactions involving all the three xylanases from T. harzianum are required to achieve maximal hydrolysis of deacetylated and acetylated xylan. Several xylose oligosaccharides containing 1,2-1inked o~-Dglucopyranosyl uronic acid residues have been isolated from enzymatic hydrolysates of different heteroxylans. Hydrolysis of corn cob arabinoxylan [162] released arabinoxylotriose which contained an interposed L-arabinose residue attached to a terminal D-xylose unit. Table 7 shows major degradation products released during the hydrolysis of xylans.

60 Table 7 Major degradation products released during the hydrolysis of xylans by endoxylanases Source of xylanase

Source of xylan

Degradation productsa

Reference

Aspergillus niger

Rice straw

X,AX2,X2, AX3,AX4 X,X2-X4, (A-X) A,Xl-X3,AX5

[184]

Corn cobs Rice straw arabinoxylan

[185] [185]

Ceratocystis paradoxa

Spear grass

X,X2-X5, AX2-AX5

[ 186]

Diploidia viticola

Corn cob

X2-X5

[186]

Stereum sanguinolentum

Rhodymem ia

X2,X3 (X4-X 10)

[ 187]

Trichoderma viride

Commercial xylan

X,X2-X5

[ 188]

Commercial cellulase

Wheat straw

X,X2-X5

[189]

"A, arabinose; X, xylose; AX, arabinoxylan; A-X, an arabinoxylo-oligosaccharide

5.2

L-Arabinnanase

L-arabinnanases are the enzymes capable of hydrolyzing L-arabinans. They hydrolyze both the 1,3-o~-L-linked L-arabinofuranosyl appendages of sugarbeet L-

61 arabinan, and the 1,5-o~-L-linked L-arabinofuranosyl residues of lower chain [174]. LArabinnanases have been reported to be produced by anaerobic bacteria, saprophytic and phytopathogenic fungi, rumen bacteria, protozoa, snails and plants [175]. Most of L-arabinnanases of fungal origin are usually secreted extracellularly into the medium in which the organism is grown, but intracellular L-arabinnanases have also been reported [176]. Two types of L-arabinnanases (exo and endo) have been characterized. Most of the L-arabinan-degrading enzymes studied have been of exo type. Endo type Larabinnanase are produced by bacteria like Clostridium felsinium and fungi such as Botrytis cineria, Gleosporium kaki and Sclerotinia sclerotium. They hydrolyze sugarbeet L-arabinan to L-arabinose and L-arabinose oligosaccharides as major products. Exo type L-arabinnanases degrade L-arabinan completely to L-arabinose. Purified enzyme preparations from A. niger[177] and Corticium rolfsii[174] were shown to hydrolyze both 1,3- and 1,5-o~-L-arabinofuranosyl residues of sugarbeet L-arabinan, and 1,3-o~-Larabinofuranosyl residues of wheat L-arabino-D-xylan and gum arabic.

5.3

D-Galactanase

D-Glactanases are capable of hydrolyzing D-galactans and L-arabino-D-galactans (Figure 8). D-Galactanases specific for 1,3- and 1,4-13-D-galactopyranosyl linkages have been reported [178]. D-Galactanases degrade D-galactan randomly to release Dgalactose and D-galacto-oligosaccharides. They are mostly endo type. Although an exogalactanase has been reported from Sclerotium rolfsii, exo type D-galactanases have not been unequivocally characterized. D-Galactanases have been reported to be produced by fungi, Bacillus subtilis and rumen anaerobic bacteria. Microbial galactanases are inducible and produced extracellularly in response to the carbon source of the culture medium. An unusual 1.4-13-D-galactanase (40kDa) isolated from B. subtilis, releasing mainly galactotetraose from 1,4-13-D-galactan, was shown to possess both exo and endo activity [179]. The purified D-galactanase (GI-III) from B. subtilis var. amylosacchariticus was found specific for 1,4-1~-D-galactopyranosyl linkages [178].

62

(

ARABINOGALACTANS"~ GALACTANS ..... j Endo-~-galactanase ~-Arabinosidase 13-Galactosidase

IG'" GALACTANoF LOW DS " ALACTOSE-OLIGOSACCHARIDES I

Endo-13-galactanase 13-Galactosidase o~-Arabinosidase ( GALACTOSE ) ARABINOSE

Figure 8. Enzymatic hydrolysis of heterogalactans

Rhizopus niveus produces four D-galactanases which hydrolyze coffee

arabinogalactan in the range of 6.5-14%. One of these F-Ill, hydrolyzes arabinogalactan to L-arabinose, D-galactose, galactobiose and a series of mixed arabinogalactooligosaccharides [180]. This indicates the multisubstrate specificity of the D-galactanase. Galactobiose is not attacked by F-Ill, showing enzyme specificity only to 1,3-13-Dgalactopyranosyl linkages. The enzyme also removed L-arabinofuranose from arabinogalactosides I, III and V but did not liberate any L-arabinose from oligosaccharide IV (arabinose:galactose, 1:3).

63

5.4

D-Mannanase

D-Mannanase (1,4-13-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-Dmannans (Figure 9). The highly purified enzyme preparations from B. subtilis and A. niger have also been shown to be 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 reported to be produced by various species of bacteria including intestinal and rumen bacteria and fungi. Microbial D-mannanases have been reported to be both inducive and constitutive, usually being secreted extracellularly [181].

GALACTOGLUCOMANNANS GLUCOMANNANS ) II

I

Endo-13-mannanase o~-Galactosidase

-

GLUCOMANNAN MANNOSE-OLIGOSACCHARIDES ) Endo-13-mannanase 13-Glucosidase oPMannosidase 13-Mannosidase

1

I GLUCOSE MANNOSE GALACTOSE

Figure 9. Enzymatic hydrolysis of heteromannans

64 Hydrolysis of plant cell wall mannans by endo-l,4-13-D-mannanase from bacterial, fungal and plant origin have been studied. Endomannanases hydrolyze 13-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 [182]. Cellobiose was not detected as hydrolysis product. Galactoglucomannans from spruce and Canadian hemlock were degraded by A. niger endomannanase to mannobiose, mannose, glucose, and mannose oligosaccharides [183]. Several mannose oligosaccharides conaining glucose and galactose have been identified in enzymatic hydrolysates of Canadian hemlock.

6

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3 Extraction of Pentosans from Lignocellulosic Materials

1

INTRODUCTION

Whether used as an energy source or chemical feedstock, optimal utilization of lignocellulosic materials would demand a pretreatment that may consist of a complete or partial fractionation [1]. Several approaches including enzymatic, physical and chemical or a combination of these have been explored in order to obtain each of the polymeric components of lignocellulosics in maximum yield and purity, and to produce low cost sugars (hexoses and pentoses) for use in biotechnological routes to fuels, chemicals and protein-rich feed for ruminants. Hemicellulose is considerably easier to hydrolyze than cellulose, therefore it is possible to hydrolyze the hemicellulose fraction selectively from the biomass [2]. The separation of lignocellulosic components is possible only when hydrogen bonds between the constituents and the hemicellulose-lignin ester crosslinkings are broken. Swelling agents such as water, alkali, ammonia, and certain salts are used to break hydrogen bonds. In the case of ester cross-links, chemical reactions involving acids or bases are used [3]. Lignocellulosic materials can be hydrolyzed by enzymatic processes to produce monomeric sugars in high yields, but the feedstock pretreatment and the enzyme production steps are currently very expensive. Acids and alkalies in particular are rather inexpensive and rapidly hydrolyze polysaccharides present in lignocellulosics. Therefore, for enzymatic processes to compete, the cost of enzymes must be low, the rate of hydrolysis must be rapid, and high yields must be achieved.

2

ENZYMATIC TREATMENT

Biotechnological exploitation of the hydrolytic products from xylanase action require appropriate functional, physical, and chemical properties. The judicious use of proper 71

72 mixes of xylanolytic enzymes could result in higher yields, and lower consumption of enzyme and energy for economically feasible industrial processes. For lignocellulose bioconversion process, maximal utilization of the various polymeric sugars is desirable. Complete xylanolytic (and cellulolytic) enzyme systems are required to achieve maximum hydrolysis of complex substrates to yield monomeric sugars. Important criteria for industrial implementation also include the availability of inexpensive and highly active enzyme preprations in bulk quantities. In certain bioconversion processes, complete hydrolysis may not be required. For instance, fermentative organisms such as Klebsiella pneumoniae can utilize disaccharides like xylobiose, and the limitation of hydrolysis associated with product inhibition can be relieved by using a sequential coculture or simultaneous saccharification and fermentation [4,5]. The accessibility of the enzymes in wood or pulp may be limited due to many factors. It has been reported that the main factors limiting the access of enzymes in woody materials are the specific surface area, fibre porosity and the median pore size of fibers [6]. In addition to these, the molecular organization of other components of the wood or pulp matrix (cellulose and lignin), may limit the accessibilty of the substrate to enzymes [7-9]. Dissolving pulp is one of the products of the pulp and paper industry used for manufacturing of rayon, cellophane, carboxymethyl cellulose, plastics and other cellulose derivatives [10]. In contrasts to pulps used for papermaking, the hemicelluloses in dissolving pulp are undesirable. They are currently removed during cooking of the wood and subsequent bleaching. However, part of the hemicelluloses remain in the pulp. Impurities are responsible for the poor cellulose derivatization. Maximum removal of residual xylan from dissolving pulp could facilitate further steps and improve the final quality of product [11]. Multiple xylanases can be used to modify pulp properties and enhance bleaching of the pulp [12-14]. Pentosans removed from the pulp can be utilized in fermentation processes. Christov and Prior [15] studied the ability of a crude enzyme preparation from Aureobasidium pullulans to hydrolyze xylan from sulfite dissolving pulp. The main degradation product was found to be xylose. The degree of pentosan removal is dependent on time and enzyme concentration, and is limited upto 31% (Table 1). Restricted pentosan removal from bleached sulphite pulps (about 25%), treated with xylanase of Schizophyllum commune (433 U/g for 24 h), has been reported [12]. The enzyme preparation of Saccharomonospora viridis solubilized only 20% of the xylan from bleached kraft pulp if three sequential 24 h xylanase treatments were applied [16]. However, trichoderma harzianum xylanase was shown to remove 54% of xylan from

73 bleached kraft pulp with 500 U/g) for 24 h [17]. There may be several factors which may affect the removal of pentosans from dissolving pulp. The more important factors may be the wood species, quantity of pentosans in pulp, the penetration capability and substrate specificity of enzymes and the linkage of xylan to the cellulose and lignin [18,19].

Table 1 Enzymatic removal of pentosans from sulfite pulp a Enzyme source

Enzyme loading (U/g)

Pentosan solubilization (%)

Reference

Aureobasidium pullulans Schizophyllum commune Saccharomonospora viridis Trichoderma harzianum

433

25

[12]

313

20

[16]

500

54

[ 17]

1500

31

[ 15]

a Hydrolysis time, 24 h

In kraft pulping process, the composition of hemicellulose is extensively modified. During the heating period of the kraft cook, when the alkali concentration is comparatively high, part of xylan is dissolved in the pulping liquor. As the cook proceeds the alkali concentration decreases and partly degraded short-chain xylan precipitates in a more or less crystalline form on the surface of cellulose microfibrils [20,21]. Application of xylanases for improving bleachability of kraft pulps has been shown to be an effective means of decreasing the consumption of chlorine chemical, increasing the final brightness and obtaining hemicellulose sugars for bioconversion [22-25]. According to

74 the proposed mechanism of enzyme-aided bleaching, xylanases are believed to act mainly on the reprecipitated xylan on the surface of the microfibrils. The removal of this xylan renders the fibre structure more permeable for extraction of lignin in the subsequent chemical bleaching. Since the pulping processes are carried out at a high temperature and high pH values, thermal and pH stable enzymes are required in order to make the enzymatic process technically and economically feasible [26]. Table 2 shows the effect of pH and temperature on the hydrolysis of different kraft pulps with thermostable xylanase preparation from Dictyoglomussp.

Table 2 Effect of pH and temperature on the hydrolysis of pine kraft pulp by thermostable xylanase preparation from Dictyoglomus sp. [26]

Variable

Composition of solubilized carbohydrates (% dry wt) Xylose

pH 6 7 8 9

0.90 0.90 0.74 0.74

Arabinose

tr tr tr tr

Glucose

0.05 0.05 0.05 0.05

Temperature 60

nd

nd

nd

70

nd

nd

nd

80 90

0.90 0.96

tr Tr

0.05 0.10

Enzyme loading, 500 nkat/g pulp; incubation time, 2 h; tr, traces; nd, not detected

75 Maximal hydrolysis of pine kraft pulp was found at 90~ The maximum amount of xylan solubilized from pine pulp were 0.96% of dry weight, corresponding to 15.5% of the pulp xylan. The main hydrolysis product from pine pulp at 80~ was xylose, whereas at 90~ the relative amount of xylooligosaccharides increased. The pulp was optimally hydrolyzed at pH 6-7, even at pH 9 the hydrolysis yield was decreased by only 18%. The degree of enzymatic solubilization of pulp xylan generally does not exceed 20% of the theoretical value due to the poor accessibility of xylan in fibrous materials [27]. Viikari et al [28] studied the hydrolysis of fibre-bound and isolated xylans from both birch and pine wood and kraft pulps using purified preparations of xylanolytic enzymes from Trichoderma reesei (Table 3). Despite high enzyme loading (5000 nkat/g xylan in the substrate), the degree of hydrolysis of fibre-bound substrates did not exceed 20% of the theoretical value. The fibre-bound xylans were equally accessible in softwood as in hardwood pulps. The isolated xylans of wood and kraft pulps could be solubilized more extensively, with a hydrolysis yield of 50-65%.

Table 3 Hydrolysis of fibre-bound and isolated xylans from birch and pine wood and kraft pulps by xylanolytic enzymes of Trichoderma reesei [28] Substrate

Type

Hydrolysis

(%) Birch

Pine

Wood Xylan from birch wood Birch kraft Xylan from birch kraft Wood Pine kraft Xylan from pine kraft

Enzyme loading, 5000 nkat/g of xylan in the substrate

5 63 20 52 1 19 55

76 3

PHYSICAL TREATMENT

Physical methods for pretreatment of lignocellulosics may be of two types: mechanical and irradiation (Table 4). Mechanical treatment methods include grinding and milling which utilize shearing and impacting forces to yield a fine substrate having a low crystallinity index [29]. A high slurry concentration can be achieved by using fine substrate which reduces the reactor volume.

Table 4 Physicochemical methods for the pretreatment of lignocellulosic biomass Chemical

Chemical

Alkali Gases Sodium hydroxide Ammonia Ammonium hydroxide Chlorine Acids Nitrous oxide Sulfuric Ozone Hydrochloric Sulfur dioxide Nitric Solvents Phosphoric Ethanol Oxidizing agents Butanol Peracetic acid Phenol Sodium hypochlorite Ethylamine Sodium chlorite Acetone Hydrogen peroxide Ethylene glycol

Physical

Thermal

Mechanical Ball milling Fitz milling Roller milling Hammer milling Weathering Irradiation Gamma Electron beam Photooxidation

Autohydrolysis Steam explosion Hydrothermolysis Boiling Pyrolysis Moist heat expansion Dry heat expansion

77

3.1

Milling

Milling helps in the distribution of reactants throughout the material [30]. Ball milling is an effective means of pretreatment [31]. In addition to reducing particle size, ball milling disrupts the crystalline structure and breaks down the chemical bonding of long chain molecules. It has been observed that the effectiveness of milling depends on the materials to be processed, softwoods are least responsive. Compression milling not only reduces the crystalline structure and crystallite size of polysaccharides but also changes specific surface area and significantly varies the degree of polymerization [32]. It also increases the accessibilty of substrate to enzymatic hydrolysis. Two-roll milling is commonly used in the rubber and plastic industries for grinding raw materials [33]. The mill consists of two cast-iron tempered surface rolls. The lignocellulosic materials are fed into the roll, masticated for a specific period of time and then the pretreated material is scrapped off. Tassinari and Macy [34] tested two-roll milling on various lignocellulosic substrates. They observed that two-roll milled maple chips yielded 17-times more reducing sugars than untreated maple. Two-roll milled newspaper exhibited a 2.5-fold increase over ball-miled newspaper. It was also found that the sedimentation volume is lower for two-roll milled newspaper than ball-milled newspaper. This allows for a higher slurry concentration in the hydrolysis vessel, thereby reducing the reactor volume and lowering the capital costs. Two-roll milling significantly decreases the degree of polymerization and crystallinity, but its effect on lignin is not well understood. Factors that control the susceptibility to enzymatic attack are the clearence between the mill rolls and the processing time. As the clearence between the rolls decreases and the processing time increases, the susceptibility to hydrolysis increases [35]. Pretreatment using colloid milling has also been attempted [36,37]. A colloid mill consists of two disk sets close to each other revolving in opposite directions while the substrate slurry is passed between the disks [35]. Mandels et al. [37] obtained modest improvements in the susceptibilty of cellulose to cellulolytic enzymes. However, operational cost makes this pretreatment uneconomical. Sweco milling has been found to be species specific, as hardwoods are somewhat more affected than softwoods [38]. Millet [39] achieved 63.9% saccharification after 24 h of hydrolysis for Sweco-milled newsprint. Ghose [40] obtained 1.7-fold increase in the reducing sugars for Sweco-milled Solka Floc over untreated Solka Floc. Vibro energy milling is similar to the ball milling except that the mill is vibrated instead of being rotated. Vibro milling provides effective

78 means for size reduction and increases digestibility of lignocellulosics [41-43]. Pew [44] observed enhanced susceptibilty of spruce and aspen woods towards the enzymatic hydrolysis. Although, hammer milling gives good size reduction and increases bulk density, hydrolysis susceptibility gain is insignificant [45]. Fine Fitz milling substantially reduces the size of substrate but it affects the crystallinity index only slightly. Fluid energy milling, at a higher energy input reduces the particle size and increases the susceptibility [46]. Wet milling is much less effective than dry milling. Although milling is considered a good way of increasing substrate reactivity to enzymatic saccharification, energy wise the methods are unattractive [47]. The major factor is the fibrous nature of lignocellulosic materials.

3.2

Irradiation

Polysaccharides of lignocellulosic materials undergo extensive depolymerization by irradiation treatment, thus increasing specific surface area for enzymatic attack. The primary effect of high energy radiation is chain cleavage. The ensuing decomposition of the formed carbohydrates results in the formation of acidic and reducing groups [48]. Irradiation also appears to affect the lignin of the lignocellulosic biomass as is evident from the increased presence of phenolic groups in irradiated wood fibre [49,50]. Radiation also causes an apparent decrease in crystallinity and increases the digestibility of lignocellulosics. High energy electron irradiation provides an effective means of enhancing the digestibility of the carbohydrates in wood [51-54]. At 100 Mrad gamma irradiation, the crystallinity index of sugarcane bagasse was found to be reduced from 66.6% to 44.4%. Studies on various lignocellulosic materials have revealed that rice straw has maximum digestibility when treated at 5X108 rad, and the optimum dose for wheat straw digestibility has been worked out to be 2.5X108 rad [55]. Lower dose (10 Mrad) of gamma irradiation did not contribute much to the pretreatment of lignocellulosic substrates [56]. The hydrolysis rate increases only when the radiation dose reaches a certain level. In addition to depolymerization, radiation produces change in the susceptibility of cellulosic materials to subsequent acid/enzymatic digestion [57]. When bagasse is acid hydrolyzed after radiation treatment, a three times higher sugar yield is obtained as compared to untreated

79 substrate [58]. Gamma irradiation is very effective in increasing specific surface area [59]. The increase in surface area is primarily due to the extensive depolymerization. Pentoses and hexoses are generated as depolymerization products [59-63]. Irradiation appears to be strongly species specific. For instance, the digestion of aspen polysaccharides is essentially completed after an electron dosage of 108 rad, while spruce is only 14% digestible at this dosage [64]. The digestibility can be increased by milling the substrate before irradiation, adding sodium salts prior to irradiation [65] or in the presence of oxygen [66]. Despite the use of irradiation pretreatment in enhancing the saccharification of polysaccharides, it holds little promise for commercial application because of high investment costs.

4

CHEMICAL TREATMENT

Chemical treatment methods have been extensively used for delignification and structural modification of lignocellulosic materials. Although, chemical treatment methods are effective, waste chemicals are difficult to recycle or dispose of.

4.1

Alkali

The dilute alkali treatment of lignocellulosic biomass causes swelling, decreases the degree of polymerization and crystallinity, separates lignin, and disrupts lignin structure [67-71]. Saponification of intermolecular ester bonds promotes the swelling of cellulose and favours enzyme penetration into the cell wall [72,73]. The concentration of NaOH in alkali treatment varies from 0.1 to 0.15 g / g solid [74-77]. However, the optimum level of NaOH in treating substrates is the subject of much controversy, as different researchers have indicated different optimum levels. It is also possible to decrease the requirement of alkali by means of presoaking the substrate [78]. Pretreatment of agricultural residues with NaOH and NH4OH decreases the neutral detergent fibre content of the material to varying degrees [79]. Hardwood hem icellu loses are more stable and hence can be removed using cold

80 alkali treatment, this treatment increases the average pore size in the cell wall structure of lignocellulosics [80]. A combination of acid (H2SO4 or HCI) prehydrolysis with alkali (NaOH) delignification has also been attempted [81]. However, this method leads to significant losses of polysaccharides. Timell [82] has described a method for isolation of softwod hemiceilulose using chiorous acid, potassium and barium hydroxide (Figure 1).

WOOD I

Chlorous acid

HOLOCELLULOSE Potassium

hydroxide

SOLUBLE HEMICELLULOSE MIXTURE Barium

MIXTURE Barium

hydroxide

ARABINOGLUCURONOXYLAN

RESIDUE

hydroxide

INSOLUBLE GALACTOGLUCOMANNAN

INSOLUBLE GALACTOGLUCOMANNAN

Figure 1. Isolation of softwood hemicellulose Cunningham et al. [83] examined the hemicellulose isolation from monocots (wheat straw and sweet sorghum bagasse) and a prolific dicot biomass source, kenaf (Table 5). Treatment of these milled materials with a 12% NaOH solution for 4 h at 80~ extracted

81 88-90% of the pentosans from wheat straw and sweet sorghum bagasse, and 80-84% from the kenaf bark and core. Ethanol precipitation of the filtrates recovered 90% of the extracted pentosans from wheat straw and sweet sorghum bagasse, and 66-76% of those extracted from kenaf bark and core. Both hemicellulose A and B could be precipitated using this method. Hemicellulose A is the fraction that precipitates by the neutralization of the alkaline extract and the hem icellu lose B fraction remains dissolved until precipitated with ethanol. This method has an advantage of recycling ethanol for further isolations.

Table 5 Characteristics of alkali-extracted agricultural residues [83] Analysis

Wheat straw

Sweet sorghum bagasse

(%) Untreated Yield b

Treated"

Untreated

34.7

Treated 32.6

Cellulose

30.3

79.5

27.2

83.2

Pentosans

25.0

7.5

23.9

8.8

Lignin

18.0

8.6

9.1

4.7

Ash 11.0 Precipitated liquor solidsc

2.5

2.3

1.3

Yield Pentosans Lignin Ash

43.4 46.3

28.0 67.4

6.2

4.5

38.5

24.3

" Substrates were treated with 12% NaOH (NaOH solution:straw, 10:1) for 4 h at 80~ b (Insoluble fraction weight/sample weight) X 100 c Solids precipitated by addition of ethanol to filtrates

82 Ammonium sulfite is conventionally used in pulping processes. Clarke and Dyer [84] developed a modified process to increase the digestibility of lignocellulosic materials for animal feed. In this process, Douglas fir is made to pulp by treating with ammonium sulfite under high pressure and elevated temperature. The resultant pulp had a residual lignin content of 15% and a dietary energy equivalent of medium quality hay when fed to steers at up to 70% of their total ration. Anhydrous ammonia in either liquid or gaseous form is a strong cellulose swelling agent [85]. It can affect a phase change in the cellulose fibre structure from cellulose I to cellulose III [86]. It can also react with lignoceilulosics by ammonolysis of the ester crosslinks of uronic acid with the xylan units, cleaving the bond linkages between hemicellulose and lignin, and cleaving the C-O and C-C bonds of lignin macromolecules to produce smaller soluble fragments [87-89]. Earlier studies on ammonia treatment mostly aimed at enhancing the substrate digestibility. Lehman [90] was first to patent this treatment method for increasing the digestibility of straw. Treatment of biomass with ammonia has been shown to significantly enhance the susceptibilty of hardwoods and softwoods to enzymatic hydrolysis by cellulases [85]. The possible determinants of wood degradability which might be altered by ammonia are: extent of wood lignification, cellulose or hemicellulose structure, extent of lignin-carbohydrate bonding, and pore size and its distribution [91]. Dale and Moreira [92] used relatively mild conditions (30-600 psi, 10-30 min) to pretreat agricultural residues with either gaseous or liquid ammonia in a closed reactor. Explosive release of ammonia from the reactor provided the substrates with an expanded fibre structure. Moore et al. [93] treated aspen with liquid ammonia and found that the percentage yield of reducing sugars increased from 11% for untreated aspen to about 36% for ammonia treated aspen. However, Waiss [94] achieved only 10% increase in the enzymatic digestibility of aspen after pretreatment with 5% ammonia at room temperature for 30 days. Supercritical fluids, often called dense gases, exist in a state above the gas-liquid critical temperature and pressure. Because of their strong dissolving and penetrating power, supercritical fluids have been utilized in extraction and liquefaction of wood at ambient temperature and pressure [95,96]. Supercritical acetone has been used to liquify cellulose with levoglucan as the major product [97]. Supercritical water exists at 374~ and 218 atm with densities varying from 0.2 to 0.7 g/ml [98]. It has been demonstrated that supercritical water, at near critical conditions, gasify and liquify wood without char formation [98]. Weimer et al. [95] demonstrated that pretreatment of hardwood biomass with supercritical ammonia remarkably enhanced the susceptibilty of substrate to

83 enzymatic hydrolysis. The treatment increased the substrate nitrogen content and total pore volume, but the chemical composition was not much affected (Table 6). Optimal supercritical ammonia treatment conditions for birch occurs at 150~ an ammonia density of 0.13 g/ml reactor volume or more, and a treatment time of 5 min or less [85]. Near theoretical conversion of cellulose and 70-80% conversion of hemicellulose to sugars from hardwood and agricultural by-products were obtained.

Table 6 Composition of supercritical ammonia-treated wood materials [95] Supercritical ammonia

% Dry wt

treatment

Hexosans

Pentosans

Lignin

Uronic acid

Soft maple

No Yes

40.5 44.3

16.7 17.1

19.4 21.9

5.3 4.8

Southern red oak Douglas fir

No Yes No Yes

49.2 45.1 54.9 57.1

19.5 17.2 4.5 3.4

21.1 24.4 33.5 32.8

3.8 2.8 1.5 1.0

Wood

Wang et al. [88] have shown that pretreatment of both hardwoods and softwoods with anhydrous ammonia under mild conditions (25~ 72 h) resulted in the formation of hemicellulose amides containing small amounts of bound lignin. Aspinall [99] has given the evidence of other types of lignin-carbohydrate bonds, e.g. between the carboxyl groups of ferulic acids and the hydroxyl functions of arabinoxylans, some of which may be susceptible to ammonolysis. Evidence for extensive ammonolysis of hemicelluloses is provided by the reported production of acetamide during tretment of wood with supercritical ammonia or by an ammonia explosion process [100]. This reaction may be viewed as deacetylation of hemicellulose, resulting in wood with increased swelling ability due to the retention of relatively insoluble, low molecular weight, deacylated

84 hemicellulose components within the wood matrix. Timell and Zinbo [101] developed a method for hemicellulose isolation from populus wood. Debarked wood was ground to in a Wiley mill and extracted with benzene-ethanol (2:1) for 24 h. After vacuum drying, extracted wood was shaken in 24% KOH for 3 h at room temperature. After filtration, solubilized fraction was precipitated with ethanol:acetic acid (15:1) and centrifuged. The hemicellulose precipitate was washed successively with 70% and 90% ethanol and dried under vacuum. The product obtained as hemicellulose fraction was ground to fine powder before acid hydrolysis.

4.2

Acids

Under acidic conditions, decomposition of sugars takes place, and xylose decomposes five times faster than glucose [102,103]. Xylose is decomposed into furfural, while glucose is chemically transformed to hydroxymethyl furfural. Both types of products are toxic to microbial fermentations [104]. Although steaming can be used to volatilize and remove furfural, the best results can be obtained by passing the hydrolysate through ion exchange columns [105,106]. Acids such as sulfuric, hydrochloric, nitric and phosphoric are generally used to prepare hemicellulose hydrolysate [72]. Datta [107] fractionated corn stover by sulfuric acid treatment. The weight loss at each fractionation step gave the weight of each of the major components in raw materials. AbduI-Halim et al. [108] examined different concentrations of sulfuric acid for solubilization of wood components and achieved best results with 72% concentration at 121~ Brownell and Nakas [109] and Singh and Ghosh [110] studied the acid hydrolysis of poplar and wheat straw hemicellulose, respectively, for further bioconversion (Table 7). Hydrolysis of oat spelt xylan with 3% sulfuric acid released 41 g/I pentose sugars out of total 43.1 g/I of reducing sugars. Hydrolysis of poplar hem icellu lose with 4% sulfuric acid at 100~ for 60 min released only 14.7 g/I reducing sugars. Singh and Ghosh [110] have reported 33.7 and 35.9 g/I fermentable sugars, mainly pentoses, after hydrolysis of wheat straw hemicellulose (100~ for 60 min) with 4% and 5% sulfuric acid, respectively.

85 Table 7 Release of sugars after acid treatment of oat spelt xylan and hemicellulose isolated from natural substrates [109,110] Sulfuric acid (%,v/v) Substrate

Oat spelt xylan Pentoses (g/I) Reducing sugars (g/I) Poplar hemicellulose Pentoses (g/I) Reducing sugars (g/I) Wheat straw hemicellulose Pentoses (g/I) Reducing sugars (g/I)

1

2

3

4

16.5 17.0

34.8 36.5

41.0 43.1

3.0

8.9

13.0

14.0

3.2

9.3

13.5

14.7

5

31.6

34.1

33.7

35.9

Acid treatment for 60 min at 100~ (liquid:solid ratio, 20:1)

An integrated process scheme (Figure 2)involving acid hydrolysis, acid recycling and fermentation of the hydrolysis products has been suggested by Ladisch et al. [111]. In this process lignocellulosic substrates are first hydrolysed by acid treatment to separate pentose and hexose sugars. Acid is recovered and recycled for further hydrolysis. Glucose and pentose sugras obtained from the enzymatic hydrolysis of cellulose and acid hydrolysis of substrate, respectively, enter into the fermentation route. Similar processes have been designed by other groups also [112,113].

86 LIGNOCELLULOSIC BIOMASS

~

Acid

ACID RECYCLE

HYDROLYSIS

LIGNIN + CELLULOSE

PENTOSES

ACID RECOVERY

Cellulase enzyme GLUCOSE + ~ RESIDUAL SUBSTRATE

FERMENTATION

Figure 2. A two-step acid hydrolysis followed by fermentation scheme for biomass processing

Chahal et al. [114] suggested a simple but lengthy procedure for the fractionation of wheat straw. Accordingly, washed straw is treated with 80% aqueous methanol followed by sodium chlorite solution at 75-80~ for 5 h. The residue (holocellulose) is washed with water and then treated with 10% NaOH to separate cellulose and hemicellulose fraction. The washed residue is collected as cellulose. Hemicellulose is recovered after acidifying the extract and precipitating the soluble hemicellulose with 4 volumes of ethanol. The material balance of this method is described in Figure 3. A similar procedure for isolation of xylan from rice straw was also described by Park et al. [115]. Rice straw is cut into small pieces and heated at 121~ for 1 h in 3% NaOH. After filtration, xylan is precipitated by the adition of an equal volume of ethanol to the filtrate followed by air drying at room temperature.

87 WHEAT STRAW (400 g) Sodium

HOLOCELLULOSE 272 g (68%)

chlorite

LIGNIN AND OTHER FRACTIONS 128g (32%)

NaOH Ethanol CELLU LO SE 1 72 g (42.5%)

Precipitation

HEMiCELLULO SE 96 g (24%)

Figure 3. Material balance for the fractionation of wheat straw into its major components by chemical treatment [114]

Ueng and Gong [116] described the preparation of hemicellulose hydrolysate by acid treatment of bagasse. Bagasse (50% moisture) is sprayed with 5.4% (w/v) sulfuric acid prior to packing into a jacketed column. Steam is injected into the column and hydrolysis is carried out at 100~ for 4 h. The hemicellulose extract is then recovered by down flow leaching at 80~ with water at a superficial velocity of 36 cm/h, and neutralized. The dilute acid treatment at high temperature hydrolyzes glycosidic bonds in hemicelluloses and leads to the solubilization of pentose sugars [117-124]. Grohman et al. [125] studied the dilute acid treatment of wheat straw and aspen wood at high (40 wt%) solid concentration (Table 8).

88 Table 8 Composition of high temperature acid pretreated wheat straw and aspen a [125] Composition

Wheat straw

Aspen

Liquor Total sugars

3.7-7.7

3.6-7.3

Acetic acid

0.7-1.4

0.8-1.6

Furfural

0.3-0.9

0.3-1.2

Relative sugar composition Xylose

60-71

74-79

Glucose

16-18

12-16

Arabinose & mannose

10-12

5-9

Galactose Total

3-4

2-4

97-105

93-108

Solid residues Dry wt losses

34-39

26-32

Lignin removal

0-5

0-5

54-57 4-5 31-33

60-66 0-3 26-29

3-5 92-100

3-5 89-103

Relative composition Anhydroglucose Anhydroxylose Klason lignin Water Total

a Solid concentration, 20-40%; acid concentration, 0.45-2.5%; temperature, 140-160~

Various kinetic models have been presented to describe the hemicellulose hydrolysis reactions [126-130]. However, these models and kinetic parameters are very specific to substrates and cover a narrow range of reaction conditions (Table 9). Kim and Lee [129] developed a kinetic model to determine the associated reaction

89 parameters specifically applicable for acid hydrolysis of hardwoods hemicellulose. The kinetics of acid catalyzed hemicellulose was investigated under low water condition of 1:1.6, solid to liquid ratio. The reactions were found to be of first order. The respective rate constants were correlated with temperature and acid concentration using Arrhenius equation with the addition of an acid term in the preexponential factor. Hydrolysis reactions were found to be more sensitive to acid concentration and reaction temperature than the decomposition reaction.

Table 9 Comparison of kinetic models and parameters in acid-catalyzed hydrolysis of pentosans Substrate,

Kinetic model

Preexponential factor ~

Activation energy b

74-147~

(Pentosan)e-->l

2.56X101sC11s

3.09X104

[126]

1-16% Cotton gin

(Pentosan)~-->2

5.75X1014C~~s

125-165~ 0.5-2% Red oak 120-140~ 1-5% Birch

(Pentosan)e-->l (Pentosan)d-->2

3.55X107C 136 10.85C ~

1.83X104 0.56X104

[128]

(Xylan)e--> 1 (Xylan)d-->2

6.00X1012Ch 1"19 1.77X10~lCh ~-~

2.82X104 2.69X104

[129]

(Xylan)e--> 1 (Xylan)e--> 1

2.26X1016Ch 1.16X10~gCh~.S4

3.74X104 2.87X104

[130]

temperature

Reference

H2SO4 (%) Buna

100-170~ 0.04-0.18 mol

Superscripts a and b are the units [(min)~(acid) N~] and (cal/g mol), respectively, whereas subscripts e and d refer to easily and difficultly hydrolyzable fractions, respectively

90 Hydrogen fluoride solvolysis, either in vapour or in solution, effectively hydrolyzes wood to produce sugar fluorides [131,132]. Removal of fluoride results in repolymerization with the formation of water soluble oligosaccharides which can be converted to monomers by weak acid hydrolysis.

4.3

Gases

Treatment with gases has the advantage in that it facilitates uniform penetration throughout the substrate. But a gaseous medium is rather difficult to handle and recovery poses more problems. Several gases such as chlorine dioxide and ozone have been used as pretreatment agents [81,133,134]. Treatment of lignocellulosics with nitrogen oxide (NO) gives a rapid rate of delignification and a high overall sugar yield [135,136]. Treatment of 100 g wheat straw with 5 g NO, followed by addition of 6 g of 02 resulted in a xylose yield of 69% based on the original xylan content of the straw [137]. Treatment of moist lignocellulosic material with SO2 gas at 120~ for 2-3 h is effective in disrupting the lignin-carbohydrate complex [138]. Reactions of various species of wood with SO2 gas for a period of 2-3 h at room temperature converts 7085% of the polysaccharides to simple sugars [38]. However, the pollution is the big problem with SO 2 treatment. Ozone may be another effective gas for treatment without producing excessive pollutants. It attacks both lignin and carbohydrates, though the rate of reaction with latter is slower [139].

4.4

Oxidizing agents

Sodium chlorite, sodium hypochlorite, peracetic acid, potassium iodate, potassium permanganate, potassium perchlorate and hydrogen peroxide are oxidizing agents which cause structural modification of polysaccharides, and carry out chemical oxidation of lignin [140]. Impressive enzymatic hydrolysis [72,141] and bioconversion of the treated substrates [142] have been obtained. Toyama and Ogawa [143] used 20% peracetic acid for delignification of corn stalks, sawdust from broad leave trees and sawdust from

91 coniferous trees. Fan et al. [144] obtained a drastic increase in the digestibility of wheat straw upon peracetic acid treatment. The action of hydrogen peroxide with Fe2§as a catalyst causes cellulosic materials to oxidize and decompose into CO2 [145-147].When wheat straw and corn stover were treated with an alkaline solution (pH 11.5) of hydrogen peroxide, about half of the lignin and most of the hemicellulose is released [113,148]. Because of the mild conditions involved and nontoxic final product, this system appears to be significantly important as a potential pretreatment method [149]. Chemical treatment is considered to be more effective than physical treatment in producing a substrate for bioconversion. However, key factors that controls the viability of the process appear to be the cost of chemical recovery and the effluent problem. Furthermore, expensive materials in the process eqiupment may also be needed [150].

5

THERMAL TREATMENT

Thermal treatment with or without steam has been used for upgrading the digestibility of various lignocellulosic materials [151-156]. Among all the pretreatment methods, steam tretament method has gained much interest and wide acceptance as a highly efficient and economically feasible method. Steam pretreatment may of two types, viz. autohydrolysis and steam explosion, depending upon the treatment conditions. Autohydrolysis uses temperatures in the range of 170-200~ whereas in steam explosion the temperature range extends to 250~ and the treatment ends with a sudden release of pressure [150]. The autohydrolysis process is more effective in the case of hardwoods than softwoods [138]. Hardwoods containing high amounts of acylated xylan have long been successfully treated [157]. Xylan solubilized by steam pretreatment gives rise mainly to oligosaccharides and relatively minor amounts of xylose monomers [158]. Lignocellulosics, when subjected to high pressure steam for a specific period followed by a sudden release of pressure result in extensive disintegration [159]. The hemicellulose fraction can then be extracted with water, alkali or solvents. The autohydrolysis reaction involves the formation of acetic acid from acetyl groups which catalyzes the hydrolysis of hemicellulose and also the breakdown of lignin-ceiluiose complex [160,161]. The release of pressure also causes lignin coalescence and

92 mechanical abrasion of the fibre. The residence time at higher temperature should be kept low to minimize the formation of inhibitory by-products [162-164]. Although, both batch and continuous systems have been developed, because of the precise control of the operating conditions and efficient steam utilization, a continuos process is considered more effective [165,166]. Steam explosion treatment completely solubilizes hemicellulose sugars and also promote enzymatic hydrolysis of cellulose [167,168]. Galbe and Zacchi [169] tested a wide range of pretreatment conditions for the steam explosion of sallow. A temperature of 220~ and a time of 15 min gave the highest yield. Hydrothermolysis has been reported to be an effective pretreatment method for poplar wood and wheat straw for enzymatic hydrolysis of lignocellulosics. In hydrothermolysis treatment, the raw material in water is subjected to high temperature but no steam appears in the process [170]. The treated substrates have been found to produce about 80-90% of the reducing sugar yield in 70 h of enzymatic hydrolysis [171]. Similar to high pressure steaming are moist-heat expansion (extrusion) and dry heat expansion (popping) treatment, both of which have been used to increase the feed efficiency of grain in animal feed [172,173]. Han and Callihan [174] showed that extrusion pretreatment is ineffective in increasing digestibility of rice straw and sugarcane bagasse. However, it has been suggested that extrusion may be a promising pretreatment method for acid hydrolysis [175]. Pyrolysis of lignocellulosic materials has also been investigated to increase the susceptibility of substrate to hydrolysis [176]. Mild acid hydrolysis of tar fractions yielded reducing sugars in the range of 80-85%.

6

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123 Grohman K, Himel M, Rivard, C, Tucker M, Baker J, Torget R, Graboski M. Biotechnol Bioeng Symp 1984; 14: 137. 124 Grohman K, Torget R, Himel M. Biotechnol Bioeng Symp 1985; 15: 59. 125 Grohman K, Torget R, Himel M. Biotechnol Bioeng Symp 1986; 17: 135. 126 Kobayashi T, Sakai Y. Bull Agri Chem Soc Japan 1956; 20: 1. 127 Oshima M. Wood Chemistry, Process Engineering Aspects, New York: NDC, 1965. 128 Beck SR, Wang T. Kinetic analysis of hemicellulose in cotton gin residues, AIChE National Meeting, Orlando, 1982. 129 Kim SB, Lee YY. Biotechnol Bioeng 1986; 17: 71.

97 130 Maloney MT, Chapman TW, Baker AJ. Biotechnol Bioeng 1985; 27: 355. 131 Murray WD. In: (Hasnain Q ed.) 5th Canadian Research and Development Seminar (Hasnain Q ed.), London: Applied Science Publishers, 1984; 255. 132 Murray WD, Asther M. Biotechnol Lett 1983; 5: 175. 133 Saarinen P, Jenson WJ, Alhojarvi J. Acta Agral Fennica 1959; 94: 41. 134 Sullivan JT, Hershberger TV. Science 1959; 130: 1252. 135 Brink DL. 157th Amer Chem Soc Meet, Los Angeles, 1974. 136 Borrevik RK, Wilke CR, Brink DL. Effect of nitrogen oxide pretreatment on enzymatic hydrolysis of cellulose, Lawrence Berkley Lab., Berkley, 1978; LBL7879. 137 Wilke CR. Process Development Studies of the Bioconversion of Cellulose and Production of Ethanol, Lawrence Berkley Lab., 1978; LBL 7880. 138 Clarke TA, Mackie KL. J Wood Sci Technol 1987; 7: 373. 139 Binder A, Pelloni L, Fiechter A. Eur J Appl Microbiol Biotechnol 1980; 11: 1. 140 Singh A, Abidi AB, Darmwal NS, Agrawal AK. MIRCEN J Appl Microbiol Biotechnol 1989; 5: 451. 141 Singh A, Abidi AB, Agrawal AK, Darmwal NS. Z Mikrobiol 1991; 146: 181. 142 Singh A, Abidi AB, Darmwal NS, Agrawal AK. In: (Bhattacharya B ed.) Biotechnology in Agricultural and Rural Development, NIRD, Hyderabad, 1988; 68. 143 Toyama N, Ogawa K. In: (Ghose TK ed.) Proc Symp Bioconversion of Cellulosic Substrate into Energy, Chemicals and Microbial Protein, New Delhi, 1978; 373. 144 Fan LT, Gharpuray MM, Lee YH. 3rd Symp Biotechnology for Energy Production, Tennessee, 1981. 145 Halliwell G. Proc Symp Bioconversion of Cellulosic Substrates into Energy, Chemicals and Microbial Protein (Ghose TK Ed.), New Delhi, 1978; 81. 146 Koenings JW. Biotechnol Bioeng Symp 1975; 5: 151. 147 Elmund GK, Grant DW, Morrison SM. Proc 3rd Int Symp Livestock Wastes, ASAE Publication, 1975; 275. 148 Vallander L, Eriksson K-E. Biotechnol Bioeng 1985; 27: 650. 149 Ghose TK. Adv Biochem Eng 1977; 4: 39. 150 Vallander L, Eriksson K-E. Adv Biochem Eng/Biotechnol 1990; 42: 63. 151 Wayman M, Lora JH. Tappi 1978; 61: 55. 152 Overend RP, Chornet E. Phil Trans Royal Soc London Ser A 1987; 321: 523. 153 Bonn G, Hormeyer HF, Bobleter O. Wood Sci Technol 1987; 21: 179. 154 Bobleter O, Bonn G, Prutsch W. In: (Focher B Marzetti A Crescenzi E eds.) Steam Explosion Techniques, Fundamentals and Industrial application, Philadelphia: Gordon and Breach, 1991;59.

98 155 Dekker RFH. In: Steam Explosion Techniques, Fundamentals and Industrial application (Focher B Marzetti A Crescenzi E eds.), Philadelphia: Gordon and Breach, 1991; 277. 156 Tanahashi M, Tamabuchi K, Goto T, Aoki T, Karina M, Higuchi T. Wood Res 1988; 75: 1. 157 Mamers H, Menz DNJ. Appita 1984; 37: 644. 158 Puls J, Poutanen K. Korner HV, Viikari L. Appl Microbiol Biotechno11985; 22: 416. 159 160 161 162 163 164 165 166 167 168

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176 Shafizadeh F. TAPPI Forest Biology and Wood Chemistry Conference, Madison, 1977.

Microbial Uptake of Pentoses

1

INTRODUCTION

Carbohydrates being the major carbon source for growth and metabolic activities of microorganisms have invigorated many researchers to unravel the molecular mechanisms underlying their transport across the plasma membrane. Amongst carbohydrates, hexoses, particularly glucose, is the most common carbohydrate known to be metabolized by a large number of microorganisms and hence its transport has been studied in greater details. On the other hand, inability of many micoorganisms to utilize pentoses, has restricted studies on microbial uptake of pentoses. However, growing concern for microbial utilization of pentoses obtained from lignocellulosic hydrolysates has led to a world wide interest in screening and improvement of strains that can assimilate and metabolize pentoses. Consequently the number of microorganisms that can utilize pentoses is increasing. A detailed list of pentose fermenting microorganisms is given elsewhere in this volume. Since metabolism of pentoses, like any other nutrient, is initiated by their uptake from the medium hence transport of pentoses across the plasma membrane is likely to play an important role in regulation of their metabolism. In fact using nuclear magnetic resonance spectroscopy it has been demonstrated that D-xylose uptake in Pichia stipitis is the rate limiting step of xylose metabolism under aerobic conditions [1]. A large number of microorganisms including both yeasts and bacteria possess inducible pentose uptake systems that are subject to regulation in the presence of other sugars [2-5]. Most of the microorganisms preferentially utilize hexoses over pentoses. Since lignocellulosic hydrolysates comprise of both pentoses and hexoses such regulatory mechanisms pose a great problem in any process optimization for maximal utilization of available sugars in lignocellulosic hydrolysates. This chapter summarizes the general mechanisms of sugar transport along with a detailed account of kinetic studies and mode of energy coupling of pentose uptake in various bacterial and yeast strains. Regulation of pentose uptake in both bacteria and yeasts has been critically 99

lOO evaluated, specially in the presence of other carbohydrates. In addition, genetic analyses of pentose uptake has been overviewed in order to provide insights into the molecular mechanisms underlying pentose uptake and regulation in pentose fermenting microorganisms.

2

MODE OF SUGAR UPTAKE

Sugar transport occurs via three different mechanisms i.e. passive diffusion, facilitated diffusion and active transport. Passive diffusion is the simplest and slowest process and is governed by the law of mass action i.e. net diffusion occurs towards the lower concentration and at equilibrium the concentration of solute on each side of membrane is equal. Thus intracellular concentration of diffusible metabolite never exceeds that in surrounding media and the process is not saturable with respect to substrate concentration. Temperature and metabolic inhibitors do not affect the transport [5,6]. Only small lipid soluble molecules such as glycerol or ethanol are known to be transported to an appreciable extent by this mechanism. The rate of transport of most sugars by this mechanism is negligible and only acyclic polyols (erythritol, xylitol, ribitol, D-arabinitol, mannitol, sorbitol and galactitol) are known to be transported by passive diffusion in yeast Saccharomyces cerevisiae [7]. Facilitated diffusion is a variation of passive diffusion in which a membrane carrier participates in the diffusion process [5,6]. Facilitated diffusion, like passive diffusion does not concentrate metabolite intracellularly nor is sensitive to metabolic inhibitors i.e. the process is independent of input of energy. Attainment of influx-efflux equilibrium is very rapid, often being attained in seconds. Since facilitated diffusion involves a membrane carrier protein, the uptake is both temperature dependent and saturable with respect to substrate concentration. This property resembles that of a simple enzyme reaction and is frequently treated by Michaelis-Menten kinetics. Facilitated diffusion is stereospecific and binding of transported compound is a prerequisite of this type of transport process so it may be competitively inhibited by structural analogs of the substrate. The carrier may also be inactivated by appropriate mutations. Facilitated diffusion is used by microorganisms for the transport of sugars. Active transport is the most common mode of sugar transport in microbial cells. Like facilitated diffusion, active transport is carrier mediated, and therefore, the transport

lOl process is temperature dependent and saturable by substrate concentration and sensitive to competitive inhibition by substrate analogs [5,6]. Mutation may inactivate the transport activity. Unlike facilitated diffusion, active transport requires energy input and thus accumulates solutes against steep concentration gradients. Due to energy coupling, active transport systems are sensitive to inhibitors of energy metabolism. Influx-efflux equilibria in active transport are attained more slowly than for facilitated diffusion, often ranging from minutes to hours. Based on the mode of energization for transport of metabolites, active transport can be further classified into chemiosmotic, direct energization and group translocation [8]. Metabolic energy for active transport is provided by establishing a membrane potential in a chemiosmotic mechanism, while the hydrolysis of ATP provides energy in direct energization mechanism. In group translocation mechanism transfer of phosphate from phosphoenol pyruvate (PEP) to the sugar substrate provides the requisite energy for the transport of metabolites. Pentoses are commonly transported by generation of membrane potential due to H§ symport [2,9-11]. The operation of H§ symport depends on the ability of the carrier molecule to become reversibly protonated or deprotonated according to the proton gradient across the membrane. Thus it becomes alternatively electrically charged and the charged carrier molecules distribute themselves across the membrane according to the membrane potential. In addition the protonated carrier possesses a high substrate affinity. Thus H* symport is an electrogenic substrate transport process. Due to co-transport of positively charged protons, the uptake of uncharged molecule causes depolarization of membrane potential. Experimentally, a transient alkalinization of unbuffered suspension is observed with the onset of sugar transport. This is due to tight coupling of substrate with influx of protons. However, this phenomenon is transient as influx of proton can be compensated soon by the stimulated pumping of protons out of the cell. Interestingly, the modes of energization of transport systems are not mutually exclusive and more than one mechanism may operate in a single microorganism.

3

PENTOSE UPTAKE IN YEAST

Although the uptake of pentose has been studied in pentose fermenting yeasts such as Pichia stipitis [1,12,13], Pichia heedii [13] and Candida shehatae [14], most of our understanding related to mechanism of pentose transport in yeasts come from the

102 studies conducted on Rhodotorula gracilis (glutinis) [15-17]. Pentose uptake in yeasts is mostly via an active symport mechanism, however, xylose uptake in Saccharomyces cerevisiae and C. shehatae via low affinity system is mediated by facilitated diffusion. Kinetic studies have revealed the presence of more than one uptake system for xylose uptake in R. glutinis [17], P. stipitis [12-13], P. heedii[13] and a high affinity system in C. shehatae [14]. R. glutinis has been employed to understand the energetics of pentose transport due to its strict dependence on energy metabolism. R. glutinis accumulates D-xylose, L-rhamnose and D-arabinose but not D-ribose by a mechanism with which D-galactose interacts [15-17]. L-Rhamnose is not metabolized by R. glutinis while D-ribose is metabolized after a period of adaptation [16,18]. Due to strict dependence of transport on energy metabolism; inhibitors of energy metabolism, proton conductors or lack of oxygen ceases net carbohydrate uptake although slow exchange of carbohydrate across the membrane has been monitored [19-20]. The transient alkalinization of R. glutinis suspensions on addition of sugar occurs due to the functioning of a proton sugar symport [21].

R. glutinis, while respiring extrudes protons and the rate is accelerated in the presence of K§ and is inhibited in cells treated with dicyclohexylcarbodiimide [21]. Accumulation of lipophilic cations tetraphenylphosphonium and triphenylphosphonium by R. glutinis has been studied under various conditions. Proton conductors, anaerobic conditions or presence of K§ lower the amount of lipophilic cation accumulation in the steady state and membrane potential is known to be dependent on' extracellular pH. This indicated the involvement of an electrogenic pump in this yeast. Onset of xylose uptake in this yeast is accompanied by about one equivalent of protons in the pH range of 3-5 [10]. Under anaerobic conditions both proton and sugar uptake is abolished. In addition, D-xylose uptake diminished the rate of uptake of lipophilic phosphonium cations. These observations gave evidence for the occurrence of an electrogenic symport in R. glutinis. Kinetic studies of xylose uptake in R. glutinis revealed the presence of two uptake systems, one responsible for high affinty uptake with an apparent Kr, of 1-2 mM, similar to that required to half saturate the proton symport, while the other low affinty system with an apparent Kr, of 18 mM near pH 5 [17] and 15 mM at pH 6.5 [10]. The presence of two affinity systems has been suggested as the high affinity system is competitively inhibited by galactose [17] and only a weak interaction of galactose with low affinity xylose carrier occurred [16]. In addition, the high affinity system is selectively derepressed in starved yeast cells. However, Hofer and Misra [10] opined a pH dependence of Km for xylose uptake in R. glutinis and suggested that the low affinity carrier corresponds to a proton symport absorbing xylose without a proton, as Km for

lO3 xylose uptake varied from 2 mM at pH 4.5 to 80 mM at pH 8.5. This change in effective carrier affinity is reversible. They also observed an apparent dissociation constant of monosaccharide carrier at pKa 6.75. At pH 8.5, when pH gradient across the cell membrane vanished no sugar accumulation was observed. Alcorn and Griffin [17] however suggested that two distinct xylose carriers are involved and their activities are additive near pH 5 where symport mechanism is expected to be fully activated. Pentose fermenting yeast Pichia has attracted considerable interest for the transport of xylose due to its efficient xylose fermenting capability. Nuclear magnetic resonance spectroscopy determined that under aerobic conditions xylose transport in P. stiptis is apparently the limiting step in xylose metabolism while under anaerobic conditions a later step in metabolism may be limiting [1]. The uptake studies of xylose revealed two transport systems i.e., low affinity system with an apparent Kr, of 2-3 mM and a high affinity system with a Krn of 0.06-0.08 mM [12]. Recently Does and Bisson [13] have studied xylose uptake in two different strains of Pichia. In addition to P. stipitis they have studied xylose uptake in P. heedii that is genetically amenable and is strict respirator. In both strains kinetic studies demonstrated at least two transport systems differing in their affinity for xylose. This was apparent on the basis of their growth in presence of xylose in the growth medium. When grown aerobically with high (2%) or low (0.05%) xylose, it was observed that while P. heedii could grow on both the concentrations P. stipitis grew only on high xylose concentration. In P. stipitis the low affinity system has an apparent Krn of about 380 mM and high affinity system has a Km of 0.9 mM when grown on high xylose concentration. Under the similar conditions the velocity of xylose uptake of 20 mM xylose solution was 20 nmol/min. P. heedii when grown on high xylose showed Km of low affinity of xylose uptake, which was 40-50 mM, however these cells grown on low xylose showed high affinity system ( i.e. 0.1 mM). Table 1 summarizes the kinetic constants for D-xylose uptake in yeasts. The uptake of glucose and xylose by P. stiptis grown on glucose and xylose, respectively, showed that an uncoupler such as 2,4-dinitrophenol (DNP) resulted in the inhibition of both xylose and glucose uptake. DNP significantly decreased the uptake of xylose in P. heedii but had no effect on glucose uptake. This differential effect of DNP on putative high and low affinity system is not clear. However, on the basis of a difference in the energy requirement of sugar uptake, Does and Bisson [13] suggested the involvement of different mechanisms for the transport of D-xylose and D-glucose in Pichia.

lO4 Table 1 Kinetic constants for D-xylose uptake in yeasts

Microorganism

Rhodotorula glutinis Rhodotorula glutinis

Mode of uptake

Transport system(s) (affinity)

Km

Active

High Low High

0.6 18.0 1.7a 2.0 b

Active

Reference

(mM)

[17] [17]

[lo] [lO] [lo]

C a

Low

[10]

15.0b Pichia

stipitis Pichia stipitis Pichia heedii Candida shehatae

Metschnikowia reukaufii Saccharomyces cerevisiae

Active, Proton symport Active Active Proton symport, Inducible, Facilitated diffusion Active, Proton symport Facilitated diffusion

apH 4.5, bpH 6.5; CpH 8.5

High Low

[lo] [lo]

83.0 ~ 0.08 3.00

[12] [12]

High Low High Low High

0.9 380.0 0.1 45.0 1.0

[13] [13] [13] [13] [14]

Low

125.0

[14]

2.0

[19]

130.0

[22]

105

Candida shehatae, known to ferment D-xylose, has also been studied for its

ability to transport sugars including pentoses [14]. Both facilitated diffusion and proton symport mechanism exist for the transport of D-xylose in this yeast. The conditions of glucose or D-xylose repression produce a facilitated diffusion in C. shehatae that accepts glucose, D-mannose and D-xylose but neither D-galactose nor L-arabinose. The apparent Km for D-xylose uptake is 125 mM and Vmax22.5 mmol/g/h. The uptake of xylose in a facultative anaerobic yeast Metschnikowia reukaufii has also suggested active, proton symport mechanism in yeasts [9]. This yeast has a constitutive mobile membrane carrier for the uptake of D-xylose (Kin, 2.0 mM), D-glucose and 3-O-methyl D-glucose. The uptake of these monosaccharides is sensitive to uncouplers like carbonylcyanide, n-chlorophenylhydrazone or dinitrophenol and no accumulation occurred in the presence of uncouplers suggesting that uptake is via an active system. In M. reukaufii the onset of sugar transport such as D-glucose, 2-deoxy-D-glucose, 3-O-methyl-D-glucose, D-xylose, D-galactose and D-fructose is known to be actively transported and resulted in a short alkalinization of unbuffered cell suspensions or a depolarization of the membrane potential in buffered cell suspensions (pH 9.0). In contrast, D-arabinose, which is not transported actively, failed to induce H§ co-transport as well as depolarization of membrane potential. The H+/ sugar stoichiometry is one H§ per sugar molecule taken up. Thus H§ symport energized by electrochemical gradient of H§ the plasma membrane is responsible for sugar transport in M. reukaufii. Studies on the uptake of xylose by S. cerevisiae and Candida utilis revealed that the uptake of xylose is less efficient (26%) than glucose transport [23]. The rate of xylose transport in S. cerevisiae is similar to that observed in C. utilis grown on glucose. The rate of sugar uptake also depends on oxygen, as uptake of both xylose and glucose by S. cerevisiae under aerobic condition is 7-10 times higher than the rates observed under anaerobic conditions [23]. The rate of xylose transport by S. cerevisiae has been reported to be similar [24] or higher [25] compared to the rate of glucose transport. The affinity of xylose transport (Kin) in S. cerevisiae is 4-fold to 25-fold [24,25] lower than glucose.

4

PENTOSE UPTAKE IN BACTERIA

The understanding of pentose transport in bacteria has been limited to a very few bacterial species and only transport of D-xylose, L-arabinose and D-ribose is known

lO6 in some detail. The existence of a transport system for xylose uptake in E. coil has been reported by David and Wiesmeyer [26]. They observed an inducible xylose permease that transported xylose against a 100-fold concentration gradient. This transport system is energy dependent and specific for xylose because D-ribose, D-arabinose and xylitol are not transported via this permease. Lam and his co-workers [2] observed that the addition of xylose to energy depleted cells of E. coil elicited an alkaline pH change which failed to appear in the presence of uncoupling agents. In addition accumulation of [14-C]xylose by energy replete cells is inhibited by uncoupling agents but not by fluoride or arsenate. Subcellular vesicles of E. coli accumulate [14-C] xylose provided ascorbate plus phenazine methosulfate are present as respiratory substrates and this accumulation is inhibited by uncoupling agents or valinomycin. These experimental findings suggest that the mechanism of energization for xylose uptake in E. coil is by a proton motive force rather than by a phosphotransferase or directly energized mechanism [2]. These workers also observed specificity of the xylose-proton symport system as L-arabinose, D-fucose D-ribose, D-lyxose and xylitol failed to promote pH changes in xylose induced E. coil strains VL17 and K10. Xylose uptake in E. coil exhibited biphasic kinetics consistent with the presence of two systems with Kr, values of 24 ~M and 110 pM [3]. Salmonella typhimurium LT2, like E. coil K12, utilizes L-arabinose, D-ribose and D-xylose as sole carbon source and hence was employed for the study of the mechanism of pentose uptake [3,27]. S. typhimurium also showed inducible uptake of xylose with 40 fold accumulation against the concentration gradient [3]. The rate of D-xylose transport in S. typhimurium was nearly half that of E. coil K-12. Contrary to E. coil K12 that exhibited two transport systems, S. typhimurium has only one transport system with an apparent Kr, of 0.41 mM. Bacteroides xylanolyticus X5-1, a strict anaerobic bacterium, that can grow on a variety of sugars including xylan has also been studied for its ability to transport Dxylose in the presence of various metabolic inhibitors [28]. Based on the specificity of D-xylose uptake and its inhibition by 2,4-dinitrophenol, mercuric chloride, arsenate and N,N'-dicyclohexylcarbodiimide it has been suggested that xylose uptake in B. xylanolyticus is an active process [28]. Prevotella (Bacteroides) ruminicola, a xylanolytic bacteria also exhibit uptake of both xylose and glucose when grown on xylose. In contrast, Fibrobacter succinogenes that is known to posses xylanolytic activity lacked the ability to utilize xylose due to the lack of xylose permease [29]. Clostridium acetobutylicum is known to ferment pentoses for solvent production. This bacteria can convert xylose to solvents with a yield of 28% which is close to the maximal value of 32% obtained with glucose. The saturation kinetics of xylose showed

107 an apparent Km of 5 mM for xylose uptake [30]. Solvent production led to a concomitant decrease in the transport activities of both glucose and xylose. This effect of end product on inhibition of transport activities has been discussed in chapter 12. Selenomonas ruminantium, a common gram negative anaerobe which is prevalent in the rumen [31], exhibits capability to ferment various carbohydrates including pentoses. It has high affinity for glucose, maltose, sucrose and xylose [32], but glucose, sucrose and xylose are preferentially utilized over maltose [33]. S. ruminantium HD strain uses phosphoenolpyruvate dependent phosphotransferase (PEP-PTS) system for glucose and sucrose uptake and maltose is utilized after hydrolysis to glucose by extracellular maltase. However, xylose is not transported via PEP-PTS system, instead a high energy phosphate compound has been implicated to be involved in xylose uptake [34]. Non-linear kinetics of xylose uptake suggested that more than one uptake systems with different affinities for xylose may be involved in its uptake in S. ruminantium [35]. The mechanism of energy coupling for pentose uptake in S. ruminantium strain HD is suggested to be electrogenic pentose proton symporters as it was possible to demonstrate pentose transport in deenergized cells by the imposition of an artificial electrical potential or chemical gradient of protons [36]. Kinetic constants for pentose uptake in a few bacteria have been listed in Table 2. L-Arabinose transport in E. coil has been studied in relatively greater detail. Early studies on L-arabinose transport in E. coil B/r have demonstrated the presence of an inducible, energy dependent uptake for this pentose sugar [37]. When the uptake via this system is abolished by mutations at a locus designated araE, it was found that E. coil B/r possesses an L-arabinose binding protein that binds L-arabinose with a Kr, of 5x10 .6 M [38]. Brown and Hogg [39] later reported that E. coil possesses two active transport systems for L-arabinose uptake. The system for high affinity has a Km of 8.3 x 10-6M, while the system for low affinity has a Kr, of 1.0 x 10-4M. These two systems showed distinct responses to analogs that act as competitive inhibitors of initial uptake. For example, D-galactose strongly inhibits L-arabinose uptake by the high affinity system but only weakly inhibits L-arabinose uptake by the low affinity system. D-Fucose, D-xylose and ~-methyl L-arabinoside competitively inhibit the uptake of L-arabinose by both the systems to approximately the same extent. Kinetic studies have shown that the Km for L- arabinose uptake by the high affinity system resembles the Kr, for binding of L-arabinose by the binding protein and both have similar K~ values for inhibitory substances. Thus on the basis of transport and inhibition kinetics and the properties of mutants lacking one or the other type of the system it has been suggested that the high affinity system involves the L-arabinose binding protein. Later the gene product of araF was characterized as L-arabinose binding protein that serves as a

lO8 component of high affinity L-arabinose transport system [40].

Table 2 Kinetic constants for pentose uptake in bacteria Substrate

D-Xylose

Microorganism

E. coil

K-12

D-Xylose D-Xylose D-Xylose

Salmonella typhimurium Clostridium acetobutylicum Selenomonas ruminantium

L-Arabinose E. coil B/r L-Arabinose E.cofi D-Ribose

E.cofi

ML 308-225

NT, Not tested.

Mode of Uptake

Transport Systems

Km

Active, proton symport, inducible Active, inducible Inducible

High Low

24 110

[3]

One

41

[3]

One

5000

Two

NT

[35,36]

One

125

[37]

High Low NT

8.3 100 NT

[39] [39] [43]

Active, inducible, proton symport Active, inducible Inducible Active, constitutive

Reference

(IJM)

[30]

lO9 Studies on D-ribose transport in E. coil started with findings that this organism is unable to ferment D-ribose unless grown in the presence of ribose even though enzymes for the fermentation of ribose are produced constitutively. This provided evidence for an inducible D-ribose transport system in E. coli [41]. But other studies suggested that both constitutive and inducible transport system for ribose transport exist in E. coil X289 [42]. Both permeases concentrate D-ribose against a gradient and transport is inhibited by sodium azide implicating involvement of active transport systems. The activity of ribose transport is severely reduced in cells subjected to osmotic shocks. In addition, this transport system is found to be absent in membrane vesicles [43]. Thus it is apparent that a binding protein is involved in D-ribose uptake in E. coil In addition isolation of ribose binding proteins from osmotic shock fluids of Salmonella typhimurium [44,45] and E. coli [45] has also been reported. Mechanism of energy coupling for transport of D-ribose was also studied in E. coil ML308-225 and its mutant DL-54 which is defective in Ca2§ Mg2§

[43]. It was observed that

substrates provided energy for the transport of ribose only when they were able to generate ATP. Further, substrates that generated ATP primarily through oxidative phosphorylation act as poor energy sources in mutant strains. In addition anaerobic conditions or uncouplers of oxidative phosphorylation proved to be ineffective on ribose transport. Thus phosphate bond energy of ATP rather than an energized membrane state has been suggested to couple the energy to ribose transport in E. coil [43].

5

REGULATION OF PENTOSE UPTAKE

5.1

Yeasts

Pentose fermenting yeasts such as Pachysolen tannophilus, Pichia stipitis and

Candida shehatae are known to ferment D-glucose, D-xylose, D-mannose and Dgalactose to ethanol. However, when these sugars are present in combination a sequential pattern of their utilization occurs, whereby the hexoses are fermented preferentially over pentoses [46,47]. This sequential sugar utilization is the major problem that limits the biotechnological use of pentose fermenting yeasts. Thus efforts have been made to understand the regulation of D-xylose uptake in the presence of

11o other sugars in pentose fermenting yeasts. Pentose fermenting yeasts are known to possess multiple uptake systems for D-xylose uptake and the type of transport system operating varies among yeasts and also depends on their nutritional status. For example, derepression of C. shehatae by starvation formed at least three sugar proton symports- one is responsible for accumulation of 3-O-methyl glucose, glucose and D-mannose while the second symport system transported D-xylose (Kr,, 1.0 mM; Vr,ax 1.4 mmol/g/h) and galactose but neither glucose, D-mannose nor L-arabinose. A third symport system is apparently used for L- arabinose transport. The stoichiometry of symport is known to be one proton for each molecule of sugar transported. Substrate of one sugar symport non-competitively inhibited the transport of substrate of the other symports. It was interesting to note that while facilitated diffusion is absent or not measureable in starved cells of C. shehatae, with glucose as substrate, it coexisted with proton symport activity when D-xylose is used as a substrate. In Pichia heedii xylose uptake occurs via low affinity system that is induced by growth on a high substrate concentration with somewhat decreased levels of high affinity uptake. Conversely, at low xylose concentration high affinity uptake is induced while low affinity uptake decreases [13]. When grown on high concentration of xylose, the xylose transport activity via low affinity system in P. heedii was 45 nmol/min/mg dry weight, and when grown on low xylose concentration the high affinity system showed an uptake of 13 nmol/min/mg dry weight. In addition while 100 mM xylose is unable to inhibit glucose uptake, xylose uptake is inhibited by the same concentration of glucose in P. heedii. This ability of glucose to inhibit xylose uptake reflects a regulatory mechanism ensuring the use of glucose prior to metabolism of xylose.

5.2

Bacteria

Like yeasts, bacteria also possess inducible pentose uptake systems. However bacteria may differ in terms of their substrate specificity. For example in E. coli, xylose is the only sugar that induces xylose uptake while cells grown on D-ribose or D-glucose are unable to induce xylose uptake suggesting that xylose transport exhibits substrate and inducer specificity [2]. D-Xylose induced transport system for the uptake of xylose is known in S. typhimurium, but significant levels of L-arabinose are also transported through D-xylose induced cells. In addition, D-xylose accumulated in L-arabinose induced cells probably through L-arabinose transport system [3]. However the fact that

111 whether these pentoses are non-specific for induction of transporter or they are nonspecific for their transport itself is not clear. Table 3 shows inducible pentose transport activities in E. coil and S. typhimurium.

Table 3 Inducible pentose transport in E. coil and S. typhimurium. Strain

Cells grown on glycerol containing

Sugar conc.

D-Xylose L-Arabinose Reference Transport Transport (nmol/mg/min)

E.cofi

D-Xylose L-Arabinose D-Ribose D-glucose

10 10 10 10

4.4 0.3 0.2 0.1

0.3 3.2 0.3 0.2

[2] [2] [2] [2]

None

-

0.01

0.8

[2]

D-Xylose D-glucose

10 mM 10 mM

6.9 0.9

0.2 0.8

[2] [2]

D-Xylose L-Arabinose None

0.2% 0.2% -

38.0 35.3 0.8

3.5 173.5 0.6

[3] [3] [3]

VL17

E.coli K10

S. typhimurium

mM mM mM mM

Although growth rate and sugar consumption of bacteria is higher in glucose grown cells, C. acetobutylicum exhibits diauxic growth in the presence of mixtures of glucose and xylose. Glucose uptake activity is observed in both glucose and xylose grown cells but growth on xylose was associated with the induction of a xylose permease activity which is repressible by glucose in xylose induced cells [30]. This is apparent from the observations that only cell suspensions of xylose grown C.

acetobutylicum incorporated xylose, whereas glucose is incorporated by cells grown on either substrate. When mixture of both xylose and glucose is inoculated with cells

112 grown on xylose, formation of the xylose uptake system is repressed and the activity is subsequently diluted until glucose is nearly exhausted from the media [30]. The fermentation kinetics of C. acetobutylicum grown on glucose, xylose and mixture of both in batch and fed-batch cultures also suggested that xylose utilization is inducible and is inhibited at glucose concentrations above 15 g/I [48,49]. In fed-batch cultures at low feeding rates with glucose concentrations below 15 g/I, glucose and xylose appear to be taken up at the same rates during the first part of the fermentation. An accumulation of xylose, when fermentation is inhibited, suggests the repression of xylose utilization, when catabolic flux of glucose alone could satisfy the metabolic activity of cells [49].

Pediococcus halophilus (soy pediococci) grow on soy maromi mash that contain a mixture of glucose, galactose, arabinose and xylose and utilizes glucose preferentially over pentoses. The accumulated pentoses during fermentation react with amino acids (from soybeans) by Maillard reaction and result in browning pigments. Thus selective utilization of pentoses from soy sauce moromi mash in fermentation process is of interest to stop the browning of pigments. Abe and Uchida [50] isolated mutant of P. halophilus-X160 that can preferentially utilize pentoses such as xylose and arabinose even in the presence of large amounts of glucose. The lack of catabolic control by glucose in these mutants has been attributed to a defect in Phosphoenolpyruvate (PEP): mannose phosphotransferase system that functions as main glucose transport system in this organism. Recent studies of Strobel [36] demonstrated that Selenomonas ruminantium strain D utilizes hexoses preferentially over pentoses when grown on a combination of glucose and xylose or arabinose. However, cellobiose and pentoses are utilized simultaneously. Continuous culture studies have shown that at low dilution rate (0.10 h-1) the organism co-utilizes glucose and xylose. This co-utilizatiton has been associated with growth rate dependent decrease in glucose phosphotransferase activity and the apparent inhibition of pentose utilization has been attributed to catabolite inhibition by the glucose phosphotransferase transport system. It appears that xylose and arabinose permease syntheses are controlled by different regulatory mechanisms in this bacteria. Xylose transport is inducible and repressed (probably via PTS mediated inducer exclusion) by growth on glucose, maltose and sucrose in S. ruminantium [36]. On the other hand arabinose transport has been found in cellobiose grown cultures of S.

ruminantium strain D. Thus arabinose permease appears to be non-inducible, and repressible in the presence of glucose [36]. Xylose uptake in glucose grown cells of Bacteroides xylanolyticus X5-1 has been reported to be very low but the uptake rate increases when incubated with a mixture

113 of xylose and glucose. The increase in the uptake rate remains unaffected by chloromphenicol indicating that protein synthesis is not required for the activation of uptake system rather a constitutive uptake system is activated [51].

6

GENETIC STUDIES ON PENTOSE UPTAKE

Genetic studies on pentose uptake have considerably enriched our knowledge of underlying molecular mechanisms of microbial pentose transport and its regulation. One of the most common approaches to genetically characterize transport system involves isolation of transport defective mutants and cloning of the gene responsible for the restoration of transport activity of mutants. Among pentoses, L-arabinose uptake system has been characterized in greater details while D-xylose transport system has also been characterized to some extent. However, such studies have been only restricted to very few bacterial species, such as E. coli and S. typhimurium. As has been dicussed earlier, L-arabinose is transported by two uptake systems in E. coil; one low affinity system and the other high affinity system. Low affinity system has been characterized by studies of mutants defective in uptake of L-arabinose through this system. Mutation causing defect in low affinity uptake is mapped to a single locus, designated araE at 60 min on E. coil chromosome [38,52]. The product of this gene has been characterized as 52 kd membrane associated protein [53,54]. The second high affinity system for arabinose uptake has been shown to be dependent on L-arabinose binding protein in E. coli [39,40]. The gene encoding L-arabinose binding protein is designated as araF and suggested to be a part of an operon located at 45 min on the E. coil chromosome [39,40]. This operon consisted of araF, araG and araH [52]. L-Arabinose binding protein is encoded by araF and localized in the periplasm, whereas araG and araH encode membrane associated proteins of 52 kd and 31 kd, respectively [55]. Expression of plasmids containing various portions of araFGH operon sequences were characterized for their ability to facilitate the high affinity L-arabinose transport in L-arabinose transport deficient mutant lacking the chromosomal copy of this operon. The capacity of these gene products to revive the high affinity transport phenotype demonstrated that the specific induction of all three operon coding sequences is essential to restore high affinity L-arabinose transport in E. coil [56]. D-xylose transport system is well characterized in Salmonella typhimurium [3] and E. coil [57]. The inducibility of D-xylose transport, D-xylose isomerase and D-xylose

114 kinase is coded by a cluster of genes xylT, xylA and xylB respectively in S. typhimurium [3]. A number of mutants of S. typhimurium including those of xylose transport defective mutants have been selected after ethylmethanesulfonate mutagenesis. These mutations mapped at 78 units on linkage map with order of gene xylT-xylR-xylB-glyS-mtlB. Shamanna and Sanderson [3] postulated gene xylR to be the regulatory gene controlling the activity of xylT, xylB and xylA through an activator A1, which converts to state A2 after interaction with inducer D-xylose. Similarly xyl- mutants of E. coil have been exploited to genetically characterize xylose transport and metabolism. Kurose and his co-workers have isolated E. coil mutants (M3C and MX-5) deficient in xylose uptake [57]. The mutant MX-5 posseses low xylose isomerase activity as compared to wild type. The gene responsible for xylose uptake in E. coil was cloned onto vector plasmid pBR 322 and the resistant hybrid plasmid was designated pXp5. This hybrid plasmid of 11.9 megadalton (Md) molecular size was a dimer, consisting of two identical DNA each containing 3.3 Md chromosomal fragment of E. coil C600 inserted into the Pst I site of pBR 322. From the foregoing discussions it is evident that the early studies of pentose uptake addressed the queries regarding the mechanism of sugar transport in bacteria and yeasts. Most of the studies employed E. coil and Rhodotorula glutinis as model organisms. Later the presence of large amounts of pentose sugars in lignocellulosic waste attracted considerable interest in microbial utilization of pentose sugars along with other hexoses for the production of liquid fuel from renewable biomass. Many studies have revealed that at the end of ferementation of lignocellulosic wastes a large amount of pentose sugars remain unutilized due to preferential utilization of hexoses followed by product inhibition. Thus the uptake of pentose in the presence of other hexoses emerged as a challenging issue for biotechnological exploitation of yeasts as well as bacteria. So far little is known about the regulation of pentose uptake in the presence of other sugars specially hexoses. Efforts have been made to isolate and clone the genes responsible for the uptake of pentoses in bacteria. Our understanding of regulatory mechanisms of pentose uptake in the presence of other sugars at the molecular level is likely to help in improvement of strains for the preferential utilization of pentoses from complex mixture of sugars.

115 7

REFERENCES

Ligthelm ME, Prior BA, du Preez JC, Brandt V. Appl Microbiol Biotechnol 1988; 28: 293. Lam VMS, Daruwalla KR, Henderson, PJF, Jones MMC. J Bacteriol 1980; 143: 396. Shamanna DK, Sanderson KE. J Bacteriol 1979; 139: 64. Webb SR, Lee H. Biotechnol Adv 1990; 8: 685. Roseman S. In: Hokin LE ed., Metabolic Transport, New York: Academic Press, 1972; 41. Cooper TG. In: Strathern JN, Jones EW, Broach JR, eds. The Molecular Biology of Yeast Saccharmyces;vol 2 Metabolism and Gene Expression, New York: Cold Spring Harbor Laboratory, 1982; 399. 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22

Canh DS, Horak J, Kotyk A, Rihova L. Folia Microbiol 1975; 20: 320. Fiechter A, Fuhrmann GF, Kappelli O. Adv Microb Physiol 1981; 22: 123. Aldermann B, Hofer M. Exp Mycol 1981;5: 120. Hofer M, Misra PC. Biochem J 1978; 172: 15. Eddy AA. Adv Microb Physiol 1982; 23: 1. Killian SG, van Uden N. Appl Microbiol Biotechnol 1988; 27: 545. Does AL, Bisson LF. Appl Environ Microbiol 1989; 55: 159. Lucas C, van Uden N. Appl Microbiol Biotechnol 1986; 23: 491. Kotyk A, Hofer M. Biochim Biophys Acta 1965; 102: 410. Janda S, Kotyk A, Tauchova R. Arch Microbiol 1976; 111: 151. Alcorn ME, Griffin CC. Biochim Biophys Acta 1978; 510: 361. Hofer M. J Membrane Biol 1970; 3: 73. Hofer M. Arch fur Mikrobiol 1971;80: 50. Hofer M. J Theor Biol 1971; 33: 599. Misra PC, Hofer M. FEBS Lett 1975; 52: 95. Kotyk A, Janacek. In : Cell Membrane Transport, New York and London: Plenum, 1975; 343.

23

Batt CA, Carvallo S, Easson DD, Akedo M, Sinskey AJ. Biotechnol Bioeng 1986; 28: 549.

24

Kotyk A. Folia Microbiol 1967; 12: 121.

25

Cirillo VP. J Bacteriol 1968; 95: 603.

26

David JD, Wiesmeyer H. Biochim Biophys Acta 1970; 201: 497.

116 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55

Shamanna DK, Sanderson KE. J Bacteriol 1979; 139: 71. Biesterveld S, Kok MD, Dijkema C, Zehnder AJB, Stams AJM. Arch Microbiol 1994; 161: 521. Matte A, Forsberg CW, Gibbins AMV. Can J Microbiol 1992; 38: 370. Ounine K, Petitdemange H, Raval G, Gay R. Appl Environ Microbiol 1985; 49: 874. Caldwell DR, Bryant MP. Appl Microbiol 1966; 14: 794. Russell JB, Baldwin RL. Appl Environ Microbiol 1979; 37: 531. Russell JB, Baldwin RL. Appl Environ Microbiol 1978; 36: 319. Martin SA, Russell JB. J Gen Microbiol 1988; 134: 819. Williams DK, Martin SA. Appl Environ Microbiol 1990; 56: 1683. Strobel HJ. Appl Environ Microbiol 1993; 59: 40. Novotny CP, Englesberg E. Biochim Biophys Acta 1966; 117: 217. Hogg RW, Englesberg E. J Bacteriol 1969; 100: 423. Brown CE, Hogg RW. J Bacteriol 1972; 111:606. Clark AR, Hogg RW. J Bacteriol 1981; 147: 920. Eggleston LV, Krebs HA. Biochem J 1959; 73: 264. David J, Wiesmeyer H. Biochim Biophys Acta 1970; 208: 45. Curtis S J. J Bacteriol 1974; 120: 295. Aksamit R, Koshland DE Jr. Biochem Biophys Res Commun 1972; 48: 1348. Willis RC, Morris RG, Cirakoglu C, Schellenberg GD, Gerber NH, Furlong CE. Arch Biochem Biophys 1974; 161:64. Detroy RW, Cunningham RL, Herman AI. Biotechnol Bioeng Symp 1982; 12: 81. du Preez JC, Bosch M, Prior BA. Appl Microbiol Biotechnol 1986; 23: 228. Fond O, Matta-EI-Amouri G, Engasser JM, Petitdemange H. Biotechnol Bioeng 1986; 28: 160. Fond O, Matta-EI-Amouri G, Petitdemange H, Engasser JM. Biotechnol Bioeng 1986; 28: 167. Abe K, Uchida K. J Bacteriol 1989; 171:1793. Biesterveld S, Oude Elferink SJWH, Zehnder AJB, Stams AJM. Appl Environ Microbiol 1994; 60: 576. Kolodrubetz D, Schleif R. J Bacteriol 1981; 148: 472. MacPherson AJS, Jones-Mortimer MC, Henderson PJF. Biochem J 1981; 196: 269. Maiden MCJ, Davis EO, Baldwin SA, Moore DCM, Henderson PJF. Nature 1987; 325:641. Horazdovsky B, Hogg RW. J Mol Biol 1987; 197: 27.

117 56

57

Horazdovsky B, Hogg RW. J Bacteriol 1989; 171"3053. Kurose N, Murata K, Kimura A. Agric Biol Chem 1987; 51" 2575.

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Microbial Metabolism of Pentoses

1

INTRODUCTION

Microorganisms are known to metabolize a wide variety of naturally occurring carbohydrates as sources of carbon and energy, consequently, they have evolved various catabolic pathways for degradation of carbohydrates. It is evident from the preceding chapters that hemicellulose consists of both hexosans and pentosans and upon hydrolysis yield D-xylose, L-arabinose and D-glucose as major components. Due to availability of these pentoses in abundance along with hexoses in renewable biomass, microbial metabolism of D-xylose and L-arabinose have attracted considerable interest. While isolation of mutants defective in metabolic pathways has helped in elucidation of these pathways, understanding of metabolism has geared up engineering of pathways for improving bioconversion and yield. For example, decreasing the formation of by-products and channelization of metabolites and energy has helped tremendously in increasing the product yield. Microorganisms differ in the metabolic pathways they employ to utilize pentoses and the end product they yield [1,2]. With the advancement in genetics and molecular biological techniques and increasing knowledge of pentose metabolism, engineering of pentose metabolic pathway has become a powerful tool for manipulating genetic traits of microorganisms. This chapter deals with our current understanding of pentose metabolic pathways in microorganisms and their regulation. Major emphasis has been given to the metabolism of naturally occurring and more abundant pentose sugar, Dxylose. Metabolism of L-arabinose, generally present in lignocellulosics has also been included. However, metabolism of other unnatural pentoses such as D-arabinose, Lxylose, L-and D-lyxose, xylitol and L-arabitol are beyond the scope of this chapter and readers are referred to [3,4].

2

METABOLISM OF D-XYLOSE

The initial metabolic pathway of D-xylose and D-xylulose in all microorganisms involves their conversion to D-xylulose-5-phosphate which is then channelled into the 119

120 pentose phosphate pathway [5-7].

2.1

Conversion of D-xylose to D-xylulose-5-phosphate

The mechanism by which coversion of D-xylose to D-xylulose-5-phosphate is achieved differs in bacteria than yeast and mycelial fungi. Bacteria generally employ the enzyme xylose isomerase, whereas yeasts and mycelial fungi employ a two step oxidation-reduction pathway [2]. The latter pathway utilizes two sets of pyridine nucleotide linked-dehydrogenases. D-Xylose is first reduced to xylitol and is then reoxidized to D-xylulose.

2.1.1 Oxidative reductive pathway

Although occasionally the presence of xylose isomerase has been reported in some yeasts and fungi, metabolism of D-xylose in these organisms mostly proceeds via a two step oxidative reductive pathway (Figure 1). D-Xylose is first reduced to xylitol by enzyme xylose reductase (alditol: NADP/NAD-l-oxidoreductase, EC 1.1.1.21 ). Xylitol is then oxidized to D-xylulose by the enzyme xylitol dehydrogenase (xylitol: NAD-2-oxidoreductase, EC 1.1.1.9). Evidence for the role of these enzymes has been provided by genetic studies. For example, a mutant of Pachysolen tannophilus deficient in NADPH-dependent xylose reductase exhibits significantly reduced growth rates on D-xylose or L-arabinose compared to its wild type [8]. Similarly, mutants of Pichia stipitis lacking either NAD(P)H-dependent xylose reductase or NAD-dependent xylitol dehydrogenase are unable to grow on D-xylose

[9]. The activity of enzyme xylose reductase has been detected in Candida albicans [10-12], Candida utilis [13-14], Geotrichum candidum [15], Pichia stipitis [16-18], Pichia quercuum [19], Pachysolen tannophilus [20-23], Cephalosporium chrysogenum [24], Melampsora lini [25], Penicillium chrysogenum [26] and Fusarium oxysporum [27,28]. The enzyme xylose reductase mostly requires reduced cofactor NADPH to carry out the electron transfer but this cofactor requirement may vary in different genera of

121 D-XYLOSE NADP XYLITOL NAD

~

D-XYLULOSE ,,~ATP

13 p

ACETYL-P <

(5)

,,, D-XYLULOS E-5-PNOSPHATE

i

ACETATE

~Pentosephosphatepathway .(4) GLYCERALDEHYDE-3-P I Glycolyticpathway PYRUVATE ' ) CO2 + H20 ~/ Citricacidcycle (6) ACETALDEHYDE

[(7) ETHANOL

Figure 1. D-Xylose metabolism by yeasts. Enzymes are indicated by numbers as follows: (1) xylose reductase; (2) xylitol dehydrogenase; (3) xylulokinase; (4) transaldolase and transketolase; (5)phosphoketolase; (6)pyruvate decarboxylase; (7) alcohol dehydrogenase.

122 yeast as well as in the same genera under different physiological conditions. The enzyme purified from Candida albicans exhibited wide specificity for a number of substrates [11 ]. Based on the relative activity of the enzyme in the presence of various substrates, it was concluded that aldose in D-glycero-configuration with hydroxyl group attached to carbon-2 acts as a good substrate of xylose reductase, while those lacking the hydroxyl group at carbon-2 are poor substrates [11]. The enzyme also catalyzed reverse reactions using polyols as substrate and NADP as the cofactor. Verduyn and coworkers [16] purified NADH-linked xylose reductase from xylose fermenting yeast Pichia stipitis. The enzyme exhibited a dual coenzyme specificity for NADH and NADPH. It has been suggested that in most yeasts which do not or only slowly ferment D-xylose anaerobically, xylose reductase activity is NADPH-linked [29,30] and they either lack or exhibit low NADH-linked activity [31]. In contrast to single xylose reductase enzyme in Pichia stipitis, multiple forms of xylose reductase have been suggested in Pachysolen tannophilus. Studies indicate the presence of three different forms of the enzyme. Under aerobic growth conditions separate NADPH specific and NADH specific forms of xylose reductase have been characterized [20,21]. However, under oxygen limited growth, NADPH-specific and dual cofactor (NADPH/NADH) forms have been identified [22]. Variation in the relative amounts of the enzymes has been observed as a function of growth conditions. As growth conditions change from aerobic to anaerobic the ratio of NADPH/NADH-iinked xylose reductase activity changes from 4:1 to 1:1 in P. tannophilus [32]. The level of total xylose reductase activity (either NADPH or NADH-linked) has been reported to be the highest in oxygen limited cells followed by anaerobic cultures. Highly aerated cultures exhibited much lower levels of total xylose reductase activity and the activity was primarily NADPH-linked. The various forms of xylose reductase from Pachysolen tannophilus have been isolated and purified. Verduyn et al. [22] isolated two forms of xylose reductase with molecular weights of 41 kDa and 37 kDa. The 41 kDa molecular weight form had dual NADPH/NADH-linked reductase activity, whereas the 37 kDa molecular weight form had NADPH-linked reductase activity. In addition to coenzyme specificity these two forms also differed in their affinities for xylose and NADPH. Ditzelmuller et al. [20,21] also reported NADPH-linked activity associated with 35 kDa to 40 kDa molecular weight but no molecular weight has been reported for NADPH-linked form of the enzyme. Contrary to the existence of two forms of enzymes with different molecular weights, Bolen et al. [23] isolated a single reductase form having NADPH-/NADHlinked activity and a molecular weight of 36 kDa. The discrepancy in the molecular weight has been ascribed to different methods of determinations, van Cauwenberge

123 et al. [32] characterized two forms of enzyme from cell free extracts of anaerobically grown cells but with the same molecular weight (i.e. 36 kDa). Although, the molecular weight of the two forms of enzyme was the same, one form required either NADPH or NADH as a cofactor and showed an isoelectric point of 5.1, whereas the other form required only NADPH as a cofactor and had an isoelectric point of 6.4. Both these forms of reductase with same molecular weight (36 kDa) has been obtained from cell free extracts of aerobically grown cells as well but NADPH-active form predominated. However, from this study it is not clear that whether the enzyme exists as two separate proteins or as a single protein that undergoes modification. Girio et al. [33] reported that under anoxic conditions NADPH is preferred as coenzyme for xylose reductase in Candida shehatae. Variations in culture conditions showed a different NADH/NADPH ratio coupled to xylose reductase. Nevertheless, presence of two independent enzymes in C. shehatae needs to be evaluated. The presence of xylose reductase activity in Saccharomyces cerevisiae has been reported though the activity was five times lower than that of Candida utilis [14]. Xylose reductase obtained from Fusarium oxysporum exhibited a pH optima at 6.5-7.2 [27]. Both NADH and NADPH-linked activity of xylose reductase has been observed in this organism. In addition, F. oxysporum also exhibited a variation in the relative amount of enzymatic activity depending upon culture conditions [28]. In addition to yeasts and fungi, some bacteria eg. Enterobacteria, Corynebacteria and Brevibacteria also employ NADPH-dependent xylose reductase for conversion of xylose to xylitol [34,35]. Further metabolism of xylitol in these organisms involves oxidation of xylitol to D-xylulose-5-phosphate in the presence of enzyme pentitol dehydrogenase [3,36]. Some bacteria employ unusual pathways for the metabolism of D-xylose and xylitol [37-40]. For example, Arthobacter sp. employs the enzyme NAD-xylose dehydrogenase, catalyzing the formation of D-xylanolactone which is subsequently hydrolyzed to xylonic acid [39]. NAD-xylose dehydrogenase D-Xylose + NAD

> D-Xylanolactone + NADPH + H§

The purified enzyme exhibits specificity for NAD and D-xylose [40]. Some strains of

Lactobacillus casei grow anaerobically on xylitol or ribitol by utilizing an unique pathway [37]. Pentitols are transported into cells by a substrate specific phosphoenolpyruvate (PEP)-phosphotransferase system which converts them to their corresponding phosphates, e.g. xylitol-5-phosphate and ribitol-5-phosphate. Pentitol

124 phosphates are then converted to pentulose phosphate in the presence of NADspecific ribitol/xylitol phosphate dehydrogenase. In general xylitol is re-oxidized by most yeasts and fungi to D-xylulose by enzyme xylitol dehydrogenase [15,26,41]. This enzyme has been reported in cell free extracts of Candida utilis [14,41,42], Candida albicans [12], Pullularia pullulans [43], Cephalosporium chrysogenum [24], Pachysolen tannophilus [18,23], Saccharomyces cerevisiae [14], and Fusarium oxysporum [27,28,44]. Xylitol dehydrogenase purified from Candida utilis showed that enzyme lacked activity towards xylitol in the presence of NADP, NADPH or NADH as a cofactor [41]. The enzyme exhibited specificity towards xylitol and D-xylulose. The enzyme has the pH optimum of 9.0 and the equilibrium of reaction at neutral pH favoured polyol formation. The enzyme has been purified from Candida albicans [12], Pichia stipitis [17] and Cephalosporium chrysogenum [24] as well. Maleszka et al. [45] isolated mutants of Pachysolen tannophilus defective in xylitol dehydrogenase. These mutants failed to grow on Dxylose or xylitol but they exhibited growth on D-xylulose [45]. This has provided genetic evidence for the involvement of xylitol dehydrogenase in D-xylose metabolism in pentose fermenting yeasts. Surprisingly xylitol that is formed as a common intermediate of xylose catabolic pathway in yeasts and fungi, acts as poor substrate for growth of these organism. This inability of yeast and fungi to utilize xylitol has been attributed to limited permeability of xylitol [46].

2.1.2 Xylose isomerase pathway

Most of the bacteria employ the xylose isomerase pathway for catabolism of D-xylose. The enzyme xylose isomerase, known to convert D-xylose to D-xylulose in one step, has been reported in many bacteria, some actinomycetes and a few yeasts. Amongst bacteria Aerobacter aerogenes [47], A. cloacae [48-50], Bacillus coagulans [51-53], B. megaterium [54,55], Bacteroides xylanolyticus [56], Brevibacterium pentosoaminoacidicum [57], Escherichia coil [58], E. intermedia [59,60], Flavobacterium arborescens [61], Lactobacillus brevis [62,63], L. gayonii [64], L. pentosus [65], L. xylosus [66], Pasteurella pestis [67], Pseudomonas hydrophila [68,69] and Salmonella typhimurium [70,71] are known to have xylose isomerase activity. Among actinomycetes, Actinoplanes missouriensis [72] and various species of Streptomyces [73-77] are known to have this enzyme. Xylose isomerase (EC

125 5.3.1.5) catalyzes reversible isomerization of an aldopentose to its corresponding ketoisomer. Thus the enzyme catalyzes isomerization of D-xylose to D-xylulose. This enzyme has also gained commercial importance for the production of high fructose syrup due to its ability to catalyze reversible reaction of glucose to fructose [78]. Xylose isomerase is known to be inducible in the presence of D-xylose. The optimum temperature ranges from 45~ for the enzyme obtained from Lactobacillus brevis [79] to 90~ for the enzyme derived from Actinoplanes missouriensis [80]. However, the enzymes obtained from most of the sources have an temperature optima greater than 65~ [81]. The optimum pH of this enzyme is generally greater than 7, except for the enzyme produced by Lactobacillus brevis that has lower pH optima of 6.5 [81 ]. Factors such as substrates, presence of cofactors or immobilization are known to alter the pH optima [81]. For example, the optimum pH of xylose isomerase derived from Bacillus coagulans varies from 7-7.5 with glucose or ribose as substrates and 8-8.5 with xylose as substrate [82]. The optimum pH range of the enzyme from Streptomyces phaeochromogenes is 9-9.5 but it can be extended to pH 7.5 in the presence of Co2§ as a cofactor [50]. The optimum pH range of the enzyme immobilized from Actinoplanes missouriensis lies in a broad pH range of 6.5 to 8 where as optimum pH of free enzyme is 7.5. This broad range of pH of immobilized enzyme has been attributed to the microenvironment surrounding the immobilized enzyme [83]. The enzyme requires divalent cations such as Mn2+,Co2§ 2+ for its activity~ The optimum concentration required is 0.2-0.7 mM of Co2§ The enzymes isolated from various sources differ in their specificity with respect to cofactor requirement. For example, Lactobacillus brevis requires Mn2§ but Co 2§ can partially substitute for Mn 2§ whereas Bacillus coagulans requires Co2§ for activity of enzyme, Mn 2§ and Mg2§ can replace it only partially [84]. The presence of inducible xylose isomerase has been reported in yeast and fungi such as; Candida utilis [85], Rhodotorula gracilis [86] and Penicillium brevicompactum [87]. However, there is little evidence to support the view that this enzyme plays a crucial role in D-xylose catabolism by yeasts and fungi. Tomoyeda and Horitsu [84], purified xylose isomerase from xylose grown C. utilis. The enzyme showed specificity for D-xylose and L-xylose but not for D-arabinose or L-arabinose and required bivalent cations e.g. Mn2§ Mg2§ and Co 2§ Mn2§is most effective of these ions followed by Co 2§ and Mg 2§ The enzyme has an optimum pH of 6-7 and an optimum temperature of 70~ The same strain has also been reported to possess Daldose reductase activity as well and at 20 fold higher specific activity [13]. In another study xylose isomersae activity in xylose grown cell free extracts of Rhodotorula

126

gracilis has been reported as an obligatory step for xylose catabolism [86]. However, other groups of workers failed to detect xylose isomerase activity in cell free extract of xylose grown Candida utilis [26,41,46] and Candida albicans [11]. Fungi, such as Penicillium chrysogenum, Aspergillus and Fusarium also lack xylose isomerase activity [81]. Thus it appears that though the possibility of direct isomerization of D-xylose to D-xylulose may exist in certain yeasts and fungi as an alternative mechanism of xylose catabolism, a definite role of the enzyme in these organisms needs to be established.

2.1.3 Phosphorylation

Once D-xylose is converted to D-xylulose either by isomerase (as in bacteria) or oxidoreductase (as in yeasts and fungi), further metabolism proceeds by a phosphorylation reaction. The reaction is catlyzed by xylulokinase (E.C. 2.7.1.17) that converts D-xylulose to D-xylulose-5-phosphate. Chiang and Knight [26] detected xylulokinase activity in cell free extracts of Penicillium chrysogenum. Xylulokinase from bacterial sources such as Lactobacillus pentosus [88] and Aerobacter aerogenes [89] has been characterized. The enzyme has also been reported in the cell free extracts of Candida utilis grown in the presence of D-xylose or L-arabinose [41 ]. Although there are very few reports on xylulokinase activity in yeasts and fungi, the ability of many yeasts to utilize D-xylulose both under aerobic and anaerobic conditions indirectly implicates the presence of this enzyme in yeasts and fungi [81,90-92]. Nonetheless, Flanagan and Waites [93] purified and characterized xylulokinase from the pentose fermenting yeast Pichia stipitis. The enzyme exhibited a Km of 5.2x10-4 M for Dxylulose, and high specificity towards D-xylulose. Xylulokinase from bacterial sources such as Aerobacter aerogenes and Lactobacillus pentosus also exhibited high specificity towards D-xylulose as substrate [88,89]. The pH optima of enzyme from

Lactobacillus pentosus and Pichia stipitis has been reported to be between 7.0 to 7.9. In general the enzyme requires Mg §247 for its activity. The molecular mass of native enzyme from Pichia stipitis appears to be 120-130 kDa and comprised of two identical subunits of 71 kDa [93]. Isolation of xylulokinase gene and inability of organisms to grow in absence of this enzyme provides genetic evidence for its involvement in Dxylose catabolism [94].

127 2.2

Conversion of D-xylulose-5-phosphate to various end products

D-Xylulose-5-phosphate is further metabolized in both yeasts and bacteria via the pentose phosphate pathway [5-7,41,95,96] or phosphoketolase pathway [97-101]. Pentose phosphate pathway utilizes enzymes transaldolase and transketolase to convert D-xylulose-5-phosphate to D-glyceraldehyde-3-phosphate and D-fructose-6phosphate. D-glyceraldehyde-3-phosphate enters the glycolytic pathway leading to the formation of pyruvate. Alternatively, enzyme phosphoketolase (E.C. 4.1.2.9) has been suggested to be involved in pentose degradation. The enzyme phosphoketolase has been found in both bacteria [99-101] and yeasts [97,98] catalyzing the cleavage of either D-xylulose-5-phosphate or D-erythrose-4-phosphate: Phosphoketolase D-Xylulose-5-P + phosphate .... > Glyceraldehyde-3-P + AcetyI-P + H20 Phosphoketolase activity was initially considered to be specific for Lactobacilfi and Leuconostoc [102,103] but was later detected in a number of bacteria such as Pediococcus pentosaceus [ 104,105], Bifidobacterium bifidum (Lactobacillus bifidus) [106], Bacteriodes ruminicola [107] and Thiobacillus [ 108,109]. The enzyme has also been reported in many yeasts and fungi including Rhodotorula graminis, Rhodotorula glutinis, Candida tropicalis, Candida humicola, Candida 107 and Fusarium sp. [97,98, 102-105]. However, exclusive metabolism of pentoses via phosphoketolase pathway has been controversial. Evans and Ratledge [97] observed the presence of pentulose (D-xylulose-5-phosphate) ketolase activity in 20 out of 25 yeasts they examined. While the enzyme activity was insignificant in D-glucose grown yeast cells, it was induced up to 70 fold in cells grown on D-xylose as sole carbon source. On the basis of high yield and biomass and ethanol obtained by pentose fermenting yeasts these authors opined that phosphoketolase plays a major role in pentose metabolism in yeasts. Nevertheless, some pentose units are channelled to pentose phosphate pathway to provide metabolic intermediates and NADPH for biosynthetic purposes. However, Ligthelm et al. [106] could not demonstrate the presence of phosphoketolase activity in Pichia stipitis cells as 13C-xylulose yielded 2-13C ethanol. Nevertheless, Girio et al. [33] suggested that in addition to the pentose phosphate pathway, phosphoketolase pathway also operates in Candida shehatae. They have suggested the importance of

128 phosphoketolase pathway for NADH dissimilation as the reduction of acetyl phosphate to ethanol oxidizes NADH produced during the conversion of xylitol to D-xylulose. Based on ~"C labeling experiments in Fusarium sp. it has been demonstrated that Dxylulose-5-phosphate is cleaved by phosphoketolase between C-2 and C-3. The product labeling pattern indicated that ethanol and CO2 are products of 3-carbon fragment [105]. However, much attention is required to unequivocally establish the importance of phosphoketolase in pentose metabolism. Pyruvate is further metabolized to various end products in different microorganisms. The nature of end product(s) and metabolic routes also depends on the physiological conditions. In yeasts, pyruvate is either converted to ethanol by action of pyruvate decarboxylase and alcohol dehydrogenase under anaerobic conditions or is oxidized to CO2 and water via citric acid cycle under aerobic conditions. The pentose phosphate pathway in yeasts is stimulated by the oxidation of NADPH [1 10,11 1]. Thus availability of NADP/NADPH appears to control D-xylose reduction as well as activation of pentose phosphate pathway. Jeffries [112], suggested that D-xylose metabolism in yeasts is coordinately regulated. The regeneration of NADPH required for reduction of D-xylose to xylitol is produced via the pentose phosphate pathway and by the subsequent oxidation of hexose phosphate. For each mole of D-xylose metabolized to CO2, 10 mol of NADPH can be generated, which in turn can be utilized for the conversion of 10 mol of D-xylose to 10 mol of Dxylulose through an intermediate formation of xylitol [81]. Under aerobic conditions many yeasts have the potential to produce polyhydric alcohols such as xylitol, glycerol, and D-arabitol from D-xylose [81,113]. Other end products of D-xylose catabolism are acetic acid [1 12], citric acid [113] and xylonic acid [1 14]. In bacteria, pyruvate is metabolized to a number of end products. For example, pyruvate is cleaved by pyruvate ferredoxin oxido-reductase in the presence of coenzyme A to yield CO2, acetyl CoA and reduced ferredoxin in solvent producing Clostridium species. Figure 2 depicts conversion of D-xylose to various solvents in Clostridium. Acetyl CoA acts as central intermediate in branched fermentation pathway. During the initial growth phase (acidogenic phase) Clostridia produces acetate, butyrate, hydrogen and CO2 that results in a decrease in pH of culture medium. However, in stationary phase of growth, the metabolism of cells shifts to solvent production (solventogenic phase) [115,116]. The carbon flow from acetyl CoA leading to formation of acids and solvents occur at three key intermediates of pathway; acetyl CoA, acetoacetyl CoA and butyryl CoA. During acidogenic phase acetyl CoA and butyryl CoA form acetate and butyrate in an analogous manner but involve different sets of enzymes. The phosphate acetyl transferase and acetate

129 D-XYLOSE

D-XYLULOSE (2)

~

D-XYLULOSE-5-P ~ entosephosphateand Glycolyticpathway

4, dox

PYRUVATE

H2

Fdr,,d ACETATE< (6) ACETYL-P< (5) ACETYL-COA

)

~(91

ACETALDEHYDE .,L,(8) ETHANOL

ACETONE (1~ -~-1)ACETOAOETAT~I~-0) ACETOACETYL-COA ISO P RO PANO i..

3-HYD ROXYB UTY RYL-C CA ,L 14) CROTONYL-COA

/

(17)

BUTYRA TE <

__('16)

(15)

"~ BUTYRYL-P~-:-- BUTYRYL-COA

~ ~

(18)

BUTYRYLALDEHYDE (19) BUTANOL Figure 2. Conversion of D-xylose to various solvents and acids by Clostridium, Products formed during acidogenic phase are shown in italics and those formed during solventogenic phase are shown in bold. Enzymes are indicated by numbers as follows: (1) xylose isomerase (2) xylulokinase (3) pyruvate ferredoxin oxidoreductase (4) hydrogenase (5) phosphate acetyl transferase (6) acetate kinase (7) acetaldehyde dehydrogenase (8) ethanol dehydrogenase (9) acetyl CoA acetyl transferase (10) acetoacetyl CoA: acetate/butyrate: CoA transferase (1 1) acetoacetate decarboxylase (12) isopropanol dehydrogenase (13) 3-hydroxy butyryl CoA dehydrogenase (14) crotonase (15) butyryl CoA dehydrogenase (16) phosphate butyryl transferase (17) butyrate kinase (18) butyraldehyde dehydrogenase (19) butanol dehydrogenase.

130 D-XYLOSE

D-XYLULOSE ~ (2) D-XYLULOSE-5-P ~ (3n)tosephosphateand Glycolyticpathway PYR~VATE (4)

CItnc CO2 + H20 ' ' acid cycle~

o~-ACETOLACTATE

ACET

2,3-

~L (5) METHYLCARBINOL

B~(6)

TANEDIOL

Figure 3. Bacterial catabolism of D-xylose to 2,3-butanediol. Enzymes are indicated by numbers as follows" (1) xylose isomerase; (2) xylulokinase; (3) transaldolase and transketolase; (4) acetolactate forming enzyme; (5) acetolactate decarboxylase (6) acetoin reductase.

131 kinase mediate the formation of acetate whereas phosphate butyryl transferase and butyrate kinase lead to the formation of butyrate [114,117-120]. During solventogenic phase ethanol and butanol are produced from acetyl CoA and butyryl CoA, with the formation of acetaldehyde and butyraldehyde in the presence of butyraldehyde dehydrogenase and then to butanol in the presence of butanol dehydrogenase [121123]. In both C. acetobutylicum and C. beijerinckiithe activity of butanol dehydrgenase has been found to be NADPH dependent [124,125]. A number of different mechanisms have been proposed for the shift in metabolism of Clostridium from acidogenic to solventogenic and have been reviewed earlier [115,116]. It has been suggested that acetate and butyrate are reassimilated by the action of enzyme acetoacetyl CoA: acetate/butyrate: CoA transferase. This enzyme can utilize either acetate or butyrate as CoA acceptor during conversion of acetoacetyl CoA to acetoacetate [121]. The acetoacetate thus produced is then decarboxylated by acetoacetate decarboxylase to form acetone [5,121]. This irreversible step has been suggested to be responsible for driving reactions towards the formation of acetoacetate [126]. In Clostridium beijerinekii acetone is further reduced to isopropanol by the action of isopropanol dehydrogenase [115,116]. In Klebsiella pneumoniae pyruvate is metabolized by the action of different sets of enzymes that lead to the formation of 2,3-butanediol as a major end product (Figure 3). Two moles of pyruvate is condensed to form acetoacetate in the presence of enzyme "pH 6-acetolactate forming enzyme" [127,128]. Acetolactate is then decarboxylated in the presence of the enzyme acetolactate decarboxylase to yield acetoin (acetyl methyl carbinol) [129]. Acetoin is finally reduced to 2,3- butanediol by the action of acetoin reductase [130]. Further studies on acetoin reductase suggest the presence of two stereospecific enzymes. One is specific for reduction of L-acetoin to L-butanediol; while the other reduces D-acetoin to meso-butanediol [131]. In Klebsiella planticola pyruvate is dissimilated by the enzyme pyruvate formate lyase to yield acetate, ethanol and formate in a molar ratio of 1:1:2, whereas, D-lactate is formed in small amounts [132].

3

REGULATION OF D-XYLOSE METABOLISM

3.1

Genetic Regulation

The metabolism of D-xylose in microorganisms starts with its transport into the

132 cell. Therefore, the initial regulatory steps of D-xylose metabolism also begin at the level of transport of D-xylose and have been discussed in Chapter 4 of this volume. Once D-xylose is taken up inside the cell further conversion of D-xylose to D-xylulose5-phosphate involves inducible enzymes. Most studies on inducibility of pentose metabolizing enzymes in yeasts and fungi have concentrated on xylose reductase and xylitol dehydrogenase. Nonetheless, inducibility of xylose isomerase has also been noted in few yeasts and has been discussed in section 2.1.2. Induction of xylose reductase and xylitol dehydrogenase has been reported in Pachysolen tannophilus, Pichia stipitis, Pullularia pullulans and Fusarium oxysporum [18,20,27,28,44,133,134]. The induction of these enzymes has been shown to be specific to D-xylose. Although L-arabinose has been found to induce both of these enzymes, in P. tannophilus, other sugars such as D-glucose, Dfructose, D-mannose, mannitol, sorbitol and glycerol failed to induce them [18,133135]. The induction of these enzymes has been demonstrated in P. tannophilus as messenger RNA encoding these enzymes are not synthesized in the presence of Dglucose [136]. In addition, enzyme phosphoketolase has also been shown to be inducible in presence of D-xylose in many yeasts [97]. D-Xylose catabolism is also subject to enzyme repression. It has been observed that when cells of Pachysolen tannophilus, Pichia stipitis, Candida shehatae, Candida utilis, Candida steatolytica are grown in the presence of both D-glucose and D-xylose, D-glucose is preferentially utilized over D-xylose [18,137-139]. This phenomenon has been attributed to catabolite repression, which is commonly exerted by D-glucose and related sugars in yeasts [140,141]. Experimentally, it has been demonstrated that in Pachysolen tannophilus and Pichia stipitis induction of xylose reductase and xylitol dehydrogenase activities are inhibited by D-glucose, D-mannose or 2-deoxyglucose, whereas cellobiose, D-galactose and L-arabinose are not inhibitory [18]. The extent of repression of these two enzymes, however, varied in Pachysolen tannophilus. Xylitol dehydrogenase activity is repressed to a greater extent by Dglucose and D-mannose than xylose reductase activity. In addition 2-deoxyglucose represses xylitol dehydrogenase activity but not the xylose reductase activity. This suggests that catabolite repression of these enzymes is not coordinately controlled in Pachysolen tannophilus. Regulation of D-xylose utilization in the presence of hexoses other than D-glucose and D-mannose varies among the yeasts [18,135,142]. In addition to induction and catabolite repression of xylose metabolizing enzymes, inactivation of xylose utilization has also been reported in some pentose fermenting yeasts. For example, addition of 8% (w/v) glucose to D-xylose grown cells of Pichia stipitis and Candida shehatae resulted in an immediate inactivation of D-

133 xylose utilization [138]. However, extent of such inactivation differs amongst the yeasts. While D-xyiose utilization is completely inhibited in Pichia stipitis, only partial inhibition is observed in Candida shehatae [138]. In bacteria, enzymes involved in early steps of xylose catabolism i.e. xylose isomerase and xylulokinase have been demonstrated to be inducible. Inducibility of these enzymes has been observed in Aerobacter aerogenes [143], E. coil K-12 [58] and Salmonella typhimurium K-12 [70]. Induction of these enzymes has been studied in Salmonella typhimurium in greater details [10]. Studies indicate that addition of inducer, D-xylose, resulted in induction of xylulokinase followed by xylose isomerase and xylose permease enzymes. All other pentoses and pentitols failed to induce xylose isomerase and xylulokinase activity [70]. The synthesis of xylose isomerase has been found to be subject to catabolite repression which is reversed by the addition of cyclic adenosine monophosphate [70]. Based on genetic studies Shamanna and Sanderson [144], suggested the presence of a regulatory gene xylR in Salmonella typhimurium, which produces an activator molecule that regulates xylose catabolizing enzymes. It has been proposed that the activator molecule possesses two sites, one site for attachment to operator type locus for the xylB (encoding xylulokinase) and for xylT (encoding xylose permease) and another site for binding xylose. Interaction of xylR gene product and D-xylose converts inactive activator to an active state, which activates xylA (encoding xylose isomerase), xylB and xylT genes. Similarly positive (activator) elements have been shown to regulate the other inducible operons of L-arabinose [145] and D-ribose [146] in E. coil as well.

3.2

Oxygenation and cofactor regulation

The role of oxygen for efficient ethanol formation during pentose fermentation by yeasts has been studied by many workers. Studies suggest that oxygen plays the following important roles in pentose metabolism by yeasts: 1

Oxygen is required to maintain the redox balance in the first step of xylose metaboilsm [16,31,1 47].

2 3

Oxygen is required for the transport of D-xylose [106,148]. Oxygen is required for growth [149].

4 Oxygen is required for unimpaired mitochondrial function [149,150]. It is known that many yeasts which can assimilate and metabolize xylose in aerobic

134 conditions are unable to ferment it to ethanol in anaerobic conditions e.g. Candida utilis, or can ferment it at very slow rates e.g. Pachysolen tannophilus. The inability of yeasts to ferment xylose anaerobically has been linked to cofactor regeneration [151]. In these genera of yeast such as Candida utilis, the cofactor requirement of xylose reductase (NADPH) differs from the cofactor requirement of xylitol dehydrogenase (NAD). Thus fermentation halts in these organisms unless NADH produced at the second step is reoxidized. However, a different mechanism exists in Pachysolen tannophilus and Pichia stipitis. These yeasts are capable of fermenting xylose to ethanol under anaerobic conditions. In both these yeasts xylose reductase and xylitol dehydrogenase can utilize NAD or NADP as cofactor [152]. Thus reduced cofactor generated in one step is utilized in the other. Nevertheless both species fail to grow in the absence of oxygen, probably due to disturbances in redox balance during these fermentations. Skoog and coworkers demonstrated that oxygenation has an influence on intracellular enzymes and metabolites during D-glucose and D-xylose fermentation [153]. Since cofactor requiring oxidative reductive pathway does not operate in glucose fermenting cells [154], it has been suggested that oxygenation does not influence the activity of xylose reductase and xylitol dehydrogenase [153,155] and requirement of oxygen in Pichia stipitis has been attributed to sugar transport, growth and proper mitochondrial function. Oxygenation has been shown to influence xylose fermentation significantly with respect to the nature of product and yield [152]. In this regard Pachysolen tannophilus has been investigated most thoroughly [156]. Pachysolen tannophilus produces cell mass under aerobic conditions, and xylitol under anaerobic conditions whereas under oxygen limiting conditions ethanol is produced [157-159]. In addition increased oxygenation leads to reassimilation of ethanol in this organism [160] and Candida tropicalis [ 161 ]. In general, it has been observed that yeasts that can produce appreciable amounts of ethanol from D-xylose exhibit increased xylitol accumulation with increasing oxygen limitation [159,162-165]. Metabolic basis for this appears to be the role of oxygen in cofactor regulation. It has been proposed that when oxygen is readily available, NADH produced by the action of xylitol dehydrogenase is rapidly oxidized to NAD by the respiratory chain. Such removal of NADH and production of NAD drives the xylitol dehydrogenase to produce D-xylulose and therefore limits xylitol accumulation. However, under oxygen limitation, the rate of NADH oxidation decreases which in turn decreases the rate of conversion of xylitol to D-xylulose. In addition, each molecule of D-xylulose entering the pentose phosphate pathway produces several NADPH molecules which tends to drive the xylose reductase

135 reaction and leads to xylitol accumulation. Efforts have been made to shift the metabolism of yeasts favouring ethanol over xylitol as end product by using metabolic inhibitors [166,167]. Early studies indicated that azide, by fine tuning of respiratory activity, shifts xylose metabolism towards ethanol formation in Candida tropicalis [166]. Similarly polyethylene glycol has been shown to shift metabolic activity of Candida tropicalis favouring ethanol formation by controlling aeration in the medium. However, later studies suggest that azide has such an effect on metabolism due to its uncoupling action [167]. Oxygen has been shown to be required by Fusarium oxysporum for D-xylose fermentation as it is known to induce key enzymes of D-xyiose catabolism in this organism [44]. The organism exhibits different fermentative capabilities when grown under different aeration conditions [44]. Highest level of ethanol and acetic acid are produced when NADH/NADPH ratio is highest, under semiaerobic conditions. Thus, oxygen plays an important role in channeling the biosynthetic intermediates of Dxyiose metabolism in fungi as well [44]. In contrast to yeasts and fungi, where initial steps of D-xylose catabolism involves cofactor dependent enzymes, bacteria employ isomerase enzyme which is independent of cofactor regulation. Thus initial steps of D-xylose catabolism in bacteria is generally not regulated by oxygenation. Nonetheless, anaerobic fermentation of pentose sugars to various solvents by bacteria has been widely acknowledged. Amongst various bacteria, Clostridium acetobutylicum has been studied in greater details. This bacteria can ferment pentose sugars to ethanol, butanol and acetone, in addition to the formation of acetate and butyrate [115,116]. Depending on the species, culture conditions and growth phase the proportion of end product varies considerably. For example, formation of ethanol by Clostridia is regulated by a number of key enzymes which in turn are regulated by end products or metabolic inhibitors [115,116]. Four oxidoreductases namely, (i) pyruvate-ferredoxin oxidoreductase: generates reduced ferredoxin during cleavage of pyruvate; (ii) NADH-ferredoxin oxidoreductase: regenerates NAD by production of reduced ferredoxin; (iii) NADPH-ferredoxin oxidoreductase: produces NADPH for biosynthesis; and (iv) ferredoxin-H 2 oxidoreductase (hydrogenase): regenerates reduced ferredoxins, play significant role in determining the reduced end product of the cell [116]. In addition acetaldehyde and ethanol dehydrogenases also play important role in determining the yield of the end product. Clostridium thermohydrosuifuricum contains both NADH-and NADPH-linked acetaldehyde and ethanol dehydrogenase which show end product inhibition at relatively low concentrations of ethanol [168,169]. Both increase in partial pressure of hydrogen and ethanol accumulation leads to increase in the pool of reduced ferredoxin

136 and

NAD(P)H

and thereby

resulting

in cessation

of carbon

flow

in

C.

thermohydrosulfuricum [169]. C. thermocellum on the other hand can dispose substantial amounts of reducing power as H2 and a large proportion of acetyl coA is converted to acetate. In addition an increase in partial pressure of H2 has little effect on growth of C. thermocellum [168,169]. This organism lacks the ability to redirect reducing power generated during pyruvate cleavage to produce ethanol and thus has very poor yield of ethanol. The presence of inducible lactate dehydrogenase also plays an important role in regulating ethanol production. Conditions favouring lactate formations such as increased NADH and fructose-l,6-diphosphate level reduces ethanol production [116]. As has been discussed earlier D-xylose is channelled to either acid or solvent production by Clostridium acetobutylicum depending on growth phase. The mechanism regulating acid or solvent production has been reviewed by Jones and Wood [115,116] and will be discussed only briefly. In C. acetobutylicum, the switch from acid production to solvent production is accompanied by decrease in activity of all the acid pathway enzymes except butyrate kinase and induction of solvent pathway enzymes [170-173]. The reassimilation of acids occurs after induction of solvent producing enzymes. The enzyme CoA transferase due to broad specificity uses either acetate or butyrate as CoA acceptor and converts butyrate and acetate to butyryl CoA and acetyl CoA favouring solvent production. This shift of acid to solvent production enables the cells to avoid the inhibitory effect due to high concentrations of acid. The re-utilization of acids during solvent production further reduces the toxic effect of acids.

4

METABOLISM OF L-ARABINOSE

The utilization of L-arabinose is of particular significance as it is the second major pentose sugar present in hydrolysates of lignocellulosic biomass. The metabolism of L-arabinose has been shown to be similar to D-xylose. Chiang and Knight [26], using cell free extracts of a mycelial fungi Penicillium chrysogenum, proposed the following pathway for L-arabinose catabolism (Figure 4).

137 L-ARABINOSE (I)

~

L-ARABITOL

L-XYLULOSE

~ ~

(3)

XYLITOL (2)

D-XYLULOSE

Figure 4. L-Arabinose catabolism in yeasts and fungi. Enzymes are indicated by numbers as follows: (1) NADPH-dependent aldopentose reductase; (2) pentitoI-NADdehydrogenase; (3) L-xylulose-NADPH reductase.

A similar mechanism of L-arabinose metabolism has also been proposed for yeasts [6]. It appears that both D-xylose and L-arabinose utilize the same enzymatic system for their metabolism in yeasts and fungi. Evidence for this comes from genetic studies. For example, a mutant of Pachysolen tannophilus deficient in NADPH dependent xylose reductase displays significantly reduced growth rates compared to their wild type on L-arabinose as well as on D-xylose [8]. L-Arabinose, on the other hand, has been shown to induce the activity of both xylose reductase and xylitol dehydrogenase in Pachysolen tannophilus [133]. Although Horitsu et al. [42] have identified and partially purified an L-arabinose isomerase from Candida utilis, the role of this enzyme in yeasts and fungi has not been confirmed. Bacteria on the other hand employ isomerase pathway for L-arabinose metabolism (Figure 5). E. coil B/r has been widely employed for the studies of Larabinose transport, metabolism, regulation and genetics [145]. Studies have shown the presence of a set of enzymes for L-arabinose metabolism in this bacteria encoded by BAD operon where araB encodes the kinase, araA encodes the isomerase and araD encodes epimerase [174-178]. The isomerase reversibly converts L-arabinose

138 to L-ribulose, which is phosphorylated to L-ribulose-5-phosphate in the presence of a kinase. L-Ribulose-5-phosphate in the presence of enzyme L-ribulose-5-phosphate epimerase is converted to D-xylulose-5-phosphate. Regulatory mechanisms involved in the L-arabinose transport and metabolism has been studied in detail [145]. Genes are transcribed in the order: araB, araA and araD. Expression of operon is under specific control of product of araC, which is autoregulatory and repressed by the regulatory protein-inducer complex and repressed by the regulatory protein alone. A similar mechanism of L-arabinose metabolism has been suggested in S. typhimurium as well [178-180].

,~(1) ~

L-ARABINOSE

L-RIBULOSE (2)

L-RIBULOSE-5-P

D-XYLULOSE-5-P

Figure 5. L-Arabinose catabolism in bacteria. The enzymes are indicated by numbers as follows: (1)isomerase; (2) kinase; (3) epimerase.

It is apparent from the foregoing discussion that considerable efforts have been made to understand the microbial metabolism of pentose. Basic studies related to pentose uptake and metabolism have provided the basis for genetic improvement of pentose utilizing microorganisms. For example, cloning and expression of xylose isomerase gene in yeast to bypass the oxidative-reductive pathway, and cloning and expression of xylulokinase gene in pentose fermenting yeasts to shift the equilibrium of conversion of D-xylose to D-xylulose-5-phosphate has significantly improved genetic traits of pentose fermenting microorganisms. In addition, cloning of genes of xylose assimilating and pentose phosphate pathway enzymes in Zymomonas mobilis has been employed to widen the substrate utilizing ability of well established ethanologenic organism. Regulatory mechanisms such as repression and inactivation of pentose metabolism in the presence of other sugars poses major limitation in efficient

139 utilization of pentose sugars from lignoceilulosic wastes. Understanding of such regulatory mechanisms are necessary for further genetic improvement of strains. While a tremendous amount of research efforts have been directed towards bioconversion of pentoses to ethanol, relatively little is known about the bioconversion of pentoses to other solvents, organic acids, xylitol, food, and feed. However, with growing knowledge of metabolism, genetics and molecular biology of thermophiles, especially Clostridium, may help in improving the process of solvent production. Research efforts to understand biochemistry and genetics of filamentous fungi is also likely to overcome some of the limitations of fermentation routes.

5

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6 Microbial Production of Ethanol

1

INTRODUCTION

Energy is the essential and basic commodity for human activities. The constant increase in energy consumption has caused a global energy crisis. According to an estimate, the world will require two third more energy in the year 2000 than it did two decade ago [1]. The United States demand in 1978 was 41% of the total world demand, and it is expected that there will be a drop by about 33% by the year 2000. The share of Europe is expected to drop, whilst no change in Japan's demand is anticipited [2]. Developing countries are projected to increase their energy demand from 19% in 1978 to 33% in the year 2000. During last 70 years, crude oil was the predominant fuel and industrial raw material. The advantages related to stability, diversity in applications, availability and relatively low price made it an ideal raw material for large organic chemical industry. The oil crisis of mid 1970s has already alarmed us of dwindling oil reserves. Fossil fuels are rapidly becoming precious commodity, which is reflected in their increasing price. Oil constituted 54% of the world energy supply in 1970s which indicates its extensive use. In spite of limited quantities of crude oil, its share in the global energy supply is expected to be predominant in the future. In order to maintain the present industrial potential of the developed countries and to allow for industrial and economic expansion of the underdeveloped countries, development of alternative fuels is required which can at least partially substitute the diminishing petroleum resources. Alcohols (ethanol and methanol) alongwith other fuel commodities have been considered to play a role of promising potential. Ethanol has almost all the advantages of liquid hydrocarbon fuels, its available enregy content per unit volume being lower than gasoline (Table 1). The energy density of ethanol is 65% that of gasoline. Ethanol also has a higher octane rating, (R+M/2=102), than does gasoline, and causes disproportionate increase in ocatane when blended with gasoline [3]. The blending octane rating is 119 for ethanol, 120 for methanol, 975 for tert-butanol, and 108 for methyl tert-butyl ether. Methanol boosts octane as effectively as ethanol, but blending methanol with gasoline results in greater difficulty of phase separation, materials 147

148

incompatibilty, and vapour lock [4,5]. Alcohol blends were used with success in over four million vehicles between World War I and II, it being generally admixed in proportions of upto 25% with gasoline. Besides being a fuel, ethanol is also a very versatile chemical feedstock. A wide variety of chemicals can be derived from ethanol. Ethanol has several other uses: (i) potable ethanol in alcoholic beverages; (ii) solvent ethanol for laboratory and pharmaceutical purposes; (iii) as an antiseptic; (iv) as a cosurfactant in oil-water microemulsions; and (v) as starting material for the synthesis of several chemicals.

Table 1 Available energy content of gasoline and alcohols Energy Fuel KJ/I (X 104)

BTU/gal

Gasoline

3.45

124,000

Ethanol

2.36

84,600

Methanol

1.80

64,800

Ethanol is produced commercially by chemical synthesis and fermentation. Practically all current ethanol is manufactured synthetically from petroleum, and beverage alcohol is produced by fermentation of molasses, cereal grains and other high starch and sugar containing materials. Industrial ethanol production based on carbohydrates from agricultural crops, was largely developed before World war II, especialy in Europe. After 1945, cheaper ethanol from petroleum products prohibited further development of fermentation ethanol. Since 1973, with the beginning of the energy crisis, more consideration has been given to the production of bioethanol, especially based on renewable resources. An advantage of using renewable resources is that they are available cheaply in large quantities which is of critical importance for large scale alcohol production. Prices of these (waste) materials are more competitive than some of the

149 conventional alcohol substrates such as grains. Availability of some of these renewable resources is even increasing. According to an estimate, wood available today in world forests is 1.5X101~ tons more (due to rising amounts of CO2 in the atmosphere and worldwide reforestation programs) than there was a century ago [6]. Considerable effort has been focussed on the microbial production of ethanol from sugars obtained by hydrolysis of lignocellulosic materials. The economics of bioconversion would be more favourable if both hexoses and pentoses could be converted to ethanol. The economic importance of obtaining ethanol from pentoses depends on the relative amounts of pentoses and hexoses. The increase in ethanol yield is expected when both sugar types are converted, over that for hexose alone, is proportional to the pentose content. Consequently obtaining ethanol from pentoses is particularly important when they are present in high amounts. An example of such feedstocks is hardwood, where Dxylose can comprise 20-34% by weight of the neutral sugars. Appreciable progress has been made recently in obtaining ethanol from pentose sugars using yeasts, moulds and bacteria. This chapter covers the state, of the art of microbial pentose fermentation to ethanol. The starting point is a description of potential microorganisms and the way they convert pentoses and natural lignocellulosics into ethanol, followed by the discussion on various factors which affect the yield and productivity of fermentation.

2

MICROORGANISMS USED FOR ETHANOL PRODUCTION

Views on the ability of yeasts to ferment aldopentoses were contradictory for a long period of time, although many of the microbial species known were able to assimilate D-xylose under aerobic conditions. However, some earlier studies have shown that a yeast could carry out the fermentation of D-xylose and L-arabinose [7,8]. During last two decades, a number of laboratories have demonstrated ethanol production from pentose sugars using different strains of yeasts, moulds and bacteria. Microorganisms differ in their ability to grow and metabolize aldopentoses. The demonstration of product accumulation from pentoses is sensitive to a number of experimental conditions which might have reflected in the origin of different views regarding the ability of microorganisms to ferment pentoses. Growth characteristics of some important pentose fermenting organisms are presented in Table 2.

150

Table 2 Growth characteristics of some important pentose fermenting organisms

Sugar type Organism

Optimum Xylose Arabinose M a n n o s e

pH

Optimum temperature

Yeast Candida

+

+

+

3-7

28-32

+

+

-

2.5-7

28-32

+

+

+

3-7

28-32

+

+

+

3-7

30-35

+

+

+

5-6

28-34

+

-

-

5-6

28-37

Paecilomyces sp.

+

+

+

2.2-7

30-37

Monilia sp.

+

-

-

5

26

+

+

+

6-8

35-37

+

+

-

4-8

65

+

+

+

5-8

60

+

+

+

4.4-9.5

6.9

shehatae Pachysolen tannophilus Pichia

stipitis Kluyveromyces marxianus

Filamentous fungi Fusarium oxysporum Neurospora crassa

Bacteria Bacillus macerans Clostridium thermocellum Clostridium saccharolyticum Thermoanaerobacter ethanolicus

151 2.1

Yeast

The applicability of yeasts in conversion of carbohydrates to ethanol has been studied extensively. Many of the yeast species are known to ferment hexoses (mainly D-glucose) to ethanol with significantly high yields. However, only few yeast strains have so far been reported which can ferment the pentoses to ethanol, even though many of them are known to metabolize pentose aerobically for growth. A great number of yeast strains have been investigated with reference to their capacity for the utilization of xylose and xylulose. With respect to xylose fermentation, yeasts which have been studied extensively are Pachysolen tannophilus, Candida shehatae, Pichia stipitis and

Kluyveromyces marxianus. Various other yeast species which have been investigated for their xylose fermenting ability include Brettanomyces, Clavispora, Schizosaccharomyces, several species of Candida viz. C. tenuis, C. tropicalis, C. utilis, C. blankii, C. friedrichii, C. solani, C. parapsilosis, C. sake, and species of Debaromyces viz. D. nepalensis, D. polymorpha. However, none of these yeast species have been found very promising. Toviola et al. [9] screened 200 strains of yeasts by employing classical method of measuring gas production as an indicator for fermenting strains. However, most of the strains selected as fermenting pentoses produced only small amounts of ethanol, and the authors concluded that criteria of gas production for screening pentose-fermenting yeasts was not reliable. Schneider et al. [10] reported that approximately 20% yeast species that give positive Durham-tube test on D-glucose, produced only about 1 g/I of ethanol from 20 g/I of D-xylose. Several positive yeasts under these test conditions were earlier not identified as xylose fermenting strains. Gong et al. [11] tested 20 strains of Candida, 21 strains of Saccharomyces and 8 strains of Schizosaccharomyces for their ability to ferment D-xylose. All of the Candida strains grew well on D-xylose, while the strains of

Saccharomyces and Schizosaccharomyces grew poorly or not at all. Xylitol was produced in the range of 10-50% by Candida strains with arabitol as the second major product. Ethanol was found to be the major product with most of the Candida strains. Schizosaccharomyces strains also produced ethanol but the concentrations were low (1-5 g/I). Different aeration conditions have been employed to the surveys designed to identify strains capable of xylose utilization. Roughly half of the yeast strains tested could utilize D-xylose for growth under aerobic conditions but not under anoxic condition [12].

152 Maleszka and Schneider [13] screened 15 yeasts for their ability to utilize D-xylose, D-xylulose and xylitol for ethanol production under aerobic, semi-aerobic (low aeration) and anaerobic conditions, with rich undefined or defined media. In almost all cases, ethanol production by P. tannophilus and species belonging to Candida and Pichia was better on rich media under semi-aerobic conditions. It was concluded that an experimental condition that is critically important in screening of pentose fermenting cultures as well as in determining the extent to which ethanol accumulates in cultures is the oxygen supply rate. Very low concentrations of ethanol will be produced when high aeration rates are provided [14-16]. Majority of yeasts can not ferment D-xylose directly [17,18]. It has been observed that instead they utilize D-xylulose, an isomer of D-xylose, both oxidatively and fermentatively. The best xylulose fermenting yeasts so far identified are species of

Brettanomyces, Candida, Hansenula, Kluyveromyces, Pachysolen, Saccharomyces, Schizosaccharomyces and Torulospora [19]. The pattern of D-xylulose utilization has been observed to be different from that of D-xylose utilization. D-xylulose can be readily utilized by Candida under aerobic or semi-aerobic conditions [11]. Ethanol concentrations from D-xylulose were generally higher and xylitol accumulation generally lower than when D-xylose was used as substrate. The pattern of D-xylulose utilization by different strains of Saccharomyces was also found similar to that of Candida. Maleszka and Schneider [13] found that P. tannophilus that ferment D-xylose, readily utilized D-xylulose. Similarly

C. blankii which readily assimilated D-xylose, and produced ethanol from D-xylulose [11]. However, S. pombe which do not assimilate D-xylose but readily utilize D-xylulose aerobically or anaerobically. Many yeasts can convert xylose to xylitol even though they are not able to grow on pentose [20,21]. Xylitol on the other hand, is generally a poor carbon source, even though it is an essential intermediate in D-xylose catabolic pathway. Maleszka and Schneider [13] tested 15 yeast strains and have reported only traces of ethanol produced from xylitol as carbon source. Out of the 16 strains tested by Barnett, only Torulopsis candida was found to have potential [22]. Pentitols are the major by-products of pentose fermentation and the ethanol production efficiency would be further increased if such compounds could be converted to ethanol. P. tannophilus can utilize xylitol but the necessary conditions differ from those of D-xylose. Pichia angophorae has been identified as the potential organism for polyol fermentation [23]. Direct conversion of hemicellulose polymer to ethanol is rare among yeasts. Of the 250 yeast strains screened, few strains of P. stiptis and C.

153 shehatae have been identified that simultaneously hydrolyse and ferment xylan to ethanol. This ability is of particular interest for developing a direct bioconversion process.

2.2

Filamentous fungi

The ability of filamentous fungi to ferment sugars has been known for about 70 years [24,25]. However, detailed studies have only recently been undertaken at several laboratories throughout the world. Several fungal species belonging to the genera Fusarium [26-31], Rhizopus [26], Monilia [32], Neurospora [33], and Paecilomyces [34] have been found to have potential for fermenting glucose as well as xylose. Fusarium oxysporum (lin1) was found to ferment glucose to ethanol and CO2 giving product yield similar to that of yeast [24]. Although xylose fermentation with filamentous fungi gives comparatively lower ethanol concentrations, appreciable yields could be obtained [35]. Suihko and Enari [36] screened 26 strains of F. oxysporum for their ability to convert D-glucose and D-xylose to ethanol. Except for strain VTT-D72014, all Fusarium strains were capable of fermenting both the sugars. Wene and Antonopoulos [37] screened over 2000 Fusarium strains, isolated from natural substrates, for their ability to utilize D-xylose. Almost all of the strains were capable of producing some ethanol ranging from 0.4 to 4.4 g/I ethanol from 10 g/I D-xylose. Besides D-xylose, many other sugars found in hemicellulose complex, such as D-glucose, D-mannose, D-galactose, and also xylitol and sucrose are fermented by Fusarium species [38]. The fermentation products formed by Fusarium sp. from pentoses or hexoses are similar consisting of equimolar quantities of ethanol and acetic acid [39]. Paecilomyces sp. NF1 was found to be able to convert all the major sugars present in the hydrolysate of plant biomass [34]. High ethanol concentrations (76 g/I) by fermentation of D-xylose can be obtained using this fungus. Yields and rates of ethanol production from D-xylose by Mucor species [40] and Neurospora crassa [33] have generally been low when compared to F. oxysporum [41]. Some potential fungal strains have been identified that ferment not only glucose and xylose but also more complex natural cellulosic substrates. Monilia sp. [42], N. crassa [43] and F. oxysporum [44-47] were found to have potential in conversion of cellu-

154 Iose/hemicellulose to ethanol/acetic acid in a single step. In this case, ethanol production is achieved by placing aerobically grown mycelia under anaerobic conditions. There are, however, several physiological parameters inherent in these fungi which makes the fungal ethanol process unattractive. These include (a)long generation and fermentation time, ranging from 4-8 days, (b) low volumetric rates of product formation, (c) growth in large clumps rather than as dispersed, single cells, (d) critical oxygenation levels (e) production of acetic acid alongwith ethanol as major by product, which results in the lower ethanol yield, and (f) low substrate and product tolerance. However, a fungal system might be of interest because of its ability of growing on natural plant biomass which a yeast system usually lacks.

2.3 Thermophilic bacteria

Thermophilic bacteria are best defined by their temperature characteristics for growth. These bacteria ferment a wide range of substrates including cellulose, hemicellulose, pectin, and starch. The different products formed include alcohols (ethanol, butanol, isopropanol and 2,3-butanediol), organic acids (acetic, butyric, formic and lactic acid), polyols (arabitol, glycerol), ketones (acetone) and gases (CO2, methane and hydrogen). The promising thermophilic pentose fermenting bacterial species include

Clostridium thermocellum, C. thermohydrosulfuricum, C. thermosaccharolyticum, C. thermosulfurogenes and Thermoanaerobacter ethanolicus [48-53]. Most of these organisms have been isolated from the hot springs of Yellowstone National Park underneath algal mats [54]. Almost all of them can be grown on defined media. They are relatively resistant to heavy metal ions [55,56], and other toxic substances and pollutants. Although they do not grow in the presence of oxygen, these anaerobes are not killed by the contact with air. Several properties of thermophilic bacteria have been identified which are advantageous for their industrial application. Some of the advantages are: Fermentation of a wide range of sugars present in cellulose and hemicellulose polysaccharides. High metabolic activity which results in faster fermentation. Less biomass formation with high specific product yield.

155 Higher temperature fermentation eleviates the risk of contamination. No oxygen requirement. Possibility of continuous product recovery by distillation of volatile compounds. The investigations on thermotolerance of the spores of thermophilic Clostridia revealed that they all have high heat resistance. Spores of C. thermohydrosulfuricum have a decimal reduction time 11 min at 121~ and over 12 h at 100~ which makes standard sterilization time greater than 30 min [57]. Controversies exist regarding the taxonomic status of some of C. thermosaccharolyticum strains [58] which do not form butyrate as is reported for the type and other strain [59,60]. A strain of C.

thermosaccharolyticum [59] synthesizes ethanol during secondary metabolism after growth is uncoupled by acid production in a manner analogous to butanol production by C. acetobutylicum. Reaction of C. thermohydrosulfuricum, C. thermosulfurogenes and C. thermosaccharolyticum are different with thiosulfate which either is reduced to H2S, deposits as sulphur, or not transformed, respectively [60]. These observations demand more taxonomic studies at molecular level.

2.4

Mesophilic bacteria

In contrast to yeast processes, bacterial systems characteristically generate multiple products. Although most yeasts and moulds can not ferment pentoses anaerobically, many bacteria readily convert xylose to a variety of products in the absence of oxygen [61]. These include Bacillus macerans, B. polymyxa, Klebsiella pneumoniae, Aeromonas

hydrophila and Aerobacter indologenes [62]. The rates yields, and product formed by these bacteria depend not only on the diverse metabolic pathways operating during anaerobic fermentation but also on the species, strains, substrates, and cultural conditions. The anaeobic fermentation of carbohydrates by B. macerans was first reported by Schradinger [63]. The Bacillus fermentation is characterized by ethanol and acetone as the main products. Weimer has studied the effect of pH of the fermentation [64]. It is chracterized by two distinct phases, where rapid growth and acetate, formate and ethanol

156 production phase precedes a slower growth and acetone production phase that is coupled with acetate and formate consumption. B. macerans can use a variety of substrates including xylan starch, major hexoses, and pentose sugars, including di- and trisaccharide derivatives. Bacterial strains of Lactobacillus and Leuconostoc convert xylose to ethanol, organic acids, and 2,3-butanediol. In general, these organisms produce more ethanol and butanediol and less acid. Unlike yeast and moulds, bacteria have, in general, a shorter generation time and higher fermentation rate. Erwinia chrysanthenn, has been found to utilize the sugars present in lignocellulosic substrate with a high ethanol tolerance capability [65].

3

KINETICS OF GROWTH AND PRODUCT FORMATION

3.1

Yeasts

Pachysolen tannophilus, Candida shehatae, C. tropicalis, and Pichia stipitis have been most studied pentose fermenting yeasts. In regard to the ethanol concentration and by-product formation, C. shehatae, P. tannophilus and P. stipitis are considered promising organism (Table 3). Woods and Millis [66] compared xylose fermentations of P. stipitis,

P. tannophilus and C. shehatae. The maximum ethanol concentration obtained were 28 g/I with P. tannophilus, 33 g/I with C. shehatae and 57 g/I with P. stiptis. The corresponding productivities were 0.11,0.26 and 0.18 g/I/h, respectively. P. stipitis was found to be less affected by higher substrate concentrations [67]. The ethanol yield from xylose with

P. tannophilus was reported to be increased by 32% when 5 g/I glucose was added during the fermentation [68]. However, the mechanism for increased yield could not be explained. P. stipitis and C. shehatae could ferment arabinose, rhamnose, galactose, mannose and xylitol in addition to xylose [69,70]. Table 4 shows fermentation balance of

P. stipitis and C. shehatae.

157 Table 3 Bioconversion of xylose into ethanol by yeasts Organism

Ethanol

Yield

Productivity

(g/I)

(g/g)

(g/I/h)

Candida sp. XF217

21.0

0.42

0.42

[32]

C. shehatae CSIR-Y492

26.0

0.29

0.65

[202]

C. blankii ATCC 18735

5.1

0.10

0.07

[11]

C. tenuis CSIR-Y566

13.3

0.25

0.42

[70]

C. tropicalis ATCC 1369

5.5

0.07

0.06

[14]

P. tannophilus NRRL-Y2460

33.4

0.21

0.12

[80]

Pichia stipitis Y7124

39.0

0.39

0.28

[67]

5.6

0.28

30.0

0.31

0.42

[8~]

5.0

0.10

0.07

[11]

Kluyveromyces marxianus K. cellobiovorus KY5199 Scizosaccharomyces pombe

Reference

[203]

Table 4 Xylose fermentation balance of pentose fermenting yeasts Product

Pichia stipitis

Candida shehatae

(mM/mM carbon) Ethanol

22.2

96

CO 2

11.2

48

Glycerol

0.3

Acetic acid

0.6

Xylitol

543

Cell mass

22.9

406

% Carbon recovered

60

117

158 In a fed-batch culture, where the sugar concentration was kept between 5 and 8 g/I, 26 g/I ethanol was obtained with P. tannophilus. The yield was increased by 41% compared with batch fermentation [71]. In continuous culture with P. tannophilus, a productivity of 2.2 g/I/h was obtained at a dilution rate of 0.12 per h [72]. A substrate concentration of 50 g/I at dilution rate of 0.34 per h resulted in a productivity of 2.1 g/I/h [19]. Generally, decrease in aeration has been found to increase the ethanol yield in yeasts. But this results in lower cell growth and risk of a wash-out. This problem may be overcome by recirculating or immobilizing the cells [73]. Cell recycling improves the fermentation productivity by virtue of increased cell concentration [74]. High cell density fermentation via continuous cell recycle may be well suited for slow fermentations such as xylose to ethanol. A bioreactor equipped with cell recycle device has been used to investigate high-cell continuous fermentation of xylose by P. tannophilus [75]. To examine the fermentation using non-growing cells (cells under nitrogen-deficient medium), continuous runs were carried out at 80 ml/min oxygen supply rate. The ethanol yield reached a maximum of 0.35 g/g xylose consumed at the dilution rate of 0.114 h1. Without nitrogen source there was little possibility for the biosynthetic process to occur. The substrate was therefore channelised almost exclusively into product formation. Ethanol productivity at the aforementioned dilution rate was 0.42 g/I/h. However, the specific productivity in non-growing cell fermentation was substantially lower than that obtained with growing cells [76]. During nitrogen starvation, cell proteins are subject to significant degradation [77]. It is concievable that enzymes involved in ethanol formation and respiration are also subject to protein degradation in non-growing cells, resulting in low specific productivity. At optimum oxygen supply rate (3.85 ml O2/g cells/min), the maximum volumetric productivity obtained was 2.43 g ethanol/I/h. The cell recycle system featuring a microprocessor controlled backflushing was also proven effective in alleviating membrane fouling and allowing long term operation of high-cell continuous fermentation. Gong et al. [11] studied the utilization and conversion of different pentose sugars by various yeast species. The majority of yeasts tested utilized xylose and produced ethanol, polyol and organic acids. The type and amount of products formed varied with the yeast strains used. Xylulose was the preferred substrate followed by xylose, arabinose and xylitol. A majority of tested yeasts utilized xylulose aerobically as well as fermentatively. Yeasts capable of fermenting xylulose are shown in Table 5. Thermotolerant strains of Candida sp. and Hansenula polymorpha have been investigated for the conversion of glucose and xylose [78]. Most strains of H. polymorpha

159 are thermotolerant [79]. Thermotolerant strains of Candida sp. exhibited Tr,axaround 48~ and were capable of fermenting at this temperature. Maximum ethanol production was obtained with Candida sp. HT4 among all the thermotolerant yeats.

Table 5 Fermentation of D-xylulose by yeast Organism

Yield

Productivity

(g/g)

(g/I/h)

Saccharomyces cerevisiae

0.46

0.15

S. saki

0.20

0.14

S. uvarum

0.45

0.32

Schizosaccharomyces pombe

0.43

0.62

Candida tropicalis

0.41

0.28

C. steatolytica

0.22

0.16

C. mogii

0.23

0.16

Hansenula anamala

0.46

0.32

Klyuveromyces fragilis

0.41

0.28

Torulopsis hensenii

0.36

0.25

Brettanomyces claussenii

0.21

0.15

Dekkera intermedia

0.31

0.22

Compiled from [11,204,205]

A significant level of aeration was found to be necessary to stimulate biomass growth and to enhance the rate of ethanol production in P. tannophilus [80]. Ethanol production appeared to be restricted by substrate inhibition at initial xylose concentration in excess of 40 g/l. Maximum ethanol yield was only 53.7% of the theoretical maximum because of the significant (up to 14% mass yield) accumulation of the by-product xylitol.

16o Out of 213 yeast species screened by Morikawa et al. [81], 23 produced more than 0.1 g/I ethanol from 20 g/I xylose. Yeast species of genera Candida and Kluyveromyces were found to be the good ethanol producers. K. cellobiovorus produced 27-30 g/I ethanol and 22-25 g/I xylitol from 100 g/I xylose in 66 h. In a mixture of glucose, xylose and cellobiose, xylose was consumed immediately after the total exhaution of glucose and most of the cellobiose was consumed after disappearence of xylose. Simultaneous utilization of glucose and xylose has been demonstrated using

Candida shehatae [82]. Cells grown aerobically on glucose and then used for anaerobic mixed sugar fermentation exhibited a sequential utilization pattern. Once glucose had been consumed, xylose utilization began. On the other hand, cells grown aerobically on xylose and then used for fermentation showed simultaneous sugar utilization. The rate of glucose consumption was higher than that of xylose. Glucose appears to inhibit xylose catabolism by repressing the induction of enzymes as has been observed in P.

tannophilus [83]. Pichia stipitis exhibits a stable ethanol yield of about 0.44 g/g when initial substrate concentration is between 20 g/I and 110 g/I [84], whereas Candida shehatae produces a stable ethanol yield of 0.4 g/g only when the initial substrate concentration does not exceed 50 g/l. The xylose conversion has been found to be inhibited completely at glucose concentration of 2.3 g/! and higher [85] which is in contradiction with the results of Panchal et al. [86] where the xylose conversion was repressed only by glucose concentrations above 20 g/l. Whole cells and a cell extract of P. tannophilus converted xylose to ethanol, xylitol and CO 2 [87]. The whole cell system converted xylitol slowly to CO 2 and little ethanol was produced, whereas the cell-free system converted xylitol quantitatively to ethanol (1.64 mol ethanol per mol xylitol) and CO2. The continued conversion of xylose to xylitol in the presence of fluoroacetate, which inhibits aconitase, demonstrated that the TCA cycle was not the source of electrons for the production of xylitol from xylose. The source of electrons was indirectly identified as an oxidative pentose hexose cycle.

161

3.2

Filamentous fungi

Species of Fusarium, Mucor, Monilia, Neurospora and Paecilomyces are known to ferment pentose sugars (Table 6). Rosenberg et al. [29] compared the xylose fermentation by Fusarium oxysporum with Bacillus macerans in a pH controlled fermentor. F.

oxysporum produced 0.41 g/g ethanol from 10 g/I of xylose with 94% of the original xylose carbon accounted for. The lost carbon was believed to be mainly in the form of CO 2. The growth of Fusarium was not exponential and specific growth rate declined with increasing cell mass. The conversion rate was too slow to be considered commercially significant. Ethanol yields of 0.32 and 0.16 g/g xylose were obtained with F. lycopercici and

Mucor 101, respectively [40]. The rate of ethanol production was also lower with Mucor sp. at high xylose concentration (20%). Suihko and Enari [36] screened 26 different

Fusarium species for ethanol production from xylose. The best strain, F. oxysporum VTTD-80134 was able to produce 21 g/I ethanol from 50 g/I of xylose in 7 days. Further optimization of the fermentation parameters resulted in the ethanol concentration of 25 g/I in 6 days, corresponding to the theoretical yield. A mixture of glucose (25 g/I) and xylose (25g/I) was also tested by these workers. Glucose utilization was completed after 1 day, and thereafter the rate of fermentation corresponded to that of xylose fermentation, yielding 25 g/I ethanol after 5 days [28].

Neurospora crassa has also been found to ferment a wide range of substrates including cellulose, xylan, starch, xylose, cellulose, galactose and arabinose. From 20 g/I xylose it produced 6.8 g/I ethanol in 7 days [33]. Mannose was metabolized rapidly with 100% conversion in 2 days, while galactose and arabinose gave 75% and 44% conversion, respectively, in 6 days. Wene and Antonopoulos [37] selected a strain of F.

oxysporum ANL 22-760 out of 2000 Fusarium isolates. This strain showed consistent yields under semiaerobic and anaerobic conditions. Under semiaeroic conditions 8.2 g/I ethanol could be obtained from 20 g/I of xylose in 72 h. Using cell recycling system fermentation time could be reduced to 48 h with almost equal yields. F. oxysporum F3 was able to produce 5 g/I ethanol from 20 g/I xylose in 6 d with a yield of 48% of theoretical [30]. F. oxysporum DSM 841 was able to produce almost equal concentrations of ethanol and acetic acid from D-xylose and D-glucose [88,89]. A cellulolytic strain of

Monilia sp. was found to convert 40% of the supplied xylose to ethanol in 7 days [42].

162 Table 6 Bioconversion of different sugars by filamentous fungi Substrate

Ethanol yield (g/g)

Glucose

0.24

[40]

Xylose

0.28

Glucose

0.18

Xylose

0.32

[4o] [4o] [4o]

Glucose

0.46

[38]

Xylose

0.50

[36]

Ribose

0.06

Galactose

0.42

Mannose

0.44

F3

Xylose

0.25

[38] [38] [38] [3o]

DSM 841

Xylose

0.20

[47]

ANL 22-760

Xylose

0.44

[37]

ATCC 10960

Xylose

0.35

[29]

F. poae

Xylose

0.26

[36]

F. clamydosporium

Xylose

0.22

[36]

F. sambucium

Xylose

0.26

[36]

Mucor sp. 101

Xylose

0.18

[4o]

Monilia sp.

Glucose

0.44

[32]

Xylose

0.20

[32]

Neurospora crassa

Xylose

0.34

[33]

Paecilomyces sp.

Xylose

0.37

[34]

Arabinose

0.28

[34]

Ribose

0.20

[34]

Galactose

0.40

[34]

Mannose

0.38

[34]

Organism/ Strain

Reference

Fusarium oxysporum

F5 ATCC 16417 VTT-D-80134

163

Paecilomyces sp. NF1 is capable of metabolizing a wide range of sugars including glucose, xylose, galactose, mannose, arabinose, ribose, maltose, cellobiose, lactose, and soluble starch [34]. Its ability to produce ethanol at a level of 73 g/I from 200 g/I xylose is the unique characteristic of this fungus. Only traces of by-products are detected at the end of fermentation. Although filamentous fungi convert xylose with high ethanol yields, the fermentation rate is slow compared with yeast and bacteria. It has been suggested that the slow rates of ethanol production with fungi may be overcome using novel mycelial columns [90]. Table 8 shows the performance of F. oxysporum in different reactor systems.

Table 7 Xylose fermentation by Fusarium oxysporum in different reactor types Reactor

Ethanol

Yield

Productivity

type

(g/I)

(g/g)

(g/I/h)

Batch

25

0.50

0.17

[28]

Continuous

4

0.41

0.04

[29]

Cell recycling

8

0.40

0.17

[37]

Immobilized cells

14

0.40

0.15

[108]

Immobilized continuous

10

0.13

0.10

[108]

Reference

3.3 Thermophilic bacteria

Thermophilic anaerobes have been shown to ferment various substrates including cellulose, hemicellulose, starch and pectin. The fastest growing thermoanaerobe yet isolated is Thermobacteroides acetoethylicus which has a doubling time of 25 min on glucose (~max=1.66). All the thermoanaerobes used for ethanol production have the same

164 end product spectrum. Besides ethanol they also produce acetic and lactic acids, CO2 and hydrogen. Thermoanaerobacter ethanolicus produces ethanol at 70~ which favours a process for simultaneous fermentation and evaporation. A mutant with high sugar tolerance has been reported [91]. Bacteroides polypragmatus can ferment xylose, arabinose and ribose to ethanol, but it also produces acetate, butyrate and H2 and CO2 gases [92]. A co-culture of Zymomonas anaerobia and Clostridium saccharolyticum was used by Asther and Khan [93]. It was observed that acetate produced from glucose by

C. saccharolyticum further inhibits the fermentation of xylose. Co-cultures of Clostridium thermocellum with Clostridium thermohydrosulfuricum were found to be effective in fermenting cellulose and hemicellulose [50]. Similarly co-culture of C. thermocellum and

T. ethanolicus on cellulose and hemicellulose demonstrated an increased rate of degradation and higher yields of ethanol by co-culture [52]. The role of the noncellulolytic bacterium in these systems includes hexose as well as pentose utilization. Table 8 exhibits potential bacterial strains for pentose fermentation.

Table 8 Xylose fermentation by different bacteria Organism

Ethanol yield (g/g)

Reference

Thermoanaerobacter ethanolicus

0.48

[52]

Clostridium thermohydrosulfuricum Clostridium thermosaccharolyticum

0.43

[50]

0.35

[49]

Clostridium thermosulfurogenes

0.20

[93]

Clostridium saccharolyticum

0.07

[93]

Zymomonas anaerobia

0.29

[93]

Bacteroides polypragmatus

0.15

[92]

Bacillus macerans

0.26

[29]

Klebsiella pneumoniae

0.31

[102]

Bacillus stearothermophilus

0.31

[206]

165

Clostridium thermocellum is unable to utilize pentose sugars formed by the breakdown of hemicellulose. The second anaerobic thermophile in the coculture, Clostridium thermosaccharolyticum utilizes pentoses to produce ethanol, acetate and lactate. The later organism also utilizes cellobiose released from cellulase enzyme action faster than C. thermocellum [94,95]. The coculture process thus has great potential but obstacles to commercialization exist. They are mainly the slow growth rate, and the byproducts such as acetate and lactate which decreases the yield of ethanol and can act as weak uncouplers [96,97]. Fermentation thermoanaerobes are shown in Table 9.

balances

of

pentoses

fermenting

Table 9 Xylose fermentation balance of thermoanaerobes Product (mM/100 mM Carbon) Organism

Reference Ethanol

Clostridium

Lactic

Acetic

acid

acid

CO2

H2

142

15

128

13.3

18.1

69

37.0

36.0

[49]

149

10.0

10.0

[52]

67

6.5

60.3

[50]

thermohydrosulfuricum Clostridium thermosaccharolyticum Thermoanaerobacter ethanolicus Clostridium

59

[207]

thermosulfurogenes

Simultaneous utilization of hexose and pentose sugars have been investigated to a limited degree in batch culture. Slaff and Humphrey [98] reported diauxic utilization of

166 glucose in preference to xylose or cellobiose for Clostridium thermohydrosulfuricum, with xylose and cellobiose consumed simultaneously in the absence of glucose. Clostridium

thermocellum also utilizes cellobiose in preference to glucose [99]. Avgeringos and Wang [100] reported 85-90% utilization of both glucose and xylose at the end of batch conversion of solvent extracted corn stover.

Clostridium thermohydrosulfuricum Rt8.B1 exhibited a continuous growth rate without diauxic, on mixed carbohydrate substrate in a pH controlled batch culture [101]. Hyperbolic grawth was observed with xylose in combination with either glucose or cellobiose. Diauxic growth was observed when the organism was grown on glucose plus cellobiose mixture. The major end products under all substrate conditions were ethanol and acetate (Table 10). Ethanol production varied depending on the substrate supplied and was always greatest on mixtures that included xylose (hyperbolic growth).

Table 10 Fermentation balance and growth rates of Thermoanaerobacter thermohydrosulfuricus Rt8.B1 [101] Growth substrate Parameter Glucose

Growth rate (h1)

0.51

Xylose

0.24

Glucose/ Cellobiose xylose 0.56

0.10

Substrate consumed (mM)

52

41

88

31

Ethanol (mM)

57

45

137

66

Acetate (mM)

26

18

12

38

Lactate (mM)

2

2

Propionate (mM)

0

4

10

4

CO 2 (mM)

83

63

149

104

Carbon recovery (%)

82

101

99

87

0

167

3.4 Mesophilic bacteria

Rosenberg et al. [29] studied the fermentation of xylose to ethanol with Bacillus macerans ATCC 8244 and compared its xylose fermentation efficiency with Fusarium oxysporum ATCC 10960. They observed that specific growth rate (0.15/h) and specific

ethanol productivity (0.8 g/g/h) of B. macerans were much higher than that of F. oxysporum. Aeromonas hydrophila, Bacillus polymyxa, and Aerobacter indologenes were found to produce ethanol at the level of 48.9, 63.0 and 55.9 mM/100 mM xylose [40]. Klebsiella pneumoniae MB-16-1048 produced 12.7 g/I ethanol with corresponding yield of 0.31 g/g xylose in 20 h [102]. This mutant strain showed 82% improvement in the

ethanol yield as compared to original strain of K. pneumoniae isolated from soil. A comparative kinetics of xylose fermentation by fungi, bacteria and yeast, is given in Table 11.

Table 11 Comparative kinetics of xylose fermentation by yeast, filamentous fungi and bacteria Organism

Specific

Ethanol

Specific

growth rate a

yield b

ethanol

Reference

production c Pachysolen tannophilus

0.24

0.22

0.12

[110]

Kluyveromyces marxianus

0.12

0.16

0.10

[208]

Candida shehatae

0.02

0.08

0.28

[70]

Pichia stipitis

0.08

0.12

0.25

[70]

Fusarium oxysporum

0.24

0.41

0.08

[29]

Bacillus macerans

0.15

0.05

0.80

[29]

0.02

0.15

[102]

Klebsiella pneumoniae Clostridium thermohydrosulfuricum

0.45

0.07

[50]

~per hour; bgrams ethanol per gram xylose; Cgrams ethanol per gram dry wt cell per hour

168 With bacteria, it is possible to obtain product concentrations, yields, and productivities comparable to those in hexose fermentations with yeasts. However, byproduct formation presents a lot of problems in the downstream processing of the product. Reported ethanol yields are low from xylose fermentation with yeast. Another difficulty in xylose utilization by yeast is to achieve acomplete conversion of xylose to xylulose economically and proper control of oxygenation. On the other hand, filamentous fungi ferment xylose with high ethanol yields, but the fermentation rate is too slow compared with yeast and bacteria and thus still need improvement in the process.

4

SIMULTANEOUS PENTOSE ISOMERIZATION AND FERMENTATION

Some organisms can not utilize D-xylose directly, instead they can utilize xylulose (produced by using D-xylose isomerase/D-glucose isomerase), an isomer of D-xylose. The ability of some yeasts and fungi to utilize D-xylulose for growth and ethanol production led to the development of two methods for producing ethanol from D-xylose by using D-xylose isomerase. One method depends on the isomerase to first isomerize D-xylose to D-xylulose which is then supplied to the cells. The other method employs recombinant yeasts that bear a bacterial gene for D-xylose isomerase. The ability to grow on D-xylose and D-xylulose may not necessarily be linked. Organisms that fail to grow on D-xylose grow readily on D-xylulose. Among the xylulose fermenting yeasts,

Saccharomyces cerevisiae and Schizosaccharomycespombe and Candida tropicalis have been most studied. These yeasts can efficiently ferment xylose in the presence of the bacterial xylose (glucose)isomerase. Wang et al. [103] compared different enzyme preparations and found Maxazyme GI the best for fermentation of xylose with Schizosaccharomyces pombe. The xylose isomerase reaction is an equilibrium reaction and at most, 20% of the xylose is converted to xylulose [19]. However, using repeated isomerization and fermentation, a transformation of 85% of xylose to ethanol could be achieved with S. pombe [104]. With Baker's yeast, 62 g/I ethanol could be achieved at 30~ using simultaneous isomerization and fermentation process [105]. The corresponding yield and productivities were found to be 0.34 g/g and 1.25 g/I/h, respectively. Linden and Hahn-Hagerdal [106] tested five yeasts,

C. tropicalis, P. stiptis, P. tannophilus, S. pombe and S. cerevisiae for the fermentation of spent sulfite liquor in the presence of commercial xylose (glucose) isomerase. The

169 maximum yield of 0.41 g/g was obtained with S. cerevisiae. When S. cerevisiae, xylose isomerase and 4.6 mM sodium azide were used for fermentation of hydrogen fluoride pretreated and acid hydrolyzed wheat straw, ethanol yield of 0.40 g/g could be obtained. Xylose utilization was found to be 84% compared to 51% in spent sulfite liquor. An important consideration in setting up the simultaneous isomerization and fermentation system is the difference in optimum operating conditions for the enzyme and cells. The optimum pH for the yeasts is below 7 and the temperature below 40~ whereas the optimum pH for isomerization is 8 and the optimum temperature is 70-80~ [107]. Thus, a compromise in the conditions must be sorted out when enzyme and cells are kept in the same reactor. Table 12 shows production of ethanol by simultaneous isomerization and fermentation process.

Table 12 Simultaneous xylose isomerization and fermentation Organism

Enzyme

Ethanol yield

Reference

source

(g/g)

Novo Sweetzyme

0.17

[104]

S. cerevisae

Takasweet

0.34

[42]

S. cerevisiae

MKC Optisweet

0.50

[103]

Schizosaccharomyces

Brocades Maxazyme

0.05

[105]

Candida tropicalis

MKC Optisweet

0.21

[106]

Pichia stipitis

MKC Optisweet

0.17

[106]

Pachysolen

MKC Optisweet

0.12

[106]

MKC Optisweet

0.31

[106]

Saccharomyces cerevisiae

pombe

tannophilus S. cerevisae + P. tannophilus

170

Using an adapted strain of S. pombe and immobilized enzyme, 45 g/I ethanol was obtained in 25-30 h from sugarcane bagasse hydrolysate with an initial sugar concentration of 105 g/I (approximately 62 g/I xylose, 28 g/I glucose, and 24 g/I arabinose), whereas using a strain of S. pombe in a pretreated bagasse hydrolysate containing approximately 180 g/I total sugars, 55 g/I ethanol was obtained [108]. Although simultaneous saccharification and fermentation can work, several questions still require consideration before its utility at the industrial level can be evaluated.Since higher amount of ethanol is produced from xylulose in oxygen-limited conditions, low or zero aeration would be optimal. However, some yeasts do not grow in the absence of oxygen raising the question of viability. Another matter of significance is the enzyme stability and cost. The possibility of inherent instability, as well as of susceptibility to proteolytic enzymes produced by the yeast is an issue in the study with soluble isomerases. The immobilized enzymes may be more stable in this respect.

5

WHOLE CELL IMMOBILIZATION

The immobilization of whole cells provides a means for the entrapment of multistep and cooperative enzyme system present in the intact cell, repetitive use and improved stability. This technique is also advantageous in separation of bioproducts from cell mass in a continuous bioconversion process. Other advantages of immobilized growing cells include: Protection of cells against unfavourable environmental factors. Changes in the permeability of the cells. Reduced inhibition by substrate and product. Reusability. Faster and easier removal of end products. Although a decrease in aeration increases the ethanol production from pentoses, it leads to lower cell growth and the risk of washout. This problem could be overcome by recirculating or immobilizing the cells [73]. Although Pachysolen tannophilus generally produces ethanol in higher concentrations and under wider range of conditions than does

171

Candida tropicalis [20], the specific rates of ethanol production by immobilized cells of the two organisms are similar in shake flasks (0.22 and 0.20 g/g dry wt cells/day, respectively). Immobilization of P. tannophilus in calcium alginate leads to a high ethanol yield (0.45 g/g) from xylose [109]. However, obtaining this yield required that cells be run continuously in a column for about 40 days using a feed solution containing 10 g/I xylose. In another study with P. tannophflus, entrapped in calcium alginate, in which the effect of several operating variables on productivity was investigated, the maximum ethanol yield was 0.35 g/g [110]. In a comparison of immobilized growing and non-growing P.

tannophilus cells, ethanol yields of 0.351 g/g and 0.308 g/g, respectively were obtained [111]. With immobilized Pichia stipitis cells, improvements in rates and yields over batch cultures were obtained [112]. Under batch conditions 22 g/I ethanol with average volumetric productivity of 0.22 g/I/h was obtained from 50 g/I xylose. However, with cells attached to a nylon net in a continuous packed-bed column reactor and 35 h residence time, a maximum volumetric productivity of 1.0 g/I/h could be achieved. In an attempt to simultaneously ferment glucose and xylose in a single reactor without the addition of oxygen, P. stipitis was immobilized in alginate beads where a low hexose concentration is expected in the core of beads, thus allowing efficient xylose conversion [113]. A continuous culture with immobilized cells of P. stipitis and suspended cells of S.

cerevisiae was studied in order to obtain a low bulk glucose concentration thereby promoting the conversion of xylose in a mixture of sugars [114]. Variuos processes for the simultaneous conversion of a mixture xylose and glucose were modeled and compared with experimental results for a better understanding of the phenomenon occurring. Simulations showed that the glucose profile in the beads were quite different for various processes. A higher xylose concentration with coimmobilization was a result of very low glucose concentration in the beads as compared to separately immobilized

P. stipitis or S. cerevisiae. Abbi [115] investigated batch and continuous fermentation of xylose and hemicellulose hydrolysate using free and immobilized cells of C. shehatae NCL-3501. A batch fermentation of 10 g/I xylose gave an ethanol yield of 0.5 g/g in 36 h as compared to 0.48 g/g by free cells. In a cell recycling system, free cells could be recycled three times to give same yield, while immobilized cells could be recycled four times. When a fed-batch fermentation of xylose with both free and immobilized cells was investigated, maximum ethanol of 26.2 g/I and 28 g/I, respectively were obtained within 72 h. C. shehatae

172 fermented acid- and auto-hydrolysate effectively with ethanol yields of 0.45 g/g and 0.5 g/g using free and immobilized cells, respectively. In a continuous immobilized cell culture using acid hydrolysate as substrate, maximum ethanol production was observed after 24 h and maintained at the rate of 8 g/I upto 9 days. Table 13 shows ethanol production by yeast in different reactor systems.

Table 13 Xylose fermentation by yeasts in different reactor systems Reactor

Organism

type

Productivity

Reference

(g/I/h)

Continuous

P. tannophilus

2.2

[72]

Continuous

P. stipitis

0.65

[85]

Fed-batch

C. shehatae

0.37

[115]

Co-culture

P. stipitis +

0.51

[117]

4.3

[118]

S. cerevisiae

Co-culture

P. stipitis + S. diastaticus

Immobilized alginate

P. stipitis

0.58

[73]

Immobilized nylon net

P. stipitis

0.54

[73]

Immobilized continuous

P. stipitis

1.0

[112]

Immobilized continuous

C. shehatae

0.33

[115]

Co-immobilized

P. stipitis +

0.84

[114]

S. cerevisiae

173 Immobilized fungal cells for xylose fermentation have not not been studied in detail. Immobilized cells of two mould cultures Mucor sp. and F. lini were compared with S.

cerevisiae for their ethanol producing ability. In shaken flasks, both fungal cultures produced ethanol at much lower rates compared to S. cerevisiae which had the best ethanol production rate of 0.3 g/I/h.

6

COCULTURE

In a coculture process, glucose and xylose are simultaneously converted to ethanol only if some conditions are fulfilled [116]. Firstly, the development of a coculture process implies that the associated yeast strains can be grown together. Furthermore, it must be performed under continuous culture conditions using a respiratory-deficient mutant of Saccharomyces to ferment glucose, in combination with a xylose fermenting yeast. Glucose concentration should be sufficiently low not to repress xylose utilization and the respiratory-deficient mutant generates oxygen profile favourable to the xylose fermenting yeast. Grootjen et al. [117] performed coculture studies in a series of reactors. The first system had three reactors with Pichia stipitis as the sole yeast, whereas the second system consisted of two reactors: Saccharomyces cerevisiae was inoculated in the first reactor to convert glucose rapidly, and the second reactor contained P. stipitis to convert pentose sugars. Almost complete utilization of glucose/xylose mixture was possible in this system. However, low dilution rates were necessary to obtain sufficiently low glucose concentration for the conversion of xylose, consequently the volumetric productivities were low (0.51 g/I/h). When a coculture of P. stipitis and a respiratory deficient-mutant of Saccharomyces

diastaticus was used to convert a mixture of glucose and xylose (glucose 70%, xylose 30%) in continuous culture, xylose was entirely consumed. However, with high substrate concentration (80 g/I), a considerable amount of substrate (20.5 g/I) was left unutilized [118,119]. At low dilution rate or low substrate concentration, P. stipitis was the dominating species in the reactor.

174 7

PERFORMANCE ON NATURAL SUBSTRATES

The ultimate goal of studying fermentation of xylose, xylulose or other pentose sugars is to be able to utilize the pentose fraction available in lignocellulosic hydrolysate. Since the insoluble raw material is inaccessible to most of the fermenting microorganisms, it has to be pre-treated and hydrolyzed. Hemicellulose can be hydrolyzed to simple sugars by using enzymes, or chemical/physical method. Some of the important results on the utilization of hemicellulose hydrolysate are summarized in Table 14.

Table 14 Performance of pentose fermenting organisms in lignocellulosic hydrolysate Organism

Substrate Treatment

Sugars

Ethanol

(g/I)

yield (g/g)

Reference

Candida sp.

Corn cob Acid

110

0.38

[32]

C. tropicalis

Aspen

Acid

80

0.15

[187]

C. shehatae

Red oak Acid

43

0.25

[168]

P. tannophilus

Aspen

Acid,

17

0.39

[201 ]

P. stipitis

Aspen

Steam, SO2

92

0.45

[202]

S. cerevisiae

Salix

Enzyme

69

0.45

[133]

S. cidri

Salix

Enzyme

69

0.47

[133]

Mucor sp.

Bagasse Acid

56

0.33

[40]

F. oxysporum

Wood

Acid

23

0.24

[125]

Z. mobilis

Salix

Enzyme

69

0.46

[133]

Z. mobilis +

Aspen

Acid

49

0.47

[209]

C. saccharolyticum

175

Several studies reported product concentrations more than 30 g/I, yields higher than 0.4 g/g and productivities higher than 0.5 g/I/h. The highest product concentration, 84 g/I with the yield of 0.47 g/g has been achieved with C. shehatae from whole barley using pre-treatment and enzymatic hydrolysis [120]. Using an adapted strain of Candida XF 217, 29 g/I of ethanol was obtained from 100 g/I of sugarcane hydrolyzate [121]. Although ethanol can be produced from D-xylose in significant yield, conversion rates are often slow. In some of the cases, fermentation times of 24 to 36 h have been reported, but most of these instances are for relatively low sugar concentrations. For higher sugar concentrations, fermentation periods are generally appreciably higher. For more rapid fermentations, high cell densities are often required [16]. For example 8.5 to 16 g/I dry weight of P. stipitis with hydrolysate prepared using a combination of steam, SO2 and enzyme [122] and 15 to 19 g/I of C. shehatae with spent sulfite liquor [123]. Intermittent feeding of cellulose hydrolysate to hemicellulose hydrolysate of hardwood resulted in improved ethanol yields, compared with fermentations of either hydrolysate alone or mixture of the two [124]. Rosenberg et al. [29] tested the performance of F. oxysporum ATCC 10960 in the acid hydrolysate of wheat straw. A total of 3.2 g/I of ethanol was produced from 11 g/I of the total sugars (2 g/I glucose + 9 g/I xylose) present, with a conversion efficiency of 29%. Ueng and Gong [40] employed species of Fusarium and Mucor for the conversion of sugarcane bagasse hemicellulose hydrolysate. The ethanol production rate with Mucor sp. was higher than with Fusarium. Fusarium F5 could ferment pure D-xylose readily but not the hydrolysate. Joshi et al. [125] attempted the utilization of sugars present in wood hydrolysate by F. oxysporum NCIM-1072 and F. oxysporum D-140. These strains were able to produce 11-13 g/I of ethanol from the hydrolysate containing about 55 g/I of the total sugars (glucose + xylose). More than 70% of Solka floc and 60% of Avicel were converted to ethanol in a single step by Monilia sp. [42]. Monilia sp. was also able to utilize hemicellulose and pectic materials. With Neurospora crassa, a conversion of 80-90% of alkali-treated cellulose powder and 60% conversion of holocellulose in steam-treated bagasse was possible in 10 days [43]. The conversion of xylan was 58% in 6 days, whereas alkalitreated bagasse was converted to ethanol in 4 days by this organism. Increase in bagasse concentration (5%) resulted in the increased ethanol yield corresponding to 75% conversion in 6 days. Table 15 shows some results on direct conversion of lignocellulosic materials to ethanol by different organisms.

176 Table 15 Direct conversion

of

lignocellulosic

materials

to

Organism

Substrate

F. oxysporum F3

Cellulose

0.48

[30]

F. oxysporum DSM841

Avicel

0.15

[46]

Potato waste

0.10

[47]

Solka floc

0.34

[32]

Avicel

0.28

[32]

Avicel

0.50

[33]

Solka fioc

0.35

[33]

Bagasse

0.22

[33]

C. thermocellum LQR1

Solka floc

0.14

[50]

Clostridium sp.

Cellulose

0.20

[121]

Xylan

0.25

[121]

T. ethanolicus

Aspen

0.15

[210]

Z.mobilis +

Aspen

0.47

[209]

Monilia sp. N. crassa NCIM 870

Ethanol yield (g/g)

ethanol

Reference

C. saccharolyticum

Fusarium oxysporum DSM 841 has potential to ferment cellulose and hemicellulose

in a single step process [47]. This organism produces cellulolytic and xylanolytic enzymes under aerobic condition and ethanol under anaerobic condition. The production of ethanol from potato wastes (cellulosic waste from potato starch industries) by F. oxysporum was improved considerably in fed-batch culture. McCracken and Gong [78] investigated the ability of a thermotolerant strain of Candida sp. HT4 to utilize sugarcane bagasse hydrolysate and cellulose to produce ethanol. Fermentation rate was slow in bagasse hydrolysate probably due to the presence of inhibitors. The rate of ethanol production on cellulose substrate increased from 0.93

g/I/h at 30~ to 1.58 g/I/h at 40~

On the other hand, Kluyveromyces cellobiovorus KY

177

5199 has been shown to produce ethanol from the enzymatic bagasse hydrolysate more efficiently than S. cerevisiae [81]. The maximum ethanol production from a total of 91 g/I sugars by K. cellobiovorus was 29 g/I in 72 h and by S. cerevisiae was 22 g/l.

Clostridium thermocellum produces very active cellulase and xylanase system and is capable of fermenting cellulose to ethanol in a single step. It produces near 1:1 ratio of ethanol and acetic acid from both Avicel and pretreated wood [126]. Ethanol:acetic acid ratios of 0.61-1.87 have also been reported for C. thermocellum fermenting lignocellulosic substrates [127,128]. The extent of substrate utilization has been found to be 86% for Avicel and 75% for pretreated wood. In a combined hydrolysis and fermentation approach, steam pretreated wheat and barley straws could yield over 0.4 g/g substrate, when cellulase preparation from

Trichoderma harzianum E58 and S. cerevisiae NRCC 202001 were used [129]. With cellulase preparations from T. reesei lower ethanol yields were obtained which were attributed to the lower amounts of 13-glucosidase detected in these preparation. Fermentation of ethanol extracted liquor xylan solids by P. tannophilus yielded 10 g/I ethanol from 43 g/I substrate, whereas ether extracted substrate resulted in 4.3 g/I ethanol from 48 g/I substrate [130]. Xylose liberated by acid hydrolysis from the hemicellulose-rich fraction of alkali pulped wheat straw produced 8.2 g/I ethanol from 47 g/I xylose in 96 h. Ethanol yields of 76-84% of theoretical value were obtained using a wild-type strain of P. tannophilus from a sugar mixture simulating spent sulfite liqour [131]. Using a mutant strain selected for more rapid growth on galactose, yields were increased to 8390%. Productivity was found to decrease on recycling of cells, suggesting that strain improvement would be a necessary step in process development. In fermentation of eucalyptus hemicellulose hydrolysate usibg P. stipitis NRRL Y7124, an initial lag phase characterized by flocculation and viability loss were observed [132]. Subsequently cell regrowth occurred with sequential consumption of sugars and production of ethanol and polyols. The fermentation was more effective at an oxygen transfer rate between 1.2 and 2.4 mM/I/h and an initial pH of 6.5. The best values obtained were, maximum ethanol concentration 12.6 g/I, sugar consumption 99%, ethanol yield 0.35 g/g sugar consumed, and volumetric ethanol productivity 4 g/I/d. The sugar consumption rates and the product formation of yeast (S. cerevisiae) and bacteria (Lactobacillus brevis, Lactococcus lactis, Escherichia coli and Zymomonas

mobilis) were investigated in spent sulfite liquor and an enzymatic hydrolysate of steam

178 pretreated salix [133]. S. cerevisiae emerged as the best candidate owing to its rapid sugar consumption rate and efficient ethanol production. Fermentation of a xylose-rich, dilute-acid-treated corn cob hydrolysate was studied using E. coil ATCC 11303, recombinant E. coil B(pL01297 and KOll), P. stipitis (CBS 5773, 6054 and R), S. cerevisiae isolate 3 in combination with xylose isomerase, recombinant S. cerevisiae (TJ1, H550 and H477) and F. oxysporum VTT-D-80134 in an interlaboratory comparison [134]. The microorganisms were studied according to three options: (A) fermentation under consistent conditions, (B) fermentation under optimal conditions, and (C) fermentation under optimal conditions of the organisms with detoxified hydrolysate. After adaptation and dilution of hydrolysate, P. stipitis CBS 5773 gave a yield of 0.34 g/g. However, all the strains of P. stipitis were highly sensitive to the inhibitors present in the hydrolysate. E. coil B,KO11 was the most promising ethanol producer {0.24 g/g (A), 0.36 g/g (B), 0.54 g/g (C)}. However, a disadvantage of bacteria in general is the requirement for a fermentation pH of 6-7, which increases the contamination risk. Under option A, both recombinant S. cerevisiae strains gave a yield of 0.07 g/g. The maximum volumetric productivity was 0.05 g/I/h for S. cerevisiae TJ1 and 0.76 g/I/h for S. cerevisiae H550.

8

FACTORS AFFECTING ETHANOL PRODUCTION

8.1

pH

If pH is uncontrolled, it changes with the extent of fermentation. Thus to assess its effect on ethanol production, the pH must be controlled at a set value over the entire fermentation period. The optimum choice of initial pH may also depend on the type of medium being fermented, type of pH control employed and the microbial strain carrying out the process. The optimum pH for yeast Pachysolen tannophilus has been found between 2.5-5.0 [135,136], for Candida shehatae between 3.5-4.5 [137], and for Pichia stipitis between 4.0-5.5 [138]. The initial pH values generally employed by various workers for fungal fermentation are within the range of 5.0-6.0 [29,36,43,46,139]. Kluyveromyces

179 cellobiovorus preferentially grow below pH 5 [81]. With C. shehatae, maximum biomass and ethanol productivity was obtained at pH 5.0, while maximum amount of polyol detected between pH 4 and 6 [115].

The optimum pH for Fusarium oxysporum lies between 5.0-5.5 and between 5.0-6.0 for Neurospora crassa. However, Paecilomyces sp. NF1 was found to have a wide range of pH optima which varied from 2.2-7.0 [34]. Ethanol production by F. oxysporum F3 is considerably affected by the pH of the aerated and nonaerated cultures. The optimum pH (6.0) for the nonaerated growth was also optimal for 13-glucosidase activity, the key enzyme in direct conversion of lignocellulosic materials to ethanol by F. oxysporum F3. Bacterial fermentation products depend upon the pH at which the fermentation is conducted. Acidic conditions favour production of neutral products at the expense of acids. The optimum pH for growth of B. macerans has been reported between 6.0-8.0 [43]. The pH range for the growth of thermophilic bacteria are those of typically neutrophilic organisms, most having optima between 6.0-7.0 and a range between 4.5-8.0. The optimum pH for recombinant strains of E. coil B (pLO297) and (KO11) were reported to be 7 and 6, respectively [134].

8.2 Temperature

Temperature has a profound effect on all the aspects of growth, metabolism, viability, and fermentability of microorganism [78]. As in the case of many other organisms, every species of yeasts has a relatively limited temperature range for growth. On the basis of temperature for growth, yeasts can be differentiated into several groups [140]. (1)

Obligate psychrophiles such as Candida frigida P8, Candida gelida P16 and Candida nivalis have a growth temperature minimum (Tmin) of -5~

(Topt) of 15~ (2)

and a maximum (mmax)of 20~

Facultative psychrophiles such as Rhodotorula spp. have a Tm~n of 0-5~ 20~

(3)

and optimum Topt of 14-

and Tmaxof 26-28~

Mesophiles such as Saccharomyces cerevisiae have a Tm~n of 5-10~

Topt of 24-

180

30~ (4)

and Tmax of 35-40~

Thermotolerant yeasts such as Hansenula polymorpha have a Tmi n of 20-26~ of 26-35~

(5)

Top t

and Trnaxof 37-45~

No true thermophilic yeasts have been described. However, certain yeasts such as

Candida slooffii and Sacharomycopsis guttulata have

a Tmi n

of 27 and 35~

respectively [141]. It has been noticed that effect of temperature depends on the microbial strain employed for fermentation [31]. The optimum temperature for most pentose fermenting yeasts is between 30-32~

[135]. However, Pichia stiptis [142] and Pachysolen

tannophilus [143,144] can grow at 37~ and 37-42~

respectively. In a simultaneous isomerization and fermentation scheme, the rate limiting isomerization step has a temperature optima of 50-70~ [145]. The use of high temperature tolerant yeasts could increase the fermentation rate thereby decreasing the fermentation time. Other benefits of using thermotolerant yeast is that cooling problems can be simplified during large scale fermentations.

Paecilomyces sp. NF1 has an optimum temperature range of 30-37~ [34], whereas Neurospora crassa has optimum range of 28-37~ [33]. Optimum temperature for most of the Fusarium strains is 30~ [31]. In mesophilic pentose fermenting bacteria, the optimum temperature range of 20-32~ for Klebseilla pneumoniae and 25~ for Citrobacter freundii were observed [102]. The optimum temperature for growth of thermophilic bacteria ranges from 60-70~ [146].

8.3

Nutrition

Besides physical factors, several nutritional factors also affect ethanol fermentation to a great extent. In Pachysolen tannophilus, growth requires a source of thiamine and biotin which can be made available by supplying yeast extract to the growth media [147,148]. Yeast extract has also been found to improve cell growth and ethanol production of Pichia stiptis and Candida tropicalis [19]. C. tropicalis requires 10-20 g/I yeast extract and peptone for ethanol production. Nitrogen and carbon sources are known

181 to influence the activity of pentose phosphate pathway enzymes and consequently the cell and ethanol yield obtained [149. However, thorough nutritional studies have not been done in yeast fermenting pentose sugars. The effects of Mg 2+ and Ca 2+ ion concentrations on P. stipitis NRRL Y-7124 growth and flocculation were evaluated by a 2X2 factorial design [150]. Negative interaction was detected between these two ions. Maximum growth rate was attained with high Mg 2+ (3 mM) and low Ca 2+ (0.34 mM) concentration levels, whereas the inverse ionic ratio was required to reach the maximum flocculation. Mahler and Guebel [151] studied the effect of Mg 2+ on P. stipitis growth and ethanol production under condition of constant oxygen uptake rate. Biomass/xylose and biomass/Mg 2+ yields increased with maximum value at 4 mM Mg 2+concentration, ethanol being the major product obtained. When P. stipitis CBS 6054 was grown in a media containing corn steep liquor as the sole source of nitrogen, amino acids, vitamins and other nutrients, the ethanol yield and fermentation rate compared favourably to those obtained with media containing more expensive sources of nutrients [152]. A variety of inorganic and organic nitrogen sources were evaluated by various workers in ethanol fermentation studies with filamentous fungi. Peptone was found to be the best organic nitrogen source. In Fusarium oxysporum, nitrogen limitation led to the death of the cells [28]. Trace elements such as Fe, Zn, Cu and Mn were found to increase the growth of Polyporus anceps in submerged culture. In spite of the pronounced effect of minerals, trace elements and growth factors on the biosynthesis of metabolites, negligible information is available with the fungal production of ethanol. Thiamine has been found essential for the growth of P. anceps in submerged culture [153]. As the concentration of thiamine increases in the media, the quantity of mycelium increases as did the substrate utilization and ethanol production. Ammonium sulphate concentration of 1 g/I was found to be the best nitrogen source for Klebsiella pneumoniae [102]. This preference may be due to the fact that this compound also provided sulfate ions which are important for the synthesis of methionine, cysteine, coenzyme lipoic acid and CoA [154,155]. Trace metal salts such as EDTA, TPP, CuSO4, CoCI 2, CaCI 2 and MnSO4 do not support ethanol formation from D-xylose by K.

pneumoniae [102]. Most of the thermophilic bacteria can be grown on defined media, but growth rates are reduced over that in complex media. Thermoanaerobacte ethanolicus requires yeast extract which could not be replaced by tryptone, casein hydrolysate, beef extract or ashed

182

yeast extract [55,56]. The requirement of thermophilic clostridia vary, Clostridium

thermocellum grow on salt medium supplemented with biotin, vitamin B12, pyridoxalamine and p-aminobenzoic acid [156]. Clostridium thermohydrosulfuricum requires yeast extract

[55]. An ethanol hyperproducing strain of C. thermocellum 1-1-B, isolated from Shibi hot spring, Kogashima was evaluated for its nutritional requirements [157]. Addition of a mixture of vitamins, yeast extract or vitamin B12 alone to the cellulose (80 g/I) medium enhanced the production of ethanol and decreased lactate formation. The organism produced 3.6 g/I ethanol, 8.5 g/I lactate, 2.9 g/I acetate and 0.9 g/I formate. Ethanol productivity was found superior to any of the wild and mutant strains of C. thermocellum reported so far.

8.4 Oxygenation

The rate of utilization of available carbohydrates and eventual conversion to ethanol is significantly affected by aeration in yeasts and filamentous fungi (Table 16). Limitation of oxygen can be associated with ethanol accumulation. With respect to oxygenation

Pachysolen tannophilus is the most thoroughly studied organism [147]. It produces cell mass under aerobic conditions, accumulates xylitol under anaerobic condition and produces ethanol under oxygen limitation [136,158]. However, there is one report where P. tannophilus was found to produce ethanol aerobically as well as anaerobically [110]. Another interesting observation with P. tannophilus is the reassimilation of ethanol when oxygenation was increased [13]. A similar observation was made with Candida tropicalis, when the xylose concentration reached a certain level, the organism preferred to utilize produced ethanol as carbon source [159]. du Preez et al. [137] compared the effect of oxygenation in P. tannophilus and Candida shehatae. With increasing oxygen limitation, the ethanol yield changed from 0.33 to 0.28 g/g with C. shehatae and from 0.24 to 0.02 with P. tannophilus, thus indicating that C. shehatae is less dependant on the degree of oxygenation. Similarly C. tropicalis was also marginally influenced by the degree of oxygenation [19].

183 Table 16 Effect of aeration condition on xylose fermentation Ethanol yield (g/g substrate) Organism

Reference Aerobic

Semiaerobic

Anaerobic

Candida shehatae

0

0.43

0.38

[70]

Candida tenuis

0

0.25

0.27

[70]

Kluyveromyces

0.27

0.30

0.10

0.28

0.26

[211 ]

0.04

0.23

0.11

[89]

0

0.43

0.38

[70]

[81 ]

cellobiovorus Pachysolen tannophilus Fusarium oxysporum Pichia stipitis

Under anaerobic conditions, absence of oxygen is the sole factor that controls the rate and extent of the cell growth. In the absence of oxygen growth either fails to occur or is restricted. However, D-xylose is still metabolized by some strains under these conditions with ethanol or xylitol as product [160,161]. In continuous culture, as the extent of oxygen limitation increased, growth decreased and ethanol production increased in P.

tannophilus [162]. This suggests that the specific rate of oxygen utilization is an important factor in determining the amount of ethanol accumulated [163,164]. Partitioning of carbon from D-xylose between growth and product formation is also affected by oxygenation. Over an intermediate range of aeration rates, as rates increase, carbon is shifted generally to growth at the expense of ethanol accumulation [165,166]. Both volumetric and specific ethanol production rates are influenced by change in aeration rates. This is particularly true with P. tannophilus and C. shehatae [137]. In a number of batch cultures, ethanol accumulation is associated with a progressive increase with time, in the extent of oxygen limitation. Ethanol can be produced by P.

184

tannophilus in media that do not support growth [164]. Limitation of growth of C. shehatae in continuous culture by a factor other than oxygen results in the production of small amounts of ethanol, but more is produced when oxygen is limited [166]. High cell density cultures have been employed in a number of studies assuming that ethanol accumulation is associated with growth limitation. Such cultures were obtained either by inoculating with relatively high cell densities [167,168] or by repetitively recycling all of the cells obtained from a culture into a comparable volume of fresh medium [170,171]. Early onset of oxygen limitation would be expected in such cultures because the aeration conditions employed are those that commonly result in oxygen limitation, even when lower cell densities are employed. in spite of the complexities of aerobiosis, several generalizations can be made that are of potential significance: Aeration rates that result in the accumulation of high ethanol concentration result in either low values for volumetric rate of ethanol production, rate of xylose utilization, or rate of growth. The presence of oxygen enhances the rate of xylose metabolism but has no significance in xylulose utilization. A similar oxygen-related effect, the "Kluyer Effect", has been described for the utilization of oligosaccharides and D-galactose by yeast [172]. This oxygen effect could result from either a direct or indirect oxygen requirement for the entry of xylose or for the activity of initial catabolic enzymes. With strains that can produce appreciable amounts of ethanol both aerobically and anaerobically, anaerobic conditions result in either poor xylose utilization and lower rates, or lower yields and ethanol production. In some cases, an increase in aeration rate improves performance in that either the rate of ethanol production increases, the yield increases or the yield of xylitol decreases. In effect, a particular range of aeration rates may be optimal for ethanol production in some systems. In filamentous fungi, ethanol accumulates only under low aeration (oxygen limited) conditions [173,174]. A series of experiments have been conducted with F. oxysporum to study the effect of aeration on ethanol production [174]. Ethanol production appeared to be growth associated and aeration rates of 0.04-0.06 vvm were found suitable for product formation. Shake flask studies on the effect of aeration rate on D-xylose fermentation and metabolism in F. oxysporum have shown that oxygen limited conditions

185 stimulate the biosynthesis of key D-xylose catabolising enzymes, consequently product formation increases several-fold. Increase in aeration rate from 0.05 to 0.08 I/I/h increases the growth of F. oxysporum VTT-D-80134, but ethanol yield is unaffected [28]. The best results are obtained using a gas mixture containing 1% oxygen. Tween-80 and ergosterol can not satisfy the oxygen requirement and ethanol production appeares to be growth associated. In F. oxysporum ANL 22-760, aeration rates higher than 0.1 vvm increase cell mass, but reduce ethanol accumulation [37]. Unlike yeast and fungi, bacterial fermentation of pentose sugars for ethanol production can be operated without the need for critical aeration. The facultative anaerobe, K. pneumoniae did not produce any solvent under aerobic condition, however, ethanol production is greatly enhanced under anaerobic conditions [173]. Banerjee [102] also observed that aerobic environment caused deleterious effects on ethanol formation by K. pneumoniae and C. freundii, though oxygen was required for the initial growth. The profile of dissolved oxygen (DO) showed that if the media is not degassed to remove oxygen, K. pneumoniae utilized the total DO within the initial 5 h of fermentation. Removal of DO from the broth using reducing agents at initial stage led to complete inhibition of growth and ethanol production. Thus DO is an important factor for initiation of D-xylose fermentation. This indicates the requirement of oxygen for the synthesis of pentose phosphate enzymes [102]. In sequence with the drop in DO level, the redox potential of fermentation broth showed a steady state decrease upto -400 mv which coincided with the maximum concentration of ethanol produced by K. pneumoniae [102]. This indicated that D-xylose fermentation by K. pneumoniae occurred under completely anaerobic condition. Such anaerobicity has also been reported in case of obligate anaerobes which exhibit a redox potential of less than -200 mv [175]. In Clostridia, oxidation of pyruvate to acetyl CoA by a coenzyme, ferrodoxin, occurred under redox potential of -400 mv [176].

8.5 Lipids

The peak concentration of ethanol and yield in the cultures of Pachysolen

tannophilus [177] was increased by the addition of low concentration of lipids (34 mg/I ergosterol, 34 mg/I linoleic acid, and 5.2 g/I Tween 80). The lipids increased the ethanol

186 concentration from 8.5 g/I to 13.25 g/I with corresponding yield of 0.2 g/g to 0.32 g/g, on the basis of D-xylose consumed. The enhanced ethanol yield was suggested to be due to the increased ethanol tolerance by addition of lipids. Neirinck et al. [178], however could not observe any effect of lipid supplementation on anaerobic incorporation with either D-xylose or D-glucose.

8.6

Metabolic inhibitors

Addition of metabolic inhibitors to the media to suppress the by-product formation and enhanced ethanol yield in yeast and filamentous fungi have been reported by several workers. Addition of sodium azide to the media increased the ethanol yield in Candida

tropicalis and Pachysolen tannophilus while the xylitol formation decreased [179,180]. Of other inhibitors tested, CCCP and dinitrophenol (DNP) also gave similar results. Addition of sodium azide, DNP and polyethylene glycol (PEG) to the medium also shifts product formation from acetate to ethanol in Fusarium oxysporum DSM 841 without affecting the utilization of xylose [88]. This strain produces ethanol and acetic acid in almost equal concentrations. Addition of azide at 0.1 to 0.3 mM improved the ethanol yield by a factor of 2, whereas acetic acid formation was inhibited by 85-95%. Quite similar effects of DNP was observed. Since both azide and DNP act at the same site of the electron transport chain, similar metabolic effects are expected [181,182]. Sodium azide represses the terminal oxidation step of the respiratory pathway and not enough ATP may be synthesized resulting in less biomass [183]. Some evidences have been presented revealing that uncoupling agents induce a rapid decrease of cytoplasmic pH which activates adenylate cyclase [182]. The cAMP increases as a consequence, stimulating several protein phosphorylation reactions and finally key enzyme of glycolytic pathway are activated. PEG is known to increase the viscosity of the medium which in turn reduce the oxygen transfer rate. With increase in PEG concentration upto 9% increase in ethanol yields and decrease in acetic acid production by Fusarium oxysporum DSM 841 was observed [88]. In Candida tropicalis, increased output of ethanol was found to be a function of decreased water activity after addition of ethanol which in turn switched to maintenance metabolism at the expense of cell growth [184]. It has also been postulated

187 that PEG being surface active compound, having chemical structure quite similar to Tween-80, lowers the surface tension of the medium and possibly acts as extractant [184].

8.7 Inhibitors present in lignocellulosic hydrolysate

Several inhibitors may be present in the lignocellulosic hydrolysate prepared by using acid catalysis (Table 7) [21,185]. Some are derived from non-carbohydrate materials and others are formed by the breakdown of carbohydrates during hydrolysis, e.g. furfural. Heavy metal ions, such as Cr, Cu, Fe, Ni etc. may also be present, probably as a result of corrosion of metal parts of the apparatus used [166]. Acetic acid is formed commonly in wood hydrolysates. Concentration of about 5 g/I acetic acid can be inhibitory to P. tannophilus [186] and greater amounts can be present in some hydrolysates. For example, an aspen hydrolysate contains 26 g/I of acetic acid [187], while sugarcane bagasse hydrolysate contains 3.9 to 10.4 g/I of acetic acid [166].

Table 17 Possible inhibitory substances present in lignocellulosic hydrolysates Carbohydrates

Extractives

Acids

Lignin

Furans

Resins

Acetic

Vanillin

Cr

Furfural

Phenols

Formic

Catechol

Cu

Hydroxymethyl

-Fats and

Levulinic

p-Hydroxybenzoate Fe

furfural Levoglucosan Sugar acids

fatty acids Tannins

Methanol Guaicol

Metal ions

Ni

188 When steam-exploded wheat straw is not water extracted, it inhibited the fermentative organisms as well as enzymatic hydrolysis step [188]. The inhibitory material was suspected to be a furan derivative. The extent of inhibition of yeast growth caused by acetic acid depends on pH. At pH 4, Candida utilis does not utilize xylose until considerable amount of acetic acid present were consumed, while at pH 6, concurrent use of acetate and xylose takes place [189]. It has been observed by several investigators that Pichia stipitis performs poorly in untreated lignocellulosic hydrolysates such as spent sulfite liquor [190,191]. In particular, acetic acid is a strong inhibitor. Steam stripping to reduce the acetic acid concentration below 0.6 g/I or pH adjustment to 5.7 improved the ethanol yield [192].

Pachysolen tannophilus appears to be a more resistant organism for the fermentation of untreated hydrolysates [191]. This organism was originally isolated from tannaries, which is as hostile an environment as an untreated lignocellulosic hydrolysate [193]. In order to obviate or minimize effects of inhibitors, hydrolysates containing xylose are treated prior to fermentation. The treatment could be:

(I) Physical treatment: ion exchange [185,187], adsorption on activated charcoal [187], solvent extraction [130], evaporation [194], and steam stripping [192,195]. (2)

Chemical treatment: adjustment of pH with calcium carbonate, oxide or hydroxide [196-201], the addition of sodium sulfite [124], and overneutralization [197].

9

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Pachysolen

tannophilus. Proc 4th Int Symp Alcohols Fuels Technoloy, Ottawa, 1984; 220. 198 Wayman M, Parekh S, Chornet E, Overend RP. Biotechnol Lett 1986; 8: 749. 199 Wayman M, Seagrave C, Parekh SR. Proc Biochem 1987; 22: 55. 200 Gong C-S, Tsao GT. Conversion of D-xylose to ethanol by yeast. Royal Soc Can int Symp Ethanol from Biomass, Winnepeg, 1982; 525. 201 Deverell KF. Biotechnol Lett 1983; 5: 475. 202 du Preez JC. Biotechnol Lett 1983; 5: 357. 203 Margaritis A, Bajpai P. Appl Environ Microbiol 1982; 44: 1039. 204 Ueng PP, Hunter CA, Gong C-S, Tsao GT. Biotechnol Lett 1981; 3: 315. 205 Suihko M-L, Poutanen K. Biotechnol Lett 1984; 6: 189. 206 Hartley BS. Paper plus liquid fuels from wood and straw. Proc Symp Present and Future uses of Straw. Society for Chemical Industry, London, 1991; 1. 207 Schink B, Zeikus JG. J Gen Microbiol 1983; 129:1148. 208 Margaritis A, Bajpai P, Cannel E. Biotechnol Lett 1981; 3: 595. 209 Murray WD, Asther M. Biotechnol Lett 1985; 6: 323. 210 Wiegel J, Mothershed CP, Puls J. Appl Environ Microbiol 1985; 49: 556. 211 Ligthelm ME, Prior BA, du Preez JC. Appl Microbiol Biotechnol 1988; 28: 63.

7 Microbial Production of Acetone and Butanol

1

INTRODUCTION

Fuel production from biomass, in the form of agricultural and forest residues, has been particularly appealing as a renewable energy source. The production of chemical feedstocks from carbohydrates encompasses a wide range of microbes and processes [1]. Acetone-butanol fermentation is one of the most important fermentations carried out by the genus Clostridium. This process, once of great industrial significance, is now carried out on industrial scale only where special conditions permit it to rival feedstock produced from a petrochemical source. Pasteur was probably the first to discover butanol as a product of microbial action. Beijerinck, Duclaux, Schradinger and Prazmowski are some most eminent names in early microbiological work on the process. Much of the early interest focussed on the possible use of butanol for conversion to butadiene, that is polymerized to give synthetic rubber. However, with the onset of World War, the demand for acetone for use in cordite manufacture and as a general solvent, has led to the setting up of a number of large plants in U.K., Canada and United States, producing acetone and butanol by fermentation of starchy materials, usually grains. The process used were pioneered by Weizmann [2,3], that produced butanol and acetone in the ratio of 2:1. However, a major difficulty arose with regard to the storage and disposal of butanol, regarded as a waste product. After the War, these plants were gradually run down with the sharp fall in the demand of acetone. Some of the plants were reopened when a use for butanol as solvent in the manufacturing of nitrocellulose lacquers, important for automobile industry, was discovered. Acetone-butanol fermentation industry expanded further, again using the classical Weizmann process, with grain as raw material. Fermentation of molasses was developed in 1930s which rapidly supressed the grain process. In 1945, 66% of the United States butanol requirement was being met by fermentation. However, after this period, the fermentation route declined sharply for several reasons, including steep rises in the price of molasses and grains and severe competition from the petrochemical industry [4]. 197

198 Newly developing economic situation and skyrocketing prices of dwindling hydrocarbon feedstocks had prompted re-examination of the old processes which used to be the sole source of industrial solvents. The synthetic route to butanol became much more expensive following increase in the price of crude oil in 1973. This triggered a greatly revived interest in the potential of fermentation. Economic pressures have led to the necessity to recover all possible products of fermentation. Thus dried biomass (high riboflavin) was used as a feed additive for ruminants, and the fermentation gases hydrogen and carbon dioxide were recovered. Such integrated plants achieved a high degree of efficiency in operation. Similarly rising molasses prices led to the investigations into alternative, cheaper sources of fermentable carbohydrates. Wood and agricultural byproducts are the most abundant renewable resource materials. Hydrolysis of these raw materials, whether enzymatic, chemical or combined with physical means, results in a variety of oligosaccharides, and simple pentose and hexose sugars. These sugars form an excellent substrate for acetone-butanol fermentation.

2

MICROORGANISM USED FOR ACETONE-BUTANOL PRODUCTION

In 1912, Weizmann [5] reported to have dicovered a bacterial culture with outstanding capabilities for anaerobic conversion of grains such as corn into acetone and butanol. The fermentation could be carried out either aerobically, which he preferred, or anaerobically. Weizmann's method of isolation involving successive heat shocking is still the method of choice [4,6]. Normally a potato based medium is used for serial subculturing of isolates obtained from soils or muds, cereals, potatoes, or roots of leguminous plants, where the organism is in loose association with nitrogen fixers. In acetone-butanol fermentation, the bacterial strains mostly involved are different species of Clostridium (Table 1). The genus is divided into five groups containing altogether 61 reference strains. Solvent producing clostridia can be found in any of the five groups. Being chemo-organotrophs, they metabolize sugars, polyalcohols, amino acids, organic acids, purines, and other organic compounds under strict anaerobic conditions. They do not reduce sulfate and some species can fix nitrogen. While a number of clostridial strains are capable of utilizing pentose sugars as carbon and energy source, the rate and efficiency of pentose metabolism differ markedly when compared to that of hexose. A serious problem stems from clostridia being rather susceptible to a

199 phage attack. The clostridial phage strain susceptibilty, however, can be minimized by production strain rotation, coupled with efficient sterilization techniques, clean operations and careful stock culture maintenance. Anaerobic production of acetone-butanol was the first large scale microbial process in which the exclusion of other microbes from the culture vessel became a factor of major importance for the success of operation.

Table 1 Solventogenic clostridia capable of utilizing pentose sugars Organism

Xylose

Arabinose

C. b u t y l i c u m

+

+

C. a c e t o b u t y l i c u m

+

+

C. beijerinckii

+

+

C. p a s t e u r i a n u m

-

+

C. felsenium

+

+

C. p r o p y l b u t y r i c u m

+

+

C. b a r k e d

+

-

C. g l y c o l y c u m

+

C. t h e r m o s a c c h a r o l y t i c u m

+

+

C. s a c c h a r o p e r b u t y l a c e t o n i c u m

+

+

McCoy et al. [7] carried out a systematic study of the 'butyl alcohol organisms' involving 11 different industrial producer strains from separate sources. The life cycle of the organisms was described in detail, placing them as members of the low acid/high alcohol-producing butyric acid bacteria. Morphologically they are rods, motile by means of a peritrichous flagella, Gram-positive, bearing granulose (a storge carbohydrate and swelling to clostridial form at sporulation (Table 2). The dimensions of the organisms were stated to vary considerably with strain, culture conditions and age.

200 Table 2 Morphological features of Clostridium acetobutylicum during acetone-butanol fermentation

[7] Time after

Cell Morphology

inoculation (h)

Dimensions (~m)

3

Young vegetative rods, granulose negative

4.7 X 0.72

27

Vegetative rods,

3.4 X 0.66

clostridia, granulose

4.7 X 1.6

Vegetative rods,

2.6 X 0.6

clostridia,

5.5 X 1.3

free spores

2.4 X 1.2

75

Clostridium acetobutylicum was found to be capable of fermenting a mixture of 50% starch ar,d upto 50% molasses [8]. Methods of isolating clostridia capable of using sugar containing substrates were developed with the substitution of invert molasses for starch [9]. Clostridium propylbutyricum has been used in a patented process involving fermentation of a 4-6% monosaccharide solution to give a 25-30% yield of solvents, mostly butanol, based on original sugar concentration [10]. Many solventogenic clostridia were isolated, producing a varied ratio of neutral products and having varied abilities to utilize a range of carbohydrate substances. However, most of them were suggested to be strains of C. acetobutylicum [11]. On the basis of differences in types of infecting bacteriophages, a different species, Clostridium saccharoperbutylicum was identified [12]. This organism produces relatively more butanol and less acetone than comparable cultures of C. acetobutylicum. Two fermentation types have been distiguished, acetonebutanol fermentation carried out by C. acetobutylicum and closely related saccharolytic strains, and the butanol-isopropanol fermentation carried out by C. butylicum [9].

201 3

KINETICS OF GROWTH AND PRODUCT FORMATION

The rate of cell growth of solventogenic species varies considerably and is influenced by medium composition and strain of the organism (Table 3). The organism quickly achieves its fastest growth rate after a short lag phase (2-3 h). The maximum specific growth rate of C. acetobutylicum in a synthetic medium has been found to be around 0.2 h1, at 6-8 h, thereafter declining slowly to around 0.15 h1 at about 11.5 h [13]. After this phase, the proportion of sporulated cells increases. The fermentability of sugars such as xylose and arabinose is much slower than glucose [14].

Table 3 Acetone-butanol fermentation of different substrates by Clostridium acetobutylicum [15] Strain

Substrate Concentration Total (%,w/v)

no.

NRRL B3179

NRRL 527

Solvents

solvents

B:A:E

(%,v/v)

(% of total)

Xylose

5

1.22

47:47:6

Arabinose

5

1.34

47:47:6

Glucose

5

1.15

47:47:6

Xylan

5

0.19

37:36:27

Cellobiose

10

2.45

79:20:1

Dextrin

5

1.75

45: 45:10

Xylose

5

0.96

83:9:8

Arabinose

10

0.51

75:14:11

Glucose

5

0.16

0:60:40

Xylan

5

0.69

56:13:31

Cellobiose

5

0.68

50:12:38

Dextrin

5

0.68

76:10:14

B:A:E, Butanol:Acetone:Ethanol

202

Compere and Griffith [15] investigated some good solvent producing clostridia namely C. acetobutylicum NRRL B527, C. butylicum NRRL B592, and C. pasteurianum NRRL B598 for the conversion of various mono- and di-saccharides. Growth and solvent synthesis rate with pentoses were comparatively slower. C. butylicum NRRL B592 was found to be particularly efficient in solvent production from xylan and cellobiose (Table 4). In an interesting experiment of Maddox [16], C. acetobutylicum was grown on pentoses in a mixed culture with Saccharomyces cerevisiae. While the S. cerevisiae converted hexoses from molasses (5% solids) to ethanol (22 g/I) in 48 h, the bacterium inoculated into the culture after 24 h utilized arabinose and xylose components (30 g/I) producing 6.6 g/I and 3.7 g/I, respectively of butanol, in another 170 h.

Table 4 Acetone-butanol fermentation of pentoses and hexoses by Clostridium butylicum [15] Strain

Substrate Concentration Total (%,w/v)

no.

NRRL B592

NRRL B593

Solvents

solvents

B:A:E

(%,v/v)

(% of total)

Xylose

5

0.54

66:18:16

Arabinose

5

1.62

81:12:7

Glucose

10

1.98

62:26:12

Xylan

10

2.59

85:14:1

Cellobiose

10

2.87

53:22:25

Dextrin

10

4.08

75:21:4

Xylose

5

0.25

76:12:12

Arabinose

5

0.34

73:6:21

Glucose

10

2.14

69:22:9

Xylan

10

1.10

85:12:3

Cellobiose

10

1.91

74:21:5

Dextrin

10

1.85

93:2:5

B:A:E, Butanol:Acetone:Ethanol

203 Utilization of different sugars for the production of acetone and butanol by C.

acetobutylicum has been investigated [17]. Maximum solvent production was obtained with glucose and cellobiose. More than 90% of the substrate was utilized with butanol production of 20 g/100 g carbon utilized (Table 5). However, in the case of pentose sugars more acids were produced than solvents and the sugar utilization was always less than 60%.

Table 5 Substrate utilization and solvent production of Clostridium acetobutylicum ATCC 824 on various sugars found in hemicellulose a [17] Sugar

Substrate

Solvents (g/100 g carbon utilized)

Utilization

(%)

Butanol

Acetone

Ethanol

Acetic

Butyric

acid

acid

Glucose

97.5

20.9

6.2

1.6

4.4

2.5

Xylose

61.0

7.1

2.3

1.3

10.0

17.6

Arabinose

54.0

9.1

4.9

1.2

16.8

13.2

Mannose

84.5

18.3

5.3

1.7

7.7

6.4

Galactose

58.0

2.2

1.3

1.4

9.3

19.7

Cellobiose

93.5

22.5

6.0

2.1

3.8

2.2

a Substrate concentration 2%, incubation at 37~ for 48h

Solventogenic clostridia are capable of fermenting media with concentration of sugars from 30 to 100 g/I [4]. Generally, initial sugar concentration of 60 g/I is used which yields about 20 g/I solution of mixed solvents. Higher concentration of butanol inhibts growth as well as product formation. The toxicity of butanol to the producing organism determines the substrate concentration that can be used economically.

204 Different substrates also affect fermentation parameters of C. acetobutylicum such as growth rate and solvent production ratio [18]. Generally, growth and substrate consumption rates with xylose, arabinose, or galactose are low when compared to glucose, mannose or cellobiose. The solvent production ratio with pentose sugars was 1:2:5 (ethanol:acetone:butanol), while ratios of 1:4:10 were obtained with the second sugar group. However in one case [19] fermentation of glucose and xylose yielded similar solvent ratios but the xylose consumption rate (0.33 g/I/h) was much lower than that of glucose (0.68 g/I/h). The fermentation of a mixture of xylose and glucose has also been attempted [20]. Presence of xylose in the medium causes a decrease in the substrate utilization (Table 6) as well as solvent yield (Tables 7).

Table 6 Substrate utilization of glucose-xylose mixtutre a % of total sugars

Substrate utilization (%)

Xylose

C. butylicum

C. acetobutylicum

[22]

[20] 80.0

Glucose

0

100

88.6

25

75

89.2

50

50

82.1

56.0

80

20

61.3

30.4

100

0

41.0

46.0

a

Initial sugar concentration 50 g/I

205 Table 7 Acetone-butanol fermentation of glucose-xylose mixture by Clostridium acetobutylicum [20] Initial

Sugar

Total

Solvent

sugar

utilization

solvents

distribution

(g/I)

(%)

(g/I)

B:A:E (% of total)

Xyl

Glu

Xyl

Glu

45

5

14.6

100

1.6

65:30:5

40

10

13.0

100

2.6

71:25:4

30

20

16.7

100

6.6

71:25:4

25

25

12.0

100

9.5

67:29:4

7.3

67:30:3

11.2

65:31:4

50

46.0 50

80

Growth and fermentability of the organism is poor at low glucose concentrations (510 g/I) with only 14% of the sugars consumed [21]. However, best fermentation yields are obtained at a glucose concentration of 40 g/l. Pure xylose is not so easily utilized by the organisms as glucose or glucose-xylose mixture. Some conclusions have been drawn while comparing the fermentation of hexoses and pentoses [22-25]: Growth and solvent synthesis rates on pentoses are substantially lower than that on hexoses. Disaccharides and oligosaccharides give higher solvent accumulation than monosaccharides. The culture may be more sensitive to higher pentose concentration than to hexoses, since even 100 g/I of glucose concentration would result result in good solvent production.

Clostridium butylicum NRRL B592 may be particularly efficient in solvent production

206 from hydrolysates of natural substrates containing xylan and cellobiose, dairy wastes, and starchy material.

4

PERFORMANCE ON NATURAL SUBSTRATES

Since the wood and agricultural residues are plentiful source of fermentable carbohydrate materials, the utilization of these substrates has attracted considerable attention. Pulp and paper industry waste water, namely spent sulfite liquor is potential material for conversion to useful products following pre-treatment to remove BOD up to 45% and supplementing nitrogen and phosphorus sources. Clostridium butylicum can utilize more than 80% of the sugars available in spent sulfite liqor [20]. Few attempts have been made to convert cellulose and hemicellulose complexes directly to acetone and butanol. Petitdemange et al. [26] suggested the use of a co-culture of Clostridium acetobutylicum with cellulolytic clostridia. However, solventogenesis could not be achieved because of low concentration of generated sugars. Similar results were obtained by sequential fermentation of cellulose using Clostridium

thermoceilum followed solventogenesis.

by

C. acetobutylicum with butyrate added to induce

Simultaneous saccharification and fermentation has also been studied to produce butanol from alkali-pre-treated wheat straw using

Trichoderma reesei and C.

acetobutylicum [27]. The final butanol concentration reached was 10.7 g/I from 140 g/I of straw. Simultaneous utilization of both hemicellulose and cellulose components was observed since pentoses did not accumulate during fermentation. C. acetobutylicum possess J}-l,4-glucan glucohydrolase and cellobiase activities, but lack an active cellobiohydrolase to hydrolyze crystalline cellulose. Studies based on a promising strain of C. butylicum, have indicated the problems associated with fermentation of wood hydrolysate [14]. Lime-neutralized hydrolysate, containing 30 g/I sugars was used to obtained the solvent yields of 25-38%. Experiments have also been conducted [28] to determine the best method of pretreating sulfite liquor (Table 8).

207 Table 8 Effect of different methods of waste sulfite liquor pre-treatment prior to acetone-butanol fermentation by Clostridium butylicum (Fitz) [26] Pre-treatment

1. Boiling, neutralization 2. Boiling, neutralization, dilution to 40% of the original strength 3. Boiling, neutralization, norite clarification

% Substrate

Fermentation

utilization

time (days)

0 62.7

0

10 7

10

4. Boiling of S02 at pH 1.5, neutralization

70.0

7

5. SO2 precipitation at pH 10

74.3

3

6. SO2 precipitation at pH 10, lignin precipitation at pH 11.5

72.6

2

7. SO2 precipitation at pH 10, lignin precipitation at pH 11.5,

74.9

2

by Ca(OH)2 neutralization

Ca precipitation with 1% Na2SO4

Bioconversion of agricultural wastes into acetone and butanol by Clostridium

saccharoperbutylicum was studied by Soni et al. [29]. Alkali-treated bagasse and rice straw were hydrolyzed by using an enzyme mixture from Trichoderma reesei and

Aspergillus wentii for 30 h. The resulting hydrolysate (6% reducing sugars) was fermented. Poor solvent production was improved by ammonium sulfate precipitation and activated carbon treatment of the hydrolysate. Sjolander et al. [30] have indicated the problems associated with bioconversion of wood hydrolysate.ln most of the cases, the inhibitory effect was accounted for by the presence of xylose decomposition product,

208

furfural. Steam stripping was found to be a simple and effective method. A 1:1 mixture of wood xylose and hexoses at pH 6.5 gave 1.7% butanol, 0.9% acetoin and 0.19% ethanol accumulation at the end of 48 h fermentation [31]. The solvent yields by C. butylicum (Prazmowski) ranged from the usual 30% to 38% of the available sugars, while butanol fraction constituted 67% of the total solvents [32].

C. acetobutylicum ATCC 824 was found to be able to utilize most of the sugars liberated from the hydrolysed hemicellulose fraction [33]. However, solvent yields were low, partly due to the poor utilization of xylose. Highest butanol yields were obtained on water soluble fraction of the solvent extracted wood (Table 9). This relatively higher yield could be the result of low molecular weight oligomers such as cellobiose, which is efficiently utilized by this organism.

Table 9 Substrate utilization and solvent production by Clostridium acetobutylicum ATCC 824 on enzymatic hydrolysate of wood hemicellulose [17] Substrate

Substrate

Solvent (g/100 g of carbon utilized)

utilization

(%)

Butanol

Acetone

Ethanol

Acetic

Butyric

Xylan

87

4.0

1.5

ND

6.3

7.9

SEW-WS1

91

13.5

1.2

ND

28.4

14.8

SEW

73

4.4

1.2

ND

44.7

5.6

SEA, solvent extracted aspen; SEW-WS1, steam exploded wood, water soluble fraction; ND, not detected

Starch bearing grains and potatoes (also contain appreciable amount of pentosans) were the first sources of carbohydrates used in acetone-butanol process [25]. In industrial practice, the grain of choice has been maize, the fermentation of which is completed in 50-60 h with an overall solvent yield of 38%. Other raw materials used for acetone-

209 butanol production are whey and domestic wastes [34-36].

5

FACTORS AFFECTING ACETONE AND BUTANOL PRODUCTION

A number of culture process parameters have been identified by several researchers which may have significant effects on acetone-butanol fermentation. In considering approaches to improve product concentration and yield, several characteristics of pentose fermenting organisms are of particular interest and require attention.

5.1

pH and temperature

Acetone and butanol are the major fermentation products of Clostridium

acetobutylicum when grown in continuous culture under phosphate limitation at pH 4.3 [37-39]. At pH values above 5.5, however, an exclusive acetate-butyrate fermentation is carried out by this organism under phosphate, ammonia or glucose limitation [40]. In batch fermentation, the concentration of acetic acid and butyric acid rise initially, at pH about 4.0, acid concentration begins to fall and the amount of acetone and butanol increases with the rise of pH. Most acetone-butanol producers are mesophilic and display temperature optima for fermentation between 30 and 37~

At lower temperature, solvent production by C.

acetobutylicum was decreased, but at elevated temperature acetone yield decreased but butanol yield remained unaffected [41]. A thermophilic strain capable of producing butanol from cellulose at 50-65~ has also been reported [42]. Optimum temperature for grain fermentation is between 36 and 37~ 34-41~

though the fermentation is normal in the range of

There is usually a loss of acetone and change in the solvent ratio at higher

temperature [25,43]. The effect of temperature on solvent yield is shown in Table 10.

210 Table 10 Effect of temperature on solvent production by Clostridium acetobutylicum Reference

Temperature

Yield

Butanol/Acetone

(~

(%)

ratio

25

29.1

3.48

[1]

30

28.4

3.70

[1]

30

31.0

[43]

33

30.0

[43]

37

25.5

37

24.0

40

24.9

4.73

[1] [43]

5.67

[1]

5.2 Repeated subculturing

Continuous subculturing of Clostridiumpasteurianum may bring about degenerative changes in the culture leading to the decreased fermentation rate and the absence of spores or clostridial forms [1]. However, subculturing of a strain of C. butylicum could be carried out 6 to 10 times without any clear physiological changes [44]. Further subculturing slows down the fermentation rate, and actively growing cultures could be maintained only by heat shocking. Table 11 shows the effect of subculturing on solvent production [8]. Interestingly, total solvent yield increases with serial subculturing, while yield of acetone decreases.

211 Table 11 Effect of subculturing followed by heat shocking on solvent yield of Clostridium

acetobutylicum [8] Serial

Total

Acetone

subculture

solvents

(% of total)

(g/I) 2

15.08

35.45

4

16.03

32.40

6

18.40

28.00

8

19.30

30.50

10

19.98

28.50

12

19.85

25.80

5.3

Production of bacteriocin

Clostridium acetobutylicum cells undergo autolysis in the stationary phase of the fermentation. A bacteriocin has been isolated from these fermentations [45]. It causes extensive lysis of the culture involving cells associated with solvent production [46]. Release of bacteriocin begins late in the exponential phase of growth (24 h). The substance has been characterized as a glycoprotein of Mr 28 kDa. The autolysin gene appeared to be chromosomal since no plasmid DNA was detected in this C.

acetobutylicum strain [47]. The relationship between high butanol levels and increased autolytic activity has been established [48].

212 5.4

Nutrition

Several nutritional factors affect acetone-butanol fermentation. Vitamin requirements of the solvent producing strains were obscured for many years by the use of complex media containing materials such as yeast extract or corn steep liquor as sources of vitamins and nitrogen. Most of the acetone-butanol producing strains require p-aminobenzoic acid and biotin for growth [21]. Thiamine has also been used in the defined medium in some cases [18]. Clostridium beijerincki required multiple amino acids and vitamins for growth [9,33]. Initial acidic fermentation products originally formed are remetabolized and converted to more reduced neutral products [49-52]. The addition of butyric acid to fermentation media increased the yield of butanol. Johnson et al. [53] reported that the additon of acetate also increased the yield of acetone. This was later confirmed by Wood et al. [54] who used 13C-labelled acids to the fermentation and demonstrated incorporation of the label into neutral products. While 85% of the butyric acid label was found in butanol fraction, only 15-19% of added acetic acid label was found in acetone and isopropanol, and 50% in butanol fraction. The addition of acetic or butyric acid (2 g/I) has been found to improve the solvent yields (Table 12) [55]. No beneficial effect on solvent production by the addition of these acids were observed to a culture grown on a glucose-based medium [56]. On the other hand, addition of acetic acid to a xylose-based medium increased solvent levels by three to four-fold.

Table 12 Effect of acetic and butyric acids on solvent production by Clostridium acetobutylicum [55] Butanol:Acetone:Ethanol ratio

Addition

Total

(2 g/I)

solvents (%)

None

32

6.0: 1.9:0.6

Acetic acid

34

6.0: 3.0:0.5

Butyric acid

35

6.0: 2.4:0.8

213 Ammonium sulfate (2-3 g/I)is the most common nitrogen source used in synthetic medium for solvent production [20,27]. Effect of different nitrogen sources on solvent yields is shown in Table 13.

Table 13 Effect of nitrogen sources on solvent yield [25] Nitrogen

Solvent

Acetone

source

yield (%)

(% of total solvents)

Ammonium sulfate

29.8

22.2

Ammonium hydroxide

31.5

22.4

Ammonium nitrate

23.6

40.9

Ammonium chloride

29.3

22.5

Ammonium acetate

30.4

27.5

The presence of MnSO 4 upto a concentration of 20 mg/I had little effect on the fermentation [21]. Levels between 0.001 and 0.05 mg/I of FeSO4 gave similar results, but in the absence of this element growth and substrate utilization was very poor. Solvent production has been found to increase with the increase in potassium concentration from 0-60 mg/l. At high levels of KCI (0.6-0.8 g/I), substrate utilization was found to be reduced but without affecting solvent production.

5.5 Oxygenation

Many steps have been taken to ensure anaerobic conditions during fermentation, ranging from the use of reducing agents, oxygen-free nitrogen, and gas-proof tubing on

214

the experimental scale to blanket of fermentation gas on the industrial scale. As obligate anaerobes, butanol producing organisms require anaerobic conditions. A low redox potential (below-250 mV)is essential for acetone-butanol fermentation [57]. Clostridium

acetobutylicum NCIB 8052 can detoxify molecular oxygen by NADH without forming H202 [58]. This organism was able to grow anaerobically and produced solvents at a redox potential of +370 mV poised by potassium ferrocyanide. Vegetative cells survived in the presence of oxygen for several hours. Work of Hongo [59] with redox dyes showed that higher yields of butanol can be achieved by the addition of 5mM neutral red into the fermentation system. This effect was later ascribed to the presence of neutral red-linked hydrogenase and pyridine nucleotide reductase activity in cells [57]. The mechanism of oxygen toxicity has been investigated in detail [59] and concluded as follows: Oxygen is itself a toxic agent. Anaerobes require low redox potentials to grow well. Organisms lacking catalase, are killed by H202 formed by reducing some of the oxygen. When growth of C. acetobutylicum was studied under anaerobic (E, -400 to -370 mV), aerated (Eh -50 to 0 mV; DO <1 I~M)), and aerobic (Eh +100 to +150 mV; DO 40-50 I~M) conditions, the specific growth rates (about 0.6 h1) of the organism were similar under aerated and anaerobic conditions [58]. Exposure of an anaerobic culture to oxygen (40-60 I~M) for periods upto 6 h was not found lethal. At high DO levels, the rate of substrate consumption decreased, and growth and net synthesis of DNA, RNA, and protein stopped. However, these consequences of oxygenation were all reversible. There was no evidence to suggest the formation of H202. Oxygen (40 ~M) inhibited growth in a medium poised at-50 mV, whereas growth was normal in an anaerobic environment poised at +370 mV. Biochemical and physiological consequences of oxygenation in acetone-butanol fermenting organisms are Decrease in the rate of substrate consumption. Cessation of growth of the organism. Cessation of net synthesis of DNA, RNA, and protein. Fall in the intracellular ATP level.

215 Increase in NADH oxidase activity. Cessation of butyrate formation. Starvation of energy and draining of reducing power.

5.6

Continuous culture

Industrially, perhaps the greatest advantage offered by continuous culture is the increased productivity often associated with this technology when compared to conventional batch fermentation. Despite this fact, continuous culture has largely remained a research tool, probably due to the practical difficulties inherent in this technique, including the difficulty in prevention of contamination and the possibility of mutation in the organism and the capital cost of conversion from batch to continuous. A continuous acetone-butanol fermentation was developed in 1932 but the process was never adopted on a commercial scale [60]. A large scale continuous acetone-butanol process has been reported by Dyr et al. [61]. The plant is described as having one seed tank and seven fermentors of 220-270 m 3, with separately adjusted feeds of wood hydrolysate/flour/molasses mixture to a concentration of 40-60 g/I sugars.The system gave a 20% increase in the productivity over the batch process, as well as substantial substrate economy. However, the normal duration of continuous fermentation was less than 90 h because of infection problem. Loss of solvent producing ability of C.

acetobutylicum NC1B8052 [50] and C.

acetobutylicum DSM1731 [40] have also been reported in nitrogen-limited chemostat cultures. Significant solvent production can be obtained when substrate is always in excess and dilution rate is low [62]. Maximum solvent production occurred with 45.5 g/I substrate at pH 5 and dilution rate 0.038 h-1. Table 14 shows some results on continuous production of acetone and butanol.

216 Table 14 Production of acetone and butanol in continuous culture of Ciostridium acetobutylicum Condition

Butanol mM

N-limited

5.7

C-limited

60.0

+ butyrate Excess carbon

5.7

Acetone g/I

mM

Reference g/I [32]

4.6

5.0

0.29

8.5 9.6

[32] [62]

4.3

[32]

Whole cell immobilization

Calcium alginate immobilized spores of C. butylicum have been used in a continuous butanol production [63]. A 10 cm long conical column was run at 37~ for 215 h with mean productivity of 1 g/I/h which was about four times higher than that of the conventional batch fermentation. Yield of butanol and isopropanol was about 30%, with final butanol concentration below 5 g/l. Calcium alginate immobilized mixture of vegetative cells and spores of C.

acetobutylicum ATCC824 has also been studied for butanol production [64]. Butanol yield was 17.6% and product formation using immobilized exponential phase cells was identical to a normal batch fermentation. When immobilized stationary phase cells in a continuous culture were used, solvent production was more rapid with butyric acid

in the feed,

though butanol yield (11%) was lower. With glucose alone as a substrate, production rate was halved but yield rose to 15.3%. On the other hand, immobilized spores yielded 20.9% butanol. Fermentation using immobilized spores could be carried out over 11 days with productivity of the system being 0.48 to 0.64 g butanol/I/h. Table 15 shows butanol production by immobilized growing cells.

217 Table 15 Production of butanol by calcium alginate immobilized whole cells of solvent producing organisms Organism

Spore/

Butanol

cell

Yield

Reference

(%) Clostridium butylicum

Spores

30.0

[63]

Clostridium acetobutylicum

Exponential

17.6

[64]

Stationary phase cells

11.0

[64]

Spores

20.9

[64]

phase cells

To develop an economically viable acetone-butanol fermentation process, a number of points are required to be considered: Any fermentation process will have to compete with the existing petroleum based process. Utilization of a fully integrated plant using the most inexpensive raw materials such as waste lignocellulosic materials. Strains with improved product tolerance will be required. At present, solvent production from pentoses is low as compared to glucose. Simultaneous efficient utilization of both sugars will have a major impact on process economy.

218 6

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Muller J. U.S. Patent 1938; 2,123,078.

11

George HA, Johnson JL, Moore WEC, Holdeman LV, Chen JS. Appl Environ Microbiol 1983; 45: 1160.

12

Hongo M, Murata A. Agri Biol Chem 1965; 29:1135.

13

Gottschal JC, Morris JG. Biotechnol Lett 1981; 3: 525.

14

Leonard RH, Peterson WH. Ind Eng Chem 1947; 39: 1443.

15

Compere AL, Griffith WL. Proc 35th Meeting of Society of Industrial Microbiology, 1979; 509.

16

Maddox IS. Biotechnol Lett 1982; 4: 23.

17

Saddler JN, Yu EKC, Mes-Hartree M, Levitin N, Brownell HH. Appl Environ Microbiol 1983; 45: 153.

18

Long S, Jones DT, Woods DR. Appl Microbiol Biotechnol 1983; 45: 1379.

19

Ounine K, Petitdemanage H, Raval G, Gay R. Biotechnol Lett 1983; 5: 605.

20

Volesky B, Szczesny T. Adv Biochem Eng/Biotechnol 1983; 27: 101.

21

Monot F, Martin JR, Petitdemanage H, Gay R. Appl Environ Microbiol 1982; 44: 1318.

22

Langlykke AF, van Lanen JM, Fraser DR. Ind Eng Chem 1948; 40: 1716.

23

Underkofler LA, Hunter JR. Ind Eng Chem 1938; 30: 480.

24

Rose AH. Industrial Microbiology, London: Butterworths, 1961; 160.

25

Beesch SC. Ind Eng Chem 1952; 44: 1677.

219 26

Petitdemanage H, Fond O, Caillet F, Gay R. Biotechnol Lett 1983; 5: 119.

27

Marchal R, Rebellar R, Vandcasteele TP. Biotechnol Lett 1984; 6: 523.

28

Wiley AJ. Ind Eng Chem 1941;33: 606.

29

Soni BK, Das K, Ghose TK. Biotechnol Lett 1982; 4: 19.

30

Sjolander NO, Langlykke AF, Peterson WH. Ind Eng Chem 1938; 30: 1251.

31

Tornescher Heffe GmbH. German Patent 1954; 920724.

32

Kalnina V. Latv PSR Zin Akad Vest Kim Ser 1961; 2: 269.

33

Mes-Hartree M, Saddler JN. Biotechnol Lett 1982; 4: 247.

34

Wilson PW, Peterson WH, Fred EB. J Biol Chem 1927; 74: 495.

35

Solomans GL. Proc Biochem 1976; 11 : 32.

36

Jean JW. U.S. Patent 1939; 2,182,989.

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Peterson WH, Fred EB. Ind Eng Chem 1932; 24: 237.

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Bahl H, Andersch W, Braun K, Gottschalk G. Biotechnol Lett 1982; 4: 29.

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Andersch W, Bahl H, Gottschalk G. Eur J Appl Microbiol Biotechnol 1983; 18: 327.

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McNeil B, Kristiansen B. Biotechnol Lett 1985; 7: 49.

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McCutchan WN, Hickey RJ. In: (Underkofler LA Hickey RJ eds.) Industrial Fermentations, New York: Chemical Pub Co., 1954; 347.

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Kutzenok A, Aschner M. J Bacteriol 1952; 64: 829.

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Finn RK, Nowry JE. Appl Microbiol 1959; 7: 29.

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Webster JR, Reid SJ, Jones DT, Woods DR. Appl Environ Microbiol 1981; 41: 371.

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van der Westhuizen A, Jones DT, Woods DR. Appl Environ Microbiol 1982; 44: 1277.

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Speakman HB. J Biol Chem 1920; 43: 401.

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Johnson MH, Peterson WH, Fred EB. J Biol Chem 1933; 101: 145.

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63

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64

Haggstrom L, Molin N. Biotechnol Lett 1980; 2: 241.

8 Microbial Production of 2,3-Butanediol

1

INTRODUCTION

Also known as 2,3-butylene glycol, dimethylene glycol or 2,3-dihydroxybutane, 2,3butanediol is a colourless and odourless liquid of high boiling point [1]. This compound can not be easily recovered by conventional distillation because of its high boiling point (about 180~ The levo-isomer form of 2,3-butanediol has a low (-60~ freezing point which forms the basis of commercial interest in the use of 2,3-butanediol as an antifreeze agent [2,3]. With a heating value of 27,200 KJ/kg, butanediol compares favourably with ethanol (29,100 KJ/kg) and methanol (22,100 KJ/kg) for use as a liquid fuel [4]. Condensation of diol to methyl ethyl ketone coupled with subsequent hydrogenation, yields ocatne isomers that can be used to produce high quality aviation fuels. Butanediol can also be employed in the production of halogenated substitutes, esters of monobasic and dibasic acids, oxides, nitrogen, ether and ketone derivatives [5]. Methyl ethyl ketone can also be used as a solvent for resins and lacquers [6]. In addition butanediol has applications in the manufacturing of high valued food additives (diacetyl), synthetic perfumes, printing inks, organic solvents, emulsifying agents, fumigants, explosives, pharmaceuticals and drugs, polyesters, plastics and amorphous resins [7-9]. The earliest report on microbial production of butanediol is that of Harden and Walpole [10] using Aerobacter aerogenes and Harden and Norris [11] using Klebsiella

pneumoniae. Later, Donker [12] investigated butanediol fermentation with Bacillus polymyxa. During next 16 years, butanediol fermentation did not attract much interest except for a report on large-scale production of butanediol [13]. Shortage of a strategic compound 1,3-butadiene during World War II stimulated studies on commercial butanediol production. Butanediol can be converted to butadiene, which is the major constituent in the production of buna-S-rubber [14]. The termination of war prevented the commercial application of the process. A large number of process were developed as a result of accelerated investigations by Canadian and United States Agencies in cooperation with private fermentation industries. Although several of these investigations led to pilot-plant operations, none reached commercial production [15]. 221

222

Microbial fermentation generate three stereoisomeric forms of 2,3-butanediol (Figure 1). In a given process, the production of a particular isomer is dependent upon the microorganism employed. Table 1 summarizes the nature of stereoisomeric forms of butanediol produced by potential microbial species.

CH3 i HO-C-H I H-C-OH I

CH3 D(-)

CH3 I H-C-OH I H-C-OH I

CH3 Meso

CH3 I H-C-OH i HO-C-H I

CH3 L(-)

Figure 1. Stereoisomeric forms of 2,3-butanediol

Table 1 Stereoisomeric forms of 2,3-butanediol produced by bacterial fermentation Microorganism

Stereoisomers

Aeromonas ( Pseudomonas) hydrophila Bacillus polymyxa

50% racemic, 48% meso, 2% levo

Bacillus subtilis Klebsiella pneumoniae

65% (D-), remainder meso

D(-) (levo) 5-14% L(+), remainder meso

(Aerobacter aerogenes) Serratia spp.

Primarily meso

Industrial scale production of solvent and chemicals via bioconversion technology requires the availability of substrates that are both abundant and inexpensive. Purified

223 carbohydrates such as glucose, sucrose and lactose, while readily fermentable, are extremely costly for this purpose. Thus the economics of butanediol fermentation limits the carbohydrate sources to industrial and agricultural residues. The waste sulfite liquor produced from sulfite pulp and paper mills besides posing a serious disposal problem for the industry is also an industrial waste material of little or no utility. This waste material has successfully been used as substrate for butanediol production by several bacterial species [16]. Interest in butanediol fermentation was renewed in the last decade mainly because of the ability of fermenting pentose sugars and lignocellulosic hydrolysate as cheap substrate [17]. However, the market for the product is still questionable [18].

2

MICROORGANISMS USED FOR 2,3-BUTANEDIOL PRODUCTION

A number of microbial species produce butanediol but only a few do so in significant quantities. Although butanediol formation has been observed in several yeasts [19] and filamentous fungi [20] like Rhizopus nigricans and Penicillium expansum, the conversion efficiency was extremely poor (approximately 0.003 g/g glucose). Thus bacteria are, at present, the only organisms of industrial importance in butanediol production. The most potential species belong to the genera Klebsiella (Aerobacter), Bacillus, Serratia, and

Pseudomonas (Aeromonas). Klebsiella pneumoniae (Aerobacter aerogenes), widely distributed in nature, is stable under a wide range of environmental conditions. Although it lacks polysaccharolytic properties, it produces high yields of butanediol with much lower yields of by-products. It retains its viability on a variety of stock culture media and may even be cultivated on media containing the fermentation substrate without loss of activity [21]. K. pneumoniae has a tendency to preferentially metabolize first glucose and then mannose among the carbohydrates present in lignocellulosic hydrolysate. Xylose, arabinose and galactose are consumed at slower rates.

Bacillus polymyxa has the ability to ferment a wide range of substrates including starch and hemiceilulose. Its amylolytic capability enables its use in the conversion of unhydrolyzed grain mashes [22,23]. B. polymyxa possess xylanase activity and can fully utilize the hemicellulosic components of natural substrates [24,25]. However, the ability

224

to ferment cellulose is rather weak or lacking. The disadvantages of this organism are the formation of large amount of ethanol as by-product and loss of fermentation activity during storage or repeated subculturing [26]. It is also very susceptible to bacteriophage attack [27]. Pilot-scale production of butanediol from agricultural residues using B. polymyxa has been attempted [28,29].

Bacillus subtilis is another species that can produce butanediol as major product of carbohydrate metabolism. However, it does not produce pure levo butanediol. Growth and bioconversion rates are weak as compared to other organisms. Production of significant amount of glycerol, at the expense of ethanol, is a common feature of this organism [30]. Serratia spp. are capable of generating good yields of butanediol, but significant levels of organic acids (formic and lactic acid) are also produced. Under anaerobic conditions, these two acids may account for 37% (molar basis) of the total products formed [31]. However, ability to ferment D-xylose is strain dependent. Most important butanediol producing species are S. marcescens, S. indica, S. plymuthica, S. anolium and

S. kiliensis [32,33]. Aeromonas (Pseudomonas) hydrophila has been used for the bioconversion of pentoses [34,35] and starch [36] into butanediol.This organism produces appreciable amounts of ethanol as by-product. However, it is able to produce butanediol in a reasonably efficient manner, without stringent pH or aeration control [35]. Although most yeasts and filamentous fungi cannot ferment pentoses anaerobically, many bacteria readily convert xylose to a variety of products in the absence of oxygen [37]. The rates, yield and products formed by these bacteria depend not only on the diverse metabolic pathways operating during anaerobic fermentation but also on the species, strains, substrates and culture conditions used [38]. The net equation for the bioconversion of xylose and glucose to butanediol are: Xylose .... > 5/3 CO 2 + 5/6 NADH 2 + 5/3 ATP + 5/6 Butanediol Glucose .... > 2 CO 2 + NADH 2 + 2 ATP + Butanediol Of the total carbon, 1/3 is lost as carbon dioxide, while 2/3 is converted to 2,3butanediol. On a mass basis, the yield of butanediol from both xylose and glucose is 50%. The theoretical maximum molar yield of butanediol from pentoses is 0.83 and from hexoses is 1.0. Table 2 shows bacterial species capable of producing 2,3-butanediol from different raw materials.

225

Table 2 Microorganism capable of producing 2,3-butanediol from different raw materials Organism

Raw material

Bacillus polymyxa

Molasses

B. subtilis Klebsiella pneumoniae Aeromonas hydrophila Serratia marcescens B. polymyxa

Whole wheat, corn starch,

Aerobacter aerogenes

acid hydrolyzed wheat, barley

K. pneumoniae

Wood hydrolysate, starch

A. hydrophila B. polymyxa

Sulfite waste liquor

B. subtilis A. hydrophila A. aerogenes Serratia spp. A. aerogenes

Sucrose

K. pneumoniae (K. oxytoca)

Pentoses

B. polymyxa

Some of the physiological characteristics inherent in these organisms include (a) the fermentation of pentose sugars without the need for critical oxygenation unlike yeast and fungi, (b) shorter generation time which is reflected as high conversion rate during

226 fermentation, (c) utilization of less proportion of substrate carbon for biomass formation, hence indicate the possibility of obtaining higher specific product yield.

3

KINETICS OF GROWTH AND PRODUCT FORMATION

Initial work on butanediol production was focussed upon glucose utilization. However, the need for evaluation of cheaper substrates for commercial feasibility was soon recognized. Consequently the principle sugars from the lignocellulosic hydrolysate (glucose, xylose, mannose, galactose and arabinose) were examined (Table 3).

Table 3 Fermentation of various monosaccharides for 2,3-butanediol production Organism

Substrate

Butanediol Reference (g/100g sugar)

B. polymyxa

Glucose

32.6

[14]

S. marcescens

Glucose

32.0

[14]

B. subtilis

Glucose

27.5

[14]

A. aerogenes NRRL-B199

Glucose

41.6

[45]

K. oxytoca ATCC 8274

Xylose

29.6

[49]

B. polymyxa

Xylose

29.0

[24]

K. pneumoniae ATCC 8274

Cellobiose

18.7

[40]

Mannose

24.8

[40]

Arabinose

22.6

[40]

Galactose

14.4

[40]

Glucose

25.4

[60]

Mannose

29.7

[37]

K. pneumoniae AU-I-d3

227 The importance of aeration in D-xylose fermentation was identified during early studies on butanediol production [15]. Yields of acetic acid frequently exceeds butanediol in the absence of oxygen. Almost equimolar concentration of ethanol and butanediol are obtained from arabinose, galactose and mannose [39]. Saddler et al. [40] investigated the utilization of sugars found in hemicellulose hydrolysate using K. pneumoniae. All the sugars were found to be completely utilized within 24 h. Maximum yield was obtained with xylose as substrate. Sugar concentration has a significant effect on butanediol production and reaction rates [13,15,41]. At higher sugar concentrations, the yield and rate both decreased. The optimum sugar concentration often depends on the particular substrate used as carbon source. Long and Patrick [15] suggested that with increase in sugar concentration, the level of accompanying toxic material also increases, resulting in poor substrate utilization. When an acid-hydrolysed wheat mesh medium was employed, the butanediol yield decreased due to incomplete utilization of sugars [42]. The apparent substrate inhibition could be explained by a decreasing water activity and altering osmolarity. To overcome these problems fed-batch and continuous fermentations have been investigated [4,39,43]. Using a defined medium in a slow feed biorector, upto 250 g/I glucose could be utilized for butanediol production [44]. With the increase in substrate concentration, increase in butanediol yield and decrease in ethanol and biomass yield of Aerobacter aerogenes NRRL B199 has been observed [45]. At substrate (glucose) concentration of 195 g/I, the butanediol yield reached 90% of its theoretical value. When fermentation begins, a large part of the substrate is used to synthesize biomass, later on, however, most of the substrate serves to produce butanediol. Reports concerning the influence of initial xylose concentration on butanediol yields are conflicting. Substrate concentrations of both xylose and glucose in excess of 20 g/I could not be efficiently utilized even after prolonged incubation [46]. Although growth and butanediol accumulation were slower in cultures containing higher substrate concentration, the final yields were not significantly different. A large number of viable cells were observed even in 30 days old culture. Reinoculation with healthy culture also failed to revitalize the process. However, complete utilization of 50 g/I xylose and glucose could be achieved with near theoretical yields of butanediol (88 and 94% from glucose and xylose, respectively) in the presence of acetic acid [47]. Higher butanediol yields were obtained when the sugars were sterilized separately before adding them to the autoclaved media. Butanediol production was also found to increase when Klebsiella pneumoniae ATCC 8724 was grown on high xylose concentration under aerobic condition [46]. Cells

228 acclimatized to high substrate concentration, could also utilize xylose efficiently [46]. On longer incubation, the level of butanediol decreased, probably due to the reoxidation of butanediol to acetoin (acetyl methyl carbinal) [48]. Table 4 shows the effect of sugar concentration on butanediol production by different strains.

Table 4 Effect of substrate concentration on 2,3-butanediol production Organism

K. pneumoniae ATCC 8724

K. oxytoca ATCC 8724

B. polymyxa

A. aerogenes NRRL-B199

Substrate

Butanediol

(g/I)

(g/I)

Reference

[46]

Xylose 50

13.9

100

23.8

150

44.3 [49]

Xylose 10

0.5

20

3.0

50

11.0

100

29.6

150

49.0 [24]

Xylose 20

4.0

40

6.6

60

6.4

100

0.8 [45]

Glucose 22

7.2

45

14.5

107

44.5

195

88.0

229 Maximum specific growth rate of K. oxytoca ATCC 8724 (1.05 h1) accurs at 20 g/I xylose concentration [49]. Kinetics of xyiose fermentation using Kiebsiella oxytoca was investigated. Bacteria begin growing exponentially after a lag time of about 3 h. During exponential growth, the carbon dioxide evolution and the oxygen uptake rates increase at the same rate, while respiratory quotient (RQ = CO2 evolution rate/O2 uptake rate) remains close to 1.0. K. oxytoca appears to have a critical DO level of 3% saturated with air, below this level metabolic changes and the cell yield decreased [50]. After oxygen limitation butanediol formation begins. The maximum product concentartion is obtained at 16.3 h. Of the total 52.6 g carbon utilized (from 132 g xylose), 35.4% goes to 2,3butanediol and 3.5% to acetoin (Table 5). The overall cell yield of 0.16 g/g xylose for this oxygen limited experiment was lower than an aerobic cell yield 0.30 g/g xyiose [51].

Table 5 Carbon balance for 2,3-butanediol fermentation of D-xylose Product

Aerobacter"

Bacillusa

Aerobacter~

Klebsieilab

hydrophila

polymyxa

indolegenes

pneumoniae

[38]

[38]

[49]

[38] Butan ed iol

39.0

38.0

44.0

35.4

Acetoin

2.6

2.5

0.9

3.5

Ethanol

48.9

63.0

55.9

2.8

Acetic acid

9.3

7.7

11.4

2.0

Succinic acid

1.1

5.5

Lactic acid

20.4

5.2

Carbon dioxide

134.7

161.0

114.0

Hydrogen

53.9

82.0

19.1

Cell mass % Carbon recovered

a

33.8 19.9

96.6

mM/100 mM D-xylose fermented

b g/100 g D-xylose fermented

92.9

98.5

97.4

230 Laube et al. [24] indicated a steady decline in butanediol yields as xylose levels were increased to 60 g/I, whereas Yu and Saddler [46] observed no significant change in butanediol yields at different initial xylose concentrations. Jansen et al [49] reported improved butanediol yields as a result of increased xylose concentration. The major factors contributing to disparity among the results of different research groups are the choice of organism and culture conditions employed. Chemically defined medium of pH 6.5 with 0.5% acetic acid in shake flask culture at 30~ was used by Yu and Saddler [46]. Jansen et al. [49] carried out fermentation at 37~ in a 7-1 bioreactor with pH-controlled automatically at 5.2. Mannose, the predominant hemicellulose sugar in southern pine water prehydrolysate has been fermented to butanediol using K. pneumoniae AU-I-d3 [37]. Fermentation kinetics was investigated using different concentrations of pure mannose. The yield of butanediol and ethanol reached maximum 29.7 and 10.7 g/I, respectively, at mannose concentration of 100 g/I, and decreased at higher mannose concentration. A comparative kinetics of 2,3-butanediol fermentation by different organisms is shown in Table 6.

Table 6 Comparative kinetics of 2,3-butanediol fermentation Parameter

Klebsiella

Klebsiella

Bacillus

Aerobacter

pneumoniae [46]

oxytoca [49]

polymyxa [25]

aerogenes [45]

Xylose (100)

Xylose (40)

Glucose (107)

Substrate (g/I) Xylose (50) ~max (h")

0.48

Butanediol (g/I)

13.9

29.6

6.6

44.5

Ethanol (g/I)

2.5

Yield (g/g)

0.35

0.30

0.26

0.42

Productivity (g/I/h)

0.29

1.35

0.085

2.02

1.98

231 4

REACTOR SYSTEMS

The operational conditions of the fermentation vessel are also very important in the establishment of an optimal process design. In batch cultures of K. pneumoniae, butanediol concentrations of 30-65 g/I with corresponding yields of 0.31-0.43 g/g substrate from various carbon sources including xylose have been reported [13,46,49,52]. However, the major disadvantage identified in batch process was the low reactor productivity, mainly because of the long fermentation period required to attain high cell densities to result in a rapid reaction rate [42]. Further, final butanediol concentration is limited by the maximum initial substrate concentration that can be tolerated by the bacteria. Such problem can be sorted by carrying out fed-batch fermentations. Butanediol concentrations of 83 g/I [53] and 99 g/I [44] with corresponding yields of 0.42 g/g and 0.37 g/g of xylose and glucose, respectively, have been reported. When a fed-batch fermentation with acclimatized cells as an inoculum was carried out, upto 160 g of xylose and 190 g of glucose could be utilized rapidly but without any significant improvement in butanediol yields. Using a double fed-batch approach (daily addition of sugars together with yeast extract), under aerobic conditions, accumulation of 88 and 113 g/I of combined butanediol and acetoin yield were obtained from 190 g/I of xylose and 226 g/I of glucose, respectively. Continuous culture techniques have been used very often for solvent production due to the inherent capacity for elevated productivities. High reactor productivities (2.7 g/I/h) has been reported in the continuous culture at a dilution rate of 0.1 per h [54]. Increase in the dilution rate to 0.2 per h further increased the productivity to 4.6 g/I/h, but butanediol concentration decreased. Lower butanediol yields obtained during continuous culture might be due to the loss of some sugars in the product stream. Moreover, a high final product concentration may be difficult since the entire process is continually subject to product inhibition to the maximum extent [42,49]. Adoption of a two-stage continuous culture approach resulted in a further increase in the butanediol production [55]. The overall butanediol productivity was 2.58 g/I/h. Butanediol production by immobilized cultures has had limited success to date. K.

pneumoniae immobilized on ~<-carrageenan produced 15 g/I of butanediol from 50 g/I of glucose at a productivity of 0.5 g/I/h [56]. Conversion of 100 g/I xylose solution at a rate three times greater that of conventional batch reaction has been achieved by immobilizing the cells on 1/2 inch Raschig rings in a closed loop reactor [57]. However, oxygen

232 transfer to the immobilized cells was problematic. Table 7 presents a comparison of 2,3 butanediol fermentation in various reactor systems.

Table 7 2,3-Butanediol production in different reactor systems Reactor

Substrate

system

Yield

Productivity

(g/g)

Reference

(g/I/h)

Batch

Xylose

0.30

1.35

[49]

Batch

Arabinose

0.26

0.36

[60]

Batch

Glucose

0.42

2.02

[45]

Batch

Sucrose

0.43

1.60

[52]

Fed-batch

Xylose

0.42

1.20

[53]

Fed-batch

Glucose

0.37

0.92

[44]

Double fed-batch

Xylose

0.43

0.56

[46]

Double fed-batch

Glucose

0.47

0.74

[46]

Continuous

Sucrose

0.48

3.28

[55]

Two-stage continuous

Sucrose

0.48

2.58

[55]

Immobilized batch

Xylose

0.33

0.60

[57]

Immobilized batch

Glucose

0.30

0.50

[56]

Immobilized continuous

Glucose

0.12

0.75

[56]

5

PERFORMANCE ON NATURAL SUBSTRATES

Several wood and agricultural residues have been considered as possible feedstock for the production of liquid fuels. Table 8 summarizes some results on butanediol production from potential natural substrates.

233 Table 8 2,3-Butanediol production from natural and complex substrates Substrate

Treatment

Sugars (g/I)

Butanediol yield

Reference

(g/g)

Beet pulp

Cellulase

11.0

0.22

[59]

Wood

Acid

46.0

0.29

[63]

Red oak

Acid

100.0

0.36

[21]

Aspen

Steam, acid

12.1

0.50

[47]

Steam, enzyme

10.0

0.22

[40]

Aspen cellulose

Acid

40.0

0.50

[60]

Southern pine

Steam, acid

21.0

0.38

[37]

Spent sulfite liquor

None

38.0

0.24

[16]

Aspen wood xylan

Xylanase

26.7

0.38

[60]

Solka Floc

Cellulase

40.0

0.26

[60]

Corn stalks

Steam,enzyme

7.7

0.31

[61]

Barley straw

Steam,enzyme

16.7

0.25

[61]

Wheat straw

Steam,enzyme

16.0

0.30

[61 ]

hemicellulose

The pulp reamins after extraction of juice from sugar beets represent about 24% of the dry weight of the beet root [58]. Fermentation of the enzymatic hydrolysate of beet pulp with Klebsiella pneumoniae resulted in butanediol yield of approximately 43% with ethanol as the major by-product. A combined enzymatic hydrolysis and fermentation (CHF) approach of wood and agricultural residues yields 40-68% greater butanediol than the values obtained with the conventional sequential hydrolysis and fermentation approach [60-62]. When xylan from aspenwood was used as a substrate in a CHF process, K. pneumoniae utilized all the released sugars to yield 0.28 g butanediol/g of reducing sugars utilized. The celluloserich water-insoluble fraction and hemicellulose-rich water-soluble fractions of wheat straw, barley straw and corn stovers can be readily utilized by K. pneumoniae. The fermentation

234

of sugars released from hemicellulose fraction of steam-exploded aspen appears to be dependent upon the method of hydrolysis employed. With acid hydrolysis, the relative production of butanediol, ethanol and acetic acid, expressed as the mole per cent of the total solvent production, was 36.2, 44.6 and 19.2%, respectively. The corresponding figures after enzymatic hydrolysis were 2.6, 14.7, and 82.7%, respectively. A marked shift to acetate production at the expense of butanediol was evident. Grover et al. [63] studied the production of 2,3 butanediol from wood hydrolysate by

K. pneumoniae. A butanediol concentrations of 12 g/I with the corresponding yield of 0.27 g/g sugars present in wood hydrolysate was achieved in 48 h. Addition of 1% (w/v) malt extract to the medium further enhanced butanediol production to 13.3 g/I (yield, 0.23 g/g). The optimal steaming duration at 240~ for solvent production from acid hydrolysed cellulose fraction of aspen was found to be 30 sec. Out of the 75% of the original wood, 62.2% was hexosan and 8% pentosan in nature. K. pneumoniaewas able to produce 20 g/I of butanediol from this substrate. Water prehydrolysate of southern pine could not be fermented efficiently unless the hydrolysate was treated with Amberlite IR-120, H§ or IR45, OH [37]. Spent sulfite liquor (35% hemicellulosic carbohydrates) is another potential substrate for bioconversion. Pretreatment of liquor prior to fermentation includes stripping of SO2 by combined boiling and aeration. Aeromonas hydrophila was found to accumulate 12.4 g/I (yield 0.33 g/g) of butanediol from 38 g/I initial sugar concentration [16].

6

FACTORS AFFECTING 2,3-BUTANEDIOL PRODUCTION

Many cultural, environmental and nutritional factors can affect the fermentative butanediol production because of the versatality of the metabolism of 2,3-butanediol producing facultative anaerobic bacteria.

235

6.1 pH

pH is a fundamental parameter in the regulation of bacterial metabolism, particularly in the synthesis of multiple end products. As a general rule, alkaline conditions favour the formation of organic acids, with a corresponding decline in the yields of such products as butanediol (Table 9). Generally, the yield of butanediol reaches maximum in the pH range 5.0-6.0 but falls to near zero above pH 7.0 [42]. The ratio of butanediol to acetoin may vary from near 25 between pH 5.2 and 6.0, to near zero at pH 7.6. Above pH 7.0, a rising formic acid concentration and falling CO 2 levels suggested that cells were able to maintain its NAD/NADH balance by reducing CO2 to formic acid when they cannot produce butanediol [42,54]. Acetic acid and lactic acid production is minimum below pH 5.0 but increases rapidly above pH 6.0 [65].

Table 9 Effect of pH on carbon balance of 2,3-butanediol fermentation [92] Product

Yield (g/100 g carbon) pH above 6.3

pH below 6.3

Butanediol

10.10

36.95

Acetoin

0.00

0.45

Ethanol

17.25

13.45

Acetic acid

32.75

1.00

Formic acid

21.85

2.85

Lactic acid

2.65

1.00

Total

87.90

55.65

Butanediol + acetoin

12.00

67.00

68.00

9.00

(% of total products) Organic acids (% of total products)

236

The pH optima also appears to be a function of the substrate used for fermentation. With glucose, optimum pH for butanediol production was 6.4, whereas it was between 5.2 and 5.4 with xylose as substrate [35].

6.2 Temperature

The strong dependence of enzymatic activity and cellular maintenance requirements upon temperature [66] makes the efficiency of bioprocess strictly temperature dependent. The temperature of fermentation affects butanediol production in a predictable manner. As the temperature is increased within the certain limits, the rate of fermentation is also increased [15]. However, this increased rate is accompanied by a decrease in the total yield of butanediol. Since butanediol fermentation is strongly exothermic, maintenance of the proper temperature is essential. Table 10 presents the optimum pH and temperature for butanediol producing organisms.

Table 10 Optimum pH and temperature of butanediol fermenting organisms Organism

Substrate

pH

Temperature

Reference

(oc) K. pneumoniae

Glucose

6.0

30

[44]

K. pneumoniae

Xylan

6.5

30

[64]

K. pneumoniae

Xylan

5.4

37

[37]

K. oxytoca

Xylose

5.2

37

[49]

A. aerogenes

Glucose

6.8

35

[45]

A. aerogenes

Sucrose

5.0-6.0

35-37

[54]

B. polymyxa

Xylose

25-35

[24]

237

With Kiebsiella pneumoniae, the optimum temperature for growth and substrate uptake rate was between 37-38~

[54,67,68]. Substrate utilization for endogenous

metabolism increased steadily as the temperature increased from 25-40~ Callow [54] obtained maximum butanediol production between 35 and 37~

Pirt and with K.

pneumoniae, whereas Olson and Johnson [44] obtained maximum yields of butanediol at 30~

6.3

Aeration

Oxygen supply rate is another factor which significantly affects the end product formation, even though 2,3-butanediol is a product of anaerobic metabolism [69]. Microbial species known to produce butanediol have been classified as facultative anaerobes, and as such can exist in the absence of oxygen. Butanediol production has been shown to be most efficient under aerobic conditions particularly when high substrate concentrations are used or during the fermentation of pentose sugars. In the absence of air, and with high substrate concentration, the growth and the metabolite production are practically non-existence [45]. Thus aeration does not only serve to agitate the medium, and to degas CO2 produced but it also plays an important role in the metabolism [70]. Limited amount of oxygen can increase the productivity as higher cell concentrations of oxygen can be maintained in chemostats. Higher level of oxygen can shift the ratio of end products from butanediol to acetoin [45]. The availability of oxygen also seems to determine the amounts of particular products excreted. For a high oxygen transfer rate (OTR) acetate is the principal product [71,72], when specific uptake rate decreases, butanediol becomes the predominant product, and when oxygen supply is cut off ethanol production increases at the expense of butanediol [42]. For every microorganism a critical dissolved oxygen (DO)level exists above which organism's respiration rate is independent of the oxygen concentration. The value of this critical DO is in the range of 0.003-0.05 mM/I for most of the organisms [73]. When the DO content of the medium exceeds the crtical level, the cell is saturated with oxygen and the process is said to be biochemically limited. If the reverse situation is observed and supply of oxygen does not meet the cellular demand, the system is said to be mass transfer limited. The critical oxygen tension for Klebsiella pneumoniae in a batch culture

238 was found to be in the range 8-15 mm Hg [74], whereas in continuous culture, a slightly lower range (2-10 mm Hg)is reported [75]. Table 11 summarizes the effect of oxygen supply on butanediol production.

Table 11 Effect of aeration on 2,3-butanediol fermentation Substrate

Aeration Oxygen supply

Yield (g/g) KLa (h 1)

Xylose

Xylose

Glucose

Mol/I/h

0.048

0.10

0.027

0.15

0.014

0.22

0.007

0.26

Finite

0.27

Anaerobic

0.04

Glucose

Reference

30

0.44

100

0.32

150

0.20

300

0.17

[49]

[39] [45]

Finite

0.21

[47]

Anaerobic

0.35

[39]

Klebsiella oxytoca exhibits exponential growth (l~=constant) under aerobic conditions and linear growth (dx/dt=constant) during oxygen limited conditions [49]. When oxygen limits growth (DO<3%), butanediol is secreted as a byproduct of the energy producing metabolism. The cell yield from substrate appears to be much lower when oxygen limits

239 growth because the ATP yield from the fermentative pathway is less than 10% of the ATP yield from the oxidative pathway, and because a large fraction of the substrate goes to butanediol [49]. During aerobic growth, the bacteria apparently obtain all the energy they need for growth and maintenance by oxidative metabolism. Regeneration of ATP and NAD is major function of molecular oxygen in the growth and maintenance of aerobic and facultative organisms [76]. Oxygen is the terminal acceptor in the production of ATP via oxidative phosphorylation. This compound subsequently provides the energy required in many of the reactions of cell synthesis, maintenance and product formation. The efficiency of producing ATP from xylose is very high when catabolism occurs via TCA cycle and the electron transport system. However, when DO falls to near zero, oxygen becomes in short supply at the terminal oxidase of electron transport system, which apparantly results in the limitation of the amount of NADH. To support further growth and to help maintain the NAD:NADH balance, K. oxytoca appears to produce 2,3-butanediol via a fermentative pathway during oxygen limited growth [49].

6.4 Water activity

Another important parameter which affects 2,3-butanediol production is water activity (~). It is an expression of the water concentration that depends on the molar concentration and activity coefficient of each solute. Increasing solute concentration decreases water activity of a solution [43]. It has been demonstrated that a number of key kinetic and bioenergetic parameters are influenced by the water activity [77]. The most important are duration of culture log phase, maximum specific growth rate, biomass to substrate yield coefficients, thermodynamic efficiency, and the maintenance energy coefficient. At a water activity of 0.985, growth rate of Klebsiella sp. was found to be 50% optimal and became lesser than 10% optimal at water activities below 0.975 [68]. Species of

Klebsiella (Aerobactor) are not as osmotolerant as some other organisms and that is the reason why very high sugar concentrations in butanediol process are not suitable. The problems associated with the bioconversion of complex substrates such as starch or wood hydrolysates may therefore result, in part, from the high solute concentrations of such feedstocks.

240

6.5

Inoculum

Inoculum size affects the initial fermentation rate of high xylose concentration without affecting butanediol production [39]. On the other hand, increasing the Bacillus polymyxa inoculum from 2.5 to 5% did not affect either the xylose consumption rate or the butanediol yield. Acclimatization of culture have been shown in many cases effective in improvement of both fermentation rates and butanediol yields (Table 12). Adaptation of

Klebsiella pneumoniaeto high substrate concentration, prior to inoculation has been found to be effective in dissimilation of 150 g/I of xylose with high butanediol yields [46]. Similar improvements were observed with acclimatized cultures of K. pneumoniae [21].

Table 12 Effect of culture acclimatization on 2,3-butanediol fermentation Organism

Bacillus polymyxa Klebsiella pneumoniae

Acclimatization

Productivity (g/I/h)

Reference

No

0.52

[24]

Yes

0.92

No

0.34

No a

0.73

Yes

1.04

[21]

Supplementation of malt sprout

6.6

Nutrient supplementation

For the growth and maintenance of microorganisms, culture media must contain all the essential nutrients. In addition to carbon and nitrogen sources, the medium may

241 include vitamins, trace metals etc., the selection of which may be determined by the nature of fermentation product. Certain nutrients and metallic cations have been found to improve the butanediol yields. Yeast extract has been found to stimulate butanediol yields. Laube et al. [24] obtained three-fold increase in the butanediol yields of Bacillus polymyxa. Similarly, yield of butanediol of Klebsiella pneumoniae was found to be improved by 51% by inclusion of yeast extract in a chemically defined medium [47]. Xylose bioconversion is facilitated by yeast extract even at low substrate concentration [24,27]. However, the cost of yeast extract prohibits the utilization of large quantities in commercial processes. Therefore, other media supplements have been examined in order to repalce yeast extract partly or completely. Addition of 1% urea to a defined medium resulted in about 25% increase in butanediol yield [47]. Proteose-peptone was found to be superior to yeast extract as a nitrogen source [78]. Trace metals have proven to be effective in nutrient supplement. Cationic species of AI, Ba, Cd, Cu, Cr, Fe, Mg, Mn, Mo, Ni, Pb, Ti, V, and Zn are present in yeast extract [79]. Supplementation of a medium containing 0.5% yeast extract to a final trace metal content equivalent to that of 1.5% extract resulted in improved butanediol yields [25]. The major stimulatory components of the trace metal mixtures were found to be Fe2+and Mn 2+ [80,81]. The Mn2+-induced enhancement of butanediol production is believed to be the result of induction of acetolactate decarboxylase enzyme [82]. However, a purified preparation of this enzyme from K. pneumoniae was not found to be stimulated in the presence of Mn 2+[83]. Phosphate has also been shown to stimulate butanediol production

[84,85]. Acetate induces the enzyme which catalyzes the breakdown of pyruvate to butanediol. Since acetate is both a metabolic product and an important medium supplement, it is of a special interest in the production of butanediol. In the presence of acetic acid K. pneumoniaecells were able to utilize more than 90% of the substrate with the yield of butanediol nearly tripled (Table 13). However, it appears to be species dependent, since no effect of acetate supplementaion on butanediol production of B.

polymyxa was observed [25].

242 Table 13 Effect of acetate supplementation on 2,3-butanediol production by Klebsiellapneumoniae Substrate

Acetate (%)

Yield (g/g)

Reference

Xylose

0

0.41

[90]

0.8

0.53

1.2

0.52

1.6

0.46

2.0

0.36

0

0.19

0.5

0.55

0

0.21

0.5

0.49

Xylose Glucose

6.7

[47] [47]

Inhibitors

Wood hydrolysate contains various components which are inhibitory to fermentative microorganisms [86-88]. Different groups of microbial inhibitors in acid-hydrolysed wood hydrolysate have been identified. Tran and Chambers [37] studied the inhibitory effects of lignin and extractive model compounds on butanediol production from mannose-rich southern pine water prehydrolysates by Klebsiella pneumoniae [37]. All tested lignin model compounds inhibited butanediol fermentation (Table 14). Protocatechuic acid and p-hydroxy benzoic acid were found to be the most toxic inhibitors, and vanilline was the weakest compound. Among the extractive model compounds linoleic acid exhibited the most adverse effect.

243 Table 14 Effect of inhibitory components (model compounds commonly found in wood hydrolysate) on 2,3-butanediol fermentation Model

Concentration Butanediol

compound

Time

(g/l)

(g/I)

(h)

None

0

29.7

46.5

Vanilline

0.030

24.3

66.5

p-Hydroxy benzoic acid

0.0430

9.4

98.3

Protocatecnic acid

0.052

9.4

45.0

Coniferyl alcohol

0.055

10.5

45.0

0

14.2

72.0

Reference

Lignin derivatives

[37]

Phenolics Syringeldehyde + Vanilline

1.0

10.7

72.0

2.5

5.6

120.0

5.0

2.9

72.0

Pinene

0.260

14.6

52.3

Limonene

0.251

13.4

95.0

Pyrogallol

0.235

28.9

121.3

Linoleic acid

0.245

10.0

75.0

Palmitic acid

0.233

14.9

71.0

Abietic acid

0.158

10.9

71.3

None

0

14.2

72.0

Furfural

1.0

12.4

72.0

2.0

12.5

72.0

0.5

13.9

120.0

1.0

12.9

48.0

[90]

Extractives [37]

Others

Sulfate

[90]

244 Nishikawa et al. [89] reported the similar results with four potential inhibitors, furfural, hydroxymethyl furfural, syringaldehyde and vanilline of aspenwood hydrolysate. Inhibition occurred in the range of 0.2 to 0.4 g/I of the inhibitor concentration. Combination of the individual materials resulted in a cumulative inhibitory effect. Prolonged incubation could alleviate some of the inhibition. Frazer and McCasky [90] studied the effect of acetate, sulfate, furfural, and phenolic compounds commonly found in hydrolysates. Phenolic compounds were the most potent inhibitors. Sulfate levels upto 0.2% (w/v) did not affect growth but reduced the butanediol yield by approximately 30%. When hydrolysate, treated with ion-exchange resins to remove phenolic compounds, was fermented with K. pneumoniae, butanediol yields exceeding 90% of theoretical were consistently achieved [91]. The production of 2,3-butanediol by bacterial species, first noted in the late ninteenth century and developed to pilot-scale in 1940s [92], continues to be of interest today mainly because of the diverse applications of this solvent. K. pneumoniae, with a broad environmental adapatability, and ability to convert all the sugars present in lignocellulosic hydrolysate into butanediol, is the most thoroughly studied organism. Osmotolerant strains resistant to inhibitors present in lignocellulosic hydrolysate may increase conversion rates. Improvement in reactor design, such as the two-stage continuous systems or the immobilized cell system, may enhance the efficiency of butanediol production.

7

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Scott WJ. Austr J Biol Sci 1953; 6: 549.

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Esener AA, Bol GA, Kossen NWF, Roels JA. Proc Int Ferment Symp 6th, 1981 ; 339.

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Mahmoud SAZ, Taha SM, EI-Sadek GM, Ali SA. Rev Microbiol 1975; 6: 77.

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81

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Sokatch JR, Bacterial Physiology and Metabolism, New York: Academic Press, 1969; 87.

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Loken JP, Stormer FC. Eur J Biochem 1970; 14: 133.

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9 M i c r o b i a l P r o d u c t i o n of O r g a n i c A c i d s

1

INTRODUCTION

in this chapter, microbial production of certain organic acids from pentoses and hydrolysates of natural lignocellulosic substrates is discussed. The demand of many of these acids is, however, low because of their very specialized fields of application and that they can be produced more economically by synthetic processes rather than by fermentations. This trend may be changed in the future due to rising cost of petrochemicals and the discovery of new applications.

2

ACETIC ACID

Acetic acid is presently produced from mineral oil or natural gas. It is an important feedstock for many chemicals such as vinyl acetate monomer, cellulose acetate, terephthalic acid, acetic acid esters and acetic anhydride [1]. Acetic acid, if obtained by fermentation of ethanol containing solutions, is called 'vinegar'. Acetic acid fermentation is an oxidative fermentation in which diluted solutions of ethanol are oxidized by bacteria to acetic acid and water. It is also possible to produce acetic acid by fermentation of carbohydrates such as glucose and xylose. In conventional vinegar production, glucose is fermented to ethanol by yeast and then ethanol oxidized by a bacterium to acetic acid. The theoretical maximum yield of this route is 2 moles of acetic acid from 1 mole of glucose or 0.67 g/g glucose. C6H1206

....... > 2C2HsOH + 2CO2

2C2HsOH + 202 ....... > 2CH3COOH + 2H20 In commercial practice, actual acetic acid yield is 0.5-0.55 g/g glucose or roughly 249

250 75-80% of theoretical [2,3]. This conversion route cannot utilize five-carbon (pentose) sugars because of the inability of the commercially available yeast, Saccharomyces

cerevisiae, to ferment pentoses to ethanol. Fermentation of pentose sugars would be important if the sugar feedstocks were obtained from hydrolysis of a lignocellulosic material. Interest in acetic acid fermentation of renewable lignocellulosic substrates has focussed on the development of an inexpensive process to produce acetic acid for subsequent use in calcium-magnesium acetate, an alternative to conventional road deicing salt [1,4].

2.1

Microorganisms used for acetic acid production

It has been demonstrated that Clostridium thermoaceticum is potentially useful for the conversion of lignocellulosic biomass because of its ability to ferment the major sugars found in the hemicellulose and cellulose fractions [1,5]. This organism was first isolated by Fontaine et al. [6]. This acetogenic, thermophilic bacteria is capable of converting glucose and xylose to acetic acid, as the only product, in near stoichiometric yields. However, it lacks polysaccharolytic enzymes to degrade lignocellulosic polysaccharides and can therefore only utilize these substrates after they have been hydrolyzed to their component sugars [1]. Fermentation with C. thermoaceticum offers a significant advantage in terms of acetic acid yield, compared to conventional vinegar fermentation. It produces 3 moles of acetic acid from 1 mole of glucose. In practice, 85% of the sugars may be converted to acetic acid and about 5% enters the cells. C. thermoaceticum can also ferment xylose and fructose. It can convert glucose and xylose almost stoichiometrically to acetic acid as follows: C6H1206 ....... > 3CH3COOH 2CsHloOs ....... > 5CH3COOH Schwartz and Keller [7] have reported the isolation of a strain of C. thermoaceticum capable of growing at pH 4.5. They also compared the performance of four other strains of C. thermoaceticum in a pH-controlled batch fermentation to determine the potential strain for improvement programme [8]. The characters determined at pH 6.0, the lowest

251 pH at which preliminary experiments showed all four strains could grow, were growth rate, tolerance to aetate, and efficiency of converting substrate to acetate and cell mass. Ljungdahl et al. [9] evaluated acetic acid production by C. thermoaceticum, C.

thermoautotrophicum and Acetogenium kivui. Both C. thermoautotrophicum [10] and A. kivui [11] have been identified as homoacetate fermenting thermophiles. The growth of A. kivui was found to be almost three times faster than other two clostridia. Filamentous fungi like Fusarium oxysporum [12] and Polyporus anceps [13] have also been reported to produce significant quantities of acetic acid from a range of substrates. However, the overall yield of acetic acid from fungi are relatively low.

2.2

Kinetics of product formation

Using Clostridium thermoaceticum ATCC 39073 [14], yield of acetic acid greater than 71% of theoretical could be obtained in 72 h from initial substrate concentration of 33 g/I xylose or less (Table 1). Maximum product concentration of 14.1 g/I was obtained when the initial xylose concentration was 20 g/l.

Table 1 Acetic acid production by Clostridium thermoaceticum ATCC 39073 Substrate

Reactor

Acetic acid

Yield

Reference

system

(g/I)

(g/g)

Xyiose

Batch

14.1

0.76

[ 14]

Xylose

Fed-batch

42.0

0.71

[14]

Glucose

Batch

13.1

0.67

[16]

Glucose

Fed-batch

56.1

0.85

[15]

252

As in the case of xylose, 20 g/I glucose has been reported to provide for maximum conversion to acetic acid [15]. Yields lesser than theoretical values of conversion may be due to the loss of carbohydrate to cell mass and thermal degradation over the course of fermentation. C. thermoaceticum grows exponentially for first 24 h, reaching a maximum specific growth rate of 0.14 per h. Acetic acid production continues until 48 h at which time over 90% of xylose had been consumed. In a fed-batch fermentation, maintaining xylose concentration between 10 and 20 g/I, a peak cell density is obtained at 30 h followed by a gradual decline until 116 h at which time no xylose is consumed [14]. Acetic acid production correlates closely to xylose consumption and reaches a final concentration of 42 g/I with a yield of 71%. In fed-batch fermenation of glucose, maintaining substrate concentration between 5 and 15 g/I, as much as 56 g/I acetic acid is produced in yields ranging from 67 to 85% [15]. In a continuous culture of C. thermoaceticum ATCC 39073, using automatic pH control, acetic acid production was found to be both growth and non-growth associated [16]. However, acetic acid yield of 2.3 mol/mol glucose consumed, were lower as compared to theoretical maximum value of 3. Continuous fermentation could be sustained for 1600 h or more without any contamination problems. Schwartz and Keller [8] compared 4 strains of C. thermoaceticum (Ljungdahl, Wood, $3 and 1745) in pH controlled batch fermentation for their efficiency of conversion of glucose to acetic acid. At pH 6 and initial acetic acid concentration of greater than 10 g/I, only two of the strains ($3 and 1745) grew. The maximum acetic acid production was 15 and 20 g/I at pH 6 and 7, respectively. C. thermoaceticum was found to be more sensitive to free acetic acid than to either acetate ion or pH. Wang et al. [17], using a different medium, reported production of 37 g/I of acetic acid at pH 7 using C. thermoaceticum (Wood). Maximum specific growth rate at pH 7 was 0.22 per h with acetate yield of 0.85 g/g glucose. An inverse relationship between both the growth rate and final cell concentration to sodium acetate concentration has also been obserevd [18].

Acetogenium

kivui

grows

faster

than

C.

thermoaceticum

and

C.

thermoautotrophicum, but is less tolerant of substrate and product concentration [9]. In a continuous culture with cell recycling system, productivities of 4.8, 4.0 and 3.8 g/I/h were obtained from A. kivui, C. thermoceticum and C. thermoautotrophicum, respectively. Hollow fiber system was found superior to Pellicon Casette system during cell recycling fermentation. With hollow fiber system, less cell lysis and less cell clogging occurred. Clogging could be prevented by reversing the flow through the system after every 12 h. Kumar et al. [19] screened various strains of Fusarium oxysporum and Monilia

253

brunnae for their ability to ferment glucose, xylose and avicel. F. oxysporum DSM 841 was selected as potential strain producing acetic acid and ethanol as major products under semiaerobic conditions. Almost equal amounts of ethanol and acetic acid are produced by this organism [20]. The product yields vary from 0.23 to 0.27 g/g for acetic acid and 0.18 to 0.22 g/g for ethanol. With cellulosic substrates, acetic acid was always the major product [21]. Singh et al. [22] obtained 12 g/I acetic acid (0.36 g/g substrate) from potato wastes using a cellulolytic strain of Fusarium oxysporum DSM 841 in batch culture. Production of acetic acid improved significantly in fed batch fermentation [21].

Polyporus anceps is capable of utilizing a range of substrates to produce acetic acid as the major product [13]. Acetic acid yields from this organism range from 0.1 to 0.2 g/g substrate. However, this organism is unable to utilize xylose as sole carbon source. Table 2 shows some results on acetic acid production by filamentous fungi.

Table 2 Acetic acid production from different substrates by filamentous fungi Organism

Substrate

Acetic acid yield (g/g)

Reference

Fusarium

Xylose

0.27

[22]

Glucose

0.23

Avicel

0.24

Arabinose

0.19

Galactose

0.22

Glucose + xylose (1:1 )

0.26

Potato wastes

0.36

[21]

Polyporus

Glucose

0.16

[ 13]

anceps

Sucrose

0.20

Fructose

0.12

Starch

0.10

oxysporum

254 Brownell and Nakas [14] carried out fermentation of carbohydrates released from pretreated oat spelt xylan and hemicellulose isolated from hybrid poplar by Clostridium thermoaceticum. During a 72 h fermentation, in which the initial pentose concentration

was adjusted to 20 g/I, concentration of acetic acid reached to 14.4 g/I on oat spelt xylan, whereas on poplar hemicellulose hydrolysate its concentration reached only 11.5 g/I (Table 3). C. thermoaceticum could consume 89 to 94% of available sugars.

Table 3 Fermentation of acid-treated oat spelt xylan and poplar hemicellulose to acetic acid by Clostridium thermoaceticum ATCC 39073 a [14]

Treatment

Oat spelt xylan

Popalar hemicellulose

(% H2SO4) Pentose

Pentose

released consumed (g/I)

(%)

Acetic

Pentose

acid

released consumed

acid

(g/I)

(%)

(g/I)

11.9

(g/I)

Pentose

Acetic

1.0

16.5

3.0

1.5

28.3

89

12.1

6.0

92

2.0

34.8

92

13.4

8.9

91

2.5

42.8

90

14.4

12.6

87

11.3

3.0

41.0

90

11.2

13.0

87

11.3

14.0

89

11.5

4.0

a

11.8

Acid treatments (solid:liquid ratio, 1:20) were carried out at 100~ for 60 min; initial

pentose concentration for fermentation, 20 g/I

Aeration rate has been found to affect significantly the acetic acid production from potato wastes by F. oxysporum DSM 841 [22]. For these experiments, fermentations were initiated with aeration rate of 0.5 vvm in aerobic phase for 72 h, thereafter brought down

255

in the range of 0.02 vvm to 0.1 vvm. The increase in aeration rate from 0.02 to 0.1 vvm significantly decreased acetic acid yields from 0.28 g/g to 0.09 g/g substrate (Table 4).

Table 4 Effect of aeration on acetic acid production from potato waste by Fusarium oxysporum DSM 841 [22] Aeration rate

Acetic acid

Yield

(vvm)

(g/I)

(g/g)

0.02

5.6

0.28

0.04

4.7

0.24

0.06

2.8

0.14

0.10

1.7

0.09

3

LACTIC ACID

Lactic acid occurs widely in nature, being found in man, animals, plants and microorganisms. It was discovered in 1780 by a Swedish chemist, Scheele, in sour milk, and Blondeau in 1847 recognized lactic acid as the final product a fermentation process [23]. The original application of lactic acid produced by microorganisms was its use as a preservative. Because of its pleasent acidic flavour and its preservative properties, lactic acid is extensively used in food industries. The acid is employed as an additive to soft drinks, essence, extracts, fruit juices, jams and syrups. Salts of lactic acid (Ca and Fe salts) have therapeutic uses.

256

3.1

Microoganism used for lactic acid production

The most important lactic acid producers belong to the family Lactobacillaceae [24] and are differentiated into various genera (Table 5). Lactic acid bacteria appear morphologically as diplococci, tetracocci, streptococci and as rods, which may be present singly or in chain. They are fastidious organisms. For normal growth, they require, besides a carbon source, nitrogen partly in the form of amino acids, several vitamins, growth substances, and minerals [25]. Twenty two species of Lactobacillus are described in Bergey's Manual, however, only three are capable of fermenting xylose to lactic acid. Of these, only Lactobacillus xylans is homofermentative [26]. Ishizaki et al. [27,28] isolated and characterized a homolactic producing strain, Lactococcus lactis IO-1 (JCM 7638). The overall stoichiometry for glucose and xylose conversion is C6H1206 ........ > 2CH3-CHOH-COOH CsHloOs ........ > CH3-CHOH-COOH + CH3COOH Small amounts of CO2 and acids may also be produced. Two moles of ATP are formed per mole of xylose or glucose consumed [29]. Alternatively, ethanol may be produced [30,31], but one less mole of ATP per mole of xylose is formed utilizing this route.

Table 5 Lactic acid bacteria and type of fermentation Genus

Type of fermentation

Configuration of lactic acid

Streptococcus

Homolactic

L(+)

Pediococcus

Homolactic

D L, L (+)

Lactobacillus

Homolactic

D(-), L(+), DL

Lactococcus

Homolactic

L(+)

Leuconostoc

Heterolactic

D(-)

257 3.2

Kinetics of product formation

Tyree et al. [29] studied the fermentation kinetics of glucose and xylose using

Lactobacillus xylosus. The maximum concentration of glucose and xylose which L. xylosus could utilize were 54 g/I and 31 g/I, respectively. L. xylosus was able to utilize glucose completely in 12 h, whereas 88% of xylose was utilized in 54 h. With xylose as substrate, the maximum cell and lactate concentrations produced were 1.3 g/I and 13 g/I, respectively (Table 6). The maximum specific growth rate of L. xylosus on glucose was 0.72 per h which was about 20% higher than the value reported for L. casei [32]. The product yield on xylose as substrate was initially 0.34 g/g, but increased as the specific growth rate declined so that consumption of 30 g/I xylose gave an overall yield of 0.41 g/g which corresponds to 0.69 moles of lactic acid per mole of xylose. Acetic acid and ethanol were also detected in xylose fermentations. L. xylosus could not utilize xylose in the presence of 5 g/I or more glucose.

Table 6 Lactic acid fermentation kinetics of Lactobacillus xylosus and Lactococcus lactis Organism

Substrate

Substrate

,/Llma , x

Lactic

Yield

consumption

(h 1)

acid

(g/g)

(%)

L. xylosus

L. lactis

Reference

(g/I)

Xylose

87

0.59

13

0.41

[29]

Glucose

100

0.72

34

0.88

[29]

Xylose

100

0.72

24

0.47

[33]

Glucose/

100

26

0.66

[33]

44

0.88

[34]

xylose (1:1 ) Glucose

100

1.25

258

Ishizaki et al [33] evaluated lactate production from xylose using Lactococcus lactis IO-1. Optimum pH and temperature for the growth and lactic acid productivity of L. lactis were 6 and 37~ respectively. Glucose-grown inoculum gave rise to a lag phase but the subsequent utilization of xylose and growth rate were similar to the fermentation with xylose-grown inoculum. Maximum yield of lactate from xylose was 0.47 g/g, whereas a yield of 0.88 g/g has been reported with L. lactis grown on glucose [34]. In a mixture of glucose and xylose, glucose was utilized with higher rate in the early stages of growth, thereafter both the sugars were utilized simultaneously.

4

CITRIC ACID

Citric acid is mainly used in the food industry with worldwide acceptability as a safe food ingredient. Due to its pleasent acid taste and high water solubility, the most extensive application is in beverages, jams and jellies as well as in sweets. Another important area of application is in pharmaceutical industry and in cosmetics. Wehemer, in 1893, was first to observe the presence of citric acid as a by-product of calcium oxalate produced by Peniciilium glaucum [35]. However, Currie [36] opened the way for the industrial production of citric acid.

4.1

Microorganismused for citric acid production

Citric acid fermentation is extremely complex, and successful process depends both on an appropriate strain and optimal fermentation parameters. Citric acid production in varying amount is fairly common among members of the genera of Aspergillus and

Penicillium. Aspergillus niger is very well known and the most studied organism with regard to citric acid production. Some other Aspergillus species known for citric acid production are: A. awamori, A. wentii, A. saitoL A. clavatus, A. fenicis and A. fonsecaeus [37]. Other moulds observed as producing relatively large amounts of citric acid are

Botrytis cineria, Mucor piriformis and Trichoderma viride.

259 With regard to yeasts, genus Candida has almost exclusively been studied. C.

lipolytica, C. tropicalis, C. guillermondii, C. parapsilosis and C. intermedia have been investigated for citric acid production. Some coryneform bacteria (Arthrobacter sp.), and related organisms (Actinomycetes) produce substantial amounts of citric acid.

4.2

Kinetics of product formation

In general, only sugars which are rapidly taken up by the organisms are useful carbon sources for citric acid fermentation. In most cases sucrose and molasses are used, but glucose (from hydrolysis of polysaccharides) or fructose have also been used. Although a lot of work has been done on fermentation of hemicellulose sugars to ethanol, acetone-butanol, 2,3-butanediol, very little information is available on production of citric acid from pentose sugars. Maddox et al. [38] investigated the ability of one strain each Aspergillus niger and Saccharomyces lipolytica for their ability to produce citric acid from sugars present in hemicellulose hydrolysate. Mannose was found to be as favourable a substrate as glucose for S. lipolytica, but neither of the pentose sugars nor galactose were metabolized by the yeast (Table 7). A. niger could assimilate both mannose and xylose readily with reasonable citric acid yield. Although A. niger readily assimilated galactose, little amount of citric acid was produced [39]. A. niger was able to utilize the mixture of sugars simultaneously, but glucose was used at a faster rate than mannose, xylose and arabinose; mannose faster than xylose; and xylose faster than arabinose. Sugar source has a marked effect on citric acid fermentation. Presence of galactose or a product of galactose metabolism has been shown to cause inhibition of citric acid production, and also glucose utilization [40]. Supplementation of methanol in galactose containing medium considerably improved the citric acid production (Table 8).

260 Table 7 Citric acid production from different organisms Organism

Aspergillus niger

Substrate

Citric acid

Yield

(g/I)

(g/g)

Xylose

22

0.31

Glucose

27

0.45

Mannose

17

0.23

Arabinose

6

0.13

Glucose/

23

0.35

11

0.17

17

0.35

Xylose/ mannose

17

0.28

Xylose/ arabinose

5

0.07

Galactose

0

0

Reference

[38]

xylose Glucose/ arabinose Glucose/ mannose

Aspergillus niger

Saccharomyces lipolytica Candida guillermondii

Sucrose

53

0.48

Fructose

23

0.25

Lactose

5

0.07

Glucose

6

0.30

Mannose

9

0.41

Xylose

0

0

Glucose

5

0.12

[39]

[39]

[77]

261 Table 8 Effect of methanol supplementation on citric acid production from galactose by different strains of Aspergillus niger [41] Strain

Control

1% Methanol

Citric acid

Yield

Citric acid

Yield

(g/I)

(g/g)

(g/I)

(g/g)

ATCC 12846

0.2

0.003

20.0

0.31

ATCC 26036

0.3

0.004

25.0

0.44

ATCC 26550

0.1

0.001

20.5

0.27

MH 15-15

0

0

12.5

0.21

IMI 83856

0.2

0.015

2.0

0.20

5

PROPIONIC ACID

Propionic acid is derived from succinate which is formed via Embden Meyerhof pathway and oxaloacetate [42] mechanism of conversion of glucose by Propionibacterium has been shown to yield two moles of propionate and one mole of acetate from 1.5 moles of glucose [43].

5.1

Microorganisms used for propionic acid production

Several species of genus Propionibacterium are known to produce propionic acid.

P. frendenreichii and P. jensenii can utilize only monosaccharides, whereas P. technicum can utilize starch, dextrin and glycogen [24]. P. acidi-propionici produces high acid

262 concentrations from both glucose and xylose [44]. Clostridium propionicum is another propionic acid producing organism but it cannot decarboxylate succinate [45]. In this case lactic acid is the intermediate, which is metabolized to propionate. The yield in acid fermentation with P. arabinosum is about 75 to 85%, consisting of 2 parts of propionic acid and 1 part of acetic acid [23].

5.2

Kinetics of product formation

Claussen and Gaddy [43] and Claussen et al. [46] investigated the kinetics of propionic acid formation of a synthetic glucose-xylose mixture, having the same concentration as the lignocellulosic hydrolysates, using Propionibacterium acidi-propionici. The purpose of their study was to examine the economic potential of producing propionic and acetic acids from orchard grass. Over a range of retention time studied, cell densities ranged from 2-5.5X10 ~ cells/ml, remaining constant for retention times greater than 10 h. Both glucose and xylose consumption increased rapidly until a retention time of 40 h, and then increased slowly. The conversion of glucose and xylose reached 87% and 83%, respectively, at retention time of 78 h. The yields of organic acids remained relatively constant at 2 moles propionic acid per mol acetic acid. The ratio of conversion of glucose to xylose was constant for all three retention times which indicates that there was no preference for glucose over xylose.

6

ITACONIC ACID

Itaconic acid (methyl succinic acid) is an important intermediate in polymer production and for the manufacturing of polyester resins or N-substituted pyrrolidones [47]. It may also be used in styrene butadiene copolymers, acryl nitrile copolymers for synthetic fiber manufacture, lattice and emulsions. The most important use of styrene copolymers is in paper coating and carpet backing.

263

6.1

Microorganisms used for itaconic acid production

This metabolic product was discovered by Kinoshita [48] in a culture of Aspergillus. It can also be produced by Aspergillus niger, Aspergillus terreus, Ustilago zeae [49], and by a number of yeast species [50]. Moyer and Coghill [51] and Lockwood and Reeves [52] screened a large number of strains discovering good producers A. terreus NRRC 265 and NRRC 1960. Today itaconic acid is produced by strains of A. terreus either by submerged (in the U.S.A. and Japan) or by surface fermentations (in the C.I.S.). In a semi-pilot plant scale, 45 to 55% yield of itaconic acid from 6% glucose has been obtained [53]. In pilot-scale of 300- 600 gallons, a maximum yield of 64.2% from 6.15% initial glucose has been achieved [54].

6.2

Kinetics of product formation

Itaconic acid can be successfully produced from various carbohydrate sources such as glucose, xylose, sucrose, beet molasses, cane molasses, and wood hydrolysate. The biosynthesis follows Embden Meyerhof pathway with the subsequent decarboxylation of the formed cis-aconitate. For itaconic acid production, pH of the medium is very important. At pH 2.1 all the glucose is metabolized to itaconic acid, but at pH 6.0 other acids are formed instead. Horitsu et al. [55] carried out itaconic acid production in a replacement-batch 400 ml glass reactor and in continuous fermentations with a maximum itaconic acid yield of 60 mg/h. In a continuous culture of immobilized A. terreus NRRL 1960, the highest itaconic acid concentration and productivity were 18.2 mg/I and 0.73 g/I/h, respectively [56]. Kautola et al. [57] investigated itaconic acid production from sucrose by immobilized A. terreus TKK 200-5-3 mycelia. A maximum itaconic acid concentration of 13.3 g/I was obtained from initial 100 g/I of sucrose. Itaconic acid production with immobilized mycelia was about two-times higher than that obtained with free mycelia. Table 9 shows itaconic acid production by immobilized cells of A. terreus in different reactor systems.

264 Table 9 Itaconic acid production by immobilized cells of Aspergillus terreus Reactor

Substrate Itaconic

Yield

Productivity

acid (g/I)

(g/g)

(g/I/h)

Reference

Batch

Xylose

30.0

0.45

0.20

[58]

Batch

Glucose

30.0

0.55

0.32

[58]

Batch

Glucose

39.4

0.64

[54]

Batch

Glucose

51.0

0.51

[78]

Batch

Sucrose

13.3

0.13

0.04

[57]

Repeated batch

Xylose

13.8

0.23

0.12

[58]

Repeated batch

Xylose

1.6

Repeated batch

Sucrose

9.7

Continuous

Xylose

5.0

Continuous

Glucose

18.2

Continuous

Xylose

8.1

[59] 0.03

[59]

0.08

0.56

[58]

0.30

0.73

[56]

0.07

[59]

Itaconic acid production from xylose and glucose by A. terreus NRRC1960 has been compared in batch, repeated batch and continuous column reactor [58]. In these experiments, A. terreus spores were entrapped in alginate gel beads or alternatively the fungal mycelia were immobilized either on Celite R-626 or in agar gel cubes. The highest itaconic acid yields obtained in a submerged culture batch fermentation was 54.5% based on total intial glucose (55 g/I) with volumetric productivity of 0.32 g/I/h, and 44.8% from xylose (67 g/I) with a productivity of 0.2 g/I/h. In a repeated batch fermentation, mycelia immobilized in agar gel had a productivity of 0.12 g/I/h. With the best immobilized system used, employing Celite R-626 as a carrier, volumetric productivities of 1.2 g/I/h from glucose and 0.56 g/I/h from xylose (both at 60 g/I substrate concentration) were obtained in continuous column operation for more than two weeks.

265

Itaconic acid production from xylose by immobilized A. terreus TKK 200-5-2 mycelia has been optimized both in repeated batch and continuous column bioreactor using statistical experimental design and empirical modelling [59]. In air-lift bioreactor, itaconic acid concentration and productivity were 2.54 g/I and 22.8 mg/I/h, respectively. However, packed-bed column reactors with a productivity of 51.7 mg/i/h at 50 g/I xylose concentration were found more suitable for itaconic acid production [60]. Itaconic acid production in different reactor systems is presented in Table 10.

Table 10 Itaconic acid production by Aspergillus terreus in different bioreactor systems Reference

Bioreactor

Itaconic

Productivity

system

acid (g/I)

(g/I/h)

Batch culture in disk bireactor

20.5

0.17

[56]

Continuous culture in disk bioreactor

18.2

0.73

[56]

Continuous culture by conventional method

7.8

0.31

[56]

Continuous immobilized cell culture

15.0

0.60

[56]

Air-lift bioreactor

2.5

0.02

[59]

Packed-bed column bioreactor

8.1

0.07

[59]

Continuous packed-bed column reactor

5.0

0.56

[58]

A. terreus needs magnesium for itaconic acid production both in growth and production medium, in order to improve acid tolerance and production [61]. The use of

266

an alkaline earth metal ion such as calcium together with copper or zinc has been found to increase itaconic acid yield about three-fold compared to that obtained without copper or zinc [62,63]. Calcium chloride at 0.3 g/I with 0.004 g/I of copper sulfate increases the itaconic acid production. It inhibits the enzyme itaconic acid oxidase, which is involved in the production of itatartaric acid, thus more substrate is available to itaconic acid [64]. Kautola et al. [65] studied the effect of trace and alkaline metals on itaconic acid production by polyurethane foam-immobilized A. terreus in repeated shake flask cultures. An increase in the copper substrate concentration increased the need for earth alkaline metals. The immobilization of mycelium increased itaconic acid production by as much as eight-fold.

7

FUMARIC ACID

Fumaric acid is used in plastic industry and to some extent in the food industry. It is also used in the manufacture of sizing resins for the paper industry. The demand for fumaric acid was increased sharply in the early 1960s. However, it was later possible to produce fumaric acid at a lower cost by catalytic oxidation of benzene, which replaced the fermentation process completely. The rising cost of petrochemicals and increasing interest in renewable resources revived the biotechnological production of fumaric acid [49].

7.1

Microorganism used for fumaric acid production

Biosynthesis of fumaric acid does not follow the regular glycolytic pathway. It is assumed that carbon dioxide fixation is of great importance in this process [49]. Rhizopus spp. accumulates fumaric acid in considerable amounts. Some species of Mucor,

Circinella, Cunninghamella, Aspergillus and Candida also produce fumaric acid. Carbohydrates are generally used as raw material.

267 7.2

Kinetics of product formation

Kautola and Linko [66] studied fumaric acid production from xylose by immobilized cells of Rhizopus arrhizus TKK 204-1 - 1a. Highest fumaric acid concentration reached with immobilized cells was 16.4 g/I with volumetric productivity of 67 mg/I/h at 100 g/I initial xylose, a C:N ratio of 160 and residence time of 10.25 days (Table 11). The volumetric productivity increased to 87 mg/I/h when residence time during the batch was 1.75 days.

Table 11 Fumaric acid production from xylose by Rhizopus arrhizus [66] Culture

Residence

C:N

Xylose

Fumaric

Productivity

time

ratio

(g/I)

acid

(mg/I/h)

(days)

(g/I)

Batch immobilized

6

157

60

10.0

69

Repeated batch

10.25

160

100

16.4

67

9

188

65

15.3

71

6

160

50

9.6

66

3

188

65

4.1

57

1.75

160

100

3.6

87

6

160

100

4.3

30

immobilized

Repeated batch free cells

About 3.5-times higher fumaric acid levels were obtained with immobilized R.

arrhizus system when compared to similar free cell system. Higher residence time also resulted in high product concentration and yield. However, high residence time is undesirable, resulting in decreased fumaric acid yields when the organism begins to

268

metabolize fuamric acid [67]. In repeated batch fermentations of 50 g/I and 100 g/I xylose using immobilized cells, nearly the same fumaric acid concentrations are obtained.

8

MIXED ACID FERMENTATION

Many bacteria produce a variety of acids during fermentation of pentoses, pentitols and polysaccharides (Table 12). The major products of these fermentations are ethanol CO2, acetic, lactic, succinic and formic acids. The relative amounts of acidic products formed depends on the type of organism, substrate and process conditions. In some cases formic acid may be replaced by an equivalent amounts of H2 and CO 2 [68].

Table 12 Organisms capable of mixed organic acid fermentation of pentose sugars Organism

Acetic

Lactic

Succinic

Formic

acid

acid

acid

acid

+

+

+

Escherichia coil Bacillus macerans

+

+

-

+

Clostridium thermocellum

+

+

+

+

Lactobacillus casei

+

+

-

+

Leuconostoc spp.

+

+

-

+

Ruminococcus albus

+

+

+

+

Spirochaeta stenostrepta

-I-

+

Facultative anaerobic bacteria, Escherichia coil can carry out mixed acid fermentation of glucose and xylose [69]. Acidic conditions were found to favour the production of

269 H2 and CO2 at the expense of formic acid. Spirochaeta stenostrepta Z1 ferments xylose, arabinose and ribose in addition to several hexoses [70,71] to acetic acid, lactic acid, ethanol, H2 and CO 2. Ruminococcus albus, a strict anaerobe, ferments cellulose and xylan to produce ethanol, acetic acid, formic acid as major products and small amounts of succinic and lactic acid [68].

Clostridium thermocellum and Clostridium thermocelluloceum also ferment cellulose, xylose and arabinose to ethanol and mixed acids like formic, lactic, acetic and succinic acid [72]. C. acetobutylicum carries out organic acid and solvent fermentation under different environmental conditions. Bacillus macerans (Bacillus acetoethylicum) conducts a modified mixed acid fermentation, of hexose and pentoses with ethanol, acetic acid and acetone as major products, and lactic and formic acids as minor products [68]. Species of Leuconostoc and Lactobacillus degrade pentose sugars through heterolactic pathway [73]. The product yields depend on specific growth conditions. A heterolactic bacterium, Thermoanaerobium brockii, was found to produce lactate as major product in high yeast extract medium and ethanol as major product in low yeast extract medium [74]. Datta [75] carried out anaerobic acidogenic fermentation of complex lignocellulosic biomass using a non-sterile mixed culture fermentation process. Acetic acid equivalents were produced at an average yield of 83.7%. During acidogenic fermentation of corn stover, all major components (cellulose, hemicellulose, lignin and pectins) were utilized. This simple fermentation scheme (non-sterile, anaerobic, unstirred, 25~ could produce a single class of useful compounds (C2-C6 volatile organic acids) directly from a complex lignocellulosic feed with high yield and specificity. Bioconversion of rice straw using mono and coculture of Fibrobacter succinogens and Treponema sp., isolated from the rumen of a sika deer, has been studied. The dry matter loss was quite high (52.1%) when a coculture was employed. The amounts of acetic, succinic and formic acids were also significantly higher for coculture when compared with that of monocultures [76].

9

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10 Microbial Production of Xylitol

1

INTRODUCTION

Xylitol, a pentavalent alcohol (pentitol) of xylose, is a common intermediary product of carbohyrates in microorganisms, human beings and animals, and is also found in many fruits and vegetables [1]. However, the concentration of xylitol is relatively low in plant sources which makes the extraction process difficult and uneconomical. Xylitol finds increased use as a sweetener in foods due to its higher sweetening power and greater negative heat of solution than other common polyols. It is 2.5-times as sweet as mannitol and has 2-times higher sweetening power than D-sorbitol [2]. Its sweetness compared to sucrose may vary from 0.85 to 1.25 depending upon pH, concentration, temperature, salt concentration and other factors [1]. A 10% solution of xylitol equals in sweetness to a 10% solution of sucrose. If sucrose is arbitrarily given a sweetening power rating of 100, the relative powers of other sweetening agents are as follows: xylitol 85-120, fructose 150, glucose 70, xylose 67, sorbitol 50, and mannitoi 40. Xylitol is an excellent, readily available source of calories- 1 g of xylitol corresponds to 4.06 kcal, similar to other completely metabolized carbohydrates. Xylitol does not need insulin for its digestion, therefore, it may be used clinically as a sugar substitute for diabetic patients or of glucose-6-phosphate dehydrogenase deficient population [3,4]. In addition to being a good anticarcinogenic sweetener, xylitoi is not utilized by microorganisms, therefore, products with xylitol are usually safe from microbial attack. It is not metabolized by the specific microorganisms (streptococci) normally present in the flora of the mouth and hence can be used for protection against dental caries. Various studies have shown about 30% reduction in dental caries in rats and humans on sorbitol and mannitol diets and virtually complete elimination of caries on xylitol diets [5,6]. Commercially xylitol is produced through chemical reduction of xylose derived from hemicellulosic hydrolysates of wood pulp or other xylose-rich materials. Since the hemicellulose fraction of these raw materials contains considerable amounts of polymers of other sugars, the process includes extensive purification and separation steps to remove these by-products from xylose or xylitol [7]. The recovered yield of xylitol is about 273

274

50-60% and xylitol production is relatively expensive. Xylitol can also be produced in high yields by fermentative and biocatalytic processes using microorganisms. Biological route for xylitol production has received much attention during last two decades. Several bacteria and yeast species have been discovered which secrete xylitol extracellularly as a metabolic by-product of ethanol or as the major product from D-xylose. In microorganisms, xylitol is formed as a metabolic intermediary product of D-xylose in two ways: D-xylose is directly converted to xylitol by NADPH-dependent xylose reductase (EC 1.1.1.21), or is first isomerized to D-xylulose by D-xylose isomerase (EC 5.3.1.5) and then reduced to xylitol by NADH-dependent xylitol dehydrogenase (EC 1.1.1.9) [8,9]. Many yeasts have NADP-dependent xylose reductase that catalyses the reduction of D-xylose to xylitol [10]. The enzymatic production of xylitol from xylose using xy!ose reductase from resting cells of Candida pelliculosa coupled with F42o-NADP oxidoreductase of Methanobacterium sp. has also been demonstrated [11]. For industrial production of xylitol, it is very important to obtain high xylitol production. Xylitol yield more than 90% of theoretical has been obtained in many cases but the productivities are low (0.32-1.67 g/I/h). For an economically feasible process, it is important to increase the production rate. This chapter describes the fermentative production of xylitol by different microorganisms and cultural variables affecting the process.

2

MICROORGANISMS FOR XYLITOL PRODUCTION

Under aerobic conditions many yeasts, especially the osmophilic yeasts, possess high potential to produce polyhydric alcohols as metabolic by-products from either glucose or xylose [12-18]. Glycerol, erythritol, D-arabitoi and mannitol are the common polyols produced by yeasts from D-glucose. Xylitol and L-arabitol are produced from D-xylose and L-arabinose, respectively. Pentitol production is observed frequently when yeasts are grown with pentoses as carbon and energy source. Xylitol can also be produced from Dxylulose by yeasts. The type and amount of polyols produced are affected by many environmental and nutritional factors. While many yeast species produce xylitol in different degrees as a by-product of xylose metabolism, the level of its formation is low [19].

275 However, several organisms have been identified that produce xylitol in substantial quantities. During last three decades, a number of laboratories have demonstrated xylitol production from pentose sugars using different species of yeast and bacteria. Microorganisms differ in their ability to grow and metabolize aldopentoses. The demonstration of product accumulation is sensitive to a number of experimental conditions which is reflected in the different views regarding the feasibility of pentose fermentation [20]. Onishi and Suzuki [21] surveyed 58 strains from various genera of yeast viz.

Saccharomyces, Debaryomyces, Pichia, Hansenula, Candida, Torulopsis, Kloeckera, Trichosporon, Cryptococcus, Rhodotorula, Monilia and Torulafor pentitol production using D-xylose, L-arabinose and D-ribose. Species of Candida were found to be the most promising, dissimilating all three pentoses and producing corresponding pentitols with the yield of 30-40% of the sugars consumed. Pichia miso, a highly osmophilic yeast isolated from miso paste [22,23], was able to dissimilate glucose to glycerol, erythritol and Darabinitol and D-xylose to xylitol, meso-glycero-ido-heptitol and D-glycero-D-ido-heptitol [24]. Gong et al. [25] screened yeast species belonging to the genera Saccharomyces,

Candida, Rhodosporidium, Saccharomycopsis, Trichosporon, Rhodotorula, baker's yeast and a respiratory deficient mutant strain of Saccharomyces cerevisiae. Significant quantities of xylitol were produced by all the yeasts except for red yeast such as

Rhodotorula. Xylitol is almost always produced during oxygen-limited or anaerobic fermentation of xylose [26-28]. McKracken and Gong [29] isolated several thermotolerant yeast strains of Candida sp., Hansenula polymorpha and Torulopsis hansenfi from sugarcane compost piles. Ethanol was the major product with glucose as carbon source, whereas xylitol was obtained as major product from xylose in the presence of D-xylose isomerase. Strains of Candida sp. were found to be better xylitol producers. Significant quantity of xylitol is produced during xylose fermentation to ethanol by

Pachysolen tannophilus [30,31]. Under aerobic conditions, most of the yeasts produce xylitol from xylose. S. cerevisiae can not utilize xylose fermentatively, but produces significant quantities of xylitol from xylose under aerobic conditions [32]. Both xylitol and ethanol are the major products of Kluyveromyces cellobiovorusferementation [33]. Nishio et al. [34] studied the continuous bioconversion of xylose into xylitol by co-immobilized cells of Candida pelliculosa and Methanobacteriumsp. A methanol yeast, Candida boindii also produces high yields of xylitol and xylulose [35]. Under microaerobic conditions, C.

guillermondii produces 0.63 g xylitol/g xylose consumed [36].

276 Some bacterial strains have also been reported to produce significant quantities of xylitol. A thiamine-requiring strain of Corynebacterium sp. grow rapidly on gluconate but not other carbohydrates [37]. However, the resting cells previously grown on a gluconatecontaining medium could utilize xylose, arabinose and ribose to produce corresponding pentitol [38]. A xylose-utilizing bacterial strain, isolated from soil, Enterobacter

liquefaciens, was found to accumulate xylitol extracellularly [39]. Izumori and Tuzaki [40] screened various strains of Mycobacterium for the production of xylitol from D-xylulose. M. smegmatis produced significant quantities of xylitol under aerobic or anaerobic conditions. Studies on xylitol production by filamentous fungi are very limited. Petromyces

albertensis accumulates large quantities of xylitol in the medium containing xylose as the sole carbon source [41]. Xylitol production by P. albertensis was found to be affected by different nutritional and cultural factors.

3

KINETICS OF GROWTH AND PRODUCT FORMATION

3.1

Yeasts

Dissimilation of xylose, arabinose and ribose for the production of pentitols by different species of 58 yeast strains was investigated by Onishi and Suzuki [21]. Among them Candida polymorpha, C. tropicalis, C. guillermondii, Pichia miso, Hansenula

subpeiliculosa, H. anamola, Torulopsis famata and Monilia sitophila produced xylitol at good yields of 35-45% of D-xylose consumed. Utilization of L-arabinose by yeast, in general, was poorer than that of D-xylulose. C. arborea, C. polymorpha, Torulopsis

halophila and Debaryomyces miso were found to be the best polyol producers from Larabinose. Only two strains of C. polymorpha and T. famata were able to utilize D-ribose for polyol production. The salt-tolerant strain of C. polymorpha could dissimilate all three pentoses with production of polyols at higher yields. Table 1 shows the potential yeast strains for the production of xylitol from D-xylose.

277

Table 1 Xylitol

production

Species

from

D-xylose

Xylitol yield (g/g)

by

Productivity

different Reference

(g/I/h)

C. polymorpha

0.39

0.23

[21]

C. arborea

0.28

0.17

[21]

C. guillermondii

0.75

0.54

[21]

C. boindii

0.31

0.33

[35]

C. blankii

0.36

0.25

[32]

C. mogii

0.56

0.39

[32]

C. parapsilosis

0.66

0.46

[32]

C. tropicalis

0.56

0.39

[32]

S. saki

0.06

0.04

[32]

S. cerevisiae

0.17

0.12

[32]

S. carlsbergensis

0.04

0.04

S. fibuligera

0.06

0.07

[25] [25]

Sch. pombe

0.07

0.05

[32]

R. toruloides

0.07

0.08

T. candida

0.03

0.04

[25] [25]

D. sake

0.22

0.13

[21]

H. anamala

0.34

0.20

[21]

P. miso

0.38

0.22

[21]

M. sitophila

0.37

0.22

[21]

T. melibiosaceum

0.12

0.12

[21]

P. tannophilus

0.15

0.11

[45]

K. cellobiovorus

0.25

0.35

[33]

yeast

species

278 All the 12 yeast strains examined by Gong et al. [25] utilized glucose, xylose, xylulose and xylitol as carbon and energy source, except a respiratory-deficient mutant of S. cerevisiae, which was not able to catabolize xylose or xylitol aerobically. Most of the yeast strains produced xylitol from both xylose and xylulose except R. toruloides. The influence of the initial substrate concentration on the fermentation kinetics of

Pachysolen tannophilus has been investigated under different cultural conditions [30]. There was always an uncoupled production of biomass and xylitol throughout the fermentation period. This phenomenon was more obvious at high initial substrate concentration (157 g/I) than at a low value. The biomass yield was more affected than that of product. Maximum xylitol yield was obtained at 40 g/I of substrate concentration. Xylitol production has also been studied from glucose in which a sequential process was employed in which glucose was first converted to xylulose by the action of a bacterium, Aerobacter suboxidans, and then xylulose is converted to xylitol by yeast [42]. A mutant strain of Candida tropicalis HXP2 yields xylitol at greater than 90% of theoretical value [43]. In contrast, the native strain of this species produces only 41.2% yield under identical conditions. The mutant strain also produces high quantities of xylitol from bagasse hemicellulose hydrolysate. The same group also examined a total of 20 strains of yeast Candida belonging to 11 species for the extent of substrate utilization and product formation under aerobic and fermentative conditions [32]. Under aerobic conditions, all the strains utilized xylose readily with xylitol as the major product followed by arabitol and ethanol. C. parapsilosis 28474 and all the five strains of C. tropicalis were better xylitol producers. However, under fermentative condition rates of yeast growth and substrate utilization were much slower. Xylitol was always the major product (1-8.4 g/I) of xylose utilization by various species of Saccharomyces tested [32]. In general, xylose utilization was poor by Saccharomyces species. On the other hand, most of the strains of Schizosaccharomyces pombe readily utilized xylose but the production of xylitol (1.13.5 g/I) was poor. The pattern of D-xylulose utilization and product formation is different from D-xylose utilization. Most of the yeast tested utilized xylulose readily under aerobic and fermentative conditions with the exception of Candida blankii [32]. Candida parapsilosis was the best xylitol producer among all the Candida species tested. Under aerobic condition xylitol was the major product, whereas arabitol was the major product under fermentative condition. Most of the Saccharomyces strains utilized xylulose readily under both aerobic and fermentative conditions. Saccharomyces uvarum 24556 produced mostly ethanol

279 under fermentative condition. Arabitol was the major product of S. cerevisiae 26497 under both aerobic and fermentative conditions. S. cerevisiae 4132 produced equal amounts of xylitol and arabitol under aerobic conditions, but produced mostly xylitol under fermentative condition. All the strains of S. pombe were found to be good producers of xylitol and arabitol from D-xylulose. Table 2 shows some results on xylitol production from D-xylulose.

Table 2 Xylitol

production

from

Species

D-xylulose

by

different

yeast

Xylitol yield

Productivity

(g/g)

(g/I/h)

C. utilis

0.04

0.05

C. blankii

O.04

0.03

C. mogii

0.20

O.14

C. steatolytica

O.08

0.06

C. parapsilosis

0.32

0.22

C. pseudotropicalis

O.16

O.11

C. tropicalis

0.04

0.03

B. claussenfi

O.03

0.02

S. uvarum

O.13

0.09

S. cerevisiae

0.39

0.27

Sch. pombe

0.23

O.14

T. candida

0.04

0.04

T. melibiosaceum

O.12

O.13

T. hensenfi

0.06

0.04

W. robertii

0.26

O.18

Compiled from references [25,32]

species

280 Most yeast strains examined for L-arabinose conversion under both aerobic and fermentative conditions produced arabitol as the major product and C. tropicalis was found to be the best arabitol producer [32]. Product concentration ranged from 4 g/I to 27 g/l. Table 3 shows arabitol production by different yeast species.

Table 3 Arabitol production by different yeast species Yeast

Debaryomyces miso Candida polymorpha Candida arborea Candida guilliermondii Torulopsis halophile Torulopsis versatilis Monilila sitophila Schizosaccharomyces pombe Pachysolen tannophilus Candida tropicalis

Arabitol

Yield

(g/I)

(g/g)

25.9

0.26

27.5

0.28

43.0

0.43

26.6

0.27

20.0

0.20

15.3

0.15

12.4

0.12

7.0

0.14

18.0

0.36

27.0

0.54

Compiled from references [21,32]

Kluyveromyces cellobiovorus, an efficient ethanol producing strain, was found to produce 25 g/I and 22 g/I xylitol from 100 g/I xylose under semianaerobic and aerobic conditions, respectively. Although xylitol production was studied as a by-product during ethanol fermentation, xylitol yields were significant [33]. A methanol yeast, Candida boindii 2201 accumulates a large amount of xylitol in

281 xylose containing medium alongwith small amounts of D-xylulose [35]. The highest amount of xylitol was obtained from cultivation on 100 g/I D-xylose. Increase in xylose concentration beyond this level results in the decrease in xylitol production, possibly due to an osmophilic effect on the cells of C. boindii or substrate repression of D-xylosemetabolizing enzymes. The low yield obtained from low xylose concentration was attributed to its use for cell mass production. Almost same amounts of xylulose accumulated at 100 and 150 g/I of xylose while slightly lesser amounts accumulated at 50 g/I xylose concentration. Under normal condition, S. cerevisiae H477 converts xylose to xylitol with a yield of 0.08 g/g xylose [44]. However, when glucose was added to the fermentation medium in a fed-batch pattern, the yield increased to 0.72 g/g. Since conversion of xylose to xylitol does not produce any energy for the cells, therefore a cosubstrate (glucose) is needed for growth and cofactor regeneration. Furaln et al. [45] studied xylitol formation by C. parapsilosis in continuous culture. When the dilution rate was varied, keeping the aeration and agitation constant at 0.3 w m and 250 rpm, respectively, the specific rate of xylitol formation was linearly dependent on the dilution rate. Xylitol formation was found to be directly coupled to growth of biomass for dilution rates below 0.14 h1. Maximum xylitol production (0.77 g/I) was obtained at D=0.055 h1 with coresponding yield of 0.16 g/g xylose (Table 4). However, feed xylose concentration (10 g/I) was not completely utilized, probably because of oxygen limitation.

Table 4 Xylitol production by Candida parapsilosis in continuous culture [45] Dilution

Biomass

Xylitol

Specific rate of

rate (h 1 )

yield (g/g)

yield (g/g)

xylitol production (g/g/h)

0.055

0.51

0.16

0.018

0.102

0.55

0.14

0.026

0.139

0.49

0.11

0.032

282

The ability of Candida guillermondii to produce xylitol from xylose and to ferment individual non-hemicellulosic derived sugars has been investigated in microaerobic conditions [36]. When the medium contained xylose as the sole carbon source, xylitol production started without lag period and was correlated with growth. Xylose was entirely consumed after 46 h of fermentation with .xylitol concentration of 14.1 g/I and specific xylitol productivity of 0.17/g/g/h. Xylitol yield was constant throughout the period and reached a value of 0.63 g/g corresponding to 69% of the theoretical value. Ethanol was detected in negligible amounts. Glucose, mannose and galactose were rapidly fermented by C. guillermondii, their specific uptake rates being 2.2-, 1.8- and 1.5-times, respectively, higher than that for xylose. However, these hexoses were found to be utilized only for growth and ethanol production, their corresponding polyols were not detected in the medium. The specific rate of arabinose uptake was very low, being one twelveth of xylose, but arabitol was accumulated at the level of 1.1 g/l. L-Arabinose can serve as an inducer and substrate for NADPH-linked xylose reductase [46,47]. Arabitol formation could consequently result from the action of a single aldose reductase with differing substrate specificities, though the existence of a separate enzyme is possible [48,49]. At a lower xylose concentration and lower aeration rate, cell concentration of

Candida tropicalis is low [50]. In these conditions, xylitol is produced at an earlier phase of cultivation. At higher xylose concentration and aeration rates, cell concentration is higher and consequently xylitol production is also higher. As a variance analysis for growth rate, aeration rate is significant at 95%. For xylitol yield, the interaction between xylose concentration and aeration rates must be admitted at the level of 95%. An increase in the initial xylose concentration from 10 g/I to 300 g/I led to the activation of xylitol production by C. guillermondii [36]. The xylitol yield increased gradually with increase in substrate concentration. The maximum xylitol concentration (221 g/I) and yield (0.75 g/g) corresponding to 82.6% of the theoretical value was obtained at a substrate concentration of 300 g/I (Table 5). In contrast to the xylitol production process, growth process of Candida guillermondii is gradually inhibited by an increase in the initial xylose concentration. However, the specificity of reduction with regard to xylose is high. This physiological property is of prime importance. The biological process could also alleviate or counteract the limiting operations which characterized the chemical process, as chromatographic fractionation with recycle steps to remove non xylitol polyols or nonxylose sugars. C. guillermondii produces xylitol with only small quantitites of ethanol and arabitol as by-products.

283 Table 5 Xylitol production and fermentation yields of Candida guillermondii grown on D-xylose under microaerobic conditions [36] Substrate Specific

Xylitol

Xylitol

Specific rate

Fermentation

conc.

growth

concentration

yield

of xylitol

time

(g/I)

rate (h 1 )

(g/I)

(g/g)

production (g/g/h)

(h)

10

0.11

6.2

0.46

0.08

46

50

0.11

30.9

0.59

0.10

165

110

0.03

68.7

0.60

0.17

238

200

0.02

151.7

0.71

0.22

291

300

0.01

221.0

0.75

0.19

406

The enzymatic production of xylitol from xylose using resting cells of Candida

pelliculosa (xylose reductase) coupled with those of Methanobacterium sp. (F42o-NADP oxidoreductase) has been demonstrated [51]. In this reaction, hydrogen serves as the substrate for reducing F42o via hydrogenase and then, the reduced F42o is coupled with oxidoreductase to reduce exogenous NADP to NADPH. Finally the NADPH produced in

Methanobacterium sp. cells could be transferred to C. pelliculosa cells to reduce xylose to xylitol. However, continuous production of xylitol was not successful because of the leakage of the related enzymes from both cells. Thus the immobilization technique for whole cells of the two organisms was used for the continuous production of xylitol. The cells of C. pelliculosa and Methanobacterium sp. were immobilized in agar, calcium alginate, k-carrageenan, photo-crosslinkable resin prepolymer, polyacrylamide or polyurethane prepolymer [34]. Among them the highest conversion (40%) was observed with benzene-treated cells coimmobilized in photo-crosslinkable resin prepolymer (ENT 2000 and 4000) followed by polyacrylamide gel (20% conversion) and calcium alginate gel (10% conversion). The conversion of xylose into xylitol was achieved using either separately immobilized cells or coimmobilized cells. In a separately immobilized cell

284

system with methyl viologen as an electron carrier, the conversion degree of xylose to xylitol reached almost 100% after 33 h of incubation when the volume ratio of immobilized methanogen to immobilized yeast was 1:2 (enzyme activity ratio, 1:4.3). The degree of conversion was found to decrease with decreasing volume ratio of immobilized bacterium and yeast cells (Table 6). In the coimmobilized cell system, the conversion degrees and rates in these volume ratios were generally higher than those in the separately immobilized system, suggesting more rapid inter-transfer of NADP(H) between methanogen cells and yeast cells. Coimmobilized cells gave stable operation for about two weeks in a continuous culture with about 35% conversion. The low conversion in continuous culture as compared to batch culture was attributed to the insufficiency of hydrogen supply under atmospheric pressure in column reactor. Although, NADP was continuously supplied to the reactor, it is essential to retain NADP(H) in the reactor so as to reuse it in the continuous reaction. It has been demonstrated that a sulfonated polysulfone membrane reactor is feasible to use for recycling NADP(H) and F,2o in the production of xylitol [52].

Table 6 Conversion of D-xylose into xylitol by immobilized cells of Candida pelliculosa and

Methanobacterium sp. HU" [34] Volume ratio of

Enzyme

Conversion

immobilized cells

activity

(%)

of bacterium and yeast

ratio b

1:2

1:4.3

100

1:1

1:2.2

85

1:0.5

1:1.1

70

"Separately immobilized (photo-crosslinkable resin prepolymer) system with methyl viologen as an electron carrier ~ h e ratio of F42o-NADP oxidoreductase to xylose reductase

285 Various thermotolerant yeasts have been evaluated for the bioconversion of xylose into xylitol [29]. Xylitol production ranged from 8.3 g/I to 46.9 g/I from 100 g/I xylose. Among different strains of Candida sp., strain HT8 produced maximum xylitol yield and productivity (Table 7). Candida sp. utilized xylose as a carbon and energy source with a generation time of about 2 h. The initial growth rate was similar when the yeasts were incubated at 35, 40 or 45~

Yeasts failed to grow when the incubation temperature was

more than 50~

Table 7 Xylitol production by thermotolerant yeast species a [29] Xylose

Xylitol

Productivity

consumption (%)

yield (g/g)

(g/I/h)

HT1

90.0

0.44

0.46

HT2

49.3

0.08

0.09

HT3

68.3

0.19

0.20

HT4

98.3

0.33

0.33

HT5

74.1

0.34

0.35

HT6

99.2

0.40

0.42

HT7

92.0

0.45

0.47

HT8

94.4

0.47

0.49

Hansenula polymorpha

75.4

0.24

0.25

Strain

Candida sp.

a

Incubation temperature, 45~

286

Repetitive isomerization of xylose followed by yeast fermentation of xylulose, and simultaneous enzymatic isomerization and yeast fermentation have also been proven to be potential methods for conversion of xylose to ethanol and xylitol. Using baker's yeast and glucose isomerase enzyme, a xylitol concentration of 33 g/I with a yield of 0.28 g/g was obtained from 120 g/I xylose at 30~ [53]. Viability of the yeast remained fairly constant and at least 90% of isomerase activity was recoverable after fermentation. Linden and Hahn-Hagerdal [54] evaluated fermentation of untreated spent sulfite liquor with five yeasts, Candida tropicalis, Pichia stipitis, Pachysolen tannophilus,

Saccharomyces cerevisiae and Schizosaccharomyces pombe, and a coculture of P. tannophilus and S. cerevisiae, in the presence of commercial xylose (glucose)isomerase and 4.6 mM azide. C. tropicalis and P. tannophilus consumed sugars quite well, whereas sugar consumption was rapidly inhibited in P. stipitis. Maximum xylitol production was obtained with P. tannophilus followed by a coculture of P. tannophilus and S. cerevisiae in 48 h of incubation. Table 8 shows some results on xylitol production from xylose by different yeasts in the presence of glucose (xylose)isomerase enzyme.

Table 8 Xylitol production from D-xylose by yeasts in the presence of glucose (xylose)isomerase Organism

Xylitol

Productivity

yield (g/g)

(g/I/h)

Baker's yeast

0.28

Candida tropicalis Pachysolen tannophilus Saccharomyces cerevisiae Saccharomyces cerevisiae + Pachysolen tannophilus

0.13

0.07

0.35

0.23

0.04

0.02

0.18

0.12

Compiled from references [53,54]

287 3.2

Bacteria

Polyol production by a gluconate-utilizing strain of Corynebacterium sp. has been studied in shake flasks [38]. In order to examine the relationship between the pentose concentration and the yield of pentitol, various concentrations of pentoses were added to the medium containing 9.6% potassium gluconate after 2 days of cultivation. Maximum yields were obtained when each of the pentose sugar (D-xylose, L-arabinose and Dribose) was added at the level of 150 g/l. Xylitol concentration was found to increase throughout the period of incubation and reached to a concentration of 66 g/I after 14 days. Arabitol production reached a concentration of 55 g/I in 14 days with 150 g/I arabinose, whereas in D-ribose medium (150 g/I), 25 g/I ribitol was obtained in 14 days.

Corynebacterium sp. was capable of rapid growth on gluconate as carbon source but at much slower rate on pentoses. Both xylitol production and the growth of Enterobacter liquefaciens were found to decrease when the initial xylose concentration in the medium was above 100 g/l. The xylitol production reached the maximum after cessation of the growth [39]. Pentitol production by different bacterial species is shown in Table 9.

Table 9 Production of pentitols a by different bacterial species Organism

Substrate

Pentitol Productivity Yield (g/g) (g/I/h)

Corynebacterium

Xylose

0.48

0.21

[38]

sp. 208

Arabinose

0.32

0.14

[38]

Ribose

0.16

0.07

[38]

0.31

0.25

[39]

0.40

0.80

[40]

0.74

0.62

[40]

Enterobacter liquefaciens Xylose Mycobacterium Xylulose smegmatis Xylulose

Reference

aXylitoI, arabitol and ribitol are the respective pentitols of xylose, arabinose and ribose

288

Mycobacterium smegmatis, grown on various carbon sources, completely utilizes D-xylulose to produce xylitol (Table 10). Cells grown on xylitol or mannitol were found to produce maximum product yield. With immobilized cells of M. smegmatis, 15 g/I of xylitol was obtained from 20 g/I xylose in 24 h.

Table 10 Xylitol production from D-xylulose by Mycobacteriumsmegmatisgrown on various carbon sources a [40] Growth

Xylitol

Conversion

substrate

(g/I)

(%)

D-Xylose

11.3

56.5

D-Galactose

12.0

60.0

L-Sorbose

11.9

59.6

Xylitol

14.8

74.2

D-Sorbitol

12.6

62.9

D-Mannitol

14.8

74.2

a Concentration of growth substrate, 10 g/I; D-xylulose concentration, 20 g/I; incubation time; 96 h; temperature, 30~

Xylitol is produced extracellularly in D-xylose containing medium during the cultivation of a fungus, Petromyces albertensis [41]. Xylitol was identified as the only polyol in HPLC analysis. The maximum amount of xylitol (36.8 g/I) was produced when the initial concentration of xylose was 100 g/l. Further increase in the substrate concentration decreased the xylitol yield due to osmotic effect on cells. The same amount of D-xylulose (2.3 g/I) was produced at 100 g/I and 150 g/I xylose concentrations.

289 4

FACTORS AFFECTING XYLITOL PRODUCTION

Bioconversion of D-xylose can be influenced by several cultural and nutritional factors. Several researchers have investigated the influence of these factors, and xylitol production has been found to increase several folds by optimizing the cultural and nutritional conditions.

4.1

pH and temperature

Initial pH of the medium significantly affects the final product yield. Kluyveromyces

cellobiovorus grow preferentially below pH 5 [33], whereas for Candida boindii, the optimum pH for growth and xylitol production were different [35]. The maximum amount of xylitol was obtained at pH 7, while the best growth was at the initial pH of 6.5. In the case of fungus, Petromyces albertensis, the maximum amount of xylitol and fungal biomass was obtained with an initial pH between 6 and 7. Temperature has a profound effect on all aspects of microbial growth, metabolism, viability, and fermentability. Microorganisms have a relatively limited temperature range for growth. Candida sp. HT4 utilized both glucose and xylose as a carbon and energy source [29]. The initial growth rate was slower when incubated at 30~ than at higher temperature but it was similar when the yeast was incubated at either 35, 40 or 45~

It

failed to grow at 50~ indicating Tmax between 45 and 50~ The optimal temperature for fermentation is usually higher than the optimal growth temperature [55]. However, due to increased rate of cell death it is more difficult to maintain the ability of yeasts to ferment at higher temperature [56]. The simultaneous enzymatic isomerization and yeast fermentation process is not considered to be efficient. Higher temperature (70~ isomerization, whereas low temperature (30~

and pH (7.5) are best for

and low pH (4.0-5.0) are optimal for yeast

fermentation. Therefore, operation of isomerization and fermentation under their respective optimal conditions would give the best efficiency. This can be done by a closed-loop recycle of sugar mixture through an external isomerase column, using a yeast settler or separator to remove yeast from the sugar solution and retain yeast in the fermentor.

290 For Enterobacter liquefaciens, optimum temperature of growth and xylitol production was 30~ [39]. Above this temperature both biomass and xylitol yields decreased considerably. Table 11 shows optimum pH and temperature of xylitol producing organisms.

Table 11 Optimum pH and temperature for xylitol producing organisms Organism

pH

Temperature

Reference

(oc) S. cerevisiae

4.0

30

[53]

Sch. pombe

4.0

30

[53]

S. carlbergensis

5.6

26

[25]

S. diastaticus

5.6

26

[25]

C. didensfi

5.6

26

[25]

C. utilis

5.6

26

[25]

T. candida

5.6

26

[25]

Candida sp. HT8

7.0

45

[29]

H. polymorpha

7.0

45

[29]

D. miso

5.0

30

[21]

P. miso

5.0

30

[21]

C. parapsilosis

4.5

30

[45]

K. cellobiovorus

5.0

28

[33]

C. tropicalis

4.0

30

[50]

M. smegmatis

7.0

35

[40]

Corynbacterium sp.

6.5

30

[38]

E. liquefaciens

7.0

30

[39]

P. albertensis

6.0-7.0

28

[41 ]

291

4.2 Oxygenation

The availability of oxygen has a significant influence on D-xylose fermentation by yeasts. Rate of xylose utilization under anaerobic condition is slow due to reduced yeast growth rate [32]. However, higher xylitol production has been observed under oxygen limited conditions. Saccharomyces cerevisiae 4132 produces approximately equal quantities of xylitol and arabitol under aerobic condition but mostly xylitol under fermentative condition [32]. Candida parapsilosis 28474 produces predominantly xylitol under aerobic conditions and arabitol under fermentative conditions. Effect of aeration on xylitol production is shown in Table 12.

Table 12 Effect of aeration on xylitol production Aeration

Xylitol

Organism

yield vvm

Pachysolen tannophilus Candida parapsilosis

Candida tropicalis

ml/min

Reference

(g/g)

0.45

0.15

0.50

0.13

1.17

0.04

0.15

0.32

0.30

0.47

0.60

0.54

1.00

0.50

2.00

0.53 400

0.43

700

0.50

1000

0.45

400 (90% 02) 0.52

[30]

[45]

[50]

292 With respect to oxygenation, Pachysolen tannophilus is the most thoroughly investigated organism. In spite of constant aeration at 0.5 vvm, there is a oxygen limitation in a typical batch culture [30]. Under completely anaerobic condition (N 2 sparged through the medium), P. tannophilus was incapable of significant growth. With the increase of aeration rate, fermentation time decreased from 68 h to 32 h. The product formation was favoured by aeration upto a certain level, while biomass production continually enhanced by increasing the aeration rate. Similar results have been reported in the case of Kluyveromyces cellobiovorus where xylitol production under aerobic and anaerobic conditions was 22 g/I and 25 g/I, respectively. Xylitol and biomass formation of C. parapsilosis was also found to be strongly influenced by oxygen consumption [45]. Oxygen consumption of the organism increased with the increase of aeration. Xylose consumption was more directed towards growth than the product formation as the aeration rate increased. Excess oxygen activates TCA cycle and thereby regenerates NAD which is used to transform xylitol into xylulose with NADH production. Xylulose is degraded through the pentose phosphate and Embden-MeyerhofParnas pathways. Thus xylitol accumulation drops with the increase in oxygen uptake. Xylitol production rate can be increased by optimizing the fermentation conditions in two ways [50]: the first, a kinetic approach, utilizes a kinetic model on xylitol production from xylose, and the second is an application of statistical method. The second method has been proved to be very useful in the optimization of biological reaction conditions [5759]. For effective xylitol production by Candida tropicalis, the first step to consider is the rapid accumulation of cells in the culture medium. Thus higher level of dissolved oxygen was kept at the earlier phase of cultivation and thereafter it decreased by the respiration of the organism at the lower level. The growth rate of C. tropicalis increased with increase in aeration rate, reaching the highest value at the aeration rate of 700 ml/min. However, xylitol production decreased when the growth rate reached to the highest level. This suggested a possible shortage of nitrogen source. An increase in concentration of yeast extract led to increased growth rate. The maximum xylitol production rate (1.78 g/I/h) was obtained at 400 ml/min aeration with 90% oxygen. When xylose concentration was increased to 150 g/I, enough cell growth with maximum xylitol productivity (2.44 g/I/h) was obtained at aeration rate of 400 ml/min with 90% oxygen. A variance analysis for xylitol production rate indicated that initial yeast extract concentration was significant at 99%, and the interaction between xylose concentration and aeration rate was admitted at a level of 95%. These results show the importance of both initial yeast extract concentration as nitrogen source and the balance between xylose concentration and aeration rate [60].

293

4.3 Magnesiumsupplementation

Influence of magnesium concentration on growth, ethanol and xylitol production of

Pichia stipitis has been studied under conditions of constant oxygen uptake rate [61]. A significant increment in specific growth rate occurred when Mg2+ level in culture media was increased from 1 mM to 6 mM. However, no significant variations were observed in the specific xylose consumption rate. Biomass/xylose and biomass/Mg 2+yields increased with the Mg 2+ upto a maximum value of 4 mM, ethanol being the major product. At low Mg 2+ concentration (lmM), 49% of carbon flux to ethanol was redirected to xylitol production, accomplished through intracellular NADH accumulation (Table 13).

Table 13 Effect of magnesium concentration on growth and product formation by Pichia stipitis [61 ] Magnesium cncentration (mM) Parameters 1

2

4

6

0.02

0.06

0.08

0.09

Specific xylose consumption (g/g/h) 0.01

0.06

0.08

0.09

Specific growth rate (h1)

Continuous culture (D=0.023) a Biomass

5.4

9.3

Ethanol

4.8

10.1

CO 2

68.4

83.9

Xylitol

17.1

a

Per cent of carbon mol of xylose consumed

0

294

4.4

Nitrogen sources and organic nutrients

Effect of different nitrogen sources and organic nutrients on xylitol production by

Candida boindii [35] and Petromyces albertensis [41] has been investigated. Ammonium acetate was found to be the most effective nitrogen source (Table 14).

Table 14 Effect of different nitrogen sources on xylitol production" Nitrogen source

Candida boindii

Petromycesalbertensis

[35]

[41]

Ammonium chloride

8.5

4.5

Ammonium sulfate

12.6

8.2

Ammonium nitrate

15.2

12.8

Ammonium phosphate

12.1

13.4

Ammonium carbonate

12.5

12.4

Ammonium acetate

18.3

16.7

Ammonium formate

12.9

11.8

Ammonium citrate

15.2

14.2

alnitial D-xylose concentration 100 g/I; nitogen source 5 g/I; cultivation time for C. boindii and P. albertensis were 4 days and 12 days, respectively.

Thiamine is an essential compound for the growth of Corynebacterium sp 208. High xylitol yields can be obtained from this organism by supplementing the medium with greater than 0.2 mg/I thiamine hydrochloride. However, corn steep liquor at the level of 6 g/I has been found to be as effective as thiamine. Inorganic ammonium salts are most favourable nitrogenous sources for xylitol production by Enterobacter liquefaciens. Of

295 various organic nutrients tested, corn steep liquor was found to be most effective nutrient for xylitol production by E. liquefaciens. Corn steep liquor concentration of 6 g/I was the optimum (Table 15). Further increase in concentration remarkably decreased the production of xylitol.

Table 15 Effect of organic nutrients on xylitol production by different organisms a Organisms

Organic

Nutrient

Xylitol

nutrient

concentration (g/I)

(g/I)

Candida

Malt extract

9

3.3

boindii

Yeast extract

6

31.4

Dried yeast

9

8.1

Peptone

9

6.8

Polypeptone

12

14.6

Malt extract

10

2.8

Yeast extract

10

30.6

Dried yeast

10

9.5

Peptone

10

6.8

Enterobacter

Corn liquor

6

30.7

liquefaciens

Malt extract

20

20.0

Peptone

20

22.1

Casamino acid 12

14.4

Yeast extract

15.2

Petromyces albertensis

4

Reference

[35]

[41 ]

[39]

alnitial D-xylose concentration 100 g/I; cultivation time for C. boindii, P. albertensis and

E. liquefaciens were 4, 12 and 5 days, respectively.

296

4.5

Methanol supplementation

The supplementation of methanol in xylose medium has been found to increase the xylitol production in yeast and filamentous fungi. Xylitol level of 45.5 g/I has been obtained with Candida boindii [35] in the medium supplemented with 2% methanol, while only 36 g/I xylitol was obtained in the medium without methanol (Table 16). However, no significant differences of growth and xylulose accumulation were observed for all cultivations. Though a little methanol was consumed in the medium, NADH for the reduction of xylulose to xylitol or xylose to xylitol could be supplied by oxidation of methanol. In addition, a small amount (0.2-0.5%) ethanol as a by-product was also detected in the methanol supplemented medium. The amount of ethanol was higher in the presence of higher concentration of methanol. Similar results of methanol supplementation in the growth medium of P. albertensis were obtained, where xylitol production increased by about 8% with 1% methanol [41]. Again, no significant differences in fungal biomass and xylulose accumulation were observed. Only a small amount (0.01%) of methanol was consumed in the medium.

Table 16 Effect of methanol supplementation on xylitol production a Organism

Candida

Methanol

Xylitol

% Increase

(%)

(g/I)

in yield

0

36.0

[35]

1

44.0

18.2

2

48.5

25.8

Petromyces

0

36.8

albertensis

1

39.8

boindii

Initial D-xylose concentration 100 g/l.

Reference

[41] 8.5

297 5

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11 Microbial Production of Single Cell Protein (SCP) and Single Cell

(sco)

1

INTRODUCTION

Harnessing the potential of microorganisms for the production of edible proteins dates back to World War I. During this period first modern efforts were made to produce torula yeast on a large scale in Germany. Subsequently, cultivation of yeasts on hydrocarbon substrate was realized to have a great potential as a source of food protein [1,2] and later microorganisms were grown on hydrocarbons for the purpose of producing proteins at a commercial scale [3]. By late 60's many countries had started producing proteins from n-paraffin feedstock and termed as petroproteins or microbial proteins which was later changed to a more aesthetic term single cell protein (SCP) [4]. The advantages of single cell protein over conventional agricultural proteins are i ii. iii.

9 SCP production is independent of the agricultural or climatic conditions. SCP production is very fast due to rapid growth of microorganisms. SCP production can be tailored for better protein composition as

microorganisms are genetically amenable. However, the major bottleneck in the production of SCP for food are high cost of raw material as substrate and acceptability of SCP as food. Thus the main market for SCP has been as animal feed [5]. A number of advances in engineering aspects such as protein recovery, heat removal, oxygen transfer and hydrocarbon transfer etc. have been employed to reduce the cost of SCP production. Alternatively, utilization of renewable lignocellulosic wastes as raw material for the production of SCP has been suggested [5-8]. Although considerable literature is available on various aspects of SCP [9-12], here a concise review of our understanding of SCP production from xylose and lignocellulosic wastes has been provided. Whilst use of cellulosic hydrolysate for various fermentation products has attracted considerable interest, SCP production has not been explored extensively probably due to their lesser profitability as compared to other fermentation products. Lignocellulosic-derived fermentative ethanol has also been suggested to be a competitive substrate for SCP production 301

302 [6,13]. Use of ethanol as substrate has several distinct advantages such as purity, acceptability, ease of storage and handling, non-toxicity, versatility as substrate, miscibility with water, relatively low oxygen demand, low heat production and high cell yield. In addition many bacteria and yeast can metabolize ethanol as sole source of carbon and energy. Due to lower cost of fermentative ethanol, it can compete with ethylene derived ethanol as a substrate for SCP production. In addition, non-food grade cellulosic wastes can be used for ethanol production which would then be suitable as a pure substrate for the production of SCP supplement for human consumption e.g. torula yeast. With increasing knowledge of our current understanding about the use of cellulolytic microorganisms as source of protein or use of cellulolytic organisms in co-culture with pentose metabolizing microorganisms may prove helpful in production of SCP. Lignocellulosics

consists of three

major carbohydrates

viz.

cellulose,

hemicellulose and lignin. It is essential from an economic point of view that all the main components should be utilized to the fullest extent because the carbon source is a major determinant of the total production costs of SCP [14]. For single cell protein production these polymers are converted to monomeric sugars, that can be used as starting material for various products by fermentation and for SCP by assimilation [7]. In practice lignocellulosics are resistant to direct fermentation and chemical or physical treatment is usually required to expose the cellulose to microbial enzymes. Even pure celluloses are resistant to microbial or enzymatic attack owing to their crystalline structure and sometimes require pretreatment. Various methods of pretreatment are described in Chapter 3. It has been observed that a large fraction of hemicellulosics remain unutilized due to inability of microorganisms to grow on pentoses. Thus a complete understanding of hemicellulosic pretreatment and subsequent utilization by microorganisms as an integrated process is essential for the development of an economically viable process. Although a number of studies related to bioconversion of pentoses to liquid fuel have been published since the late 70's, only very little is understood about pentose utilization for SCP production. The success of processes for the production of microbial proteins (SCP) as specific process and as adjunct to waste processing scheme over the traditional method of their production has generated interest for harnessing the potential of microorganisms for the production of single cell oil (SCO) [15]. Organisms grown for SCO have dual advantages as in addition to SCO production, defatted biomass residue may contain up to 40% protein [15]. Thus SCO production has an edge over that of SCP. However, for obvious economic reasons, these microbial oils should be of higher value added types rather than resembling the bulk low priced soybean or

303

sunflower oil [16]. The ease with which microorganisms are amenable to genetic manipulation provides opportunity for the production of value added lipids. These properties of microbial lipids have been explicitly summarized by Ratledge [17]. These important properties are easy extractability, high content of triacyiglycerol, no deleterious fatty acyl residues in lipids, resistance to autoxidation, and an ability to grow in continuous fermentation. A large number of yeasts and fungi are known to accumulate as much as 30-75% of total cell weight as lipids but their use for the development of commercial process has been limited so far mainly due to the cost factor. The major cost factor is substrates utilized by oleaginous microorganisms. This has awakened an interest in utilization of cheaper substrates or waste products for SCO production [17]. A large number of substrates such as molasses, whey, starch and ethanol have been tested for SCO production [17]. Hydrolyzed plant waste material such as wood and straw containing mixture of hexoses, mainly glucose, from cellulose and pentoses, mainly xylose from hemicellulose can provide economically viable substrates for SCP and SCO production, however, utilization of pentoses and lignocellulosic waste for both SCP and SCO production have not been studied in great details. Thus a survey of microorganisms utilizing both pentoses and hexoses simultaneously for SCP and SCO production has a great potential in lowering the cost of substrate. Therefore a summary of microorganisms utilizing pentoses for the production of SCO is also included in the present chapter.

2

MICROORGANISMS

USED FOR SCP P R O D U C T I O N

Although bacteria, algae, fungi and yeasts all are potential sources of SCP, Wilkinsion [18] suggested the following desirable features for an organism suitable for the production of SCP i inexpensive or no nutrient requirement ii high protein productivity i.e. high content of protein and growth rate iii

suitable amino acid composition

iv v vi vii . , ,

VIII

good nutritional availability high yield lack of toxicity ease of harvesting low risk of contamination

304 Krug and co-workers [19] have compared various characteristics of bacteria and yeasts for their suitability as producers of SCP. Both yeasts and bacteria have their advantages and disadvantages. While bacteria have advantage over yeast by virtue of possessing higher protein content, faster growth rate, high cell yield and no requirement of growth supplementation, yeasts on the other hand have some distinct advantages over bacteria such as low contents of nucleic acid, lower cost of cell recovery due to larger sizes, lower risk of contamination as they grow well at pH 3-4 and have better public acceptance. Fungi on the other hand offers certain advantages for the production of SCP due to their fibrous nature. It is apparent that they can be harvested with ease and provide a texture for the fabrication of food analog. The problem of acceptability of fungi as food product is also low due to their long association with food products such as mushrooms. Low content of nucleic acid in fungi also renders them suitable for SCP production. In addition fungi are known to possess cellulolytic activity and thus can use lignocellulosic wastes. The major disadvantage of using fungi for SCP production appears to be their slow growth rates. Yeasts and filamentous fungi have commonly been employed forthe production of SCP, however, some processes using mesophilic and thermophilic bacteria have also been described [6]. Commercial scale SCP processes have mainly employed hydrocarbon and alcohol fermentation by yeasts. Alternatively production of SCP from carbohydrate waste has also attracted considerable interest [5]. This section elucidates the potential microorganisms employed for bioconversion of pentoses and lignocellulosic materials containing pentoses to SCP. Production of fodder yeasts on hydrolysates of plant material e.g. agricultural, food, timber, paper and wood processing wastes have been employed in USSR since mid thirties however, little was known until early eighties about the pentose utilization and assimilation by yeasts [6]. At present a number of Candida sp. including Candida blankii, C. maltosa, C. utilis, C. scottii and Trichosporon cutaneum, Hansenula polymorpha and Pichia pinus are known to assimilate sugars from lignocellulosic hydrolysates and produce SCP. Although it is generally accepted that Saccharomyces cerevisiae is unable to assimilate D-xylose [20], van Zyle and co-workers [21] have observed that four strains of Saccharomyces cerevisiae viz. ATCC 26602, ATCC 26603, CBS 1907, CSIR-Y2 utilize xylose aerobically at different efficiencies in the presence of a mixture of substrates. The degree of xylose utilization by S. cerevisiae ATCC 26602 depends upon the presence of other substrate or yeast extract. The utilization of xylose is observed to be greatest when sugar substrates such as Dribose are co-metabolized [21]. Various yeast species belonging to Candida have been screened for their ability

305 to assimilate sugars from beech wood hydrolysates [22]. Beech wood hydrolysate obtained by phosphoric acid treatment contain xylose as a major sugar (55-97%), while other sugars viz. arabinose, ribose, galactose, mannose, glucose and rhamnose are also found in the mixture. The assimilation of individual sugars by Candida sp. has been tested for over 72 h in cultures grown at 37~ C. maltosa 18, C. utilis 66-30, C.guilliermondii 63-7, C. blankii 87-3 and 87-4 and C. scottii 79-46 assimilated glucose, galactose, xylose, mannose, L-and D-arabinose, L-rhamnose and mixtures of these sugars to different extent. C. blankii 87-3 and 87-4 exhibited both maximum assimilation of sugars from beech wood hydrolysate as well as good growth and have been designated as the best strains. Strain 87-4 assimilated about cent percent of xylose, glucose and 92% of other sugars present in the hydrolysate and yielded 51% dry weight of biomass [22]. In another study, Jwanny et al. [23] tested a number of yeasts viz. Hansenula polymorpha, various strains of Candida e.g.C, boidinii Y4235 and 4232, C. maltosa R-42, C. tropicalis. C. guilliermondii 42075, C. lipolytica, Pichia

pinus and Saccharomyces cerevisiae 8112 for their ability to grow on vegetable processing waste. Of these strains tested P. pinus growing on 3% mango peel extract was selected as it produced 3 g biomass/g reducing sugar after 48 h fermentation and SCP production ranged from 66-70% of the dry weight. H. polymorpha has also been considered to be a suitable microorganism for SCP production from lignocellulosic materials due to its ability to utilize both glucose and xylose [24]. However this organism fails to utilize arabinose as a carbon source. SCP production by H. polymorpha from xylose on synthetic medium in batch culture has led to optimization of the process for the use of acid hydrolysate solution as raw material for SCP production. Wheat straw hydrolysed at 90~ using dilute sulfuric acid provided growth of H. polymorpha at pH 4.8 without any supplementation. In continuous culture at a dilution rate of 0.13h -1 the yield of 0.61 g biomass per gram ylose consumed and a biomass productivity of 0.74 g/I/h has been reported [24]. Meyer and co-workers [25] have isolated 26 yeasts strains from a large number of samples for their ability to utilize D-xylose and L-arabinose in vitamin-free medium at 36~ Six of these isolates belonging to C. blankii have shown rapid growth at 38~ on both D-xylose and L-arabinose in similar media. These isolates show highest growth in D-glucose followed by D-xylose and L-arabinose. All the six isolates had similar protein contents and grew on D-xylose at pH as low as 3. Some of these isolates even showed maximum growth at pH 3.5-4. The protein content of C. blankii isolates is comparable with C. utilis. All the isolates of C. blankii grew at 44~ while C. utilis fails to grow at this temperature. Interestingly, C. blankii isolates utilized all the carbon sources present in sugarcane bagasse hemicellulose hydrolysate (including

306 sucrose and acetic acid). This led to critical evaluation of C. blankii UOVS 63.1, UOVS 64.1, UOVS64.2 and CSIR-ESP 94 strains for their ability to grow and produce SCP on a simulated sugar cane bagasse hydrolysate medium [26]. Fermentation carried out at 38~ with agitation at 200-600 rpm showed that utilization of D-xylose, L-arabinose and acetic acid are delayed by the presence of D-glucose. However, after glucose depletion the other carbon sources are utilized simultaneously. A maximum specific growth rate of 0.36h -1 and cell yield of 0.47 g cells/g carbon source has been obtained [26]. Later, the growth of a C. blankii isolate, UOVS 64.2, in bagasse hemicellulose hydrolysate was evaluated in carbon-limited continuous culture [27]. C. blankiigrew satisfactorily on bagasse hemicellulose hydrolysate pre-treated by adding ammonium hydroxide, which provided the nitrogen source and raised the pH. During chemostat cultivation of C. blankii in hydrolysate diluted 50% with distilled water, the steady state values of dry mass increased markedly as the dilution rate increased from 0.05 to 0.15h 1, however further increase in dilution rate showed no effect on the growth. The biomass productivity increased linearly with increasing dilution rate up to 0.3h -~ with maximum productivity of 2.72 g/I/h. The cell protein and RNA contents also increased with dilution rate whereas DNA content decreased. Cultivation of C. blankii in undiluted bagasse hemicellulose hydrolysate showed critical dilution rate at 38~ and pH 5.0 to be 0.21 h-~ in contrast to 0.35h -1 in 50% diluted hydrolysate under similar conditions. However, cell yield, protein yield and protein contents were comparable in the two growth media. The protein yield was also comparable at pH 4 and 5 in undiluted hydrolysate although maximum specific growth decreased at pH 4. C. blankii thus proved to be very good yeast for SCP production as it utilizes all the different carbon sources including L-arabinose present in the bagasse hemicellulose hydrolysate. The relatively high optimum temperature of 38~ and ability to grow on undiluted bagasse hemicellulose hydrolysate at pH 4 has several advantages for commercial scale biomass and SCP production from this yeast. Under these conditions no strict aseptic condition is required and a large temperature difference between culture medium and cooling water reduces the cooling cost, which is estimated to be 2-5% of the total operating cost for SCP production [14]. In addition, the use of undiluted hydrolysate facilitates a more economical cell harvesting due to high cell concentration. Yinbo et al. [28] screened twenty five strains of yeasts and filamentous fungi for their ability to grow on steam exploded hemicellulose autohydrolysate liquor (SEHAL). The SEHAL is obtained from hard woods or agricultural residues by hot water extraction and contains mainly xylose and its oligmers. Screening on SEHALagar plate showed that amongst yeasts Trichosporon cutaneum 851 and Pichia stipitis

307 NRRL Y7124 exhibit characteristic higher growth rate. T. cutaneum 851 exhibited relatively higher protein content as well. The quality of biomass as feed from T. cutaneum has been further improved by UV and HNO 2 mutagenesis. Mutants with

decreased content of glycogen or lipids have been selected on to media enriched with high concentration of inhibitors such as sodium vanadate, lithium sulfate, thiourea or ferrous sulfate. By this method mutant strains with higher content of protein have been selected [28]. A mutant strain designated as 851S exhibited growth rate similar to parent and contained higher protein content (47%). The amino acid composition of protein revealed that 851S contained methionine 27.7 mg/g biomass which is higher than the normal feed yeasts [29]. At optimal culture conditions T. cutaneum 851S produced a biomass of 10.5 g/I with an yield of 0.75 g/g total sugar in less than 16 h. Continuous fermentation of T. cutaneum 851S in the SEHAL medium showed best results at a dilution rate of 0.33h -1 with a yield of 0.72 g/g total sugar, a cell concentration of 9.2 g/I and a productivity of 3.0 g/I/h. Alternatively steam-exploded waste material from paper mills, which requires no supplementation of nutrient can also be extracted by spent ammonium sulfite liquor (SASL). However, this medium contains some compounds inhibitory to microorganisms, therefore, growth of T. cutaneum 851S on SASL-extracted SEHAL is not observed at the beginning. An

acclimated strain of T. cutaneum 851S-6B-1, has been isolated after adapting yeast repeatedly on SASL-SEHAL agar plates and increasing SASL concentration followed by continuous adaptation in a chemostat. This acclimated T. cutaneum 851S-6B-1 grew well in all waste medium and gave a similar result in continuous fermentation with less risk of contamination at a lower medium cost [28]. As discussed earlier yeasts generally lack the ability to utilize oligosaccharides directly and enzymatic hydrolysis is required, whereas fungi can grow on oligosaccharides directly but their growth rate and protein contents are relatively low. This has prompted researchers to use mixed cultures of yeast and fungi. Using such an approach Peitersen [30], employed Trichoderma viride and the yeast Saccharomyces cerevisiae or Candida utilis. Fermentation carried out in an aerated 5 I fermenter with sodium hydroxide treated barley straw as substrate using culture of T. viride and yeast, revealed that the production time for maximum yield of cell protein is reduced by several days compared to fermentation with T. viride alone. The biomass has 22% of protein content and amino acid composition resembled that of T. viride alone. The presence of yeast in the mixed culture does not increase the

utilization of polysaccharide in straw, rather it enhances the production of protein and enzyme

degrading

cellulosics.

Mixed

cultures

of two

kinds

of cellulolytic

microrganisms, Cellulomonas sp and Aspergillus faecalis has shown an increased

308 growth rate [31-33]. Mixed culture of Cellulomonas sp and Candida utilis using barley straw pretreated with caustic soda has also been reported [34]. Cellulomonas sp. degrade the cellulose substrate to cellobiose, which is then consumed by the other organism in a mixed culture. Thus the inhibition of cellulase induction by cellobiose is avoided. Thus mixed culture provides a way of improving fermentation of cellulosic wastes. Advantages of using fungi for SCP production are obvious and have been discussed earlier. Fungi, most commonly employed for the production of SCP are Fusarium oxysporum, Trichoderma viride, T. reesei, Chaetomium cellulolyticum and

Aspergillus terreus. Conversion of lignocellulosic materials to SCP by some yeasts and fungi are listed in Table 1. Early studies have suggested Fusarium oxysporum var. lini as a source of SCP with high nutritive value [35]. Later this organism grown in a semicontinuous process on vinasse, a by-product from distillation of fermented sugar, exhibited a mycelium yield of 15.8 g/I after 24 h [36]. The mycelia from this organism contain 50% crude protein with nutritive value comparable to casein [36]. Trichoderma viride QM 9123 and QM 6a grown on barley straw pretreated with NaOH under high pressure have also been studied [37]. The production of enzyme cellulase and SCP is observed to be better using strain QM 9123. The protein content of biomass varied between 21-26% and up to 70% of straw is utilized in 48-144 h [37]. Production of SCP from cellulose by a thermotolerant strain of Aspergillus terreus in batch cultivation showed that 80-88% of available cellulose is utilized in 30-36 h and the biomass contains 23-38% protein. In semi-continuous cultivation studies in which 90% of the biomass is withdrawn at the end of growth cycle, 84% of added cellulose is utilized with the biomass containing 32% crude protein [38]. Chaetomium cellulolyticum has also been employed for the production of SCP [39-42]. Pretreatment of substrates show marked effects on SCP production from C. cellulolyticum. Chahal et al [39] obtained 0.074 g protein /I/h from growth of C. cellulolyticum using 1% wheat straw pretreated with caustic soda and peracetic acid. The growth of this organism on alkali pretreated hardwood saw dust solids has also suggested that pretreatment results in an increase in SCP production [41]. In an another study it has been shown that pretreatment of 1% corn stover with nbutylamine resulted in an increase in protein content of C. cellulolyticum from 0.8 g/I to 1.58 g/I with protein productivity of 0.088 g protein/I/h [42]. Chahal [40] has suggested an integrated process for food/feed and fuel production from biomass. In this process surplus hemicelluloses are converted by fungi T. reesei, C. cellulolyticum and Pleurotus sajor-caju in SCP. All these fungi utilized sugars found in hemicellulose fraction at pH 6.0. The SCP produced by these fungi contained 36-48% crude protein

309 on a dry weight basis which can be used as animal feed.

Table 1 Conversion of lignocellulosic material to SCP Microorganism

Substrate

Biomass

Protein

(g/I)

content

Reference

(%) Hansenula polymorpha

Synthetic mediaa

2.6

NT

[24]

Wheat straw

4.1

NT

[24]

15.9

47

[27]

7.4

35

[28]

SEHAL

12.1

28

[28]

SEHAL

12.3

28

[28]

Vinasse

15.8

50

[36]

Wheat straw hydrolysate

6.8

44

[40]

Wheat straw

7.0

36

[40]

4.9

48

[40]

hydrolysate

Candida blankii Trichosporon cutaneum 851 Aspergillus niger 426A Penicillium decumbens

Bagasse hemicellulose hydrolysate SEHAL b

114-2

Fusarium oxysporum var lini Chaetomium cellulolyticum Pleurotus sajor-caju Trichoderma reesei

hydrolysate Wheat straw hydrolysate

mixture of xylose, glucose and arabinose b steam exploded hemicellulose autohydrolysate liquor

31o Production of SCP using thermophilic microorganisms may offer some advantages as fermenter operation in a thermopilic temperature range prevents accumulation of bacterial and fungal contamination, results in low cooling and downstream processing costs, besides having faster growth rates. Humphrey and coworkers [43] cultured Thermoactinomyces sp. batchwise at high temperature (55~ and obtained a high growth rate. However, use of thermophiles has attracted relatively less attention for SCP production due to their lower biomass yield than mesophiles on comparable growth substrates.

3

MICROORGANISMS USED FOR SCO PRODUCTION

A large number of microorganisms are known to accumulate 20% or more of their biomass as lipid and have been termed oleaginous [44]. However for the biotechnological exploitation of microbial lipids it is necessary that it should be cheaper than the equivalant plant lipids or must offer some novel property not realizable in existing oil commodities. Alternatively, microbial lipids could be produced as an adjunct to another process e.g. waste processing [17]. Although algae, bacteria, yeast and fungi are all known to accumulate lipids in their biomass and have been described in detail by Ratledge [17], the present section summarizes those microorganisms that can accumulate fat, utilizing pentoses and pentose containing substrates. Table 2 lists some of the yeasts and fungi accumulating lipids utilizing xylose and other lignocellulosic materials. Yeasts are known to utilize hydrolysates of rice hull, rice straw [45,50], fruit and vegetable wastes [51], and domestic wastes [52], and thus can be employed for the production of SCO from these waste streams. Amongst oleaginous yeasts Rhodotorula gracilis [45] and Candida curvata [46,47] are known to utilize xylose as a carbon source. Yoon and coworkers [45] evaluated R. gracilis for its ability to utilize different carbohydrates as carbon source under same carbon:nitrogen ratio for the production of lipids. The lipid yield obtained was in the order glucose > sucrose > maltose > xylose > cellobiose. R. gracilis grown on cellulase hydrolysate of alkalitreated rice straw yielded relatively higher lipid however, the organism failed to grow when rice straw treated only with alkali is used as a carbon source [45].The lipid yield obtained from cellulase hydrolyzates of alkali-treated rice straw indicated that agricultural cellulosic wastes which contain appreciable quantities of pentose sugars such as xylose, arabinose and cellobiose can be utilized by this organism as cheap raw material for microbial lipid synthesis.

311 Table 2 Conversion of xylose and other lignocellulosic material to SCO Cell Biomass (g/I)

Lipid content (%)

Reference

Xylose Batch, Cellulose 60 h Rice straw"

4.5 6.0 9.0

62.1 39.0 53.9

[45]

Candida curvata D

Xylose

Batch, 90 h

9.9

48.6

[46]

Candida curvata D

Xylose

Continuous, 15.0 dilution rate 0.05h -1

37.0

[46]

Candida curvata D

Xylose

Continuous, dilution rate 0.06h -1

8.2

30.0

[47]

Cryptococcus albidus

Xylose

Batch

33.0 b

33.0

[48]

Aspergillus niger AS-101

Xylose Avicel Bagasse

Batch, 144 h

1.7 2.0 1.9

15.2 14.9 13.6

[49]

Microorganism

Carbon source

Rhodotorula gracilis NRRL-1091

Culture conditions

``Pretreated with 0.25 N NaOH solution by autoclaving at 121~ for 1 h, ground by ball mill and saccharified by cellulase. bBiomass yield g product/100 g substrate utilized.

Candida curvata, another oleaginous yeast, has been reported to efficiently convert glucose, lactose, xylose and ethanol into lipids and biomass [46]. Maximum lipid accumulation in batch culture with xylose as a carbon source on nitrogen limited medium has promoted further studies on the lipid accumulating ability of this organism [47]. Of ten oleaginous yeasts examined, only Candida curvata D was found

312 to have the ability to utilize glucose and xylose simultaneously. Thus this oleaginous yeast is capable of growing on hydrolyzed wood and straw wastes and can be used as a potential source of SCO [47]. In a single stage chemostat C. curvata D produced similar biomass yields, lipid contents and fatty acids on glucose and xylose mixed in varying proportions. Fatty acyl composition of Candida curvata D in batch and continuous culture has been studied and showed predominance of palmitic acid and oleic acid [46,47]. Cryptococcus albidus also exhibited a similar fatty acid composition of SCO. Fatty acyl composition of some of yeasts and fungi grown on xylose or lignocellulosic materials have been listed in Table 3.

Table 3 Fatty acid composition of SCO produced by yeasts and fungi Microorganism

Carbon source

Culture conditions

Relative fatty acid composition (%) 16:0 18:0 18:1 18:2 18:3

Reference

Candida curvata D Candida curvata D

Xylose

Batch, 96 h Continuous, dilution rate 0.04h -1 Continuous, dilution rate 0.06h -1 Batch

41

14

43

4

[46]

30

15

45

4

[46]

22

17

45

12

22

4

55

14

Batch, 144 h

8 9 10

6 6 5

25 23 26

49 52 51

Xylose

Candida curvata D

Xylose

Cryptococcus albidus Aspergillus niger AS-101

Xylose

Xylose Avicel Bagasse

4

[47]

[48]

5 6 5

[49]

Since hemicellulose is principally composed of pentosans such as xylan that represents 20-40% of most lignocellulosic agricultural residues [53], a series of lipid accumulating yeasts including Candida, Cryptococcus, Lipomyces, Rhodosporidium,

313

Rhodotorula and Trichosporon have been examined for their potential to saccharify xylan and accumulate triacylglycerol [54]. Of the genera tested only Cryptococcus and Trichosporon isolates saccharified xylan. Strains of Cryptococcus albidus have been employed for one step saccharification of xylan coupled with triacylglycerol synthesis. C. terricolus, a strain constitutive for lipid accumulation, lacked extracellular xylanase but assimilated xylose and xylobiose. Thus continuous conversion of xylan to triacylglycerol has been possible in cultures of C. terricolus supplemented with xylanase [54]. Although lipid accumulating ability of a number of fungi is well known [17], their ability to produce lipid, utilizing pentose sugars and cellulosic waste material is not well studied. Aspergillus niger AS-101, which utilizes different iignocelluiosic substrates efficiently in submerged as well as in solid state culture for the production of cellulase and SCP [55-58], has also been employed to study lipid accumulation [49]. The amount of lipid accumulated ranged from 14-17% on various carbon sources, namely glucose, xylose, avicel (microcrystalline cellulose) and bagasse (a natural lignocellulosic substrate). Interestingly unsaturated fatty acid comprised around 80% of the total fatty acid with predominance of linoleic and oleic acid (Table 3). Thus A. niger has potential for the conversion of lignocellulosic material into SCO in addition to SCP. Conversion of lignocellulosic materials to SCP has attracted considerable interest and many processes are known for SCP production using yeasts and fungi as described in Chapter 14 of this volume and elsewhere [11,40]. However, successful production of SCP invokes many problems. The major problems associated with SCP production from lignocellulosic substrates are the recovery of pretreatment reagents, production of enzymes with high activity for the hydrolysis of lignocellulosics and their reusability, efficient assimilation of pentoses by yeasts and effective use of lignin [11]. With growing concern for environmental pollution, it is necessary to effectively utilize lignin. Thus a combination of lignin assimilation by microorganisms with SCP production can provide an economically viable and environmental friendly process for SCP production from lignocellulosics. Although at present the main market for SCP is animal feed, our increasing knowledge of ethanol production from lignocellulosic materials is likely to contribute to SCP production for human consumption, as ethanol is an acceptable substrate for SCP that can be used for human consumption. It is well established that microorganisms accumulate lipids in their biomass, however the idea of production of SCO is relatively new and till now no commercial

314 addition, genetic manipulation of microorganisms may lead to production of tailor made lipids.

4

5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21

REFERENCES

Davis JB, Updegraf DM. Bacteriol Rev 1954; 18: 215. Beerstecher E. In: Petroleum Microbiology, Houston: Elsevier, 1954. Champagnat A, Vernet C, Laine B, Filsoa J. Science 1963; 197: 13. Mateles RI, Tannenbaum S. (eds)In: Single Cell Protein, Cambridge: MIT Press, 1968. Forage AJ, Righelato RC. Prog Ind Micobiol 1978; 14: 59. Laskin AI. Ann Rep Ferm Proc 1977; 1:151 Flickinger MC, Tao GT. Ann Rep Ferm Proc 1978; 2: 23. Cousin MA. Ann Rep Ferm Proc 1980; 4: 31. Davis P. (ed). In: Single Cell Protein, London: Academic Press, 1974. Tannenbaum SR, Wang DIC (eds) In: Single Cell Protein II, Cambridge: MIT Press, 1975. Tanaka M, Matsuno R. Enzyme Microb Technol 1985; 7: 197. de Pontanel GH. (ed.) In: Proteins from Hydrocarbons, New York: Academic Press, 1972. Watteeuw CM, Armiger WB, Ristroph DL, Humphrey AE. Biotechnol Bioeng 1979; 21: 1221. Moo-Yong M. Process Biochem 1977; 12: 6. Ratledge C. Enzyme Microb Technol 1982; 4: 58. Ratledge C. J Am Oil Chem Soc 1987; 64: 1647. Ratledge C. In: Microbial Lipids vol 2 (Ratledge C, Wilkinson SG eds.) London: Academic Press Ltd, 1989; 567. Wilkinsion JF. 21st Symposium of the Society of General Microbiology, New York: Cambridge University Press, 1971;15. Krug ELR, Lim HC, Tsao GT. Ann Rep Ferm Proc. 1979; 3: 141. Kreger-van Rij NJW. In Yeasts: a Taxonomic Study, 3rd Edn. Amsterdam: Elsevier Science Publishers, 1984; 45. van Zyl C, Prior BA, Kilian SG, Kock JLF. J Gen Microbiol 1989, 135: 2791.

315 22

Kostov V, Ratchev R, Lazarova G, Russeva L, Krasteva J, Ivanova V. Acta

23 24 25 26 27 28 29

Microbiol Bulg 1991;28: 51. Jwanny EW, Rashad MM, Moharib SA. J Basic Microbiol 1989; 29: 581. Feliu JA, Gonzalez G, de Mas C. Process Biochem Int 1990; 25: 136. Meyer PS, du Preez JC, Kilian SG. Syst Appl Microbiol 1992; 15: 161. Meyer PS, du Preez JC, Kilian SG. J Ind Microbiol 1992; 9: 109. Meyer PS, du Preez JC, Kilian SG. Biotechnol Bioeng 1992; 40: 353. Yinbo Q, Hongzhang C, Peiji G. J Ferm Bioeng 1992; 73: 386. Sahy LK, Wegner EH, Reiter SE. Dev Ind Microbiol 1983; 24: 305.

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Peitersen N. Biotechnol Bioeng 1975; 17: 361. Han YW, Dunlop CE, Callihan CD. Food Technol 1971; 25: 130. Han YW, Callihan CD. Appl Microbiol 1974; 27: 159. Han YW. J Ferm Technol 1982; 60: 99. Kristensen TP. Eur J Appl Microbiol Biotechnol 1978; 5: 155. Vinson LJ, Cerecedo LR, Mull RP, Nord FF. Science 1945, 13: 388. Silva MEST, Nicoli JR. J Ferm Technol 1985; 63: 91. Peitersen N. Biotechnol Bioeng 1975; 17: 1291. Miller TF, Srinivasan VR. Biotechnol Bioeng 1983; 25: 1509. Chahal DS, Swan JF, Moo-Young M. Dev Ind Microbiol 1977; 18: 433. Chahal DS. Biotechnol Bioeng Symp 1984; 14: 425. Pamment NB, Moo-Young M, Hsieh F-H and Robinson CW. Appl Environ Microbiol 1978; 36: 284. Tanaka M, Robinson CW, Moo-Young M. Biotechnol Lett 1983; 5: 597. Humphrey AE, Moreira A, Armiger W, Zabriskie D. Biotechnol Bioeng Symp 1977; 7: 45. Thorpe RF, Ratledge C. J Gen Microbiol 1972; 72: 151. Yoon SH, Rhim JW, Choi SY, Ryu DDY, Rhee JS. J Ferm Technol 1982, 60: 243. Evans CT, Ratledge C. Lipids 1983; 18: 623. Heredia L, Ratledge C. Biotechnol Lett 1988; 10: 25.

48 49

Hansson L, Dostalek M. J Am Oil Chem Soc 1986; 63:1179. Singh A. Experentia 1992; 48: 234.

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Watanabe D. Hakko Kyokaishi 1974; 32: 62.

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Glatz BA, Floetenmeyer MD, Hammond EG. J Food Protection 1985; 48: 574. Fustier P, Simard RE. Can Inst Food Sci Technol 1976; 9: 182. Saddler JN, Yu EKC, Mes-Hatree M, Levitin N, Brownell HH. Appl Environ Microbiol 1983; 45; 153.

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Fall R, Phelps P, Spindler D. Appl Environ Microbiol 1984; 47: 1130. Singh A, Abidi AB, Darmwal NS, Agrawal AK, Srivastava S. Ind J Biol Res 1988; 6: 1. Singh A, Abidi AB, Darmwal NS, Agrawal AK, MIRCEN J Appl Microbiol Biotechnol 1988; 4: 473. Singh A, Abidi AB, Darmwal NS, Agrawal AK, MIRCEN J Appl Microbiol Biotechnol 1989; 5: 451. Singh A, Abidi AB, Darmwal NS, Agrawal AK, World J Microbiol Biotechnol 1990; 5: 333.

12

Microbial Tolerance to Solvents and Organic Acids

1

INTRODUCTION

Adverse effects of fermentation end products on the performance of many bioprocesses constitute not only problems of fundamental biological interest but also has a practical dimension of considerable importance due to economic interest. Selection or construction of strains resistant to end product or manipulation of environmental factors contribute to the process optimization. Pentose metabolism yields a broad spectrum of products including ethanol, acetone, butanol, butanediol and organic acids. The nature and composition of these end products depend on the type of microorganism employed as well as the culture conditions used. While ethanol tolerance has received considerable scientific attention, our knowledge of microbial tolerance to other solvents is limited. Therefore, an overview of our general understanding of microbial tolerance to fermentation end products such as solvents and organic acids will be made in this chapter. Due to vast literature available on ethanol tolerance of yeasts, it is aimed to briefly discuss biophysical and biochemical mechanisms underlying toxic effects of ethanol. For detailed studies on ethanol tolerance, readers are referred to [1-3]. In addition, toxic effects of other end products such as butanol, butanediol and organic acids will be encompassed. Special attention is given to our current understanding of end product tolerance in pentose fermenting microorganism. Adaptive mechanisms leading to product tolerance are also discussed. Manipulation of plasma membrane of microorganisms by supplementing broth has yielded promising improvement in fermentation processes and has been included here along with strategies for genetic improvement of strains by selection of highly tolerant strains.

317

318 2

EFFECT

OF

SOLVENTS

AND

ORGANIC

ACIDS

ON

CELLULAR

PHYSIOLOGY OF MICROORGANISMS

End products of a fermentation exert a series of physical and biochemical effects on the catalytic activity of microorganisms leading to growth inhibition [1-3]. The major end products of pentose fermentation include ethanol, butanol, butanediol and organic acids. These organic solvents mainly influence membrane physiology of microorganisms by partitioning in lipid bilayers and then interfering with lipid-lipid and lipid-protein interactions [4-5]. The toxicity of a solvent depends on the polarity and the molecular weight as a solvent of low polarity and high molecular weight is expected to have the least toxicity [6]. Another more direct approach to establish a correlation between nature of solvent and its toxicity has been based on Iogarithum of partition coefficient, log P, of a given compound in the standard octanoi-water two phase system [7]. Cells are minimally inactivated by solvents with a log P> 4 [8-10]. In addition, water soluble and water immisible solvents exert different effects on microbial activities [1 1]. In order to reduce solvent toxicity, extractive fermentation has been employed for removal of fermentation products in situ [12-13]. In extractive fermentation, dilute solvent is stripped from the fermentative broth by continuous contact with an organic solvent into which the product can be absorbed. However, the extractant used should be selective for the product being recovered, immiscible in water, non-toxic to the microorganism, cost effective and should have a higher volatality as compared to the product [14]. Such systems have been employed for the production of ethanol [15-16], acetone-butanol [14], and organic acids [17-18]. Other strategies employed to reduce solvent toxicity are use of membranes to separate solvents from the cell containing broth [19-21], immobilization of microbial cells to reduce the contact of the immiscible solvents with microbes [22] and vacuum fermentation [23]. While the development of technical know how to overcome solvent toxicity is of considerable importance, the tolerance of strains capable of significant solvent production can not be ignored. In a fermentative process increasing the final concentration of solvent produced has attracted considerable interest as the energy consumed for distillation is a major factor in economic improvement of solvent production. With conventional technology of distillation, recovery of solvent from fermentation broth containing less than 6-8% ethanol or 3-4% butanol is not optimal in terms of energy recovery [24]. Table 1 lists various effects of solvents and organic acids on the cell physiology of microorganisms. In this section emphasis will be laid

319 upon the solvent tolerance of pentose fermenting microorganisms.

Table 1 Effect of various solvents and organic acids on cellular physiology of microorganisms Solvents/ organic acids

Organism

Effect on cellular physiology

Ethanol

Escherichia coil

Inhibits growth and cell division Promotes leakage of protons and nucleotides Inhibits lactose permease Increases membrane fluidity Inhibits growth and viability Inhibits fermentative activity Increases leakage of protons magnesium and nucleotides Increases membrane fluidity Inhibits growth and viability Alters lipid composition of cell membrane Inhibits glycolysis Inhibits growth and viability Alters fatty acid and sterol composition Increases membrane fluidity Inhibits ammonium, sugar and amino acid uptake Enhances passive proton influx and decreases glucose induced proton efflux

Ethanol

Ethanol

Ethanol

Zymomonas mobilis

Clostridium thermocellum

Saccharomyces cerevisiae

Reference

[25-27] [28] [25-29]

[28] [30] [31] [31-32] [33] [34] [35] [36] [3] [37-38] [39-40] [41-45]

[46-48]

320 Table 1 (Contd.) Effect of various solvents and organic acids on cellular physiology of microorganisms

Ethanol

Ethanol

Ethanol

Saccharomyces cerevisiae

Candida shehatae Pichia

stiptis Ethanol Butanol

Fusarium oxysporum Clostridium acetobutylicum

Leads to transient PM hyperpolarization and transient efflux of K§ Causes leakage of amino acids, nucleosides and nucleotides Decreases cytoplasmic pH Inhibits PM H§ activity Inhibits fermentative activity Leads to accumulation of cytochrome P450 Induces stress proteins Decreases the mean cell volume and total cell volume as well as viability Inhibits growth and production but ethanol yield is less sensitive Inhibits growth and fermentation Inhibits growth and fermentation Reduces cellular ATP content Alters phospholipid and fatty acid composition Increases membrane fluidity Inhibits uptake of glucose and xylose Inhibits activity of proton translocating ATPase

[49]

[5O] [47,51]

[52] [2] [53-55] [56]

[57]

[58] [59] [60,61] [62] [63-65] [63] [66]

[67]

321 Table 1 (Contd.) Effect of various solvents and organic acids on cellular physiology of microorganisms

Butanediol

Klebsiella pneumoniae Bacillus polymyxa

Acetic acid

Clo s tridiu m a ce to butylicu m

Acetic

C los tridiu m

acid Butyric acid

acetoaceticum Clostridium a ce to butylicu m

Inhibits biomass production but not the product yield Inhibits biomass production but not the product yield Inhibits growth Acts as uncoupler and allows protons to enter the cell Increases H§ activity and reduces cellular ATP content Inhibits growth Inhibits growth Diminishes pH gradient

[68] [69,70]

[71-74] [75]

[60-62]

PM; Plasma membrane

Yeasts are most tolerant to ethanol, but generally lack the ability to ferment pentoses. Pentose fermenting yeasts on the other hand are relatively less tolerant to ethanol. For example, pentose fermenting C. shehatae exhibits less tolerance to ethanol compared to other ethanol producing strains. C. shehatae grown on minimal medium with vitamins tolerated 5% added ethanol on D-xylose and 6% on glucose [76], whereas S. cerevisiae had tolerance to 11%, Kluyveromyces fragilis 8% [77-78] and Candida wickerhamfi 7.4% [79]. In another study it was observed that ethanol completely inhibits growth of C. shehatae under aerated conditions at a concentration of 37.5 g/I at 30~ [80]. However, under oxygen limited conditions addition of 25 and 50 g/I ethanol completely inhibited fermentation of D-xylose in C. shehatae and also resulted in a decline in cell viability [57]. Wayman and Parekh [81], have reported higher values of yield and ethanol tolerance when C. shehatae ATCC 2298 was used under semiaerobic conditions to convert whole barley hydrolysate, containing approximately 70% glucose and 30% xylose. Bioconversion of 260 g/I sugar solution

322 ceased at ethanol concentration of 100 g/I, but resumed once ethanol was removed by vacuum distillation. Bioconversion of 180 g/I sugar solution to 84 g/I ethanol within 72 h has been reported [81]. This suggested that yeast can tolerate 8% ethanol when xylose was substrate as all glucose is consumed before bioconversion of xylose starts. Ethanol at a concentration of 20 g/I begins to inhibit ethanol productivity and xylose consumption of Pachysolen tannophilus [82-83]. However, the concentration of ethanol that stops ethanol production is much higher than that is growth inhibitory viz. 42 g/l. Ethanol added at a concentration of 80 g/I results in specific productivity around 0.03 g/g/h which is half that observed in the absence of added ethanol. Slininger et al. [82] observed that although P. tannophilus can tolerate ethanol up to 100 g/I, they generally accumulate a maximum of 30 g/I ethanol in cultures even with excess of xylose. This suggested that ethanol toxicity is not the factor limiting ethanol accumulation when xylose is substrate. Influence of substrate on ethanol production is obvious as P. tannophilus produces more than 50 g/I of ethanol when glucose rather than xylose is used as substrate [84]. Based on kinetic studies it was noted that 64.3 g/I ethanol is the maximum concentration that allows cell growth of Pichia stipitis in the presence of xylose at 40 g/I [85]. The tolerance of P. stipitis Y7124 to added ethanol was evaluated in anaerobic and microaerobic conditions during the fermentation of a sugar mixture consisting of D-glucose 20%, D-xylose 75% and L-arabinose 5% [58]. It appears that the presence of oxygen plays an important role in determining ethanol tolerance. In microaerobic conditions, ethanol up to 20 g/I had no inhibitory effect on the fermentation capability of P. stipitis and it produced ethanol with yield up to 0.4 g/g and a specific rate of production of 0.1 g/g/h. An increase in the initial concentration of ethanol decreased the rate of ethanol production but yield appears to be less sensitive to ethanol inhibition. In anaerobic conditions, maximum fermentative ability is obtained at zero initial level of ethanol in culture. When initial ethanol concentration increases growth and ethanol production declines [58].

Fusarium oxysporum ATCC 10960 showed similar growth rates when ethanol was added either before inoculation or at the mid point of fermentation [86]. Ethanol induced inhibitory effects are seen at an ethanol concentration of 15 g/I and growth was inhibited above 42 g/I ethanol. However, substrates also influence inhibitory effects of ethanol as it was inhibitory to glucose fermentation by F. oxysporum VTTD80134 above 4.5% (w/v) and xylose fermentation above 3.5% (w/v) and the maximum ethanol concentrations achieved were 6.0 and 4.1%, respectively [87]. An increase in substrate concentration also results in decreased ethanol production. For example, 20% (w/v) glucose decreased fermentation rate with final yield of 4.5% in

323 7 days and 30% glucose resulted in 4.3% ethanol yield in 14 days [88]. On the other hand xylose at concentrations ranging from 15 to 20% increased the log phase but the fermentation rates remained unaffected [88]. F. oxysporum DSM 841 produced 3.6 g/I of ethanol on potato waste with an yield of 0.1 g/g. The exogenous addition of ethanol showed that above 30 g/I ethanol is growth inhibitory and about 50% growth inhibition was observed at 50 g/l. Only slight growth was observed at a concentration above 100 g/I [59]. Thermophilic bacteria known to ferment pentoses are less tolerant to solvents including ethanol. For example, clostridia generally produce less than 4% ethanol. This is mainly because they are extremely sensitive to solvent inhibition. Ethanol at 0.5% (w/v) causes 50% reduction in growth rate at 60~

in C. thermocellum [35] and

ethanol at 1.5% (w/v) concentration causes the same growth inhibition in C.

thermosaccharolyticum under similar conditions [89]. This low ethanol tolerance of thermophilic bacteria in both C. thermocellum and C. thermohydrolyticum is the major limitation in their usage for industrial ethanol production. Thermoanaerobacter ethanolicus JW 200 grown at 20% (w/v) sugar concentration ceases fermentation when ethanol concentration reaches 0.5% but cells do not die, they tolerate ethanol up to 9% (w/v) after accomodation [90]. It has been even found to grow at this concentration, after accomodation, although ethanol is not produced. C. thermocellum and C. thermosaccharolyticum can accumulate at least 3% ethanol in the medium when grown in co-culture. However, substantial amounts of acetate and lactate are also formed during such fermentations [91-92]. Co-culture of T. ethanolicus and C. thermocellum exhibited low ethanol tolerance and ethanol production over 1% in the medium has not been observed [90]. A number of ethanol-induced effects on the cellular physiology of clostridia are listed in Table 1. Butanol, another product of bacterial fermentation, is known to be highly toxic to cells and inhibits fermentation at concentration as low as 2%. In the industrial acetone-butanol fermentation process, solvent production ceases when concentration of butanol reaches 13 g/I [93]. It has been noted that in acetone-butanol fermentation, butanol is the primary toxic substance as acetone or ethanol do not reach the inhibitory

level during fermentation

[94]. The

higher toxicity

of butanol to

microorganisms has been attributed to its greater hydrophobicity. The addition of ethanol or acetone causes 50% growth inhibition of Clostridium at 40 g/I and cell growth completely ceases at 70 g/! concentration of acetone and 50-60 g/I ethanol [60,61]. Exogenous addition of 7-13 g/I butanol to cultures growing on hexoses resulted in 50% growth inhibition and at 12-16 g/I butanol completely inhibited growth. However, below threshold level, which appears to be 4-4.8 g/I growth inhibitory effect

324 of butanol was not observed [95]. The growth inhibitory effect of butanol was much severe when cell were grown on xylose as growth was inhibited at concentration 8 g/I butanol [66]. Butanediol is known to mainly inhibit biomass production and not the product yield. Klebsiella pneumoniae that is known to produce butanediol exhibits maximum growth rate at a butanediol concentration below 1 g/I and at higher concentrations specific growth rate decreases rapidly. However, butanediol concentration up to 80 g/I have little effect on specific rate of its formation [68]. Another study on shake flask with 8% initial xylose concentration showed that diol concentration above 65 g/I resulted in complete growth inhibition [96]. The influence of butanediol on Bacillus

polymyxa is of the same nature as on K. pneumoniae. Butanediol up to 20 g/I had no effect on the yield of diol from xylose in B. polymyxa [69]. Butanediol at higher concentration strongly inhibited growth of organisms but metabolic steps leading to its fermentation remain unaffected [70]. However, other end products of diol pathway such as ethanol and acetic acid at 1% concentration inhibited butanediol formation [97]. The production of alcohols is often accompanied by various weak organic acids including acetic, lactic, succinic and formic acid, in addition, production of fumaric and itaconic acid has also been occasionally reported [98]. Since these acids are present in fermentation broth, they contribute to overall inhibition of growth and metabolism. Analysis of end product of anaerobic fermentation of C. thermocellum has shown that alcohols and lactate decrease the optimum growth temperature but no such effect was observed with acetate, butyrate and beta hydroxy butyrate [99]. Although ethanol tolerant mutants exhibited slight tolerance to propionate, butyrate, lactate and beta hydroxy butyrate, they lacked resistance to acetate. This has suggested that mechanism of inhibition of organic acid is different from alcohols [99]. The cell membrane is freely permeable to weak acids in their undissociated form and act as uncouplers which allows protons to enter the cell from medium [71-74]. At sufficiently high concentrations undissociated acids result in collapse of the pH gradient across the membrane. The uncoupling action of these weak acids is counteracted by increased ATPase activity of cell thus leading to depletion of ATP reserve of cell which inhibits all the metabolic activity of the cell. In fact, C. thermocellumincubated with 0.8 M acetate has shown depletion of cellular ATP content [35]. Measurement of internal pH of C. acetobutylicum at different stages of growth has also revealed interesting results [62]. During log phase of growth, over 0.1 M total weak acids are produced and extracellular pH decreased from 6.0 to 4.8, but internal pH remained unchanged at pH 6.2 indicating that internal pH is not influenced by the normal levels of acid

325 produced. However, exogenously added butyric acid (0.17 M) diminished the pH gradient [62]. Butyrate at concentration 0.07 M [61] and 0.16 M [60] was found to inhibit growth of C. acetobutylicum by 50%. The observed difference in the two studies has been attributed to the use of buffered medium by Costa and Moreira [61]. The inhibitory effect of butyrate and alcohol appeared to be additive. The switch from acidogenic to solventogenic phase has been suggested to be an adaptive detoxification mechanism of cell [100]. In fact lack of this shift in metabolism leads to death of C. acetobutylicum due to the toxic effects of acummulated acid as end product [101]. In C. thermoaceticum which produces only acetic acid, cell death has been reported due to build up of end product to toxic level [102]. Acetic acid, both in ionized form (acetate) and in undissociated form (acetic acid) is growth inhibitory to C. thermoaceticum. This organism in a pH-controlled fermentation (with sodium hydroxide at pH 6.0) produces 56 g/I acetic acid, while in absence of pH control, pH decreases to 5.4 and maximum acetic acid produced was only 15.3 g/I [75]. To understand the mechanism of organic acid induced inhibition in

C. thermoaceticum, Wang and Wang [75] studied effect of various salts on growth rate. An inverse linear relation between the cell growth rate and the final cell concentration to sodium acetate concentration was monitored. Effect of various exogenously added salts on the relative growth inhibition had suggested that growth inhibitory effect of various anions is in order of acetate > chloride > sulfate and that of cations is in order of ammonium > potassium > sodium. It was observed that undissociated acetic acid at concentration between 0.04-0.05 M or ionized acetate at 0.8 M completely inhibits growth of C. thermoaceticum. Thus undissociated acetic acid is much more inhibitory than ionized acetate ion. In batch cultures of C. thermoaceticum using glucose as substrate it was observed that acetic acid levels above 10 g/I cause a reduction in specific growth rate and product formation [75,103104]. Inhibitory effects of acetic acid on xylose utilization and acetic acid production has also been demonstrated [105]. C. thermoaceticum grown in non-pH controlled batch culture at 55~ under head space of 100% carbon dioxide typically produced 14 g/I acetic acid during a 48 h fermentation in a medium contatining 2% xylose. However, in fed-batch fermentation 42 g/I acetic acid is produced by the organism after 116 h when concentration of xylose was maintained 2% and pH was controlled at pH 7.0. This has further provided evidence that main toxic form of acetic acid is its undissociated form. In addition to the effect of acetic acid on the cellular physiology of acetic acid producing organisms, its inhibitory effects have also been observed in yeasts including those known to ferment pentoses such as Pichia stipitis [106,107], Candida blankii

326 [108] and C. shehatae [109]. Inhibition of pentose fermentation by acetic acid is of great concern mainly due to the fact that hydrolysis of hemicellulosic part of lignocellulosic waste yields appreciable amounts of acetic acid as a decomposition product of acetylated sugars [110-112] and acetic acid is one of the major component present in hemicellulosic hydrolysate that inhibits pentose fermentation by yeasts. The toxic effects of acetic acid on yeast is common [113], because at the pH optimal for yeast fermentation (pH 4-5), acetic acid largely exists as undissociated acid and causes uncoupling effects, van Zyle et al. [107] observed that Pichia stipitis CSIRY633 (CBS 7126) is inhibited by acetic acid depending upon pH, acetic acid concentration and aeration. At pH 5.1 no ethanol production occurred when 10 g/I acetic acid was added while at pH 6.5 fermentation was only partially inhibited. A 50% inhibition of the volumetric rate of ethanol production occurred at acetic acid concentrations of 0.8 and 13.8 g/I at pH 5.1 and 6.5, respectively. While at pH 6.5, 15 g/I acetic acid reduced the ethanol production by 50%, similar inhibition was caused by 1 g/I acetic acid at pH 5.1. Tran and Chambers [111,114], however, observed that Pichia stipitis CBS 5776 is more tolerant to acetic acid as 11.9 g/I acetic acid reduced ethanol production by 76% at pH 5.0. This difference in the sensitivity to acetic acid has been attributed to the difference in the experimental conditions as under oxygen limited conditions used in the latter case some of the acetic acid may be utilized [107]. Lee and McCaskey [115] reported that 5 g/I acetic acid completely inhibited the growth of Pachysolen tannophilus at pH 3.0, 4.2 and 5.2 while Watson et al. [116] found that 7.4 g/I acetic acid caused only 50% inhibition of specific growth rate of this organism at pH 5.4. Candida shehatae growing on a minimal medium with vitamins and Dxylose as sole carbon source of energy tolerated acetic acid up to 0.4% (v/v) at pH 4.5 [109]. Under these conditions the temperature range of growth of C. shehatae squeezed from 5-34~ to 21-27~ and growth yield on D-xylose decreased to 64%. In addition tolerance to ethanol also dropped from 5% (v/v) to 2% (v/v) [109]. Based on the growth rates of C. blankii in carbon limited chemostat cultures du Preez et al. [108] suggested that inhibitory effect of acetic acid on growth is determined in part by ratio of xylose to acetic acid. Fusarium oxysporum 841 exhibited relatively higher tolerance to acetic acid as about 50% growth inhibition was observed at 40 g/I acetic acid and 100 g/I acetic acid completely inhibited growth of this fungi [59]. Lactate, which is the end product of glucose or xylose fermentation by Lactococcus lactis is known to inhibit lactate production depending on the substrate utilized [117].

327 3

ADAPTIVE MODIFICATIONS SOLVENT TOLERANCE

IN

MICROORGANISMS

LEADING

TO

Solvents produced by microorganisms after achieving higher concentrations act as chemical stress and adversely affect growth and metabolism. Therefore, microorganisms have developed various mechanisms to offset the deleterious effects of solvents present in the surrounding medium. One of the most common adaptive mechanisms involves alteration of membrane lipid composition which in turn influences physical properties of membrane and permits the survival of microbes in the presence of higher concentrations of solvents. Another mechanism involved in combating tolerance to solvents is induction of specific proteins termed as "stress proteins". Alternatively, microorganism may develop mechanisms to metabolize the solvents produced at a higher rate. Studies on various microorganisms suggest that adaptive mechanisms have been evolved by microorganisms that frequently encounter the presence of solvents in their natural environment. In fact some of these microorganisms have acquired specific membrane lipid composition and thereby tolerance to solvents.

3.1

Modification of lipid composition

Ethanol, which constitutes major solvent produced has been shown to elicit changes in membrane lipid composition of a number of microorganisms [118,128-131]. Table 2 lists adaptive modifications of lipid composition leading to solvent tolerance. E. coil has been employed as a model organism to understand the fundamental action of ethanol on microorganism. In addition, E. coli itself produces ethanol as a major fermentation product [132-133] and is thereby expected to evolve some resistance to the potential chemical stress of ethanol. E. coli grown in the presence of ethanol synthesizes membranes enriched in acidic phospholipids [26,119]. In addition the membrane lipid composition of ethanol sensitive mutant of E. coil has lower levels of acidic phospholipid while ethanol resistant mutant have shown higher proportions of acidic phospholipids [26].

328 Table 2 Adaptive modifications of lipid composition leading to solvent tolerance in microorganisms Product

Microorganism

Adaptive modifications

Reference

Ethanol

E. coil

Increase in acidic phospholipids and cisvaccenic acid and decrease in palmitic acid Decrease in phospholipid:protein ratio Induces stress proteins Posseses higher contents of cis-vaccenic acids and unusual hopanoids Alters lipid:protein ratio Induces stress proteins Posseses a novel C-30 dicarboxylic fatty acid Increase in the proportion of long chain cismono unsaturated fatty acids Decrease in saturated fatty acids (palmitic acid) and increase in unsaturated fatty acid (oleic acid) Decrease in total ergosterol Alterations in oleic acid contents Increase in phospholipid:protein ratio Increases ratio of saturated/unsaturated fatty acids

[26,118-119]

Z. mobilis

C. thermohydrosulfuricum Lactobacillus strains S. cerevisiae

Schizosaccharo -myces pombe

Butanol

C. acetobutylicum

[28] [120-121]

[122] [30] [123-124] [89] [125-126]

[37]

[38] [127]

[63]

329 These observations have implicated the possible role of acidic phospholipids in ethanol tolerance. Besides phospholipids, fatty acyl composition of E. coli grown in the presence of ethanol is also altered. An increase in the proportions of cis-vaccenic acid followed by corresponding decrease in palmitic acid was noted in E. coil exposed to ethanol suggesting involvement of cis-vaccenic acid in ethanol tolerance [118]. The role of fatty acids in ethanol tolerance was further substantiated by the observations that E. coli mutants defective in synthesis of cis-vaccenic acid were hypersensitive to killing and growth inhibition by added ethanol and this hypersensitivity was prevented by subsequent incorporation of cis-vaccenic acid to the E. coil membrane [134]. Incorporation of palmitic acid in the plasma membrane, contrary to ethanol induced changes, resulted in hypersensitivity to ethanol. It appears that ethanol-induced increase in cis-vaccenic acid fluidizes the membrane due to increased proportions of cis-unsaturated fatty acids as well as increases thickness of membrane that serves as an adaptive response to the organism [134]. These changes in fatty acyl composition of E. coil due to exposure to ethanol has been attributed to a shift in the synthesis of membrane lipids from saturated to unsaturated fatty acids which is reversed as soon as ethanol is removed. Ethanol induced changes in fatty acyl composition are also offset by supplementing saturated fatty acid such as palmitic acid. On the other hand unsaturated fatty acids such as palmitoleic acid (16:1) or oleic acid (18:1) supplementation could not offset ethanol-induced decrease in synthesis of palmitoyl residue. These observations suggested that reduced levels of saturated fatty acid on exposure to ethanol are due to decreased de novo biosynthesis rather than acylation/deacylation of existing lipids [135-136]. Evidence has also been provided that overall reduction in phospholipid:protein ratio observed in E. coil is beneficial to the organism for adaptation to ethanol induced fluidization [28]. Zymomonas mobilis, capable of rapid and efficient conversion of sugars to ethanol, can tolerate 12% (w/v) ethanol and exhibits a distinct plasma membrane composition. Its membrane lipid contains exceptionally higher amounts of cis-vaccenic acid and unusual hopanoids [122]. Contrary to E. coli, which on exposure to ethanol showed increased content of cis-vaccenic acid, Z. mobilis, under similar conditions did not show any dramatic change in phospholipid or fatty acyl composition in response to ethanol [30]. It was suggested that lipid composition of Z. mobilis has optimally evolved for survival in the presence of ethanol and thus presents a classical example of adaptation to ethanol [30]. However, different strains of Z. mobilis varying in their tolerance to ethanol exhibit differences in their fatty acyl composition [137]. For example, highly ethanol tolerant strain ZM481 contains low levels of short chain saturated fatty acids that are absent in less tolerant strains ZM1 and ZM4 and also

330 contains lower levels of cis-vaccenate. Strain ZM1, in contrast, contains higher levels of cis-vaccenate than both ZM4 and ZM481. These results have clearly indicated a correlation between strains of Z. mobilis exhibiting higher ethanol tolerance and those possessing increased levels of shorter saturated fatty acids compared to longer unsaturated fatty acids. These differences were accentuated in stationary phase cultures due to high ethanol contents [137]. Thermophilic ethanol producing bacteria Clostridium thermocellum and Clostridium thermohydrosulfuricum also exhibit ethanol-induced changes in their membrane lipid composition. C. thermocellum grown in the presence of ethanol synthesize shorter chain length monounsaturated and ante-iso branched chain fatty acids. However, analysis of fatty acyl composition of an alcohol resistant mutant of Clostridium suggested that increase in shorter chain length fatty acid is maladaptive [35]. C. thermohydrosulfuricum which is relatively more tolerant to ethanol as

C. thermocellum exhibits specific membrane lipid composition that consisted of a novel C30 dicarboxylic fatty acid [89]. Ethanol-dependent modifications in phospholipid fatty acyl composition has also been demonstrated in Saccharomyces cerevisiae [37]. Increasing concentrations of ethanol ranging from 0.5 to 1.5M leads to a progressive decrease in proportions of saturated fatty acids (mainly palmitic acid) and a corresponding increase in unsaturated fatty acid (mainly oleic acid) resulting in an increased fluidity of membrane as judged by their unsaturation index [37]. Ethanol-induced alterations in plasma membrane fatty acyl composition was also shown by Schizosaccharomyces pombe, although such changes were influenced by culture conditions as well. While ethanol grown aerobic cultures exhibit decreased contents of oleic acid, an increase in oleic acid content was monitored in ethanol grown anaerobic cultures [127]. Phospholipid: protein ratio increase under both anaerobic and aerobic conditions, probably via increased synthesis of phosphatidylinositol. These findings suggested that ethanol tolerance in Schizosacharomyces pombe is associated with higher content of oleic acid and ability to maintain higher rates of phospholipid synthesis [127]. Alterations in membrane sterols of S. cerevisiae in response to growth in the presence of ethanol may also be considered as an adaptive modification. S. cerevisiae exposed to ethanol exhibits a significant reduction in its total ergosterol content [38]. However, the percentage of ergosterol increase is due to decrease of other minor sterols such as zymosterol, fecosterol and lanosterols. Besides ethanol other end products of fermentation like acetone and butanol also induce alterations in the membrane lipids of fermenting microorganism for better survival in the solvent enriched environment. For example, Clostridium acetobutylicum, compared to

331 which is most extensively employed for acetone-butanol fermentation and therefore, has developed a mechanism to adapt to higher concentrations of butanol. Butanol-induced changes in membrane lipid of C. acetobutylicum exhibits an increase in the ratio of saturated to unsaturated fatty acids in both stationary phase solvent producing cells and vegetative cells grown in the presence of butanol (0.5 to 1.0%, v/v) [63]. This finding was further accentuated by the fact that a butanol tolerant mutant exhibits relatively lower levels of palmitic acid (C16:0) and higher levels of palmitoleic acid (C16:1) [138]. The increased ratio of saturated fatty acid in the membrane appears to be an adaptive response of cell to counter effect increase in membrane fluidity. The overall changes in membrane lipid composition of C.

acetobutylicum brought about during acetone-butanol fermentation are largely accounted for by solvent production [138]. However, pH also plays an important role in altering the lipid composition. A decrease in pH results in a decrease in unsaturated to saturated fatty acid ratio and an increase in cyclopropane fatty acid. This implies that end products of acetone-butanol fermentation have cumulative effects which lead to adaptation of the organism.

3.2

Induction of stress proteins

Amongst various solvents, ethanol accumulation in broth represents the most common natural environmental stress for fermenting microorganisms. Therefore, many organisms on exposure to high concentrations of ethanol trigger the synthesis of stress proteins [120,121,123,124,139]. Although stress proteins are known to be synthesized in all organisms under environmental stress and help to protect cells from potentially lethal challange, their precise mechanism of action is still not clear. Recent findings have suggested that all stresses may act through a common mechanism possibily by accumulation of aberrant proteins or misfolding of proteins [140-142]. Ethanol-induced synthesis of stress proteins has been demonstrated in E. coli [120,121], Z. mobilis[123,124] and S. cerevisiae[56,143]. In E. coli htpR is one ofthe regulatory genes involved in induction of stress proteins. Interestingly, mutation of this gene blocks both ethanol and thermal stress response [120-121] as well as reduces both thermal and ethanol tolerance suggesting a direct role of stress proteins in growth and survival [120]. Similar findings have been reported in yeast, S. cerevisiae where ethanol and heat shock induce a similar set of proteins [56]. In addition,

332 induction of heat shock proteins results in acquisition of thermotolerance, even if the proteins are induced by ethanol [143]. The presence of heat shock proteins also increases the viability of S. cerevisiae in the presence of ethanol [133]. Besides S.

cerevisiae, a number of other fungi are known to induce stress proteins on exposure to ethanol [139]. Another mechanism evolved to overcome ethanol stress involves induction of cytochrome P450 and increased ethanol metabolism [139].

MANIPULATION OF MEMBRANE LIPID COMPOSITION AND TOLERANCE TO SOLVENTS

That the plasma membrane is the first cellular component to come in contact with solvents produced, coupled with observations that a good correlation exists between hydrophobicity and solvent tolerance, together suggests plasma membrane lipids as prime target(s) of solvent toxicity [1]. This has led to a number of experiments exploring influence of added membrane lipid on solvent toxicity. Considerable evidence suggest that lipids such as phospholipids, sterols and fatty acids act as modulators of ethanol tolerance [144], tolerance to other solvents such as butanol has however attracted limited attention [67]. Table 3 depicts the role of membrane lipids as modulators of ethanol tolerance. The importance of lipids as modifier of ethanol tolerance in yeasts has come from a series of studies carried out with S. sake [145-150]. S. sake when grown in the presence of Aspergillus oryzae exhibits increased tolerance to ethanol [145]. In later studies the factor present in A. oryzae was identified as proteolipids and its phospholipid fraction was found to confer increased ethanol tolerance [145,146,149]. Jin and co-workers observed that addition of protein-phospholipid complex enhances the fermentation productivity of S. cerevisiae [151 ]. The role of individual phospholipids in enhancing resistance towards ethanol has come from studies on S. cerevisiae modified in plasma membrane phospholipid composition [39]. Amongst various phospholipids studied, phosphatidylserine was found to confer resistance to S.

cerevisiae cells with

regard to their fermentation

capability.

Probably the

anion:zwitterion ratio of plasma membrane phospholipid contributes towards ethanol sensitivity of yeast cell [39].

333 Table 3 Membrane lipids as modulators of ethanol tolerance Organism

Lipids supplemented

Product tolerance

Reference

Saccharomyces sake

Proteolipids

Increased growth, fermentative activity, and endurability Increased growth and endurability Increased growth and fermentative activity Increased ethanol production Increased growth and endurability but not the fermentative activity No effect on growth[149] endurability and fermentative activity Increased growth, endurability and fermentative activity Increased fermenta-tive activity and reduced fermentation time

[145,147, 148]

Increased ethanol productivity

[151]

(Aspergillus oryzae) Crude egg Yolk PC Purified PC

PC-albumin complex Ergosteroyl oleate or ergosterol + oleic acid Ergosterol or Oleic acid

Ergosteroyl oleate + egg yolk PC

Tween 80, ergosterol + albumin

S. cerevisiae

Protein-phospholipid complex

[149] [149]

[145] [149]

[149]

[158]

334 (Table 3 Contd.) Membrane lipids as modulators of ethanol tolerance

S. cerevisiae

PC, Palmitic acid and cholesterol

Increased growth

[159]

PS

Increased alanine uptake, proton efflux, fermentative

[39]

Ergosterol or Campesterol + linoleic acid Linoleic acid

Oleic acid or Linoleic acid or Linolenic acid

S. cerevisiae NSI113

S. uvarum Kluyveromyces fragilis

Linseed/cotton seed or soyabean oil or their fatty acid extract Linoleic acid or Tween 80 Ergosterol + oleic and linoleic acid

activity Increased viability and nutrient uptake Increased viability and nutrient uptake Seqential increase in alanine uptake, proton efflux and fermentative activity Increased fermentation rates

[152]

[41]

[40]

[163]

Increased ethanol production Increased growth rate and biomass production

[160]

[162]

Wine yeast

Yeast hull

Enhanced growth

Pachysolen tannophilus

(mixture of sterols and UFA Ergosterol, linoleic acid

rate and fermentative activity Increased ethanol production

[161]

[156]

and Tween 80 PC, phosphatidylcholine; PS, phosphatidylserine; UFA, unsaturated fatty acids

335 Fatty acids which form the hydrophilic core of membranes are also known to provide resistance towards ethanol [40,41,152]. Hayashida and his co-workers observed that egg-yolk phosphatidylcholine containing unsaturated fatty acids promote ethanol tolerance of S. sake, while dipalmitoylphosphatidyl-choline (with saturated fatty acid) are unable to do so [148-149]. Anaerobically growing S. cerevisiae, enriched with linoleic acid also acquire greater resistance towards ethanol compared to those enriched in oleic acid [41,152]. Using unsaturated fatty acid auxotrophic strains of S. cerevisiae and growing them aerobically in the presence of various unsaturated fatty acids, the importance of unsaturated fatty acyl residues in rendering cells tolerant to ethanol had been observed [40]. It is apparant that polyunsaturated fatty acids that provide greater fluidity to membrane are important determinants of ethanol tolerance in yeasts. The importance of optimal membrane fluidity in ethanol tolerance of yeast has been implicated by others using passive permeability of acetic acid as an index of fluidity [153]. Sterols have also been shown to be important in conferring ethanol tolerance to yeast cells [152]. Under anaerobic conditions, yeast cells, grown in the presence of unsaturated alkyl chain containing sterols (namely ergosterol and stigmasterol) were resistant to ethanol compared to those grown in the presence of saturated chain containing sterols (namely cholesterol and campesterol) [152]. This was attributed to the greater efficacy of barrier forming ability of unsaturated alkyl chain containing sterol molecules against the entry of ethanol into cells. Sterols are also known to provide endurability to yeast cells [149]. It is noteworthy that the viability of yeast cells in the presence of ethanol could be directly correlated to the presence of ergosterol in the membrane [154-155]. Thus it appears that ergosterol present in the membrane prevents the cells from deleterious effects of ethanol. Pachysolen tannophilus, one of the pentose fermenting yeasts, also exhibits enhanced ethanol production when nutrient media is supplemented with exogenously added lipids such as a mixture of ergosterol, linoleic acid and Tween 80 [156]. The maximum ethanol yield obtained for lipid supplemented culture was 0.32 g ethanol/g xylose consumed compared to 0.20 g ethanol/g xylose consumed in a control culture in which no lipids were added but grown under similar conditions [156]. In addition to ethanol, tolerance to butanol has also been observed in

Clostridium acetobutylicum grown in the presence of supplemented lipids [157]. The cell membrane fatty acyl composition of C. acetobutylicum was manipulated by supplying exogenous fatty acids in biotin deficient media. Cells grown in paimitic acidsupplemented media acquired hypersensitivity to butanol. Elaidic acid and eicosaenoic

336 acid supplemented cultures had greater and lesser tolerance to butanol, respectively. However, butanol tolerance was similar for oleic acid supplemented control cultures. Supplementation of saturated fatty acids to the medium that increased the ratio of saturated fatty acid in the membrane was found to increase butanol tolerance up to two folds and cell growth and ATPase activity were also increased [157]. This has provided clue that butanol tolerance can be modified by altering the membrane fatty acyl composition.

5

GENETIC BASIS OF TOLERANCE TO SOLVENTS AND ORGANIC ACIDS

Although pentose fermenting microorganisms yield a wide array of solvents and organic acids, our understanding of the genetic basis of product tolerance has been very limited. Since yeasts have been employed for ethanol production from time immemorial, most of the studies have concentrated on ethanol tolerance of yeasts. It is well known that various yeast strains differ in their tolerance to ethanol. For example some of the yeast strains used in sake fermentation are known to be more tolerant to ethanol than others. Although such observations have led to an understanding that ethanol tolerance is an intrinsic property of yeast but it is becoming increasingly apparent that a number of environmental factors also influence it [3]. In an attempt to understand the mechanism of ethanol toxicity and tolerance in yeast, analysis of single site mutations which confer ethanol sensitive phenotype has been carried out [164]. However, a detailed systematic analysis of such mutation sites is required to delineate the genetic basis of ethanol sensitivity. Further, genetic studies indicate that ethanol tolerance/sensitivity in yeast is controlled by a number of genes [165-169]. This polygenic control of ethanol tolerance was suggested on the basis of observations that segregants derived from diploid S. cerevisiae differ in their ability to tolerate ethanol. While none of haploids exceeded parental levels of tolerance, some of the crosses between haploids exceeded parental levels of tolerance [164]. At least four different genes have been implicated to be involved in ethanol tolerance [164]. Polygenic control of ethanol tolerance has hampered the isolation of ethanol tolerant yeast mutants as mutations in a number of genes is required to improve ethanol tolerance. This has restricted conventional use of agar plate screening procedure. In addition yeasts being highly ethanol tolerant is liable to only relatively small improvements in their tolerance [170]. Other approaches such as selection in continuous culture and

337 hybridization has been frequently used to generate ethanol tolerant mutants. A continuous feed back control system for isolation of ethanol tolerant yeast has been devised [171]. In this selection procedure the fermentative activity of culture is monitored continuously by using infrared analyzer to determine the concentration of CO2. This signal is fed to a potentiometeric controller which regulates a peristaltic pump that added 70% (v/v) ethanol into culture vessel, on production of more CO 2. Mutants are isolated from culture vessels by selecting cells which form colonies on 12% ethanol plates. An interesting aspect of this procedure is that the selection regime never exposed the cultures to ethanol concentrations above 5% (w/v). Nonetheless, mutants selected survived and fermented at enhanced rate in the presence of far higher concentrations of ethanol [171 ]. Another approach for selecting highly tolerant wine yeast was followed in pH-regulated continuous culture [172]. Hybrids between naturally occuring wine yeast strain and a laboratory strain were subjected to competition experiments in pH-controlled continuous cultures with increasing concentrations of ethanol over a wide range. The continuous culture system was obtained by controlling dilution rate of chemostat connected to a pH meter. The nutrient pump of chemostat was switched on and off in response to the pH of culture, which was kept near a critical external pH. Under these conditions, when the medium was supplemented with ethanol, the ethanol concentration of culture increased with each pulse of dilution. Using this method a highly tolerant hybrid strain of yeast was selected which was able to grow up to 16% (v/v) ethanol [172]. Protoplast or spheroplast fusion technique was employed by a number of workers to enhance the ethanol production capability [173-176], but this method has gained very limited success in obtaining both high ethanol producing capability and enhanced tolerance [177]. Genetic characteristics of hybrids selected between homothallic and heterothallic strains have led to some interesting insights on the ethanol production and tolerance [167]. The most tolerant spores did not generally produce ethanol at high concentration and on the other hand ethanol sensitive strains produced high levels of ethanol. This has led to the conclusion that ethanol tolerance and ethanol producing ability of yeast are independent from each other and these characteristics segregate independently [167]. In addition to yeasts, ethanol resistant mutants of E. coil [25,26], Z. mobilis [137] and Clostridia [34,89,178], are known. Some of the solvent tolerant mutants are listed in Table 4. In E. coli improvements in resistance to growth inhibition were quite modest and mutants were unable to grow on ethanol concentration above 5.5% (w/v) [25,26]. Ethanol tolerant mutants of Z. mobilis were isolated by Rogers and co-workers using parent strain ZM4 by two different strategies [137]. In one of the experimental

338 scheme, exponentially grown ZM4 strain was exposed to N-methyI-N'-nitro-Nnitrosoguanidine (NTG) for a period of 30 minutes and after washing the cells were inoculated into glucose medium and continuous cultures were carried out at 30~ with a dilution rate of 0.1h 1. Samples drawn from continuous culture were plated on glucose-agar medium supplemented with ethanol. Based on this experiment, strain ZM444 was selected which showed prolonged survival at higher ethanol level compared to ZM4. The other scheme followed two sequential NTG mutagenesis and selection of strains on agar plates containing ethanol up to 120 g/l. The selected strain ZM481 showed increased viability compared to ZM4. Thermophilic anaerobes C. thermocellum and C. thermosacchrolyticum were improved for the production of ethanol from cellulosic biomass [179]. C. thermocellum ferments hexoses to ethanol in low yields producing large quantities of lactic and acetic acid. The growth of C. thermocellum ATCC 27405 is inhibited by 50% in the presence of 8 g/I ethanol. Adaptation and selection of strains for ethanol tolerance was achieved by serial transfer during log phase of growth of strain fermenting cellobiose, xylose or solka floc with increasing concentrations of ethanol up to 40 g/l. Using this approach ethanol tolerant strain S-4 [180] and S-6 [179] have been isolated. In addition to higher ethanol tolerance these isolates showed that ratio of ethanol to acetate was favourably increased. S-6 strain was further improved by selecting lactate dehydrogenase negative mutants after NTG mutagenesis. C. thermosacchrolyticum (HG-2) was initially isolated as stable contaminant in a culture of C. thermocellum. This organism has ability to metabolize pentoses but shows poor ethanol tolerance and productivity. However, adaptation and selection for increased ethanol resistance by serial transfer on ethanol containing media yielded a superior alcohol resistant strain (HG-3) [179]. This selection strategy also improved the ratio of ethanol to acetate produced by these strains grown on glucose from 1:1 to 2:1 in case of HG-3 and selection for low acid producing strains allowed isolation of HG-4 which produced ethanol: acetic acid in a ratio of 4:1. Further selection for mutants resulted in a new strain

HG-6

which

forms

lower

amounts

of

acidic

products

[91].

C.

thermosaccharolyticum strain HG-6 and its derivative HG-6-610 produce up to 27 g/I of ethanol, less than 2.6 g/I acetic acid and less than 2 g/I lactic acid [92]. Thus it was possible to select high ethanol tolerant and high ethanol yielding clostridial strains. Herrero and Gomez [181] also isolated ethanol resistant strain C9 from wild type C. thermocellum ATCC 27405 by enrichment technique. Parent cultures grown on cellulose broth with 5g/I ethanol for 120 h at 60~ were selected on the basis of their higher growth and subjected to sequential transfer to fresh medium with increasing concentration of ethanol. After ninth transfer a strain capable of growing in ethanol at

339 Table 4 Solvent / organic acid tolerant mutants Solvent/ Organic acid

Organism

Method for selection/ isolation

Mutants/ isolates

Reference

Ethanol

Z. mobilis

NTG mutagenesis and continuous culture Two sequential NTG mutagenesis Adaptation and selection

ZM444

[137]

ZM481

[137]

s-4

[18o]

S-6 C-9 HG-3 HG-4 HG-6 HG-6-610 JW200

[179] [181] [179]

SB154 SB155 SB159 SB160 FDHS

[171]

[172]

SA- 1

[185]

904

[186]

lyt-1

[187]

S-3

[103]

1745

[103]

Z. mobilis Clostridium thermocellum ATCC 27405 C. thermosaccharo-

Adaptation and selection

lyticum UV-mutagenesis Thermoanaerobacter ethanolicus Selection in Saccharomyces continuous cerevisiae culture Wine yeasts Butanol

Clostridium acetobutylicum ATCC 824 C. acetobutylicum 903

Acetic acid

Clostridium thermoaceticum

Selection in continuous culture Selection in increasing concentration of butanol NTG mutagenesis EMS mutagenesis Adaptation and selection EMS mutagenesis

[91] [90]

NTG, N-methyI-N'-nitro-N-nitrosoguanidine; EMS, ethyl methane sulfonate; UV, Ultraviolet.

340 25 g/I was obtained [181]. Mutants of Thermoanaerobacter ethanolicus JW200 were selected after exposure to UV-light and to media supplemented with high iron. These mutants produce up to 3% ethanol from 7% starch and tolerate up to 10% ethanol with only 50 to 60% reduction in growth rate after a lag period. Addition of 1 to 2% ethanol did not prolong the lag period and growth proceeded at faster rate than control cultures [90]. Tolan and Finn [182] have studied ethanol production and tolerance of pentose fermenting Erwinia species. Screening of a large number of Erwinia species has shown that E. chrysanthemi B374 is most ethanol tolerant species and can tolerate 4% ethanol. Genetic improvement of this species for ethanol production and tolerance was attempted by expressing pyruvate decarboxylase (pdc) gene of Z. mobilis. Transconjugants showed an increased ethanol yield from 0.72 to 1.45 mole/mole of xylose and a decrease in production of toxic products such as formate. However, ethanol tolerance decreased from 4% to 2%. Similar results were obtained when Z.

mobilis pdc gene was expressed in E. coil as ethanol yield increased from 0.44 mol/mole to 1.66 mol/mole glucose but level of ethanol tolerance was as low as 0.23%. Nonetheless, when pdc gene of Z. mobilis was expressed in enteric bacteria Klebsiella planticola, a marked concurrent decrease in yields of formate, lactate, acetate and butanediol was observed and transconjugants tolerated ethanol up to 4% [183]. Butanol tolerant mutants of Clostridia have also been obtained using chemical mutagenesis. NTG and EMS were employed to isolate mutants of C. acetobutylicum [184]. Unfortunately mutants isolated in the presence of inhibitory concentrations of butanol could not produce higher concentrations of butanol in the non-growing phase [95]. A butanol tolerant mutant (SA-1) of C. acetobutylicum ATCC 824 [185], had characteristics similar to autolysis deficient (lyt-1) mutant. Although this mutant produced more butanol than parental strain, the acetone production was lower. Thus the overall solvent production decreased. Using chemical mutagenesis, Hermann et al. [186] isolated butanol resistant strains from C. acetobutylicum 903. One of the mutant 904 produced 30 to 40% higher concentration of solvents. Interestingly, this mutant was stable and maintained butanol tolerance and higher yields over a period of several years even in the absence of selection pressure. An acetate tolerant strain $3 was derived from C. thermoaceticum (Wood) culture after adaptation and selection on sodium acetate without mutagenesis. Another strain 1745 reported to be acetic acid tolerant was derived from the C. thermoaceticum (Ljungdahl) culture after EMS mutagenesis and selection on 2% sodium acetate [103]. In the wild type C. thermoaceticum, biomass production stopped

341 when gross acetic acid level (initial acetic acid plus net acetic acid production) reached about 10 g/I (9.3-12.5 g/I) whereas acid tolerant $3 strain at pH 6 and initial acetic acid concentration 12.5 g/I attained a gross concentration of 18 g/l. Another strain 1745 at pH 6 with initial acetic acid concentrations of 5.3 g/I attained gross concentration of acetic acid 13.8 g/I whereas at pH 7 and no added acetic acid it attained gross concentration of 14 g/l. In this strain acetic acid production continued even after net growth stopped and reached a level of 15 g/I at pH 6 and about 20 g/I at pH 7 [103]. The higher concentration observed at pH 7 has been interpreted as a greater sensitivity of the strain to acetic acid as compared to acetate ions. However, a detailed programme of selection of strains tolerant to acids is required for genetic improvement of strains for acid production. In retrospect, it is apparent that while various technical methods employed to overcome the toxicity of solvents produced during fermentation processes have resulted in increased production of solvents, improvement of strains with higher tolerance to solvents is also necessary for greater product formation. The search for the microorganisms tolerant to solvents and organic acids has geared up mainly due to economic reasons. In fermentative processes the energy consumed for distillation is the major economic factor and with conventional technology of distillation recovery of solvents from fermentation broth with less amounts is not optimal in terms of energy recovery and thus necessitates use of microorganisms with higher tolerance for solvent production. Studies on the effects of end product on cellular physiology of solvent producing microorganisms provide the basic understanding about the mechanisms underlying growth and product inhibition. While effects of ethanol on yeast are fairly known, effects of other solvents like butanol, butanediol, acetone etc. are still far from clear. The complexity of the mechanisms of solvent induced toxic effects on microbial cells has limited the development of a common approach to the development of solvent tolerant microorganisms. Environmental factors such as temperature and osmotic pressure play a crucial role in determining ethanol tolerance of yeast and other microorganisms, nevertheless yeasts are most tolerant to ethanol. However, pentose fermenting yeasts are not as tolerant to ethanol as bakers yeasts and show a great dependence on the type of substrate, substrate concentration, temperature and oxygenation conditions. Amongst mycelial fungi Fusarium oxysporum exhibit considerable tolerance to ethanol as well as acetic acid. Thermophilic bacteria exhibit poor tolerance to ethanol and other solvents like butanol and butanediol. Butanol production from bacteria has been limited mainly due to low tolerance of bacteria to these alcohols. Butanediol, another product of bacterial fermentation inhibits biomass production but not the product yield. Organic acids produced during

342 many bacterial fermentations also have an inhibitory effect on cellular physiology of the organism. Organic acid induced inhibition has been observed in the organisms producing these acids as well as in a large number of yeasts. This has acquired greater significance in case of fermentation of hemicellulosic hydrolysates which are known to contain significant amounts of these acids and thus prove to be inhibitory to pentose fermenting organisms. Microorganisms traditionally employed in solvent production develop adaptive mechanisms to tolerate these solvents. These adaptive mechanisms generally lead to modifications in plasma membrane lipids of the organism. The supplementation of lipids in fermentation media has yielded significant improvement in tolerance to ethanol and product yield. However, this approach has not been widely employed for improving tolerance of other solvents. Efforts have also been made to genetically improve strains for their tolerance to solvents and a few genetically improved resistant strains that can tolerate higher concentrations of ethanol or butanol are known. However, the genetic complexity due to polygenic control of ethanol tolerance/sensitivity has hindered the development of new strains as any genetic approach employed needs to modify a number of genes to improve ethanol tolerance. Our understanding of the genetic mechanisms involved in the tolerance of other solvents is not clear and needs to be investigated. With our practical knowledge of the use of lipids in the improvement of ethanol tolerance and increasing knowledge of lipid biosynthetic pathways it may be possible to clone genes of fatty acids and other lipids known to improve ethanol tolerance. Acid tolerance of the organism may also be improved with our basic understanding of the mechanisms involved in maintenance of intracellular pH. Plasma membrane ATPases known to pump protons have been suggested to be important in controlling acid tolerance. With the advent of molecular genetic approaches and increased understanding of biochemical principles underlying the solvent tolerance it is hoped to develop strains highly tolerant to solvents.

6

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13 Genetic Improvement of Pentose Fermenting Microorganisms

1

INTRODUCTION

Pentose fermentation suffers from a number of limitations, the major limiting factors being the rate of bioconversion, product yield and tolerance to solvent produced [1,2]. In addition, a number of ethanologenic microorganisms lack the ability to utilize pentoses. Thus improvement of microbial strains is necessary to make the fermentation process economically competitive. Strategies for the improvement of strains for pentose fermentation constitute selection and isolation of mutant strains as well as recombinant DNA techniques to restructure the metabolic network for the bioconversion of pentoses to desired products. So far major efforts have been directed towards genetic improvement of strains for the bioconversion of pentoses to ethanol. Studies have also been made, albeit to a lesser extent, on genetic improvement of strains for the bioconversion of pentoses to other solvents such as acetone and butanol. However, studies on genetic improvement of microbial strains for the bioconversion of pentoses to xylitol, organic acids, single cell protein (SCP) and single cell oil (SCO) have been very limited. The major targets for genetic improvement of microorganisms employed for pentose bioconversion are: 1 Improve efficiency of pentose bioconversion. 2 Increase productivity by: a Rate of substrate conversion. b Raising fermentation temperature. c Improving tolerance to solvents. d e

Increasing utilization of pentoses in the presence of other hexoses. Minimizing the by-product formation.

3 4

Increase resistance to natural inhibitors present in lignocellulosic hydrolysates. Broadening the pentose utilizing ability of solventogenic microorganisms. Above mentioned approaches are general and genetic improvement(s) of these targets, individually or in combination with others result in improvement of strains. However, approaches to improve strains for pentose bioconversion to ethanol or 351

352

acetone-butanol or butanediol or organic acids or SCP and SCO may involve some additional strategies and have been included in this chapter. Genetic studies to understand pentose uptake and their regulation in the presence of other hexoses are described in Chapter 4 of this Volume. Solvent tolerance of microorganisms and strategies employed for their improvement have been discussed in Chapter 12. The thrust of this chapter is to emphasize research advances in classical and molecular genetics aimed at the improvement of pentose fermenting microbial strains by redesigning the metabolic pathways. Basic approaches employed to improve pentose fermentation are screening, mutation, recombination and gene cloning. Each method has its distinct advantage and in some cases combination of more than one method may be employed to improve the strain. For example, mutation is the simplest of all approaches and requires little knowledge of the genetics and physiology of pathways involved. It predictably leads to rapid improvement of strains. Genetic recombination is another approach widely used as an adjunct to mutagenesis when several lineages of mutants are established. It provides a valuable tool to construct strains with different combinations of mutations that influence product yield. Protoplast fusion is a relatively simple approach to recombine the properties of a wide variety of microorganisms for which biosynthetic pathways and genetics are relatively less understood. Gene cloning is employed for those microorganisms for which biochemical pathways and genetics is relatively well understood. Various procedures used to carry out mutation, recombination and gene cloning in pentose fermenting microorganisms are included in this chapter.

2

SCREENING AND MUTAGENESIS

Screening is an essential, most direct and the least expensive means of improving microorganisms for industrial purposes. Screening is a general method and does not require biochemical and genetic information of the organism. Enrichment of cultures provides a suitable environment for the growth and reproduction of specific microorganisms while at the same time being inhibitory or lethal for non-target microorganisms [3,4]. Various methods of selective enrichment of microorganisms have been employed for isolation of microbes for lignocellulosic bioconversion. Such approaches have been discussed in this section. In general, the screening protocol

353 involves choice of organism, induction of genetic variability in the cell population followed by fermentation in small scale of many individuals from the population and then assay of fermentation product to identify an improved strain [3,4]. The same general protocol of selection is followed for many survivors of mutagenesis as chances of finding a more productive variant exist. In designing a mutation protocol for a particular microorganism, it is important to choose an effective mutagen. For each mutagen and each organism, there is a combination of mutagen concentration, time of exposure and conditions of treatment which produces the highest proportions of a particular class of mutants [3]. Both direct and indirect mutagens are employed for such purposes. While direct mutagens cause mutation by mispairing mechanisms involving either template or nucleotide precursors, indirect mutagens act by inducing a post replication repair system prone to error [5]. After mutagen treatment cells are allowed to undergo a period of DNA replication and cell growth so as to convert the damaged DNA into stable, altered DNA encoding reproducible and inheritable mutations. Table 1 lists characteristics of some of the microbial mutants isolated for bioconversion of lignocellulosics to solvents. Using ultraviolet-irradiation mutagenesis (indirect mutagen), Gong and coworkers [6], isolated a mutant of Candida sp., Candida XF217 which produces five times more ethanol than the parental strain. The parent strain Candida sp. C2, produces xylitoi as the major product whereas the mutant strain accumulated ethanol as the major product at the expense of xylitol [7]. Candida sp. XF217 produces ethanol from xylose both aerobically as well as under oxygen limiting conditions. However, oxygen is required for cell growth and xylose consumption [8]. The specific activities of enzymes of xylose catabolism, such as xylitol dehydrogenase and xylulokinase is increased in mutant strain as compared to parental strain whereas the activity of xylose reductase remains unaltered. It appears that increased xylitol dehydrogenase and xylulokinase activities of the mutant enabled the shift of the catabolic pathway from xylitol to ethanol production. Thus instead of excreting xylitol as the final product, these mutants convert more xylitol to D-xylulose and ultimately to ethanol.

Pachysolen tannophilus, a pentose fermenting yeast has been studied in relatively greater details for the genetic improvement of strain for bioconversion of pentose to ethanol. Jeffries [9] isolated mutants of P. tannophilus by UV mutagenesis followed by enrichment in nitrate and xylitol as sole source of nitrogen and carbon respectively. Isolated mutants exhibit a higher rate of ethanol production and yield from xylose, however, their performance also depends on the oxygenation conditions. Under aerobic conditions mutants produced ethanol from xylose twice as faster as

354 their parent strain whereas under anaerobic conditions producion rate is only 50% faster than the parent. The yield of ethanol from xylose also differs under these conditions. While mutants have better yield than parent under aerobic conditions, the yield remains unchanged under anaerobic conditions.

Table 1 Characteristics of some microbial mutants isolated for bioconversion of lignocellulosics to solvents Mutant

Method of mutagenesis/ selection

Characteristics of Mutants

Reference

Candida sp.

UV

Accumulates ethanol at the expense of xylitol

[6,7]

UV, Enrichment with nitrate and xylitol UV, Enrichment with ethanol Chemical

Higher rate of ethanol production

[9]

Enhanced ethanol accumulation Increased ethanol production and yield

[10]

XF 217

Pachysolen tannophilus Pachysolen tannophilus Klebsiella pneumoniae

[12]

MB-16, MB16-1048

Clostridium acetobutylicum SA-1

Clostridium acetobutylicum SA-3

Clostridium acetobutylicum

Selective enrichment with butanol Selective enrichment with butanol and heat shock Tn 916

Butanol tolerant Butanol producer

[13]

Butanol tolerant Butanol producer

[14]

Granulose and sporulation negative

[15]

355 UV-induced mutants of P. tannophilus have also been selected on the basis of diminished growth on ethanol [10]. The strategy behind the selection procedure was based on the enhanced ethanol accumulation by minimizing losses that occur due to its oxidation. Eleven independent mutant loci that conferred the ethanol defective phenotype have been identified and three of them viz. eth 1-1, eth 2-1 and eth 1 1 sup produce significantly more ethanol than the wild type in aerobic batch cultures [10]. One of these mutants, eth 2-1 produced less xylitol and lacked the malate dehydrogenase activity which is required for metabolism of two carbon compounds. In a subsequent study, using UV-mutagenesis, James et al. [11] isolated another class of P. tannophilus mutants. These mutants, selected on the basis of their defective growth on D-xylose retained their ability to grow normally on D-glucose. Of thirty mutants isolated, ninteen were genetically analyzed. Classical genetic analysis by back crossing mutants with the wild type and their segregants suggested that mutations occurred at nine distinct loci and many more loci are susceptible to mutation. Apparently, these mutations are pleiotropic in nature, and the expression of some of them is susceptible to nutritional conditions and genetic background. Interestingly, mutations at several loci resulted in poor growth of organism on at least one compound that is either an intermediate of tricarboxylic acid cycle such as succinate or 2-oxoglutarate, or are metabolized via this cycle such as ethanol or glycerol. Thus growth on D-xylose involves oxidative metabolism in which tricarboxylic acid plays a role. Biochemical characterization of these mutants suggest defect in one or more xylose catabolizing enzymes such as xylose reductase, xylitol dehydrogenase and xylulokinase [11]. The xyll and xyl2 alleles affected the level of xylitol dehydrogenase activity. Mutants exhibited less than 9% xylitol dehydrogenase activity compared to that of the wild type. These mutants grew normally on D-xylulose but failed to grow on D-xylose and xylitol. This inability of mutants to grow on D-xylose has been attributed to a defect in utilization of xylitol, which is a product of D-xylose reductase [11]. Since xylitol is accumulated as by-product of ethanol during pentose fermentation, xylitol dehydrogenase negative mutants are expected to improve the productivity of strains as mutants do not produce xylitol at the expense of ethanol. Strains of P. tannophilus bearing xy113 mutations had significantly lower activity of the D-xylose reductase enzyme and were further characterized by Schneider et al. [16]. These mutants exhibit slower growth rates (one fifth) as compared to the wild type on D-xylose and L-arabinose as both pentoses require D-xylose reductase for their catabolism [17,18]. However, they grew at the same rate as wild type on xylitol and D xylulose that enters the xylose catabolic pathway subsequent to D-xylose reductase

356 Selective enrichment of growth medium with increasing concentrations of butanol has been a method of choice for isolating Clostridium acetobutylicum with greater butanol producing abilities (Table 1). In addition a number of mutants of clostridial strains have been isolated by using chemical mutagenesis [19,20]. It appears that indirect mutagens such as UV and mitomycin are ineffective in producing mutations in C. acetobutylicum. The inability of UV radiation to cause mutations in clostridial strains has been attributed to lack of error prone repair system in Clostridium that is required for indirect mutagenesis. Ethyl methanesulphonate (EMS) has been widely employed as a mutagen for isolating variety of C. acetobutylicum mutants with characteristics such as acid tolerance, autolysis resistance, allyl alcohol resistance, lack of sporulation etc. and have been described elsewhere [19,20]. Mutants resistant to solvents have been discussed in Chapter 12. The conjugative transposon Tn 916 has also been employed for mutagenesis of C. acetobutylicum ATCC 824 [15]. Screening of mutants for loss of granulose synthesis exhibited five classes of mutants that contained a single transposon insertion and differed in their solvent producing abilities. Class 1 mutants lacked the activity of enzyme induced during solventogenesis and failed to produce acetonebutanol and were assigned as regulatory mutants. Class 2 mutants did not produce acetone but synthesized small amounts of butanol and ethanol. Class 3 mutants produced low levels of all solvents. Class 4 and 5 mutants produced essentially the same or higher amounts of solvents than the parent strain. Thus multiple regulatory elements are required to induce solvent production and sporulation in C.

acetobutylicum. Mutants of Klebsiella pneumoniae have also been isolated in an attempt to improve kinetics of ethanolic fermentation of D-xylose. Mutants MB16 and MB 16-1048 showed a high ethanol productivity and a short fermentation period compared to wild type [12]. Mutants isolated for the bioconversion of lignocellulosic biomass to single cell protein or single cell oil have been described in Chapter 11 of this volume.

3

GENETIC RECOMBINATION

3.1

Hybridization

Genetic recombination of S. cerevisiae has been well studied, however, our understanding of such studies on pentose fermenting yeasts are relatively poor. From

357 the published data it is evident that in principle, hybridization or cross-breeding is possible that has been widely employed for improvement of brewing yeasts [21]. The major problems associated with non-conventional yeasts are poor mating ability, poor sporulation, spore viability, homothallism, aneuploidy, polyploidy and polygenic control [21]. Nevertheless, classical hybridization techniques have been developed to increase chromosome number in pentose fermenting yeasts as an approach for improvement of strains. James and Zahab developed techniques for construction of diploids [22] and polyploids [23] of P. tannophilus a homothallic organism with predominant haploid phase. The technique for producing polyploids involved prototrophic selection and interruption of normal sequence of events leading from nuclear fusion to meiosis [23]. Maleszka et al. [24] observed that increasing the chromosome number improved ethanol production by P. tannophilus from several carbon sources, notably xylose. In addition the level of by-product formation from D-xylose such as xylitol decreased. Increase in ploidy also increased the growth rate of the organism on D-galactose but not appreciably on D-xylose. Some of these effects observed on increasing chromosome number have been attributed to complex physiological phenomena. For example, the rate of ethanol production from xylose increases without significant changes occurring in the activities of xylose catabolizing enzymes such as xylose reductase, xylitol dehydrogenase and alcohol dehydrogenase [24]. It appears that improvement in the yield of ethanol production from D-xylose in P. tannophilus results from an increase in the number of one or more particular chromosome rather than multiplication of entire genome. However, similar studies in bacteria and fungi for improving pentose utilization have not been reported.

3.2

Protoplast fusion

Protoplast fusion has been used as a general technique for genetic recombination of a variety of industrial microorganisms, particularly for microorganisms which have not been subjected to extensive genetic analysis. Protoplast fusion in bacteria and fungi have been reviewed earlier [25-27]. In the presence of a fusogenic agent such as polyethylene glycol (PEG) protoplasts are induced to fuse and form transient hybrids or diploids. It is presumed that during this hybrid state the genome or chromosomes reassort and lead to genetic recombination. Protoplast fusion provides characteristic advantages such as promotion of high frequencies of genetic

358 recombination between organisms for which poor or no genetic exchange has been demonstrated or which are genetically uncharacterized. Intraspecific, interspecific or intergeneric fusions involving two or more complete parental genomes have been demonstrated. Thus desirable genes from divergent strains can be introduced. Mating type do not inhibit hybrid formation in this method. Protoplast fusion of a number of industrial microorganisms has been described by Matsushima and Baltz [28]. Spheroplast fusion has been studied by a number of workers in order to improve ethanol production of yeast [29-34] and has been reviewed earlier [35]. Studies also indicate that spheroplast fusion may be employed for construction of yeast strains with new capabilities for utilizing new substrates [21]. However, Stewart et al. [36] have opined that spheroplast fusion is not specific enough to modify genetic characteristics of industrial yeast strains in a predictable fashion. The yeast Candida blankii is known to utilize xylose as substrate. Strain ESP94 of this yeast has been isolated for the production of SCP on bagasse hydrolysate. However, the major drawback in commercial utilization of this strain for SCP production has been small cell size that complicates cell harvesting during a continuous fermentation process. Hence Gericke and van Zyle [37] employed intraspecific protoplast fusion of auxotrophic mutants of strain ESP-94 in order to increase the cell volume of strain ESP-94. They obtained six genetically stable fusants with larger cell volume and higher DNA contents. One of the fusants, fusant F17, exhibited thrice as much cell volume as that of ESP-94 and showed similar growth rates on xylose as the carbon source. Strains of Zygosaccharomyces fermentati, a thermotolerant yeast, with the ability to utilize cellobiose have been fused with ethanologenic S. cerevisiae. The intergeneric hybrid produced with the traits of both parents was very stable and had the ability to grow on either cellobiose or lactic acid as the carbon source [38]. Johanssen et al. [39] have employed protoplast fusion for construction of polyploids of C. shehatae. They observed that increase in ploidy leads to small increases in rate of ethanol production from xylose. However, attempts to construct polyploids of Pichia

guilliermondii by protoplast fusion method for higher biomass or ethanol production have not be successful [40]. Interspecific

protoplast

fusion

of

pentose

fermenting

clostridia,

C.

acetobutylicum P262 has been shown to yield a stable fusant [41]. This has suggested that C. acetobutylicum can undergo homologous recombination. Protoplast fusion has great promise for genetic improvement of clostridial strains. However, detailed studies are required for improving these strains for pentose bioconversion.

359 4

GENE CLONING, EXPRESSION AND CHARACTERIZATION

4.1

Yeasts

Pentose fermenting yeasts particularly, Pachysolen tannophilus, Candida

shehatae and Pichia stipitis are known to ferment xylose to ethanol but they lack in efficiency. In addition they are relatively ineffective in fermenting glucose [42]. On the other hand, brewers yeast, Saccharomyces cerevisiae, which is one of the most ethanol tolerant yeasts, is unable to produce ethanol from xylose. However, a ketoisomer of xylose, xylulose can be utilized by many Saccharomyces sp. for ethanol production [43-45]. One of the approaches followed to construct Saccharomyces sp. suitable for pentose bioconversion to ethanol involves cloning of both xylose reductase and xylitol dehydrogenase genes from naturally xylose fermenting yeasts such as Pichia. In another approach the xylose isomerase gene of bacteria such as E. coil has been cloned and expressed in yeast. Using the first approach Takuma and coworkers [46] cloned the xylose reductase gene from Pichia stipitis and expressed it in Saccharomyces cerevisiae but recombinants failed to metabolize xylose. Hallborn et al. [47] obtained efficient conversion of xylose to xylitol by transforming Saccharomyces cerevisiae with gene encoding xylose reductase of Pichia stipitis CBS 6054. The recombinants showed 95% conversion of xylose to xylitol. Kotter et al. [48] and Tantirungkij et al. [49] have also shown that the cloned intact xylose reductase and xylitol dehydrogenase gene from Pichia stipitis can be expressed in S. cerevisiae. The recombinant strain also metabolized xylose albeit with much lower efficiency. It has been suggested that the expression of these genes in S. cerevisiae by their natural genetic elements (promoter and ribosomal binding sites) may not be effective [50]. In order to improve the efficiency of yeast transformants, overexpression of these genes have been sought. Chen and Ho [50] demonstrated that expression of Pichia xylose reductase can be improved nearly 20-folds by fusion of xylose reductase structural gene to the 5'noncoding sequence of yeast alcohol dehydrogenase (adcl) containing the intact genetic elements for gene expression and 3'-noncoding sequence of yeast xylulokinase gene. Alternatively, mutants of xylose assimilating recombinant S. cerevisiae carrying xylose reductase and xylitoi dehydrogenase genes on plasmid pEXGD8 have been selected after ethyl methanesulfonate treatment for their rapid

360

growth on xylose containing medium [51]. The fastest growing mutant strain IM2 showed a lower activity of xylose reductase but a higher ratio of xylitol dehydrogenase to xylose reductase and higher xylulokinase activity than the parent strain. In batch fermentation under oxygen limitation mutants showed higher yield (1.6 times) and improved production rate (2.7 times) than the parent. In fed-batch culture with slow feeding of xylose and appropriate oxygen supply ethanol yield has been reported to be further increased while production rate decreased [51]. However, the fact that most yeasts do not efficiently utilize xylose due to cofactor (NADPH/NADP) regulation [52] the second approach involving cloning of bacterial isomerase gene has been most popular for improving S. cerevisiae for pentose fermentation. Most bacteria, such as E. coli, Bacillus subtilis, convert xylose directly to xylulose by single enzyme xylose isomerase, which requires no cofactor for its action [53,54]. Thus efforts have been made to circumvent xylose reductase-xylitol dehydrogenase pathway of yeasts by cloning and expression of bacterial xylose isomerase gene [55-62]. Lawlis et al. [56] cloned a 4.2 kilobase pair fragment of E. coil chromosome which contained the gene xylose isomerase and xylulokinase into plasmid pBR 322 by complementation of a mutant deficient in xylose utilization. Ho and Chang [58] purified and characterized xylose isomerase gene from E. coli hybrid plasmids bearing different sizes of insert. E. coil xyl A gene has been expressed both in Saccharomyces cerevisiae [59] and Schizosaccharomyces pombe [60]. The transformed Schizosaccharomyces pombe exhibited growth on xylose as sole source of carbon [60]. In subsequent studies it has been observed that the limiting factor for D-xylose utilization by the transformed yeast is low activity of xylose isomerization [61 ]. The low activity of xylose isomerase in transfomed yeast has been attributed to either low expression of xyl A gene or proteolytic activity. In vitro studies however demonstrated proteolytic activity in transformed yeast but the problem of low expression should not be ignored [61]. In another study Stevis and Ho [62] observed that overproduction of xyl A gene can not be accomplished by cloning the intact gene on a high copy number plasmid alone. This was probably due to the fact that expression of gene through its natural promoter is tightly regulated in E. coil Thus xyl A gene has been fused with other strong promoters such as tac and lac to construct a number of fused genes. E. coil transformants containing the fused gene, cloned on high copy number plasmids showed 20-fold overproduction of xylose isomerase. Attempts have also been made with xylose isomerase genes from Bacillus subtilis and Actinoplanes missourienis [63]. In addition to xylose isomerase genes, cloning and expression of xylose uptake gene from E. coil[64], xylulokinase gene from

361 Pachysolen tannophilus [65] and Saccharomyces cerevisiae [58,66] have also been attempted.

4.2

Bacteria

Efforts have also been made to improve bacterial strains known to utilize both pentose and hexose sugars. For example, E. coli and many enteric bacteria such as Klebsieila, Erwinia etc. are known to ferment both types of sugars and have been characterized as having mixed acid type of fermentation. They dissimilate xylose to yield pyruvate via pentose phosphate and Embden-Meyerhof pathways. Under anaerobic conditions, pyruvate is degraded by pyruvate-formate lyase to yield acetate, ethanol and formate as main fermentation products in the ratio 1:1:2, respectively. These enteric bacteria have a remarkable trait that they can metabolize all the sugar constituents in lignocellulosic material but they do not convert these sugars to any single product of commercial value. On the other hand Z. mobilis, an anaerobic Gramnegative bacterium is known to be a potent producer of ethanol from glucose. The organism employs Entner-Doudorff pathway in conjugation with the enzyme pyruvate decarboxylase and alcohol dehydrogenase [67]. Z. mobilis possesses many of the traits sought in an ideal biocatalyst for fuel ethanol production. It shows higher ethanol productivity (3 to 5-fold) than yeast [68,69] with an ethanol yield from glucose upto 97% of the theoretical maximum yield [68]. Other favourable traits are ability to ferment at low pH, high sugar and ethanol tolerance, and tolerance to inhibitors present in lignocellulosic hydrolysate. However, the organism can only utilize glucose, sucrose and fructose as carbon and energy sources [70]. Thus, Z mobilis has the potential to become a superior organism than traditional yeasts for the fuel ethanol production from iignocellulosics, if its narrow substrate range could be overcome [68,70]. Alternatively these genetic traits of Z. mobilis can be transferred to E. coil in order to reduce the spectrum of fermentation product of enteric bacteria to mainly ethanol, two basic approaches have been followed [71]: 1 2

Insertion of a Z. mobilis gene encoding pyruvate decarboxylase alone in enteric bacteria with reliance on the host organism for alcohol dehydrogenase. Insertion of an artificial operon containing the Z. mobilis gene for both pyruvate decarboxylase and alcohol dehydrogenase in enteric bacteria. In early studies expression of Z. mobilis pdc gene in E. coli has been shown

362 to cause an increase in ethanol production [72]. Apparently due to low endogenous levels of native alcohol dehydrogenase only low levels of ethanol were monitored in the recombinants. However, Ingram and co-workers [73] observed that E. coil mutants, hyper-expressive for native alcohol dehydrogenase produced ten-fold higher levels of ethanol from glucose upon the insertion of Z. mobilis pdc gene. Tolan and Finn [74] transferred the pdc gene of Z mobilis into K. planticola wild type cells which ferments hexoses and pentoses to acetate, ethanol, formate, lactate and 2,3-butanediol. They observed that yield of ethanol is increased from O.7M to 1.3 M per mole of xylose and levels of other catabolic end products such as acetate and formate decreased at low pH. In another study using a similar approach Z. mobilis pdc gene was inserted in Erwinia chrysanthemi [75]. The recombinant E. chrysanthemi produced 7.4 g ethanol/I from xylose. This shift in metabolism has been attributed not only to the level of pdc gene expression but also to the strong affinity of pyruvate decarboxylase for pyruvate rather than for pyruvate formate lyase or lactate dehydrogenase. However, such recombinants were not satisfactory for pentose bioconversion as they required slow feeding of nutrients for higher yields and fermented mixed substrates very slowly. In addition, at higher growth rates and sugar uptake, organic acid and butanediol accumulated instead of ethanol. Therefore, efforts have been made to isolate mutants of Klebsiella planticola deficient in pyruvate formate lyase. Feldmann et al. [76] isolated such mutants that produced more than 70% lactate with residual acetate, 2,3 butanediol and traces of ethanol, formate and C02. Further they constructed a recombinant strain from these mutants by introducing plasmids carrying the pdc gene from Z. mobilis. The recombinant strain was an efficient ethanol producer and produced 387mM ethanol from 275mM xylose in 80h (about 83% of theoretical yield). The recombinant strain utilized more than double the amount of xylose compared to wild type. However, they showed poor ethanol tolerance and this trait has limited the practical usage of this recombinant. Using another approach, insertion of an artificial operon, pet operon (for producton of ethanol), containing Z. mobilis gene for both pyruvate decarboxylase (pdc) and alcohol dehydrogenase (adhB) has been investigated [73,77,78]. The pet operon has been constructed by deleting the native promoter region from both genes along with 3'-terminal sequence of pdc gene which is presumed to act as a transcriptional terminator [71]. Both genes show high levels of expression under the control of E. coil lac promoter. The recombinant strain efficiently produced high concentration of ethanol (56 g/I) from xylose and showed volumetric productivities up to 1.4 g ethanol/I/h. In addition it exhibited an ability to efficiently ferment all other sugar constituents of lignocellulosic material [78]. Using hemicellulose hydrolysate

363 from Pinus sp. an ethanol yield of 91% of the maximum theoretical yield has been reported in 48 h [79]. Recombinants of E. coli carrying the pet operon on plasmid PLOI279 converted hemicellulosic hydrolysate to ethanol at an efficiency of 94% of theoretical maximum [80]. The efficiency was 15% better than the highest efficiency reported for pentose utilizing yeast in similar system. Optimization of fermentation and nutritional supplementation showed that recombinants can convert 100% xylose from Aspen prehydrolysate fortified with tryptone and yeast extract with volumetric productivity of 0.29-0.76 g/I/h. Recombinants also produced ethanol from a nutrient supplemented newsprint prehydrolysate medium but with 20% lesser yield and productivities than softwood hemicellulose hydrolysate media [81]. The final ethanol concentration reached upto 14.6 g/I with a conversion efficiency of 74.5% of theoretical maximum. In addition recombinants showed the ability to ferment mannose as well with 90% efficiency [82]. Ohta et al. [83] have studied expression of pdc and adh genes from Z. mobilis into another enteric bacteria Klebsiella oxytoca. The recombinant strain containing only the pdc gene exhibited more than twice the parental level of ethanol produced. Efficient ethanol production by the recombinant with both pdc and adh genes has been observed. The maximum volumetric productivity obtained was 2.1 g/I/h for both glucose and xylose. Interestingly, plasmids carrying the two genes were stable and maintained in K. oxytoca in the absence of antibiotic selection. Fermentation of various sugars such as arabinose, xylose and glucose by recombinant K. oxytoca has been evaluated by Bothast et al. [84]. The organism produced 0.34-0.43g ethanol/g sugar at pH 6.0 and 30~ on 8% sugar substrate. Preferential utilization of glucose followed by arabinose and xylose has also been monitored in a pH controlled batch fermentation. However, under similar conditions the ethanol production followed xylose > glucose > arabinose. Alternatively, metabolic engineering of xylose fermentation in Z. mobilis is an essential step towards its development as a biocatalyst for fuel ethanol production from lignocellulosic feedstocks. Worldwide attempts to transform Z. mobilis into an efficient ethanol producer from abundant renewable carbon sources are being carried out [85,86]. Although Z. mobilis fails to grow on xylose [70], xylose transport appears to be mediated by an indeginous glucose facilitated transport system [87]. Thus xylose isomerase and xylulokinase were thought to be required to convert pentoses into intermediates of pentose phosphate pathway [67]. Approaches followed to engineer the metabolic pathway of Z mobilis involves: 1

Insertion of xylose isomerase and xylulokinase gene from enteric bacteria to

364 Z. mobilis. 2 Insertion of pentose assimilating operon and pentose phosphate pathway operon to Z. mobilis. Following the first approach, Liu et al. [88] transferred xylose isomerase (xylA) and xylulokinase (xylB) genes from Xanthomonas campestris XAI-1 to derivatives of Z. mobilis strain ZM6. Transconjugant Z. mobilis strains expressed both xylose isomerase (0.645 U/mg) and xylulokinase (0.145 U/mg) at rates similar to those found in E. coilwild type growing on xylose. But they failed to grow on xylose as sole source of carbon. Similarly Feldmann et al. [89] studied expression of these two genes from Klebsiella pneumoniae 1033 in Z. mobilis strains CP4 and ZM6 on shuttle plasmids carrying a strong pdc gene promotor from Z. mobilis [90]. Recombinant strain expressed both xylose isomerase (0.15 U/mg) and xylulokinase (1.6 U/mg), but again failed to grow on xylose and could not produce ethanol from pentose sugar. The presence of low 6-phosphogluconate dehydrogenase and transketolase activities has been demonstrated but transaldolase activity was undetected suggesting a lack of complete pentose phosphate pathway in Z. mobilis [89]. Recently Zhang and co-workers [91] engineered the metabolic pathway of Z. mobilis using the second approach i.e. inserted both pentose assimilating operon and pentose phosphate pathway operon. They cloned E. coil xylA and xylB genes under the control of a strong constitutive Z. mobilis glyceraldehyde-3-phosphate dehydrogenase promoter [92] by polymerase chain reaction-mediated overlap extension [93]. The resulting xylose assimilating operon was transferred into Z. mobilis CP4. Although both genes were functionally expressed, transformants failed to grow on xylose probably due to lack of transaldolase and sufficient transketolase activity. Therefore, Zhang et al. [91] synthesized an open reading frame postulated to encode a transaldolase homolog at 0 to 2.4 min on the E. coil chromosome and subcloned it under the control of Z mobilis enolase promoter by PCR-mediated overlap

expression. They also synthesized the transketolase gene (tktA) from E. coil W3110 genomic DNA and subcloned it downstream of transaldolase homolog translation termination codon to form an operon encoding the non-oxidative portion of pentose phosphate pathway. These two operons consisting of two xylose assimilating and two pentose phosphate pathway genes were simultaneously transferred in Z. mobilis CP4 on a chimeric shuttle vector constructed from a 2.7 kb Z. mobilis native plasmid and pACYC 184. The recombinant Z. mobilis CP4 (pZB5) grown on a glucose based medium demonstrated the presence of xylose isomerase (0.11 U/mg), xylulokinase (1.5 U/mg), transaldolase (0.88 U/mg) and transketolase (0.16 U/mg) activities. This recombinant strain demonstrated the capability to grow on xylose as sole source of

365 carbon and efficiently produced ethanol as the principal fermentation product. The recombinant showed cell growth at a rate 0.057 h1 and an ethanol yield of 0.44 g/g xylose, approximately 86% of theoretical yield. The recombinant also achieved 94% of theoretical yield from glucose within 16 h. A mixture of both glucose and xylose was fermented to ethanol at 95% of theoretical yield within 30 h. This high yielding capability of recombinant on a mixture of hexose and pentose sugar offers great potential for advanced process design that needs co-fermentation of mixed sugar feedstocks. It is apparent that approaches employing both classical genetics and genetic engineering has led to a certain level of success for the improvement of strains for lignocellulosic bioconversion to various fermentation products. However, the major focus in so far has been the improvments in bioconversion of lignocellulosics to ethanol. Screening and mutagenesis has met with limited success in isolating strains with decreased by-product formation, higher ethanol production and product tolerance. Attempts have been made to improve pentose fermentation by yeasts by cloning both xylose reductase and xylitol dehydrogenase genes from naturally xylose fermenting yeasts. However, cofactor requirement by these enzymes has limited bioconversion by transformants. Another approach has been to circumvent xylose reductase and xylitol dehydrogenase pathway of yeast by cloning and expression of bacterial xylose isomerase gene. Alternatively, bacterial pentose fermenting ability has also been improved by cloning the pdc gene either alone or with the pet operon from Z. mobilis. Recently, Z. mobilis pentose pathway has been engineered by cloning two operons consisting of xylose assimilating and pentose phosphate pathway gene simultaneously. Genetic improvement of ethanologenic organisms employed for the bioconversion of lignocellulosics have already provided efficient strains with high yielding capabilities. Nevertheless, genetic improvement of strains for bioconversion of lignocellulosics to other solvents such as acetone, butanol, butanediol needs further attention. With our increasing knowledge of genetics and molecular biology of clostridial strains employed for acetone-butanol fermentation, such improvements are likely to be achieved. In addition increasing knowledge of fungal genetics holds promise to improve strains for direct bioconversion of lignocellulosics to various products.

366 5

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14 Process evaluation and bioengineering

1

INTRODUCTION

Lignocellulosic residues are sources of cheap raw materials for the production of a variety of value-added chemicals and protein-rich food and feed materials. However, the processing of biomass into chemicals via fermentations is at primitive stage of development as compared to the chemical processing of petroleum and natural gas [1]. Plans for the future research in this area and evaluation of results already obtained, are directly concerned not only with the efficiency of process variables but also with the energy and environmental aspects. The current industrial activity of lignocellulosic biomass fermentation is limited mainly because of two reasons. First, the cost of raw materials processing is a critical factor. Lower substrate pretreatment cost and maximum carbon yields will improve the economic viability of the fermentation process. Second, present fermentation productivities are low, when compared to chemical processes for fuel and chemical production. The economic importance of utilization of natural biomass mostly depends on the bioconversion of both hexose and pentose sugars present in the hydrolysate. Since fermentation of pentoses is slower than that of hexoses, improvement in pentose utilizing organisms, in terms of yield, productivity, and end product tolerance are likely to be concerned in the overall process. Direct fermentation of lignocellulosic materials by certain microbial systems that produce solvents and polysaccharidases is an attractive and exciting approach. The advantages of direct fermentation include the use of a single bioreactor that simplifies the process, reduces the capital cost and increases the overall rate of conversion. In addition to the increased efficiency in the use of raw materials, value of by-products is increased by producing commercial polysaccharidases and microbial biomass that have higher value-added as either a feed because of its protein, lipid or vitamin content, or as biocatalyst for further biochemical synthesis of and transformation. The technology for biological prodution of chemicals from pentoses present in lignocellulosic biomass involve the successful interlinking of four system components: 371

372 Pretreatment and preparation of substrates. Design of fermentation process. Product recovery, including by-products. Processing of by-products and wastes. The capabilities and limitations of microbial catalysts are key to process design, as this markedly affects the types of pretreatment processing [2]. For example, wood hydrolysis to sugar is required when noncellulolytic microorganisms are used for fermentation. End product inhibition limits the productivity of all solvent-producing fermentations. The end product recovery is governed by the process temperature and final concentration of the end product achieved. The need for waste treatment and byproduct utilization are key components of overall process costs. Thus successful combination of all four system components would comprise an economic process. An ideal solvent production process must have low operating costs (continuous process, low energy input, use of low cost substrates, near or complete utilization of substrate, use or elimination of by-products), and low capital costs (small reactors with mechanical simplicty and high productivity).

2

PRETREATMENT OF SUBSTRATE

Insoluble substrates like lignocellulosic materials must usually undergo some pretreatment prior to fermentation. In general, the pretreatment of a fermentation substrate must allow the maximum utility of the substrate, and any process wastes will have deleterious effect on overall process [3-7]. In wood pulping processes, cellulose is the valuable product, while hemicellulose and lignin end up as waste by-product. Alternative pulping strategies are being developed so that wood can be pulped by new chemical or biological means that will conserve the hemicellulose fraction as a fermentation feedstock and improve the polymeric properties of processed lignin for applications in adhesive, plastics and other polymers [7]. Wood can be steam-exploded and both the hemicellulose and cellulose are fermentable [8-11]. An important limitation of many pretreatment technologies is their ineffectiveness against softwood substrates. However, progress is being made in this area [12].

373 Current techniques of sugar extraction from biomass favour harsh conditions with strong acids in which much of the hemicellulose content is lost, largely through degradation to furfural. Furfural could be used as a chemical feedstock, and has received attention as a by-product to improve the economics of acid hydrolysis based ethanol production [13-15]. However, considering several cost and utility factors, it appears that xylose conversion into ethanol is more attractive than conversion to furfural [16]. On the other hand, mild acid hydrolysis has several advantages including the prevention of decomposition of xylose to furfural (inhibitory to microbial growth) and it also limits the production of by-products. Kinetics of acid hydrolysis of pentosans has been investigated by several researchers [17,18]. Three phases exist in the process. An initial random attack on the hemicellulose chains by the acid results in the formation of oligomers of various degrees of polymerization. These are then split into monomerss with subsequent degradation to furfural. Simple steam treatment, without aid of acid catalysts (autohydrolysis) has been found to be effective in liberating sugars from hemicellulosic materials. Autohydrolysis occurs at least in part via acetic acid formed by the cleavage of acetyi side chains present in pentosans. A two step process developed at Indian Institute of Technology, Delhi involves autohydrolysis of rice straw followed by solvent treatment for the fractionation of rice straw [7]. This process allows high yields of the each major fraction (cellulose, hemicellulose, and lignin) of rice straw. Enzymatic hydrolysis is another alternative for pentose recovery. The main advantages of this approach are greater specificity and reduced formation of degradation by-products. Important drawbacks include high cost and slow rates of hydrolysis. An organism capable of simultaneously producing cellulase and xylanase enzymes would be desirable for the hydrolysis of holocellulose. Relatively few organisms have the ability to hydrolyze xylan and ferment the resultant xylose. To overcome this problem, two or more organisms may be used in a coculture. Several researchers have pointed out the difficulties involved with analysing lignocellulosic hydrolysate due to the complex matrix of hydrolysate, lack of suitable analytical instruments and lack of trained personel for interpretation of chromatograms [19]. A separation method for the analysis of lignocellulosic hydrolysate using spent sufite liquor as the model substrate has been developed [20]. The separation of glucose, xylose and arabinose is performed using a precolumn and two HPX-87H (BioRad) columns coupled in series to enhance the resolution. When a hydrolysate also contains galactose and mannose, the sugars must be separated on an HPX-87P column (BioRad).

374

3

FERMENTATION DESIGN

The design of the biorector system depends on the physiology of the organisms employed. Numerous systems are available which are either based on batch, fed-batch or continuous culture fermentation. All the products discussed in previous chapters can be produced in batch cultures and the technology involved is well known. Batch processes still dominate the fermentation industries with relatively low productivity and high capital cost. New process are now evolving that use continuous open systems with large increase in productivity and with reduced running and capital costs. There are two basic approaches to the conversion of xylose to ethanol by yeasts. One is to use a two-stage system based on xylose isomerase and yeast; the other is to use a single-stage system with a selected yeast species such as Pachysolen tannophilus,

Candida shehatae or Pichia stipitis. The two-stage system has the advantages of a higher overall rate of conversion and probably a higher ethanol yield as well [21-23]. However, an additional mitigating factor in the two-stage system is the cost of D-xylose isomerase enzyme. Several systems based on chemostat principle have been developed that improve productivity by maintaining close control of the environmental conditions. The chemostat or continuous stirred tank reactor (CSTR) is a significant improvement over batch culture for the production of primary metabolites using both yeast and bacteria [24,25]. In these systems, the organisms are normally substrate limited, thus substrate inhibition can be avoided and simultaneous use of mixed substrate is possible. One limitation of continuous culture is retention of the biocatalyst. If high inhibitory end product concentration exist, then productivity is reduced in line with the cell growth rate [26]. The temporal and spacial homogeneity of chemostats can also be a problem for multiphase fermentations common to acetone-butanol. This is because a rapidly growing acidogenic phase is required before a slow growing solventogenic phase intiates. Growth inhibition can be relieved and much higher cell densities can be obtained by operating ethanol fermentation under vacuum [27]. The continuous process conducted under vacuum enables rapid and complete conversion of concentrated sugar solutions. Two-times increase in ethanol productivity has been found in continuous vacuum fermentation performed with cell recycling [28]. Oxygen has an important influence on viability of cells and their activity and must be present in small concentration. The optimum oxygen feed rate for ethanol production

375 is about 0.1 vvm. Higher oxygen concentrations decreases ethanol productivity. Cell recycling and immobilization techniques have been applied successfully to yeast and bacterial fermentations [29,30]. Immobilization of solvent fermentations may stabilize the process, increase product tolerance, improve substrate utilization, and increase productivity. Various methods for immobilization of solvent producing microorganisms have been studied. Cell entrapment in gel-forming polysaccharides such as alginate and carrageenan are the most popular methods. Immobilized yeast cell entrapped in alginate have been used at pilot scale for ethanol production where efficient conversion was achieved for over 4000 hours of operation [31]. Extractive fermentations offer a method by which the toxic fermentation products may be removed from the system in situ. it allows continuous process with reduced product inhibitory effects. It also permits the use of more concentrated sugar solutions, thereby reducing the amount of water needed in the process [32]. The use of reduced pressure within the fermentation system achieves volatilization of solvents to form highly enriched vapours which are then concentrated by distillation [33,34]. These include Vacuferm and Flashferm processes where a flashing vessel is incorporated. This technique is of particular interest in the case of thermoanaerobes where the lower ethanol concentration can be removed from the media because of the high fermentation process temperature. In a conversion scheme developed by Sitton et al. [35], it has been shown that 188 kg of ethanol can be produced per ton of corn stalks. At a predicted consumption rate of 220 tons of corn stalks per day, the process would generate 41,360 kg of ethanol daily. Scheme of the process design is given in Figure 1. Separate microbial systems are used for ethanol production from glucose- and xylose-rich streams. A fixed film reactor of Saccharomyces cerevisiae is suggested for hexose conversion, while the pentose mixture is fermented by immobilized Fusarium oxysporum.

376

CORN STALKS]

PREHYDROLYSlS (4.4% 1-12SO 4) SOLIDS IMPREGNATION (85%H2S04)

STREAM I----" XYLOSE-RICH

-I

DILUTE ACID HYDROLYSIS

GLUCOSE-RICH STREAM

(8%HzS04)

IELECTRODIALYSISI ~-. - I XYLOSE I

GLUCOSE l-..

~ S. cerevisiae

~ F. oxyspor u m

ETHANOL (2.1%)

I

ETHANOL

(0.5%)

"- IDISTILLATION l ETHANOl_ (95%)

Figure 1. Process for the conversion of corn stalks to ethanol

I

377

An integrated process for refining and bioconversion of rice straw to ethanol and coproducts (single cell protein and lignin) has been developed at Indian Institute of Technology, Delhi. In this process rice straw is fractionated into cellulose, hemicellulose and lignin by a two-stage treatment process (Figure 2). Cellulose is used as substrate for the production of cellulolytic enzymes as well as ethanol in a simultaneous saccharification and fermentation system using Candida acidothermophilum. Pentose sugar stream generated after autohydrolysis of rice straw is used for single cell protein production using Candida utilis.

IRICE STRAW ! CELLULOSE +

! AUTOHYDROLYSIS] I

":,~

" 1 HEMICELLULOSE !

LIGNIN

ETHANOl_ TREATMENT ~ _ _

=Ic ,,u,os I

LIGNIN ETHANOL

SIMULTANEOUS SACCHARIFICATION FERMENTATION

l. cov .Y1

Yeast

ETHANOL

RESIDUE

Cellulase enzyme

Yeast

Ir

i-LIGNIN !

l ETHANOL I

'

'MIXED FODDER

SINGLE CELL PROTEIN

Figure 2. An integrated scheme developed at Indian Institute of Technology, Delhi for the refining and bioconversion of rice straw to ethanol and co-products

378 Detroy et al. [36,37] have designed a process for ethanol production from wheat straw. Three methods of straw pretreatment were proposed: (a) autohydrolysis followed by ethanol extraction; (b) autohydrolysis with subsequent ether extraction; and (c) alkali extraction of substrate. About 41% of wheat straw pentosans were extracted by autohydrolysis. Suitability of each pretreatment method was assessed by fermenting hemicellulose hydrolysate using Pachysolen tannophilus. Incomplete utilization of the sugars derived by autohydrolysis and ethanol extraction of wheat straw was noticed. On the other hand, fermentation of ether- and alkali-extracted substrate did result in complete utilization of xylose in 7 and 6 days, respectively. Deverall [38] evaluated hydrolysates of a hardwood (aspen) and a softwood (pine) for ethanol production. P. tannophilus was able to utilize 96% of xylose and almost all glucose and mannose in 35 h of fermentation with ethanol yield of 84% of theoretical. For pine hydrolysate, a two-stage process has been implemented where the substrate was first inoculated with Saccharomyces cerevisiae. A 24 h batch culture resulted in the accumulation of 16.5 g/I of ethanol. Centrifugation of broth produced a 'beer' containing 7.8 g/I of unconsumed carbohydrates (pentosans and galactose accounted for 69.2% and 26%, respectively). Inoculation of beer with P. tannophilus resulted in an improvement of ethanol yield by 9%, giving a final ethanol concentration of 18 g/l. Nolan et al. [39] described University of Pennsylvania/General Electric Company Process for complete utilization of biomass (Figure 3). Butanol-water at 160~ was used to reduce the lignin content. When butanol phase containing most of the lignin is cooled, excess lignin would separate leaving a saturated butanol-lignin slurry which is potentially useful as diesel fuel. The hemicellulose is hydrolyzed by the pretreatment process and is extracted into the aqueous stream. Simultaneous saccharification and fermentation was suggested using Thermomonospora enzyme to produce sugars and fermenting sugar to ethanol using an appropriate Clostridium sp. at 60~ Similarly butanol can also be produced by simultaneous saccharification and fermentation of cellulose. However, after fermentation, butanol is present at a level of only 1.5 to 2% and the distillation recovery is very enthalpy intensive. Fermentation of pentosans results in additional butanol production.

379

I

BIOMASS (WOOD CHIPS)

I SUGARSYRUPI I HYDROLYSIS !

~1 BUTANOL I

FELLULOSEI

ITREATMENT

I BUTANOL PHASE I

BUTANOL I RECYCLING LIGNIN RECOVERY

SOLVENT RECOVERY

SIMULTANEOUS SACCHARIFICATION FERMENTATION

ILIGNINI

/

! BUTANOLI

II

IAQUEOUS PHASE I

I~COSEI IFERMENTATIONI

1

IETHANOL I

!

SIMULTANEOUS SACCHARIF ICATION EXTRACTIVE 9FERMENTATION

Figure 3. University of Pennsylvania/General Electric process for total biomass utilization

A classical batch process is difficult to control and it cannot cope with the toxicity of the accumulating solvent products. It also does not utilize the biocatalyst to the full extent. The newly developed process should make the full use of most of the organisms by removing the accumulated solvents as they are produced to prevent their toxic effect

[40].

380 Multiphase systems are useful in fermentations that involve two phases such as that observed in acetone-butanol fermentation [41,42]. In a two-stage acetone-butanol process, the first vessel produces the catalyst in acidogenic phase, which is then fed into a second vessel where solventogenesis occurs under slower growth conditions. In tube or tower fermentation system, it is possible to retain biomass by allowing sedimentation to occur within the fermentor. Volumetric productivities can be increased several-fold. Application of continuous fermentation to commercial scale has not yet been developed due to degeneration of cellular acetone-butanol producing activity. Generally little or solvents are produced in chemostats with carbon/energy or nitrogen limitation. Nongrowing immobilized cells do not lose their solvent-producing capacity when the viability of cells are maintained by pulse-wise addition of nutrients to the reactor [43]. Higher butanediol product concentrations are obtained in the fed-batch mode due to inhibition by high substrate concentration. However, the yields are reduced compared to fermentations where substrate and product concentrations are kept low [44,45]. A successful compromise between product concentration, yield and high productivity can be achieved using a two-stage process [46]. Processes for the pilot- and commercial scale production of butanedioi from molasses and cereal grains (wheat and barley) have been developed [47]. However, such processes are not available for the utilization of pentose sugars. Several agricultural residues have been considered as possible feedstocks for butanediol production. Environmental factors have been shown to significantly influence diol production by bacteria. The operational conditions of the fermentor are similarly important in the establishment of an optimal process design. While a concentrated product stream is desirable in any bioconversion scheme, it is essential in the butanediol process. A minimum of 80 g/I has been estimated to be required for economically feasible recovery [48,49]. The employment of cultures of Klebsiella

pneumoniae that had been acclimatized to high substrate concentrations, resulted in the accumulation of 106 g/I of butanediol from 225 g/I of glucose, and 81 g/I of butanol from 189 g/I of xylose [50]. Claussen et al. [51] designed a fermentation process for the production of organic acids from hydrolysate of lignocellulosic materials using Propionibacterium acidi-

propionicL This organism has been shown to produce propionic and acetic acids from a mixture of glucose and xylose without inhibition. As a design basis, a plant utilizing 200 metric tons of orchard grass per day was chosen. The orchard grass is first hydrolyzed to produce glucose and xylose. The acid production unit consists of a 4.65 million-gallon steel reactor and a 40,000 gallon culture tank. Propionic and acetic acid that are

381

produced in the reactor are then separated and purified for direct sale. In an integrated process for food/feed and fuel (ethanol) production from biomass (Figure 4), it has been suggested the hemicellulose fraction can also be utilized for the production of active mycelial inoculum to be used for the production of cellulase [52].

i FOREST BIOMASS I

ICELLULOSEI

ILI~NIN!

,/%

IHEMICELLULOSE I

I ADHESIVES I CHEMICALS COMBUSTION

CELLULOSE IHYD LYSIS! |ENZYMF | / AEROBIC O ~91--I PRODUCTION/ ~1"-I FERMENTATION I MICROBIAL 1 T. reesei / l INOCULA

AEROBIC FERMENTATION

C. cellulolyticum

|RESIDUAL V

ANAEROBIC

!~

FERMENTATION

IF~ THANOLI

SINGLE CELL I

PROTEIN

Figure 4. An integrated process for food/feed and fuel (ethanol) production from biomass

A large portion of glucose (produced from hydrolysis of cellulose) can be saved in above process which could be used for the production of additional ethanol. The inocula of other fungi, Chaetomium cellulolyticum and Pleurotus sajor-caju can also be produced on this fraction. The surplus hemicellulose can be converted into single cell protein by

382 these fungi. The single cell protein product of these fungi contains 40-47% crude protein on a dry weight basis which can be used as an animal feed.

4

DOWNSTREAM PROCESSING

Product recovery is considered to be the most critical stage in the overall bioprocesses. Solvents are volatile compounds and distillation offers the most obvious way of recovery. However, high solvent concentrations are required for distillation to become economical. A significant part of energy expenditure of the whole process is spent on product recovery [53]. The use of bioprocess alcohol as a liquid fuel has been questioned on the basis that the energy required for distillation is equal to the total combustion energy of the alcohol product (21 .lXl 0e Joules/I). New methods of distillation are being developed and alternatives to distillation has been investigated which offer more efficient separation processes. Low-boiling organic solvents like ethanol are relatively easy to separate by distillation due to large boiling point and volatility difference. However, the separation of dilute alcohol-water mixture into pure alcohol and pure water is difficult as alcohol and water form an azeotrope at 95.7 wt% (89 mol%) alcohol concentration [33]. Anhydrous ethanol is usually produced by azeotropic distillation with an added entrainer. The benzene azeotropic distillation is best known [54]. Production of anhydrous alcohol from azeotrope using benzene as an entrainer requires 1 kg steam/I of product. Other entrainers used are trichloroethylene, n-pentane and ether. A variety of techniques have been proposed for dehydration and molecular sieve dehydration appear to be particularly promising [55,56]. Current practice for the energy-efficient distillation of ethanol is based on vapour recompression heat operating between the overhead vapour and the reboiler and also heat integration between columns operating at different pressures [57]. Extractive distillation offers the dual advantages of low reflux ratios and therefore low energy requirements and also elimination of the azeotrope [58,59]. Lynd and Grethlein [60] have designed an ethanol distillation process specifically for separating ethanol from dilute broth. This process uses intermediate heat pumps with optimal sidestream return (IHOSR)in conjunction with extractive distillation [61,62]. This process was later modified [12] in which condensation of the overhead vapour from the extractive

383 column provides heat for the evaporator and the stripping column, which is operated at a lower pressure. A summary of the energy demands for the various processes is given in Table 1 [63].

Table 1 Energy requirement for the separation of ethanol from aqueous solutions Process

Final ethanol

Energy requirement

concentration

(BTU/gal)

(%) Simple distillation

95

18,000

Azeotropic distillation

100

9,400

Simple distillation and azeotropic distillation

100

27,400

Multiple effect distillation

95

10,000

Vapour recompression distillation

95

1,930

Vapour recompression distillation and absorption

100

3,930

Supercritical extraction

91

2,850

Absorption water

100

2,000

Vacuum dehydration

100

37,300

Absorption dehydration

100

13,000

Vacuum distillation can also be used to advantage in producing a product of normal (atmospheric pressure) ethanol/water azeotropic composition. Under vacuum, the required reflux may be reduced. For a 13 wt% alcohol feed, producing 95 wt% alcohol product, the

384

energy requirement for a single column distillation is reduced from 7X106 Joules/I to 2.41X106 Joules/I of product [64]. Ladish and Dyck [65] suggested a process for ethanol dehydration by vapour phase water absoption which may be used beneficially at industrial scale. In this process distillation is conducted only to produce 85 wt% ethanol product. These vapours are then passed through a bed of absorbant material to produce pure alcohol. Laboratory scale tests indicate that dry corn starch will selectively absorb all the remaining water and essentially no alcohol. Alternatively, molecular sieves [66] or polystyrene resins [67-69] can be used to preferentially absorb ethanol from dilute feeds. Activated carbon columns have been used to remove organic solvents by passing stripped vapours through the column [70]. The use of supercritical carbon dioxide has been proposed to extract organic materials from aqueous solution. This method is based on the principle that organics may be extracted into liquid CO 2 at high pressure and ambient temperature. The process operates at 1000 psi at 25~ Analysis of pilot plant systems has shown that the energy requirements are significantly lower for supercritical extraction than that for distillation methods [71]. However, the major disadvantage is the high capital cost of equipment. Distillation costs for butanol and butanediol are very high and in the case of butanediol distillation appears to be impractical method. These compounds are less volatile than water, the boiling point of butanol is 118~ and that of butanediol is 184~ As a consequence alternative product recovery systems have been developed. Luyben's group [72-79] has investigated and compared five technologies for butanol recovery on the basis of design parameters and energy efficiency viz. stripping, adsorption, liquid-liquid extraction, pervaporation and membrane solvent extraction. The ease of operation in conjunction with a fermentation was an important criterion in the choice of these methods. In situ product recovery can improve the performance of a butanol fermentation. In general two different types of integrated processes can be distinguished: fermentation with product recovery integrated in the fermentor, and fermentation and product recovery in closed loop. A system with a loop may be more flexible than a system with recovery in the fermentor. However, the recirculation stream needed may be high and logistic problem may occur. On a large scale, it is possible that a process with recovery in the fermentor may be realized only with liquid-liquid extraction as the separation step, or adsorption in a chromatographic type of reactor [73]. Table 2 shows several technologies for in situ recovery and purification of butanol and acetone/butanol/ethanol mixtures.

385 Table 2 Technologies for in situ butanol recovery, and downstream acetone/butanol/ethanol recovery and purification In situ butanol

Acetone/Butanol/Ethanol

recovery

recovery and purification

Gas stripping

Conventional beer stripping

Liquid-liquid extraction

Modified distillation

Adsorption

Three-phase distillation

Pervaporation

Distillation/extraction

Membrane solvent extraction

Liquid-liquid extraction

Reverse osmosis

Supercritical extraction

The simplicity of evaporation method has been shown in Biostill process [80]. Pervaporation is already being used commercially for the dehydration of alcohol [81] and is likely to be developed further with respect to the optimization of membranes, modules and process design. In liquid-liquid extraction, problems with fouling and operation were encountered but the high capacity and high selectivity of solvents, and the possibility of performing the separation in the fermentor, e.g. stirred tank [82] or loop reactor [83], make the extraction an attractive method for in situ product recovery. Attempt has also been made to calculate the potential of the fine recovery technologies on a large scale with respect to equipment size, based on an identical butanol production rate [72]. In general, size of equipment may be reasonable with stripping, adsorption and liquid-liquid extraction. However, the design of an apparatus for liquid-liquid extraction is a tedious task. The energy requirements with pervaporation are lower, but still relatively high compared to conventional downstream techniques (Table 3). In adsorption and stripping, the processes with a low selectivity, the energy requirements are high. With liquid-liquid extraction, the energy requirements are also relatively high, but the main advantage is the high selectivity of alcohol/water separation.

386 Table 3 Energy demand in the integrated butanol recovery processes [72] Estimated heat of recovery (MJ/kg ABE)

Technique

Product a

Stripping

B, and AE mixture

21

Adsorption

B, AE mixture

33

Extraction and perstraction

ABE mixture

14

Pervaporation

B, and AE mixture

9

aA, acetone; B, butanol; E, ethanol

The physical and chemical properties of 2,3-butanediol (high boiling point, hygroscopicity etc.) make its recovery from fermentation broth an extremely difficult task by conventional methods. Solvent extraction has been found to be an effective means of butanediol recovery. Solvent extraction can achieve separation of butanediol from water soluble impurities such as sugars and protein. This enables the isolation of purer end product. Suitable solvents include diethyl ether and n-butanol [84,85]. About 75% diol present in a fermentation broth could be recoverd by a single extraction with diethyl ether [86]. Recoveries of co-products such as acetoin, ethanol and diacetyl were found to be 65, 25, and 75-90%, respectively. Othmer [87] has proposed process schemes for recovery of butanediol with butanol or butanediol diacetate. Butanediol can be easily adsorbed on active carbon [88]. Better results can be obtained if this operation precede solvent extraction. Diol recoveries of 100% have been obtained using acetone or dioxane as the extracting agent. However, the most practical method of diol recovery appears to be countercurrent steam stripping. Successful pilotscale steam opertaions were established in the mid-1940s by both the Northern Regional Research Laboratories, Peoria [89] and National Research Council of Canada [90]. Busche et al [91] developed a process for the recovery of acetic acid from dilute solutions. The process was developed using broth fermented

by Clostridium

thermoaceticum which converts glucose and xylose to acetic acid in near quantitative yields. In this process, after separating the product effluent from the cells, the solution is

387 pumped into a multistage extracter operating at 55~ (commensurate with the fermentor operating temperature), and 75 atm CO2 pressure to provide sufficient CO2 in solution to effect acidification of the feed. Acetate concentration in the feed was found to be the most important operating parameter. All utilities demand decrease in inverse proportion to increase in acetate concentration. Not only does the product concentration affect the cost of utilities, it also affects the investment required for all the manufacturing facilities.

5

ECONOMIC EVALUATION

In the economic evaluation of biological production of fuel and chemicals, by far the most important factor is the total yield, as the raw material cost is of prime importance [92,93]. Yields should be compared to the theoretical yield. The total volumetric productivity determines the capital investment costs, i.e the size of fermentors. The specific productivities, on the other hand, gives a measure of the efficiency of the fermenting organism. Not only yield and productivities are of importance, but also chemical and process steps that increase production costs play a significant role in process economics. A low media pH and a fast production time reduces the contamination risk and therefore contribute to the process economy [94,95]. The use of medium sterilization and sterile techniques add to extra cost to the process [93]. Lignocellulosic hydrolysates are detoxified using chemicals, molecular sieves, mixedbed resins, ion-exchange and adsorption resins, calcium hydroxide and extraction with organic solvents [96,97]. The calcium hydroxide treatment is not economically sound since both chemicals and equipment add costs. Detoxification of the hydrolysate costs US$ 0.052 in the chemical cost alone of calcium hydroxide which should be compared with the world market price of ethanol, US$ 0.34-0.44 [93]. Detoxification methods such as organic solvent extraction and ion-exchange resins have been considered to be even more expensive than calcium hydroxide treatment. Sitton et al. [35] carried out studies on process design and economic aspects of ethanol production from corn stalks, keeping in view the utilization of both hexoses and pentoses. Almost 50% of the capital investment was found to be in the acid hydrolysis and acid recovery processes. Raw material constitute 36% of the operating cost. The breakeven price for ethanol was calculated to be US$ 298.55 m3.

388 Lynd [12] studied the economic impact of the distinguished features of thermophiles (cellulase enzyme production, pentose utilization) for ethanol production from lignocellulosics as compared to a base case of ethanol production from wood using yeast and enzymatic hydrolysis [98]. In situ cellulase production in the case of thermophiles, eliminates or greatly reduces the cost of production and utilization of enzymes. Pentose utilization eliminates furfural production and processing as well as associated storage and waste treatment steps. Pentose utilization in combination with cellulase production increases the ethanol output by 47%. As compared to base case utility related operating and labor costs are lower in thermophilic case. The selling prices for ethanol including return on capital investment, worked out to be US$ 0.52/I for the base case and US$ 0.28/I for thermophilic case. Relative to the base case, the impact of cellulase production considered individually is to reduce the selling cost by 19 cts/I ethanol or 37%, and the impact of pentose utilization is to reduce the selling cost by 12 cts/I ethanol or 23%. Hinman et al. [99] evaluated economic impact of xylose bioconversion to ethanol for a wood-to-ethanol plant. A base case, without xylose fermentation, and alternative cases, with xylose fermentation were examined. For the base case, in which none of the xylose is converted to ethanol, the price of ethanol was US$1.65/ga1. The economic impact of xylose conversion depends on three key parameters: yield, ethanol concentration, and productivity. The highest yield obtainable is 100% theoretical or 0.51 g of ethanol per g of xylose. In addition, the highest ethanol concentration attainable in the xylose conversion unit is 30 g/I from a 60 g/I xylose feed. The capital cost per annual gallon for high productivity values was found to have a minimum value of US$ 0.25/annual gal. Accordingly, the maximum potential reduction in the price of ethanol was calculated using yield at 100% theoretical, ethanol concentration at 30 g/I, and capital cost per annual gallon at US$ 0.25. Using these values, the price of ethanol with xylose conversion is US$ 1.23/gal, which represents a US$ 0.42 or 25% reduction in the price of ethanol from base case. On the basis of performance data existing for three xylose fermenting yeasts, i.e.

Pachysolen tannophilus, Candida shehatae and Pichia stipitis, representative prices were calulated (Table 4). Pichia stipitis and Candida shehatae are better strains, since they are currently capable of achieving 70% of the maximum possible ethanol price reduction. At present fungi, bacteria and xylose isomerase-yeast combination do not appear to be capable of attaining or surpassing the performance of best yeasts [99].

389 Table 4 Ethanol prices calculated for different biocatalysts in xylose fermentations [99] Case

Ethanol cost (US$/gal)

Base case a

1.65

Maximum potentiaP

1.23

Pachysolen tannophilus c

1.48

Candida shehatae c

1.36

Pichia stipitis c

1.37

aNo xylose is converted to ethanol bXylose is converted to ethanol using following performance parameters: 100% yield, no ethanol inhibition, high productivity CXylose is converted to ethanol with published performance parameters for each type of biocatalysts

The three key parameters associated with xylose bioconversion that have impact on the economics of a wood to ethanol bioconversion plant are yield, ethanol concentration, and productivity. The study of Hinman et al. [99] has shown that ethanol yield and concentration are the most important parameters, whereas productivity has a relatively minor impact. Yield has importance because at a given wood feed rate, each increase in yield translate directly into an increase in revenue. If ethanol concentration is not high enough, it is necessary to add dilution water to the feed stream to the xylose conversion unit in order to achieve the maximum potential yield. Addition of water increases the size of distillation and concentration units, and the waste treatment unit. The load on the utility systems also increases. The productivity is of minor importance because it has only impact on the size of the xylose conversion unit, which is relatively small percentage of the total capital cost of the plant. It was found that a four-fold improvement in the capital cost per annual gallon from US$1.00 to US$ 0.25 reduces the price of ethanol US$ 0.05/gallon. On the other hand, four-fold improvement in yield from 20% to 80% at 30 g/I ethanol concentration reduces the price of ethanol US$ 0.20/gallon, and a three-fold

390 improvement in ethanol concentration from 10 g/I to 30 g/I at 100% yield reduces the price of ethanol US$ 0.35/gal. Acetone and butanol production was priced out of the market by rapidly increasing substrate prices and harsh competition based on cheap acetone and butanol from petrochemical sources. The inherent flexibility of a large petrochemical plant, unlike most fermentation plants, can vary the composition of product range to suit more closely market demand, making the synthetic route even more competitive [100]. The economics of an acetone-butanol fermentation plant utilizing wood chips as raw material has been illustrated by Gibbs [101]. Total value of products for one week's operation, producing 2 tons of acetone, 4 tons of ethanol, and 24 tons of butanol was 15,660 pound sterlings, whereas total cost was estimated to be 10,683 pound sterlings per week, utilizing 200 tons of saw dust or wood chips. No allowance was made for H2 and CO2 production and disposal or use of dried biomass. It seems likely that any revival of the fermentation process will depend on the availability of wastes and their efficient utilization. Claussen et al. [51] studied the aspects of process design and economics of a plant utilizing 200 metric tons of orchard grass per day for organic acid production. Hydrolysate containg glucose and xylose was used to produce propionic and acetic acids. About 90% of the operating cost was found to be due to the production of glucose and xylose by acid hydrolysis of orchard grass. The cost of dilute mixed acids projected in this study was US$ 0.108/kg to break even as compared to market price of acetic and propionic acids US$ 0.58 and 0.73 per kg, respectively. If sold in the open market, the acids would have to be separated and concentrated. Solvent extraction with trioctylphosphine oxide followed by distillation was found to be a feasible recovery method for acid concentrations from 0.1 to 5% [102]. The projected cost of recovery with this method was calculated to be US$ 0.139/kg.

6

FUTURE PROSPECTS

Notwithstanding short term economic factors and trends in the attention of scientists and policy makers, the prospect of decreasing and ultimately vanishing supplies of oil is a real problem today. It is becoming increasingly apparent that renewable energy sources must be developed to compliment existing supplies. Lignocellulosic biomass has been

391 shown to represent an enormous reservoir from which liquid fuels, chemical feedstocks and protein-rich feed materials can be generated in a realtively simple and cost effective manner. While the technology for hexose (glucose) bioconversion is well known, the high proportion of pentose sugars in biomass makes utilization of hemicellulose fraction an essential factor in the management of this resource. The integration of new ideas with existing technologies must soon result in the development of processes for the manufacture of chemicals from the total sugar content of biomass. As well as increased efficiency in the use of raw materials, by-product credits are increased by producing commercial depolymerases and microbial biomass that has higher value added as a feed because its vitamin content. The cost of raw materials and product yields are the major factors governing solvent production processes. The limitation of a process (i.e., its cost) are usually governed by two major factors: Engineering factors involving process design, substrate preparation, reactor design, and product recovery. Biocatalyst factors involving substrate range, substrate conversion, productivity, product yield and product tolerance. How to successfully integrate these engineering and biocatalyst factors into improving solvent fermentations, is a tremendous challenge to process development. Newly developed bioprocess technology lends itself to application of increasingly more powerful, readily available, and cheaper electronics control and microprocessor hardware. On-line process control and optimization is a reality with an unprecedented potential. The key elements for the control/optimization purposes are sensors for various key parameters [40]. Fast and reliable analytical sensors are available. By both understanding of the process kinetics and the interdependence of individual parameters, predictions can be made as to the process behaviour and/or response to control parameter changes. In bioprocesses, the key element is the microorganism. Its performance in accumulating maximum product concentration before shutting off its biosynthetic activity, which is critical to product yield, is unsatisfactory in most cases due to product inhibition. A microbe with increased product tolerance is essential. Another feasible approach is to improve maximum specific rate of substrate-product transformation. Improvement in the rate of flux of carbon can be made in a number of ways [2]:

392

Increasing the amount of key rate-limiting enzymes present in the organism by genetically increasing gene dosage or changing the controlling mechanisms for regulation of enzyme synthesis. Enhancing enzyme synthesis by removal of inhibitors produced during fermentation. Modification of enzymes by site-specific mutagenesis which alter kinetics of substrate utilization and end product formation. This overall approach for strain improvements is more promising and requires intensive research on the physiology, biochemistry and genetics of solvent producing microorganisms. Tolerance to substrate is not as significant in open continuous systems as substrate levels are usually low. In batch culture, however, it can be a significant factor when high product concentrations are required. When complex substrates such as wood hydrolysates are used, toxic products can exist and inhibit microbial growth significantly. Identifying the exact biochemical basis for toxicity is a prerequisite to develop strategies for strain improvement. The fermentation biocatalyst employed need to completely utilize substrate, prevent formation of by-products, and produce a final product concentration that allows for economic recovery. Thus overall emphasis of future genetic engineering research in this area would be to produce better fermentation biocatalysts with wider substrate ranges, and the ability to function under environmental conditions more suited to bioprocess engineering.

7

REFERENCES

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397

Index

Acclimatization, 240 Calcium magnesium acetate, 250 Acetic acid, 249-255,321,325,326 Candida Acetogenium kivuL 252 acidothermophilum, 377 Acetoin, 130,228 arborea, 280 reductase, 130,131 blankii, 305,306 Acetone, 198 boindii, 277,289 Acid hydrolysis, 86 curvata D, 311 kinetic model of, 89 guillermondii, 260,282,283 Active transport, 100,101,104 mogii, 279 Adaptive modifications, 327,328 parapsilosis, 281 Aeration conditions, 183 pelliculosa, 284 Aeromonas hydrophila, 224 shehatae, 104,150,156,157 Agricultural residues, 6,81 tropicalis, 156,169,286 Alkali treatment, 79 utilis, 377 Anti-freeze agent, 221 Carbohydrates, 4 Arabinans, 44 uptake, 102 Arabino-4-O-methyl-glucuronoxylan, 48 Catabolite repression, 132,133 L-Arabinose Cell recycling, 158 binding protein, 107 Cellulase, 52 metabolism, 136-138 Cellulose, 13-15 transport, 107 Chaetomium cellulolyticum, 381 Arabinoxylan, 42 Citric acid, 258-261

Aspergillus niger, 260,261,263,265,31 3 oryzae, 332 terreus, 263,264 Autohydrolysis, 91 Autolysin, 211

Bacillus macerans, 164 polymyxa, 223 stearothermophilus, 164 Bioengineering, 371 Biomass, 2,24,25,381 Bioreactor, 265 2,3-Butanediol, 130,131,221 fermentation, 229,230 production, 223-224 stereoisomers, 222

Clostridium acetobutylicum, 106,130,199-212 beijerinckii, 130 butylicum, 202,204,207,208 propionicum, 262 saccharolyticum, 150 thermoaceticum, 250,325 thermocellum, 150,165,323,324, thermohydrosulfuricum, 164,330 thermosulfurogenes, 164 Coimmobilized cells, 284 Conjugative transposon, 356 Continuous culture, 215,231 Corynebacterium sp., 287 Dissolved oxygen, 237 Distillation, 383 Downstream processing, 382

398 Economic evaluation, 387 Energy demand, 1 Enrichment, 352,354,356 Enterobacter liquefaciens, 287 Enzymatic analysis of hemiceilulose structure, 52 Enzymatic hydrolysis of heterogalactans, 62 heteromannans, 63 heteroxylans, 57 Enzyme-aided bleaching, 74 Ethanol, 147 production, 149 tolerance, 317,323,329,333-335 Extractive fermentation, 375 Extraneous materials, 22,23

Genetic recombination, 356 Glucans, 17 Glucomannan, 50,51 Growth characteristics, 150 Growth kinetics, 156,201,221

Hansenula polymorpha, 285,305,309 Hardwoods, 10 Hemiceilulolytic enzymes, 55 Hemicellulose, 16 biodegradation, 54 biosynthesis, 33 chemistry, 43 isolation, 80 Hemicellulosic substances, 33 Hybridization, 356 Hydrolysate bagasse, 177 cellulose, 175 hemicellulosic, 175 lignocellulosic, 174 wood, 234

Facilitated diffusion, 100,102,104 Fermentation acetone-butanol, 197 2,3-butanediol, 221 design, 374 ethanol, 147 Fibrobacter succinogenes, 106,269 Fodder yeasts, 304 Immobilized cells, 163 Forest residues, 10 Industry Fuel, 197 fermentation, 197 Fumaric acid, 266-268 food, 266 Fungal strains, 153 petrochemical, 197 Furfural, 373 plastic, 266 Fusarium oxysporum, 123,135,150,322 pulp and paper, 206 Inhibitors, 187,242 Itaconic acid, 262-266 Galactanases, 61,62 Galactans, 51,52 Galactoglucomannan, 49 Kinases, 35 Gasoline, 147,148 Klebsiella pneumoniae, 130, 167,180 Gene Kluyveromyces alcohol dehydrogenase, 361,362 cellobiovorus, 157 pyruvate decarboxylase, 361,362 marxianus, 167 xylitol dehydrogenase, 359 xylose isomerase, 360,363 xylose reductase, 359 Lactic acid, 255-258 xylulokinase, 360,363

399

Lactobacillus lactis, 257 xylans, 256 xylosus, 257

Nuclear magnetic resonance, 52,103 Nutrition, 180,212

Lignin, 20-22 Lignocellulosic materials, 9,25 resources, 3 wastes, 7 Lipid composition, 327-330

Oleaginous yeasts, 311 Organic acids acetic, 249 citric, 258 formic, 268 fumaric, 266 itaconic, 262 lactic, 255 propionic, 261 succinic, 268 tolerance to, 336 Oxidative reductive pathway, 120-123 Oxidoreductase, 135 Oxygenation, 133,182,213,214,291 Oxygen transfer rate, 237

Mannanase, 63 Mannans, 50-51 Materials extraneous, 22 lignocellulosic, 9 starch containing, 2 sugar containing, 2 Membrane lipids, 332 potential, 101 tolerance to solvents, 332 Mesophilic bacteria, 155 Metabolic inhibitors, 186 Metabolism endogenous, 237 xylose, 119 Methanobacterium sp., 284 Methyl ethyl ketone, 221 Milling, 77 Mixed acid fermentation, 268,269 Monilia sp., 162 Mutagenesis uv, 353-355 chemical, 354 Mutants tolerant to acetic acid, 339,340 butanol, 339,340 ethanol, 337,339 Mycobacterium smegmatis, 287

Pentosan composition of hardwoods, 46 softwoods, 48 Pentose isomerization, 168 Pentose phosphate pathway, 121,128 Pentose uptake bacteria, 105-108 genetic studies, 113-114 regulation, 109-113 yeasts, 101 - 105 Pet operon, 362,363 Petromyces aibertensis, 288 Phosphoketolase, 126 Phosphorylation, 126

NAD-xylose dehydrogenase, 123 Neurospora crassa, 162

Pichia heedii, 103,104,110 miso, 275 stipitis, 103,104,132,156

Pachysolen tannophilus, 132,156,286 Paecilomyces sp., 162 Passive diffusion, 100 Pdc gene, 362

Pediococcus halophilus, 112 Penicillium chrysogenum, 136

400 Plant cell wall, 12 polysaccharide, 44 structure, 12 Plasma membrane, 99,317 Pleurotus sajor-caju, 308,309,381 Polyporus anceps, 253 Pretreatment chemical, 79 irradiation, 78 physical, 76 thermal, 91-92 Process evaluation, 371 Propionibacterium acidi-propionici, 262 jensenii, 261 Propionic acid, 261,262 Protoplast fusion, 337,357,358 interspecific, 358 Pulp dissolving, 72 kraft, 74 sulfite, 73 Pyruvate ferredoxinoxidoreductase,128 Reactor systems cell recycling, 163 continuous, 172,215,264 fed-batch, 172,231,251 immobilized, 163,216 membrane, 284 repeated batch, 264 Repititive isomerization, 286 Residues agricultural, 6 industrial, 8 lignocellulosic, 11 Rhizopus arrhizus, 267 Rhodotorula glutinis, 102 Ruminococcus albus, 269

Saccharomyces cerevisiae, 159 lipolytica, 260

Salmonella typhimurium, 106,108,111 SASL, 3O7 Schizosaccharomyces pombe, 159 Screening, 352 SEHAL, 306,307,309 Selenomonas ruminantium, 107,108 Serratia marcescens, 225 Single cell protein (SCP) 301,302,351 production from, filamentous fungi, 304,308,309 yeasts, 304,307,309 Single cell oil (SCO) 301,302,351 production from, fungi, 312 yeasts, 311,312 Softwoods, 10 Solvent effect on cellular physiology, 318 toxicity, 318 Solventogenic clostridia, 199,203 Stereoisomers, 222 Straw, 11 Stress proteins, 331 Sugar nucleotides interconversion, 38 polymerization reaction, 40 synthesis, 34 Sulfite waste liquor, 225 Supplementation, lipids, 185,333-336 magnesium, 293 methanol, 296 Symport, 101-106 Thermoanaerobacter ethanoficus, 164 Thermoanaerobes, 165 Thermophilic bacteria, 154,163 Thermotolerance, 155 Tolerance acetic acid, 321,326 2,3-butanediol, 321 butanol, 320,335 ethanol, 317,323,329,333-335 genetic basis, 336

4Ol Trace metals, 241 Transaldolase, 131 Transketolase, 131 Uptake L-arabinose, 107 D-ribose, 109 D-xylose, 99,103-105 Waste agricultural, 207 sulfite liquor, 207 Water activity, 239 Whole cell immobilization, 170 Xylans, 17,46-50 isolated, 75 Xylanase, 56-59 Xylitol, 273 Xylitol dehydrogenase, 132 Xylonic acid, 123 D-Xylose, 159 oligosaccharides, 59 Xylose isomerase, 124,125,133 Xylose reductase, 120-122, 132 Xylulokinase, 126,133 D-Xylulose, 159 D-Xylulose-5-phosphate, 120,126,127 Yeast strains, 151

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