Microbial Pentose Utilization: Current Applications in Biotechnology

Microbial Pentose Utilization

Current Applications in Biotechnology

Vol. Vol. Vol. Vol. Vol. Vol.

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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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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

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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

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

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 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 0.1

Hardwoods

Black cherry

45

20

21

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

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

HO- I .

CHzOH

L-Arabinose

IHO H- -OH H- IOH

H- I-OH

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

IFERMENTAT~)N I

I HYDROGENATION,,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

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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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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

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29 67

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

68

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69

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73

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74

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79

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8O

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84

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86

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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 H~

O-UDP

HO

O-UDP II

4-Epimerase

OH

HO

H0~ O N O

O--UDP

CHzOH HO

O-UDP

(b)

(a)

HO

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

COO-

2 NAD§ 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 (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

Arabinose

Glucose

0.90 0.90 0.74 0.74

tr tr tr tr

0.05 0.05 0.05 0.05

60

nd

nd

nd

70

nd

nd

nd

80 90

0.90 0.96

tr Tr

0.05 0.10

Temperature

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

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

5 63 20 52 1 19 55

Pine

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

(%)

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 Untreated

Yield b

Sweet sorghum bagasse 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

No Yes

49.2 45.1

19.5 17.2

21.1 24.4

3.8 2.8

Douglas fir

No Yes

54.9 57.1

4.5 3.4

33.5 32.8

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

REFERENCES

Ghose TK, Ghosh P. Proc Biochem 1975; 6:611. Lee YY, Lin CM, Chambers RP. Biotechnol Bioeng Symp 1979; 8: 75. Ghosh P, Singh A. Adv Appl Microbiol 1993; 39: 291. Yu EKC, Deschalteles L, Louis-Seize G, Saddler JN. Appl Environ Microbiol 1985; 50: 924. Yu EKC, Saddler JN. Trends Biotechnol 1985; 3: 100. Wong KKY, Deverall KF, Mackie KL, Clarke TA, Donaldson LA. Biotechnol Bioeng 1988; 31:447. Viikari L, Kantelinen A, Ratto M, Sundquist J. ACS Symp Ser 1991; 460: 12.

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10 11 12 13 14 15 16 17 18 19

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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

Low

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

[10]

a

[lo] [lo]

15.0b 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

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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
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

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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.

30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47

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.

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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

Ethanol Butanol

Saccharomyces cerevisiae

Candida shehatae Pichia

stiptis

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

Inhibits biomass production but not the product yield Inhibits biomass production but not the product yield Inhibits growth Acts as uncoupler and allows

[68] [69,70]

protons to enter the cell

Acetic

C los tridiu m

acid Butyric acid

acetoaceticum Clostridium a ce to butylicu m

Increases H§ activity and reduces cellular ATP content

[71-74]

Inhibits growth

[75]

Inhibits growth Diminishes pH gradient

[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]

Z. mobilis Clostridium thermocellum ATCC 27405 C. thermosaccharo-

lyticum

Adaptation and selection

ThermoanaeroUV-mutagenesis bacter ethanolicus Selection in Saccharomyces cerevisiae continuous 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

S-6 C-9 HG-3 HG-4 HG-6 HG-6-610 JW200

[179] [181] [179] [91] [90]

SB154 SB155 SB159 SB160 FDHS

[171]

SA- 1

[185]

904

[186]

lyt-1

[187]

S-3

[103]

1745

[103]

[172]

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)

-I

STREAM I----" XYLOSE-RICH

DILUTE ACID HYDROLYSIS

GLUCOSE-RICH STREAM

(8%HzS04)

IELECTRODIALYSISI ~-. GLUCOSE l-..

- I XYLOSE I

~ 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

":,~

LIGNIN

" 1 HEMICELLULOSE !

=Ic ,,u,os I

ETHANOl_ TREATMENT ~ _ _

LIGNIN ETHANOL

SIMULTANEOUS SACCHARIFICATION FERMENTATION

Yeast

ETHANOL

l. cov .Y1

Cellulase enzyme

RESIDUE

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.

I

379

BIOMASS (WOOD CHIPS)

I SUGARSYRUPI I HYDROLYSIS !

~1 BUTANOL I

FELLULOSEI

ITREATMENT

I BUTANOL PHASE I BUTANOL I RECYCLING LIGNIN RECOVERY

ILIGNINI

SOLVENT RECOVERY

SIMULTANEOUS SACCHARIFICATION FERMENTATION

/

! 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

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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

Lignin, 20-22 Lignocellulosic materials, 9,25 resources, 3 wastes, 7 Lipid composition, 327-330 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 NAD-xylose dehydrogenase, 123 Neurospora crassa, 162

Nuclear magnetic resonance, 52,103 Nutrition, 180,212 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

Pachysolen tannophilus, 132,156,286 Paecilomyces sp., 162

Passive diffusion, 100 Pdc gene, 362

Pediococcus halophilus, 112 Penicillium chrysogenum, 136

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

Pichia heedii, 103,104,110 miso, 275 stipitis, 103,104,132,156

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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