Advances in Applied Microbiology, Volume 39

ADVANCES IN

Applied Microbiology VOLUME 39

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ADVANCES IN

Applied Microbiology €Edited by SAUL NEIDLEMAN Vacaville, California

ALLEN I. LASKIN Somerset, New Jersey

VOLUME 39

Academic Press, Inc. A Division of Harcourt Brace S.Company

San Diego New York Boston London Sydney Tokyo Toronto

This book is printed on acid-free paper. @

Copyright 0 1993 by ACADEMIC PRESS, INC. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher.

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United Kingdom Edition published by

Academic Press Limited 2 4 2 8 Oval Road, London NW1 7DX International Standard Serial Number: 0065-21 64 International Standard Book Number: 0-12-002639-2 PRINTED IN THE UNITED STATES OF AMERICA

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CONTENTS

Asepsis in Bioreactors

M . C. SHARMA AND A . K . GURTU I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11. “Invaders” . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Consequences . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

IV . Sources ............................................................................... V . Approaches . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI . Overcautious Approaches ......................................................... VII . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1 3 3 4 6 24 25 25

Lipids of n-Alkane-Utilizing Microorganisms and Their Application Potential

SAMIR S . RADWANAND NASERA . SORKHOH I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I1. n-Alkane-Utilizing Microorganisms ....................................... 111. Total Lipid Contents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV . Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V . Acylglycerols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI . Sterols ................................................................................. VII . Fatty Alcohols, Ketones, and Epoxides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VIII . Hydrocarbons and Waxes .......................................................... IX . Phospholipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . X . Glycolipids and Peptidolipids ..................................................... XI . Biolipid Extract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . XI1. Environmental Considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

29 30 35 42 59 61 63 66 67 73 76 78 81

Microbial Pentose Utilization

PRASHANTMISHRAAND AJAYSINGH I. 11. 111. IV . V.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pentoses from Natural Sources . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pentose-Fermenting Organisms ................................................... Pentose Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Production of Solvents and Organic Acids ......................................

V

91 93 94 101 112

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CONTENTS

VI . Factors Affecting Pentose Fermentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII . Product Tolerance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VIII . Strain Improvement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IX . Future Prospects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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136 139 142 143

Medicinal and Therapeutic Value of the Shiitake Mushroom

S. C. TONG

AND J

. M . BIRMINGHAM

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I1. Medicinal and Therapeutic Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Patented Products and Processes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV . Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

153 154 172 175 177

Yeast Lipid Biotechnology

Z . JACOB I. I1. 111. IV . V. VI . VII .

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yeasts as Potential Sources of Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Importance of Yeast Lipids in Beverages and Foods . . . . . . . . . . . . . . . . . . . . . . . . Medical Importance of Yeast Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Modification of Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Commercial Significance of Yeast Lipid Biotechnology ....................... Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

185 186 187 191 192 204 207 208

Pectin. Pectinase. and Protopectinase: Production. Properties. and Applications

TAKUOSAKAI. TATSUJI SAKAMOTO. JOHAN HALLAERT. AND ERICKJ . VANDAMME I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Review of Pectin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Classification of Pectic Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Role of Pectic Enzymes in Phytopathogenesis .................................. Applications of Pectinases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Protopectin-Solubilizing Enzyme (Protopectinase) ............................. References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

I1. 111. IV . V. VI .

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Physicochemical and Biological Treatments for Enzymatic/ Microbial Conversion of Lignocellulosic Biomass

PURNENDU GHOSHAND AJAYSINGH I. I1. I11. IV . V. VI . VII .

Introduction .......................................................................... Structure of Lignocellulosic Biomass ............................................. Physical Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Thermal Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chemical Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biological Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Epilogue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . OF PREVIOUS VOLUMES. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . CONTENTS

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Asepsis in Bioreactors M. C. SHARMA~ AND A. K. GURTU’ SOL Antibiotics Limited Hyderabad 500 482, India

I. 11. 111. IV.

Introduction “Invaders” Consequences Sources A. Inoculum B. Nutrient Medium C. Bioreactor System D. Air/Liquid Transfer E. The “Rogue” V. Approaches A. Sensitive Sterility Assessment Methodology B. Certified Aseptic Laboratory Inoculum C. Autoclavable Bioreactor D. Sterile Mediurn/Feed E. Aseptic Bioreactor System F. Maintenance of Asepsis during Fermentation G. Protected Fermentation H. Product Changeover I. Schedules and Procedures VI. Overcautious Approaches VII. Conclusion References

I. Introduction

The key process of biotechnology is fermentation, and the key equipment, or the heart of the process, is the bioreactor-fermentor. The fermentation process generally employs pure culture as the biocatalyst; the success of the process depends, to a large extent, on ensuring asepsis in the fermentation system and in the process. Although the terms “asepsis” and “sterility” in fermentation are microbiologically incorrect, they have been generally accepted. In fact, the term “monosepsis” would be more correct. Asepsis in biotechnology means freedom from unwanted microorganism(s), just as in clinical medicine it means freedom from pathogenic microorganism(s) (Bull et a]., 1983). Absolute sterility is a concept in probability and is an unattainable ideal in

’

Present address: Biotech International Ltd., VIPPS Centre, Masjid Moth, Greater Kailash-11, New Delhi 110 048, India. * Present address: J. K. Pharmachem Ltd., Madras 600 014, India. 1 ADVANCES IN APPLIED MICROBIOLOGY, VOLUME 39 Copyright 0 1993 by Academic Press, Inc. All rights of reproduction in any form reserved.

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practice. A low-level, contaminated batch in a bioreactor, with normal product biosynthesis, achieves the empirical sense of “sterility,” though not the absolute one (Reisman, 1988). There are many industrial fermentations, e.g., ethanol, baker’s and fodder yeasts, and vinegar, where no serious attempt is made to maintain asepsis either in the fermentor medium or in the subsequent conduct of the process. In fact, such attempts are not warranted because of rapid culture proliferation, rapid metabolic transformations, and the resultant environment being generally nonconducive to contaminant growth (Herold and Necasek, 1959; Bailey and Ollis, 1986). It was during the first world war that Weizmann made the pioneering efforts to establish the first truly aseptic acetone-butanol fermentation (Hastings, 1978). However, the inactivation of penicillin by penicillinase-producing microbes necessitated engineering developments aimed at carrying out biotechnological processes with absolute exclusion of foreign microbes. The fermentation industry does not publish figures on the rate of nonsterile operations, yet a figure of 5-30’/0 nonsterility of bioreactors is realistic (Saudek, 1956; K. Gerlach, unpublished communications, 1986). At times the rise in figures warrants being called a “wave” of contamination. Economic considerations indicate that a contamination probability of 1in a 100 is acceptable for batch fermentations, considering the norm of 1 in 1000 as the probability of contamination commonly employed in design calculations for a sterilization process (Banks, 1979). A nonsterility rate of 1% or less is often regarded as a commendable performance (Soderberg, 1983). Artificially selected industrial microbes generally used in biotechnology endeavours are at risk of being overwhelmed by competing wild organisms. Mammalian/animal/plant cell cultures are especially prone to microbial contamination because of a long process cycle (20 or more days] and a relatively slow growth rate (doubling time as long as 100 hr). These cell cultures can be compared to an artificial organ without autoimmune protection, lacking any defense system, and therefore, extremely vulnerable to a breach of sterility (Knight, 1989).Yet the industry now operates large-scale animal cell cultures routinely with contamination rates of just 2% (Spier, 1988), even with culture lengths of several months and intermittent additions of fresh medium. The physicochemical environment generally maintained in a bioreactor is optimal for a host of microorganisms. Very rarely is the medium “protected”, i.e., selectively utilizable by a limited range of microbes (Stanbury and Whitaker, 1984) or else has an antimicrobial added to it. The intended metabolite, even if an antimicrobial, is normally produced late (in idiophase). Very often, the bioreactor in trophophase

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would therefore be an ideal incubator for numerous microorganisms, if provided an opportunity of entry. II. “Invaders”

The microorganisms that invade the sterile fermentation process are bacteriophages, mycoplasma, bacteria, or fungi. Phages normally infect through air and the development of phage-resistant strains has resulted in the rarity of such infections, particularly in view of the host specificity of phages. Mycoplasma and viral infections are more common in animal cell cultures through serum and can be eradicated easily (Arathoon and Birch, 1986). The microsize, omnipresence, and faculty of utilizing widely varying substrates for nutrition make the bacteria capable of causing widespread infections and maximum damage to the fermentation process. While gram-positive bacteria have air as their main source, gram-negative bacteria are transmitted through liquids, particularly water. Among the fungi, yeasts may originate mainly from insufficient sterilization of substrates. Filamentous fungi have air as their main source. Fungal infections occur rarely (Herold and Necasek, 1959). 111. Consequences

The invasion of a fermentation process in a bioreactor by a foreign microorganism can result in a variety of consequences: 1. A fast growing, wild contaminant may outcompete the normally slow growing desired strain, deplete the nutrients, and fatally interfere with the chemistry of the process and the final product. 2. The invader may not outgrow the desired strain, but may cause minor to appreciable alterations in the physicochemical characteristics of ongoing fermentation. The contaminant may also produce undesirable and possibly toxic metabolites that may lead to lower yields and productivity. 3. The contaminant may grow to a certain level and subsequently be inhibited by the metabolite(s) produced by the desired strain. The fermentation may continue to its logical end, as at times in the case of broad spectrum antibiotics. 4. Contaminants, i.e., phages, could result in the lysis of the desired microbe in a bacterial/actinomycete fermentation. 5 . Mucilage/slime produced by the contaminant may choke the filter pores to varying degrees, which in a worst-case scenario would ruin the entire batch on hand.

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6. Another consequence of nonsterility could be the degradation/ racemization of the desired metabolite, e.g., the enzymatic degradation of p-lactam antibiotics by p-lactamase-producing bacteria, leaving behind a totally unproductive batch or producting DL- or D-amino acids in an L-amino acid fermentation. 7. Undesirable moieties produced by the contaminant may lead to interference in the downstream recovery of the product, resulting in not only lower yields but a substandard product as well. The processing of the product may cause increased production costs. 8. The contaminant may render the final product unusable, e.g., single cell proteins where the cells constitute the product. 9. Not every contaminant at the fermentation stage exerts detrimental effects. The contaminant has been reported to increase fermentation yields and to better downstream processing (Reisman, 1988) due to the presence of useful enzymes like proteases and lipases.

Irrespective of the consequences of contamination, preventing the entry of contaminants is necessary. The attempt, therefore, should be to identify the sources of contamination and to conduct fermentation processes in an aseptic manner. IV. Sources

The sources of contamination in a fermentation process can be broadly ascribed to several factors. A. INOCULUM Contamination at the inoculum development stages assumes greater significance in view of the consequent nonavailability of quality “starter” material for the bioprocess and therefore the opportunity loss. The presence of a contaminant in laboratory-grown inoculum, often in concentrations low enough to escape detection up to seed bioreactor maturity, can lead to the subsequent manifestation of nonsterility in the fermentation bioreactor. In view of the relatively small volumes used for sterility testing, sensitive methodologies are obligatory. The sources of laboratory inoculum nonsterility in turn could be autoclaving deficiencies, “lumpy” medium, inadequate maintenance of sterile chambers, ineffective ultraviolet irradiation from germicidal lamps, use of inefficient germicidal solutions, wetting of plugs, and insufficient/ prolonged stocking of sterilized media/glasswares. Inadequately screened cell banks are potential sources of viral contamination in animal cell culture processes (Arathoon and Birch, 1986).

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B. NUTRIENT MEDIUM Nonsterility of the nutrient medium used in the seedifermentation bioreactor may culminate in a contaminated operation. An inadequate sterilization operation, “lumpy” medium, and the bioreactor itself may contribute to medium nonsterility. The choice between batch and continuous sterilization and their design depends on the scale of operations and the characteristics of the medium. In the case of separate medium sterilization, the tank/continuous sterilizer, piping, and receiving vessel could be sources of nonsterility. Animal sera used for cell culture processes may sometimes carry viral and other contaminants and render the process nonsterile (Arathoon and Birch, 1986; Maurer, 1986; Elander, 1989).

C. BIOREACTOR SYSTEM A continuous stirred tank reactor is the most commonly used bioreactor in view of its versatility and flexibility of operations therein. The sterility considerations of such a bioreactor satisfactorily cover most of the other types of bioreactors as well. The design, material, and fabrication of the bioreactor are important factors contributing to the sterility of operations. Apart from these factors, aberrations in fermentor sterilization, air supply, agitation system, sampling and monitoring ports, and uncontrolled foaming could lead to invasion of the bioreactor boundary by undesired microorganisms. The agitator shaft entry and the air exit are the most vulnerable points in a bioreactor. All types of seals, except a double mechanical seal, provide gaps as entry points for contaminants (Steel and Miller, 1970; Bull et al., 1983; Aiba et al., 1986; Reisman, 1988). Intermediate bearings on shafts, improper impeller hubs and keyways (Reisman, 1988), and pipe in pipe connections provide unhygienic places to harbor contaminants. Too many interior fittings create pockets likely to conceal microbes, are difficult to sterilize, and can result in contamination of the bioreactor. Stress corrosion/cracking in the vessel and internal coil leads to repeated contaminations. Hammering caused by steam during batch sterilization causes stress on spargerflanged joint components and creates nonuniform steam distribution-related sterility hazards. The numerous ports, pipes, and valves required in a fed-batch process are further sources of microbial entry if not properly selected or designed. Flangedhhreaded joints, pervious sealing materials, defective slopes of pipes, dead ends, pockets, indentations, crevices, solid depositions, stagnant layers, rising stem valves, leaking or “weeping” pipes/ flangeshalves, and the absence of steam seals/crosses are potential

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causes of contaminations. The continuous presence of humidity/condensed moisture and entrained medium in the air exhaust area provides an environment conducive to microbial growth at this boundary point of direct contact between sterile and nonsterile zones. Uncontrolled foam generation during fermentation causing “foaming out” through the air exhaust increases the chances of fermentor contamination (Solomons, 1967; Ghildyal et al., 1988). D. AIRILIQUID TRANSFER

Contamination through the depth filter is possible because of inadequate filter packing, channeling, free moisture, or fiber fragmentation. Individually or collectively, these factors contribute to filter inefficiency. In the case of the membrane filter, damaged 0 ringsimembrane cause losses in filter integrity. The air filter, soaked in nutrient media because of backflowloverflow, can act as an incubator for contaminating microbes. The liquid transfer systems, including those for the inoculum, feeds, and supplements, may also contaminate the process because of system deficiencies described earlier and/or inadequate sterilization. A common system for transfers to several fermentors, if having such deficiencies, would be a catastrophe.

E. THE“ROGUE” For all practical purposes even the very rare appearance of a nonproducing wild “rogue,” which occurs due to reversion mutation of the production culture during the fermentation process, leads to contamination of the bioreactor. There are no precautionary or remedial measures against such mutant appearances and early downstream processing may be warranted for salvage, if any. V. Approaches

Ever since the aseptic submerged culture technique for penicillin production was introduced, there have been attempts to perfect fermentation techniques, design, and systems in achieving asepsis. Coordinated efforts of microbiologists, technologists, engineers, and biochemists are required to evolve various bioreactor contamination control strategies. The successful commercial scale production of numerous fermentation products is a measure of the magnitude of such achievements. Exhaustive reviews on such techniques have been made by Rhodes and Fletcher (1966), Solomons (1969, 1971), Augurt (1983), Wallhausser (1985), and Bailey and Ollis (1986).

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Aseptic design development is based on several norms by which the intrusion of foreign microbes into a defined fermentation system can be precluded. A closer look at the behavior of a microbe at different types of boundary layers reveals the following criteria (Lundell and Laiho, 1976) relevant to asepsis: 1. Nonpenetration of homogeneous solids by a microbe. 2. No movement or growth of a microbe through holes smaller than

its own dimensions. 3. No propulsion of a microbe against the flow of carrier medium. 4. No movement of a microbe on dry surfaces without external forces. 5. No growth on a surface with temperatures exceeding a microbe’s maximum growth temperature. 6. No growth on nonmetabolizable/hydrophobic/toxicmaterials. 7. Inactivation of a microbe by high temperature, toxic chemicals, and irradiations. 8. A characteristic doubling (reproduction) period of each microbe. Most of the aseptic design considerations involve any or all of the combinations of these criteria. The sterile design of a bioreactor normally adds 15-25% to the cost toward its design, purchase, and installation (Reisman, 1988). The following approaches constitute some of the techniques essential for achieving aseptic bioprocesses:

STERILITY ASSESSMENT METHODOLOGY A. SENSITIVE The absence of a contaminant(s) in the culture inoculum, sterilized seed/fermentation media, and equipment needs to be ascertained at every stage in order to avoid nonsterility and to detect the stage at which the contaminant invaded the process. Often the results of conventional sterility checks may not be available before the culture has reached the production bioreactor or before the contaminant has reached a growth level capable of disturbing the desired fermentation. Speed of detection is germane to the salvage of a batch. A sensitive sterility assessment methodology with accelerated results would be useful for such applications. The use of a variety of nutrient media in tubes (broth, slants, and stabs) and flasks incubated at varying incubation temperatures under statidshaken conditions would be ideal in covering a range of growth conditions required by different contaminants (Soderberg, 1983). The use of thioglycolate, with a small amount of agar (0.05%), and oxidoreduction dyes like methylene bluekesazurin have been recommended for the fast detection of aerobes as well as anaerobes in a single medium (U.S. Pharmacopoeia, 1980).

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M. C. SHARMA AND A. K. GURTU

Membrane filter discs could prove useful for sterility checks of broths containing antimicrobials. Penase (a potent lactamase preparation) is recommended for the inactivation of penicillin/p-lactams during sterility testings (Difco Laboratories, 1985). Sterilized filter assemblies with membrane filter discs/glass wool pads find application in sterility checks of air/liquids passed into the fermentation equipment. Trial runs with uninoculated lean nutrient media are used to ascertain the sterility status of fermentation equipment, transfer manifolds, and air systems during validation. The use of selected radioactive carbon sources in sterility check media has been recommended under the “Bactect” system for a fast detection of contaminants (McLaughlin et a]., 1983). The mass spectrophotometric identification of 3-hydroxymyristic acid, a characteristic of gram-negative bacteria, has been applied for a rapid sterility assessment by Elmroth et al. (1990).The techniques of sampling for sterility checks were described by Elsworth (1960), and newer techniques (Charton, 1990) involving the use of thermoplastic elastomer tubings for improving sterile access to bioreactors have been described. A mobile flexible film containment cabinet attached to the bioreactor is used in sampling recombinant DNA-based fermentation processes (Hambleton et al., 1991). Elander (1989) described the use of a sterile stainless steel container with a small sterilizing filter on its vent that was attached to the fermenter, union sterilized, sample drawn, and the union finally resterilized before disconnection and further processing of the sample drawn. For animal cell cultures, the establishment and testing of suitable cell banks with screening for freedom from viruses and other adventitious agents is a major exercise (Lubiniecki and May, 1985). Further work on accelerated sterility assessment methodology in the fermentation industry is needed. B. CERTIFIED ASEPTICLABORATORY INOCULUM

The inoculum preparation room needs to have a clean room design and scheduled validation/checks of this sterile room through exposure of plates. Schedules for area fumigation and in situ integrity testing of HEPA filters by aerosols/dioctyl phthalate/dioctyl sebacate ensure sterility in these rooms. The area used for inoculum preparation needs to be isolated from that, used for in-process sterility assessment. Only validated disinfectant dilutions should be permitted in sterile rooms. The rotation of disinfectants to avoid buildup of resistant microflora is necessary and must be practiced rigidly. Principles and practices of laboratory management related to facilities, design, decontamination, access to work place, personal hygiene, apparel, sanitation, ventilation,

ASEPSIS IN BIOREACTORS

9

and safety are amply described (Rhodes and Fletcher, 1966; Soderberg, 1983; Wallhausser, 1985; Scheirer, 1987). The careful planning of premises, air circulation, and pressure differentials has recently become important, particularly in culturing animal cells and genetically engineered microbes (Scheirer, 1987; Knight, 1989). Finch (1958), Sykes (1958), Borick (l968), Benarde (1970), and Wallhauser (1985) have discussed the utility of various disinfectants and chemical sterilants for such purposes. “Biosafety in Microbiological and Biomedical Laboratories” published by the U.S. Department of Health and Human Services (1984) describes the standard and special microbiological practice guidelines and designs for aseptic laboratory operations. The proper storage of biodegradable raw materials like corn-steep liquor, soya flour, corn flour, and seed meals under hygienic conditions and preferably at reduced temperatures and humidity minimizes the further increase of microbial load. Fine powders, presoaking and preboiling, proper batching sequence, and straining help prevent the “lumpy” medium threat to aseptic processing (Soderberg, 1983). Validation and proper operation of autoclaves used for media/glassware sterilization is mandatory for pure inoculum propagation. The proper venting of air and steam ensures the absence of air pockets necessary for attainment and sensing of correct uniform temperature in the autoclave. Air, being heavier than steam, must also be expelled from the lowest level in the autoclave. Using a jacket steam ejector to remove leftover steam in the autoclave chamber after sterilization is advisable in order to minimize moisture condensation on sterilized wares (Wilkinson and Baker, 1964). Autoclave performance may be checked using “biological indicators” (Banks, 1979) or sterilizing temperature indication stickers/tapes, like those supplied by the 3M Corporation, to confirm proper autoclave operation. Such indicators have been reviewed by Augurt (1983) and Wallhausser (1985). Procedures obligatory for efficient performance of autoclaves have been described by Rhodes and Fletcher (1966), Stumbo (1976), Augurt (1983), and Wallhausser (1985). To avoid dirt accumulation, it is advisable to use rimless glassware and presterilized cotton/synthetic hydrophobic plugs. Needless to say, wrapping with waterproof papers/aluminum foil, vacuum drying through steam ejection, and hot air oven drying of autoclaved materials is very useful. Incubation of sterilized media at different temperatures for 72 hr and subsequent checks ensure their sterility before use. A need-based planning of sterile glassware and media is necessary to avoid the “emergency” use of freshly autoclaved material or prolonged stocking of autoclaved material that could be a “sterility risk,” particu-

10

M. C. SHARMA AND A. K. GURTU

larly in a humid climate. Longer necks and optimum degree of filling of containers prevent the plugs from getting wet during incubation on shaker. Attention to these details results in significant improvements in sterility at the laboratory stage. C. AUTOCLAVABLE BIOREACTOR

Smaller glass or stainless steel bioreactors (1-5 liter capacity) containing medium can be sterilized by autoclaving and later connected aseptically to various nutrients, air, pH, and pressure maintenance lines for operations. Bioreactors with capacities higher than 5 liters are difficult to sterilize in an autoclave, and the possibility of sepsis is increased if connections of larger diameters are used (Solomons, 1969). D. STERILE MEDIUMIFEED

The precautions mentioned earlier for laboratory medium preparation are necessary for bioreactor medium preparation as well. Medium mixing facilities are often neglected in the fermentation industry. Tank design for total drain out, smooth internal finish, high-pressure water jet cleaning after each operation, and chemical cleaning at regular intervals eliminate the formation of dried medium crusts and discourage microbial buildup in the tank. Sterilization of industrial media is not possible through filtration [which in fact is not recommended because suspended matter is a desirable ingredient of the medium) and is unreliable through irradiation or chemical sterilants. Steam sterilization is, therefore, the only choice and can be carried out in the bioreactor itself or in a separate pressure vesselfcontinuous sterilizer. Aseptic considerations warrant a preference for steam sterilization over other alternatives. 1. Batch Sterilization

Indirect heating through a jacket, external or internal coils, hollow baffles, and steam sparging through an air delivery system and dip pipes or any combination of the aforementioned are used for batch sterilization. Efficient mixing and circulation of the medium promote the efficiency of heat transfer and ensures the uniform heating essential for proper sterilization. In such a case the vessel also gets sterilized along with the medium. It is imperative that steam is supplied to the bioreactor through all dip pipes or ports in direct contact with the medium [Bull et a]., 1983; Stanbury and Whitaker, 1984). In fact, it is desirable to continue regulated steam supply at these points throughout the sterilization cycle since the stagnation of improperly heated medium

ASEPSIS IN BIOREACTORS

11

inside the pipe section, external to the bioreactor, is a serious sterility hazard. Relative merits of in situ sterilization and the use of separate pressure vessel have been discussed in detail by Richards (1966,1968). More often it is the in situ sterilization that is preferred. 2. External Continuous Sterilization

The external continuous sterilizer involving high temperature short time treatment (Baily and Ollis, 1986) is the method of choice when large volume bioreactors are used. The advantages of continuous sterilization of initial medium as well as nutrient feeds (Aunstrup et a]., 1979) over batch sterilization have been comprehensively described by Solomons (1969, 1971), Augurt (1983), Banks (1979), Cooney (1985), Wallhausser (1985), and Aiba et al. (1986). A shorter process time, better heat recovery, better medium quality, and cost effectiveness are ensured. The design of a continuous sterilizer is extremely important. Plate heat exchangers are suitable for media containing low levels of suspended solids. For media containing substantial solids or for viscous media, tubular heat exchangers with high flow rates and turbulent flow are used (Bull et al., 1983; Cooney, 1985). The sterilization of oils and viscous antifoams has to be planned carefully (Bader et al., 1984). Oils free of moisture are often difficult to sterilize. It is preferable to mix water in oils/antifoams before sterilization and to use a tubular continuous sterilizer. A spiral heat exchanger and an injector-flash cooler sterilizer (Banks, 1979; Stanbury and Whitaker, 1984) are useful versions of continuous sterilizers. The replacement of process water, held up inside the heat exchanger, with demineralized water prior to system sterilization with steam or superheated water (125°C) is necessary for eliminating stress corrosion cracking (Soderberg, 1983). Designing the sterilizer with near plug flow, proper retention, automatic control of the sterilization temperature, monitoring of inlet as well as outlet temperature at the holding section, and automatic switch over to the recirculation mode with a simultaneous shut off of the delivery line to the bioreactor when the sterilization temperature drops are mandatory features for aseptic performance of the system. Figure 1 represents a standard external continuous sterilization system for medium and feeds for bioreactors. Installation of a conductivity probe in the cooling water exit enables instant detection of a leak in the system. 3. Sterilization by Filtration

Sterilization through filtration by “depth” or membrane filters is a choice for clear media, especially where volumes are relatively low [Bull

M. C. SHARMA AND A. K. GURTU

12

Sterilized

* productto

bioreactor

Water detergent

Cooling Water

II Raw Product FIG.1. A typical external continuous sterilization bioreactor system. (1) Balance tank. (2) Flow control. (3) Interchanger. (4) Heater. (5) Holding tubes. (6) Cooler. (7) Controls. (8) Steam.

1 1

et al., 1983; Reisman, 1988).Membrane filters have superior operational performance because of a fixed, well-defined pore size. Sterilizable, cleanable, and reusable effective filter components have been developed and are used for plant and animal cell cultures to avoid loss of nutrients in media due to steam sterilization. Cross flow membrane cassettes, plate, disc, or cylindrical cartridge units, and even ceramic cartridges are now available for repeated sterilization operations. Their life can be prolonged by employing coarse prefilters or screens. Advance sterilization of the bioreactor and transfer lines is a prerequisite for medium sterilization through a separate pressure vessel, continuous sterilization, or filtration.

E. ASEPTIC BIOREACTOR SYSTEM In systems with strict aseptic requirements the material of construction and design requires detailed attention. The necessity of maintaining sterility has a far reaching impact on equipment design, piping, and layout (Elsworth, 1960; Soderberg, 1983; Bjurstrom, 1985; Reisman, 1988). 1. Bioreactor Internah

An aseptic bioreactor requires internals without niches and crevices to ensure steam availability to the entire surface. Internals with smooth contours ensure cleanability and complete drainage to exclude areas where condensates collect and to insulate microbes from sterilizing temperatures. Repeated sterilizations should leave no pits or a corrosive buildup in the vessel. The choice of material for such fermentation systems is restricted to borosilicate glass or stainless steel 316L (less than 0.03% carbon) with 150 grit or electro-polished internal finish

ASEPSIS IN BIOREACTORS

13

(Bull et al., 1983; Bjurstrom, 1985; Reisman, 1988). Fibreoptics and diamond sensor instruments devised by reputable bioreactor manufacturers are currently in use for inspection of welded joints and internal surface profiles. High surface qualities with grain 400 and roughness values of less than 0.06 p m are requested presently. The entries into the fermentor should be restricted to the bare minimum (Bjurstrom, 1985; Van Brunt, 1987; Reismann, 1988). The air supply pipe does not need to run through the top-head to down below the bottom impeller but instead can be introduced through the bottom cylindrical portion, This avoids “cake” formation on the pipe above the agitated broth level, particularly in the case of maintenance of the elevated inlet air temperature, thus avoiding a potential contamination pocket. Similarly, internal cooling coils should be avoided as much as possible to reduce brackets, supports, and weld joints. Internal fittings, when absolutely necessary, need to have a clear gap from the fermentor wall to facilitate proper cleaning. The use of pipes for brackets, ladders, and supports is forbidden. The ring type air sparger should have holes on its lower side for drainage of broth when the vessel is emptied. 2. Agitator Shaft and Seal

A stirrer shaft seal is the most difficult to construct in a fermentor. A side or bottom entry shaft (Richards, 1968) is undesirable since the bearings would be submerged. Eighty percent of all fermentor installations have top entry agitators (Sittig, 1982). Simple stuffing box and bush seals are potential sterility hazards because of frictional damage to packing, uncertain sterilizations, and oozings. Stellite hardening of the agitator shaft in the stuffing box region helps in maintaining the shape of the shaft, which normally gets worn and leads to increased leakages, Chemicals are sometimes used to disinfect the area. A double mechanical stainless steel 316 seal that can be sterilized with live steam and lubricated with cooled steam condensate is always preferred (K. Gerlach, unpublished communications, 1986; Liberman et al., 1986; Reisman, 1988). Seals able to withstand live steam/hot steam condensate lubrication have a decided edge over others. Polyethylene tetrafluoroethylene (PTFE) secondary seal, ethylene propylene dimer (EPDM), fluorocarbon and viton “0” rings, resin impregnated carbon/ tungsten carbide faces with a ceramic seat are the state of the art components of such seals (Fig. 2) like those from A.W. Chesterton Co. (Stoneham, MA) and John Crane, U.K. Ltd. (Slough, Berks, U.K.). Use of a back pressure indicator and regulator in the seal lubricant exit line helps ensure sterility and also acts as a forewarning device for loss of seal integrity. A low seal pressure/low level alarm system with an

14

M. C. SHARMA AND A. K. GURTU

Tungsten-Catbide stationary ring Cahn

Condensate out FIG.2. A state of the art double mechanical seal for a bioreactor agitator.

automatic stirrer switch off are features of recent versions (K. Gerlach, unpublished communications, 1986). Another attempt in solving the problems related to the sealing of the impeller shaft involved the use of a high-torque magnetic drive system (Cameron and Godfrey, 1969; Bull et aI., 1983). The driven magnet is on one end of the impeller shaft and the driving magnet is outside the vessel. Such magnetic drives, without piercing for an agitator shaft, can only be used for small-sized fermentors (Knight, 1989). Using neodymium-iron-boron supermagnets, up to 50 horsepower can be delivered to the agitator. The air lift bioreactor does not involve any moving parts within the bioreactor and obviates the need for piercing the shell for agitation. Such bioreactors are useful in animal cell cultures and in single cell protein fermentations (Smith, 1980; Arathoon and Birch, 1986). 3. Bioreactor Ports

Every port of entry and exit on the fermentor is a potential source of contamination and therefore has to be properly designed to avoid

15

ASEPSIS IN BIOREACTORS

stagnancy of the medium therein. It should have a positive sealing penetration capable of maintaining its integrity when subjected to repeated sterilizations and chemical exposure. The bottom discharge valve in a batch sterilization fermentor has to have a flush bottom valve with steam seal 0 rings to avoid the risk of becoming a repeated source of sepsis. A totally bottom closed bioreactor is preferable except for operational complications (K. Gerlach, unpublished communications, 1986). Headplates of small bioreactors and the manhole lid of large industrial fermentors need to be steam sealed. Entry ports, when not permanently piped to addition vessels, seed tanks, or feed manifolds, need to be sterilized initially by heat conducted from the shell of the fermentor or heated by a double valve and steam bleed arrangement. The entry ports may have needle-penetrable self-sealing diaphragms or a special connection with quick connecting valves (Bull et al., 1983). These connections have to be made sterile by standard aseptic measures (steam, flaming, and/or chemical disinfection). The ports for probes-pH, redox, and dissolved oxygen-should be slanted, with 0 rings at a point closest to the inner surface of the fermentor wall (K. Gerlach, unpublished communications, 1986; Reisman, 1988). Figure 3 shows a resterilizable side-mounted process port with a steam lock for aseptic operations. Application of hydrophobic toxic grease between two 0 rings has been recommended by Lundell and Laiho (1976).Double 0 ring seals with steam tracings between the seals have been recommended by Hambleton et al. (1991) for lid fittings and filter housings

Steam in

0

Out

FIG.3. A typical resterilizable side-mounted process port with steam lock.

16

M.C. SHARMA AND A. K . GURTU

of containment requiring bioreactors. Triple elastomer seals are used where steam tracing is not practical. The inspection glasses located at the dome of the bioreactor have to be cleaned quite often with a jet of steam to wash down broth splashings. It is important that only live steam is delivered and not the steam condensate. Steam sealing of sight and light glasses has been suggested by Lundell and Laiho (1976). The circular shape of sight glass offers improved seal characteristics as compared to other shapes (Hambleton et al., 1991). Asepsis in samples drawn from bioreactors is usually accomplished by providing a fixed sample line through a permanent penetration. Two valves in tandem on the pipe with regulated steam flow help maintain sterility between sampling operations. A sterilizable sampling device with piston valves having an 0 ring seal and flush with the inside vertical wall of the fermentor in the closed position eliminates the dead space that may cause sterility problems. Encasing the sampling pipe with a removable screw cap with a pinhole ensures that external as well as internal pipe surfaces are continually exposed to live steam between samplings. The importance of aseptic sampling systems can be judged from the variety of systems proposed for bioreactors (Heatley, 1950; Chain et al., 1954; Heden, 1958),including automated, computercontrolled sterilizable sampling systems (Ghoul et al., 1986; Seifert and Mattaeu, 1988). The air exhaust region has been the focus of attention recently because of its vulnerability to back-in infection. Protection of the exit point by an absolute membrane filter is recommended, although it has practical difficulties of membrane clogging because of “entrainment” of the medium. Mechanical foam separators like “Turbosep” [Anonymous, 1990; Hambleton et al.,1991) offer some relief by ensuring effective separation of foams, aerosols, and liquids from the air stream. Liquid drops into the bioreactor and the exit gas heated by 10-15°C passes through the exit filter. The incineration of effluent gas is a recommended step (Melling and Allner, 1981; Bull et al., 1983) in containing organisms as well. Systems of double inlet and outlet air filters permitting isolation, replacement, and sterilization without undue interference in the ongoing fermentation are now available (Hambleton et al., 1991). The positioning of an air exit at the highest elevation on the top dome, an inverted “U” configuration of the air exhaust line, and an exit valve positioned on the downward leg of the loop are sometimes adopted. This creates a sufficient distance between the fermentor and the outside nonsterile environment. It also prevents the “fall back” of collapsed foam or overflown broth from the exhaust valve into the bioreactor,

ASEPSIS IN BIOREACTORS

17

thus avoiding direct physical contact between the sterile and nonsterile environments. To discourage microbial growth in the air exhaust, the pipe immediately after the valve, as well as the air filter, if used, is kept sufficiently heated through a steam jacket. Disinfectants sometimes become trapped in the air exhaust line. The air exhaust line after the valve should be easy to dismantle for the upkeep of internal hygiene. 4. Piping and Valves In a typical fed batch fermentation process requiring the addition of four nutrient feeds, pH maintenance, and partial withdrawals, the bioreactor needs about 10 ports, 60 valves, and a considerable length of pipe connections. The design of this ancillary pipeware is crucial for asepsis. Sterile piping is usually of welded construction with internally and externally polished welds and has the minimum possible of flanged connections (Perlman, 1950; Smith, 1980; Stanbury and Whitaker, 1984; Bjurstrom, 1985; Bailey and Ollis, 1986; Van Brunt, 1987; Reisman, 1988). Threaded fittings are not acceptable in aseptic services. Sharp turns, indentations, and dead legs in lines should be avoided (Reisman, 1988). Any upturn in a sterile line should have a drain valve and steam sealing on the downstream side. The lines should also be sloped slightly for free flow and complete drainage (Stanbury and Whitaker, 1984).Of special significance is the worksmanship during erection. Misaligned pipe sections bolted under tension invariably have flanges as a perpetual source of leakage (Soderberg, 1983).Cutting stainless steel pipe with a hacksaw and making a V-notch prior to argon arc welding are necessary. Sterile sections of pipe should be separated from nonsterile sections by a barrier such as a sterile filter or a live steamheated section of pipe (Bailey and Ollis, 1986; Reisman, 1988). Gaskets and 0rings made from elastomers, e.g., EPDM, viton, silicone rubber, or high temperature-resistant Teflon, are ideal for sterile operations since they are impervious, long lasting, and easily cleanable (Bull et a]., 1983; Reisman, 1988). The selection and placement of the valves used for a sterile system need special care. Microorganisms have been known to grow through closed valves under conditions suited to them (Bjurstrom, 1985). Valves must meet cleanliness, maintenance, and sterility requirements. Diaphragm and pinch valves with Teflon sealing are ideal for aseptic operations (Stanbury and Whitaker, 1984; Bjurstorm, 1985; Threfall and Garland, 1985; Reisman, 1988). Diaphragms made out of high temperature butyl rubber are advantageous because of their long life expectancy under most operating conditions, including liquids containing solids and abrasives. Ball valves are also used while butterfly valves are rarely used. Globe and gate valves are

18

M. C. SHARMA AND A. K. GURTU

considered unsuitable because of internal crevices and the inherent lack of cleanability. Valves should be designed and positioned to permit the total drainage of materials. Most of the valves in a bioreactor system must be operated during sterilization and any mistake may lead to contamination. The microprocessor controlled operation of valves in the right sequence, activating all individual valves, is desirable for aseptic processing (K. Gerlach, unpublished communications, 1986). 5. Air System

The production of sterile air is currently based on “depth” or membrane types of filters. With depth filters, the design is very crucial (Aiba et al., 1986). Long staple glass fibers packed in pressure vessels were initially used, and prepacked cartridges (Perkowski, 1983) are now available in a wide range of sizes for this purpose. Generally, glass fibers with a diameter less than 10 p m are packed to a minimum density of 180 kg/m3. A packing density of 300 kg/m3 is achieved by high compaction under wetted conditions. Incorporation of a built-in mechanical bed compaction device in the filter should take care of the sagging packed bed without opening the lid. The filter bed should be perfectly dried out before being used as an air supply to a sterilized bioreactor. Excessive heating with a steam jacket in packed regions should be avoided to save glass fibers from fragmentation and the consequent reduction of filter efficiency. To maintain an elevated air temperature, air should be passed through a heater prior to entry into the filter in order to inactivate the microorganisms trapped in the filter bed while effectively drying the packing. However, this requires increased energy consumption for heating and the subsequent cooling of air to make it acceptable for the bioprocess. The relevance of high air temperature may be judged from the fact that for every cubic meter per hour of influent air flow, approximately 66 million microbes can be expected to challenge the filter annually, even with rather clean air (1700 organisms/m3 of air). Membrane filters have increased acceptance, and membranes made out of PTFE and polyvinyl difluoride, incorporating 0.1 pm pore size, repeated sterilizability, and high void volumes (80%), permit air flow with low pressure drops (Smith, 1981; Stanbury and Whitaker, 1984). Polyhexamethylene adipamide, polyamide, cellulose nitrate, cellulose acetate, and regenerated cellulose membranes are also used. The superiority of a membrane filter system and criteria for selecting fermentation air filters have been reviewed extensively by Leahy and Gabler (1984), Conway (1984, 1985), and Hambleton et al. (1991). The advantages described also include in situ validation of the integrity of filters by a

ASEPSIS IN BIOREACTORS

19

forward flow system. Such a validation is impossible in the case of depth filters, and hence a nagging doubt persists about the depth filter being the cause of nonsterility. Provision of moisture and oil-removing prefilters enhances the performance of membrane filters. Special attention needs to be paid to mechanical damages to the 0 rings and the membranes during cleaning/assembling. Strict adherence to the prescribed norms and limits of thermal and pressure differentials is helpful in maintaining integrity and a long operational life of sterile membrane filters. Even the steam used for their sterilization should be filtered. It is essential that each bioreactor has an individual sterile filter on its air supply line. A common storage tank for a sterile feed to many bioreactors should preferably have two membrane air filters in series as a precautionary measure. 6. Liquid Transfer System

When transferring liquids from one bioreactor to the other, sterilization of intervening piping, prior to passage of liquid, is invariably done by steam. Short flexible pipes with sanitary quick connecting ends/ valves may be used after autoclaving or in situ sterilization. The application of stericonnectors for the sampling and transfer of liquids has been emphasized by Heden (1958) and Steel and Miller (1970). Tolbart and Feder (1982) described an air-shielded quick connecting system for the sampling and transfer of liquids for aseptic processes. In fixed piping systems, steam supply and condensate trappings are a must on every section of sterile operation lines. Separate sections of the plant, with a double block protection, should be sterilizable without interfering with other ongoing operations. Lines with double/triple valves and steam bleeding are necessary for the maintenance of sterile zones at desired places (Bull et a]., 1983). Sterilizable diaphragm metering pumps are recommended for feed transfers instead of glass tube rotameters. Various devices have been suggestedhsed for the aseptic transfer of inoculum from the laboratory to the seed culture reactor as well as from one bioreactor to another. These transfers have to be made while maintaining a differential in positive pressure in the donor and recipient vessels. The inoculation port should be equipped with a steam supply. Some of these systems have been described by Parker (1950), Jackson (1958), Steel and Miller (1970), and Meyrath and Suchanek (1972). For laboratory inoculum transfer, the system could have a “pressure flask” with a side nozzle fitted with silicone tubing with a needle fixed at the distal end. The transfer of inoculum is aided by a peristaltic pump, after the sterile needle pierces the presterilized silicone/rubber diaphragm on the inoculation port of the seed bioreactor. Another possibility could

20

M. C. SHARMA AND A. K. GURTU

be to use a metallic pot, filled with inoculum in sterile room, clamped onto a vertical steam cross system on the bioreactor, and sterilized at the junction by steam with transfer effected through a pressure differential manipulated in the pot and bioreactor. Any inoculation procedure requiring total depressurization of the bioreactor is not advisable. The pooling of inoculum out of many containers at any stage should be avoided as much as possible. For continuously fed industrial bioreactors, the considerations of investment, space, energy, and process economics determine the setup of feed systems and the choice between a dedicated/common feed tank or a dedicated point of use continuous sterilizer for each feed. Because of feeds, the dedicated feed tank is ideal for containing large-scale outbreaks of nonsterility. When in use, concentrated acid and alkali solutions and anhydrous ammonia gas would normally be considered self-sterilizing. However, the air supply to the acid and alkali tanks must be filtered via a membrane filter, the feed line to the bioreactor must hold the acidlalkali for a certain amount of time prior to the addition, and the stream should remain unbroken during a fermentation run (Hambleton et al., 1991). Many of the aforementioned concepts necessary for aseptic operations of bioreactors have been incorporated in Fig. 4 . F. MAINTENANCE OF ASEPSIS DURING FERMENTATION The maintenance of asepsis is necessary throughout the complete process cycle. During the cool down from the sterilization temperature, the systems should be pressurized with sterile air to avoid pulling a vacuum and drawing in contaminating organisms. Positive pressures have to be maintained throughout all processes (Stanbury and Whitaker, 1984; Bjurstrom, 1985;Reisman, 1988).Air pressure fluctuations should be minimized to avoid the back flow of nutrients into the air filters. The availability of a standby automatic changeover, captive power generation unit, compressed air buffer reservoir, and the automatic closure of air inlet and outlet valves at a preset air flow/pressure fall, could help in avoiding the depressurization of bioreactors. The use of a nonreturn valve on the air line is also helpful. Special attention needs to be paid in eliminating thermal and pressure differential shocks to membrane filters through check valves and proper operations. Effective foam control through foam breakers (Hall et al., 1973; Viesturs et al., 1982), foam probes, and antifoam additions help in aseptic fermentations. A new breed of foam sensors for accurate foam level detection and control has been developed with corrosion-resistant construction material without ridges or crevices to prevent the growth of organisms on the probe

ASEPSIS IN BIOREACTORS 20

8

21

8

17

t:

9

11

24

FIG.4. A state of the art bioreactor system with asepsis concepts. (1)Bioreactor with SS 316 L material, electropolished internal finish, internal fittings with gaps, and total drain concept. (2) Manhole with steam seal. (3) Viewglass with steam seal. (4) Double mechanical agitator seal, steam lubricated, with level indicator. (5) Steam filter. (6) Air filter. (7) Inoculum vessel. (8) Steam inlet. (9) Steam/condensate outlet. (10)Steam trap. (11) Air from prefilter. (12) Inverted “U” loop on air supply line. (13) Inverted “U” loop on air exhaust line. (14) Steam heated jacket. (15) Exhaust air filter bypass. (16) Vessel pressure controller. (17) Air exhaust. (18) Pressure gauge. (19) Quick connecting aseptic coupling. (20) Inlets for medium, feed, and antifoam. (21) To cleanable, selfdraining sparger. (22) Flush bottom valve. (23) To harvest. (24) Drain. (25) Sampling point. (26) Filter cover for sampling point. (27) Limpet coil/jacket. (28) Exhaust cooler condenser.

(Russell and O’Hare, 1991). The choice of antifoams is a subject in itself (Solomons, 1967; Ghildyal et a]., 1988). The maintenance of aseptic transfer lines is a process needing care and attention. The sequencing of valve operations to ensure the bleeding off of the condensate and the heating up each portion to sterilization

22

M. C. SHARMA AND A. K. GURTU

temperature is necessary. The use of “Tempilistiks” (Soderberg, 1983) or thermocouples, color-changing adhesive tapes or Browne’s tubes (Threfall and Garland, 1985) is recommended to ensure proper sterilization. The use of a dedicated steam trap on every sterile process line is necessary to build up pressure. A common steam trap serving several pipe lines reduces sterilizing efficiency (Bull et a]., 1983) and results in incapacitating a number of lines, even if only a single component fails. Once the transfer from a donor vessel is finished, the pipe line must be steam sterilized prior to receiving the contents of another donor. G. PROTECTED FERMENTATION

The application of antimicrobial substances for the protection of fermentations has been in practice since 1923, particularly in manufacturing alcohol, yeast, and antifungal antibiotics (Hayduck, 1923). Herold and Necasek (1959) have written a comprehensive review on “Protected Fermentation.” Bisulfites, formic acid/formaldehyde, boric acid, picric acid, pentachlorophenol (Underkoffler and Hickey, 1954), and antibiotics like polymyxin, penicillin, and chlorotetracycline (Strandskov and Bockelmann, 1953; Day et a]., 1954; Borzani, 1956; Borzani and Aquarone, 1957; Herold and Necasek, 1959) have been used as antimicrobials in maintaining asepsis in beer/alcohol fermentations. The addition of neomycin for prophylaxis in fermentations of nonantibacterial products was suggested by Bull et aI. (1983). The present day animal cell cultures are usually protected by “antimicrobials’’ (Arathoon and Birch, 1986). Perlman (1979) reviewed the use of antibiotics in cell culture media and recommended using penicillin/ streptomycin, chloramphenicol, or tetracycline against bacterial contaminants; gentamicin or tylosin against mycoplasma; and amphotericin B against yeasts. Lambert and Birch (1985) advocated the use of penicillin, gentamicin, amphotericin B, or nystatin for contamination control in cell growth media. Despite the threat of development of antimicrobial-resistant contaminants, the protected fermentations are currently in use. As it may be unethical for surgeons to disregard aseptic techniques in surgery, it would be unprofessional for biotechnologists to cover the inefficiencies of techniques and equipments by protecting the fermentation process with antimicrobials (Herold and Necasek, 1959). However, the use of antimicrobials in lowering the cost of production, simplifying equipment and maintenance, or salvaging a contaminated batch may at times become necessary. Bull et al. (1983) consider such prophylactic measures as a last resort and regard them as poor substitutes for proper equipment maintenance and operating practices.

ASEPSIS IN BIOREACTORS

23

H. PRODUCT CHANGEOVER In the event of a “wave” or siege of contamination in the production unit, the advisable recourse, pending investigational findings, is to temporarily switch over to the production of any other fermentation product, if available, or to substitute the strain with an immunehesistant strain, especially in case of a phage spread over (Soderberg, 1983).There have been cases of stoppages of production in the case of single product plants whenever such contamination “waves” appeared.

I. SCHEDULES AND PROCEDURES Routine and preventive maintenance of vessels and systems plays a crucial role in asepsis. Checklists for equipment elements to be inspected after every harvest should be maintained and adhered to (Reisman, 1988). Separate checklists for weekly/monthly inspections also should be made. A third checklist should include a thorough internal inspection of the bioreactor during downtime for preventive maintenance (once or twice a year). A special checklist has to be made for a postcontamination check. A record of the checks made and adjustmentirectification carried out helps in correcting measures in the future. Leak tests are generally performed during preventive maintenance or if the contamination frequency is high. A fixed schedule for the hydraulic pressure testing of the vessel, jacket, and coil should be adhered to irrespective of the sterility status in the past. The use of an iodine solution with a starch indicator for detecting leaks in cooling coils may prove useful. Perkowski et al. (1984) recommend the use of a popping sound generated by a patented liquid leak amplifying chemical in conjunction with ultrasonic waves in detecting microscopic leaks in cooling coils that could not be detected even with a halogen leak detector after Freon-12 pressurization in coils. Radiographic detections have been described as very reliable in testing leaks. Biological tracers (Bacillus subtilis var. niger spores) have been recommended by Hambleton et al. (1991) for the microbiological assessment of the integrity of fermentors and their components. Nutrient feed lines should be checked for hermeticity prior to each sterilization. Control instruments validation and calibration for pressure and temperature should also be included in the maintenance schedule. The inspection of a bioreactor should cover the shell, dome, agitator seal, air exhaust line, nozzles, sight glasses, 0 rings, valves (inputs and effluents), air filters, probes, shaft, keyways, impellers, hubs, hangers, brackets, coils, sparger, ladder, thermowells, dip pipes, inoculum header, feed lines, and sampling line (Reisman, 1988).

24

M. C. SHARMA AND A. K. GURTU

The use of a high pressure water jet during regular postharvest cleaning operations should ensure a thorough cleaning and removal of undesired materials (Reisman, 1988). Vessels should be boiled with a 5% alkali solution and the feedlines and air exhaust system should be cleaned with a hot alkali solution at regular intervals. In case of severe contamination, checks and cleaning should be rigorous. Leakages, however minor, should not be overlooked. No transfer line should be in use for repeated aseptic transfers for prolonged periods without steaming in between for sterilization. The good housekeeping of fermentation areas and proper disinfection of contaminated batches, prior to discharge, helps in the maintenance of asepsis in bioreactors. Initiating and adhering to standard operating procedures, training personnel (Soderberg, 1983),rigorous checking, following proper procedures, obtaining feedback on observations, ideas, and errors, and communicating at regular/informal technical discussions are essential for good operating practices. The logging of data, deviations, and infrequent observations do forewarn and indicate the areas for corrective/remedial measures. VI. Overcautious Approaches

Driven by the fear of adverse economic consequences of nonsterility in the bioreactor, the fermentation industry often tends to adopt rather overcautious approaches. Some such practices include: 1. Sterilization of emptying vessel prior to batch sterilization 2. Incorporating chemical disinfectants like formaldehyde during

empty vessel sterilization by steam 3. Prolonged maintenance of medium at around 100°Cprior to raising to sterilization temperatures 4. In situ batch sterilization of insoluble medium ingredients coupled with external continuous sterilization of soluble components 5 . Exceeding the classical “121°C for 30 min” sterilization regarding temperature and/or time in case of equipment, piping, air filter, and thermostable feeds for the process 6. Providing a standby fiber-packed filter and switch over before the designed filter span 7. Prolonging drying of the fiber-packed air filter 8. Maintaining a high air temperature even when using a validated membrane air filter 9. Using an air sampler or bubbler on the sterile air system for detecting contaminants, despite knowing that the sample tested is a very minute part of the total air supplied to the bioreactor (Soderberg, 1983)

ASEPSIS IN BIOREACTORS

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10. Using laminar air flow benches inside sterile rooms/chambers for microbiological handlings.

It is a matter of debate whether the practice of such techniques is necessary, desirable, or avoidable. VII. Conclusion

Asepsis in a bioreactor can be achieved through integrated efforts on system design, materials and layout, effective validation and operating procedures, scheduled checks and maintenance, and trained motivated personnel. The effectiveness of these efforts depends on the stringent adherence to schedules and procedures and on the limits of sensitivity of sterility assessment methodology. Every effort made on the design, operation, and maintenance of a bioreactor system for minimizing opportunities of invasion by unwanted microbes is worth the trouble. REFERENCES Aiba, S., Humphrey, A. E., and Millis, N. F. (1986). “Biochemical Engineering.” Academic Press, New York. Anonymous (1990). “Process Filtration News,” Biopharm Edition. Domnick Hunter Filters Ltd., Durham, England. Arathoon, W. R., and Birch, J. R. (1986). Science 232, 1390-1395. Augurt, T. A. (1983). Kirk-Othmer Encycl. Chem. Technol. 3rd Ed. 21,626-644. Aunstrup, K., Andresen, O., Falch, E. A., and Nielsen, T. K. (1979). In “Microbial Technology” (H. J. Pepler and D. Perlman, eds.), 2nd ed., Vol. 1, pp. 282-309. Academic Press, New York. Bader, F. G., Boekeloo, M. K., Graham, H. E., and Cagle, J. W. (1984). Biotechnol. Bioeng. 26, 848-856. Bailey, J. E., and Ollis, P. R. (1986). “Biochemical Engineering Fundamentals.” McGrawHill International Edition, Singapore. Banks, G. T. (1979). Top. Enzyme Ferment. Biotechnol. 3, 170-266. Benarde, M. A. (1970). “Disinfection.” Dekker, New York. Bjurstrom, E. E. (1985). Chem. Eng., (N.Y.) Feb. 18, pp. 126-158. Borick, P. M. (1968). Adv. Appl. Microbiol. 10,291-312. Borzani, W.(1956). Bol. Dep. Quim. Esc. Politec., Univ. Sao Paulo 2, 1-4. and Aquarone, E. (1957). J. Agric. Food Chem 5, 612-616. Borzani, W., Bull, D. N., Thoma, R. W., and Stinner, T. E. (1983). Adv. Biotechnol. Proc. 1, 1-30. Cameron, J., and Godfrey, E. I. (1969). Biotechnol. Bioeng. 11, 957-985. Chain, E. B., Paladino, S., Ugolini, F., Callow, D. S., and Van der Sluis, J. (1954). Rend. 1st. Super. Sanita (Engl. Ed.) 17, 61-86. Charton, R. (1990). Am. Biotechnol. Lab. Dec., p. 60. Conway, R. S. (1984). Biotechnol. Bioeng. 26, 844-847. Conway, R. S. (1985). In “Comprehensive Biotechnology” (M. Moo-Young, ed.), Val. 2, pp. 279-286. Pergamon, Oxford and New York. Cooney, C. L. (1985). In “Comprehensive Biotechnology” (M. Moo-Young, ed.), Vol. 2, pp. 287-298. Pergamon, Oxford and New York.

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Day, W. H., Serjak, W. C., Stratton, J. R., and Stone, L. (1954).J. Agric. Food Chern. 2, 252-258.

Difco Laboratories (1985). “Difco Manual,” lo th ed. Difco Laboratories Inc., Detroit, MI. Elander, R. P. (1989).In “Genetic Engineering Technology in Industrial Pharmacy: Principles and Application” (J. M. Tabor, ed.). pp. 115-129. Dekker, New York. Elmroth, I., Valeur, A., Odham, G., and Larsson, L. (1990). Biotechnol. Bioeng. 35, 787-792.

Elsworth, R. (1960). Prog. Ind. Microbiol. 2, 103-130. Finch, W. I. (1958). “Disinfectants, Their Values and Uses.” Chapman & Hall, London. Ghildyal, N. P., Lonsane, 8 . K., and Karanth, N. G. (1988). Adv. Appl. Microbiol. 33, 173-222.

Ghoul, M., Ronet, E., and Engasser, 3. (1986). Biotechnol. Bioeng. 28, 119-121. Hall, M. J., Dickinson, S. D., Pritchard, R., and Evans, J. I. (1973). Prog. Ind. Microbiol. 12,169-234.

Hambleton, P., Griffiths, J. B., Cameroon, D. R., and Melling, J. (1991). J. Chern. Tech. Biotechnol. 50, 167-180. Hastings, J. J. H. (1978).In “Economic Microbiology” (A. H. Rose, ed.), Vol. 2, pp. 31-45. Academic Press, London. Hayduck, F. (1923). U.S. Pat. 1,449,112. Heatley, N. G. (1950).J. Gen. Microbiol. 4, 410-412. Heden, C. G. (1958). Nord. Med. 7, 1090. Herold, M., and Necasek, J. (1959). Adv. Appl. Microbiol. 1, 1-21. Jackson, T. (1958). In “Biochemical Engineering” (R. Steel, ed.), pp. 183-222. Heywood, London. Knight, P. (1989). Bio/Technology 7(5), 459-461. Lambert, K. J., and Birch, J. R. (1985).In “Animal Cell Biotechnology” (R. E. Spier and J. B. Griffiths, eds.), Vol. 1, pp. 85-122. Academic Press, London. Leahy, T. J., and Gabler, R. (1984). Biotechnol. Bioeng. 26, 836-843. Liberman, D.I., Fink, R., and Shalfer, F. (1986). In “Manual of Industrial Microbiology and Biotechnology” (A. L. Demain and N. A. Solomons, eds.), pp. 402-409. Am. SOC.Microbiol., Washington DC. Lubiniecki, A. S., and May, L. H. (1985). Dev. Biol. Stand. 60, 141-146. Lundell, R., and Laiho, P. (1976). Process Biochem. 11, 13-17. Maurer, H. R. (1986). In “Animal Cell Culture: A Practical Approach” (R. I. Freshner, ed.), pp. 13-31. I.R.L. Press, Oxford and Washington, DC. McLaughin, J., Bruno, C. F., and Forrest, T. (1983). Biotechnol. Bioeng. 25, 1229-1236. Melling, J . , and Allner, V. (1981). In “Essays in Applied Microbiology” ( J . R. Norris and M. H. Richmond, eds.), Chapter 11, p. 1. Wiley, New York. Meyrath, J. and Suchanek, G. (1972). Methods Microbiol. 7 , 159-209. Parker, A. (1950).“Recent Advances in Fermentation Industry.” Royal Institute of Chemistry, London. Perkowski, C. A. (1983). Biotechnol. Bioeng. 25, 1215-1222. Perkowski, C. A., Daransky, G. R., and Williams, J. (1984). Biotechnol. Bioeng. 26, 857-859.

Perlman, D. (1950). Bot. Rev. 16, 449-523. Perlman, D. (1979). In “Methods in Enzymology” (W. B. Jakoby and I. H. Pastan, eds.), Vol. 58, pp. 110-116. Academic Press, New York. Reisrnan, H. B. (1988). “Economic Analysis of Fermentation Processes.” CRC Press, Boca Raton, FL. Rhodes, A., and Fletcher, D. L. (3966).“Principles of Industrial Microbiology.” Pergamon, Oxford and New York.

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Richards, J. W. (1966). Process Biochem. 1(1), 41-46. Richards, J. W. (1968).“Introduction to Industrial Sterilization.” Academic Press, London. Russell, M., and O’Hare, D. (1991). Am. Biotechnol. Lab. July, p. 30. Saudek, E. C. (1956). Bacteriof Rev. 20, 279-281. Scheirer, W. (1987). Trends Biotechnol. 5(9), 261-265. Seifert, G. K. E., and Mattaeu, P. (1988). Biotechnof. Bioeng. 32, 923-926. Sittig, W. (1982).J. Chem. Tech. Biotechnol. 32, 50. Smith, S. R. L. (1980). Philos./Trans. A. SOC.London, Ser. B 290, 341-354. Smith, S. R. L. (1981). In “Microbial Growth in C1 Compounds” (H. Dalton, ed.), pp. 342-348. Heyden, London. Soderberg, A. C. (1983). In “Fermentation and Biochemical Engineering Hand Book” (H. C. Vogel, ed.), pp. 111-117. Noyes, Data Corp., Park Ridge, NJ. Solomons, G. L. (1967). Process Biochem. 2, 47-48. Solomons, G. L. (1969). “Materials and Methods in Fermentation.” Academic Press, New York. Solomons, G. L. (1971). Adv. Appl. Microbiol. 14, 231-248. Spier, R. (1988). Trends Biotechnol. 6, 2-6. Stanbury, P. F., and Whitaker, A. (1984).“Principles of Fermentation Technology.” Pergamon, New York. Steel, R., and Miller, T. L. (1970). Adv. Appl. Microbiol. 12, 153-188. Strandskov, F. B., and Bockelmann, J. B. (1953). J, Agric./Food Chem. 1, 1219-1223. Stumbo, R. (1976). In “Industrial Microbiology” (B. M. Miller and W. Litsky, eds.), pp. 412-450. McGraw-Hill, New York. Sykes, G. (1958). “Disinfection and Sterilization.” Spon, London. Threfall, G., and Garland, S. G. (1985). In “Animal Cell Biotechnology” (R. E. Spier and J. B. Griffiths, eds.), Vol. 1, pp. 123-140. Academic Press, London. Tolbart, W. R., and Feder, J. (1982). Biotechnol. Bioeng. 24, 1885-1887. Underkoffler L. A., and Hickey, R. J. (1954). “Industrial Fermentation,” Vols. 1 and 2. Chemical Publications, New York. U.S. Department of Health and Human Services (1984). “Biosafety in Microbiological and Biomedical Laboratories.” HHS Publication, Washington, DC. U.S. Pharmacopoeia (1980). XX Revision, NF XV. USP Convention Inc., Rockville, MD. Van Brunt, J. (1987). Bio/Technology 5(11), 1133-1138. Viesturs, U. E., Kristapsons, M. Z., and Levitans, E. S. (1982). Adv. Biochem. Eng. 21, 169-224.

Wallhauser, K. H. (1985). In “Biotechnology” (H. J. Rehm and G. Reed, eds.), Vol. 2, pp. 699-724. Verlag Chemie, Weinheim. Wilkinson, G. R., and Baker, C. L. (1964). Prog. Ind. Microbiol. 5, 237-283.

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Lipids of n-Al kane-Utilizing Microorganisms and Their Application Potential

s.RADWAN’” AND NASERA. SORKHOHt * Institut fur Mikrobiologie, Universitat Munster,

SAMIR

D-4400Munster, Germany f

Department of Botany and Microbiology, Faculty of Science, University of Kuwait, safat 13060, Kuwait

I. Introduction 11. n-Alkane-Utilizing Microorganisms

111. Total Lipid Contents A. Lipids and Fats B. Basic Studies C. Biotechnological Considerations IV. Fatty Acids A. Basic Studies B. Biotechnological Considerations V. Acylglycerols VI. Sterols VII. Fatty Alcohols, Ketones, and Epoxides VIII. Hydrocarbons and Waxes IX. Phospholipids X. Glycolipids and Peptidolipids XI. Biolipid Extract XII. Environmental Considerations References

I. Introduction

Until the beginning of the 1970s the results of many studies on hydrocarbon-utilizing microorganisms were covered by patents. Such studies comprised the isolation of microorganisms, oxidation and hydroxylation of hydrocarbons, biomass production, biosynthesis of proteins, carbohydrates, “fats,” vitamins, enzymes, nucleotides, antibiotics, and organic acids, as well as recovery procedures for these products. During the past two decades the number of original studies published in scientific journals continuously increased, and it now exceeds by far

’

Present address: Department of Botany and Microbiology, Faculty of Science, University of Kuwait, Safat 13060,Kuwait. 29 ADVANCES IN APPLIED MICROBIOLOGY. VOLUME 39 Copyright 0 1993 by Academic Press, Inc. All rights of reproduction in any form reserved.

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SAMIR S. RADWAN AND NASER A. SORKHOH

the number of patents. As far as the history of studies on lipids of hydrocarbon-utilizing microorganisms is concerned, a parallel development occurred. In the beginning of the 1970s there were numerous patents covering results on “fat” production by oil-utilizing yeasts. Within the past 25 years much original research work has been published on the lipids of microorganisms that utilize hydrocarbons, especially n-alkanes, as substrates; a review of these studies appears to be timely. Earlier reviews on the microbial degradation of hydrocarbons (Ratledge, 1978, 1980; Rehm and Reiff, 1981; Fukui and Tanaka, 1981; Biihler and Schindler, 1984) and on lipids of oleaginous microorganisms (Boulton and Ratledge, 1984; Ratledge, 1986) devoted parts of their discussion to lipids and/or fatty acids of n-alkane-utilizing microorganisms. This article presents a comprehensive review of this subject, referring to the potential commercial values of various lipid classes and fatty acids. In addition, reference is made to environmental considerations associated with the proposed application of microorganisms in controlling oil pollution and in enhanced oil recovery. During many of such processes microorganisms liberate surfactive lipids, whose impact on the environment is not known, so far. That lipids of n-alkaline-utilizing microorganisms should be expected to differ from lipids of the same organisms grown on conventional substrates is apparent from the following arguments. 1. n-Alkanes, being water insoluble, expectedly induce in cell membranes alterations that allow for their enhanced active transport. Such alterations may involve the membrane lipids which contribute to about 50% of the membrane weight. 2. Alkanes, themselves lipids, are taken up, chemically unchanged, and thus directly contribute to the total cell lipids. 3. Initial phases of n-alkane metabolism involve oxidation of these substrates to fatty alcohols and fatty acids which become, in part, incorporated into complex cell lipid compounds. 4. Several n-alkane-utilizing microorganisms reveal cytological entities that are associated with their growth on n-alkanes as substrates. Thus, certain bacteria produce intracytoplasmic membranes, and yeasts produce peroxisomes. Like other biological membranes and organelles, these cytological entities are expected to be rich in lipids. I I. n-Alkane-Uti lizi ng Microorganisms

This subject has been repeatedly reviewed (Klug and Markovetz, 1971; Levi et a ] . , 1979; Einsele, 1983) along with the physiology of

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n-alkane-utilizing microorganisms and alkane metabolism (Rehm and Reiff, 1981; Fukui and Tanaka, 1981; Boulton and Ratledge, 1984). The most interesting microbial genera capable of utilizing n-alkanes as sole sources of carbon and energy are listed in Table I. This list, which may grow in the future, includes microorganisms reported in earlier reviews together with additional members reported in original research publications. It is apparent that the utilization of n-alkanes as substrates is a widely distributed activity among microorganisms. This activity is achieved by both prokaryotes and eukaryotes, including mainly organotrophs and a few photoautotrophs. Although Table I comprises a relatively large number of genera, most studies in the literature have been done on TABLE I MICROBIAL GENERACONTAINING HYDROCARBON-UTILIZING SPECIES OR STRAINS' Prokaryotes Photoautotrophic: Rhodospirillum, Rhodopseudomonas, Oscillatoria [Cerniglia et al. (1980a)l Organotrophic cocci: Acinetobacter, Micrococcus, Sarcina Curved rods: Vibrio, Azospirillum [Roy et al. (1988)] Gram-negative rods: Aeromonas, Alcaligenes, Chromobacterium, Flavobacterium, Klebsiella, Pseudomonas [Klug and Markovetz (1971)l Gram-positive rods: Bacillus [Loginova et al. (1981)], Bacillus stearothermophilus [Sorkhoh et al. (1993)j Actinomycetes and related organisms: Arthrobacter, Brevibacterium, Corynebacterium, Rhodococcus [Egorov et 01. (1986)], Mycobacterium, Actinomyces, Nocardia, Streptomyces Eukaryotes Photoautotrophic: Chlorella [Schroeder and Rehm (198l)], Scenedesmus [Schroeder and Rehm (1981)l Organotrophic: Yeasts: Candida, Debaryomyces, Endomyces, Leucosporidium, Lodderornyces, Metschnikowia, Pichia, Rhodosporidium, Rhodotorula, Saccharomycopsis, Schwannio-myces, Selenotila, Sporidiobalus, Sporobolomyces, Torulopsis, Trichosporon, Wingea Filamentous fungi: Absidia [Hoffmann and Rehm (1978)], Aspergillus, Aureobasidium, Beauveria [Davies and Westlake (1979)], Botrytis, Cephalosporium, Cladosporium, Corellospora [Kirk and Gordon (1988)], Cunninghamella, Dendyphiella [Kirk and Gordon (1988)], Fusarium, Hormodendrum [Lin et al. (1971a,b)] Lulworthia [Kirk and Gordon (1988)], Mortierella, Mucor, Penicillium, Phialophora, Phoma [Davies and Westlake (1979)], Scedosporium [Ornodera et al. (1989)], Scoleobasidium [Davies and Westlake (1979)], Sporotrichum. Varicosporino [Kirk and Gordon (1988)], Verticillium a

Unless otherwise specified, the information is according to Levi et al. (1979)

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SAMIR S. RADWAN AND NASER A. SORKHOH

much fewer ones. Thus the prokaryotic genera most frequently studied include Acinetobacter, Pseudomonas, and the actinomycetes Arthrobacter, Corynebacterium, Rhodococcus, Mycobacterium, and Nocardia. In this context, a problem related to the nomenclature should be mentioned. Thus, for example, Acinetobacter used to be called Micrococcus cerificans in earlier studies (see Makula and Finnerty, 1970). Similarly, many of the Nocardia species in the Eighth Edition of “Bergey’s Manual of Determinative Bacteriology” are classified in the more recent first edition of “Bergey’s Manual of Systematic Bacteriology,” as Rhodococcus species (see Nakajima and Sato, 1983). Koronelli (1988) used the term “saprophytic mycobacteria” for Rhodococcus, Corynebacterium, and related genera. In view of the striking morphological similarity among such actinomycetes (Sorkhoh et al., 1990a),misleading identities of these organisms should be expected, especially in very early publications. In the genus Bacillus, only B. stearothermophilus has been reported to utilize n-alkanes (Loginova et al., 1981); other reports (Kachholz and Rehm, 1978) were not confirmatory. It appears, however, that the ability to utilize n-alkanes as sole sources of carbon and energy is lacking among mesophilic Bacillus species (Kvasnikov et al., 1973; Kachholz and Rehm, 1977). The yeast genus most frequently studied is Candida; fewer studies have been done using Lodderomyces, Rhodotorula, and Torulopsis. Among the filamentous fungi, Aspergillus, Cladosporium, Cunninghamella, Fusarium, and Penicillium received most of the researcher interest. In this context, it is noted that Lindley and Heydeman (1985) emphasized the importance of an extended lag phase when assessing substrate optima for alkane utilization by filamentous fungi. Reportedly, failure to take the progressively longer lag phase (in respect to carbonchain length) into consideration may have led workers, who used single point biomass measurements as an indicator of growth, to underestimate the potential of fungi to grow on alkanes such as octadecane. Of course, none of the microorganisms are capable of utilizing all nalkanes; each organism can utilize only a certain range of compounds. But collectively, all compounds from the gaseous low molecular weight (van Ginkel et al., 1987; Ornodera et al., 1989) up to the medium and high molecular weight constituents (Demanova et al., 1980b) can be attacked by microorganisms. This activity is maintained during immobilization of both unicellular (El-Aassar et al., 1988) and filamentous (Heinrich and Rehm, 1981) microorganisms. Among the interesting alkane-utilizing microorganisms are the thermophiles Thermus ruber (Loginova et al., 1981), Thermoleophilum album (Zarilla and Perry, 1984) and Bacillus stearothermophilus (Sorkhoh et al., 1993), and the nitrogen-fixing Azospirillum sp. (Roy et al., 1988).

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Mixtures of n-alkane are much more efficiently oxidized via the activities of microbial associations than in pure cultures. This has been demonstrated experimentally using an association of Pseudomonas and Mycobacterium (Koronelli et al., 1984). Particularly interesting is the ability of the cyanobacterium Oscillatoria sp. and the green algae Chlorella vulgaris and Scenedesmus obliquus to utilize n-alkanes as sole sources of carbon and energy (Cerniglia et al., 1980a,b; Schroeder and Rehm, 1981; Zawdzki and Langowska, 1983). Oscillatoria sp., diatoms, as well as green, red, and brown algae can also oxidize naphthalene to at least six metabolites including 1-naphthol. The degradation of hydrocarbons by associations of cyanobacteria and organotrophic bacteria has been documented (Gusev et al., 1981, 1982; Sorkhoh et al., 1992).

Microorganisms may exhibit, during n-alkane utilization, characteristic morphological and cytological features (see Table 11). One of these features is hydrocarbon inclusions. The production of intracytoplasmic inclusions by oil-utilizing microorganisms has been documented by early investigators working on bacteria (Scott and Finnerty, 1966; Atlas and Heintz, 1973; Kennedy and Finnerty, 1975a) and filamentous fungi (Cundell et al., 1976; Koval and Redchitz, 1978: Redchitz, 1980). These inclusions may occupy up to 40% of the total cell volume (Griffin and Traxier, 1983). On conventional carbon sources these inclusions are absent in most cases or, in a few cases, have only a very minute size. Another characteristic is intraplasmic membranes. Such membrane systems have been observed in alkane-utilizing bacteria, viz. Acinetobacter sp. (Kennedy and Finnerty, 1975b) and Rhodococcus rhodochrous (Ivshina et al., 1982).Volutin inclusions have been observed in the hyphae of Aspergillus spp. (Redchitz and Koval, 1979) grown in the presence of hydrocarbons and Rhodococcus rhodochrous incubated in an atmosphere of propane (Ivshina et al., 1982). In addition, Penicillium sp. grows in shaken cultures in media containing n-hexadecane as hollow mycelial balls enclosing hydrocarbon droplets, whereas the mycelial balls were solid in media containing peptone as substrate (Cundell et al., 1976). Alkane-grown cells, but not glucose-grown cells of Candida tropicalis possess a mannan-fatty acid complex on their surfaces (Kappeli et al., 1978). Also, alkane-grown cells show a radial arrangement of the wall polymers, with protruding parts, in contrast to the smooth surfaces of glucose-grown cells. The authors believe that the mannan-fatty acid complex may be involved in the cell binding to alkanes. Omar and Rehm (1980) noticed that n-tetradecane- and npentadecane-grown cells of Candida parapsilosis produce much more pseudomycelia and show much higher catalase activity than glucosegrown cells. According to Fukui and Tanaka (1981) the conspicuous

SAMIR S. RADWAN AND NASER A. SORKHOH

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TABLE I1 CYTOLOGICAL CHANGES INDUCEDDURING MICROBIAL GROWTHON HYDROCARBONS Microorganism Acinetobacter sp.

Acinetobacter calcoaceticus Arthrobacter sp.

Substrate Paraffinic hydrocarbons Hexadecane or hexadecene Hexadecane Hexadecane

Flavobacteriurn, Brevi bacterium

Crude oil

Rhodococcus rhodochrous

Propane

Candida lipolytica

Hexadecane

Candida tropicalis

n-Alkanes

Aspergillus oryzae, Aliphatic A. effusis, A. hydrocarbons ochraceus, A. sydowi, A. niger Cladosporium Hydrocarbons resin ae

Penicillium sp.

n-Hexane

Cytological changes

Reference

Intracytoplasmic hydrocarbon inclusions Intracytoplasmic membrane Thin fimbriae

Kennedy and Finnerty (1975a); Scott and Finnerty (1966) Kennedy and Finnerty (197 5b) Rosenberget al. (1982)

Intracytoplasmic hydrocarbon inclusions Intracytoplasmic hydrocarbon inclusions Volutin, hydrocarbon inclusions, intracellular membrane system Complex membrane and vesicles on the outer cell surface, peroxisomes Protruding parts on the outer cell surface, peroxisomes Fatty inclusions, volutin inclusions

Griffin and Traxier (1983)

Thinner walls, vacuoles, microbodies, increased catalase Grows as hollow balls enclosing hydrocarbon, hydrocarbon inclusions

Atlas and Heintz (1973) Ivshina et al. (1982)

Gutevskaya and Shishkanova (1982)

Kappeli et al. (1978); Fukui and Tanaka (1981) Koval and Redchitz (1978); Redchitz and Koval (1979); Redchitz (1980) Smucker and Cooney (1981)

Cundell et aI. (1976)

appearance of peroxisomes in the cells is one of the specific features of alkane-utilizing yeasts. Smucker and Cooney (1981),investigating the cytological changes in Cladosporium resinae, shifted from a glucose to a hydrocarbon medium and found that the cell walls become thinner (in hyphae and spores), large vacuoles appear, enrichment with micro-

n-ALKANE-UTILIZING MICROORGANISMS

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bodies occurs and catalase activity increases. Rosenberg et al. (1982) presented evidence that special thin fimbriae on the cell surfaces of Acinetobacter calcoaceticus are the agents that mediate the adherence of this bacterium to hydrocarbon droplets. Reportedly, only strains capable of adhering to hydrocarbon droplets, and consequently of utilizing these compounds, possess such thin fimbriae. Neufeld et al. (1983) observed that cells of Acinetobacter sp. lose their structural integrity when cultivated on hydrocarbon substrates, probably because of extraction of lipophilic surface components by the hydrocarbon. The plasma membrane of hydrocarbon-grown Candida lipolytica becomes thicker and contains deep projections (Ludvik et al., 1968). Ill. Total Lipid Contents

A. LIPIDSAND FATS Sometimes, especially in early publications, the terms “lipids” and “fats” have been used synonymously. It is, however, well known that the term “fats” is the common name of the chemical lipid class of triacylglycerols. In contrast, “lipids” is a collective term which comprises several chemical classes that possess the common property of being soluble in lipophilic solvents. The most common lipid classes in biological materials, including microorganisms, are triacylglycerols, sterols and steryl derivatives, phospholipids, and glycolipids. Triacylglycerols are storage products in the cell, whereas the other classes are constituents of cell membranes and organelles, where they have both structural and physiological functions.

B. BASICSTUDIES Although research on lipids of alkane-utilizing microorganisms dates back to the 1950s and 1960s, most of our current information about this subject has been published in the past two decades. However, many of these publications unfortunately were not concerned with a “classical” analysis of lipids, but were devoted primarily to the fatty acid composition of the total lipids. This is understandable in view of the early recognized fact that n-alkanes are oxidized to fatty acids during their assimilation. Nevertheless, this fatty acid-oriented interest was at the expense of the interest in elementary information about the total lipid contents and lipid composition, which were not as extensively investigated. Table 111 presents the total lipid contents of some alkane-utilizing microorganisms. Whenever available, the lipid contents of the same

36

SAMIR S. RADWAN AND NASER A. SORKHOH TABLE 111 TOTALLIPIDCONTENTS OF ALKANE-UTILIZING MICROORGANISMS" Microorganism

Substrate

n-Hexadecane Ahodococcus rubropertinctus Micrococcus n-Alkanes freu denreich i i Mycobacterium Acetate convolutum M. convolutum M. convolutum M. convolutum M. convolutum M. convolutum M. convolutum M. convolutum C28 Nocardia sp. n-Alkanes Pseudomonas Glucose aeruginosa P. aeruginosa "-Cm Candida 107 n-Alkanes Candida tropicalis Glucose C. tropicalis n-C16 C. tropicalis n-Alkanes C. tropicalis n-Alkanes C. tropicalis n-Alkanes Candida lipolytica Glucose C. lipolytica n-Alkanes C. lipolytica n-Alkanes C. lipolytica n-Alkanes C. lipolytica n-Alkanes Candida rugosa Glucose C. rugosa n-Alkanes Candida Glucose parapsilosis C. parapsilosis CI, Candida maltosa n-Alkanes Mycotorula japonica n-Alkanes Pichia vonriji n-Alkanes Rhodotorula glutinis n-Alkanes Rhodotorula gracilis n-Alkanes Absidia spinosa Glucose A. spinosa c 1 2 A. spinosa c13 A. spinosa n-Paraffin

Total lipids 14.0-22.0

Reference Egorov et al. (1986)

5 .O-12.4

Kvasnikov et al. (1977a)

0.19

Hallas and Vestal (1978)

0.55 0.55 0.49 0.60 0.53

Hallas and Vestal (1978) Hallas and Vestal (1978) Hallas and Vestal (1978) Hallas and Vestal (1978) Hallas and Vestal (1978) Hallas and Vestal (1978) Hallas and Vestal (1978) Raymond and Davies (1960) Koronelli et al. (1982a)

0.49

0.56 56.0 6.7 7.0 26.0 2.8-5.0 6.5-12.5 17.0 6.0 10.0

6.0-8.5 12.0-17.0 27.0 17.0 47.0 5.5-6.5 16.0-19.0 9.4 12.3 11.0-16.5 15.0

Koronelli et al. (1982a) Thorpe and Ratledge (1972) Hug et al. (1974) Hug et 01. (1974) Mishina et al. (1977) Thorpe and Ratledge (1972) Hug and Feichter (1973) Mishina et al. (1977) Mishina et al. (1977) Pelechova et al. (1971) Nyns et al. (1968) Jwanny (1975) Iida et 01. (1980) Iida et al. (1980) Omar and Rehm (1980)

35.0 32.0 25.8 15.6 14.9 48.6

Omar and Rehm (1980) Blasig et al. (1989) Yamaguchi and Kurosawa (1976) Pelochova et al. (1971) Pelochova et al. (1971) Pelochova et al. (1971) Hoffmann and Rehm (1978) Hoffmann and Rehm (1978) Hoffmann and Rehm (1978) Hoffmann and Rehm (1978)

38.4

Hoffmann and Rehm (1978)

20.0

(c13-c17)

Cunninghamella echinulata

Glucose

n-ALKANE-UTILIZING MICROORGANISMS

37

TABLE 111 (Continued) Microorganism C. echinulata C. echinulata C. echinulata

Substrate

Total lipids

Reference

29.9 49.4 52.5

Hoffmann and Rehm (1978) Hoffmann and Rehm (1978) Hoffmann and Rehm (1978)

27.2

Hoffmann and Rehm (1978)

c 1 2

17.1

Cl, n-Paraffin

10.9 22.7

Hoffmann and Rehm (1978) Hoffmann and Rehm (1978) Hoffmann and Rehm (1978)

c 1 2

CI, n-Paraffin (c13-c17)

Mortierella isa bellina M. isabellina /M. isabellina M. isabellina

Glucose

(C13-C17)

Data are expressed in percentage of dry biomass except for Mycobacterium convolutum whose data are given in mg lipidlmg protein.

organisms growing on conventional substrates such as glucose, acetate, or peptone are also given, for the purpose of comparison. Alkane-utilizing bacteria have a lower lipid content than yeasts and filamentous fungi. Oleaginous microorganisms are much more frequent among eukaryotes than prokaryotes. As a rule, the growth on alkanes as sole sources of carbon and energy is associated with increased lipid content of the microorganisms, as compared with the values recorded when conventional carbon sources are used. This fact was realized by investigators in the 1960s (Johnson, 1964; Mizuno et al., 1966; Dunlap and Perry, 1967; Nyns et al., 1968; Koronelli, 1968). Microorganisms grown on alkanes and on conventional carbon sources have similar lipid compositions, with only quantitative differences (see Table IV). However, sometimes there may be qualitative differences. Dodecanegrown cells, but not glucose-grown cells of Rhodococcus rhodochrous contain sterols, diacylglycerophosphocholines, and an unidentified glycolipid (Sorkhoh et a]., 1990b). The lipid classes responsible for increased total lipid content during the shift to alkane utilization are listed in Table V. Makula and Finnerty (1970), through phosphorus analysis of the total lipids, concluded that Acinetobacter HO1-N grown on hexadecane contain 50% more phospholipids than when the cells are grown on glucose. Also, Mycobacterium vacca grown on propane contain 66% more phospholipids than on acetate (Vestal and Perry, 1971). Kvasnikov et al. (1974, 1977b] found that lipid synthesis by Micrococcus freudenreichii grown on n-alkanes is much more active than on glucose; it was twice as active during slow growth and six times more active during rapid growth. Reportedly, free

38

SAMIR S. RADWAN AND NASER A. SORKHOH TABLE IV LIPIDCOMPOSITION OF Acinetobacter

SP.

HO1-N GROWNON n-HEXADECANE'

Intracellular lipids (pmol/g dry cells) Nutrient broth- yeast extract

Lipid classes Phospholipids Triacylglycerols Monoacylglycerols diacylgl ycerols Free fatty acids Free fatty alcohols Wax esters

+

Extracellular lipids (Fmol/liter)

Hexadecane

46.0 1.8

0.4

Nutrient broth-yeast extract

Hexadecane

129.0 2.5 6.8

0.0

0.0

2.4

25.6 410.0

8.2

4.0 0.0 0.0

7.5 Trace 11.5

2.6

18.0

0.0

60.0 0.5

280.0

"Results from Makula et al. (1975)

TABLE V LIPIDCLASSESREPORTEDTO ACCUMULATE IN MICROORGANISMS SHIFTED TO n-ALKANE UTILIZATION Microorganisms Acinetobacter sp.

Lipid classes

Phospholipids, mono- and diacylglycerols A. lwoffi Phospholipids Mycobacterium vacca Phospholipids Micrococcus Phospholipids, free fatty acids, waxes freudenreichii Arthrobacter ceroformans Wax esters Phospholipids Mycobac terium convolutum Sterols, Rhodococcus monoacylglycerols, rhodochrous unknown glycolipids Candida tropicalis Phospholipids C. tropicalis Phospholipids, fatty acids C. Iipolytica Phospholipids C. guilliermondii Triacylglycerols, wax C. rugosa Ergosterol C. albicans Total sterols Cladosporium resinae Hydrocarbons Paecilomyces persicinus Triacylglycerols, free fatty acids

Reference Makula and Finnerty (1970) Makula et al. (1975) Vachon et al. (1982) Vestal and Perry (1971) Kvasnikov et al. (1974, 1977b) Koronelli et al. (1978) Hallas and Vestal (1978) Sorkhoh et al. (1990b)

Mishina et al. (1977) Kvasnikov et al. (1977b) Mishina et al. (1977) Demanova et al. (1980a) Iida et al. (1980) Sorkhoh et al. (1991) Walker and Cooney (1973) Boyer and Pisano (1974)

39

n-ALKANE-UTILIZING MICROORGANISMS

fatty acids make up about 60% and phospholipids about 20% of the total lipids from cells grown on glucose. On alkanes, the phospholipid content increased to 46% and the cells accumulated wax esters (20-47% of the total lipids). The total lipid content of Mycobacterium convolutum grown on n-alkanes is two to five times higher than on acetate (Hallas and Vestal, 1978). The newly synthesized lipids have their origin in the alkane substrate as shown by results of [14C]acetate incorporation into cellular lipids and proteins (Table VI). Less than 10% of the labeled acetate is incorporated into lipids of n-alkane-grown cells compared to acetate-grown cells, whereas incorporation into proteins is not lower. This result indicates that in n-alkane-grown cells the lipids should have been synthesized mainly from the alkane substrates. In contrast, a few authors failed to find any substantial difference in the total lipid content between alkane-grown cells and cells utilizing conventional carbon sources. Koronelli et al. (1982a), working with Pseudomonas aeruginosa, analyzed “free” and “bound” lipids and found that hexane- and glucose-grown cells contain 7.0 and 6.7% free lipids, respectively. The values for bound lipids are 6.7 and 5.6%, respectively, of the dry biomass. There are comparatively more reports on total lipids of yeasts than of bacteria and filamentous fungi. This is apparently because only yeasts have been successfully used for biomass production as fodder, with hydrocarbons as substrates. In the majority of these studies Candida yeasts were used. Shigyo and Takeuchi (1972) developed a technique for the complete extraction and separation of lipids from hydrocarbongrown yeast. Hug et al. (1974) observed that lipid synthesis by Candida TABLE VI INCORPORATION OF [14C]ACETATEINTO TOTALCELL LIPIDSAND PROTEINS OF Mycobacterium convolutuma Substrate

Lipid incorporation (pmol acetatdpg lipid)

Acetate Nonadecane (C1J Eicosane (Czo) Decosane (C,) Tricosane (C,,) Tetracosane (Cz4) Hexacosane (Cz6) Octacosane (C2J From Hallas and Vestal (19781.

Protein incorporation (pmol acetatdpg protein)

315

86

24 10

15

84 80 80

11

97

29

76

18

65 82

15

40

SAMIR S. RADWAN AND NASER A. SORKHOH

tropicalis is activated by the presence of the hydrocarbon, and that little substrate uptake is possible until the lipid concentration in the cells is sufficiently high. These authors confirmed an earlier viewpoint (Dunlap and Perry, 1967; Vestal and Perry, 1971) that high lipid content is necessary for hydrocarbon uptake, and is not merely a reflection of the lipophilic nature of the substrate. Mishina et a]. (1977) reported that the total lipid content of Candida tropicalis almost doubled when shifted from glucose to alkane utilization. The chain length of the nalkane substrate did not exert any obvious effect on the total lipid content. However, these authors found in the same study that the lipid content of another species, C. lipolytica, comprised about 4% of the dry biomass, irrespective of whether the substrate was glucose or an nalkane. Kvasnikov et al. (1977b) reported that the activated lipid synthesis by C. tropicalis on hydrocarbons occurs primarily at the expense of the substrate. Davidova et al. (1978), working on C. tropicalis and using I4C substrates, compared the kinetics of label incorporation from noctadecane and glucose into the main groups of organic substances, viz. proteins, nucleic acids, polysaccharides, lipids, free amino acids, organic acids, free carbohydrates, and nucleotides. When the cells utilize n-octadecane, the radioactivity in the lipid fraction is higher than when they are grown on glucose. A few investigators working on yeasts failed to observe any striking effect of the alkane substrate on the total lipid content. Reference has already been made to C. lipolytica, which had a similar lipid content when grown on glucose or on n-alkane. Only small quantitative differences in the total lipid content of C. tropicalis were observed when the cells were grown on n-alkanes, glucose, or glycerol (Giewicz et al., 1983). Moreover, there is some contradiction in the literature regarding the effect of the alkane-chain length on the total lipid content. While Mishina et al. (1977) did not find any effect of the alkane-chain length on the total lipids of C. tropicalis, a result which was later confirmed on Mycobacterium convolutum (Hallas and Vestal, 1978), Demanova et al. (1980a) found that the lipid content of C. guilliermondii grown on n-octadecane (C18)is three times higher than that of cells grown on . et al. (1980) showed that the lipid content of C. docosane ( C Z 2 )Iida rugosa significantly increases from 16 to 19% of the dry biomass when the n-alkane chain length increases from C,, to Cz0. The effect of the alkane substrate concentration also has been investigated. Zalashko et al. (1983) reported that the optimum concentration of n-hexadecane for growth and lipid production by C. tropicalis is 1-2%; higher concentrations are inhibitory. The optimum paraffin concentration for growth and lipid synthesis by C. maltosa is 1.5-2.5% (Maksimova eta]., 1988).

n-ALKANE-UTILIZING MICROORGANISMS

41

Andreevskaya and Zalashko (1984) studied the effect of temperature on biomass and lipid production by C. tropicalis grown on C, to C,, nalkanes. The highest lipid content was obtained at 10°C, although this temperature was not optimal for growth. A few studies on total lipid contents and lipid composition of nalkane-utilizing filamentous fungi have been published. Walker and Cooney (1973) showed that the total lipid content of Cladosporium resinae grown on n-alkanes as substrates is higher than that on glucose or glutamic acid. Similarly, Paecilomyces persicinus grown on nhexadecane contains more total lipids than when glucose is utilized as substrate (Boyer and Pisano, 1974). The increase was particularly noticeable in the triacylglycerol and free fatty acid fractions, which were, respectively, three and five times higher in alkane- than in glucosegrown mycelia. Three oleaginous fungi belonging to the mucorales behaved differently on n-paraffin with C,,-C,, alkanes (Hoffmann and Rehm, 1978). Absidia spinosa and Cunninghamella echinulata grown on n-paraffin contain more total lipids than on glucose, whereas Mortierella isabellina contain less total lipids. The lipid content of the three fungi grown on n-dodecane is obviously lower than on glucose. C. BIOTECHNOLOGICAL CONSIDERATIONS

Although there are numerous patents covering the production of useful compounds by n-alkane-utilizing microorganisms, there are so far no industrial fermentation processes based on hydrocarbon substrates. According to Shenman (1984), even gas oil and n-paraffin single cell protein projects are changing to methanol or carbohydrate feedstocks. The prediction, made two decades ago, that some of the world's supplies of oil and fats could be produced microbiologically using n-alkanes as a starting material (Ratledge, 1970) remains too optimistic. n-Alkane-utilizing microorganisms appear to be the least suitable as fat sources, particularly for food and feed purposes, for two main reasons: (1)Fats, i.e., triacylglycerols, as demonstrated in this article, frequently are not a major lipid class in such microorganisms. The increased total lipids during shift to n-alkane utilization is primarily due to increased biosynthesis of phospholipids, fatty acids, monoacylglycerols, and wax esters, but rarely of triacylglycerols (see Table V). (2) Even if it is possible to enrich such microorganisms with triacylglycerols, for example, by nitrogen and other element starvation (for review, see Ratledge, 1986) and/or by genetic manipulation, a problem regarding the suitability of the fat product to be returned into the food chain remains unsolved. Because of the relatively high content of odd-chain

42

SAMIR S. RADWAN AND NASER A. SORKHOH

fatty acids (see Section XI) associated with microbial growth on nalkane mixtures, the fat product cannot be recommended as a nutrient. However, the chance for lipids from n-alkane-utilizing microorganisms to find application in the diverse field of oleochemical industry (for reviews, see Richter and Knaut, 1984; Leonard and Kopald, 1984) is rather good. Some aspects of these applications are discussed in the following sections dealing with individual lipid classes of n-alkaneutilizing microorganisms. IV. Fatty Acids

Fatty acids occur in all biological systems, mainly in complex lipids such as triacylglycerols, wax esters, steryl esters, phospholipids, and glycolipids. Sometimes they occur, although only in small amounts, in the free form. Fatty acids perform important physiological functions in the living cell. Acyl moieties, stored, for example, in triacylglycerols, may be degraded to acetyl-CoA, and via the citric acid and glyoxylate cycles can provide the cell with energy and cell material. The acyl moieties of phospholipids and glycolipids in cell membranes and organelles exhibit different degrees of unsaturation and thus determine the degree of fluidity and consequently the active transport properties of these membranes and organelles. The usual fatty acids in lipids of biological origin are predominantly those with 16 and 18 (in marine eukaryotes also 20) carbon chains. They may be saturated or unsaturated, with one or more double bonds per molecule. As far as the microbial fatty acids are concerned, differences exist between prokaryotes and eukaryotes. Unlike yeasts and filamentous fungi, bacteria are usually low in polyunsaturated fatty acids with two or more double bonds. Unusual fatty acids among microorganisms include the hydroxy acids, branched acids, cyclic acids, di- and tricarboxylic acids, and mycolic acids. In this section major emphasis is put on the usual fatty acids. Some of the unusual fatty acids, e.g., dicarboxylic acids and mycolic acids, will be considered elsewhere in this chapter. A. BASIC STUDIES Most studies have been done on the constituent fatty acids of total lipids from n-alkane-utilizing microorganisms. Only rarely were the constituent fatty acids of individual lipid classes investigated. Various studies had two different, yet not contradictory, objectives. The first is rather practical; n-alkane-utilizing microorganisms were investigated as potential sources of fatty acids and fats. The second objective is

n-ALKANE-UTILIZING MICROORGANISMS

43

academic; fatty acids were studied as a means for elucidating the biochemical mechanism(s) by which n-alkanes are microbiologically oxidized. Oxidation mechanisms leading to fatty acids are particularly relevant to the present section. However, for more detailed information the reader may refer to specific reviews (Klug and Markovetz, 1971; Einsele and Fiechter, 1971; Rehm and Reiff, 1981; Fukui and Tanaka, 1981; Buhler and Schindler, 1984). There are three different mechanisms known so far for the initial attack on n-alkanes by microorganisms. (1)Hydroxylation by a monooxygenase system in which cytochrome P450 may or may not be involved; (2) dehydrogenation leading to the n-alkene, which is subsequently hydrated; and (3) hydroperoxidation through a free-radical mechanism followed by reduction to the corresponding alcohol. The alcohol produced by any of the three mechanisms is then oxidized to the correspondingfatty acid. Here too are different pathways: (1)The monoterminal oxidation pathway in which one of the terminal methyl groups is oxidized leading successively to the corresponding 1-alkanol, 1-alkanal, and monocarboxylic fatty acid; this pathway is prevailing in many bacteria, yeasts, and fungi; (2) the diterminal oxidation pathway involves the oxidation of both methyl groups leading to the corresponding a,w-dicarboxylic (dioic) acid; (3) the subterminal oxidation pathway involves the oxidation of methyl groups leading successively to the corresponding secondary alcohol, ketone, and fatty acid. The composition of fatty acids and oxidation intermediates in the biomass and medium are indicative of the oxidation pathway prevailing in the culture. Cooney (1979) and Rehm and Reiff (1981) tabulated results of total fatty acid analysis for numerous microorganisms growing on n-alkanes [and n-alkenes) as substrates. In reviewing this subject, only representative data are tabulated, and readers interested in more details can refer to the reviews cited previously. Studies on bacteria date back to the sixties. Dunlap and Perry (1967) cultivated Mycobacterium sp. OFS on n-alkanes with 13, 14, 15, 16, or 1 7 carbon chains as sole sources of carbon, and analyzed the total cell fatty acids after 3 days. These authors showed that the cells accumulated in their total lipids fatty acids with chains equivalent in length to those of the substrates (Table VII). Cells grown on odd-chain alkanes tridecane [C13),pentadecane (CJ, and heptadecane (C17)have larger proportions of fatty acids with 1 7 carbon chains than cells grown on even-chain nalkanes. The same authors extended their work a year later (Dunlap and Perry, 1968) to cover the additional n-alkanes C,,, C,,, C,,, C,,, C,,, and C,,, and also studied Mycobacterium sp. 7EIC grown on C,,, Corynobacterium sp. grown on C17,and Brevibacterium sp. JOB5 grown

TABLE VII CONSTITUENT FATTYACIDSOF TOTALLIPIDSFROM Mycobacterium AND Micrococcus GROWNON II-ALKANES WITH DIFFERENT CHAIN LENGTHS' Mycobocterium sp. OFS Fatty acids 1o:o 11:0 12 : O 13:O 14:O 15:O 15:l 16:O 16:l 17:O 17:l 18:O 18:l

GIb

C,2b

-

-

2.1 5.1 0.4 4.1 1.1 Trace 3.3 7.0 2.9 1.4 0.4 0.1 30.6 27.5 18.1 13.8 1.2 0.5 2.0 0.6 0.8 20.7 17.5

C13c C,,'

C,,"

Micrococcus cereficons

GGC C,,c

CIab

-

-

-

-

-

-

1.4 0.9 21.4 1.8 6.6 1.7 15.3 14.3 3.6 9.8

1.0 2.9 0.9 39.0

0.5 1.0 0.2 5.9 1.5

-

7.2 4.0

-

50.0 28.2

-

0.8 5.7

-

-

-

9.6

19.4

-

2.5 0.3 2.3 0.5 18.1 2.7 3.8 2.5 19.4 27.2

-

-

0.8 0.5 6.9 0.9 74.0 8.9

-

5.0

-

19.5

2.9 1.3 0.4 28.3 15.5 0.4 0.5 1.9 26.3

Data expressed in relative percentage of the total fatty acids Data from Dunlap and Perry (1968). Data from Dunlap and Perry (1967). Data from Makula and Finnerty (1968).

C,,b

C2,b -

-

3.0

Trace Trace 22.0 18.0

-

0.1 0.3 3.0 10.4 2.5 21.8 13.8

Trace Trace

-

30.0

21.6

C2ab Clod -

3.8

0.4 0.8 3.1 6.0 21.1 3.7 15.8 10.6 3.0 8.5

-

13.8

C,,d

C,,d

0.5

7.6

0.7 0.9 2.6

-

Trace

-

4.4

1.7 1.9

-

1.6 1.7 7.2 7.7 2.2 1.2

Trace

-

Trace

18.7 14.2 11.2 15.6 6.3 26.1

23.3 14.3

17.3 11.6 7.7 7.7 6.7 27.4

39.5 14.5

6.7 23.6

-

6.6 0.8

-

6.6 47.5

C,,d 1.7

7.3

28.4

10.9 16.9

11.6 23.2

CIjd 1.1 1.2 1.3 1.1 0.2 32.6 36.2

Trace 4.5 3.3 3.8 11.3 3.3

CIGd C,,d 0.2

Trace

-

2.0 1.4 1.8 0.4 6.6 3.6 2.7 1.6 21.0 49.9 1.6 7.5

7.1

2.0 -

29.6 43.3

-

7.1 2.0

C,,d Trace

-

3.9

-

1.9 -

16.3 20.8 -

10.4 41.6

n-ALKANE-UTILIZING MICROORGANISMS

45

on C,, and C,,. The results (Table VII) also confirmed that cells grown on odd-chain alkanes accumulate larger proportions of fatty acids with 17 carbon chains and to a lesser extent 15 carbon chains in their lipids. Interestingly, cells grown on the very long chain alkanes, C,,, C,,, and C,,, do not accumulate any fatty acids with the same chain length and contained instead the usual fatty acids with 16 and 18 carbon chains, in addition to considerable concentrations of fatty acids with 15 and 17 carbon chains. The accumulation of C,, and C,, fatty acids indicates that very long even-chain alkanes are also attacked by mid-chain oxidation. Makula and Finnerty (1968), working on Micrococcus cerificans (Acinetobacter sp.) grown on C,,, C,,, C,,, C,,, C,,, C,,, C,,, C,,, and C,, n-alkanes, confirmed these results (Table VII). The data in Table VII indicate that the tendency to accumulate fatty acids with chains equivalent in length to those of the n-alkane substrates in cell lipids is valid within the alkane chain range C,,-C,,, even if the alkane is odd or even chained. Total lipids from Nocardia salmonsicolor PSU-N-18 grown on nhexadecane contain hexadecanoic (46%),hexadecenoic (4.5%),and octadecenoic (14.5°/0)acids as predominant fatty acids (Abbott and Casida, 1968). Killinger (1970) showed that total lipids from Pseudomonas sp. grown on acetate, propionate, or n-alkanes with different chain lengths contain fatty acids predominantly with 15 to 18 carbon atoms, and that fatty acids with odd chains accumulate in cell lipids after growth on propionate or odd-chain n-alkanes (Table VIII). Reportedly, the carbon chains of the fatty acids are synthesized mostly de novo, but can also be taken unshortened from the substrate. Confirming these results on Mycobacterium, Micrococcus (Acinetobacter), Pseudomonas, and other genera, Yano et al. (1971) found that lipids of Arthrobacter simplex contain significant amounts of odd-chain fatty acids only when grown on odd-chain n-alkanes (Table VIII). On the other hand, Edmonds and Cooney (1969) showed that lipids of Pseudomonas aeruginosa grown on n-tridecane contain stearic, palmitic, and palmitoleic acids as predominant fatty acids but only small proportions of pentadecanoic and heptadecanoic acids and no tridecanoic acid at all (Table VIII). bassilnikov et al. (1972)found that various strains of Mycobacterium lacticolum var. aliphaticum grown on n-hexadecane accumulate in their total lipids only hexadecanoic, tetradecanoic, and dodecanoic acids. Similarly, Koronelli et al. (1981) reported that total lipids of two Arctic, pigmented Mycobacterium strains grown on n-hexadecane contained palmitic acid (16 : 0)as a predominant usual fatty acid. Mycobacterium vaccae JOB5 grown on n-pentadecane accumulates large pro-

TABLE VIII CONSTITUENT FAITY ACIDSOF TOTALLIPIDSFROM Pseudomonos AND Arthrobacter GROWNON n-ALKANES WITH DIFFERENT CHAIN LENGTHS' Arthrobacter simplex

Pseudomonas sp. Fatty acids Acetateb Propionateb

1o:o

4.0

1l:O 12:o 13:O 14:O 15:O 15:l 16:O

-

16:l 17:O 17:l 18:O 18:l >I8 a

5.6

4.0

29.4 23.5

-

1.8 1.9 3.5 0.4 0.5 6.6 1.2 7.0 6.3 28.4 22.4

Glob

Cllb

18.3

0.7 2.6 2.4 1.6 Trace 1.4 2.6 2.4 6.6 3.6 15.8 32.6 15.3 1.5 16.5 2.3 13.8 44.0 18.4 7.4

1.5

-

2.0

-

23.4 29.8

-

-

-

34.5

19.6

25.0

-

-

-

-

C13'

-

Data expressed in relative percentage of the total fatty acids. Data from Killinger (1970). Data from Edmonds and Cooney (1969). Data from Yano et al. (1971).

Cl,b

CISb Cl,b

1.4

1.0 2.8 1.8 3.3 1.6 2.4 1.3 1.1 0.4 0.9 15.8 6.2 3.1 9.1 5.1 7.8 18.3 12.2 12.0 17.6 15.4 21.6

-

4.8 -

8.3

37.7 19.8

Cl,d

Cl,d

C,,d

2.0

-

-

5.5

-

-

-

-

5.1

7.0

1.1

0.5 6.9 1.9

1.5 33.4

1.5 2.5

-

-

C,,b

-

-

5.8

-

-

-

35.6 15.9

9.9 1.2 0.4 2.9 1.7 54.5

-

-

_

-

28.0

18.1

20.8

7.5 26.5

20.4 7.7 2.9 17.4 2.0 37.7

-

-

-

-

-

-

-

C,,d

1.1 Trace

10.0 38.9 53.0 2.3 2.3 Trace 1.1 29.1 1.4 Trace 18.8 3.0

-

-

CITd

Cl,d

C,,d

-

-

-

Trace Trace 1.8 Trace

-

Trace Trace

9.7

0.9

9.6

-

-

-

1.8 0.2 22.6 46.4

30.3 6.8 0.9

Trace

Trace

Trace

19.2

5.0 54.3

-

-

2.0 21.2 3.4 0.7 13.1 39.9

n-ALKANE-UTILIZING MICROORGANISMS

47

portions of pentadecanoic (51%) and pentadecenoic (14.4%) acids in its total lipids; cells grown on n-heptadecane contain high concentrations of heptadecanoic (22.6%), heptadecenoic (19.1%),and pentadecanoic (17.9%) acids in their total lipids (King and Perry, 1975). Reportedly, cells grown on n-tetradecane, n-hexadecane, and n-octadecane contain only minute amounts of odd-chain fatty acids, if at all. Makula et al. (1975) found that Acinetobacter sp. grown on n-pentadecane and n-heptadecane contain relatively large proportions of fatty acids with 15 and 1 7 carbon chains. On the other hand, cells cultivated on nhexadecane do not contain any odd-chain fatty acids. In the total lipids of glucose-grown cells of Corynebacterium cyclohexanicum, methyl tetradecanoic (35%) and methyl pentadecanoic (35%) acids are the major constituent fatty acids, whereas in the lipids of cells grown on cyclohexanecarboxylic, m-hydroxybenzoic, butyric, and acetic acids, methyltetradecanoic acid is the major fatty acid, making up 65-81Yo of the total fatty acids (Kaneda, 1983). In addition to n-alkanes, bacteria can also terminally oxidize 1alkenes (Dunlap and Perry, 1968; Makula and Finnerty, 1968), cyclohexylalkanes (Beam and Perry, 1974),and chlorinated alkanes (Murphy and Perry, 1983). Table IX indicates that Mycobacterium convolutum can oxidize 1-chlorohexadecane to 1-chlorohexadecanoic acid, which it can then desaturate, shorten, elongate, and incorporate into different lipid classes, including diacylglycerophosphoinositolmannosides,the predominant phospholipid (Murphy and Perry, 1987). Koronelli et al. (1988) suggested the use of tritium-labeled n-alkanes in estimating the activity of alkane-oxidizing bacteria. Oxidation of octadecane in the unlabeled CH, group leads to labeled stearic acid, whereas its oxidation in the labeled CH, group results in the loss of radioactivity. In their comparative study they concluded that Rhodococcus erythropolis is 70 times more active in alkane oxidation than Pseudomonas aeruginosa. Yeasts and filamentous fungi are eukaryotes; they differ from bacteria in possessing organelles. Therefore, early studies on n-alkane-utilizing yeasts have debated about whether the substrate oxidation occurs in mitochondria, cytoplasmic membranes, or elsewhere in the cell (Van der Linden and Huybregste, 1967; Lebeault et al., 1970;Liu and Johnson, 1971). The whole subject has been reviewed by Fukui and Tanaka (1981). In Candida yeasts, n-alkanes are first hydroxylated to fatty alcohols in microsomes (see also Blasig et al., 1988). These alcohols are subsequently oxidized to fatty acids via aldehydes in microsomes, mitochondria, and peroxisomes. Further P-oxidation of the fatty acids occurs exclusively in peroxisomes, whereas fatty acids produced in microsomes and mitochondria are incorporated into various complex lipids.

48

SAMIR S. RADWAN AND NASER A. SORKHOH TABLE IX CONSTITUENT FATTY ACIDS OF TOTAL LIPIDS AND

DIACYLGLYCEROPHOSPHOINOSITOLMANNOSIDESOF Mycobocterium C O n V O l U t U m GROWN ON n-HEXANE AND 1-CHLOROHEXADECANE"

Total lipids Fatty acids 14:O 15:O 16:O 1 7 : Obr 16:l 9 1 6 : l 11 18:O 1 9 : Obr 18:l CI 1 2 : o c1 13:O C1 14:O C1 15:O Cl 1 5 : l Cl 16:O C1 17:Obr c1 1 6 : l c1 1 8 : O C1 1 9 : Obr Cl 1 8 : l

Hexane

Chlorohexane

Trace Trace

Hexane

Chlorhexane

2.0

6.2

Trace

Trace 48.6 3.8 35.8 3.1

Diacylglycerophosphoinositolmannosides

3.9

Trace 1.1

Trace Trace Trace Trace 1.6

Trace

10.6 1.2

52.1 30.4 6.2 5.8

Trace Trace

-

-

3.0

1.0

Trace Trace Trace

26.2 4.2

19.7

Trace

-

38.1 4.4 14.2

29.7 17.3 4.2

-

-

1.6 4.5

7.2 1.1

8.0

Data from Murphy and Perry (1987). Values are expressed in relative percentage of total fatty acids.

In this context, Ermakova and Lozinov (1976) isolated 13 species belonging to seven genera of yeasts, which could neither grow on liquid paraffin nor oxidize n-octadecane, but grew well on tridecanol, and oxidized various aliphatic alcohols and fatty acids. These authors concluded that the only reaction that can be considered typical of n-alkane-utilizing yeasts is the oxygenase reaction leading to the fatty alcohols. Schunck et al. (1983) isolated and reconstituted the alkane monooxygenase systems of the yeast Lodderomyces elongisporus. On the cytological level, the possession of large numbers of peroxisomes is considered to be one of the specific features of alkane-utilizing yeasts (Fukui and Tanaka, 1981). As has been demonstrated in bacteria, yeasts also tend to accumulate fatty acids with chains equivalent in length to those of the alkane sub-

n-ALKANE-UTILIZING MICROORGANISMS

49

strates in their total lipids (e.g., Klug and Markovetz, 1967a,b). This is, however, particularly valid for n-alkanes with C,, to C,, chains. Before reviewing these studies, attention should be directed to some contradictions in the literature probably because of strain and/or cultural condition variations. Such contradictions are obvious in studies concerned with the proportions of saturated and unsaturated fatty acids in Candida yeasts. A group of earlier workers reported that lipids from yeast cells grown on n-alkanes contain larger proportions of unsaturated fatty acids at the beginning of the incubation period than at the end (Dyatloviskaya et a]., 1965; Pelechova et al., 1971).Another group of authors conversely showed that the proportion of unsaturated fatty acids is low in the beginning and increases at the end (Hug and Fiechter, 1973; Volvova and Pecka, 1973). A third group demonstrated that the proportion of unsaturated fatty acids remains rather constant throughout the incubation period (Mishina et al., 1973, 1977). Rattray et al. (1975) realized that yeast cells grown on odd-chain nalkanes contain large proportions of odd-chain fatty acids in their total lipids. Mizuno et al. (1966) showed that Candida petrophilum grown on n-hexadecane contains in its total lipids almost exclusively C,, and C,, fatty acids. In contrast, n-tridecane-grown cells contain, in addition to these acids, considerable proportions of heptadecenoic (38.8%)and pentadecanoic (9.3%) acids. The results of Klug and Markovetz (l967b), Mishina et al. (1973), and Jwanny (1975) on Candida Iipolytica grown on different n-alkanes are summarized in Table X and the results of Hug and Fiechter (1973) and Mishina et al. (1973) on C. tropicalis are presented in Table XI. Despite obvious differences in the values given by various authors, it may be concluded that yeast cells grown on n-alkanes with chains between C,, and C,, accumulate fatty acids with equivalent chain lengths in their total lipids. Moreover, the largest proportions of oddchain fatty acids are present in cells grown on odd-chain n-alkanes. Similar results have been recorded for Candida rugosa grown on nalkanes with C,, up to C,, chains (Iida et al., 1980) and Mycotorula japonica grown on C,,, C,,, C,,, C,,, C,,, and C,, n-alkanes (Yamaguchi and Kurosawa, 1976). Souw et al. (1976) showed that tetradecanoic acid is produced by Candida sp. through monoterminal oxidation of n-tetradecane with ntetradecanol as an intermediate, and demonstrated the desaturation of the fatty acid to its monoenoic homolog. No odd-chain fatty acids are present in the cell lipids. Lipids from n-tetradecane-grown cells of C. parapsilosis have similar fatty acid patterns to lipids from glucosegrown cells (Omar and Rehm, 1980).Similarly, it has been demonstrated

TABLE X CONSTITUENT FATTYACIDSOF TOTALLIPIDSFROM Candida lioolvtica GROWNON ALKANES WITH DIFFERENT CHAINLENGTHS' Fatty acids

Cllb

CIzb

C13b

C,,b

C,,"

Cl,b

CISc

Ul

0

11:o 12:o 13 : O 14:O 14:l 15:O 15:l 16:O 16:l 17:O 17:l 18:O 18:l 18:2

1.1

-

-

11.7

0.3

-

Trace -

5.0 0.6

2.5

Trace

-

-

13.4 14.6

10.4 17.4

Trace

-

5.7 2.4 47.9 12.1

0.9 3.1 43.1 7.9

Trace 8.5 0.3

16.6 1.8 2.0 6.1 1.5 31.6 0.6 20.6 10.5

C,,b

C,,'

-

Trace Trace

-

Trace

Trace Trace Trace

19.5 2.6

3.0

0.3

0.8

0.1

5.0 5.0

-

-

-

-

-

Trace Trace

4.5

-

8.5 24.3

13.5 7.5

Trace

Trace Trace

20.0 2.9 0.3 1.4 1.5 56.5

20.8 1.7 2.6 4.3 0.8 25.4 12.3 8.6 19.7

1.0 0.8 29.6 13.5

44.8 10.1 14.9

Data expressed in relative percentage of the total fatty acids. From Mishina et al. (1973). From Klug and Markovetz (1967b). From Jwanny (1975).

-

14.3 7.2

-

0.6

5.0

-

-

30.2 27.7

30.0 20.0

-

Trace Trace

1.0 2.1 30.0 8.3

Cl,C

CIBb

C1,C

-

0.8

-

Trace Trace -

16.8 12.3

21.4 6.7

-

Trace

3.7 7.9 41.2 16.7

5.3 6.7 16.7 34.8

~

Trace Trace

-

Cl,b

C,,d

~~~~

~~

10.0 25.0 -

-

-

1.3

1.7 2.9 2.0 3.0 10.0 3.7 26.7 9.0 2.9

-

-

0.2 0.4

2.0

-

Trace -

-

5.6 17.0 6.3

-

2.1 0.8 0.7 0.6 4.6 77.9

-

0.5

3.9 3.9 5.2 2.6 14.3 36.6 19.6 3.9 5.2

-

2.6 4.0

TABLE XI CONSTITUENT FATTY ACIDSOF TOTALLIPIDSFROM Candida t r o p i c a h GROWNON ALKANES WITH DIFFERENT CHAINLENGTHSO ~

Data from Hug and Fiechter (1973) Fatty acids Glucose

L ul

ll:o 12:o 13:O 14:O 15:O 15:l 16:O 16:l 17:O 17:l ia:o ia:i ia:z i8:3 19:o

0.5 Trace 0.6

1.3 0.4 13.4 12.0 0.8

2.2 14.6 23.0 30.5

Acetate

CI2

0.5 Trace 1.5 1.3 0.7 16.0 11.4 1.3 2.o 14.3 22.2 28.5

24.4 Trace 1.4 Trace

C,,

C,,

0.5 2.4 0.4 28.4 Trace 0.8 33.3 7.2 Trace 0.4 10.4 9.6 7.3 6.8 5.0 11.8 Trace 1.3 Trace 2.0 5.5 5.3 12.5 14.1 5.3 33.5 24.7 19.1 8.4 7.0 4.5 Trace Trace

Data expressed in relative percentage of the total fatty acids.

~~

~

Data from Mishina et al. (1973)

C15

C,,

1.2 0.9 0.5 0.5 22.8 4.5 2.6 2.6 2.0 15.6 18.6 16.9 4.6 5.4

-

0.1 Trace 0.4 0.1 Trace 28.9 27.7 Trace 5.5 8.3 21.2 7.6 -

-

C17

Cll

C12

1.0 1.3 1.2 1.4 1.9

-

0.5

-

1.4 Trace _ _ _ 2.7 3.8 10.6 0.2 0.8 Trace 0.4 11.5 Trace 18.3 0.5 0.4 0.9 17.2 2.0 15.1 0.8 33.8 0.5 12.5 2.3 24.9 1.4 33.7 0.3 3.4 0.5 6.0 Trace 17.8 2.4 49.2 2.1 50.9 1.1 65.6 0.6 0.6 2.9 Trace 40.5 17.9 28.1 18.3 23.0 12.5 16.2 7.1 14.0 4.5 6.0 1.8 1.8 0.8 3.9 1.2 1.0 0.8 -

1.9

8.1 5.8 5.1 2.1 17.1 8.6 25.3 30.0 24.8 1.3 3.4 27.0 0.8 20.3 - 2.9 5.5 -

C13

C14

CIS

C16

C,,

C18

7.2 1.3 0.3 5.6 67.0 15.8 2.7

52

SAMIR S. RADWAN AND NASER A. SORKHOH

that n-pentadecane is monoterminally oxidized to pentadecanoic acid via pentadecanol by Candida sp. (Souw et al., 1977). Moreover, evidence for diterminal and subterminal oxidation, as well as for the desaturation of pentadecanoic acid, is presented. Blasig et al. (1984) incubated glycerol-grown cells of Lodderomyces elongisporus for a few hours with n-hexadecane and n-heptadecane as sole carbon sources, and demonstrated that the levels of hexadecanoic and heptadecanoic acids in the cell lipids increase dramatically (Table XII). Table XI1 indicates that these fatty acids are actively desaturated by the yeast cells to the corresponding monoenoic acids (16 : 1 and 17 : 1).These findings were confirmed by studying the distribution of radioactivity of [l -l4C] hexadecane, as a sole carbon source, among the fatty acids of total lipids from this yeast. The results (Table XIII) indicate the occurrence of both fatty acid elongation and @-oxidation.The latter pathway is substantiated by the appearance of [14C]myristic(14 : 0)and -1auric (12: 0 ) acids. There is little information about the oxidation of very long n-alkanes with C,, and longer chains by yeasts. The only available study is that of Blasig et al. (1989) who demonstrated that Candida maltosa could TABLE XI1 CONSTITUENT FAITY ACIDSOF TOTALLIPIDSOF Lodderomyces elongisporus BEFORE AND AFTER INCUBATION WITH n-ALKANES' ~

4

Fatty acids 12:o 14:O 15:O 16:O 16:l 16:2 17:O 17:l

Before incubation 0.57 0.94 18.84 9.35 0.52

hr after incubation with C,,

C,,

0.05

0.05 0.22 1.36 7.72

0.99

0.35 68.28 41.31 2.18

-

-

18:l 18:Z 18:3

1.73 24.97 1.16 0.36

20:o

-

0.15 29.20 20.31 5.30

18:O

Total

58.41

?

6.02

167.91 ? 24.06

-

58.52 27.65 0.35 8.25 0.45 1.09 105.65 ? 22.96

Values are expressed in f i g fatty acid per mg protein. Cells were grown on glycerol as a carbon source before they were incubated with alkanes. Results from Blasig et of. (1984).

53

n-ALKANE-UTILIZING M I C R O O R G A N I S M S T A B L E XI11

DISTRIBUTION OF RADIOACTIVITY AMONG FATTY ACIDS OF Lodderomyces e l o n g i s p o r u s AFTER INCUBATION WITH [1-'4C]HEXADECANEa Fatty acids

0.2

hr

-b

12:o 14:O 16:O 16 : 1 16:2

10.1 1.97 ? 0.22 10.1

18:O

<0.25

18:l 18:2 18:3

-

Total

2.11 t 0.22

a

-

1 hr

2 hr

4 hr

10.01 CO.1 5.61 t 0.99 1.34 2 0.22 10.1 10.25 <0.25

10.1 <0.25

<0.1 10.25 13.73 f 0.24 7.45 t- 0.14 1.03 f 0.03 0.39 i 0.01 1.39 f 0.04 1.19 ? 0.06 0.25 f 0.01 25.55 f 0.29

<0.1

-

10.01 7.41 ? 1.01

Values are expressed in Not detectable.

13.64 ? 0.03 5.01 i 1.07 0.40 i 0.17 0.29 f 0.05 0.67 ? 0.14 0.56 ? 0.22 10.1 20.72

f

1.11

8

hr

10.1

10.25 34.56 f 2.43 26.69 t 0.70 2.43 t 0.46 0.81 t 0.39 8.02 f 0.13 3.98 t 0.31 0.90 f 0.29 77.68 f 2.64

20

hr

10.1 10.25 31.15 i 0.59 29.27 f 0.91 1.13 ? 0.97 0.90 f 0.17 19.61 f 0.88 3.28 ? 0.04 0.45 i 0.04 86.03 i 1.71

lo3cprn/rng protein. Results from Blasig et al. (1984).

assimilate C,o-C2, n-alkanes via monoterminal oxidation. However, the corresponding very long chain fatty acids do not accumulate in the cell lipids in any significant amounts. Such fatty acids seem to be immediately chain shortened by C, units down to an optimal range of chain length from C,, to C,,, and incorporated into cell lipids, directly or after desaturation. Even- and odd-chain fatty acids predominate in experiments with even- and odd-chain n-alkanes, respectively. The total fatty acid patterns of certain filamentous fungi grown on nalkanes seem to differ from those of n-alkane-grown yeasts and bacteria. The results of studies published so far indicate that fungi grown on C,,, C,,, and C14 n-alkanes tend to accumulate the usual fatty acids with 16 and 18 carbon chains in mycelial lipids. Fatty acids with chains equivalent in length to those of the n-alkanes may or may not increase. Fungi grown on C,,, C,,, and C,, n-alkanes are reported by some investigators to accumulate mainly odd-chain fatty acids while others report even-chain fatty acids. In many of the studies it has been observed that fungi grown on C,, and C17n-alkanes surprisingly accumulate decanoic (10 : 0)and undecanoic (11:0) acids, respectively (Table XIV) in their lipids, an observation not recorded in bacteria and yeasts. Cooney and Proby (1971) showed that the total lipids of Cladosporium resinae grown on C,,, C,,, C,,, C,,, and C,, n-alkanes exhibit similar fatty acids patterns; the C,, and C,, fatty aci+ (saturated and unsaturated) predominated. Lin et al. (1971a) found:interesting differences in the total fatty acid patterns of n-alkane-grown Cladosporium sp. and Homo-

TABLE XIV CONSTITUENT

FATTYACIDSO F TOTAL LIPIDS FROM FILAMENTOUS FUNGI GROWNON n-ALKANES

O F DIFFERENT CHAIN

LENGTHSO

Cunninghamella Cladosporiurn ~ p . ~

Horrnodendrum hordeib

Absidia spinosaL

echinulataC

Mortierella isabellinoc

Fatty acids

Glucose

C,,,

C,,

C,,

C,,

Glucose

Clo

1o:o

Trace

32.4

-

Trace

15.5

0.6

6.8

-

1.4

4.6

23.5 -

-

11:o

83.0

-

-

16.2

73.3

12:o

0.6 -

Trace

1.7

Trace

Trace

4.1

13.0

Trace

13:O

-

-

-

-

-

-

Trace

14:O

Trace

Trace

Trace

-

0.2

Trace

15:O

0.9

-

0.4

-

0.4

Trace

-

Trace

16:O

27.7

29.7

42.3

36.3

1.4

36.5

16:l

2.1

17:O

Trace

C,,

C,,

C,,

Glucose

C,,

C,,

Glucose

C,,

-

C,,

Glucose

C,,

C,,

-

-

Trace

-

-

-

0.2

8.6 -

0.1

01

10.0

8.6

Trace

-

10.1

0.1

-

24.0

Trace

Trace

14.1

0.3

0.9

3.9

0.6

0.5

2.9

0.4

Trace

3.0

0.9

0.3

0.2

0.4

23.2

Trace

0.3

170

-

1.o

12.8 11.1

-

-

04

01

0.1

33.1

26.1

6.3

15.4

25.3

6.3

20.2

24.4

5.6

21.7

23.8

-

5.0

14.9

4.7

2.6

4.2

1.1

1.6

2.8

0.9

3.6

6.1

1.7

Trace

0.3

Trace

0.3

8.2

-

0.2

8.5

Trace

0.6

4.3

-

3.1 -

-

Trace

0.1

14.1

-

0.2

16.6

Trace

0.7

11.9

3.1

-

-

-

-

18:O

Trace

Trace

Trace

Trace

Trace

-

-

4.2 -

4.7

1.7

0.6

4.6

3.9

0.9

3.1

1.1

0.9

18:l

11.9

10.8

9.2

19.0

18.3

5.8

2.6

-

47.3

18.8

4.6

52.8

27.0

9.0

54 8

25.4

12.5

18:Z

53.6

22.7

39.4

20.8

34.9

33.1

20.3

1.3

14.3

11.9

4.9

11.3

16.2

8.8

11.5

12.9

8.8

18:3

5.2 -

0.5

2.1

Trace

12.3

8.7

11.8

1.4

-

16.5 -

11.8

-

14.4 -

24.6

-

10.0 -

17:l

9.0 -

1.7 -

-

-

0.2

19:l

-

-

3.8 -

Trace

-

-

2.4

-

-

20:o

0.6

0.1

-

Trace

0.2

0.2

0.1

Trace

19:o

" Data expressed

0.9

in relative percentage. From Lin et al. (1971a), similar results were also recorded for Penicillium lilocinum and Aspergillus versicolor (Lin et of., 1971a) From Hoffmann and Rehm (1978).

0.1

0.8

Trace

n-ALKANE-UTILIZING MICROORGANISMS

55

dendrum hordei at different growth phases. Thus, n-decane (Clo) and n-undecane (C,,)-grown fungi have higher levels of decanoic (10:0) and undecanoic (11: 0)acids, respectively, in early than in later growth phases. In contrast, n-hexadecane (C,,)-grown fungi have much higher concentrations of palmitic acid (16 : 0) in later than in early growth phases. Fungi grown on n-heptadecane (C17)have, at all phases of growth, relatively high concentrations of undecanoic acid (11: 0) in their total lipids. In another study, the same authors (Lin et al., 1971b) reported that the major mycelial fatty acids of Penicillium lilacinum and Aspergillus versicolor grown on n-decane and n-undecane are the usual c,, and C,, fatty acids. On n-hexadecane and n-heptadecane, P. lilacinum contains decanoic and undecanoic acids, respectively, as major fatty acids. Pelz and Rehm (1973) demonstrated that the oxidation of various n-alkanes by Cunninghamella echinulata, Absidia glauca, and Mucor sp. to the corresponding fatty acids is achieved monoterminally. The same authors found, however, that subterminal oxidation occurs in species of Aspergillus, Penicillium, and Verticillium leading to fatty acid patterns that do not clearly depend on the identity of the n-alkane substrate. Cerniglia and Perry (1974) showed that the predominant fatty acid in the total lipids of Penicillium zonatum grown on ntridecane, n-tetradecane, and n-pentadecane is linoleic acid (18 :2), and that the mycelia accumulated considerable amounts of fatty acids equivalent in chain length to the n-alkane substrate. Gerasimova and Lin (1975) observed that Cunninghamella elegans grown on n-undecane, n-dodecane, n-tetradecane, and n-hexadecane accumulate mainly even-chain fatty acids in mycelial lipids; C,, and C,, fatty acids predominate in all cases. In contrast, n-tridecane-, n-pentadecane-, and n-heptadecane-grown mycelia accumulate predominantly odd-chain fatty acids. A few investigators studied the assimilation of chlorinated n-alkanes by yeasts and filamentous fungi. Murphy and Perry (1984) cultivated Candida lipolytica, Cunninghamella elegans, and Penicillium zonatum on 1-chlorohexadecane and 1-chlorooctadecane as sole sources of carbon, and analyzed the biomass fatty acids. Sixty to 70% of the total fatty acids from C. elegans and about 50% of total fatty acids from C. iipolytica were chlorinated. Penicillium zonatum contained 20% 1chlorohexadecanoic acid after growth on either substrate, but did not incorporate any C,, chlorinated fatty acids. Fatty acids accumulate in n-alkane-utilizing microorganisms, not only intracellularly, but may also be excreted into the growth medium (for bacteria, see Romero and Brenner, 1966; Makula et al., 1975; Kachholz and Rehm, 1977; for yeasts, see Klug and Markovets, 1967b; Yama-

SAMIR S. RADWAN AND NASER A. SORKHOH

56

gushi and Kurosawa, 1976; Souw et al., 1977; and for filamentous fungi, see Siporin and Cooney, 1975).These extracellular fatty acids, however, are not produced from the n-alkanes extracellularly. The n-alkane is first taken up by the cell, where it is metabolized. The accumulation of fatty acids (and other lipid classes) into the aqueous medium may be due to their solubility in the n-alkane substrate which extracts them from the cells. In addition, many alkane-utilizing microorganisms excrete biosurfactants into the medium which may emulsify and extract fatty acids and lipids from the cells. There is not much information available on the fatty acids of photosynthetic microorganisms grown in the presence of n-alkanes. The only available publication is that of Schroeder and Rehm (1981)who reported on the fatty acids of Chlorella vulgaris grown in the light with and without n-tridecane. The results (Table XV) indicate that incubation with the alkane is associated with an increased concentration of the polyunsaturated fatty acids, linoleic (18:2) and linolenic (18:3) acids, in the total lipids of this alga. However, more work is still needed before final conclusions regarding the effect of n-alkanes on algal fatty acids can be drawn. B. BIOTECHNOLOGICAL CONSIDERATIONS

Greater amounts of fatty acids, 5- to 15-fold or more, accumulate in microorganisms during growth on n-alkanes than the amounts produced during growth on conventional carbon sources (Makula and Finnerty,

TABLE XV CONSTITUENT FATTYACIDSOF TOTALLIPIDS FROM Chlorella vulgaris GROWNIN THE LIGHTWITH AND WITHOUT n-TRIDECANE' ~~~~

~

Fatty acids

Without n-tridecane

With n-tridecane

ll:o

2.6 5.1 66.2 6.8 3.9 8.1 7.3

1.7 2.9 32.4

(Y

14:O 16:O 18:O 18:l 18:2 18:3

17.8 23.2 22.0

~~

Data from Schroder and Rehm (1981). Values are expressed in relative percentage of total fatty acids.

n-ALKANE-UTILIZING MICROORGANISMS

57

1972; Boyer and Pisano, 1974). The amount of fatty acids produced can be correlated with the biomass and can be used as an index of the extent of n-alkane utilization (Levkina and Rebrikova, 1976). Miyakawa et al. (1984) showed that a mutant of Candida lipolytica can also excrete fatty acids into the medium up to 1 mg/ml. These facts imply that nalkane-utilizing microorganisms may be suitable tools for the biotechnological production of fatty acids. However, one of the problems associated with biomass production using n-alkane mixtures as substrates is the relatively high content of odd-chain fatty acids in the product (Rattray et al., 1975). These compounds result from terminal oxidation of odd-chain alkanes normally present in the oil-distillation fractions used as substrates. To solve this problem, it was suggested to use microbial strains that fail to accumulate odd-chain fatty acids, even in the presence of odd-chain alkanes (Fukui and Tanaka, 1980). Such strains are found among Candida yeasts. The attack on this problem was made possible after it was realized that n-alkane-grown cells of c. lipolytica contain two acyl-CoA synthetases; synthetase I is involved in the production of acyl-CoA, which incorporates acyl moieties into complex lipids, and synthetase I1 is linked to the p-oxidase system which degrades acyl moieties to acetyl-CoA (Kamiryo et al., 1977; Mishina et al., 1978a,b). Mutants defective in synthetase I were isolated by Kamiryo et al. (1977) and were cultivated by Tanaka et al. (1978) on odd-chain n-alkanes with 11 to 1 7 carbon atoms. For comparison, the wild-type strain was also studied. The analysis of total fatty acids of the cells showed that in the wild strain the odd-chain fatty acids made up 98-99% of the total fatty acids, whereas in the mutants the proportion of odd-chain fatty acids did not exceed 12-13% (Table XVI). It is quite obvious that the mutant strains synthesize their even-chain fatty acids de novo from acetyl-CoA units produced from odd-chain fatty acids by synthetase 11. Such mutants may be useful for the production of biomass, low in oddchain fatty acids, from oil-distillation fractions. Microorganisms active in the diterminal oxidation of n-alkanes are potentially suitable for the biotechnological production of dicarboxylic acids. Many of the long-chain dicarboxylic acids are used as raw materials in the perfume industry (Uchio and Shiio, 1974). In addition, they represent basic substrates for the synthesis of plasticizers, lubricants, and other products (Buhler and Shindler, 1984).Most studies on dicarboxylic acids used yeasts such as Candida, Pichia, and Torulopsis (Ogino et al., 1965; Iizuka et al., 1966; Okuhara et al., 1971; Shiio and Uchio, 1971; Uchio and Shiio, 1972a,b; Souw et al., 1977; Yi and Rehm, 1982a,b,c, 1988a,b, 1989), and a few investigators used fungi (Lin et al., 1971a,b).Bacteria have not frequently been reported to produce large

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TABLE XVI

FAITY ACIDSOF TOTALLIPIDSFROM Candida CONSTITUENT 1ipOlytiCa GROWNON n-PENTADECANE' Fatty acids

Wild-type 0.1 0.2

13:O 14:O 14:l 15:O 15 : 1 16:O 16:l 17:O 17:l 17:2 18:O 18:l 18:2 19:o 19:l

Trace 22.9 2.5 0.3 0.4 5.2 55.2 9.9 Trace Trace 1.3 Trace 2.0

Total oddchain fatty acids

97.8

Mutant L-5

Mutant L-7

0.1 0.3 0.3 7.6 0.1 12.2 18.7 3.6

Trace 0.6 0.3 5.8 0.2 12.9 17.8 Trace 2.9

0.7 28.9 26.9

1.0 31.3 27.3

11.4

8.9

Trace

Data from Tanaka et al. (1978). Values are expressed in relative percentage of total fatty acids. (I

amounts of dicarboxylic acids, although Klein et al. (1979) described a series of long-chain high-molecular weight dicarboxylic acids with vicinal dimethyl branching as major components of the lipids of Butyrivibrio spp. However, these authors assume that such dicarboxylic acids are produced by the union of two fatty acid chains and not by diterminal oxidation of n-alkanes. Studies on yeasts repeatedly demonstrate that odd- and even-chain fatty acids, as well as saturated and mono-unsaturated dicarboxylic acids, are produced from the appropriate substrates. However, usually a mixture of compounds with chains shorter than that of the alkane substrate is produced instead of the dicarboxylic acid with the carbon skeleton identical with that of the substrate (see Ogata et a]., 1973). To solve this biotechnical problem mutants lacking the ability to assimilate dicarboxylic acids can be created and applied in the fermentation process (see, for example, Uchio and Shiio, 1972b). Another potential field of application is the microbiological production of short-chain fatty acids, which command much higher prices

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than the long chain fatty acids with 16 and 18 carbon chains. This idea has been considered since early stages of petroleum microbiology. Ratledge (1968) cultivated Candida sp. on a fraction of n-alkanes predominating in tridecane in a fermentor and found after 108 hr that about 54% of the total fatty acids of the cells were shorter in chain length than CI6; with glucose as a substrate, this value did not exceed 2%. Maximum conversion was 71.5% (w/w)for alkanes into cells and 24.8% for alkanes into fatty acids. Ratledge (1970) showed that Candida 107 could achieve a 25% conversion of a fraction of n-alkanes with C,, to C,, to fatty acids. Another proposed field of study that has not been considered so far is the possible application of n-alkane-utilizing microorganisms in the biosynthesis of polyunsaturated C,, and C,, fatty acids. These fatty acids, e.g., arachidonic acid (20:4) and eicosapentaenoic acid (20:5), are valuable because of their unique physiological activities and as precursors of prostaglandins and other eicosanoids (Pace-Asciak and Wolfe, 1971; Horton, 1972; Deby, 1988). They are obtained from protozoa and fish oil, which makes their large-scale production rather uneconomical; therefore alternative sources are recommended (Korn et al., 1965; Iizuka et al., 1979). Several potential sources have been suggested (Radwan, 1991a), e.g., fungi growing on fatty acids with shorter chains (Radwan and Soliman, 1988) and on hydrocarbons (Iizuka et al., 1979), algae maintained under specific light colors (Radwan et al., 1988), and mosses (Al-Hasan et al., 1989). Shimizu et 01. (1989) described the conversion of linseed oil to an eicosapentaenoic acid containing oil by Mortierella alpina. Marine macroalgae that are active in the synthesis of these polyunsaturated fatty acids may be enriched with them, when they are incubated with specific n-alkanes under controlled environmental conditions. In this context, it should be reiterated that Chlorella vulgaris incubated with n-tridecane produce more polyunsaturated C,, fatty acids than control cells (Schroeder and Rehm, 1981). V. Acylglycerols

Bacteria and yeasts growing on n-alkanes contain smaller amounts, or at most slightly higher concentrations, of triacylglycerols than cells grown on conventional carbon sources (Davis, 1964;Makula et al., 1975; Thorpe and Ratledge, 1972). Alkane-utilizing bacteria are, as a rule, not rich in triaclglycerols, although they may produce large amounts of partial acylglycerols, particularly monoacylglycerols. These latter compounds are amphipathic and consequently surfactive, and this may explain why some alkane-grown bacteria accumulate them, especially

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extracellularly. Makula et al. (1975) showed that Acinetobacter sp. grown on n-hexadecane contains only traces of monoacylglycerols and diacylglycerols intracellularly, but produces extracellularly about 18fold more of these compounds than cells grown in nutrient broth. Hallas and Vestal (1978) found that the neutral lipid fraction of Mycobacterium convolutum made up about 70% of the total lipids from acetate-grown cells. When the cells were grown on C,,, C,,, C,,, C,,, C,,, C,,, and CZ8 n-alkanes, the neutral lipid fraction decreased from 66 to 48%. These authors did not identify individual neutral lipid classes which may comprise, in addition to acylglycerols, wax esters, steryl esters, sterols, and fatty acids. Because of the relatively low concentration of triacylglycerols in nalkane-utilizing bacteria only limited attention has been given to the study of their constituent fatty acids. Romero and Brenner (1966) found that triacylglycerols of Pseudomonas aeruginosa grown on n-hexadecane contained almost exclusively even-chain fatty acids with 18, 16, and 14 carbon chains. The predominant acids were linoleic (35.2%), palmitic (22.3%), and octadecenoic (19.7%) acids. Egorov et al. (1986) reported that triacylglycerols represent a main lipid class in Rhodococcus rubropertinctus grown in n-hexadecane, and that myristic acid (14:O) is the predominant fatty acid in this lipid class. Candida yeasts, which include oleaginous species (Pedersen, 1962; Gill et al., 1977; Moon and Hammond, 1978;Eroshin and Krylova, 1983), also seem to have only a limited capacity to accumulate triacylglycerols when they utilize n-alkanes as substrates. Thorpe and Ratledge (1972) found that n-alkane-grown yeasts contain smaller proportions of triacylglycerols than cells grown on conventional carbon sources. Shigyo and Takeuchi (1972) calculated that the total lipids of n-alkane-grown yeasts contain only about 25% simple and derived lipids, which include acylglycerols, fatty acids, sterols, and waxes. Mishina et al. (1977) observed that triacylglycerols are only one of several major classes in the total lipids of Candida tropicalis and C. lipolytica grown on n-alkanes. The total lipid contents were obviously low; about 4% of the dry biomass for C. lipolytica and 12-17% for C. tropicalis. According to Ratledge (1986), oleaginous microorganisms are designated as such when they produce at least 20% total lipids. There have been few studies on the effect of the chain length of the n-alkane substrate on the triacylglycerol content of microorganisms. Demanova et al. (1980a) showed that the total lipid content of Candida guilliermondii grown on n-octadecane (C18)was three times higher than when the cells were grown on n-docosane ( C Z 2 ]and , that the difference was mainly due to increased proportions of triacylglycerols and waxes.

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Little research has been done on acylglycerols of n-alkane-grown filamentous fungi. Siporin and Cooney (1975) observed that triacylglycerols are the only extracellular neutral lipid class detected in cultures of Cladosporium resinae grown in n-hexadecane, n-dodecane, and glucose. The constituent fatty acids of these extracellular triacylglycerols contain predominantly (more than 66%) lauric acid (12:0), in addition to lesser proportions of myristic (14:0),palmitic (16:0),and stearic (18:O) acids. This result is interesting in view of the fact that commercially, lauric acid commands higher prices than fatty acids with longer chains. Obviously, more work needs to be done on acylglycerols in n-alkaneutilizing microorganisms. Nevertheless, it appears that the presence of lipophilic compounds (alkanes) as substrates would direct the metabolic activity toward the production of more surfactive lipids. In other words, n-alkane-utilizing microorganisms may produce more partial acylglycerols, as mentioned earlier, and more phospholipids and glycolipids (see below), but not more triacylglycerols. VI. Sterols

Relatively little attention has been given to the study of sterols from nalkane-utilizing microorganisms, although this subject appears to have considerable biotechnological potential. According to Fukui and Tanaka (1980),n-alkanes might be an excellent carbon source for ergosterol production because a sufficient amount of acetyl-CoA and a lipophilic environment can be provided using this substrate. As a rule, prokaryotes have only a very limited capacity, if at all, to synthesize sterols. The only available report on sterols from alkanegrown bacteria is that of Sorkhoh et al. (1990b). These authors showed that n-dodecane-grown cells of Rhodococcus rhodochrous contain sterols, whereas glucose-grown cells lack this lipid class. The major sterol in the few yeasts and filamentous fungi studied so far is ergosterol. Thus, Shirota et al. (1970) found 1% ergosterol in nalkane-grown cells of Candida petrophilum, a value which was higher than the ergosterol content of glucose-grown cells. Tanaka et al. (1971) reported that C. tropicalis produces 70 mglliter ergosterol, and that nalkanes are better substrates than glucose. Sica et al. (1982) showed that ergosterol is the major sterol in n-alkane-grown cells of C. tropicalis. In addition, there are small amounts of (22E)-ergosta-5,7,9(11)22tetraen-3P-01, ergosta-7-en-3P-01, (22E)ergosta-7,22-dien-3P-ol, ergosta7,24(28)-dien-3@01,and cholesta-8,24-dien-3p-o1. Sica et al. (1984) also found that ergosterol is the major sterol in n-alkane-grown cells of C. lipolytica, in addition to small quantities of ergost-7-en-3/3-01,ergosta-

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7,22-dien-3/3-01, ergosta-7,24(28)-dien-3@-01,and ergosta-5,7,9(11),22tetraen-3P-01. Ergosterol is also the major sterol in n-alkane-grown Pichia spp. (Jeong et al., 1975). Lin et al. (1971a,b) reported that Cladosporium cladosporioides, Hormodendrum hordei, Aspergillus versicolor, and Penicillium lilacinum grown on n-dodecane and n-undecane contain larger proportions of ergosterol than when grown on glucose. Nakao et al. (1973) showed that n-alkane-grown Fusarium sp. excrete up to 260 mg/liter ergosterol into the medium. More total sterols are produced with short-chain than with long-chain alkanes as substrates (Mishina et al., 1977; Sorkhoh et a]., 1991). The role of n-alkane-utilizing microorganisms in biotransformation reactions of sterols and steroidal compounds is of particular biotechnological value. Microbial biotransformations have been repeatedly reviewed (Marsheck, 1971; Smith, 1974, 1984; Heftmann, 1975; Martin, 1977, 1984; Schoemer and Martin, 1980). Hence the objective of the present discussion is to underline the fact that many of the valuable biotransformations are in fact achieved by n-alkane-utilizing microorganisms. Pharmaceutically active steroids are produced either chemically or microbiologically. The microbiological production involves partial synthesis from naturally occurring steroids such as diosgenin, stigmasterol, p-sitosterol, campesterol, and cholesterol. One important biotransformation approach involves the elimination of a saturated aliphatic C-17 side chain of plant sterols leading to 17-ketosteroids or bisnorcholanic acid derivatives. Equally important is the partial degradation of the steroid ring system leading to hexahydroindan propionic acid derivatives which can produce steroids with unnatural configurations. Microorganisms involved in such biotransformations include those that are known for their activity in hydrocarbon utilization and oxidation, for example, the bacterial genera Arthrobacter, Brevi bacterium, Corynebacterium, Flavobacterium, Mycobacterium, Nocardia, and Rhodococcus, and the fungal genera Cunninghamella and Rhizopus. Reviewing the microbial conversion of n-alkanes and fatty acids, Ratledge (1984) observed that hydrocarbon-utilizing microorganisms can partially oxidize an even greater range of lipophilic compounds. Interestingly, metabolic pathways of steroidal biotransformations may be similar to those of nalkane utilization. Thus, during transformation of plant sterols to 17ketosteroids the C-17 side chain is shortened by C,, hydroxylation followed by oxidation to a C,, carboxylic acid, which is further shortened by a mechanism similar to &oxidation, leading finally to a C,,-keto compound. To substantiate these biotransformations, a few relevant studies that have been published in the past decade are summarized.

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Shah et al. (1980) studied the transformation of phyto-sterol to androsta 1,4 diene-3,17 dione by Arthrobacter simplex and suggested that this organism be explored further for the production of the C,, series of steroidal hormones on an industrial scale. Schoemer and Wagner (1980) transformed p-sitosterol by Nocardia sp. M29-40 to steroid-ringp-seco acid which is a valuable precursor in chemical steroid synthesis. Mulheim and van Eyk (1981) studied the oxidation of steroid hydrocarbons by numerous fungi and bacteria. In this context, Pseudomonas sp. NCIB 9872 grown on cyclopentanol is found to preferentially biotransform norbornanone to an equivalent lactone of potential use in the synthesis of prostanoid analogs (Sandey and Willetts, 1989). Jawoski et al. (1982) used spores of Cunninghamella elegans to hydroxylate cortexolone to cortisol and epicortisol. Nakamastu et al. (1983) used mutants of Nocardia carollina to convert 20 g/liter soybean oil into 2.8 g/liter HIL [3a-H-4a-(3’-propionic acid)-5a-hydroxy-7/3-methylhexahydro-lindanone-b lactone] which is an intermediate in the chemical synthesis of 19-norsteroids. A similar transformation was achieved by a mutant strain of Rhodococcus australis using cholesterol, stigmasterol, or psitosterol as starting materials (Ferreira et al., 1984). Spassov et al. (1983) investigated the transformation of pregnenolone triacetate to Reichstein-S-17a-acetate by Flavobacterium dehydrogenans and of hydrocortisone-17a-acetate to prednisolone-17a-acetate by Arthrobacter simplex. Arinbassova et al. (1985) found that the activity of 3ketosteroiddl-dehydrogenase in cells of Arthrobacter globiformis is the major factor that controls the direction of the biotransformation of hydrocortisone. Thus, under aerobic conditions, i.e., high enzymatic activity, the end product of hydrocortisone transformation is prednisolone or its 20P-hydroxyderivative. Microbial hydroxylation reactions have been also suggested for the biosynthesis of nonsteroidal, medicinally and technologically valuable compounds (Golbeck and Cox, 1984; Sakaki et al., 1987; Yoshida et al., 1990). For example, Yoshioka et al. (1990) suggested a Fusarium verticilliides-mediated hydroxylation of N-acetyl-0-toluidine to 4‘hydroxy-N-acetyl-0-toluidine,which is an important intermediate in the synthesis of thermosensitive dyes (Sat0 et al., 1989). VII. Fatty Alcohols, Ketones, and Epoxides

During the course of n-alkane oxidation to fatty acids, microorganisms may accumulate varying amounts of one or more of the reaction intermediates, either intracellularly or in many cases extracellularly, in the growth medium. Sometimes these intermediates are of higher commer-

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cia1 value than the n-alkane substrate and/or the corresponding fatty acid. Hence, effort has been devoted to biosynthesizing these compounds, not only from n-alkanes, but also from other natural products such as plant oils. Whether n-alkane-oxidation occurs terminally or subterminally, the immediate intermediates are primary and secondary alcohols, respectively. Further oxidation of secondary alcohols leads to ketones. Early investigators have observed the transient accumulation of fatty alcohols by n-alkane-grown microorganisms. Thus, in the course of a study on glutamic acid production by Arthrobacter parafineus from nparaffins, Suzuki and Ogawa (1972) noticed an accumulation of primary fatty alcohols in the paraffin layer with carbon chains corresponding in length to those of the n-paraffins used. Investigating the biodegradation pathway of n-dodecane and n-tridecane using several members of the mucorales, Hoffmann and Rehm (1976) observed in all cases the extramycelial accumulation of primary and secondary alcohols and ketones with carbon chains equivalent in length to those of the n-alkane substrate. Egorov et al. (1976) reported that six species of Mycobacterium, three strains of Pseudomonas sp., and Cladosporium resinae produced cetyl alcohol from n-hexadecane. The mycobacteria produced the largest amount of alcohol, about 120 pg/mg protein, or 2-4 mg/ml medium. Markovetz (1978) found primary alcohols, isomeric alcohols, and their corresponding ketones, diols, and epoxides among the intermediates of n-alkane and n-alkene oxidation using various bacteria, yeasts, and filamentous fungi. Fish et al. (1982) studied the induction of n-alkane hydroxylase activity in Pseudomonas putida with the goal of producing primary alcohols, particularly n-nonanol from n-nonane. The application potential of fatty alcohols involves the synthesis of vitamin E and K, from precursors such as pristanol (and pristanic acid, see Akutagawa et a]., 1978),which accumulates as an oxidation intermediate in cultures of Nocardia sp. BPM 1613 grown on the branched as a sole source of alkane pristane (2,6,10,14-tetramethylpentadecane) carbon (Nakajima et al., 1974; Nakajima and Sato, 1981).This stimulated interest in studying the fermentative bioconversion of pristane to pristano1 (Kuriyama et al., 1982; Kuriyama and Fukuoka, 1983). There are numerous patents covering the production of fatty alcohols with up to 20 carbon chains from alkanes using various species of the bacterial genera Acinetobacter, Arthrobacter, Brevibacterium, Corynebacterium, Micrococcus, Nocardia, and Pseudomonas and of the yeast genera Candida and Pichia. Methylotrophic bacteria are particularly active in the production of very short-chain alcohols and ketones. These patents have been tabulated by Buhler and Shindler (1984). In this context, it

n-ALKANE-UTILIZING MICROORGANISMS

65

should be mentioned that alkane-utilizing Candida yeasts have been successfully converted from the citrate to the polyol process, with the objective of producing polyhydroxy alcohols such as mannitol, erythritol, and arabitol from alkanes (Tabuchi and Hara, 1973; Hattori and Suzuki, 1974a,b,c). After realizing that methyl ketones produced by Penicillium roqueforti constitute the major material responsible for the special flavor of blue-veined cheeses (Kinsella and Hwang, 1976; Rothe et al., 1982), more attention was devoted to the microbiological production of fatty ketones as flavors and aromas in the food industry. The large-scale manufacture of food products such as salad dressings, soups, crackers, and cakes flavored with compounds simulating the blue cheese taste stimulated studies on the biosynthesis of methyl ketones exhibiting the natural aroma. However, in most of these investigations, free fatty acids or fatty acids esterified in plant oils are used as starting material. Kinderlerer (1987) described the conversion of coconut oil to methyl ketones by two species of Aspergillus. Yagi et al. (1989) reported the synthesis of keto-alkanes from fatty acid esters by Trichoderma. Larroche and Gros (1989) described the batch and continuous production of 2-heptanone from octanoic acid using entrapped spores of Penicillium roqueforti. Yagi et al. (1990) found that species of Penicillium, Aspergillus, Cladosporium, Fusarium, and Trichoderma are capable of accumulating methyl ketones in cultures containing synthetic tricaprin or palm-kernel oil. Epoxides are usually toxic; they are powerful alkylating agents. Hence, low-molecular weight epoxides are used in industry for sterilization purposes. Epoxides are produced microbiologically via oxygenation of alkenes; the enzyme-catalyzed epoxidation has been reviewed by May (1979) and Guengerich and MacDonald (1984). Examples of studies on low-molecular weight epoxides are those of Wingard et al. (1985) and Brink and Tramper (1986) who described the production of propylene oxide from propylene by immobilized cells of Nocardia carollina and Mycobacterium sp., respectively. Similarly, Hou et al. (1983) and de Smet (1983) reported on the epoxidation of short-chain alkenes by 16 species of bacteria belonging to the genera Actinobacter, Actinomyces, Arthrobacter, Brevibacterium, Corynebacterium, Mycobacterium, Nocardia, and Pseudomonas. The production of long-chain epoxides has also been considered. Schwartz and McCoy (1976) described the production of 7,8-epoxy-loctene, 1,2-7,8-diepoxyoctane, and 1,2-epoxyoctane from 1-octene by Pseudomonas oleovorans. Ohta and Tetsukawa (1978) reported on the synthesis of optically pure (R)( +)long-chain epoxides from the corre-

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SAMIR S. RADWAN AND NASER A. SORKHOH

sponding terminal olefins by Corynebacterium equi (IF0 3730). The same organism produced 1,2-epoxyhexadecane from 1-hexadecene (Ohta and Tetsukawa, 1979). Terminal olefins other than 1-hexadecene are also assimilated and the corresponding epoxides are produced from substrates with chains longer than C,,. VIII. Hydrocarbons and Waxes

Total lipid extracts of microorganisms usually contain only small proportions of hydrocarbons, unless they are grown on n-alkanes as substrates. Tornabene (1977), in his review on microbial formation of hydrocarbons, reported on the hydrocarbon contents of numerous bacteria, fungi, cyanobacteria, microalgae, and plankton. A process for the continuous conversion of glucose to hydrocarbons has been described by de Boks et al. (1982). It comprises the fermentation of glucose to ethanol, followed by the catalytic conversion of ethanol, in the presence of water over zeolite H-ZSM-5, to hydrocarbons. The microbial production of C, and C, hydrocarbons (Fukuda et al., 1984, 1985) and olefinic hydrocarbons (Fujii et al., 1985) has also been considered. Olama et al. (1990) found that Candida guilliermondii grown on an oil distillation fraction contains about 40% hydrocarbons in its total lipids. It is tentatively assumed that microbial cells growing on long-chain alkanes as substrates contain these alkanes as predominat constituents of their hydrocarbon fraction. Little experimental work on this subject has been done, and the few results recorded indicate that the previous assumption is valid for some, but not all, alkane substrates. Walker and Cooney (1973) studied the hydrocarbon composition of Cladosporium resinae grown on glucose, glutamic acid, and individual n-alkanes as substrates. Glucose-grown mycelia contain C,-C,, hydrocarbons; pristane (tetramethyl pentadecane) and n-hexadecane together make up 98% of the total. On glutamic acid the biomass contains C7-C,, hydrocarbons; n-tridecane, n-tetradecane, n-hexadecane, and pristane make up 74%. n-Decane (C,,)-grown mycelia contain C,-C,, hydrocarbons; n-hexadecane makes up 96%. Mycelia grown on individual n-alkanes from C,, to C,, contain C,,-C,, hydrocarbons, and mycelia grown on n-hexadecane contain C,,-C,, hydrocarbons. In C,,-grown mycelia, nhexadecane and pristane make up 92%; on C,,-C17 n-alkanes, the major mycelial hydrocarbon is the one which the organism utilized. The authors concluded that mycelia cultured on C,, or longer n-alkanes accumulate these compounds prior to their oxidation. Waxes result from the condensation of fatty acids and fatty alcohols; therefore they are expected to occur in considerable amounts in n-

n-ALKANE-UTILIZINGMICROORGANISMS

67

alkane-utilizing microorganisms. Indeed some authors reported on the increased synthesis of waxes as a major reason for the increased total lipid content in response to n-alkane utilization (Kvasnikov et a]., 1977b; Koronelli et al., 1978; Demanova et al., 1980a). Wax production by hydrocarbon-utilizing microorganisms has been known for more than three decades. Stewart et al. (1959) identified cetyl palmitate in Micrococcus cerificans (Acinetobacter sp.) grown on n-hexadecane. Raymond and Jamison (1971) reported on the wax composition of hydrocarbon-grown Nocardia spp. Krassilnikov et al. (1973, 1974) found that the wax of Mycobacterium ceroformans and M. lacticolum grown on n-hexadecane is almost pure cetyl palmitate, whereas the wax produced on media with n-heptadecane is a mixture of esters of several fatty acids, notably margaric acid. Bacchin et al. (1974) reported on didecyldecane-1,lO-dioate from n-decane-grown cells of Corynebacteriurn sp. 7ElC. Makula et a ] . (1975) identified several monoesters excreted extracellularly by Acinetobacter sp. grown on n-hexadecane and n-heptadecane. Koronelli et al. (1978) showed that various strains of Arthrobacter cereformans grown in an n-hexadecanecontaining medium produce high proportions of wax (up to 70% of the total lipids), with cetyl palmitate contributing to about 95% of the total wax. Knox and Cliffe (1984) studied the production of octyl oleate and cetyl oleate using the mycelia of Rhizopus arrhizus in a loop reactor consisting of a packed bed-glass column. Especially interesting are the mono- and diunsaturated waxes identified in Acinetobacter sp. HO1-N grown on C,, to C,, n-alkanes and which are similar to the waxes in jojoba oil (Dewitt et al., 1982).Jojoba oil is used in the cosmetic industry. IX. Phospholipids

Almost all studies that are concerned with phospholipid analysis report on increased concentrations of these compounds in response to microbial growth on n-alkanes as substrates. Phospholipids are major constituents of biological membranes and organelles. Hence, their increase in Acinetobacter species (Makula and Finnerty, 1970; Makula et al., 1975), for example, obviously relates to the induction of intracytoplasmic membranes, also associated with growth on n-alkanes (Kennedy and Finnerty, 1975b). Similar intracellular membranes are also observed in other bacterial genera, e.g., Rhodococcus (Ivshina et al., 1982). Yeasts utilizing n-alkanes produce special organelles, peroxisomes (Fukui and Tanaka, 1981),which probably contain phospholipids. However, increased phospholipid levels are also reported in many

68

SAMIR S. RADWAN AND NASER A. SORKHOH

bacteria and filamentous fungi which do not produce intracytoplasmic membranes or peroxisomes on n-alkanes as substrates. An acceptable explanation in such cases should consider the surfactive property of these amphipathic molecules. Additional phospholipids probably play some role in enhancing the uptake of n-alkanes by the cells. Some experimental evidence that may support this hypothesis has been provided by Miller and Bartha (1989), who studied the uptake of noctadecane (C18)and n-hexatriacantane {&) by Pseudomonas sp. Radioactively labeled substrates were encapsulated into bilayers (liposomes) of diacylglycerophosphocholines, and the uptake rates of encapsulated and unencapsulated alkanes were compared. The results indicated that the uptake rate of phospholipid-encapsulated alkanes was 18 times higher than that of the unencapsulated alkanes. There is some contradiction in the literature regarding the identity of a few phospholipid fractions in bacteria as well as in the composition of the phospholipid fatty acids (Tables XVII and XVIII). Makula and Finnerty (1970) reported that the predominant phospholipids of n-hexadecane-grown cells of Micrococcus cerificans (Acinetobacter sp.) are diphosphatidyl glycerols and diacylglycerophosphoglycerols. Hallas and Vestal (1978) found that diacylglycerophosphoethanolamines, diphosphatidyl glycerols, and diacylglycerophosphoserines are the predominant phospholipids in Mycobacterium convolutum grown on Cl,-C,, n-alkanes. Murphy and Perry (1987), also working on M. convolutum, but grown on n-hexadecane and 1chlorohexadecane, identified diacylglycerophosphoethanolaminesand diphosphatidyl glycerols (cardiolipins) as predominant phospholipids, but no diacylglycerophosphoserines; instead they identified diacylglycerophosphoinositolmannosides (Table XVII). Koronelli et al. (1982a) identified the major phospholipids of glucose- and n-hexadecane-grown cells of Pseudomonas aeruginosa as diphosphatidyl glycerols, diacylglycerophosphoethanolamines, diacylglycerophosphoglycerols, and diacylglycerophosphocholines. In all yeasts studied diacylglycerophosphocholines and diacylglycerophosphoethanolamines are the major phospholipids. Minor phospholipids are diacylglycerophosphoinositols and diacylglycerophosphoglycerols. Shigyo and Takeuchi (1972), studying cell lipids in relation to the nutritive value of hydrocarbon-grown yeast, found that phospholipids make up about 75% of total lipids and consist almost exclusively of diacylglycerophosphoethanolamines.Fodder yeast obtained from growth on purified liquid n-paraffins also contains the latter phospholipids (Goskaya et a]., 1978).These two phospholipids also predominate in Candida tropicalis and Candida Jipolytica grown on glucose or n-

TABLE XVII PHOSPHOLIPID COMPOSITION OF II-ALKANE-UTILIZING MICROORGANISMS~ ~

PC Microorganisms Acinetobacter HO1-N Acinetobacter sp. Acinetobacter sp. Micrococcus freudenreichii M. freudenreichii Mycobacterium convoluturn M. convolutum M. convolutum M. convolutum M. convolutum M. convolutum M. convolutum M. convolutum M. convolutum (30°C) M. convolutum (3OOC)

Substrate

PE

Total phospholipids

PS

PI

PIM

c16

Glucose n-Alkanes Acetate

C, c20 c22

c23 c24

c26 c2* c16

CI.C,,

f + C

46.0' 129.00 20.0% 28-46% 30% 34% 33% 44% 49% 51% 50% 52%

DPG

51.0 48.8 48.2 35.9 56.2 49.5 50.2 47.9 34.6 21.0

8.5 10.6 8.3 17.9 17.5 18.0 19.1 22.0 36.7 49.3

~

~

~~~~~

PA

oh of total phospholipids

Cl, Peptone

PG

Reference

++

37.7 35.0 37.2 19.5 21.5 25.5 22.5 23.7 28.6 30.2

Trace 1.8 1.4 1.5 Trace 1.0 Trace 1.1

Makula and Finnerty (1970) Makula et al. 1975) Makula et al. (1975) Kvasnikov et 01. (1974) Kvasnikov et 01. (1974) Hallas and Vestal (1978) Hallas and Vestal (1978) Hallas and Vestal 11978) Hallas and Vestal (1978) Hallas and Vestal (1978) Hallas and Vestal (1978) Hallas and Vestal (1978) Hallas and Vestal (1978) Murphy and Perry (1987) Murphy and Perry (1987) (continues)

TABLE XVII (Continued) PC

Microorganisms

u

0

M . convolutum (ZOC) M. convolutum (Zo"C] Pseudomonas aeruginosa Fodder yeast Fodder yeast Condido tropicolis C. tropicolis C. Iipolytico C. lipolytica Cladosporium resinaeb C. resinaeb

Substrate

++

++

++

51.0

49.0

+d

++ ++ ++ + +

PI

44.3 30.5

74.0%

++

PS

PIM

PG

DPG

Yo of total phospholipids

C16

CI.C,, Glucose or Clb n-Alkanes n-Alkanes Glucose n-Alkanes Glucose n-Alkanes Glucose C,, or C,,

PE

Total phospholipids

++

++

++

++ ++ +

+

++

++

++ ++ ++

+

+

+

+ +

+

+

+

Reference

30.4

26.2

52.1

17.4

++

+ + + +

PA

++

+ +

Murphy and Perry (1987) Murphy and Perry (1987) Koronelli et al. (1982a) Shigyo and Takeuchi (1972) Goskaya et al. (1978) Mishina et 01. (1977) Mishina et al. (1977) Mishina et 01. (1977) Mishina et al. (1977) Siporin and Cooney (1975) Siuorin and Coonev (19751

PC, diacylglycerophosphocholines; PE, diacylglycerophosphoethanolamines;PS, diacylglycerophosphoserines; PI, diacylglycerophosphoinositol; PIM, diacylglycerophosphoinositolmannosides;PG, phosphatydylglycerols; DPG, diphosphatidylglycerols; PA, phosphatidic acids. pg molig dry cells, otherwise Yo of dry biomass. ' Extracellular lipids. Major fraction. Not a major fraction.

TABLE XVIII CONSTITUENT FATTYACIDSOF TOTALPHOSPHOLIPIDS AND INDIVIDUAL PHOSPHOLIPID CLASSES FROM

Mycobacterium convolutum GROWN

ON n-ALKANES' ~~~

PE"

sn-1 Total phospholipidsb Fatty acids

U Y

13:O 14:O 14:l 15:O 15:l 16:O 16:l:

30°C

c,,

c,,

c,,

Trace

-

-

-

-

5.5

6.1

-

_

-

3.4

34.2

9.5

12.6

29.7

-

-

-

-

-

29.5 15.5

4.5 1.5

26.5 15.1

14.0 9.0

17.1 12.3

Acetate

c,,

3.4

c,,

-

6.4 28.5

-

4.5

-

in:i

26.7

18br 19 : 1 19br

-

1.5 3.8

2.7 29.7

-

12.4 4.3

-

17.0

6.1 12.4

20°C

30°C

c16

c16

20°C

30°C

Sn-2 20°C

30°C

20°C

30°C

20°C

cz

c16

-

1.9

-

-

18.8

3.2 10.8

-

5.3

Trace

Trace

4.4

-

-

-

-

Trace

1.1

1.1

-

-

-

-

-

-

-

-

-

25.9

20.5

33.6

26.5

10.6

17.7

42.7

26.4

55.5

52.1

-

5.8 2.0

0.0 1.1

8.3 14.5

3.1 3.1

37.6 4.1

1.7 11.0

31.6 24.6

6.6 1.6

6.2 5.8

56.1

52.2

40.5 1.3

67.7

30.3 1.7

-

-

Trace

Trace

6.7 1.1 3 .O

30.4

Trace Trace

24.5 1.2 4.8

28.7

1.8

-

-

5.9

12.9

4.5

4.0

Trace

4.0

3.0

_

-

42.0 3.0

44.8

7.5

9.7 4.3

4.4

-

3.0

-

1.0

-

15.4

5.3

9.4

6.6

5.4

-

-

10.0

24.4 5.7

3.8 4.4 1.9 3.5

2.6 7.6

-

54.2 6.9 3.8

4.3 5.8

8.3

6.6

7.7

Sn-1

Sn-2

c,,

9

11 17:O 17:l 17br 18:O

PIMC

CL'

-

9.7

c16

c16

c ~ 6

-

-

-

-

-

-

-

3.6

1.5

5.1

5.9

4.5

2.0

-

-

-

-

3.0

-

-

Trace

Trace

1.8

1.3

Trace

Trace

c16

3.4

c16

c16

c16

Data expressed i n relative percentage of total fatty acids. PE, diacylglycerophosphoethanolarnines;CL, cardiolipins; PIM, diacylglycerophosphoinositolrnannosides. From Hallas and Vestal (1978). From Murphy and Perry (1987).

72

SAMIR S. RADWAN AND NASER A. SORKHOH

alkanes; diacylglycerophosphoinositols and diacylglycerophosphoglycerols are minor components (Mishina et al., 1977). Although Makula et al. (1975) did not find any phospholipids produced extracellularly by n-hexane-grown cells of Acinetobacter s ~ . , Siporin and Cooney (1975) found several phospholipid classes, viz. diacylglycerophosphocholines, diacylglycerophosphoserines, diacylglycerophosphoethanolamines, and cardiolipins in the medium that supported the growth of Cladosporium resinae on glucose, n-dodecane, and n-hexadecane. Little work has been done so far on the effect of environmental variables on the phospholipid composition of n-alkane-utilizing microorganisms. Murphy and Perry (1987) found that Mycobacterium convolutum grown on n-hexadecane and 1-chlorohexadecane at 20°C contains larger proportions of diacylglycerophosphoethanolamines but lower concentrations of cardiolipins than at 30°C. The growth on 1chlorohexadecane as substrate at both temperatures is associated with increased proportions of diacylglycerophosphoinositolmannosides in the cells. A few studies have been published on the composition of constituent fatty acids in total phospholipids and in individual compounds from n-alkane-utilizing microorganisms. Makula and Finnerty (1972) demonstrated that the chain length of fatty acids in total phospholipids of ntetradecane-, n-pentadecane-, n-hexadecane-, and n-heptadecanegrown cells of Micrococcus cerificans (Acinetobacter sp.) reflects the chain length of the n-alkane substrates. Patrick and Dugan (1974) confirmed this result on Acinetobacter sp. using the substrates ntetradecane, n-hexadecane, and n-octadecane. Nonane-grown cells reportedly contain both odd- and even-chain fatty acids in their polar lipids. Growth on dotriacontcantaine (C3J, l-chlorohexadecane, 1chlorododecane, and 1-phenyldodecane is associated with the accumulation of significant amounts of odd-chain fatty acids in the polar lipids. In this context, it should be mentioned that 1-chlorohexadecane, as substrate, leads to the accumulation of chlorinated fatty acids in the total lipids as well as individual phospholipids of Mycobacterium convolutum (Murphy and Perry, 1987; see Table IX). This bacterium, when grown on C,, to C,, n-alkanes, does not incorporate any fatty acids with the corresponding chain length in its lipids (Table XVIII). Instead, the phospholipid fraction contains up to 44.8% pentadecanoic acid (15 : o), indicating that subterminal oxidation near the middle of the n-alkane chains may occur in this genus. (See also the patterns of fatty acids of total lipids from Mycobacterium sp. OFS in Table VII.) As should be expected, individual phospholipids from n-hexadecane-grown cells of

n-ALKANE-UTILIZING MICROORGANISMS

73

M. convolutum are poor in pentadecanoic acid (Hallas and Vestal, 1978; Table XVIII). However, this table shows contradictory data regarding acetate-grown cells (Murphy and Perry, 1987) and n-hexadecane-grown cells (Hallas and Vestal, 1978), particularly with respect to the fatty acids 17 : 1 and 17-branched. In the individual extra-mycelial phospholipids of Cladosporium resinae, the major constituent fatty acid is lauric acid (12 :0)(Siporin and Cooney, 1975). Phospholipids from Acinetobacter Iwoffi grown on nhexadecane contain predominantly palmitic (16 : o) and palmitoleic (16 : 1)acids in addition to small proportions of C,, fatty acids (Vachon et a]., 1982). The application potential of phospholipids from n-alkane-utilizing microorganisms is discussed later in section XI. X. Glycolipids and Peptidolipids

Glycolipids and peptidolipids are surfactive amphipathic substances that are frequently produced and excreted into the medium by n-alkaneutilizing microorganisms. The subject of biosurfactants is directly related to the uptake mechanism(s) of hydrophobic substances by microorganisms. It was early realized that the water-soluble fraction of alkanes is too low to satisfy microbial requirements for growth (Aibo et al., 1969). Hence, n-alkane utilization by microorganisms is made possible either by direct physical contact between the cells and the alkane droplets (Mimura et al., 1971; Miura et al., 1977; Neufeld et al., 1983) or by production of biosurfactants. These result in the formation of minute (
74

SAMIR S. RADWAN AND NASER A. SORKHOH

uptake by the cells because these compounds are sometimes intensively produced in the absence of n-alkanes (Cooper et al., 1982; Cooper and Paddock, 1983). Glycolipids produced by n-alkane-utilizing microorganisms contain mono-, di-, or polysaccharides as the hydrophilic moiety and usually fatty acids as the lipophilic moiety. Examples of low-molecular weight carbohydrate moieties are rhamnose (Syldatk et al., 19853, sophorose (Asmer et al., 1988), cellobiose (Frautz et a]., 1986), trehalose (Rapp et al., 1979; Kretschmer and Wagner, 1983), and mannosyl-erythritol (Kretschmer et al., 1982).Examples of polysaccharide-containing glycolipid biosurfactants are emulsan (Gutnick et al., 1981), liposan (Cirigliano and Carman, 1984), and a pentasaccharide lipid (Powalla et a]., 1989). Rhamnolipids are produced by Pseudomonas aeruginosa grown on n-alkanes such as n-hexadecane, or water-soluble substrates such as glycerol and glucose. The surfactant molecule contains either one or two rhamnose residues; the lipid moiety is a P-hydroxy or monoenoic fatty acid or fatty acid dimer. Pseudomonas aeruginosa 44T1 produces rhamnolipids when grown on n-dodecane but not any other n-alkane (Robert et al., 1989).The best yield is obtained with olive oil as a carbon source. Since phosphates inhibit rhamnolipid production, a biphasic fermentation process was suggested by Ramana and Karanth (1989). In the first phase, resting cells of P. aeruginosa CFTR-6 were produced in a phosphate-containing medium with glucose as a carbon source. In the second phase, the cells were incubated in the same medium without phosphate. Rhamnolipids were continuously produced by P. aerugipurity) was isolated nosa on a pilot plant scale and the compound by adsorption chromatography followed by anion-exchange chromatography (Reiling et al., 1986). Linhardt et al. (1989) suggested fermentatively produced rhamnolipids as a source of rhamnose. Hirayama and Kato (1982) isolated methyl rhamnolipids viz. L-rhamnopyranosylp-hydroxydecanoyl-P-hydroxydecanoic acid methyl ester and Lrhamnopyranosyl-L-rhamnopyranosyl-P-hydroxydecanoyl-~-hydroxydecanoic acid methyl ester, with cytotoxic effects. Sophorose lipids are produced by yeasts, such as Candida and Torulopsis, from n-alkanes or water-soluble substrates such as glucose. The lipid moiety is a hydroxy fatty acid that may be monounsaturated (Goebbert et a ] . , 1984). Trehalose lipids are produced by several members of the actinomycetes viz. Rhodococcus, Corynebacterium, Arthrobacter, Mycobacteriurn, and Nocardia grown on n-alkanes. Many of these biosurfactants are trehalose mycolates. Trehalose dimycolates, commonly known as

n-ALKANE-UTILIZING MICROORGANISMS

75

the cord factor, trehalose trimycolates, and trehalose tetraesters are produced by Mycobacterium smegmatis (Kilburn et al., 1982), Rhodococcus aurantiacus (Tomiyasu et al., 1986), and R. erythropolis (Ristau and Wagner, 1983; Kim et al., 1990), respectively. Interestingly, the triester isolated from R. aurantiacus contains up to C,, polyunsaturated mycolic acids. The tetraester may be tetrahalose corynomycolates (Kim et al., 1990),or the four acyl residues may be two decanoic, one octanoic, and one succinic acid (Ristau and Wagner, 1983). When resting cells of Arthrobacter sp. DSM2567 are incubated with mono-, di- or trisaccharides, they produce up to eight glycolipids containing the corresponding carbohydrate moiety and one, two, or three branched, p-hydroxy fatty acids (Li et al., 1984). Among these compounds were cellobiose and maltose monocorynomycolates, which reduced the interfacial tension from 42 to only 1 mN/m at critical micelle concentrations below 20 mg/liter. Obviously, many of the glycolipids produced by the actinomycetes contain a-branched, P-hydroxy fatty acids, i.e., mycolic acids. These fatty acids as such are surfactive (see Haferburg et al., 1986).Also, many of the peptidolipids produced by n-alkane-utilizing actinomycetes contain these unusual fatty acids. Batrakov et al. (1981a) isolated seven acyl-peptide derivatives from Mycobacterium paraffinicum; in three of them the peptide moieties consisted of L-valine, L-threonine, and Lleucine (2 : 1: I), whereas in the other four the peptide consisted of glycine, L-leucine, D-alloisoleucine, L-threonine, t-serine, L-homoserine, and D-alanine (3 : 3 : 2 : 2 : 2 : 1: 1). Five of these lipopeptides contain 0mycolyl substituents; three of them are glucosylated. Batrakov et al. (1981b) also isolated an N-acyltetrapeptide from M. paraffinicum with a mycolic acid residue. Koronelli et al. (1982b) isolated a pigmented peptidolipid from marine n-alkane-utilizing mycobacteria and found the lipid moiety to consist of usual and mycolic acids. The latter were predominantly C34 : 0 and C34 : 1. A well-known peptidolipid biosurfactant is surfactin, whose peptide moiety consists of seven amino acids bonded to the carboxyl and hydroxyl groups of the C,, acid (Arim et al., 1968). Interestingly, this biosurfactant is produced by Bacillus subtilis. As mentioned earlier, the genus Bacillus is not known to have n-alkane utilization activity. Mulligan et al. (1989) reported on activated surfactin biosynthesis in a mutant strain of B. subtilis. Cooper et al. (1979) observed that the surface activity of a medium supporting the growth of Corynebacterium lepus is due to corynomycolic acids at early growth phases, and to a peptidolipid containing corynomycolic acids at the end of fermentation. The peptide part consisted of alanine, glutamic acid, glycine, leucine, serine, threo-

76

SAMIR S. RADWAN AND NASER A. SORKHOH

nine, phenylalanine, aspartic acid, proline, methionine, isoleucine, valine, and lysine. Koronelli et al. (1983) characterized a peptidoglycolipid produced by Pseudomonas aeruginosa in a medium containing n-hexadecane. The peptide part consisted of lysine, aspartic acid, glutamic acid, serine, proline, valine, and leucine, and the sugar moiety of rhamnose. The lipid part was a mixture of fatty acids with 11-18 carbon chains, predominantly unsaturated. Reportedly, this biosurfactant stimulated nalkane assimilation by the bacterial cells. XI. Biolipid Extract

Biolipid extract is the single-cell “oil” obtained as a by-product during the production of fodder yeast grown on oil-distillation fractions. In order to rid the biomass of residual, not utilized hydrocarbons, the product is extracted with organic solvents, which also co-extract cell lipids (Bauch et a]., 1977). The biolipid extract has been extensively studied by scientists in “East” Germany and the “Soviet Union,” where hydrocarbon-based fodder yeast production is being achieved. The only comprehensive report published on these lipids is perhaps that of Voigt and Seidel (1990),which is written in German. Biolipid extract is a dark-brown, heavy fluid with a characteristic odor and high interfacial activity. Chemically this product consists (Voigt and Seidel, 1990) of 45.6% hydrocarbons, 20-30% phospholipids, 10-20% acylglycerols, 5-10’/0 fatty acids, 0.5-0.7% sterols, and about 0.1% coenzyme Qg. Hydrocarbons, the major constituent class, are obviously the residual, not utilized, substrate. Other lipid classes are cellular lipids. Because of the relatively high phospholipid content, the product is surfactive. The phospholipids consist predominantly of diacylglycerophosphocholines and diacylglycerophosphoethanolamines; minute amounts of lysophospholipids and cardiolipins are also present. Triacylglycerols predominate in the acylglycerol fraction. The constituent fatty acids of this product range in chain length between C,, and C,, (Table XIX). The relatively high content of the odd-chain fatty acid heptadecenoic acid (17 : 1) is characteristic. As mentioned earlier, most of the surface activity is because of phospholipids; the role of fatty acids in this respect appears to be negligible (Bormann et al., 1979). The applications of biolipid extract are largely based on its phospholipid content. Several techniques have been suggested for improving the properties of this product, and consequently its application potential. Thus, by

n-ALKANE-UTILIZING MICROORGANISMS

77

TABLE XIX CONSTITUENT FATTYACIDSOF

Fatty acids

BIOLIPIDEXTRACT^

Total fatty acids (Yo)

12:o

1.0

13:O 14:O

2.0 3.0 7.0 12.0 10.0 9.0 26.0 7.0 12.0 9.0 2.0

15:O 16:O 16: 1 17:O 17:l

18:O 18:l

ia:z 18:3

Results from Voigt and Seidel (1990).

reducing the water content and the concentration of low-boiling point constituents through evaporation, the odor of the product and its stability during storage can be improved. Through reduced pressure distillation, a part of the constituent hydrocarbons can be eliminated and thus a technically more useful product is obtained (Seidel et a]., 1988). This viscous, very dark brown fluid contains only 15-25% hydrocarbons; the proportions of other lipid classes are correspondingly higher than in the biolipid extract. Phospholipids in the hydrocarbon-poor residue can be precipitated with acetone and separated. The acetone-soluble fraction can be saponified with alcoholic KOH, the nonsaponifiable fraction can be extracted, and after treating the aqueous phase with mineral acids, the fatty acids can be precipitated in a rather pure form. Techniques for isolation of constituent lipid classes of biolipid extract have been described by Worbs et al. (1984) and Voigt et a]., 1984a,b). Many fields of application, e.g., in agrochemistry, mineral flotation and bitumen production and processing, have been suggested for biolipid extract (Voigt and Seidel, 1990). Some aspects of agrochemical applications are mentioned in this chapter. Potentially the product may be used as an emulsifying and dispersing agent during formulating herbicide, pesticide, and growth regulator preparations. Inclusion of phospholipids in these preparations facilitates penetration of the active substance into the plant tissues (Holz, 1985),making it possible to apply only very low concentrations of these substances (Seidel et a]., 1990). The constituent fatty acids of biolipid extracts have antiphytoviral and

78

SAMIR S. RADWAN AND NASER A. SORKHOH

antifungal activities, and therefore can be applied in controlling plant diseases (Voigt et al., 1985). These fatty acids also increase the stress tolerance of plants, leading to higher yields after seasons of physiological drought (Bergmann et al., 1987).

XII. Environmental Considerations

Crude oil and its technical derivatives are currently global environmental pollutants of marine, terrestrial, and atmospheric habitats. Microbial hydrocarbon degradation is the most important natural process for removing these pollutants. Microbial seeding and environmental modification (bioremediation) are management options that are in part currently in operation. These and related environmental aspects have been the subject of several reviews (Atlas, 1978, 1980, 1981; Cane et al., 1983; Bartha, 1986). Even halogenated hydrocarbons, which are particularly hazardous synthetic products, can frequently be dehalogenated by oil-utilizing microorganisms (Omori and Alexander, 1978a,b; Yokota et al., 1986; Vandenbergh and Kunk, 1988). Nevertheless, although the major part of contaminating oil is relatively rapidly biodegraded, some constituent compounds can persist for a long time (Schaeffer et al., 1979; Oudot et al., 1981). Through genetic manipulation (Williams, 1978; Fennewald et al., 1978; Fall et al., 1979; Porits et al., 1983; Amund, 1984; Painceira and Molo, 1988),efforts are being continuously devoted to creating microbial strains with a wide range of biodegradation activity that can be applied to control oil pollution (for reviews, see Devereux and Sizemore, 1981; Singer and Finnerty, 1984; Chakrabarty, 1985). Some of these mutant strains have been patented ( e g , Hala, 1985; Vandenbergh, 1990). However, there is reason to believe that the application of such mutant strains in nature would face and even cause a number of ecological and environmental problems (Radwan, 1991b). Experience with attempts to seed other microorganisms, e.g., nitrogen fixers, show that the inoculated cells may fail to establish themselves in the prevailing ecosystem. If, however, seeded oildegrading microorganisms would succeed in overcoming the competition stress exerted by the indigenous strains, their activity may pave the way for new, unpredicted problems. As mentioned earlier, hydrocarbon-utilizing microorganisms occasionally excrete a variety of biosurfactants, whose impact on the environment has not been adequately studied. That the natural microbial equilibrium would be seriously distorted in the concerned ecosystem by these substances (for review, see Traxier et al., 1983) is just one of the least hazardous effects. It

n-ALKANE-UTILIZING MICROORGANISMS

79

is tentatively assumed that biosurfactants, being natural products, are biodegradable, and consequently environmentally safe. Yet, it has been frequently demonstrated that biosurfactants produced by a given hydrocarbon-utilizing isolate may dramatically inhibit another hydrocarbon-utilizing isolate. Thus, a lactonic sophorolipid produced by Torulopsis bombicola was found to inhibit 19 species of Candida, Pichia, Debaryomyces, Saccharomycopis, and Lodderomyces, and only when these organisms were grown on n-alkanes as sole carbon sources [Ito et al., 1980). Similar results were recorded using other microorganisms and biosurfactants (Nakahara et a]., 1981; Pines and Gutnick, 1986; Foght et al., 19891, although in a few cases, stimulatory effects were also observed (Goclik et a]., 1990). These results clearly show that the application of biodetergents and/or biodetergent-producing microorganisms for the purpose of controlling oil pollution may potentially result in inhibiting the natural population involved in this process. Another suggested field of application for biodetergents and biodetergent-producing microorganisms is enhanced oil recovery. Such suggested applications also may be associated with unpredictable environmental problems. The general principles of microbial recovery methods involve processes such as microbial decomposition of nonhydrocarbon organic matrix of the rock, dissolution of the sedimentary rocks, detergent action on the rocks, gas-drive and increased mobility of the oil, and others. Several reviews have been published on the potential use of microorganisms and biosurfactants in enhanced oil recovery (e.g., Zajic et al., 1983; Grula and Grula, 1983; Finnerty and Singer, 1983; Gutnick, 1984; Moses, 1984,1987). There are also patents covering such applications (e.g., McInerney et a]., 1985). As already shown, many of the glycolipid biosurfactants and peptidolipids produced by the actinomycetes contain mycolic acids, which also are surfactive. There are also monoacylglycerols produced by actinomycetes, in which the acyl moieties are mycolic acids (Ioneda and Lopes Silva, 1979; Lopes Silva and Ioneda, 1980). Before discussing the potential impact of mycolic acids on the environment, a brief introductory word concerning these unusual fatty acids is given. Mycolic acids are a-alkyl, p-hydroxy very long chain fatty acids; they are considered the most characteristic cell wall component of some actinomycetes. Mycolic acids contribute to some characteristics of the cell, such as acid fastness, hydrophobicity, adherability, and pathogenicity. Mycobacterium possesses mycolic acids with the longest chains (C60-C80),whereas mycolic acids from Nocardia (C4,+&), Rhodococcus [C34-C46),and Corynebacterium [CZz--C3Jare shorter (see Goodfellow and Minnikin, 1980).

80

SAMIR S. RADWAN AND NASER A. SORKHOH

There is reason to believe that enriching waters and soils with longand short-chain mycolic acids may be potentially hazardous. Trehalose mycolates produced by oil-degrading actinomycetes may contain mycolic acids with up to 74 carbon atoms (Tomiyasu et al., 1986). The relation of such very long-chain mycolic acids to pathogenesis of Mycobacterium tuberculosis has been demonstrated (e.g., Toubiana et al., 1979; Khuller et al., 1982). Daffe et al. (1988) reported trihalose polyphthienoylate as a specific glycolipid in virulent strains of M. tuberculosis. Phthienoic acids constitute a family of dextrorotary odd-numbered unsaturated fatty acids. Kaneda et al. (1986) found that granuloma formation and hemopoiesis could be induced by C,,-C,, mycolic acidcontaining glycolipids from Nocardia rubra. Granuloma formation is also induced by mycolic acid-containing glucose and trehalose glycolipids isolated from more than 10 strains of Nocardia, Rhodococcus, and related actinomycetes (Sawai et al., 1987). Glycolipids containing mycolic acids longer than C,, have a stronger granulomagenic activity in mice than glycolipids with shorter chains (Yano, 1988). Not only glycolipid biosurfactants from actinomycetes, but also those from other bacterial taxa may be of public health concern. Methyl rharnnolipids from Pseudornonas aeruginosa have cytotoxic effects (Hirayama and Kato, 1982). Lipopolyglycans from mycoplasmas have endotoxic properties, potentially inducing procoagulant activity in human leukocytes (Miragliotta et al., 1987). The toxicity and antigenic properties of mycobacterial glycolipids are known (Jardine et al., 1989). In this context, it should be mentioned that potentially pathogenic mycobacteria such as Mycobacteriurn aviurnintracellulare, M. scrofulaceum, and M. fortulitum occur in water polluted with industrial and domestic residues (Cardoso and Filho, 1979). The structure of mycolic acids in Mycobacteriurn (Toriyama et al., 1980; Toriyama, 1982) and Nocardia (Tomiyasu et al., 1981;Tomiyasu, 1982) changes according to the prevailing environmental conditions, such as temperature and medium composition. These results are adequate to demonstrate that unpredictable environmental hazards may be associated with extensive application of biosurfactants and biosurfactant-producing microorganisms in controlling oil pollution and in enhanced oil recovery. This subject merits systematic study. ACKNOWLEDGMENTS Sincere thanks are due to Professor Dr. H.-J. Rehm for his interest and for critically reading the manuscript. The authors’ work cited in this review has been supported by Kuwait University, Research Grant No. SO 042 and SO 049. Samir S. Radwan is the recipient of an Alexander von Humboldt stipendium, October-December 1990.

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Microbial Pentose UtiIizatio n PRASHANT

MISHRAAND AJAYSINGH

Biochemical Engineering Research Centre Indian Institute of Technology New Delhi 110 016, lndia

I. Introduction 11. Pentoses from Natural Sources 111. Pentose-Fermenting Organisms A. Yeast B. Filamentous Fungi C. Mesophilic Bacteria D. Thermophilic Bacteria IV. Pentose Metabolism A. Yeast and Fungi B. Bacteria V. Production of Solvents and Organic Acids A. Ethanol Production B. Acetone and Butanol Production C. 2.3-Butanediol Production D. Organic Acids Production VI. Factors Affecting Pentose Fermentation A. Ethanol Production B. Acetone and Butanol Production C. 2,3-Butanediol Production D. Organic Acids Production VII. Product Tolerance VIII. Strain Improvement IX. Future Prospects References

I. Introduction

An interest in transforming vast reserves of renewable plant biomass to fuel and chemical feedstock was generated by the oil crisis of the mid 1970s (Rosenberg, 1980). This approach has led researchers to direct technologies toward the utilization of lignocellulosic biomass from crop residues, forest product residue, as well as other industrial waste carbohydrate streams (Vallander and Eriksson, 1990). Another issue that arises in this context is that estimates of conventional feedstock costs for the production of fuel and chemicals are in a conversion range from 30 to 70% of the product selling price (Schneider, 1989a,b). Thus much of the current interest has been focused on the processes that are based on cheaper cellulosic and hemicellulosic feedstocks.

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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 operates at lower temperature and produces specific products in high yields with fewer by-products (Stewart et al., 1984). It is possible to generate useful products from wastes by the effective recycling of materials with the consequent reduction of overall process cost using fermentation technology (Volesky and Szczesny, 1983). Any waste stream containing carbohydrates can be utilized for the generation of useful chemicals (Schneider et a ] . , 1983). Crops, trees, and other plants that are grown for food and other economic purposes also generate hundreds or thousands of millions of tons of lignocellulosic wastes. Most of these wastes are not utilized properly. One great disadvantage lies in the fact that since these wastes are generally in a solid form and are spread thinly over the land surface, the cost of transportation is too high, making this a prohibitory factor (Hahn-Hagerdal et al., 1991). However, there are some convenient waste streams generated from manufacturing processes such as those from pulp and paper industries, wood-based industries, and food 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 (Lovitt et a]., 1988). When the alternative technologies of oil-based feedstocks are compared, biologically produced chemicals compare favorably with the chemical industry (Wiegel, 1980). Fermentation involves simpler technology, and the by-products are mostly nontoxic, unlike those from chemical plants (Palson et al., 1981). A fermentation plant can be smaller and dispersed to meet social needs; nevertheless, these feasibilities are still not economical with present oil prices. 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. The economics of bioconversion would be more feasible if both hexose and pentose sugars present in lignocellulosic materials could be utilized (Skoog and Hahn-Hagerdal, 1988). While the technology employing yeast and bacteria to make chemicals from hexoses is well known, the ability of these organisms to ferment pentoses has been considered problematical (McCracken and Gong, 1983; Kurtzman, 1983). The pentose sugars in question are D-xylose and L-arabinose, which comprise up to 30% of the neutral carbohydrates derived from

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agricultural crop residues, wood, and other plant materials ( Jeffries, 1983; Gong, 1983). These sugars, along with D-glucose, D-mannose, and D-galactose, are derived from cell wall middle lamella that are often referred to as “hemi-cellulose.” Hemicelluloses and hemicellulosederived carbohydrates have many potential uses. Hemicellulose can be converted by microorganisms to various value-added products such as ethanol, sugar alcohols, solvents, and organic acids (Lovitt et a]., 1988). The nature and amount of conversion products depends on the type of pentose sugar, metabolic efficiency of the microorganism, and cultural conditions employed. This chapter examines recent microbiological, biochemical, and biotechnological findings in relation to our previous understanding of pentose fermentation, metabolism, and utilization. It reviews the regulatory aspects of pentose metabolism, genetic modifications, and improvements of pentose fermenting microorganisms. A detailed account of production of various solvents and organic acids is provided along with suggestions for future research. II. Pentoses from Natural Sources

Wood and agricultural by-products, the most abundant renewable resource materials, are virtually inexhaustible based on the photosynthetic process (Hall and Slessor, 1976). These by-products are composed of three major components: cellulose, hemicellulose, and lignin. Cellulose and hemicellulose are found in the secondary wall of the cell. Crystalline microfibrils of cellulose are surrounded by amorphous hemicellulose, and the whole is embedded in a matrix of lignin (Aspinall, 1959). Cellulose, the chief constituent of the cell wall, is a linear polymer composed of D-glucose units linked with P-glycosidic bonds. Unlike the orderly crystalline structure of cellulose, hemicellulose exhibits variability in both structure and sugar constituents. Hemicelluloses are composed of neutral sugars, uronic acids, and acetal groups, all present as their respective anhydrides, i.e., xylan, araban, glucan, galactan, and mannan (Table I). The common hemicellulose components are Dxylose, D-glucose, D-mannose, L-arabinose, D-galactose, 4-O-methylD-glucuronic acid, and occasionally a few other sugars. As the anhydrides, hemicellulose averages about 26% of hardwood, 22% of softwood, and about 30% of various major agricultural residues (Brasch and Wise, 1956; Rydholm, 1965; Krull and Inglett, 1980; Gordon et al., 1983). The content of hemicellulose sugars varies greatly with the plant species (Table I). The xylan content of hardwood is generally much higher (11-25%) than that of softwood (3-8%). In hardwood, P(1 + 4)linked xylan contains side chains of 4-O-methyl-~-glucuronicacid and

PRASHANT MISHRA AND AJAY SINGH

94

TABLE I

DISTRIBUTIONOF HEMICELLULOSICS IN LIGNOCELLULOSIC RESIDUES'

Percent of total dry weight ~~~

Residue

Hemicellulose sugars

Glucan

Galactan

Mannan

Arabinan

Xylan

Agricultural Hardwoods Softwoods

25 26.2 22.3

36.5 50 46

1.2

0.8 1.4

1.1 2.5 11.2

2.5 0.5 1.0

16.2 17.4 5.7

~~

Adapted from Jeffries (1983).

acetyl groups, whereas glucomannans are the predominating hemicellulose in softwood. Glucomannans are the linear polymer of p ( 1 + 4)linked glucose and mannose. The xylan content of soft wood is lower than hardwood, and the lignin content is higher. In conifers, the predominant hemicellulosic sugar is D-mannose, which as mannan forms about 11%of the total dry weight (Sjostrom, 1981). D-Xylose is the predominant hemicellulose sugar of deciduous trees and agricultural residues. Pectic polysaccharides, mainly composed of D-galacturonic acid and L-arabinose are usually not isolated in hemicellulose fractions (Volesky and Szczesny, 1983). For a detailed structure and chemistry of hemicelluloses, readers are referred to an excellent review by Wilkie (19 79).

Whether used as an energy source or as chemical feedstock, pretreatment of the substrate is necessary for the optimal utilization of lignocellulosic materials. The processes presently available include high temperature, pressure, and chemical treatment. These processes produce hydrolysates containing a variety of carbohydrates (fermentable or nonfermentable) and lignin-derived compounds. These compounds may be inhibitory to the microorganisms and consequently to the fermentation processes. One of the best examples is spent sulfite liquor from the pulping and paper industries. Pentose sugars are easily recovered from lignocellulosic substrates but are difficult to ferment. Both from economical and environmental points of view this fraction must be fermented to derive useful chemicals. As such the cost estimates for lignocellulosederived solvents and chemicals indicate the significant impact of the price of raw material (Hahn-Hagerdal et a]., 1991).

I l l . Pentose-Fermenting Organisms Views on the ability of yeasts to ferment aldopentoses were contradictory for a long period of time, although many of the microbial species known are able to assimilate D-xylose under aerobic conditions. How-

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MICROBIAL PENTOSE UTILIZATION

ever, some early studies have shown that a yeast could carry out the fermentation of D-xylose and L-arabinose (Plevako and Czeban, 1935; Karczewska, 1959). During the last decade, a number of laboratories have demonstrated the solvent and organic acid production of pentose sugars using different strains of yeasts, molds, and bacteria. Microorganisms differ in their ability to grow and metabolize aldopentoses. The demonstration of product accumulation from pentoses is sensitive to a number of experimental conditions which might have been reflected in the origin of different views regarding the feasibility of pentose fermentation. General characteristics of pentose fermenting organisms are discussed here and in Table 11. TABLE I1 GROWTH CHARACTERISTICS OF PENTOSE-FERMENTING ORGANISMS Sugar type’ Organism Yeast Saccharomyces cerevisiae Klyuveromyces marxianus Pachysolen tannophilus Candida shehatae Pichia stiptis

Glu

+

+ + + +

Xyl Ara

-

+ +

+

+

-

+ +

+

+

Temperature optimum

Man Cel

+ +

-

-

-

+

+

(“C)

-

30-35 30-35 28-32 28-32 28-32

3-7 3-7 2.5-7 3-7 3-7

28-34 28-37 26 30 30-37

5-6 5-6 5 5.4 2.2-7

5.5-8 6-8 5.5-8 5-6 4-8

-

-

Filamentous fungi Fusarium oxysporurn Neurospora crassa Monilia sp. Mucor sp. Paecilomyces sp.

+ + + + +

+ + + + +

+ +

+ + - + - + - + -

Mesophilic bacteria Bacillus polymyxa B. macerans Aerobacter hydrophila KJebsieJla pneurnoniae Clostridiumacetobutylicum

+ + + + +

+ + + + +

+ + + +

+

+ + + + +

+

35-37 35-37 35-37 35-37 30-37

Thermophilic bacteria Clostridium thermocellum C. thermohydrosulfuricum C. therrnosaccharolyticurn C. therrnosulfurogenes Thermoanaerobocter ethanolicus

+ + + + +

+ + + + +

+ + + + +

-

+

65 65 60 60 69

a

+ +

PH optimum

-

-

Glu, glucose; Xyl, xylose; Ara, arabinose: Man, mannose: Cel, cellulose

4-8 4.7-8 5-8 4.5-7.5 4.4-9.5

96

PRASHANT MISHRA AND AJAY SINGH

A.

YEAST

The applicability of yeasts to convert carbohydrates to ethanol has been studied extensively. Many of the yeast species are known to ferment hexoses (mainly D-glucose) to ethanol with significantly high yields. However, only a few yeast strains have been reported so far that can ferment pentoses to ethanol, even though many can metabolize pentose aerobically for growth. Many yeast strains have been investigated with reference to their capacity for the utilization of xylose and xylulose. With respect to xylose fermentation, yeasts that have been studied extensively are Pachysolen tannophilus, Candida shehatae, Pichia stiptis, and Kluyveromyces marxianus. Other yeasts investigated for their xylose-fermenting ability include Brettanomyces, Clavispora, Schizosaccharomyces, several species of Candida viz. C. tenuis, C. tropicalis, C. utilis, C. blankii, C. friedrichii, C. solani, C. parapsilosis, C. sake, and species of Debaryomyces viz. D. nepalensis and D. polymorpha. However, none of these yeast species have been found to be very promising. Toviola et al. (1984) screened 200 strains of yeasts by employing the classical method of measuring gas production as an indicator for fermenting strains. However, most of the strains selected as fermenting pentoses produced only small amounts of ethanol, and the authors concluded that criteria of gas production for screening pentosefermenting yeasts was not reliable. Schneider et al. (1981)reported that approximately 20% of the yeast species that give a positive Durham tube test on D-glucose produce only about 1 g/liter of ethanol from 20 g/liter of D-xylose. Several yeasts that tested positive under these conditions were previously not identified as xylose-fermenting strains. Gong et al. (1983) tested 20 strains of Candida, 21 strains of Saccharomyces, and 8 strains of Schizosaccharomyces for their ability to ferment D-xylose. All of the Candida strains grew well on D-xylose, while the Saccharomyces and Schizosaccharomyces strains grew poorly or not at all. Xylitol was produced in the range of 10-50°/0 by Candida strains with arabitol as the second major product. Ethanol was found to be the major product with most of the Candida strains. Schizosaccharomyces strains also produce ethanol but the concentrations are low (1-5 g i liter). Different aeration conditions have been employed in the various surveys designed to identify strains capable of xylose utilization. Roughly half of the yeast strains tested utilize D-xylose for growth under aerobic conditions but not under anoxic conditions (Barnett, 1976). Maleszka and Schneider (1982) screened 15 yeasts for their ability to utilize

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97

D-xylose, D-xylulose, and xylitol for ethanol production under aerobic, semiaerobic (low aeration), and anaerobic conditions, with rich undefined or defined media. In almost all cases, ethanol production by P. tannophilus and species belonging to Candida and Pichia was better on rich media under semiaerobic conditions. Thus it was concluded that the experimental condition that is critically important in the screening of pentose-fermenting cultures as well as in determining the extent to which ethanol accumulates in cultures is the oxygen supply rate. On the one hand, very low concentrations of ethanol are produced when high aeration rates are provided (Jeffries, 1981; Watson et al., 1984a; Schneider, 1989b). On the other hand, all the yeasts tested to date failed to grow anaerobically on D-xylose to any considerable extent (Baillargeon et al., 1983; Sreenath et a]., 1986). The majority of yeasts cannot ferment D-xylose directly. It has been observed that instead they utilize D-xylulose, an isomer of D-xylose,both oxidatively and fermentatively. The best xylulose-fermenting yeasts so far identified are species of Brettanomyces, Candida, Hansenula, Kluyveromyces, Pachysolen, Saccharomyces, Schizosaccharomyces, and Torulospora (Skoog and Hahn-Hagerdal, 1988). The pattern of Dxylulose utilization has been observed to be different from that of Dxylose utilization. D-Xylulose can be readily utilized by Candida under aerobic or semiaerobic conditions (Gong et a]., 1983).Ethanol concentrations from D-xylulose are generally higher, and xylitol accumulation is generally lower than when D-xylose is used as substrate. The pattern of D-xylulose utilization by different strains of Saccharomyces is also similar to that of Candida. In contrast, Maleszka and Schneider (1982) found that P. tannophilus, which ferments D-xylose, less readily utilized D-xylulose, while C. blankii, which readily assimilates D-xylose, also produces ethanol from D-xylulose (Gong et al., 1983). Another example is provided by S. pombe which does not assimilate D-xylose but readily utilizes D-xylulose aerobically or anaerobically. Many yeasts can convert xylose to xylitol, even though they are not able to grow on pentose (Jeffries, 1982, 1984). Xylitol, for example, is generally a poor substrate, even though it is an essential intermediate in the D-xylose catabolic pathway. Maleszka and Schneider (1982)tested 15 yeast strains and reported only traces of ethanol produced from xylitol as a carbon source. Only Torulopsis candida was found to have potential out of the 16 strains tested by Barnett (1968). Pentitols are the major by-products of pentose fermentation, and ethanol production efficiency would be further increased if such compounds could be converted to ethanol. Pachysolen tannophilus utilizes xylitol but the conditions necessary differ from those of D-xylose. Pichia angophorae has

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PRASHANT MISHRA AND AJAY SINGH

been identified as a potential organism for polyol fermentation (Lee and Schneider, 1987). The direct conversion of the hemicellulose polymer to ethanol is rare among the yeasts. Of the 250 yeast strains screened, few strains of P. stiptis and Candida shehatae have been identified to simultaneously hydrolyze and ferment xylan to ethanol. This ability is of particular interest in developing a direct bioconversion process. B. FILAMENTOUS FUNGI

The ability of filamentous fungi to ferment sugars has been known for approximately 70 years (Anderson, 1924; Letcher and Willmann, 1926). However, detailed studies have only recently been undertaken at several laboratories throughout the world. Several fungal species belonging to the genera Fusarium (Perlman, 1950; Cochrane, 1958; Rosenberg et al., 1981; Enari and Suihko, 1984; Christakopoulos et a]., 1989; Singh and Kumar, 1991), Rhizopus (Perlman, 1950), Monilia (Gong et al., 1981a) Neurospora (Deshpande et al., 1986), and Paecilomyces (Wu et al., 1986) have been found to have potential for fermenting glucose as well as xylose. Fusarium oxysporum (lini) was found to ferment glucose to ethanol and CO, giving a product yield similar to that of yeast (Anderson, 1924). The observation that the fermentation of xylose, although giving a comparatively lower ethanol yield, but still appreciable, was made and led scientists to use this organism for fermenting wood wastes (White and Williams, 1928). Suihko and Enari (1981) screened 26 strains of F. oxysporum for their ability to convert D-glucose and D-xylose to ethanol. Except for F. oxysporum VTT-D72014, all Fusarium strains are capable of fermenting both of the sugars. Wene and Antonopoulos (1988) screened over 2000 Fusarium strains, isolated from natural substrates, for their ability to utilize D-xylose. Almost all of the strains are capable of producing some ethanol, ranging from 0.4 to 4.4 g/liter ethanol to 1 0 g/liter D-xylose. In addition to D-xylose, many other sugars found in the hemicellulose complex, such as D-glucose, D-mannose, D-galactose, and also xylitol and sucrose, are fermented by Fusarium species (Suihko, 1983). The fermentation products formed by Fusarium sp. from pentoses or hexoses are similar, consisting of equimolar quantities of ethanol and acetic acid (Gibbs et a]., 1951). Paecilomyces sp. NF1 was found to be able to convert all the major sugars present in the hydrolysate of plant biomass (Wu et al., 1986). High ethanol concentrations (76 g/liter) by fermentation of D-xylose can be obtained using this fungus. Yields and rates of ethanol production

MICROBIAL PENTOSE UTILIZATION

99

from D-xylose by Mucor species (Ueng and Gong, 1982) and Neurospora crassa (Deshpande et al., 1986) have generally been low when compared to F. oxysporum (Singh et al., 1992a). Some potentially useful fungal strains have been identified that ferment not only glucose and xylose but also more complex natural cellulosic substrates. Monilia sp. (Gong et al., 1981b), N. crassa (Rao et al., 1983; Deshpande et a]., 1986), and F. oxysporum (Antonopoulos and Wene, 1984; Christakopoulos et al., 1989, 1990; Kumar et al., 1991a,b) have the potential for the direct conversion of cellulose/hemicellulose to ethanollacetic acid in a single step. In this case ethanol production is achieved by placing aerobically grown mycelia under anaerobic conditions. There are, however, several physiological parameters inherent to these fungi which makes the fungal ethanol process unattractive. These include (a) a long generation and fermentation time, ranging from 4 to 8 days; (b) low volumetric rates of product formation; (c) growth in large clumps rather than as dispersed, single cells; (d)critical oxygenation levels; (e) production of acetic acid along with ethanol as a major byproduct, which results in the lower ethanol yield; and (f ) low substrate and product tolerance. However, a fungal system might be of interest because of its ability to grow on natural plant biomass which yeast systems usually lack. C. MESOPHILIC BACTERIA

Although most yeasts and filamentous fungi cannot ferment pentoses anaerobically, many bacteria readily convert xylose to a variety of products in the absence of oxygen (Rosenberg, 1980).These include Bacillus macerans, B. polymyxa, Klebsiella pneumoniae, Clostridium acetobutylicum, Aeromonas hydrophila, Aerobacter sp., Erwinia sp., Leuconostoc sp., and Lactobacillus sp. (Rosenberg, 1980; Gonget al., 1983;Detroy and Bolen, 1983). A variety of products are formed by the bacterial fermentation of pentoses. The rates, yields, and products formed by these bacteria depend not only on the diverse metabolic pathways operating during anaerobic fermentation but also on the species, strains, substrates, and cultural conditions used. Bacterial strains of Lactobacillus sp. and Leuconostoc sp. convert D-xylose to acetic acid, ethanol, and CO,. Bacillus macerans ferments D-xylose into a mixture of ethanol, acetic acid, and acetone, whereas A. hydrophila produces a mixture of ethanol and 2,3-butanediol from D-xylose (Flickinger, 1980). Strains of K. pneumoniae produce ethanol and/or 2,3-butanediol (Jansen and Tsao, 1983; Banerjee, 1985). In general, species belonging to the genera

100

PRASHANT MISHRA AND AJAY SINGH

Bacillus, Aerornonas, and Kiebsielia produce more ethanol and butanediol and less acid. The proportions of the solvent and acid are greatly influenced by the pH of the fermentation broth. Some of the physiological characteristics inherent in these organisms include (a) the fermentation of pentose sugars without the need for critical oxygenation, unlike yeast and fungi; (b) a generally shorter generation time which is reflected in the high conversion rate during fermentation; and (c) utilization of a lesser proportion of substrate carbon for biomass formation, hence promising the possibility of obtaining a higher specific product yield. Some mesophilic bacterial species have the ability to utilize a broad range of substrates, including hexoses and pentoses. For example, K. pneurnoniae, which is easy to cultivate, grows rapidly in simple media and can metabolize a wood hemicellulose hydrolysate containing D-glucose, D-mannose, D-cellobiose, D-xylose,Dgalactose, and L-arabinose (Anderson and Wood, 1962; Yu and Saddler, 1982). Another gram-negative facultative anaerobic bacterium, Erwinia chrysanthenn, has also been found to utilize the sugars present in a lignocellulose substrate with a higher ethanol tolerance capability (Tolan and Finn, 1987).

D. THERMOPHILIC BACTERIA

Thermophilic bacteria are best defined by their temperature characteristics for growth. These bacteria ferment a wide range of substrates, including cellulose, hemicellulose, pectin, and starch. Different products formed by these bacteria include alcohols (ethanol, butanol, isopropanol, and Z,%butanediol), organic acids (acetic, butyric, formic, and lactic acid), polyols (arabitol, glycerol), ketones (acetone), and gases (CO,, methane, and hydrogen). Promising therrnophilic pentosefermenting bacterial species include Ciostridiurn therrnocellum, C. thermohydrosulfuricurn, C. therrnosaccharolyticum, C. therrnosulfurogenes, and Therrnoanaerobacter ethanolicus (Rosenberg, 1980; Avgeringos et al., 1981; Ng et al., 1981; Ben-Basset and Zeikus, 1981; Wiegel et al., 1983; Sonnleitner, 1983). Most of these organisms have been isolated from the hot springs of Yellowstone National Park, underneath algal mats (Brock, 1978). Most of them can be grown on defined media and are also relatively resistant to heavy metal ions (Wiegel et al., 1979; Wiegel and Ljungdahl, 1981) and other toxic substances and pollutants. Although they do not grow in the presence of oxygen, these anaerobes are not killed by the contact with air. Several properties of thermophilic

MICROBIAL PENTOSE UTILIZATION

101

bacteria have been identified which are advantageous for their industrial application. These advantages include Fermentation of a wide range of sugars present in cellulose and hemicellulose polysaccharides High metabolic activity which results in faster fermentation Less biomass formation with high product yield High temperature fermentation alleviates the risk of contamination No oxygen requirement Possibility of continuous recovery by distillation of volatile compounds Investigations on the thermotolerance of the spores of thermophilic Clostridia reveal that they all have high heat resistance. Spores of C. thermohydrosulfuricum have a decimal reduction time of 11 min at 121°C and over 1 2 hr at 100°C which makes the standard sterilization time greater than 30 min (Hyun eta]., 1983). Controversies exist regarding the taxonomic status of some of the C. thermosaccharolyticum strains (Wang et al., 1983) which do not form butyrate as is reported for the type shown and other strains (Hsu and Ordal, 1970; Landuyt and Hsu, 1985). A strain of C. thermosaccharolyticum (Hsu and Ordal, 1970) synthesizes ethanol during secondary metabolism after growth is uncoupled by acid production in a manner analogous to butanol production by C. acetobutylicum. C. therrnohydrosulfuricum, C. thermosulfurogenes, and C. thermosaccharolyticum reactions are different with thiosulfate which is reduced to H, S, is deposited as sulfur, or is not transformed, respectively (Landuyt and Hsu, 1985). These observations demand more taxonomic studies at a molecular level. IV. Pentose Metabolism

A. YEASTAND FUNGI

While the metabolism of hexoses has been extensively studied in yeasts, our understanding of pentose metabolism in these organisms is relatively limited. A number of yeasts that can assimilate aldopentoses oxidatively lack the ability to ferment them to ethanol (Barnett, 1976). However, with the upsurge of interest in pentose utilization for production of ethanol, a number of yeast and fungi have been tested. The transport of pentoses across the cell membrane constitutes the first step in their metabolism. Xylose transport has been studied in Saccharomyces cerevisiae (Kleinzeller and Kotyk, 1967; Batt et a].,

102

PRASHANT MISHRA AND AJAY SINGH

1986),Rhodotorula gracilis ( Janda et al., 1976; Alcorn and Griffin, 1978; Hofer and Misra, 1978; Heller and Hofer, 1978),Pachysolen tannophilus (Neirinck et al., 1984), Candida shehatae (Lucas and van Uden, 1986), and Candida utilis (Batt et al., (1986). The transport of D-xylose in S. cerevisiae, which cannot metabolize D-xylose, appears to take place via a facilitated diffusion process (Kleinzeller and Kotyk, 1967). The initial rate of xylose uptake in glucose-grown cells of S. cerevisiae and C. utilis is similar (Batt et a]., 1986). In uptake studies, Rhodotorula is widely used, and these studies indicate that xylose transport occurs by an active process (Hofer and Misra, 1978; Heller and Hofer, 1978) that is energized by an electrochemical gradient of Hf across the plasma membrane (Hofer and Misra, 1978). Studies by Lucas and van Uden (1986) suggest that facilitated diffusion as well as sugar proton symport mechanisms exist in C. shehatae. Candida shehatae repressed by growth in glucose or D-xylose exhibits a facilitated diffusion of D-xylose (K, k 125 mM, V, 22.5 mmol/g/hr).However, derepressed (starved) cells exhibit sugar proton symport for D-xylose uptake (K, 1.0 mM, V,, t 1.4 mmol/g/hr). While facilitated diffusion was absent or not measurable in starved cells of C. shehatae, when glucose was a substrate, the facilitated diffusion system coexisted with proton symport activity when D-xylose was the substrate (Lucas and van Uden, 1986). Once xylose is taken up by the cells, it is first converted to D-xylulose and is then subsequently phosphorylated. Pentose metabolism by yeasts is summarized in Fig. 1. Yeast and mycelial fungi differ from bacteria in the mechanism by which D-xylose is converted to D-xylulose. Bacteria generally achieve this conversion by employing the enzyme xylose isomerase (Horecker, 1962; Chen, 1980a,b),whereas yeast and mycelial fungi employ a two-step reduction and oxidation (Chiang and Knight, 1960) utilizing two sets of pyridine nucleotide-linked dehydrogenases. Pentoses are first reduced to their corresponding pentiols by a pyridinelinked dehydrogenase and then reoxidized to the corresponding pentuloses. Although a number of yeasts are capable of assimilating Dxylose under aerobic conditions (Barnett, 1976), only a few can utilize D-xylose anaerobically. D-Xylulose, on the other hand, is used anaerobically by many yeasts (Wang et al., 1980a,b; Ueng et al., 1981). The enzymes involved in early steps of xylose metabolism in yeasts and mycelial fungi are xylose reductase, polyol dehydrogenase (Arcus and Edson, 1956; Chiang and Knight, 1960; Veiga et a]., 1960; Chakravorty et al., 1962; Moret and Sperti, 1962; Veiga, 1968a,b),xylose isomerase (Tomoyeda and Horitsu, 1964; Hofer et al., 1971),and xylulokinase (Chakravorty et a]., 1962; Hofer et al., 1971).

*

*

103

MICROBIAL PENTOSE UTILIZATION

L-ARABINOSE

L-ARABITOL

+ XY L I T O L

L-XYLULOSE

p NAD 6 NADH

2

1 ACETATE

6

1'

'* 8

4I

(TCA

CYCLE^

ACETALDEHYDE

ETHANOL

FIG.1. Pentose metabolism by yeasts. (1) Aldose reductase; (2) xylitol dehydrogenase; (3) xylose isomerase; (4) xylulose kinase; (5) phosphoketopentoepimerase;(6) transaldolase and transketolase; (7) phosphoketolase; (8) pyruvate decarboxylase;(9) alcohol dehydrogenase; (10) erythritol dehydrogenase.

104

PRASHANT MISHRA AND AJAY SINGH

Although the oxidoreductive pathway seems to be an obligatory pathway for pentose metabolism in yeasts, the presence of an inducible enzyme xylose isomerase (D-xylose ketol-isomerase, EC 5.3.1.5) has also been reported in Candida utilis (Tomoyeda and Horitsu, 1964), Rhodotorula gracillis (Hofer et al., 1971),and Penicillium brevicompactum (Ziegal and Kobzeva, 1980). The purified xylose isomerase from xylose-grown C. utilis is specific for D-xylose over L-xylose but not for D-arabinose or L-arabinose and has a requirement for bivalent cations, e.g., Mn2+,Mg2+,and Co2+.Mn2+ is the most effective of these ions followed by Co2+ and Mg”. The enzyme has an optimum pH of 6-7 and an optimum temperature of 70°C. However, other groups of workers have failed to detect xylose isomerase activity in cell-free extracts of xylose-grown C. utilis (Chakravorty et al., 1962) and C. albicans (Veiga, 1968a). Since the presence of an inducible enzyme D-xylose isomerase has been demonstrated in some yeasts, the direct isomerization of Dxylose to D-xylulose may occur in certain yeasts. It appears that the first step of D-xylose metabolism in many yeasts and fungi is catalyzed by aldose reductase (NADP-linked polyol dehydrogenase; EC 1.1.1.21). The enzyme acts on L-arabinose and other aldoses as well as on D-xylose (Chiang and Knight, 1959; Veiga et al., 1960; Scher and Horecker, 1966a; Veiga, 1968a; Horitzu et al., 1968; Bolen and Detroy, 1985).The enzyme catalyzes the reduction of aldoses to corresponding polyols and requires reduced NADP to carry out the electron transfer. This enzyme has been reported in many yeasts and mycelial fungi, including C. albicans (Veiga et al., 1960;Veiga, 1968a,b), C. utilis (Horitzu et al., 1968; Batt et al., 1986), Geotrichum candidum (Moret and Sperti, 1962), Pichia quercuum (Suzuki and Onishi, 1973), Pichia stiptis (Verduyn et al., 1985a; Rizzi eta]., 1988),Cephalosporium chrysogenum (Birken and Pisano, 1976), Melampsora lini (Clancy and Coffey, 1980), Penicillium chrysogenum (Chiang and Knight, 1959), Pachysolen tannophilus (Smiley and Bolen, 1982; Verduyn et al., 1985b),and Fusarium oxysporum (Suihko et a]., 1983; Singh and Schiigerl, 1992). The enzyme purified from C. albicans (Veiga, 1968a) has a wide specificity for a number of substrates. However, based on the relative activity in the presence of various substrates, it was concluded that aldose in the D-glycero configuration with the hydroxyl group attached to carbon 2 acts as a good substrate of D-xylose reductase, while those lacking the hydroxyl group at carbon 2 are poor substrates. Reverse reactions using polyols as the substrate and NADP as the cofactor have also been reported (Veiga, 1968a). NADH-linked xylose reductase has been purified from the xylose-fermenting yeast Pichia stiptis and has been shown to exhibit a dual coenzyme specificity for NADH

MICROBIAL PENTOSE UTILIZATION

105

and NADPH (Verduyn et al., 1985a). In most yeasts which do not or only slowly ferment xylose anaerobically, xylose reductase activity is NADPH linked (Scher and Horecker, 1966b; Bruinenberg et al., 1983) and they either lack or exhibit low NADH-linked activity (Bruinenberg et al., 1983).Multiple forms of xylose reductase in Pachysolen tannophilus CBS 4044 have been reported by Verduyn et al. (1985b) who observed both NADPH- and NADH-dependent xylose reductase activity in xylose-grown P. tannophilus. The ratio of these activities varies with growth conditions. These two xylose reductase enzymes are separated by affinity chromatography. One enzyme is active with both NADPH and NADH, while the other is specific for NADPH. In addition to coenzyme specificity, these two enzymes also differ in their affinities for xylose and NADPH. NADH-linked xylose reductase activity is very low in aerobic cultures of P. tannophilus grown in xylose-containing media. It has been suggested that NADH-linked xylose reductase activity is required for the anaerobic fermentation of xylose and that such fermentation via NADPH-linked reductase leads to an imbalance of the NAD+/ NADH redox system (Bruinenberg et al., 1983,1984).Therefore, oxygen limitation introduces the need for NADH-linked xylose reductase in order to avoid accumulation of NADH. Batt et al. (1986) reported the presence of xylose reductase in S. cerevisiae, although its activity was found to be fivefold less than that of C. utilis. The enzyme is inducible by xylose and repressible by glucose in S. cerevisiae (Batt et al., 1986). The inducibility of this enzyme has also been reported in the presence of xylose in F. oxysporum (Suihko et al., 1983; Singh and Schugerl, 1992). Yeasts also metabolize L-arabinose, most likely by utilizing the same enzyme aldose reductase (Bolen and Detroy, 1985) which reduces L-arabinose to L-arabitol (Fig. 1). In the next step the reoxidation of xylitol to D-xylulose is catalyzed by xylitol NAD-2 oxidoreductase which is also known as xylitol dehydrogenase (NAD polyol dehydrogenase, EC 1.1.1.9) (Chiang and Knight, 1960; Moret and Sperti, 1962; Chakravorty et a]., 1962). This enzyme has been reported in cell-free extracts of C. utilis (Chakravorty et al., 1962; Horitzu and Tomoeda, 1966; Batt et al., 1986),C. albicans (Veiga, 1968b), Pullularia pullulans (Sugai and Veiga, 1981), Cephalosporium chrysogenum (Birken and Pisano, 1976),Pachysolen tannophilus (Simley and Bolen, 1982), S . cerevisiae (Batt et al., 1986), and F. oxysporum (Suihko et al., 1983; Singh and Schiigerl, 1992). Studies on purified enzymes from C. utilis showed that the enzyme lacked activity toward xylitol in the presence of NADP, NADPH, or NADH as a cofactor (Chakravorty et al., 1962). The enzyme has a pH optimum of 9.0, has specificity toward xylitol and D-xylulose, and the equilibrium of reac-

106

PRASHANT MISHRA AND AJAY SINGH

tion at neutral pH favors polyol formation. The enzyme has also been purified from C. albicans (Veiga, 1968b),Cephalosporium chrysogenum (Birken and Pisano, 1976),and Pichia stiptis (Rizzi et al., 1989).Genetic evidence for the involvement of xylitol dehydrogenase in D-xylose metabolism in yeast has been provided by the isolation of mutants of P. tannophilus with a defect in xylitol dehydrogenase. These mutants are able to grow on D-xylulose but fail to grow on D-xylose or xylitol (Maleszka et al., 1983a). It is surprising to note that xylitol, which is produced as a common intermediate of pentose metabolism, does not act as a good substrate for microbial utilization (Gong et a]., 1983). This finding has been attributed to the limited permeability of xylitol (McCracken and Gong, 1983). In the next step, D-xylulose is metabolized by a phosphorylation reaction. D-Xylulokinase (EC 2.7.1.17) catalyzes the phosphorylation of D-xylulose to ~-xylulose-5-phosphate.Information available on its activity in yeasts and mycelial fungi is limited. Chiang and Knight (1960) detected D-xylulokinase activity in cell-free extracts of P. chrysogenum. The activity of this enzyme was later reported in cellfree extracts of C. utilis (Chakravorty et a]., 1962) grown in the presence of D-xylose or L-arabinose. Further, the presence of D-xylulokinase in yeast has been implied by the ability of many yeasts to utilize D-xylulose under aerobic or anaerobic conditions (Wang et a]., 1980a,b: Gong et a]., 1981c: Gong, 1983). ~-Xylulose-5-phosphateis further metabolized via the pentose phosphate pathway which utilizes the enzymes transaldolase and transketolase to convert ~-xylulose-5-phosphateto ~-glyceraldehyde-3phosphate (Chakravorty et al., 1962). ~-Glyceraldehyde-3-phosphate either produces ethanol via entry into the EMP pathway followed by action of pyruvate decarboxylase and alcohol dehydrogenase or alternatively is oxidized to pyruvate and then to CO, and water via the TCA cycle. In yeasts, the pentose phosphate pathway is stimulated by the oxidation of NADPH (Holzer and Witt, 1960: Osmond and ApRees, 1969). Thus availability of NADP/NADPH may control xylose reduction and activation of the pentose phosphate cycle. Jeffries (1982) suggested that the metabolism of xylose in yeast is coordinately controlled. The regeneration of NADPH required for the reduction of xylose to xylitol is produced via the pentose phosphate pathway and by the subsequent oxidation of hexose phosphate. For each mole of xylose metabolized to CO,, 10 mol of NADPH can be generated, which in turn can be utilized for conversion of 10 mol of xylose to 10 mol of xylulose through xylitol as an intermediate (Gong, 1983). Under aerobic conditions many yeasts have the potential to produce polyhydric alcohols such as xylitol, glycerol, and D-arabitol from xylose

MICROBIAL PENTOSE UTILIZATION

107

(Onishi and Suzuki, 1966; Gong, 1983). Other products of xylose metabolism are acetic acid (Jeffries, 19821, citric acid (Onishi and Suzuki, 1966), and xylonic acid (Kiessling et aJ., 1962). In addition to the pentose phosphate pathway, the enzyme phosphoketolase has been found in several yeasts, including Rhodotorula graminis, Rhodotorula glutinis, Candida tropicalis, Candida humicola, and Candida 107 (Sgorbati et al., 1976; Ratledge and Botham, 1977; Botham and Ratledge, 1979). Phosphoketolase (EC 4.1.2.9) catalyzes the cleavage of either ~-xylulose-5-phosphateor ~-fructose-6-phosphate to form glyceraldehyde-%phosphate or ~-erythrose-4phosphate: ~-xylulose-5-P+ phosphate + glyceraldehyde-3-P + acetyl phosphate + H,O Based on 14C labeling experiments it has been found that D-xylulose phosphate in Fusarium species is cleaved between carbon atoms 2 and 3 rather than being metabolized by the pentose phosphate and Embden Meyerhof pathways. The product labeling pattern indicates that acetic acid is derived from the two carbon fragment and that ethanol and CO, are products of the three carbon fragment (Gibbs et al., 1954) The enzyme involved is phosphoketolase. It has been noted that F. oxysporum produces significant quantities of acetic acid and ethanol but this mold when growing produces higher levels of CO, and much less acetate. Thus it is likely that either growing cells use a different metabolic scheme or they reduce acetate to ethanol. Further, based on NMR studies, Ligthelm et al. (1988) could not demonstrate the presence of phosphoketolase activity in Pichia stiptis cells as [13C]xyloseyielded only [2-13C]ethanol.If phosphoketolase is active, then [2-13C]ethanol (which is formed from acetyl phosphate) should have also been detected. However, Girio et al. (1989), using Candida shehatae, have suggested that the operation of phosphoketolase, in addition to the classical pentose phosphate pathway, is essential for NADH dissimilation as the reduction of acetyl phsophate to ethanol oxidizes NADH produced during the conversion of xylitol to xylulose. Although much attention is required to unequivocally establish the importance of phosphoketolase, it has the potential to act as a key enzyme for xylose fermentative metabolism in yeasts. B. BACTERIA

Pentose metabolism in bacteria has been studied in detail (Horecker, 1962). As discussed earlier the major difference in early steps of pentose metabolism in bacteria is how they convert xylose to xylulose. Most of the bacteria are known to employ the xylose isomerase pathway instead

108

PRASHANT MISHRA AND AJAY SINGH

of the oxidoreductase pathway for this purpose. Many bacteria such as Aerobacter aerogenes (Mortlock and Wood, 1964a,b) and Salmonella typhimurium (Shamanna and Sanderson, 1979a,b) possess a set of inducible enzymes. These enzymes are xylose transport enzymes, xylose isomerase, and xylulokinase. The presence of these enzymes in S. typhimurium has been demonstrated by the sequential appearence of xylose transport enzymes, xylose isomerase, and xylulokinase (Shamanna and Sanderson, 1979a). The involvement of these enzymes was further confirmed by isolating mutants defective in these enzymes. The metabolism of other pentoses in most bacteria also involves an isomerization step (Mortlock and Wood, 1964a). Thus, L-arabinose, L-xylose, and Darabinose are isomerized to their corresponding pentuloses which are subsequently phosphorylated. The enzyme D-xylulokinase catalyzes the phosphorylation of D-xylulose to ~-xylulose-5-phosphate. However, Dribose is metabolized by a direct phosphorylation reaction by the formation of ~4bose-5-phosphatein E. coli (David and Wiesmeyer, 1970). In some bacteria, e.g., Enterobacteria, Corynebacteria, and Brevibacteria, the reduction of xylose to xylitol is catalyzed by an NADPHdependent xylose reductase (Yoshitake et al., 1973a,b).Further metabolism involves the oxidation of xylitol to xylulose-5-phosphate. Pentiols, e.g., xylitol, ribitol, and arabitol, are oxidized to their corresponding pentuloses, e.g., xylulose, ribulose, and xylulose, in the presence of the enzyme pentiol dehydrogenase (Mortlock and Wood, 1964a; Mortlock, 1976).The bacteria also employ some unusual pathways for the metabolism of D-xylose and xylitol (London and Chace, 1977,1979;Yamanaka et al., 1977; Yamanaka and Gino, 1970). The enzyme NAD-D-xylose dehydrogenase has been described in Arthobacter species (Yamanaka et al., 1977), catalyzing the formation of D-xylonolactone which is subsequently hydrolyzed to xylonic acid:

D-xylose

+ NAD+

NAD-xylose dehydrogenase w

D-xylonolactone

+ NADH + H+

The purified enzyme exhibits specificity for NAD and D-xylose (Yamanaka and Gino, 1979). Some strains of Lactobacillus casei grow anaerobically on ribitol or xylitol by utilizing a unique pathway (London and Chase, 1977). Pentiols are transported into cell by a substratespecific PEP-phosphotransferase system which converts them to their corresponding phosphates, e.g., xylitol-5-phosphate and ribitol-5phosphate. Pentiol phosphates are then converted to pentulose phosphates in the presence of NAD-specific ribitol or xylitol phosphate dehydrogenases.

MICROBIAL PENTOSE UTILIZATION

109

After the formation of xylulose-5-phosphate, its metabolism proceeds along the two major routes (1)by cleavage of pentoses with the formation of a three carbon unit and another two carbon unit compound using the enzyme phosphoketolase (Fred et a]., 1922; Johnson et a]., 1931; Racker, 1948) and (2) by the pentose phosphate pathway leading to the formation of hexose phosphate or other intermediate compounds capable of entering the glycolytic pathway using the enzymes transaldolase and transketolase (Lipmann, 1936; Dickens, 1938). Some bacteria, e.g., K. penumoniae, completely convert xylulose-5-phosphate to a triose via the transketolase and transaldolase enzymes of the pentose phosphate pathway. The metabolic pathway in this organism has been deduced by labeled pentoses (Neish and Simpson, 1954; Altermatt et a]., 1955). In this process 3 mol of pentose yields 5 mol of Dglyceraldehyde-3-phosphatewith the consumption of 5 mol of ATP. Glyceraldehyde is converted to pyruvate by enzymes of the glycolytic pathway. Pyruvate is metabolized by the tricarboxylic acid cycle or is converted to various products of anaerobic fermentation. For example, in K. penumoniae, 2,3-butanediol is the major product of the fermentative pathway (Fig. 2). Two moles of pyruvate is condensed to form acetolactate in the presence of enzyme “pH 6 acetolactateforming enzyme” (Stormer, 1968a; Johansen et a]., 1975). Acetolactate is then decarboxylated (Loken and Stormer, 1970) in the presence of the enzyme acetolactate decarboxylase to yield acetoin (acetylmethylcarbinol). In the final step, acetoin is reduced to 2,3-butanediol in the presence of acetoin reductase (Larsen and Stormer, 1973). Purification of the enzyme acetoin reductase shows that two stereospecific enzymes exist. One specifically reduces L-acetoin to L-butanediol, while the other reduces D-acetoin to meso-butanediol (Voloch, 1981). In strict anaerobic bacteria, for example clostridia, pyruvate is converted to acetyl-CoA with the reduction of ferredoxin, which in turn is oxidized by the activity of a hydrogenase enzyme (Dellweg, 1981), which disposes of excess electrons in the form of molecular hydrogen, thus ensuring high energy production by controlling the flow of electrons in substrate metabolism (Fig. 3). Ferredoxin indeed limits pyruvate oxidation (Valentine, 1964). Under iron deficient conditions, in sacchrolytic clostridia, hydrogen is not produced but instead a shift toward acetate production occurs. With a pH shift to the acid region it becomes progressively more difficult for the cell to reoxidize reduced NADH to NAD+. Therefore the organism opts for the production of butyrate which has a lower acidic end product than acetate. Once accumulation of butyrate lowers the pH to 4.0, the organism favors the production of acetate and converts butyrate to butanol. Clostridium

110

PRASHANT MISHRA AND AJAY SINGH

Lrl 1 Q 1 XYLOSE

X y l o s e isomerase

XY LULOSE

Xylulokinase

IXYLULOSE-5-PHOSPHATE I

1

T r a n s a l d o l a s e and T r a n s k e t o l ase

IPENTOSE PHOSPHATE PATHWAY I

9 GLYCOLYTIC PATHWAY

PYRUVATE

enzyme

ti TCA CYCLE

Acetol a c t a t e d e c a r b o x y l ase

Acetoi n reductase

FIG.2. Catabolism of xylose to 2,3-butanediol.

acetobutylicum possesses a transferase system that can divert the production of /?-hydroxyl-CoA to acetoacetate, and on decarboxylation, acetoacetate forms acetone (Fig. 3). This diversion in the metabolic pathway eliminates two NAD+-generating steps. Thus production of acetate is the key point in the control of metabolic diversions for produc-

I

1 XYLULOSE-5-PHOSPHATE

I

PENTOSE PHOSPHATE PATHWAY

GLYCOLYTIC PATHWAY

1

Hz

LrJ ACETYL CO A

F-7 ACETALDEHYDE

Acetate 6 ACETYL-P

ACETOACETYL CO A

BUTYRYL COA

Acetate NADH NAD

A c e t y l GO A

11

91;:

+

0 23 A I ACETOACETATE

12

B-OH BUTYRYL CO A

I

, k::,

ACETONE

CROTONYL CO A

BUTYRALDEHYDE]

ISOPROPANOL

4

13

NAD

FIG.3. A general anaerobic pentose metabolic scheme for the synthesis of various solvents. (1) Hydrogenase; (2) aldehyde dehydrogenase; (3) ethanol dehydrogenase; (4) phosphoketolase; (5) acetoacetyl-CoA thiolase; (6) acetyl-CoA transferase: (7) transferase; ( 8 )acetoacetate decarboxylase; (9)hydroxy butyrate dehydrogenase; (10)crotonase; (11) oxidoreductase; (12)butraldehyde dehydrogenase; (13)butanol dehydrogenase.

PRASHANT MISHRA AND AJAY SINGH

112

tion of various solvents. (Volesky and Szczesny, 1983). In another species, klebsiella planticoja, pyruvate is dissimilated by the enzyme pyruvate formate lyase to yield acetate, ethanol, and formate in a molar ratio of 1 : l : Z . D-Lactate and 2,3-butanediol are formed in small amounts (Tolan and Finn, 1987). V. Production of Solvents and Organic Acids

A. ETHANOLPRODUCTION

Some examples of pentose fermentation to ethanol by various yeast, fungi, and bacteria are presented in Tables 111, IV, and V. 1. Utilization of Pentose Sugars

Yeasts. Pachysolen tannophilus, Candida shehatae, C. tropicalis, and Pichia stiptis are the most studied pentose-fermenting yeasts (Table 111). In regard to the ethanol concentration and by-product formation, C. shehatae and P. stiptis are considered promising organisms. Wood and Millis (1985) compared xylose fermentations of P. stiptis, P. tan-

TABLE I11 BIOCONVERSION OF XYLOSEINTO ETHANOLBY YEAST

Organism Candida sp. XFZ17 C. shehatae CSIRY492 C. shehatae Y-12856 C. tenuis CSIR-Y566 C. tropicalis ATCC 1369 C. tropicalis ATCC 32113

Pachysolen tannophilus

Ethanol (giliter)

Yield (gig)

Productivity (glliterihr)

Xylitol (g/liter)

21 26

0.42 0.29

0.42 0.65

-

24 13.3 5.5

0.45 0.25 0.07

0.20 0.42 0.06

9.3

-

5.8

0.11

0.03

3.4

1.8

0.06

0.04

22.9

33.4

0.21

0.12

0

Schvester et al.

39 22.2 5.6

0.39 0.43 0.28

0.28 0.79

0.9 0

Silinger (1985) du Preez et al. (1989) Margaritis and Bajpai

4

1

Ref Gong et al. (1981a) du Preez (1983)

Silinger (1985) du Preez et al. (1989) Jeffries (1981) Skoog and HahnHagerdal (1988) Debus et al. (1983)

IFGBOlOl

P. tannophilus NRRL Y2460

Pichia stiptis Y-7124 P. stiptis CSIR-Y633 K1yuvermyces marxianus SUB80-S

(1983)

(1982)

MICROBIAL PENTOSE UTILIZATION

113

nophilus, and C shehatae. The maximum ethanol concentrations obtained were 28 g/liter with P. tannophilus, 33 g/liter with C.shehatae, and 57 g/liter with P. stiptis. The corresponding productivities were 0.11, 0.26, and 0.18 g/liter/hr, respectively. Pichia stiptis was the least affected by the higher substrate concentrations (Silinger, 1985). The ethanol yield from xylose with P. tannophilus was reported to be increased by 32% when 5 glliter glucose was added during the fermentation (Jeffries, 1985). However, the mechanism for increased yield could not be explained. Du Preez et al. (1986) observed that P.stiptis and C. shehatae could ferment arabinose, rhamnose, galactose, mannose, and xylitol in addition to xylose (du Preez et al., 1989). In a fed-batch culture, where the sugar concentration was kept between 5 and 8 g/liter, 26 g/liter ethanol was obtained with P. tannophilus. The yield was increased by 41% as compared to batch fermentation (Neirinck, 1985). In continuous culture with P. tannophilus, a productivity of 2.2 g/liter/hr was obtained at a dilution rate of 0.12/hr (Linko, 1986). A substrate concentration of 50 g/liter at a dilution rate of 0.34/hr resulted in a productivity of 2 . 1 g/liter/hr (Skoog and HahnHagerdal, 1988). Generally a decrease in aeration increses the ethanol yield in yeasts, but this leads to lower cell growth and the risk of washout. This problem could be overcome by recirculating or immobilizing the cells (Maleszka et al., 1983b). Pichia stiptis immobilized in alginate gel resulted in a productivity of 0.58 g/liter/hr with a substrate concentration of 50 g/liter (Linko, 1986).With nylon net immobilization the productivity was 0.54 g/liter at the same substrate concentration. Filamentous fungi. Species of Fusarium, Mucor, Monilia, Neurospora, and Paecilornyces are known to ferment pentose sugars (Table IV). Rosenberg et al. (1981) compared the xylose fermentation of F. oxysporum with €3. macerans in a pH-controlled fermenter. Fusarium oxysporurn produced 0.41 g/g ethanol from 10 giliter of xylose with 94% of the original xylose carbon accounted for. The lost carbon was believed to be mainly in the form of CO,. The growth of Fusarium was not exponential, and the specific growth rate declined with increasing cell mass. The conversion rate was too slow to be considered commercially significant. Ethanol yields of 0.32 and 0.16 g/g xylose were obtained with F. lycopercici and Mucor 101, respectively (Ueng and Gong, 1982). The rate of ethanol production was also lower with Mucor at a high xylose concentration (20%). Suihko and Enari (1981) screened 26 different Fusarium species for ethanol production from xylose. The best strain, F. oxysporum, was able to produce 2 1 g/liter ethanol from 50 g/liter of xylose in 7 days. Further optimization of the fermentation

PRASHANT MISHRA AND AJAY SINGH

114

TABLE IV

FERMENTATION OF PENTOSES BY FILAMENTOUS FUNGI ~~~

~

Organism

Fusarium oxysporum F5 F. oxysporum

Substrate

Ethanol Yield Productivity (glliter) (g/g) (g/liter/hr)

Ref.

Xylose Xylitol Xylose

14 2 25

0.28 0.04 0.50

0.15 0.02 0.17

Christakopoulos et al.

Xylose Xylose Xylitol Xylose

11 8 7 6.8

0.22 0.16 0.14 0.34

0.11 0.08 0.07 0.05

Suihko and Enari (1981) Ueng and Gong (1982)

73.5 10.2 13.8

0.37 0.20 0.28

0.44 0.11 0.14

Wu et al. (1986)

(1989)

Suihko and Enari (1981)

VTT-D-80134

F. solani Mucor 105

Neurospora crassa Paecilomyces sp. Xylose Ribose Arabinose

Deshpande et al. (1986)

parameters resulted in the ethanol concentration of 25 g/liter in 6 days, corresponding to the theoretical yield. A mixture of glucose (25 g/liter) and xylose (25 g/liter) was also tested by these workers. Glucose was completely utilized after 1 day, and thereafter the fermentation rate corresponded to that of xylose fermentation, yielding 25 g/liter ethanol after 5 days (Enari and Suihko, 1984). The Monilia sp. also ferments a wide range of substrates, including cellulose, xylan, starch, xylose, cellulose, galactose, and arabinose. A concentration of 20 g/liter xylose produced 6.8 g/liter ethanol in 7 days (Deshpande et al., 1986). Wene and Antonopoulos (1988) selected the strain F. oxysporum AL 22-760 out of 2000 Fusarium isolates. This strain showed consistent yields under semiaerobic and anaerobic conditions. Under semiaerobic conditions 8.2 g/liter ethanol could be obtained from 20 g/liter of xylose in 72 hr. Using a cell recycling system, the fermentation time could be reduced to 48 hr with comparable yields. Fusarium oxysporum F3 produced 5 g/liter ethanol from 20 g/liter xylose in 6 days with a theoretical yield of 48%. (Christakopoulos et al., 1989). Fusarium oxysporum DSM 841 produced almost equal concentrations of ethanol and acetic acid from D-xylose and D-glucose (Singh et al., 1992c,d). Bacteria. Table V presents ethanol yields of some pentose-fermenting bacteria. Rosenberg et al. (1981) studied the fermentation of xylose to ethanol with B. macerans ATCC 8244 and compared its xylose fermentation efficiency with F. oxysporum ATCC 10960. They observed that specific growth rate (O.l5/hr) and specific ethanol productivity 10.8 g/

115

MICROBIAL PENTOSE UTILIZATION TABLE V ETHANOL PRODUCTION BY D-XYLOSE-FERMENTING BACTERIA

Organism CIostridium thermohydrosulfuricum C. thermosaccharolyticum C. thermosulfurogenes C. saccharolyticum Thermoanaerobacter ethanolicus Zymomonas anaerobia Klebsiella pneumoniae Bacteroides polypragmatus Bacillus macerans

Ethanol (dliterl

Yield (dg)

Ref.

13

0.43

Ng et al. (1981)

27

0.35 0.20 0.07 0.48 0.29 0.31 0.15 0.26

Avgeringos et al. (1981) Schinck and Zeikus (1983) Asther and Khan (1985) Carreira et al. (1983) Asther and Khan (1985) Banerjee (1985) Patel (1984) Rosenberg et al. (1981)

8.5 4.8 37.3 12.7 6.5 -

g/hr) were much higher than that of A. hydrophila, B. polymyxa, and Aerobacter indologenes found to produce ethanol at levels of 48.9, 63.0, and 55.9 mmol/lOO mmol xylose (Rosenberg, 1980). Klebsiella pneumoniae MB-16-1048 produced 12.7 g/liter ethanol with a corresponding yield of 0.31 g/g xylose in 20 hr (Banerjee, 1985). This mutant strain showed 82% improvement in the ethanol yield as compared to the original strain of K. pneumoniae isolated from soil. Thermophilic anaerobes have been shown to ferment various hexoses and pentoses, including cellulose, hemicellulose, starch, and pectin. The fastest growing thermoanaerobe yet isolated is Thermobacteroides acetoethylicus which has a doubling time of 25 min on glucose (pmax 1.66). All of the thermoanaerobes used for ethanol production have the same end product spectrum. Besides ethanol they also produce acetic and lactic acids, CO,, and hydrogen. Thermobacteroides ethanolicus produces ethanol at 70°C which favors a process for simultaneous fermentation and evaporation. A mutant with high sugar tolerance has been reported by Carreira et al. (1983). Bacteroides polypragmatus can ferment xylose, arabinose, and ribose to ethanol, but it also produces acetate, butyrate, and H, and CO, gases (Patel, 1984). A coculture of Zymomonas anaerobia and C. saccharolyticum was used by Asther and Khan (1985).Acetate produced from glucose by C. saccharolyticum further inhibits the fermentation of xylose. Cocultures of C. therrnocellum with C. thermohydrosulfuricum are effective in fermenting cellulose and hemicellulose (Ng et al., 1981). Similarly, C. thermocellum and T. ethanolicus on cellulose and hemicellulose showed an increased rate of degradation and higher yields of ethanol by coculture (Wiegel et a]., 1983).

PRASHANT MISHRA AND AJAY SINGH

116

With bacteria, it is possible to obtain product concentrations, yields, and productivities comparable to those in hexose fermentations with yeasts. However, by-product formation presents a problem in the downstream processing of the product. Reported ethanol yields are low from xylose fermentation with yeast. Another difficulty in xylose utilization by yeast is achieving a complete conversion of xylose to xylulose economically. Also, proper control of oxygenation is required. On the other hand, filamentous fungi ferment xylose with high ethanol yields, but the fermentation rate is too slow as compared with yeast and bacteria and still needs improvement. The comparative kinetics of xylose fermentation by fungi, bacteria, and yeast is given in Table VI. 2. Use of o-Xylose Isomerase

Some organisms cannot utilize D-xylose directly, so instead they utilize xylulose, an isomer of D-xylose (Table VII). The ability of some yeasts and fungi to utilize D-xylulose for growth and ethanol production led to the development of two methods for producing ethanol from D-xylose using the D-xylose isomerase. One method depends on the isomerase to first isomerize D-xylose to D-xylulose which is then sup-

TABLE VI

COMPARATIVE KINETICSOF GROWTHAND PRODUCTFORMATION OF YEAST,FILAMENTOUS FUNGI, AND BACTERIA ~~

Organism Pachysolen tannophilus Kly uvemmyces marxianus Candida shehatae Pichia stiptis Fusarium oxysporum Bacillus macerans Klebsiella pneumoniae Clostridium therrnohydrosulfuricum

~

Specific growth ratea

Cell Ethanol yieldb yieldC

Specific ethanol productd

Ref.

0.24

0.22

0.34

0.12

Silinger et a1. (1982)

0.12

0.16

0.28

0.10

Margaritis et al. (1981)

0.02 0.08 0.24 0.15

0.08 0.12 0.41 0.05 0.02 0.07

0.34 0.43 0.41 0.26 0.31 0.43

0.28 0.25 0.082 0.80 0.15

du Preez et al. (1989) du Preez et al. (1989) Rosenberg et al. (1981) Rosenberg et al. (1981) Banerjee (1985) Ng et al. (1981)

-

0.45

Per hour. Grams dry weight per gram xylose. Grams ethanol per gram xylose. Grams ethanol per gram xylose per hour.

-

117

MICROBIAL PENTOSE UTILIZATION TABLE VII FERMENTATION OF D-XYLOSE WITH XYLOSE Organism Saccharomyces cerevisiae S. cerevisiae Schizosaccharomyces pombe

Candida tropicalis Pichia stiptis Pachysolen tannophiius S. cerevisiae + P. tannophilus

Enzyme Novo Sweetzyme MKC Optisweet Brocades Maxazyme MKC Optisweet MKC Optisweet MKC Optisweet MKC Optisweet

ISOMERASE A N 0 YEAST

Ethanol (g/liter)

Yield (g/g)

Productivity (g/liter/hr)

20.9

0.17

0.44

Chiang (1981)

24.0

0.50

1.o

2.3

0.05

0.009

Wang et nl. (1980b) Hahn-Hagerdal et al. (1986)

7.9

0.21

-

6.3

0.17

-

5.0

0.12

-

12.6

0.31

-

Ref.

Linden and HahnHagerdal (1989) Linden and HahnHagerdal (1989) Linden and HahnHagerdal (1989) Linden and HahnHagerdal (1989)

plied to the cells. The other method employs recombinant yeasts that bear a bacterial gene for the D-xylose isomerase. The ability to grow on D-xylose and D-xylulose may not necessarily be linked. Organisms that fail to grow on D-xylose grow readily on D-xylulose. Among the xylulose-fermenting yeasts, S. cerevisiae, S. pombe, and C. tropicalis have been studied the most. These yeasts can ferment xylose in the presence of the bacterial xylose (glucose) isomerase. Wang et al. (1980a) compared different enzyme preparations and found Maxazyme GI the best for fermenting xylose with S. pombe. The xylose isomerase reaction is an equilibrium reaction, and at most, 20% xylose is converted to xylulose (Skoog and Hahn-Hagerdal, 1988). However, using repeated isomerization and fermentation, a transformation of 85% xylose to ethanol could be achieved with S. pombe (Chiang, 1981). With Baker’s yeast, 62 g/liter ethanol could be achieved at 30°C using a simultaneous isomerization and fermentation concept (HahnHagerdal et al., 1986). The corresponding yield and productivities were found to be 0.34 g/g and 1.25 g/liter/hr, respectively. Linden and HahnHagerdal (1989) tested five yeasts, C. tropicalis, P. stipfis, P. tannophiIus, S. pombe, and S. cerevisiae, for the fermentation of spent sulfite liquor in the presence of commercial xylose (glucose) isomerase. A maximum yield of 0.41 g/g was obtained with S. cerevisiae. Some authors also used S. cerevisiae, xylose isomerase, and 4.6 mM sodium

118

PRASHANT MISHRA AND AJAY SINGH

azide for the fermentation of hydrogen fluoride pretreated and acidhydrolyzed wheat straw. In this instance an ethanol yield of 0.40 g/g was obtained. The xylose utilization was found to be 84% compared to 51% in spent sulfite liquor. An important consideration in setting up the simultaneous isomerization and fermentation system is the difference in optimum operating conditions for the enzyme and cells. The optimum pH for yeasts is below 7 and the optimum temperature is below 40°C, whereas the optimum pH for isomerization is 8 and the optimum temperature is 70-80°C (Olivier and du Toit, 1986). Thus, a compromise in the conditions must be sorted out when enzyme and cells are kept in the same reactor. 3. Performance in Lignocellulosic Hydrolysate The ultimate goal in studying the fermentation of xylose, xylulose, or other pentose sugars is utilizing the pentose fraction in lignocellulosic hydrolysate. Since the insoluble raw material is inaccessible to most of the fermenting microorganisms, it should be pretreated and hydrolyzed. Hemicellulose can be hydrolyzed to simple sugars using enzymes or chemical or physical methods. Some of the important studies made on the utilization of a hemicellulose hydrolysate are summarized in Table VIII. Several studies reported the product concentration at more than 30 g/liter, yields higher than 0.4 g/g, and productivities higher than 0.5 giliterihr. The highest product concentration, 84 g/liter with a yield of 0.47 g/g, was achieved with C. shehatae from whole barley using pretreatment and enzymatic hydrolysis (Wayman and Parekh, 1985). Using an adapted strain of Candida XF 217, 29 g/liter of ethanol was obtained from 100 g/liter of a sugarcane hydrolyzate (Lodics and Gong, 1984). Although ethanol can be produced from D-xylose in significant yields, conversion rates are often slow. In some cases, fermentation times of 24 to 36 hr have been reported, but most of these instances are for relatively low sugar concentrations. For higher sugar concentrations, fermentation periods are generally appreciably higher. For more rapid fermentations, high cell densities are often required (Schneider, 1989a); for example, 8.5 to 16 giliter dry weight of P. stiptis with a hydrolysate prepared using a combination of steam, SO,, and enzyme (Parekh et al., 1986) and 15 to 19 g/liter of C. shehatae with spent sulfite liquor (Yu et al., 1987). Intermittent feeding of a cellulose hydrolysate to a hemicellulose hydrolysate of hardwood resulted in improved ethanol yields compared with fermentations of either hydrolysate alone or a mixture of the two (Beck, 1986).

119

MICROBIAL PENTOSE UTILIZATION TABLE VIII

PERFORMANCE OF XYLOSE-FERMENTING ORGANISMS IN A LIGNOCELLULOSIC HYDROLYSATE Fermentable Ethanol sugars yield Substrate Treatment (glliter) (g/g) (I

Organism Candida sp. XF217 C. tropicalis C. shehatae Pachysolen tannophihs Pichia stipis Saccharomyces cerevisiae Mucor sp.

Corn stover Aspen Red oak Aspen

Acid

SOX, 20A

0.38

Gong et al. (1981a)

Acid Acid Acid, Na,

69X, 11H 40X, 3G 5X, 10G, 2M

0.15 0.25

Fein et al. (1984) Jeffries (1985) Deverall (1983)

so3

Steam SO, 28X, 64G Aspen Softwood Na,SO, 27.5X. 6G, 4A Acid 43X, 13G Bagasse

Wood Fusarium oxysporum Clostridium Aspen saccharolyticum + Zymomonas mobilis

Ref.

0.39 0.45 0.34 0.33

Acid

18.5X, 4.5G

0.24

Acid

15X, 34G

0.47

Parekh et al. (1986) Linden and HahnHagerdal (1989) Ueng and Gong (1982) Joshi et al. (1990) Murray and Asther (1984)

A, arabinose: G , glucose: H, hexose: M, mannose: X: xylose.

Rosenberg et al. (1981)tested the performance of F . oxysporum ATCC 10960 in an acid hydrolysate of wheat straw. A total of 3.2 g/liter of ethanol was produced from 11 g/liter of the total sugars (2 g/liter glucose 9 g/liter xylose) present, with a conversion efficiency of 29%. Ueng and Gong (1982) employed species of Fusarium and Mucor for

+

the conversion of sugarcane bagasse hemicellulose hydrolysate. The ethanol production rate with Mucor was higher than with Fusarium. Fusarium F5 could ferment pure D-XylOSe readily but not the hydrolysate. Recently Joshi et al. (1990) attempted the utilization of sugars present in a wood hydrolysate by F . oxysporum NCIM-1072 and F. oxysporum D-140. These strains produced 11-13 g/liter of ethanol from the hydrolysate containing about 55 g/liter of the total sugars (glucose + xylose). B.

ACETONE AND BUTANOLPRODUCTION

Several species of saccharolytic clostridia are known to produce acetone and butanol, in addition to volatile fatty acids and gaseous products from carbohydrates. In some cases, acetone is further reduced to isopro-

120

PRASHANT MISHRA AND AJAY SINGH

panol. An attractive feature of these organisms is their capacity to utilize pentose sugars and certain other complex carbohydrates (Table IX). Compere and Griffith (1979) investigated some good solvent-producing clostridia, namely C. acetobutylicurn, C. butylicum, and C. pasteurianum, for the conversion of various mono- and disaccharides. Growth and solvent synthesis rates with pentoses were comparatively slower. In contrast, di- and oligosaccharides have higher solvent accumulations and are more sensitive to higher concentrations of pentose. C. butylicum NRRL B592 was more efficient in solvent production from xylan and cellobiose. In an interesting experiment by Maddox (1982), C. acetobutylicum was grown on pentoses in a mixed culture with S. cerevisiae. While the S. cerevisiae converted hexoses from molasses (5% solids) to ethanol (22 g/liter) in 48 hr, the bacterium was inoculated into the culture after 24 hr and utilized arabinose and xylose components (30 g/liter), producing 6.6 and 3.7 g/liter, respectively, of butanol in another 170 hr. The higher butanol production from arabinose reflected higher conversion rates observed with this pentose sugar. Since wood and agricultural residues are plentiful sources of fermentable carbohydrate materials, the utilization of these substrates attracts considerable attention. Pulp and paper industry wastewaters, namely spent sulfite liquors, are potential material for conversion to useful products. Following pretreatment to remove biological oxygen TABLE IX BY ACETONE-BUTANOL PRODUCTION

Organism

Substrate (glliter)

Clostridium acetobutylicum ATCC 824

Glucose (70) Xylose (70) Arabinose (70) + acetate (30 mM) C. acetobutylicum Complex extruded corn SA-I

CLOSTRIDIUM

Butanol Acetone (glliter) (glliter) 8.9

15 3.3

-

SPECIES

Ethanol (giliter)

Ref. Ounine et al (1983)

10.5

4.4 0.9 4.5

13.9

6.3

1.5

Lovittetal. (1988)

10.3

7.4

0.1

1.5

-

2.1

14.8

2.3

1.1

Marchal et al. (1984) Idemitsu Kosan Co. (1983) Soni et al. (1982)

-

(100)

C. acetobutylicum Wheat straw IFP 921 (156) Clostridium sp. Complex AH-1 cellulose (10) Saccharoperbu tyl Cellulosic acetonicum hydrolysate (60)

MICROBIAL PENTOSE UTILIZATION

121

demand to 45% and supplementing nitrogen and phosphorus sources, C. butylicum was able to utilize more than 80% of the sugars (Volesky and Szczesny, 1983). Earlier studies, based on a promising strain of C. butylicum, indicated problems associated with the fermentation of wood hydrolysate (Leonard and Peterson, 1947). A limeneutralized hydrolysate (pH 6 . 5 ) ,containing 3% sugars, was used with solvent yields of 25-38Oh. Few attempts have been made to convert cellulose and hemicellulose complexes directly to acetone-butanol. Petitdemanage et al. (1983) suggested the use of a coculture of C. acetobutylicurn with cellulolytic clostridia. However, solventogenesis was not achieved because of low concentrations of generated sugars. Similar results were obtained by sequential fermentation of cellulose using C. thermocellum followed by C. acetobutylicum with butyrate added to induce solventogenesis. A simultaneous saccharification and fermentation has also been studied to produce butanol from alkali-pretreated wheat straw using Trichoderma reesei and C. acetobutylicum (Marchal et al., 1984). The final butanol concentration reached was 10.7 g/liter from 140 g/liter of straw. A simultaneous utilization of both hemicellulose and cellulose components was observed since pentoses did not accumulate during fermentation. C. acetobutylicum has a ,&-1,4-glucanglucohydrolase and cellobiase, but lacks an active cellobiohydrolase to hydrolyse crystalline cellulose. rDNA technology and protoplast fusion could be applied to transfer the genes of this enzyme. The key element in any bioprocess is the microorganism. The maximum accumulation of 2.5%total solvent before shunting of bacterial biosynthetic activity indicates solvent inhibition. Thus a microbe with increased product tolerance is essential.

c. 2,3-BUTANEDIOL PRODUCTION Much of the work on butanediol fermentation was performed during World War I1 when it was discovered that 2,3-butanediol could be chemically converted to P,S-butadiene, a major constituent of bunarubber, and several other chemicals (Rosenberg, 1980). Interest in butanediol fermentation was renewed in the last 10 years because of the microorganism’s ability to ferment pentose sugars and the availability of lignocellulosic hydrolysate as a cheap substrate. However, the market for the product is questionable (Lovitt et aI., 1988). Bacillus polymyxa is capable of fermenting starch, hexoses, and pentoses, while K. Pneurnoniae can only utilize monosaccharides. Bacillus polymyxa produces large amounts of ethanol and is also unstable, losing its ability to produce butanediol (Ledingham and Neish, 1954).

122

PRASHANT MISHRA AND AJAY SINGH

In contrast, Klebsiella is capable of producing higher yields and more stable fermentations (Jansen and Tsao, 1983; Lovitt et a ] . , 1988). Butanediol exists in three stereoisomeric forms D( -), L( +), and meso, and different organisms yield one or more of these isomers in fermentations. The physical and chemical properties and chemical reactions of 2,3butanediol are discussed extensively by Ledingham and Neish (1954). Net equations for the conversion of xylose and glucose are: Xylose -+$20, + gNADH, Glucose --+ 2C0, + NADH,

+ 9ATP + Sbutanediol + 2ATP + butanediol

On a mass basis, the butanediol yield from both xylose and glucose is 50%. The theoretical maximum molar yield of butanediol from pentoses is 0.83 and hexoses is 1.0. Table X presents a comparison of 2,3-butanediol fermentations in various reactor systems. In batch cultures of K. pneumoniae, butanediol concentrations of 30-65 g/liter with corresponding yields of 0.310.43 g/g substrate from various carbon sources, including xylose, have been reported (Fulmer et al., 1933; Freeman and Morrison, 1947). However, the major disadvantage identified in the batch process was the low reactor productivity, mainly because of the long fermentation period required to attain high cell densities to result in a rapid reaction rate (Jansenand Tsao, 1983). Further, the final butanediol concentration is limited by the maximum initial substrate concentration that can be

TABLE X 2,3-BUTANEDIOLPRODUCTION BY PENTOSE-FERMENTINGBACTERIA Organism

System

Substrate

Butanediol (giliter)

Yield (gig)

Ref. ~

Aerobacter aerogens

Klebsiella pneumoniae

Batch

Xylose

29

0.29

Batch

G1ucose

15

0.33

Batch

Xylose

30

0.31

Batch

Wood hydrolysate Glucose

13.3

0.29

99

0.37

83

0.42 0.33

Fed-batch Fed-batch Immobilized cells

Xylose Xylose

31

~

~~~~

Jansen et a]. (1984b) Sablayrolles and Goma (1984) Jansen (1982) Grover et al. (1990) Olson and Johnson (1948) Jansen (1982) Chambers et 01. (1979)

MICROBIAL PENTOSE UTILIZATION

123

tolerated by the bacteria. Such problems can be sorted out by carrying fed-batch fermentations. Butanediol concentrations of 83 and 99 g/liter with corresponding yields of 0.42 and 0.37 g/g from xylose and glucose, respectively, have been obtained by Jansen (1982) and Olson and Johnson (1948). Chambers et al. (1979) studied the pentose fermentation in a closed loop-immobilized batch cell reactor. They were able to achieve the conversion of a 100 g/liter xylose solution at a rate three times greater than that needed for a conventional batch reaction by immobilizing the cells on 4 inch Raschig rings. However, oxygen transfer to the immobilized cells was the problem. Much higher reactor productivities were reported by Pirt and Callow (1958) in continuous culture. These authors reported a butanediol productivity of 2.7 g/liter/hr at a dilution rate of o.l/hr. Increasing the dilution rate to 0.2/hr further increased productivity to 4.6g/liter/hr, but the butanediol concentration fell to 23 g/liter. Lower butanediol yields obtained during continuous culture might be due to the loss of some sugars in the product stream. Moreover, a high final product concentration may be difficult since the entire process is continually subjected to product inhibition to the maximum extent (Jansen and Tsao, 1983; Jansen et al., 1984a). Recently, Grover et al. (1990) studied the production of 2,3-butanediol from a wood hydrolysate by K. pneumoniae. They achieved a butanediol concentration of 1 2 g/liter with a corresponding yield of 0.27 g/g sugars present in 48 hr. Adding 1%(w/v) malt extract to the medium further enhanced the butanediol production to 13.3 glliter (yield, 0.23 g/g). Earlier A. aerogens had been reported to utilize all major sugars (hexoses, pentoses, and certain disaccharides) and uronic acid derived from hydrolysates of hemicellulosic and cellulosic materials to produce 2,3butanediol (Yu et al., 1982; Saddler et al., 1983, 1984). D. ORGANIC ACIDPRODUCTION

Many bacteria produce a variety of acids during the fermentation of pentoses, pentitols, and polysaccharides (Table XI). The major products of these fermentations are ethanol, CO,, and acetic, lactic, succinic, and formic acids. The relative amounts of acidic products formed depend on the type of organism, substrate, and process conditions. In some cases formic acid may be replaced by equivalent amounts of H, and CO, (Rosenberg, 1980). Facultative anaerobic bacteria such as Escherichia coli can carry out a mixed acid fermentation of glucose and xylose (Reynolds and Werkman, 1937). Acidic conditions favor the production of H, and CO, at the

124

PRASHANT MISHRA AND AJAY SINGH TABLE XI ORGANISMS CAPABLE OF FERMENTING PENTOSESUGARS TO ORGANIC ACIDS

Organism Bacteria Bacillus macerans Clostridium thermocelllum C. acetobutylicum C. thermoceticum Lactobacillus casei Leuconostoc sp. Ruminococcus albus Therrnoanaerobiurn brocki Fungi Aspergillus terreus Fusarium oxysporum Rhizopus orrhizus

Acetic acid

Lactic acid

+ +

+ +

Succinic acid

Formic acid

Fumaric acid

Itaconic acid

+ +

expense of formic acid. Spirochaeta stenostrepta Z1 ferments xylose, arabinose, and ribose in addition to several hexoses (Canale-Parola et al., (1967; Hespell and Canale-Parole, 1970) to acetic acid, lactic acid, ethanol, H,, and CO,. Ruminococcus albus, a strict anaerobe, ferments cellulose and xylan, a polymer of D-xylose, to acetic acid, ethanol, formic acid, and small amounts of succinic and lactic acid (Bryant et al., 1958). Clostridium thermocellum and C. thermocelluloceum also ferment cellulose, xylose, and arabinose to ethanol and mixed acids like formic, lactic, acetic, and succinic acid (McBee, 1950). Clostridium acetobutylicum carries out organic acid and solvent fermentation under different environmental conditions (Rosenberg, 1980). Bacillus macerans (B. acetoethylicum) conducts a modified mixed acid fermentation of hexose and pentoses with major products, ethanol, acetic acid, and acetone, and minor products, lactic and formic acid (Rosenberg, 1980). Species of Leuconostoc and Lactobacillus degrade pentose sugars through a heterolactic pathway (Doelle, 1975;Gottschalk, 1986).Ethanol is formed from acetyl phosphate and lactate is derived from the Embden-Meyerhof pathway. The product yields depend on specific growth conditions, and acetate and formate are also formed. De Vries et al. (1970) studied Lactobacillus casei for the production of lactate, acetate, formate, and ethanol in a continuous culture system. A heterolactic bacterium, Thermoanaerobium brockii, produces lactate as the

125

MICROBIAL PENTOSE UTILIZATION

major product in a high yeast extract medium and ethanol as the major product in a low yeast extract medium (Lamed and Zeikus, 1980). Datta (1981) carried out an anaerobic acidogenic fermentation of complex lignocellulosic biomass into acetic acid using a nonsterile mixed culture, fermentation process. Acetic acid equivalents were produced with an average yield of 83.7%. During the acidogenic fermentation of corn stover, all major components (cellulose, hemicellulose, lignin, and pectins) are utilized. This simple fermentation scheme (nonsterile, anaerobic, unstirred, 25°C) could produce a single class of useful compounds (C2-C6 volatile organic acids) directly from a complex lignocellulosic feed with high yield and specificity. Recently Brownell and Nakas (1991) carried out a homoacetate fermentation of pentose sugars with C. Thermoaceticum (Table XII). In batch culture, C. thermoaceticum produced 14 g/liter of acetic acid with a corresponding yield of 0.76 g/g xylose in 48 hr at 55°C under a head space of 100% CO,. In fed-batch fermentations, this organism produced 42 g/liter acetic acid after 116 hr. When the concentration of xylose was maintained at 2% from an acid hydrolysate of oat spelt xylan and poplar hemicellulose, C. thermoaceticum produced 14.4 and 11.5 g/Iiter acetic acid in 72 hr. Fusarium oxysporum DSM841 utilized a wide range of substrates, including cellulose, hemicellulose, starch, xylose, arabinose, galactose, and glucose, to produce ethanol and acetic acid (Kumar et a]., 1991a,b). Fusarium oxysporum DSM 841 produced 1 2 g/liter acetic acid and 3.6 gl/ethanol from potato wastes (cellulosic waste from starch industries) with corresponding yields of 0.36 and 0.1 g/g, respectively (Singh

TABLE XI1 BIOCONVERSION OF PENTOSE SUGARS TO ACETICACID

Organism

Substrate

Acetic acid Yield (g/liter) (g/g)

Productivity (g/liter/hr)

Ref.

~

Clostridium thermoaceticum

Fusarium oxysporum

Xylose

14.1

0.76

0.20

Oat spelt xylan Popler hemicellulose hydrolysate Xylose

14.4 11.5

0.72 0.58

0.20 0.16

8.8

0.24

0.12

7.0 8.0

0.19 0.22

0.09 0.11

Brownell and Nakas (1991)

Singh et al. (1991)

Arabinose Galactose

126

PRASHANT MISHRA AND AJAY SINGH TABLE XI11

FUMARIC ACIDPRODUCTION FROM XYLOSE BY IMMOBILIZED CELLS OF Rhizopus arrhizusa ~

~

~

Xylose concentration

~~~~~~~

Residence time

C : N ratio

I%)

Fumaric acid (giliter)

Productivity (mg/liter/hr)

10.25 9 6 3 1.75

160 188 160 188 160

10 6.5 5 6.5 10

16.4 15.3 9.6 4.1 3.6

67 71 66 57 87

Adapted from Kautola and Linko (1989).

et al., 1992b). In a fed-batch culture, the maximum concentration, yield, and productivities of acetic acid were 22.5 g/liter, 0 . 3 8 g/g, and 0.09 g/ literihr, respectively (Kumar et a]., 199'lb). Fumaric acid, used in the manufacture of sizing resins for the paper industry, can be produced biochemically from pentoses. Kautola and Linko (1989) studied fumaric acid production from xylose using immobilized cells of Rhizopus arrhizus (Table XIII), The highest fumaric acid concentration with immobilized cells reached 16.4 g/liter at loo/o initial xylose, had a C :N ratio of 160, and a residence time of 10.25 days with a volumetric productivity of 6 7 mg/liter/hr. The volumetric productivity increased to 8 7 mg/liter/hr when the residence time during the batch was 1 . 7 5 days. About 3.5 times higher fumaric acid levels were obtained with immobilized R. arrhizus system when compared to a similar free cell system. TABLE XIV ITACONIC ACIDPRODUCTION FROM XYLOSE AND GLUCOSE BY IMMOBILIZED CELLS OF Asergillus terreus Reactor

Substrate

Itaconic acid (g/liter)

Yield (g/g)

Productivity (g/liter/hr)

Ref.

Batch Batch Batch Batch

Xylose Glucose Glucose Glucose

30 30 39.4 51

0.45 0.55 0.64 0.51

0.20 0.32

Repeated batch Continuous

Xylose

13.8

0.23

0.12

Kautola et al. (1985) Kautolaet 01. (1985) Pfeifer et a!. (1952) Waszczuk and Marciniak (1974) Kautola et al. (1985)

Xylose

5

0.08

0.56

Kautolaet al. (1985)

-

-

MICROBIAL PENTOSE UTILIZATION

127

Kautola et al. (1985) used immobilized Aspergillus terreus cells for the production of itaconic acid, an important intermediate in polymer production, in a batch, repeated batch, and a continuous column reactor (Table XIV). The maximum volumetric productivities with D-xylose as substrate obtained in a batch, repeated batch, and continuous culture were 0.20, 0.06, and 0.56 g/liter/hr, respectively. VI. Factors Affecting Pentose Fermentation

A number of cultural process parameters have been identified by several workers that have significant effects on solvent or organic acid production. In considering approaches to improve product concentration and yield, several characteristics of pentose-fermenting organisms are of particular interest and require attention. A. ETHANOL PRODUCTION Various physical, nutritional, and other parameters have been studied that may have appreciable effects on the ethanol production by yeast, fungi, and bacteria. 1. pH

If pH is uncontrolled, it changes with the extent of fermentation. Thus to assess its effect on ethanol production, the pH must be controlled at a set value over the entire fermentation period. The optimum choice of initial pH may also depend on the type of medium being fermented, the type of pH control employed, and the microbial strain carrying out the process. The optimum pH value for yeast P. tannophilus is between 2.5 and 5.0 (Silinger et al., 1982; Debus et al., 1983), between 3.5 and 4.5 for C. shehatae (du Preez et al., 1984), and between 4.0 and 5.5 for P. stiptis (Dellweg et al., 1990). The initial pH values generally employed by various workers for fungal fermentation are within the range of 5.0-6.0 (Suihko and Enari, 1981; Batter and Wilke, 1977; Rosenberg et al., 1981; Ueng and Gong, 1982; Rao et a]., 1983; Kumar et a]., 1991a). The optimum pH value for F. oxysporum lies between 5.0 and 5.5 and between 5.0 and 6.0 for N. crassa. However, Paecilomyces sp. NF1 has a wide range of pH optima which varies from 2.2 to 7.0 (Wu et a]., 1986). Bacterial fermentation products depend on the pH at which fermentation is conducted. Acidic conditions favor the production of neutral products at the expense of acids. The optimum pH for B. macerans growth has been reported to be between 6.0 and 8.0 (Rosenberg, 1980). The pH range for the growth of thermophilic bacteria are those

128

PRASHANT MISHRA AND AJAY SINGH

of typically neutrophilic organisms, with optima between 6.0 and 7.0 and a range of 4.5-8.0. 2. Temperature

The effect of temperature depends on the microbial strain employed for the fermentation (Singh et al., 1992a). The optimum temperature for most pentose-fermenting yeasts is between 30 and 32°C (Silinger et al. 1982; du Preez et al., 1984). However, P. stiptis (Kurtzman, 1984) and P. tannophilus (Barnett et al., 1983; Kurtzman, 1984) can grow at 37°C and 37-42"C, respectively. The Mold Paecilomyces sp. NF1 has an optimum temperature range of 30-37°C (Wu et al., 1986), whereas ethanol production by N. crassa is optimum between 2 8 and 37°C (Deshpande et al., 1986). The optimum temperature for most of the Fusarium strains is 30°C (Singh and Kumar, 1991). In mesophilic pentosefermenting bacteria, the optimum temperature range is 20-32OC for K. pneumoniae and 25°C for C. freundii (Banerjee, 1985). The optimum temperature for thermophilic bacteria growth ranges from 60 to 70°C (Wiegel, 1980). 3. Nutritional Factors

Besides physical factors like pH and temperature, several nutritional factors also affect ethanol fermentations to a great extent. In P. tannophilus, growth requires a source of thiamine and biotin which can be made available by supplying yeast extract to the growth media (Dellweg et aI., 1984; Silinger et a]., 1987). Yeast extract also improves cell growth and ethanol production of P. stiptis and C. tropicalis (Skoog and HahnHagerdal, 1988).Candida tropicalis requires 10-20 g/liter yeast extract and peptone for ethanol production (Skoog and Hahn-Hagerdal, 1988). Nitrogen and carbon sources influence the activity of pentose phosphate pathway enzymes and consequently the cell and ethanol yield obtained (Bruinenberg et a]., 1983). However, thorough nutritional studies have not been done on yeast-fermenting pentose sugars. A variety of inorganic and organic nitrogen sources were evaluated by various workers in ethanol fermentation studies with filamentous fungi. Peptone was found to be the best organic nitrogen source [Gong et al., 1981d; Rao et al., 1983; Christakopoulos et al., 1989; Kumar et al., 1991b). In F. oxysporum, nitrogen limitation leads to cell death (Enari and Suihko, 1984). Trace elements such as Fe, Zn, Cu, and Mn and growth factors like thiamine increase the growth of P. anceps in submerged culture (Perlman, 1949). An ammonium sulfate concentration of 1 g/liter is the best nitrogen source for K. pneumoniae (Banerjee, 1985). This preference may be due to the fact that this compound also provides sulfate

MICROBIAL PENTOSE UTILIZATION

129

ions which are important for the synthesis of methionine, cysteine, coenzyme lipoic acid, and CoA (Thiamine, 1963; Tempest, 1981).Trace metal salts such as EDTA, TPP, CuSO,, CoCl,,CaCl,, and MnSO, do not support ethanol formation from D-xylose by K. pneumoniae (Banerjee, 1985).Most of the thermophilic bacteria can be grown on defined media, but growth rates are reduced over that in complex media. Thermoanaerobacter requires yeast extracts which could not be replaced by tryptone, casein hydrolysate, beef extract, or ashed yeast extract (Wiegel et a]., 1979; Wiegel and Ljungdahl, 1981). The requirements of thermophilic clostridia vary. Clostridium thermocellum grow on a salt medium supplemented with biotin, vitamin B,,, pyridoxamine, and p-aminobenzoic acid (Ng et al., 1977). Clostridium thermohydrosulfuricum requires yeast extract (Wiegel et a]., 1979). 4. Oxygenation

The rate of utilization of available carbohydrates and the eventual conversion to ethanol are significantly affected by aeration in yeasts and filamentous fungi (Table XV). Limitation of oxygen can be associated with ethanol accumulation. With respect to oxygenation, P. tannophilus is the most thoroughly studied organism (Silinger et al., 1987). It produces cell mass under aerobic conditions, accumulates xylitol under anaerobic conditions, and produces ethanol under oxygen limitations (Debus et al., 1983; Schvester eta]., 1983; Schneider et a]., 1985). However, there is one report where P. tannophilus produced ethanol aerobically as well as anaerobically (Silinger et al., 1982). Another interesting observation with P. tannophilus is the reassimilation of ethanol when oxygenation is increased (Maleszka and Schneider, 1982). A similar observation was made with C. tropicalis. When the xylose concentration reaches a certain level, the organism prefers ethanol produced as a carbon source (Hahn-Hagerdal et al., 1985). Du Preez et al. (1984) compared the effect of oxygenation in P. tannophilus and C. shehatae. With an increased oxygen limitation, the ethanol yield changes from 0.33 to 0.28 glg with C. shehatae and from 0.24 to 0.02 with P. tannophilus, thus indicating that C. shehatae is less dependent on the degree of oxygenation. Similarly, C. tropicalis is also marginally influenced by the degree of oxygenation (Skoog and Hahn-Hagerdal, 1988).

Under anaerobic conditions, the absence of oxygen is the sole factor in controlling the rate and extent of cell growth. In the absence of oxygen, growth either fails to occur or is restricted. However, D- xylose is still metabolized by some strains under these conditions with ethanol or xylitol as the product (Jeffries, 1982, 1983; Bruinenberg et al., 1984;

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PRASHANT MISHRA AND AJAY SINGH

Amin et a]., 1988; Prior et al., 1988).In a continuous culture, as oxygen limitation increases, growth decreased, and ethanol production increases in P. tannophilus (Mutze and Wandrey, 1983; Amin et al., 1988). This suggests that the specific rate of oxygen utilization is an important factor in determining the amount of ethanol accumulated (Mahmourides et a]., 1985; Chung and Lee, 1986). Partitioning of carbon from D-xylose between growth and product formation is also affected by oxygenation. Over the intermediate range of aeration rates, as rates increase, carbon is shifted generally to growth at the expense of ethanol accumulation (Schneider et al., 1983; Schvester et al., 1983; Watson et al., 1984b). Both volumetric and specific ethanol production rates are influenced by changes in aeration rates. This is particularly true with P. tannophilus and C. shehatae (Baillargeon et a]., 1983; du Preez et a]., 1984). Table XV shows the effects of aeration on product yield in yeasts and the filamentous fungus, Fusarium. In filamentous fungi, ethanol is accumulated only under low aeration [oxygen limited) conditions (Singh and Kumar, 1991; Singh et a]., 1992b). A series of experiments conducted with F. oxysporum studied the effects of aeration on ethanol production (Enari and Suihko, 1984; TABLE XV

EFFECTSOF AERATION ON D-XYLOSEFERMENTATION BY YEAST AND MOLD Parameter/ condition Biomass yield Aerobic Semiaerobic Anaerobic Ethanol yield Aerobic Semiaerobic Anaerobic Xylitol yield Aerobic Semiaerobic Anaerobic Acetate yield Aerobic Semiaerobic Anaerobic Reference

Pichia stiptis

Candida shehatae

Candida tenuis

0.51 0.12 0.06

0.49 0.08 0.02

0.31 0.09 0.02

0.25 0.014 0.015

0.38 0.13 0.07

0 0.43 0.38

0 0.43 0.38

0 0.25 0.27

0.10 0.28 0.26

0.04 0.23 0.11

0 0

0 0.13 0.23

0.03 0.21 0.23

0.17 0.30 0.30

-

-

-

0 0.24 0.16

Ligthelm et al. (1988)

Singh et al.

0.09

du Preez et 01.

du Preez et al.

du Preez et al.

(1989)

(1989)

(1989)

-

Pachysolen tannophilus

-

Fusorium oxysporum

(1992c)

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Singh et a]., 1 9 9 2 ~ )Ethanol . production appears to be growth associated and aeration rates of 0.04-0.06 vvm are suitable for product formation. Shake flask studies on the effect of aeration rates on D-xylose fermentation and metabolism in F. oxysporum show that oxygen-limited conditions stimulate the biosynthesis of key D-xylose-catabolizing enzymes, thereby product formation increases severalfold. Unlike yeast and fungi, bacterial fermentation of pentose sugars for ethanol production can be operated without critical aeration. The facultative anaerobe K. pneumoniae does not produce any solvent under aerobic conditions; however, ethanol production is greatly enhanced under anaerobic conditions (Yu and Saddler, 1982). Banerjee (1985) also observed that an aerobic environment causes deleterious effects on ethanol formation by K. pneumoniae and C. freundii, although oxygen is required for initial growth. The profile of dissolved oxygen (DO) shows that if the media is not degassed to remove oxygen, K. pneumoniae utilizes the total DO within the initial 5 hr of fermentation. Using reducing agents at the initial stage to remove DO from the broth leads to the complete inhibition of growth and ethanol production. Thus DO is an important factor in initiating D-xylose fermentation. This indicates the requirement of oxygen for the synthesis of pentose phosphate enzymes (Doelle, 1975). In sequence with the drop in the DO level, the redox potential of fermentation broth shows a steady-state decrease up 400 mV which coincides with the maximum concentration of ethanol produced by K. pneumoniae (Banerjee, 1985). As a result, the D-XylOSe fermentation by K. pneumoniae occurs under completely anaerobic conditions. Such anaerobicity has also been reported in the case of obligate anaerobes which exhibit a redox potential of less than -200 mV (Jorlik and Willet, 1976). In Clostridia, the oxidation of pyruvate to acetyl-CoA by the coenzyme ferrodoxin occurs under a redox potential of -400 mV (Volesky and Szczesny, 1983). 5. Lipids

The peak concentration and yield of ethanol in cultures of P. tannophilus (Dekker, 1986) are increased by the addition of a low concentration of lipids (34 mg/liter ergosterol, 34 mglliter linoleic acid, and 5.2 g/ liter Tween 80). Lipids increase the ethanol concentration from 8.5 to 13.25 g/liter with corresponding yields of 0.2 to 0.32 g/g, on the basis of D-xylose consumed. This enhanced ethanol yield is suggested to be due to the increased ethanol tolerance by the addition of lipids. Neirinck et al. (1984),however, did not observe any effect of lipid supplementation on anaerobic incorporation with either D-xylose or D-glucose.

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6. Metabolic Inhibitors

Adding metabolic inhibitors to the media to suppress the by-product formation and to enhance ethanol yields in yeast and filamentous fungi have been reported by several workers. Adding sodium azide to the media increases ethanol yields in C. tropicalis and P. tannophilus while xylitol formation decreases (Hahn-Hagerdal et al., 1985; Lohmeir-Vogel and Hahn-Hagerdal, 1985; Manderson and Newland, 1987).Other inhibitors tested, trichlorophenol and dinitrophenol, also resulted in similar observations. Adding azide, dinitrophenol, and polyethylene glycol (PEG) to the medium also shifts product formation from acetate to ethanol in F. oxysporum without affecting the utilization of xylose (Singh et al., 1991). 7. Inhibitors Present in the Lignocellulosic Hydrolysate Several inhibitors may be present in the lignocellulosic hydrolysate prepared using acid catalysis (Jeffries, 1984;Tran and Chambers, 1985). Some are derived from noncarbohydrate materials and others are formed by the breakdown of carbohydrates during hydrolysis, e.g., furfural. Heavy metal ions such as Cr, Cu, Fe, and Ni may also be present, probably as a result of corrosive metal parts of the apparatus used (Watson et ai., 1984a). Acetic acid is formed commonly in wood hydrolysates. A 5 g/liter concentration of acetic acid can be inhibitory to P. tannophilus (Lee and McCasky, 1983) and greater amounts can be present in some hydrolysates. B. ACETONE AND BUTANOLPRODUCTION 1. pH

Acetone and butanol are the major fermentation products of C. acetobutylicum when grown in a continuous culture under a phosphate limitation of pH 4.3 (Peterson and Fred, 1932; Bahl et a]., 1982; Andersch et al., 1983). At pH values above 5.5, however, an exclusive acetate-butyrate fermentation is carried out by this organism under phosphate, ammonia, or glucose limitation (Bahl et al., 1982; Andersch et a ] . ,1982). In batch fermentation, the concentration of acetic acid and butyric acid rises initially, at a pH of about 4.0, the acid concentration begins to fall, and the amount of acetone and butanol increases with the rise of pH. 2. Temperature Most acetone-butanol producers are mesophilic and display temperature optima for fermentation between 30 and 37°C. At lower tempera-

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tures, solvent production by C. acetobutylicum decreases, while at elevated temperatures, the acetone yield decreases but butanol yields remain unaffected (McNeil and Kristensen, 1985). A thermophilic strain capable of producing butanol from cellulose at 50-65°C has also been reported (Idemitsu Kosan Co., 1983). 3. Nutritional Factors

Several nutritional factors affect the acetone-butanol fermentation. Most of the acetone-butanol-producing strains require p-aminobenzoic acid and biotin for growth in addition to sugar containing a mineral salts medium (Monot et al., 1982). Thiamine has also been used in a defined medium in some cases (Long et al., 1983). Clostridium beijerincki requires multiple amino acids and vitamins for growth (Prescott and Dunn, 1940; Mes-Hartree and Saddler, 1982). Initial acidic fermentation products formed are remetabolized and converted to more reduced neutral products (Speakman, 1920; Rosenberg, 1980). Adding butyric acid to the fermentation media increases the yield of butanol. Johnson et al. (1933) reported that the addition of acetate increases the yield of acetone. This was later confirmed by Wood et al. (1945) who added I3C-labeled acids to the fermentation and demonstrated incorporation of the label into neutral products. While 85% of the butyric acid label is found in the butanol fraction, only 15-19% of the added acetic acid label is found in acetone and isopropanol and 50% in the butanol fraction. Different substrates also affect fermentation parameters of C. acetobutylicum such as growth rate and solvent production ratio (Long et al., 1983). Generally, growth and substrate consumption rates with xylose, arabinose, or galactose are low when compared to glucose, mannose, or cellobiose. The solvent production ratio with pentose sugar is 1 : 2 : 5 (ethanol : acetone :butanol), while ratios of 1 : 4 : 10 are obtained with the second sugar group. However, in one case (Ounine et al., 1983) glucose and xylose were fermented and yielded similar solvent ratios but the xylose consumption rate (0.33 giliterihr) was much lower than that of glucose (0.68 g/liter/hr). 4. Oxygenation

As obligate anaerobes, butanol-producing organisms require anaerobic conditions. A low redox potential (below -250 mV) is essential for the acetone-butanol fermentation (Lovitt et al., 1988). Clostridium acetobutylicum NCIB 8052 can detoxify molecular oxygen by NADH without forming H,O, (O’Brien and Morris, 1971). This organism grows anaerobically and produces solvents at a redox potential of + 370 mV when poised by potassium ferrocyanide. Vegetative cells survive for

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several hours in the presence of oxygen. Hongo’s work (1957)with redox dyes showed that higher yields of butanol can be achieved by adding 5 mM neutral red into the fermentation system. This effect was later ascribed to the presence of neutral red-linked uptake hydrogenase and pyridine nucleotide reductase activity in cells (Lovitt et a ] . , 1988).

c. 2,3-BUTANEDIOL PRODUCTION Many environmental factors affect butanediol production because of the versatality of the metabolism of 2,3-butanediol-producingfacultative anaerobic bacteria. 1. pH

The fermentation balance is significantly influenced by pH. Generally the yield of butanediol reaches maximum in the pH range of 5.0-6.0 but falls to near zero above pH 7.0 (Jansen and Tsao, 1983).The ratio of butanediol to acetoin varies from near 25 between pH 5.2 and 6.0to near zero at pH 7.6.Above pH 7.0,there is a rising formic acid concentration and a falling CO, level. This suggests that the cell maintains its NAD/NADH balance by reducing CO, to formic acid when it cannot produce butanediol (Neish and Ledingham, 1949;Pirt and Callow, 1958; Jansen and Tsao, 1983).Acetic acid and lactic acid production is minimum below pH 5.0 but increases rapidly above pH 6.0 (Neish and Ledingham, 1949). 2. Temperature

With K. pneumoniae, the optimum temperature for growth and substrate uptake rate is between 37 and 38°C (Pirt and Callow, 1958;Topiwala and Sinclair, 1971;Esener et a]., 1981). Substrate utilization for endogenous metabolism increases steadily as the temperature increases from 25 to 40°C.Pirt and Callow (1958)obtained maximum butanediol production between 35 and 37°C with K. pneumoniae, whereas Olson and Johnson (1948)obtained maximum yields of butanediol at 30°C. 3. Nutritional Factors

Sugar concentration has a significant effect on butanediol production and reaction rates (Fulmer et al, 1933;Long and Patrick, 1963;Esener, 1981).At higher sugar concentrations, both yield and rate decrease. The optimum sugar concentration often depends on the particular substrate used as the carbon source. Long and Patrick (1963)suggested that as the sugar concentration in raw material increases, the level of accompa-

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nying toxic material also increases, resulting in poor substrate utilization. When an acid-hydrolyzed wheat wash medium is employed, the butanediol yield falls and carbohydrate utilization becomes incomplete (Jansen and Tsao, 1983). The apparent substrate inhibition could be explained by a decreasing water activity and changing osmolarity (Esener, 1981). To overcome these problems, fed-batch and continuous fermentations have been investigated (Pirt, 1975; Flickinger, 1980; Yu and Saddler, 1982). The exogenous addition of acetate has been found to improve butanediol yields, presumably because it induces and is utilized by the butanediol-forming pathway (Stahly and Werkman, 1942; Stormer, 1968b). Oxygen supply rate is another factor which significantly affects end product formation, even though 2,3-butanediol is a product of anaerobic metabolism (Stahly and Werkman, 1942; Ledingham and Neish, 1954). Limited amounts of oxygen increase the productivity as higher cell concentrations of oxygen are maintained in chemostats. Higher levels of oxygen shift the ratio of end products from butanediol to acetoin (Sablayrolles and Goma, 1984). Jansen (1982) carried out a study on the effect of aeration rates on cell yield and butanediol production. At a maximum oxygen transfer rate (OTR), the cell mass is highest and butanediol yield is lowest. By decreasing OTR to zero, butanediol increases toward the theoretical maximum while the butanediol reaction rate significantly decreases due to diminishing cell yields (Jansen and Tsao, 1983). Yu and Saddler [1982) reported that K. pneurnoniae metabolizes glucose anaerobically, but that some oxygen is required for xylose fermentation. The availability of oxygen also determines the amounts of particular products excreted. With a high OTR, acetate is the principal product (Pirt, 1957; Harrison, 1967). When the specific uptake rate decreases, butanediol becomes the predominant product (Pirt and Callow, 1958). When the oxygen supply is cut off, ethanol production increases at the expense of butanediol (Jansen and Tsao, 1983).

Another important parameter affecting 2,3-butanediol production is water activity. It is an expression of the water concentration that depends on the molar concentration and activity coefficient of each solute. Increasing the solute concentration decreases the water activity of a solution (Pirt, 1975). At a water activity of 0.985, the growth rate of Klebsiella sp. is 50% optimal and becomes less than 10% optimal at water activities lower than 0.975 (Esener, 1981). Species of Klebsiella [Aerobactor) are not as osmotolerant as some other organisms (Scott, 1953) and that is the reason why very high sugar concentrations in butanediol processes are not suitable.

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D. ORGANIC ACIDPRODUCTION

Organic acid fermentations are significantly affected by the pH of the culture medium. Generally an acidic condition favors the production of neutral products at the expense of organic acids, while alkaline conditions have the opposite effect (Rosenberg, 1980; Volesky and Szczesny, 1983). Acetic acid production is both growth and nongrowth associated when either xylose or glucose is used as a substrate (Sugaya et a]., 1986). However, product yield significantly decreases after a certain substrate level. Maximum substrate conversion to acetic acid is obtained at 2% glucose (Sugaya et a]., 1986). A further increase in substrate concentration decreases substrate conversion as well as acetic acid yield. The utilization rate of available carbohydrates and the eventual conversion to products are considerably affected by aeration (Skoog and Hahn-Hagerdal, 1988). As discussed previously, ethanol production by yeast and fungi is associated with the rate of oxygen supply. The production of acetic acid by F. oxysporum is greatly affected by aeration rates. A decrease in aeration rate from 0.1 vvm to 0.02 vvm. significantly increased the acetic acid yields from 0.09 gtg substrate to 0.28 g/g substrate. Similarly, increasing the amount of oxygen (1%)aeration significantly increases ethanol with a decrease in acetic acid formation in F. oxysporum VTT-D-80134 (Enari and Suihko, 1984). Fumaric acid production by immobilized cells of R. arrhizus is found to be much better than with free cells (Kautola and Linko, 1989). Fumaric acid yields and productivities are considerably influenced by the C :N ratio and residence time in repeated batch fermentations of xylose. Including MgSO, in the fermentation media is necessary for the stable production of itaconic acid by A. terreus (Kautola et a]., 1985).In continuous cultures with the same organism, ammonium nitrate is a preferable nitrogen source over ammonium sulfate. VII. Product Tolerance

The fermentation rates in many bioprocesses depend on the cellular resistance of microbes to the end product. Pentose metabolism yields a broad spectrum of products, including ethanol, acetone, butanol, butanediol, and organic acids, which depends on the type of microorganism employed as well as the culture conditions used. The effect of ethanol on the inhibition of growth and metabolism of bacteria and yeasts has been widely studied (Ingram and Buttke, 1984; Ingram, 1986;

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van Uden, 1989). Ethanol tolerance depends on the type of microorganism (Ingram, 1986).Lactobacillus homohiochii and Lactobacillus heterohiochii are the most alcohol-tolerant microorganisms and are capable of growth above 16% (w/v) (Kitahara et al., 1957; Demain et a]., 1961). Saccharomyces and Zymomonas also tolerate ethanol at relatively high concentrations (8-12Y0, w/v),whereas E.coli tolerate ethanol at concentration up to 5% (w/v). However, thermophilic bacteria, e.g., Clostridium thermohydrosulfuricum, tolerate ethanol up to only 1.5% (w/v) (Ingram, 1986). In addition, ethanol tolerance also varies within species of the same genera. For example, ethanol tolerance among the genus Saccharomyces is sake yeasts > wine yeasts > distillers yeast > brewers yeast (Casey and Ingledew, 1986; Rose, 1987). There are two basic hypotheses for the mechanism of alcohol inhibition of fermentation: (1)damage to cell membrane and (2) end product inhibition of the glycolytic enzymes. It is apparent from in vitro studies that enzymes of the glycolytic pathway are resistant to the ethanol concentration produced during fermentation (Miller et aI., 1982). In addition, for many ethanologenic organisms the potency of alcohol as an inhibitor has been correlated with lipid solubility (Ingram and Buttke, 1984), implying that the hydrophobic site of the membrane is a prime target of ethanol inhibition. Results obtained so far also accredit the cell membrane as the major cause of ethanol inhibition in ethanologenic mesophilic organisms, e.g., Saccharomyces and Zymomonas. However, this does not appear to be the case for some ethanologenic thermophilic bacteria. Among various functions of the membrane, alcohol inhibits the uptake of various nutrients viz. glucose, ammonium ions, and amino acids; changes the physicochemical properties of membrane; and also causes leakage of various essential cofactors (Ingram, 1986; D’Amore and Stewart, 1987). So far, only limited information is available on the ethanol tolerance of xylose-utilizing yeasts. An ethanol concentration of 20 g/liter begins to affect specific ethanol productivity and xylose consumption in the pentose-utilizing yeast P. tannophilus (Silinger et al., 1982; Watson et al., 1984b). The concentration of ethanol that stops ethanol production is much higher than the concentration that is growth inhibitory (42 g/liter). Ethanol added at a concentration of 80 g/liter results in specific productivity around 0.03 g/g/hr which is about half that observed in the absence of added ethanol. Results indicate that although P. tannophilus can tolerate ethanol up to 100 g/liter, a maximum of 38 g/liter ethanol accumulates in cultures even with an excess of xylose (Silinger et al., 1982). Thus it is apparent that ethanol toxicity is not the factor limiting ethanol accumulation when xylose is the substrate.

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This is in accordance with the observation by Jeffries (1985) that P. tannophilus is able to produce more than 50 g/liter ethanol when glucose rather than xylose is the substrate. Candida shehatae is found to tolerate 80 g/liter ethanol when a mixture of hexose and pentose sugars is used as the substrate (Wayman and Parekh, 1985). Ethanol above 15 g/liter inhibits the growth of Fusarium oxysporum ATCC 10960; however, no growth has been reported at concentrations above 42 g l liter (Rosenberg et a]., 1981). Further, ethanol has no inhibitory effect on xylose fermentation by F. oxysporum VTT D-80134 at 3.5 to 4% (w/v) (Suihko, 1983). Butanol is the most hydrophobic fermentation product of acetonebutanol and ethanol-producing bacteria, and it is generally thought to be the most toxic major product in limiting fermentation. The growth of Clostridia is inhibited by butanol concentrations of less than 1%. The half-maximum growth inhibitory concentration of butanol is seen at a concentration of 0.15-0.18 M in C. acetobutylicum (Leung and Wang, 1981; Costa and Moreira, 1983). Butanol toxicity in mesophilic C. acetobutylicum appears to be related to membrane damage, while ethanol inhibition in thermophilic clostridia, e.g., C. therrnocellurn and C. thermohydrosulfuricum, is due to the direct inhibition of glycolysis. In C. acetobutylicum, growth inhibitory concentrations of butanol dissipate the pH gradient across the plasma membrane and partially inhibit glucose transport and ATPase activity (Bowles and Ellefson, 1985). Efforts have been made to isolate mutants resistant to butanol (Lin and Blaschek, 1983; Hermann et al., 1985). Butanol-resistant mutants lose their sporulation ability which may be due to the pleotropic nature of mutations (Hermann et al., 1985). Butanol-tolerant mutants of C. acetobutylicum do not show increased tolerance to either acetone or ethanol, suggesting that although these products may damage the membrane, their specific action must be somewhat different. In addition to alcohols, weak organic acids are also produced during the fermentation of pentoses. The effect of weak acids, such as acetate and butyrate, on C. acetobutylicum has been studied (Herrero et a]., 1985; Bowles and Ellefson, 1985). These weak acids act as uncouplers of proton transport across the cell membrane. Thus high levels of accumulated weak acids increase ATPase activity, leading to the loss of cellular ATP. Herrero et al. (1985) observed depleted cellular ATP contents in cells of C. thermocellum incubated with 0.8 M acetate. In another study, the total acid produced (0.1 M ) by C. acetobutylicum under normal growth conditions had no effect on cellular physiology but butyric acid added at a concentration of 0.17 M acted as an uncoupler (Bowles and Ellefson, 1985). The half-maximum growth inhibition by

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butyric acid is 0.07-0.16 M in C. acetobutylicum (Leung and Wang, 1981; Costa and Moreira, 1983). It is apparent from these studies that weak acids are toxic products and lead to end product inhibition of the C. acetobutylicum fermentation. In fact, a switch from acid production to solvent production has been regarded as a detoxification mechanism (Rogers, 1986). Cultures of C. acetobutylicum that fail to switch from acidogenesis to solventogenesis die because of acid accumulation (Gottwald and Gottschalk, 1985). Similarly, cultures of acidogenic bacteria, e.g., C. thermoaceticum (which produces only acetic acids), also do not survive once acid is accumulated. Murray et al. (1983) isolated mutants of C. saccharolyticum with less acetic acid production which showed an increased ethanol resistance. Alcohol-tolerant mutants of CIostridium have also been reported by exposure to increasing concentrations of alcohols (Herrero and Gomez, 1980; Lovitt et al., 1984; Bowles and Ellefson, 1985).These mutants may prove to be useful for higher alcohol production in Clostridia. VIII. Strain Improvement

From the aforementioned material, it is apparent that the available yeasts or bacteria are not completely satisfactory for the bioconversion of D-xylose to ethanol. There are various limiting factors like conversion rate, product yield, and low ethanol tolerance. Thus these strains are amenable to strain selection and genetic improvement using recombinant DNA technology. One approach in improving a strain for xylose utilization involves the construction of yeasts that convert D-xylose into ethanol using genetic engineering methods. Brewers yeast, Saccharomyces cerevisiae, which is one of the most ethanol-tolerant yeasts, is unable to produce ethanol from D-xylose; however, a ketoisomer of xylose, xylulose, can be utilized by many Saccharomyces species for ethanol production (Wang et aI., 1980a,b; Gong et al., 1983). Thus these species have the potential to produce ethanol, if they are transformed with a gene coding for the enzyme that can convert xylose to xylulose and is expressed in the yeast. However, most yeasts do not efficiently utilize D-xylose because of cofactor (NADPH/NADH)regulation (Batt et a]., 1986). Hence practical strategies in developing yeast strains to utilize xylose for ethanol production involve attempts to circumvent the xylose reductase-xylitol dehydrogenase pathway. The cloning and introduction of genes from a pentose-metabolizing yeast (e.g., P. tannophilus, C. shehatae, or P. stiptis) in S. cerevisiae would not alleviate the problem of cofactor limitation. Alternatively, transformation with a xylose isomerase gene

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from another source for the direct cofactor free conversion of D-xylose to xylulose is the most useful approach. This involves isolation and characterization of the E. coli gene for D-xylose isomerase. Hence the xylose isomerase gene from E. coli is purified and characterized (Ho and Chang, 1989). Several hybrid plasmids bearing this gene have already been isolated bearing different sizes of the insert. Using this approach the E. coli gene has been expressed both in S. cerevisiae and Schizosaccharomyces pombe (Sarthy et al., 1987; Chan et al., 1989). Schizosaccharomyces pombe has been transformed with hybrid plasmid pDB 248 XI which contains the xylose isomerase gene from E. coli. In transformed yeasts two enzymes are involved in the conversion of D-XylOSe to ~-xylulose-5-phosphate(Chan et a]., 1986). The xylose isomerase converts D-xylose to D-xylulose without NADH or NADPH as a cofactor and xylulokinase converts D-xylulose to ~-xylulose-5phosphate with ATP as a cofactor. Although cloned strains grow on xylose as the sole carbon source, ethanol production is slow (3%, w/ v) and xylitol production is very active. A later study indicated that the low isomerization of xylose in the transformed yeast is the limiting step for D-xylose fermentation. Although yeast proteases decrease xylose isomerase activity in vitro, this finding needs to be confirmed using protease-negative mutants. Xylitol, a by-product of D-xylose fermentation, has no effect on the activity of xylose isomerase activity (Chan et ai., 1989).A low activity of xylose isomerase in transformed yeast might also be due to a low expression of the xylose isomerase gene. Hence construction of an isomerase gene under control of a highly active yeast promoter is likely to improve the expression of the xylose isomerase gene (Chan et al., 1989). Attempts have also been made with xylose isomerase genes from Bacillus subtilis and Actinoplanes missourienis (Amore et al., 1989). In order to increase the production of ethanol from xylose, another approach employed cloning and the expression of the xylose uptake gene from E. coli (Kurose et ai., 1987) and the xylulokinase gene from P. tannophilus (Stevis et al., 1987) and S. cerevisiae (Ho and Chang, 1989).Some of the yeasts like Candida and Rhodotorula which are thought to contain the D-xylose isomerase enzyme are potential organisms in conversion. However, the disadvantage lies in xylitol production. This trait can be altered through conventional mutagenesis. Attempts have also been made to increase ethanol production using genetic methods. For example, xylose consumption and ethanol production by E. coli are increased by a coordinate expression of Zymomonas mobilis pyruvate decarboxylase (pdc) and adh I1 genes (Ingram and Conway, 1988; Neale et al., 1988). Similarly, Klebsiella planticola, which produces acetate, formate, lactate, ethanol, and CO, as end prod-

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ucts of hexose and pentose metabolism, has been used to improve ethanol production. In this organism, pentoses and hexoses are degraded to pyruvate which is dissimilated by the enzyme pyruvate formate lyase to yield acetate, ethanol, and formate (1: 1:2) (Tolan and Finn, 1987). A mutant of Klebsiella lacking pyruvate formate lyase was selected, which accumulated 70% lactate with residual acetate and 2,3butanediol and traces of ethanol. These mutants were further transformed using a plasmid carrying the pyruvate decarboxylase gene from 2. mobilis (which has a highly active pdc system but is incapable of fermenting pentose sugars). These transformed Klebsiella mutants show efficient ethanol production (Feldmann et al., 1989). Various approaches using mutants have been developed to improve the performance of microbes for pentose utilization. Using ultraviolet irradiation mutagenesis, mutants are selected for their ability to utilize xylose for the production of various products (Gong et al., 1981a; McCracken and Gong, 1983). One of the mutants of Candida, Candida SP XF217, which produces 31 g/liter of ethanol from 100 g/liter Dxylose, shows the maximum increase in ethanol production (McCracken and Gong, 1983). In these mutants the specific activity of xylitol dehydrogenase and xylulokinase is increased; however, D-xylose reductase activity remains the same. The increased xylitol dehydrogenase and xylulokinase activity of the mutant enabled them to shift 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. Mutants of P. tannophilus were selected on plates containing nitrate and xylitol as the sole source of nitrogen and carbon (Bolen and Detroy, 1985). These mutants were able to use nitrate for rapid growth. Mutants of P. tannophilus, which can not grow on ethanol as the sole carbon source, also accumulate more ethanol and less xylitol (Lee et al., 1986). These mutants lack enzyme activity for the further degradation of ethanol. For example, mutant eth-2-1 is deficient in the enzyme malate dehydrogenase required for the metabolism of a two carbon compound either by the TCA or by glyoxalate cycles (Lee et aI., 1986).Further hybridization of the eth 2-1 mutant with another mutant, a rapid grower in nitrate-xylitol, improves ethanol production from Dxylose (Clark et a]., 1986). Another approach in obtaining improved strains of P. tannophilus is through the construction of polyploids (Jamesand Zahab, 1982,1983). Pachysolen tannophilus is homothallic in nature and is usually diploid for only a brief period between mating and meiosis. However, auxotrophic haploids of this strain are produced by ultraviolet irradiation of vegetative cells. The diploids are induced to undergo mitosis rather than meiosis. Using this approach a series of

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triploids, tetraploids, and aneuploids were developed (James and Zahab, 1982,1983). A correlation between ploidy and production of ethanol and xylitol is observed in P. tannophilus (Maleszka et al., 1983b). In general, an increase in the chromosome number above the haploid level shows an increase in ethanol production and a decrease in xylitol production, but the level of by-products, e.g., acetic acid and arabitol, follows no pattern (Maleszka et a]., 1983b). An increase in the ploidy of C. shehatae leads to a small increase in ethanol production from Dxylose. The reason for the increase-in-ploidy-enhanced ethanol formation from xylose seems to be due to complex physiological changes that are still not clear. Efforts using a combination of both classical genetics and modern genetic engineering techniques are required to further improve strains for increasing pentose utilization and ethanol production. IX. Future Prospects

The current and projected scarcity of liquid and gaseous fossil feedstocks has promoted renewable (biomass) resources for the production of substitutes. These substitutes include sugar, starch, and lignocellulosic residues. Traditional starch and sucrose-based fermentations can be expanded to increase the supplies of liquid fuels. However, utilization of these agricultural commodities competes with food production because of their use in human and animal food, resulting in a less favorable energy balance. In contrast, lignocellulosic residues are cheap raw materials, containing glucose and pentose sugars. Sugars derived from cellulose and hemicellulose components of biomass are more attractive substrates for the microbial production of solvents. However, the expense involved in converting hemicellulose components has been responsible for the limited success in developing industrial processes. Thus the economic importance of lignocellulose utilization depends primarily on the bioconversion of both hexose and pentose sugars. If these pentoses could be converted to ethanol, some 4 billion additional gallons of ethanol could be obtained in addition to that derived from D-glucose. In developing a simple industrial process for solvent productivity, a biological system that could ferment both types of sugars simultaneously, with high product yields and higher bioconversion rate, is essential. Fermentation of the pentoses is generally slower than that of the hexoses. These rates could be improved through process optimization, but it is likely that the slow rates of pentose fermentation result from the biochemical pathway employed. Yeasts, in general, are the best choice in fermenting pentoses to ethanol. Basic studies related to the regulation of pentose metabolism using

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molecular biological techniques will help in developing new organisms devoid of limitations of ethanol production from pentoses. The cloning of the n-xylose isomerase gene in yeast to by-pass the oxidoreduction steps; the cloning of transhydrogenase (EC 1.6.1.1)to carry out regeneration of NADPH through the interconversion of “NADP+ + NADH NADPH + NAD+;” and the cloning of n-xylulokinase to shift the equilibrium of conversion of D-xylose to ~-xylulose-5-phosphate are among the possible approaches (Gong, 1983). Some pentose-fermenting bacterial strains produce organic acids in addition to the neutral products such as acetone, butanol, isopropanol, butanediol, and ethanol. These acids are usually not desirable products. Knowledge of the regulation of these pathways should suggest biochemical strategies for minimizing acid production in the fermentation. Thermophilic bacteria like C. thermoaceticurn have the potential for homoacetate fermentation of pentose sugars present in the hydrolysate of natural substrates (Brownell and Nakas, 1991).However, the acetic acid fermentation process is hampered by the low end product tolerance. Acetic acid levels above 10 g/liter are reported to stop cell growth and product formation (Wang and Wang, 1984; Sugaya et a]., 1986; Brownell and Nakas, 1991). Thus improvement in pentose-fermenting organisms in terms of yield, productivity, and end product tolerance is likely to be involved in the overall process. These improvements could be achieved by searching for new isolates with desirable properties, understanding the physiology of the strains, and making them accessible to genetic manipulation using advanced recombinant DNA techniques in manipulating carbon flow to desirable products. Therefore, the overall emphasis of future genetic engineering research in this area would be to produce better fermentation biocatalysts with wider substrate ranges and improved kinetics of product synthesis. In addition, improved methods for separating these products from fermentation broths should be sought. The large-scale production of solvents and chemicals from pentose sugars in addition to products derived from hexoses found in agricultural wood and industrial residues should be realized in the near future. Last, but not the least, careful consideration should be given to the idea that the organisms that have been described in the past are not the only candidates of particular interest, new organisms exhibiting desirable new fermentation pattern remain to be discovered.

*

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Medicinal and Therapeutic Value of the Shiitake Mushroom

s.c.JONG AND J. M. BIRMINGHAM Mycology and Botany Department American Type Culture Collection Rockville, Maryland 20852

I. Introduction 11. Medicinal and Therapeutic Properties

A. B. C. D. E.

Hypolipidemic Activity Anti-thrombotic Activity Antibiotic Activity Antiviral Activity Anti-cancer/Anti-tumor Effects F. Lentinan, a Biological Response Modifier 111. Patented Products and Processes A. Anti-hypertensive and Anti-cholesteremic Compositions B. Antibiotics C. Viricides Including Anti-AIDS Agents D. Neoplasm Inhibitors E. Immunoregulatory Substances F. Anti-ulcer Composition G. Anti-clotting Composition H. Anti-asthma Composition I. Bone Formation Accelerator J. Dermatological Compositions K. Postoperative Treatment L. Assay Processes IV. Discussion References

I. Introduction

The shiitake mushroom, Lentinula edodes (Berkeley) Pegler [Lentinus edodes (Berkeley) Singer], is the second most popular edible mushroom in the global market. According to ancient Chinese medical theory, consumption of the shiitake was recommended for long life and good health. Wu Juei, a Chinese physician of the Ming dynasty (1368-1644), claimed that it preserved health, improved stamina and circulation, cured colds and, in modern terms, lowered blood cholesterol (Mori, 1974). Many Japanese people believe that the shiitake is an elixir. In order to explore and possibly exploit the shiitake myth, many scientists have attempted to document its traditional therapeutic value. 153 ADVANCES IN APPLIED MICROBIOLOGY, VOLUME 39 Copyright 0 1993 by Academic Press. Inc. All rights of reproduction in any form reserved.

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The shiitake mushroom contains proteins, fats, carbohydrates, vitamins, and minerals (Breene, 1990). Hypolipidemic and anti-thrombotic substances have been identified; the nucleic acids induce interferon production (Vo, 1987; Eder and Weig, 1988; Chang and Miles, 1989). Reputed antitumor components vary in chemical nature, but the most important may prove to be a polysaccharide that acts as a host defense potentiator (Jong et al., 1991). II. Medicinal and Therapeutic Properties

A. HYPOLIPIDEMIC ACTIVITY

The ability of the shiitake to lower blood cholesterol was first reported by Kaneda and Tokuda (1966), who found that a diet supplemented with the dried ground sporophores of L. edodes lowered average plasma cholesterol when fed to rats. The active principle was identified as an amino acid and named lentinacin by Chibata and co-workers (1969), and lentysine by Kamiya and co-workers (1969).Tokita and co-workers (1972) isolated two closely related compounds from the dried mushroom. The main and active component, 2(R),3(R)-dihydroxy-4-(9adenylj-butyric acid, was called eritadenine, the name currently in use. The minor component, 2(R)-hydroxy-4-(9-adenyl)-butyric acid, had no effect. Eritadenine lowers all lipid components of serum lipoproteins in both animals and humans (Takashima et a]., 1973; Tokuda et a]., 1976; Tokuda and Kaneda, 1979; Suhadolnik, 1979). It has very low toxicity in rats and is effective when administered orally, although only 10°/o is absorbed from the intestinal tract. The effect continues even when it is removed from the diet (Yamamura and Cochran, 1976a). Intravenously administered eritadenine is ineffective; it is rapidly cleared from circulation and excreted through the kidneys. Of the 1 2 4 derivatives of eritadenine that have been synthesized and tested, the most active are carboxylic acid esters with short-chain monohydroxy alcohols. It appears that a carboxyl group and one hydroxyl group, along with an intact adenine ring, are necessary for biological activity. Kabir and co-workers (1987) examined the effect of dried shiitake on the blood pressure and plasma lipids of spontaneously hypertensive rats (SHRs), and found that it decreased both the VLDL- and HDLcholesterol levels. In human testing S. Suzuki and Ohshima (1976) reported that serum cholesterol was decreased in groups of women fed fresh, dried, or UV-irradiated shiitake. A similar experiment, conducted on people 60 years or older, showed that serum cholesterol decreased after 1 week.

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Because shiitake mushrooms are a rich source of dietary fiber, Kurasawa and co-workers (1982) fed rats a standard control diet, and one containing cholesterol, along with whole shiitake or neutral detergent fiber (NDF) extract. The eritadenine-free NDF had a cholesterollowering effect distinct from that of eritadenine, which was attributed to its ability to bind to cholic acid salts. B. ANTI-THROMBOTIC ACTIVITY

Hokama and Hokama (1981) discovered that low molecular weight compounds extracted from some mushrooms, believed to be nucleosides and/or other nucleic acid derivatives, were capable of inhibiting aggregation of blood platelets. The highest yield of the inhibitors was obtained from L. edodes with an IC,, (inhibition concentration) of 80 pglml.

C. ANTIBIOTIC ACTIVITY Bianco (1981) reported that L. edodes was active against Candida albicans, Staphylococcus aureus, and Bacillus subtilis.

D. ANTIVIRAL ACTIVITY Goulet and co-workers (1960) were the first to show that antiviral substances were present in mushrooms. Tsunoda and Ishida (1969) found that an aqueous extract of the fruiting body and spores of the Donko variety of L. edodes was effective against influenza A/SW15 virus infection in mice. The active principle, identified as doublestranded RNA (ds-RNA), originated from attached virus-like particles (Ushiyama et al., 1971; Takehara et al., 1979) and induced interferon production (Kleinschmid, 1972). F. Suzuki and co-workers (1976) extracted ds-RNA from spores; a single dose produced a survival rate of 60% in rabbits infected with influenza virus. Virus-like particles in three basic shapes, spherical (S), filamentous (F),and rod-shaped (R), were detected in normal mycelia and fruiting bodies by Mori and Mori (1976). The S and F particles induced interferon production in sera of rabbits after intravenous (i.v.) injection. Takehara and co-workers, (1979, 1981, 1984) and Toyomasu and coworkers (1986) purified S and F particles and extracted ds-RNA from the S particles. In vitro tests with rabbit kidney cells (RK-13) showed that all induced interferon production. S-derived RNA was the most effective and F particles the least effective. In vivo studies indicated that a single intraperitoneal (i.p.) administration of S particles, prior to

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virus challenge, significantly reduced the mortality of mice infected with western equine encephalitis virus; F particles were considerably less effective. RNA with the same molecular weight as that extracted from the S particles of the spores could be obtained from fruiting bodies. Viral growth was inhibited in vitro, but less effectively than the spore RNA; the same was true for in vitro induction of interferon. When administrated i.p., purified S and F particles and the extracted ds-RNA demonstrated antitumor activity against Ehrlich ascites carcinoma in mice. Antiviral and antitumor activity appears to overlap. A peptidomannan (KS-2) extracted from cultured mycelia grown on stillage from whiskey manufacture (T. Fujii et a]., 1978) also exhibited antiviral activity (F. Suzuki et al., 1979). KS-2, composed of a-linked mannose and a small amount of peptide, has a molecular weight of -6 x lo4-9.5 x lo4.When administrated orally or i.p. to mice infected intranasally with influenza virus, KS-2 afforded therapeutic as well as prophylactic protection through its interferon-inducing activity. Yamamura and Cochran (1976b) determined that compound Ac2P isolated from the aqueous extract of dried shiitake was effective against the viral disease scrapie. Ac2P is a high molecular weight polysaccharide composed mainly of pentose sugars. IR vitro and in vivo tests in mice showed it to be a selective inhibitor of orthomyxoviruses, such as influenza viruses.

E. ANTI-CANCER/ANTI-TUMOR EFFECTS Ikekawa and co-workers (1968, 1969) found that an i.p. injection of an aqueous extract of L. edodes greatly inhibited growth of tumors (81%) arising from sarcoma 180 ascites cells implanted in Swiss albino mice. The active principle, a polysaccharide, was isolated and named lentinan by Chihara and co-workers (1969,1970),who observed complete regression with no toxicity. Additional data (Maeda and Chihara, 1971) showed that lentinan strongly inhibited the growth of transplanted tumors, but it had no effect on spontaneous mammary adenocarcinoma in mice when applied after reimplantation of autologous tumor tissue (Tokuzen and Nakahara, 1971). Because of its clinical and commercial importance, lentinan is considered separately in the following sections. T. Fujii and co-workers (1978) found that when given either orally or i.p. the polysaccharide KS-2 suppressed the growth of Ehrlich ascites tumors as well as sarcoma 180 tumors in mice. Sugano and co-workers (1982) obtained a water-soluble fraction (LEM) and two alcohol-insoluble fractions (LAP and LAP1) from the culture medium of mycelia with activity against Ehrlich ascites carci-

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noma in mice. LAP and LAP1 contained 58 and 65% sugar and 25 and 34% protein, respectively. The major sugar was xylose (>30%) with appreciable amounts of glucose, galactose, and arabinose (13-20%), about 9% mannose, and 1-2% each of fructose and rhamnose. Until the 1980s most of the research on the anti-tumor activity of mushrooms involved administration by injection to test animals. Lentinan and various other polysaccharides were shown to be ineffective when administered orally. More recent work (Mori et a]., l983,1987a,b; Nanba and Kuroda, 1987; Nanba et al., 1987) involved oral administration of powdered, dried mushroom fruiting bodies and powders from which the carbohydrate fraction (P-glucan) and/or lipid fraction was removed. Results of tests with L. edodes indicated that tumor growth could be inhibited 67% by the whole powder, 57% by defatted powder, 39% by polysaccharide-free powder, and 0% by powder free of both lipid and carbohydrate. Addition of extracted lipid elevated inhibition by 25%. The inhibition rate increased when mushrooms were fed over a longer period of time.

F. LENTINAN,A BIOLOGICALRESPONSEMODIFIER 1. Chemical Structure

Lentinan is a P-glucan (Hamuro et al., 1976) with a backbone of P-~-(l+3)-glucanand side chains of both p-~-(143)and P - ~ - ( l + 6 ) linked D-glucose residues, together with a few internal p-~-(1+6)-linkages (T. Sasaki and Takasuka, 1976). There are two P-D-(1+6)-glucopyranoside branches for every five linear /3-~-(1+3)-glucopyranoside linkages (Chihara, 1990). Based on crystalline structure studies (Bluhm and Sarko, 1977a,b), the probable structure of lentinan was determined to be a right-handed triple helix. Saito and co-workers (1977, 1979) concluded that the ordered conformation of both the p-n-(1+3)-linked main chain and side chains is a single-helix conformation which tends to form multiple helices as junction zones for gel structure. High resolution solid-state 13C NMR studies of the secondary structure (Saito et al., 1987; Saito, 1988) indicated that lentinan takes the curdlane-type single-helix conformation and is converted to the triple-helix form by lyophilization after dissolution in a 8 M urea solution and dialysis against distilled water. N. Suzuki and co-workers (1982) studied the hydrodynamic behavior of lentinan and established the relationship between the molecular weight and the diffusion coefficient. The anti-tumor fraction of lentinan is of high molecular weight (T. Sasaki et al., 1976);the average weights

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of two samples were 6.9 x lo4 and 5.2 x lo4 (R. Sakamoto, 1982). Maeda and co-workers (1988) investigated the correlation between higher structure and biological functions. Denaturation and renaturation of lentinan using urea and DMSO were associated with the decrease and recovery of anti-tumor activity against P-815 mastocytoma and vascular dilation and hemorrhage-inducing activity, which are Tcell-mediated responses. The change of the higher structure did not affect the increase of serum acute phase proteins, a non-T-cell-mediated response. 2. General Mode of Action

Lentinan appears to be a potent host defense potentiator which improves homeostasis of the host against cancer or infection (Chihara et al., 1989). It has no direct cytotoxicity to target cells; its action is host mediated. It activates the classical and alternative pathways of the complement system and augments the responsiveness of the host through maturation, differentiation, and proliferation of lymphoid and other physiologically important cells. The fact that lentinan is a T-celloriented adjuvant, in which macrophages play some part, distinguishes it from other well-known immunopotentiators (Hamuro et al., 1976). Although it does not specifically accelerate the production of interleukin-2 (IL-2) from helper T cells, it potentiates the induction of different types of anti-tumor effector cells, such as killer T cells, NK cells, and cytotoxic macrophages (Chihara, 1983; Chihara et aI., 1987). The effector cells may act either selectively or nonselectively on target cells. Various kinds of bioactive serum factors appear immediately after the administration of lentinan, most induced by macrophages. They act on lymphocytes, hepatocytes, vascular endothelial cells, or synovial fibroblasts, causing the many host defense reactions associated with inflammation and immunity. 3. Role of the Thymus and Cell Response

The action of lentinan is part of a thymus-derived immune mechanism (Maeda and Chihara, 1971,1973b; Maeda et a]., 1971,1973). Haba and co-workers (1976) showed that the selective suppression of T cell activity in both sarcoma 180 tumor-bearing mice and cell-free Ehrlich ascitic fluid-treated mice can be prevented by treatment with lentinan. In contrast to strong anti-tumor activity in vivo, lentinan showed no inhibition of sarcoma 180 cell cultures and failed to inhibit tumor growth in thymectomized mice bearing subcutaneously transplanted sarcoma 180 cells, or to stimulate conventional immune responses, such as antibody formation or phagocytosis. Shiio and co-workers ( 1 9 8 7 ~ ) demonstrated that the administration of Thy 1.2 antibody prevents lenti-

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nan suppression of tumor growth in C3H/He mice with sarcoma 180 solid tumor, and diminished its effect on tumor growth suppression in BALB/c nu/nu mice with grafted sarcoma 180 solid tumor. Coadministration of thymus homogenate or extract greatly enhanced its effect. Arai and co-workers (1971) found the anti-tumor activity of lentinan in mice with transplanted sarcoma 180 was reduced if the animals were x-irradiated, given benzylthioguanosine, an immunosuppressive agent, or blocked by injection of anti-lymphocyte serum after tumor transplantation. Dennert and Tucker (1973) showed that lentinan has only minor effects on the plaque-forming cell response to sheep red blood cells, but significantly stimulated antibody dependent, cell-mediated immunity. It did not increase sensitization of T killer cells in an allogeneic system. A simple assay system for ascertaining the cellular orientation of adjuvants and the action of lentinan on T cells using mice was described by Dresser and Phillips (1973, 1974). Maeda and Chihara (1973a) found that lentinan activated the antitumor effect of peritoneal-exudate cells in rats against sarcoma 180 in vivo. Hamuro and co-workers (1979, 1980) showed that it may potentiate cellular-immune responses by reducing synthesis of immunesuppressive prostaglandins from peritoneal-exudate cells. Injection of lentinan (i.p.) rendered murine peritoneal-exudate cells highly cytotoxic which may be related to their ability to activate the alternative path of the complement system. Zakany and co-workers (1980a,b) studied the effect of lentinan on the retardation and regression of transplanted tumors in murine allogeneic- and syngeneic-tumor hosts and concluded that a tumor-induced immunosuppression can be overcome, most likely through enhancement of the migration inhibitory factor (MIF) production. Izawa and co-workers (1982) proposed that lentinan enhances formation of lymphocyte-activating factor (interleukin-1 or IL-I), which results in the accelerated maturation of cells into effector cytotoxic T lymphocytes and natural killer cells. Lentinan augmentation of macrophage reactivity to macrophage-activating factor appears to result in enhanced formation of effector macrophages. Fruehauf and co-workers (1982) found that lentinan augmented IL-1 production by human monocytes and was able to stimulate IL-1 production by the leukemic cell line K-562, while Masuko and co-workers (1982) noted that it restored and potentiated the delayed hypersensitivity reaction in MM46 mammary carcinomabearing C3H/He mice. Sendo and co-workers (1981)observed that administration of lentinan to BALB/c mice enhanced natural killer cell activity in vitro against a radiation-induced lymphoma and a Molony murine leukemia virus-

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induced lymphoma, and natural cytotoxic activity against a methylcholanthrene-induced fibrosarcoma. Fachet and co-workers (1986) examined the influence of lentinan on the oxazolone-specific antibody response and delayed-type hypersensitivity reaction of BALB/c mice. Both the IgM and IgG humoral-immune responses were increased, but the delayed-type hypersensitivity reaction was only moderately enhanced. Lentinan treatment resulted in a considerable decrease in lethality by anaphylactic shock in both tumor-bearing and control animals. The effect of lentinan on granulopoiesis in BABL/c nude mice was investigated by Matsuo and co-workers (1987a,b).They concluded that mature T cells participate in regulation of granulopoiesis in vivo, and lentinan augments granulopoiesis, at least in part, via mature T cell populations. The in vitro studies of Abel and co-workers (1986, 1989) investigated the effect of lentinan on the pinocytotic and phagocytic activity of macrophages. Pinocytosis of HRP (horseradish peroxidase) and dextran by the murine macrophage cell line CaM4, which exhibits a lower basic pinocytic activity than peritoneal cells, was augmented up to 310 and l Z O % , respectively. Microbead phagocytosis by mouse peritoneal macrophages was amplified u p to goo%, suggesting a P-glucan receptormediated activation of pinocytosis and phagocytosis by lentinan. Cawley and co-workers (1987) studied the effect of lentinan on a number of early-phase host-defense mechanisms and on several clinically relevant sublethal infections in rats. It stimulated an increase in peripheral neutrophil numbers, accompanied by a decreased mobilization of these cells, and it demonstrated anti-inflammatory properties. Shiio and co-workers (1988d)found that lentinan administered subcutaneously (s.c.)to ICR/CRJ mice bearing sarcoma 180 solid tumors caused a regression of tumor growth and increased neutrophils, the footpad reaction to tumor antigen, and T cell and neutrophil chemotaxis. Thyroid-follicular epithelial cells (thyrocytes) have been shown to express a number of functions similar to monocytes. In addition to their functions as endocrine cells, they may also participate in the local immune responses under appropriate conditions. Although spontaneous production of thymocyte-stimulating activity (TSA) was not detected by Hirose and co-workers (1987) when grown in culture medium, TSA was demonstrated in culture supernatants after stimulation with the lentinan. Gergely and co-workers (1988) studied the in vitro effects of lentinan on cytotoxic functions of human lymphocytes in patients with solid tumors and chronic lymphocytic leukemia. Lentinan did not influence blastogenesis and lectin-dependent cell-mediated cytotoxicity, but did increase natural cell-mediated cytotoxicity of tumor-bearing subjects.

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4. Effect on Serum Proteins

A distinct correlation was seen by Chihara and Maeda (1982) between the anti-tumor effect of lentinan and the increase in serum protein components in mouse strains that are high responders to lentinan, but not in BALB/c mice which are low responders. The tumor-inhibition ratio for lentinan was 98.0% (Maeda et al., 1974).Maeda and co-workers (1986) separated and purified the inducing factors for acute-phase proteins and vascular dilatation and hemorrhage from lentinan. The former had a molecular weight of about 1.4 x lo5 and the latter consisted of two components with molecular weights of 3.4 x lo5 and 2.5 x lo5. Biological activities were markedly reduced by treatment with proteinase K or trypsin, indicating that they contain a peptide chain as an active part. One of the three kinds of mouse-serum proteins, ceruloplasmin, similar to human ceruloplasmin, was increased by the administration of lentinan (Itoh et al., 1980). A bioactive factor capable of stimulating the production of the acutephase transport proteins, haptoglobin, hemopexin, and ceruloplasmin was found by Suga and co-workers (1986) in mouse serum soon after lentinan treatment. The acute-phase transport protein-inducing factor (APPIF), which appears to be a peptide compound, was produced by macrophages and may regulate the productions of acute-phase transport proteins in hepatocytes. Appearance of APPIF is considered to be one of the earliest manifestations of the mode of action of lentinan, in addition to its augmented production of vascular dilatation and hemorrhage-inducing factor and IL-1. 5 . Effect on Enzyme Activity

Serum X-prolyl dipeptidyl-aminopeptidase activity, which is depressed in cancer patients, is clearly reduced in mice with Ehrlich carcinoma and sarcoma 180 and is slightly reduced in mice with methylcholanthrene-induced sarcomas. The reduced activity is completely reversed during tumor regression of sarcoma 180 by administration of lentinan (Kato et al., 1979). K. Sasaki and co-workers (1982, 1985, 1986) investigated the effect of lentinan on the hepatic drug-metabolizing enzymes in mice and found that in vivo it decreased cytochrome P-450 content and the activities of aminopyrine N-demethylase, aniline hydroxylase, and 7ethoxycoumarine 0-deethylase in the hepatic microsomes. In vitro it had no effect on aminopyrine N-demethylase or aniline hydroxylase activities. The depressing effect of lentinan on the increase in cytochrome P-448 content induced by 3-methylcholanthrene was much greater than its depression on the increase in cytochrome P-450 content induced by phenobarbital.

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The depression of hepatic microsomal enzyme systems varied with the strains of mice used. The potential use of lentinan as a hepatoprotectant was considered by Lu and Fang (1985). They found that serum glutamic-pyruvic transaminase (GPT) was inhibited in vivo after injection in lab animals and in vitro in the blood of humans and rabbits. Feher and co-workers (1989) studied the effects of lentinan on enzyme-induced lipid peroxidation, xanthine-xanthine oxidaseinduced cytochrome c reduction, and superoxide dismutase (SOD) enzyme activity. In low concentration it decreased SOD activity of lymphocytes and erythrocytes from healthy subjects. In higher concentrations, it increased the low superoxide dismutase activity of erythrocytes and lymphocytes of patients with cirrhosis of the liver. No antioxidant effect was observed in NADPH-induced and Fe3+-stimulatedlipid peroxidation and in a xanthine-xanthine oxidase system. Chen and Lu (1989) found that lentinan was a reversible inhibitor of ornithine decarboxylase. 6. Tumor-Host Systems

Using different murine hosts, Suga and co-workers (1984) confirmed the anti-tumor effect of lentinan in syngeneic and autochthonous tumor-host systems and its suppressive effect on 3-methylcholanthrene (MC)-induced carcinogenesis. They found DBA/2, SWM/Ms, and A/J mice suitable hosts for lentinan treatment. Possibly these strains of mice are most sensitive to delayed-type hypersensitivity and/or cytotoxic T cell response in which T cells and lentinan play an important role. Later investigations (Suga et al., 1989) showed the preventive effects of lentinan on metastasis or recurrence of DBAI2.MC.CS-1 and DBA/ 2.MC.CS-T fibrosarcoma, MH-134 hepatoma, and other murine tumors. Yoneda (1984) examined the effect of lentinan on pulmonary metastases in syngeneic mice bearing Lewis lung carcinoma (3LL). Shiio and co-workers (1987a) found that i.v. injection inhibited pulmonary metastases of 3LL, melanoma (BE),and fibrosarcoma (MC-CS-1)in mice transplanted with the tumor cell S.C.or i.v. as evaluated by tumor weight and number of metastasis in the lung. Rose and co-workers (1984) considered the effect against Lewis and Madison 109 (M109) lung carcinomas implanted in the footpads of syngeneic mice. They had greater success with lentinan alone, or lentinan combined with surgery, in treating the M109 lung carcinoma than in treating 3LL. Jeannin and co-workers (1988)tested the effect of lentinan in a model of colon cancer in rats. Lentinan inhibited the growth of carcinomatosis and increased life span. The effectiveness of lentinan was dependent on the number and frequency of the injections and the dose.

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Miyakoshi and co-workers (1984) indicate that lentinan can inhibit cancer in humans. The activation of killer T cells by a mixed lymphocyte culture was accelerated only when responder cells were mixed with both a suboptimum number of stimulator cells and lentinan. The interferon level in the peripheral blood circulation of cancer patients was elevated following lentinan administration, and natural killer activity of peripheral mononuclear cells was enhanced. Miyakoshi and Aoki (1984) found that augmentation of DNA synthesis of peripheral mononuclear cells (PMNC) occurred both in vitro and in vivo by adding or injecting lentinan. The coexistence of T cells, B cells, and adherent cells (mainly monocytes) was essential. 7. Toxicity and Age Dependence of the Host

Lentinan has only a slightly toxic side effect in in vivo application to animal models and human subjects. O’Hara (1980a,b) traced the fate of 3H-labeled lentinan after injection into mice, rats, and dogs. The radioactivity was predominantly incorporated in the liver, the spleen, and the mesenteric lymph nodes. Toxicity studies of lentinan have been performed by Moriyuki and Ichimura (1980),Ishii and co-workers (1980), and Shimazu and co-workers (1980) using rats and mice. The i.v. LD,, of lentinan in male and female rats was 250-500 mg/kg. Oral and S.C. LD,,s were > 2500 mglkg. The higher i.v. doses produced cyanosis, convulsions, and death. Other evidence of toxicity included enlargement of the spleen, nodules on kidneys, erythema of the ears, hemorrhages in the lungs and abdomen, enlargement of the mesenteric lymph nodes, and edema of the diaphragm and intestine. Kosaka and co-workers (1982) determined that lentinan administered to adrenalectomized and oophorectomized patients had no side effects and increased the patients’ survival; there were no toxic effects on adrenalectomized rats. The effect of lentinan on fertility and general reproductive performance of the rat and on pregnancy of the New Zealand white rabbit has been studied by Cozens and co-workers, (1981a,b,c,d). Reactions were generally dose related with no significant effects on the offspring. Toxiciticy studies of lentinan on the rhesus monkey (Sortwell et a]., 1981) and the beagle dog (Chesterman et a]., 1981) showed that a dose level of 0.5 mg/kg/day was without adverse effect. Shiio and co-workers (1987b) studied the effect of age on the antitumor activity of lentinan and found that it enhanced delayed cutaneous hypersensitivity similarly in aged as well as young mice, and is as effective as an anti-cancer immunopotentiator in aged as well as young animals.

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Tsukagoshi (1988) reported that in studies where lentinan was combined with tegafur for the treatment of inoperable and recurrent gastric cancer, the side effects included eruption and redness, mild pressure in the chest, nausea and vomiting, headache, sweating, flushing, dizziness, feeling of throat obstruction, and decrease of white and red blood cell counts and hemoglobin. All side effects were mild and transient. 8. Use of Lentinan in Combination Therapies According to Shiio and Yugari (1981) lentinan is a strong nonspecific immunopotentiator that will act effectively in living things with observed tumor immunity. However, the anti-tumor effects of lentinan vary with the experimental tumor, whether lentinan is used in combination with a therapeutic agent or another procedure, and the timing of administration in the course of treatment. Chang’s study (1981) of the protection against vesicular stomatitis virus (VSV), the Abelson virusinduced tumor, and the allogeneic trophoblastic tumor in mice underscored the importance of selecting the correct strain for study, and the necessity of determining the optimal conditions for enhancement, especially for boosting natural killer activity. a. Combination with Other Therapeutic Agents. Badger (1984) showed that lentinan in combination with chemotherapy increased survival 50% when compared to chemotherapy alone. Combination therapy (S. Abe et a]., 1982a,b, 1983, 1985) with bacterial lipopolysaccharide (LPS) was very effective against Ehrlich carcinoma in ddY mice and syngeneic mammary carcinoma MM46 in C3HIHe mice, but it was only slightly effective on solid-type MH134 hepatoma and colon 38 adenocarcinoma, and ineffective on ascitic L1210 in CDF, mice. On the other hand, combination with cyclophosphamide (CY) strongly inhibited the growth of solid-type MH134 and colon 38 adenocarcinoma even when administered after tumor inoculation. A combination of lentinan with LPS and streptococcus preparation OK-432 showed that all three components were needed for maximum anti-tumor activity in solid-type tumor MH134. Use of OK-432, CY, and/or lentinan plus LPS against Lewis lung carcinoma in C57BLi6 mice provided a model for combination therapy against weakly immunogenic tumors. Moriya and co-workers (1983, 1984) studied the anti-tumor effect of LPS alone and with lentinan using C3H/He mice bearing MI3134 tumor. In combination, the tumor growth was significantly inhibited as compared to that in the delayed-type hypersensitivity in tumor-bearing mice. Moriyama (1982) and Moriyama and co-workers (1981, 1982) found anti-tumor activity on C3H/He mice with MH134 related to dos-

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age and time of administration. Lentinan with mitomycin-C (MMC), 5fluorouracil (5-FU), cytosine arabinoside (ara-C), and particularly LPS inhibited the proliferation better than lentinan or chemotherapy alone. Ishimura and co-workers (1975) tested Corynebacterium parvum and lentinan, serotonin, or thyroid hormone for potentiation of host resistance. Shiio and co-workers (1983) reported on a synergistic effect with marinactan in inhibiting methylcholanthrene A-induced sarcoma (Meth A) growth in mice. Inagawa and co-workers (1986) showed that the combination of OK-432 and lentinan can be practically applied for endogenous tumor necrosis factor (TNF) induction in clinical trials of cancer therapy. Akimoto and co-workers (1984)found that administration of lentinan following a toxic dose of 5-FU induced protection from mortality in C3H/He mice. Interferon inducers apparently play an important role in the prevention of side effects of cell cycle-specific cytotoxic drugs like 5-FU without decreasing their anti-cancer activity. Matsuo and coworkers (1987a,b) suggested that lentinan could contribute to the recovery from some of the hematopoietic depression in clinical chemoimmunotherapy. Injection of lentinan 1 day after 5-FU resulted in prompt restoration of the leukopenia through the recovery of neutrophils, monocytes, and lymphocytes as well as prompt and marked rebound of granulocyte-macrophage progenitor cells. Jiang and co-workers (1985) observed a positive correlation in sarcoma 180 tumor-bearing mice of increases in CAMPlevel in the spleen, blood, and tumor tissue with tumor inhibition by lentinan (alone or in combination with sheep spleen RNA), Tricholoma matsutake polysaccharide, and sheep spleen RNA, in addition to their enhancement of the immunity function. S. Yamasaki and co-workers (1985) determined that nonspecific and specific immune effector induction was synergistically augmented by lentinan and purified recombinant IL-2. In vivo application of lentinan augmented the in vitro IL-2-triggered induction of lymphokineactivated killer cells (LAK) as well as the similarly induced natural killer cell (NK) activation against a wide range of murine solid tumors. Lentinan also increased the generation of cytotoxic T lymphocytes (CTL) from thymocytes against alloantigens in synergy with IL-2. K. Yamasaki and co-workers (1989) induced significant IL-2-activated killer activity in spleen cells of C57BL/6N mice bearing lung metastasis by injecting a combination of IL-2 and lentinan. Minaguchi (1986) found that lentinan decreased the incidence of cancer in rats treated with N-ethyl-N’-nitroso-N-nitroguanidine (ENNG). It also restored the chemotaxis by peritoneal macrophages,

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lymphocyte blastogenesis in the peripheral blood and mesenteric lymph nodes, and the percentage of T cells in the mesenteric lymph nodes. Shiio and Yugari (1987) studied the suppressive effects of combined use with a chemotherapeutic agent on pulmonary metastasis in mice (Lewis lung carcinoma) after surgical removal of the carcinoma. They found that combined therapy with tegafur or cyclophosphamide with lentinan was most effective when the former was given prior to surgery and the latter following surgery. In contrast, combined treatment with bleomycin and mitomycin C was most effective after surgery. Haranaka and co-workers (1987) found that the anti-tumor activity of recombinant human tumor necrosis factor (rhTNF) against Meth A sarcoma in mice and human tumors in vivo was enhanced when combined with lentinan. Takahashi and co-workers (1988) studied the local induction of a TNF-like cytotoxic factor (CF) in murine tissues (MH134 hepatoma) after administration of anti-tumor polysaccharides, including lentinan. Their findings suggest that CF induction is correlated with anti-tumor activity. Komatsumoto and co-workers (1988) found that a combination therapy of lentinan with UFT [2,4(1H,3H)-pyrimidinedione,5-flUOrO-l(tetrahydro-2-furany1)- with 2,4(1H,3H)-pyrimidinedione]was more effective than UFT alone in preventing the metastasis of primary mammary adenocarcinoma to the lungs of rats after surgical excision of the primary site. Shiio (1988a,b,c)studied the administration of a combination of lentinan and cyclophosphamide in the early growth stages of sarcoma 180 solid tumor in mice and found that the combination gave greater inhibition than either agent alone. An additive anti-tumor effect with cisplatin, adriamycin, carboquone, UFT, tegafur, and bleomycin was also observed. When lentinan was administered with cyclophosphamide, 5FU, or tegafur simultaneously, the suppressed growth of Lewis’s lung carcinoma was stronger than when treated with any of the agents alone. Similar results were observed in B,, melanoma and MM102 mammary carcinoma systems. Studies of lentinan and cyclophosphamide, or lentinan and 5-FU, in C3H/He mice bearing an autochthonous tumor showed neither singular nor simultaneous use was effective. However life was prolonged when administration of lentinan was started after cyclophosphamide or 5-FU treatment. Hasegawa and co-workers (1989) studied the effect of lentinan on the inhibition of mitomycin C-induced sister-chromatid exchanges in mouse bone marrow cells and determined that 23% could be inhibited. Lentinan is not only useful for cancer treatment as an immunopotentia-

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tor in combination with anti-cancer drugs, but may also prevent the increase of chromosomal damage induced by anti-cancer drugs. b. Combination with SurgicallEndocrine Therapy. Kosaka and coworkers (1984, 1985, 1986, 1987) studied the cumulative effects of lentinan and endocrine therapy on the growth of 7,lZ-dimethylbenzanthracene (DMBA)-induced mammary tumors of rats. Tumor growth was inhibited when lentinan injections were combined with surgical-endocrine therapy (adrenalectomy and ovariectomy) but not when combined with medicinal-endocrine therapy (tamoxifen treatment). Surgical-endocrine therapy was associated with an infiltration of macrophages and T lymphocytes into the mammary tumors, depletion of estrogen receptors and progesterone receptors in the tumors, and the lowering of blood prolactin levels. This combined therapy is being used clinically with women who have recurrent breast cancer. Patients treated with lentinan showed longer disease-free intervals and a much higher survival rate than controls. Shiio (1988d,e) found that the administration of lentinan after surgical tumor resection suppressed the natural metastasis of B,, melanoma, L1210, and LSTRAm tumors, and showed better results than those observed with the administration of lentinan before tumor resection. Postoperative administration of lentinan was also effective in prolonging the life of mice bearing a second tumor transplant in a nonnatural metastatic tumor system, such as MM102 and colon 26. Lentinan did not prolong life in mice with ascites sarcoma 180 tumor; when ascites tumor cells were transplanted i.p. into mice with solid tumor, it prolonged life and increased the immunity induced by the sarcoma 180 solid tumor, and the residual immunity after resection of the solid tumor. In this system, lentinan suppressed recurrence when administered either before or after surgery. c . Combination withX-Ray Therapy. Shiio and co-workers (1988a,b,c) found that X-ray irradiation and lentinan treatment of mice bearing solid-type sarcoma 180 had an additive effect with the administration of lentinan before or after X rays. C3H/He mice with syngeneic MM102 tumor transplanted S.C. in the footpad were used to study the timing of administration of lentinan. In combination with 2000-3000 rads of irradiation, tumor growth was decreased compared to groups that received radiotherapy or lentinan alone. Lentinan administration before or after irradiation had similar effects. A combination with X-ray therapy prolonged the life of BDF, mice bearing L1210 leukemia and sup-

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pressed the growth of KLN205 squamous cell carcinoma and the metastasis of Lewis lung carcinoma in mice. 9. Other Beneficial Effects a. Anti-parasite Activity. Byram and co-workers (1979) observed that administration of lentinan to thymus-intact mice by i.p. injection resulted in the formation of conspicuously enlarged lung granulomas in response to either Schistosoma mansoni or S. japonicum eggs or to antigen-coated beads. White and co-workers (1988) studied its effect on the resistance of CBA/H mice to Mesocestoides corti. Increasing prophylactic and therapeutic doses resulted in a marked reduction in the numbers of parasites in the peritoneal cavity, particularly in those mice that received lentinan therapeutically. Encapsulated parasites were observed to be dead or dying, and damage appeared to be mediated by increased numbers of macrophages and giant cells.

b. Antibacterial Activity. Sakamoto and co-workers (1983) investigated the effect of lentinan treatment on host resistance of malnourished rats against Listeria monocytogenes infection. Administration induced complement C3 elevation in vivo, and the C3 enhancement increased the resistance to infection. Iguchi and co-workers (1985) studied bacterial infections in mice with neutropenia caused by cyclophosphamide or fluorouracil. Administration strengthened resistance against infections with Escherichia coli, Pseudomonas aeruginosa, and Klebsiella pneumoniae by 10 to 100 times. Lentinan is effective in augmenting resistance against bacterial infections in a host immunocompromised by anti-cancer agents. Kawanobe and co-workers (1988) examined the ability of lentinan to increase host resistance to certain bacterial infections in malnourished rats through complement C3 activity. The administration of Znchlorophyllin and 2 mg/kg of lentinan enhanced C3 levels and C3b and C3bi formation. Apparently, phagocytic activity or the clearance capacity of macrophages is augmented through the interaction of macrophages and increased C3bi formation. Yano and co-workers (1989) found that lentinan could enhance the resistance of carp Cyprinus carpi0 to experimental bacterial infection of Edwardsiella tarda by activating the nonspecific immune system. c. Anti-fungal Activity.

H. Chen and co-workers (1987) investigated

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the therapeutic effects of lentinan against Candida infection. The peritoneal macrophages (PECs) from lentinan-treated mice showed a strong inhibitory activity against germ tube formation of C. albicans. Ohno and co-workers (1986) showed that it was effective in the augmentation of candidastatic activity of the murine macrophage cell line J177.1 in vitro. d. Antiviral Activity. Lentinan used in combination with 3'-azido3'-deoxythymidine (AZT) suppressed the surface expression of human immunodeficiency virus (HIV) antigen more strongly than AZT alone. Tochikura and co-workers (1987) showed that it can enhance the effect of AZT on replication of HIV in various human hematopoietic cell lines in vitro. Additional investigations (Tochikura et al., 1988, 1989) measured the effects of lentinan sulfate and E-P-LEM against human retroviruses. The substances almost completely blocked cell-free infection of HIV-1 and HIV-2 and inhibited cell-to-cell infection by HIV-1, HIV-2, and HTLV-I. Moreover, reverse transcriptase activity of avian myeloblastosis virus was inhibited. Yoshida and co-workers (1988) found that sulfated lentinan inhibited HIV-induced cytopathic effect and viral antigen synthesis in HIVinfected MT-4 cells. It showed >98% reduction of reverse transcriptase activity of avian myeloblastosis virus. Hatanaka and Uryu (1989) showed that sulfonated lentinan had anticoagulant and antiviral (HIV) activities, but complete sulfonation of the C-6 carbons and a high molecular weight were necessary for the biological activities. Lentinan sulfate with an S content of >13.9% effectively prevented HIV-induced cytopathic effects in an HTLV-Icarrying cell line (MT-4) in vitro at concentrations of >3.3 pgiml (Hatanaka et al., 1989). Tochikura and co-workers (1988) fractionated an extract of culture medium (LEM) and both the resulting product (E-P-LEM) and LEM were studied for their effect on the activity of HIV in vitro. The experiments were performed using either a cell-free infection system with MT-4 cells, or a cell-to-cell infection system with MOLT-4 cells, which induces multinucleated giant cells. E-P-LEM almost completely blocked both the cytopathic effect of giant cell formation and specific antigen expression due to HIV, whereas LEM before ethanol precipitation blocked the expression of HIV antigen in MT-4 cells only at a high concentration. Pretreatment of the virus with E-P-LEM before infection blocked HIV infection in the target cells. Thus, the inhibitory effect on HIV could be due to a blocking of the initial stages of HIV infection.

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H. Suzuki and co-workers (1989) showed that fractions from an extract of the mycelium culture medium (LEM) activated mouse macrophages, caused proliferation of bone marrow cells, and inhibited the replication of HIV virus in vitro. The EPS4 fraction was composed of water-soluble lignins containing minor amounts of protein (3.2%) and sugars (12.2%). The active principle in the fraction EPS3 is a highly condensed and polycarboxylated lignin which is denatured and solubilized by L. edodes from bagasse (H. Suzuki et ~ l . 1990). , e. Effect on Low Natural Killer Syndrome. Low natural killer syndrome (LNKS)is a newly proposed category of immune disorders being characteristically diagnosed by lowered NK cell activity against K562 target cells as a definite laboratory abnormality, with general clinical symptoms of remittent fever and uncomfortable fatigue, persisting without explanation for more than 6 months. It is independent of AIDS or the AIDS-related complex. LNKS patients responded well to the administration of lentinan, despite no responses to conventional fever treatments (Aoki et al., 1987)

f. Anti-diabetic Activity. Satoh et al. (1988) studied the effects of various biological response modifiers, including lentinan, on insulindependent (Type I) diabetes mellitus in nonobese diabetic (NOD) mice. Lentinan inhibited development. g. Radioprotection. Matsubara and co-workers (1988) studied the radioprotection action of metallothionein induction with lentinan in mice exposed to X rays. Increases in survival induced by pretreatment with heavy metals and immunostimulants were similar, but Zn used in combination with an immunostimulant appeared to produce optimal protection. Further studies (Matsubara et d., 1989) substantiated that lentinan enhanced survival of male ICR :Jcl mice subject to whole body X-irradiation. Chirigos and Patchen (1988) surveyed the ability of biological response modifiers to restore and/or counteract the suppressive effects of the radiomimetic drug cyclophosphamide and total-body irradiation. Patchen and co-workers (1988) studied the potential use of 1 7 immunoregulators for radioprotective efficacy in female C3H/HeN mice. Significant radioprotection, based on enhanced survival following wholebody irradiation, was observed for lentinan, which also stimulated he-

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TABLE I MEDICINAL BENEFITSOF THE SHIITAKE MUSHROOM Benefit

Compound

Antibiotic Anti-parasitic

Polysaccharide lentinan

Antibacterial

Polysaccharide lentinan

Anti-fungal

Polysaccharide lentinan

Anti-tumoI

Polysaccharide lentinan Nucleosides and/or Nucleic acid derivatives Polysaccharide lentinan

Antiviral

peptidomannan (KS-2) Double-stranded RNA Polysaccharide LAP1 Double-stranded RNA

Anti-diabetic Anti-thrombotic

Peptidomannan (KS-2) Polysaccharide AcZP Polysaccharide lentinan Lentinan sulfate LEM

Hy pocholesteremic (Hypolipidemic)

E-P-LEM EPS3 and EPS4 Eritadenine

Immunomodulatory

Polysaccharide lentinan

Radioprotection

Polysaccharide lentinan

Reference Byram et a]., 1979 White et al., 1988 M. Sakamoto et al., 1983 Iguchi et al., 1985 Kawanobe et al., 1988 Ohno et al., 1986 Chen et al., 1987 Satoh et al., 1988 Hokama and Hokama, 1981 Chihara et al., 1970 Dennert & Tucker, 1973 Hamuro et al., 1976 Fujii et al., 1978 Takehara et al., 1981 Sugano eta]., 1982 F. Suzuki et al., 1976 Takehara et al., 1979 F. Suzuki et al., 1979 Yamamura and Cochran, 1976b Tochikura et al., 1987 Tochikura et al., 1988, 1989 Tochikura et al., 1988 H.Suzuki et al., 1989 Tochikura et al., 1988, 1989 H. Suzuki et al., 1990 Kaneda and Tokuda, 1966 Tokita et al., 1972 Yamamura and Cochran, 1976a Suzuki and Ohshima, 1976 Tokuda and Kaneda, 1979 Kurasawa et al., 1982 Tsunoda and Ishida, 1969 Suzuki et al., 1976 Hamuro et a)., 1976 Chirigos and Patchen, 1988 Patchen et al., 1988

matopoiesis. Results indicated the potential use of immunomodulators for protection against acute radiation injury and hematopoietic enhancement alone may not be sufficient to enhance survival following irradiation.

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Ill. Patented Products and Processes

A. ANTI-HYPERTENSIVE AND ANTI-CHOLESTEREMIC COMPOSITIONS A germanium-rich beverage from L. edodes is both anti-hypertensive and anti-cholesteremic (Iizuka, 1982). A health food preparation containing shiitake has an anti-cholesteremic effect (Y. Abe and Kaneda, 1986). Manufacture of cholesterol-lowering and immunoactivating eritadenine was accomplished by fusing L. edodes and Collybia velutipes (Nippon-Food, 1987). B. ANTIBIOTICS

Various antibiotic substances have been obtained from Lentinula (Shiio et al., 1973). Lentinan can be used for Pseudomonas infection control in animals. Administration increased their survival 70% (Ajinomot0 Co., Inc., 1985). Lentiallexine (octa-7-en-3,5-diyn-l-o1) is obtained when L. edodes is cocultured with Trichoderma sp. (Mitsubishi Chem. Ind. Co., Ltd., 1988). C. VIRICIDESINCLUDING ANTI-AIDSAGENTS An anti-tumor viricide and fungicide for the treatment of verruca and collagenosis is produced from the mycelium and used culture medium of Basidiomycetes, preferably L. edodes. Xylose is the main component (Noda-Inst., 1983). Soluble protein extracted from L. edodes fruiting body (FBP)is useful as a viricide (Nikken Chem. Co., Ltd., 1986). A nontoxic extract from the nutrient medium and tissue medium of Basidiomycetes, such as L. edodes, is effective against viral hepatitis. The active components are a mixture of polysaccharides and cytokinin substances, mainly zeatin and zeatin riboside (Iizuka, 1986). Several compositions are used as anti-AIDS drugs. LEM-HT, extracted from cultured Lentinula mycelium, can control the reduction of T lymphocytes due to viral infection by stimulating macrophages and IL-1 activity (Iizuka and Maeda, 1988).Another is obtained from Basidiomycetes, especially L. edodes (Iizuka et al., 1990, 1992). A method for the inhibition of virus infection involves sterilization of devices with a dilute mycelial extract from L. edodes, which acts as a therapeutic agent against HIV virus and hepatitis B virus (Noda-Food, 1989). Lentinan sulfate was shown to be active against HIV when tested in MT-4 cells infected with HTLV-I11 (Yamamoto et al., 1989).A viricide composition to prevent the multiplication of HIV virus, herpes virus, and hepatitis

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B virus was obtained by fermentation of vegetable fiber by Basidiomycetes, especially L. edodes. The product of the extraction is in the form of the brown powder LEM (Noda-Food, 1990a). The polysaccharide fraction of an aqueous extract of a mycelial culture of L. edodes with a molecular weight of 1 x lo5-1 x lo6 is used in preparation of a drug for prevention or therapy of herpes simplex virus, cytomegalo virus, or Epstein-Barr virus infections (Nippon Chem.: Noda-Food, 1990; Koga et al., 1991). A similar compound suppresses replication of HIV virus, inhibits the adsorption of HIV virus onto host cells, and inhibits the activity of reverse-transcriptase. It is nontoxic with no side effects (Noda-Food, 1990b).

D. NEOPLASM INHIBITORS The first neoplasm inhibitor was identified as a glucan (Chihara et al., 1972). Emitanin (T. Suzuki and Ikegawa, 1977) and emitanin-1 (Yamamoto and Ikegawa, 1980) also have anti-cancer properties. An anti-tumor polysaccharide can be obtained cultivating L. edodes on bagasse (Japan Synthetic Rubber Co., Ltd., 1978). Anti-cancer polysaccharides are stabilized and anti-cancer activity is increased synergistically when the polysaccharides are dissolved in water in the presence of water-soluble high molecular weight compounds and monosaccharides. The addition of a water-soluble dextran increases the solubility of lentinan (M. Fujii et al., 1980). Liposaccharides isolated from bacteria (Proteus vulgaris) have been combined with lentinan for use as synergistic neoplasm inhibitors (Ajinomoto Co., Inc., 1981). E. IMMUNOREGULATORY SUBSTANCES

KS obtained from the cultured mycelium of L. edodes and refined KS-2-A enhance the host defense function (Ishida et al., 1979a; Kirin Brewery Go., Ltd., 1980). KS-2-B (Ishida et a]., 1979b, 1981) and KS-2D (Kirin Brewery Co., Ltd., 1981) are also effective interferon-inducing substances. A combination of human interleukin 2 (IL-2) purified from various cell lines and lentinan is useful as an anti-tumor agent (Yoshimoto et al., 1983,1988).Lentinan has been used as an immunotherapeutic agent for the control of neoplasm and infection (Ajinomoto Co., Inc., 1984). A tumor therapy has been described using murine monoclonal antibodies to carcinoma, melanoma, and pancreatic carcinoma as anti-tumor agents and lentinan as a macrophage activator to enhance the anti-tumor activity of the antibodies. Antibody-dependent macrophage-mediated

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cytotoxicity has been demonstrated using the antibodies and the lentinan-stimulated macrophages against various human tumors in vitro (Herlyn, 1986, 1992). An immunostimulant comprising an N-containing low molecular substance and sugar protein containing xylose is obtained from the mycelium culture medium of a Basidiomycete, such as L. edodes (NodaFood, 1984a; Sugano et al., 1984). This composition is useful for the treatment of cancer by intraperitoneal or oral administration. An extract composed of two peptoglycans from the mycelium and culture liquid enhances humoral and cell-mediated immunity to a wide range of diseases, e.g., hepatitis, influenza, and herpes virus infections, cancer, immunodeficiency disorders, and mycotic infections (Sugano et al., 1985). A preparation for treatment of kidney inflammation without reducing immunity, which consists of a saccharide and a protein, is obtained from the mycelium culture broth of Basidiomycetes, such as Lentinula, by fermentation in a culture medium rich in xylose (NodaFood, 1986). A high molecular weight immunostimulant containing mostly neutral sugar and protein extracted from the spawn of the fruit body of L. edodes grown in a solid medium containing cellulose is useful for the treatment of chronic-type hepatitis B and other immunodeficiencies (Noda-Food, 1987). F. ANTI-ULCER COMPOSITION

Ulcer-suppressing agents containing an extract of the mycelium of L. edodes may be administered orally, as suppositories, or by injection (Mitsubishi, 1983). G. ANTI-CLOTTING COMPOSITION

Pharmaceuticals contain lentinan for the prevention and therapy of disseminated intravascular clotting. The efficacy was demonstrated in rats with experimentally induced disseminated intravascular clotting (Res. Dev. Corp. of Japan, 1987).

H. ANTI-ASTHMA COMPOSITION A drug for asthma and treatment for cancer and skin disease contains the extract of a cultivated mycelium of a Basidiomycete, such as Lentinula. The mycelium cultured on a medium containing horse excrement contains saccharides, primarily xylose, and protein (Noda-Food, 1984b).

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I. BONE FORMATION ACCELERATOR An accelerator of bone formation contains lentinan. The pharmacological effects were shown in rats with bone damage (Res. Dev. Corp. of Japan, 1988).

J. DERMATOLOGICAL COMPOSITIONS A bathing composition employs the mycelium of a Basidiomycete, such as L. edodes. The method of preparation allows all of the pharmaceutically active ingredients contained in the mycelium, to be utilized (Nikkei Co., Ltd., 1986).A composition for external application to regenerate and revitalize damaged cells has been found useful in the cosmetic and pharmaceutical fields (Yamada and Yamada, 1992).

K. POSTOPERATIVE TREATMENT A prophylactic and therapeutic treatment of complications after lensectomy utilizes lentinan (Taiho Pharmaceutical Co. Ltd., 1990). L. ASSAYPROCESSES

P-1,3-Glucan has been used as part of the chain reaction-triggering substance in liposomes for a simple and highly sensitive lysis immunoassay. The change in viscosity was measured for detection of p-1,3glucan release from the liposomes to determine the antibody detection (Seiko Instruments & Electronics, Ltd., 1989). An anti-lentinan antiserum has also been employed in immunoassays (Ajinomoto Co., Inc., 1992). The concentration of endotoxin in an unknown sample can be determined by using the reaction of a horseshoe crab hemocyte lysate with endotoxin in a solution with a water-soluble polysaccharide containing p-1,3-glucosidic linkage, or a derivative containing the linkage, such as that found in lentinan (Matuura and Tsuchiya, 1993). IV. Discussion

In carefully controlled laboratory studies the shiitake mushroom has been shown to contain components effective in the treatment of cancer, heart disease, and diseases caused by viral infections. Animal models have demonstrated that these biologically active principles may exert totally different effects depending on the dose, route of administration, and the condition of the host. It is also apparent that similar types of

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S. C. JONG AND J. M. BIRMINGHAM TABLE I1 PATENTED PRODUCTS AND PROCESSES Product or process

Anti-asthma composition Antibiotics

Anti-clotting composition Anti-hypertensive/ anti-cholesteremic compositions Anti-tumor compositions (neoplasm inhibitors)

Anti-ulcer composition Antiviral compositions [including anti-AIDS agents)

Bone formation accelerator Dermatological compositions Immunoregulatory substances

Postoperative treatment Assay processes

Patent Noda-Food, 1984b Shiio et aI., 1973 Ajinomoto Co., Inc., 1985 Mitsubishi Chem. Ind. Co., Ltd., 1988 Res. Dev. Corp. of Japan, 1987 Iizuka, 1982 Abe and Kaneda, 1986 Nippon-Food, 1987 Chihara et al., 1972 T. Suzuki and Ikegawa, 1977 Japan Synthetic Rubber Co., Ltd., 1978 Yamamoto and Ikegawa, 1980 Ajinomoto Co., Inc., 1981 Fujii et ol., 1980 Mitsubishi, 1983 Noda-Inst., 1983 Nikken Chem., Co., Ltd., 1986 Iizuka, 1986 Iizuka and Maeda, 1988 Noda-Food, 1989, 1990a, 1990b Yamamoto et al., 1989 Nippon Chem.: Noda-Food, 1990 Iizuka et al., 1990, 1992 Koga et al., 1991 Res. Dev. Corp. of Japan, 1988 Nikkei Co., Ltd., 1986 Yarnada and Yamada, 1992 Ishida et al., 1979a, 1979b, 1981 Kirin Brewery Co., Ltd., 1980, 1981 Yoshimoto et ol., 1983, 1988 Ajinomoto Co., Inc., 1984 Noda-Food, 1984a, 1986, 1987 Sugano et al., 1984, 1985 Herlyn, 1986, 1992 Taiho Pharmaceutical Ltd., 1990 Seiko Instruments ?i Electronics, Ltd., 1989 Ajinomoto Co., Inc., 1992 Matuura and Tsuchiya, 1993

effects can be elicited by structurally diverse molecules. Two of the most promising and effective principles isolated are lentinan and LEM. Lentinan first extracted from the fruiting body of L. edodes is a pure P-1,3-glucan containing only glucose. It appears to act as a host-defense potentiator which can improve the physiological constitution of the

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host against cancer, as well as various kinds of infections, and restore or augment the ability of responsiveness of the host to bioactive substances, such as lymphokines or cytokines. Lentinan has been shown to exert prominent anti-tumor effects in murine allogeneic, syngeneic, and autochthonous hosts, to prevent chemical and viral carcinogenesis, to increase host-resistance to bacterial, viral, and parasitic infections, and is effective against HIV or AIDS infections. Remarkable life span prolongation has been achieved in patients with advanced and recurrent stomach, colorectal, and breast cancer. Lentinan is commercially available for clinical use. In 1987 it was the eighth top-selling anti-cancer drug in Japan with a 2.2% share of the market valued at $3 million (Fukushima, 1989). The whole extract of L. edodes mycelial culture (LEM)and its purified fractions have antiviral activities and immunomodulating functions. The active principle in the EP3 fraction and its lower molecular weight fraction (EPS4) has been identified as a highly condensed and carboxylated lignin. LEM inhibits the infectivity of HIV and cytopathic effects on virus-infected cells in vitro, enhances IL-1 production, activates murine macrophage functions, promotes proliferation of murine bone marrow cells, suppresses proliferation of rat ascite hepatoma AH414, and promotes seroconversion from HBe antigen to anti-HBe antibody in chronic hepatitis B patients. LEM has been shown to be effective in AIDS therapy and hepatitis B therapy by oral administration. The use of cultivated edible mushrooms, such as L. edodes, as a source of biologically active principles offers obvious advantages. Edibility increases the likelihood of a safe, tolerated principle, while cultivability assures an adequate supply. However, cultivation of the mushroom need not be a limitation when active principles can be derived from its mycelial cultures, as is the case with LEM. The biologically active components isolated and identified in edible fungi show great promise and should be further exploited for their therapeutic effects, either as dietary components or as purified drugs. The cure for the diseases that plague mankind, particularly cancer and AIDS, may lie within the biochemistry of edible mushrooms.

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Yeast Lipid Biotechnology Z. JACOB' Fermentation Technology and Bioengineering Discipline Central Food Technological Research Institute Mysore 570 013, India

I. Introduction 11. Yeasts as Potential Sources of Lipids 111. Importance of Yeast Lipids in Beverages and Foods A. Beer and Wine B. Dairy and Baked Products C. Oriental Foods and Pickles IV. Medical Importance of Yeast Lipids V. Modification of Lipids A. Fermentative Synthesis and Modification of Lipids B. Genetic Engineering Aspects of Yeast Lipid Modification C. Chemical and Biochemical Interesterifications VI. Commercial Significance of Yeast Lipid Biotechnology VII. Conclusion References

I. Introduction In vitro tissue culture (somatic, meristem, and shoot tip) techniques are now available for the clonal propagation of oil seed plants (Pandey, 1989a). In addition to the above, there are many examples in plant science of the genetic modification of oil seeds (rape, flax, sunflower, safflower, soybean) and transgenic seeds, which are now available to produce lipids of desired composition (Khatoon, 1991; Voelker et a ] . , 1992). Certain yeasts are also considered as potential lipid producers. Recent developments in yeast biotechnology are the results of the search for novel compounds, life-saving biopolymers, and modified food components and medicine (carbohydrates, protein, and lipids),which otherwise may not easily be synthesized by chemical pathways (Hodgson, 1991; Dixon, 1991).The integrated approaches of biotechnology, recombinant DNA technology, and fermentation technology have made the area of research on lipid biotechnology more challenging and attractive. During fermentation of various bioorganic substrates, the yeasts synthesize and store lipids intracellularly. Some species of yeasts such as Rhodotorula, Lipornyces, and Candida produce lipids closer to vegeta-

' Present address: Department of Medicine, Division of Oncology, V.C. 12-238,College of Physicians and Surgeons of Columbia University, New York, New York 10032. 185 ADVANCES IN APPLIED MICROBIOLOGY, VOLUME 39 Copyright 0 1993 by Academic Press, Inc. All rights of reproduction in any form reserved.

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ble oils and fats. Tapping of such alternate sources has assumed greater importance in the last few decades because of the widening gap between the total demand and production of vegetable edible oils and fats in many developing countries. Although many efforts have been made, limited success has been achieved in this regard mainly because of the lower productivity of the oleaginous biomass and because of many technological constraints in the downstream processing of the product. When comparing the time-consuming process of plant oil seed production to that of the yeast biomass, the latter has an edge over the former in the ease of cultivation in a shorter amount of time. However, yeasts and their biotechnological application in obtaining desired lipids have not received adequate attention as compared to the oil seeds. While granting the inherent loopholes of the use of yeasts for mass scale production of useful lipids, there has been a reorientation in thinking toward the production of value-added lipids or novel lipids, using the integrated approach of biotechnology. Recent attempts of insertion and expression of desired intraspecific traits have not fully fulfilled the goal of higher productivity of edible oils and modified lipids. This chapter considers the state of the art role of yeasts in lipid biotechnology and discusses their prospects in food and medicine. II. Yeasts as Potential Sources of Lipids

These eukaryotes produce beneficial products such as alcohol, beverages, single cell protein (SCP), and many biochemicals used in the food and pharmaceutical industries. As noted earlier, some yeasts also produce beneficial lipids and lipid-containing emulsifying compounds (Jacob 1989; 1992). The advantages of using yeasts as lipid producers are that (1) they produce lipids similar to vegetable oils and fats, (2) they can be grown reasonably well on cheap agroindustrial and food industrial wastes, (3) their lipids can be produced at a faster rate in bulk in large capacity reactors than the usual time-consuming agricultural practices, and (4) most of the potential lipid producers and their products seem to be relatively nontoxic to humans. Yeasts use carbohydrates as precursors to synthesize lipids by enzymatic pathways (Ratledge, 1982; Ratledge and Evans, 1988; Guerzoni et a]., 1985; Rattray, 1988; Holdworth et al., 1988). The lipid composition, quality, and amount vary from species to species according to the growth stage, optimal availability of essential nutrients, and the conditions of the reactor. However, normally, the ability to catabolize a specific substrate or precursor to a desired lipid component is limited. Increased productivity of a specific lipid component requires the control of specific enzymatic

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pathways. Specific lipid modifications may be possible by (1)rerouted metabolic pathways (stimulatory or inhibitory) under optimized conditions of growth in the presence of higher amounts of precursors or suitable substrates, (2) intraspecific cloning and expression of desired genes in the organism, (3) biochemical conversion of the postharvest lipids by enzymatic conversions or direct fermentation with a suitable microorganism, and (4) chemical conversion. Once the modifications are made by biochemical or chemical means the total lipid can be extracted by a suitable solvent system that is similar to vegetable oil extraction and from it desired components may be isolated by fractional extraction using supercritical CO, or by column separation. Ill. Importance of Yeast Lipids in Beverages and Foods

A. BEERAND WINE The primary contribution of brewer’s yeast to beverages is the production of ethanol and organoleptic and quality determining compounds such as aldehydes, ketones, lower fatty acids, and esters (Johnson et al., 1958; Wiseblatt, 1960). The quality of beer and wine depends on the type of fermentation, substrate supplied, and the yeast growth conditions (Kirsop, 1977, 1988; Berry and Watson, 1987). During vinification of grape juice, different species of Saccharomyces [chevaliere, carlsbergiensis, fructum) produce varying quantities of acetic, n-butyric, ncaprioc, n-caprylic, n-capric, 9-decenoic, succinic, formic, propionic, isobutyric, 2-methyl butyric, isovaleric, lactic, 2-hydroxycaproic, nperlargonic, and malic acids, higher alcohols (fuse1 oil), and esters (Margalithi and Schwartz, 1970; Berry and Watson, 1987). Many metabolic changes related to lipids also occur. During champagnization the wine yeasts proliferate and liberate diverse lipids, thus enriching the wine lipids (Kishkovskii et al., 1986). The improvement of sparkling and foaming properties of sparkling wine depends on the type of yeast and the chemical properties of the surfactants used for wine fortification (Razmadze, 1985). The performance of pitching yeasts during brewing also depends on the intracellular lipid content (Sayle, 1986). Optimum aeration results in the synthesis of an amount of lipid necessary for desirable fermentation (Ohno and Takahashi, 1986).Wort aeration, temperature, and oxygen supply strongly affect the lipid composition (Ohno and Takahashi, 1983). Oxygen is essential for unsaturation and cyclization of squalene to lanosterol. In the absence of oxygen, no phospholipids and triacylglycerols are synthesized. In oxygen-limited fermentations the toxic decanoic fatty acids are adsorbed by the yeasts (Munoz

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and Ingledew, 1989). Pitching yeasts are rich in unsaturated fatty acids like palmitoleic and oleic acids. The synthesis of oleic acid requires more oxygen and time (Ahvenainen, 1982). During beer fermentation, the intracellular unsaturated lipids help maintain the cell viability as well as the ethanol tolerance of the yeast (Rose, 1978). Taylor et al. (1979) reported that beer fermentation requires externally supplied lipids such as sitosterol, unsaturated fatty acids of spent grain lipids, and medium chain fatty acids as well as a higher oxygen supply. These factors cause favorable changes in the patterns of fatty acid composition, sterols, decreased content of esters, medium chain fatty acids, and increased fuse1 alcohol content. During wort production, the brewer’s yeast liberates more lipids with unsaturated fatty acids and sterols (Pfisterer et a]., 1977). The influence of yeast lipids on beer flavor is substantial and is growth related. Freshly synthesized fatty acids may be excreted and impart caprylic flavor to the beer and are primarily controlled by the oxygen supply. Aries et al. (1977) reported that in the presence of oxygen the fermented wort (produced by pitching yeast) contains a complex mixture of unsaturated fatty acids of varying chain lengths. During anaerobic fermentation, increased medium chain fatty acids are produced and a major fraction of them are excreted. Acetoin is also produced when ergosterol is fed in the presence of oxygen during brewing (Haukeli and Lie, 1976). In wine fermentation the lipid content of the yeast increases, regardless of the method of fermentation. However, maximum accumulation was noticed during the aerobic process and was the lowest in pressurized CO, conditions. Wine lipids contain C14 to C24 fatty acids. The major fatty acids are palmitic, stearic, palmitoleic, oleic, linoleic, and linolenic acids. The degree of anaerobiosis, temperature, CO, concentration, and ethyl alcohol formation affects the lipid content and its degree of unsaturation (Portnova, 1981). Abdurazakova et al. (1982) reported that the presence of unsaturated long chain fatty acids stimulates the production of lipase in Saccharomyces vini (wine yeast) and that maximum activity is observed at the exponential phase of growth. Chemiluminescence as well as the lipid content of the wine are also found to be affected by the oxygen supply during wine fermentation using S. vini (Magomedov and Portnova, 1977). Sparkling wine yeasts transfer lipids and proteins to the external medium and these depend on the nitrogen supply which affects the foaming characteristics (Razmadze et al., 1980). Commercially, the flavor of apple wine is enhanced by adding lipases obtained from Candida sp. or Rhizopus delamar (Tanabe Seiyaku Co., 1970). During wort fermentation, using S. cerevisiae, the extent of yeast growth is related to the cellular sterol levels, and the

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survival of yeasts is correlated with unsaturated fatty acid residues which in turn affect the cell physiology, particularly the membrane structure and function (Day et a]., 1976). B. DAIRYAND BAKEDPRODUCTS

As mentioned earlier, microorganisms are also responsible for desirable flavors in fermented and baked foods. Microorganisms utilize the supplied substrates and precursors and transform them to organic acids, aldehydes, ketones, alcohols, esters, and many other compounds, which, in totality, makes the dairy and baked products more appealing and acceptable. Bacteria and yeasts that possess lipolytic (fat splitting) activity are found in many proteinaceous and fatty foods (Davis, 1970; Scheibner, 1970). The action of bacteria is quite evident in imparting good sensory feeling to most of the fermented products, while the role of yeasts seems to be minimal. Little work has been reported so far on the contribution of different yeast species to the final flavor-determining components of dairy and baked products. Microbial lipases play an important role in the development of cheese flavors (Posorske, 1984). In situ synthesis of bacterial lipases has been reported in Dutch, Swiss, Gouda, and cheddar cheeses (Seitz, 1974). In Roquefort cheese, molds like P. roqueforti and Mucor mehii give a characteristic flavor and odor. The spores of P. roqueforti split the triacylglycerol to fatty acids which are further transformed to methyl ketones by an oxidase enzyme. It is the lipase activity of P. roqueforti that yields caprylic, capric, and caproic acids which may then be involved in the formation of methyl ketones (cheddar cheese flavor). However, in the ripening of Camembert cheese, various film yeasts (yeast-like fungus) belonging to the genera of Geotrichum apparently contribute a thin surface growth and reduce the acidity of the cheese before the main mold P. camemberti establishes its acivity. However, the involvement of yeasts and their production of flavor components (using the supplied lipids or precursors) in cheeses are relatively unknown. Except for a few instances, it is believed that yeasts have a smaller role than molds and bacteria. This observation does not seem to be correct. It was reported that lower fatty acids are Eormed as a result of carbohydrate metabolism. Higher fatty acids are produced by lipolysis by Micrococci, gram-negative bacteria (Reiter et a]., 1967), and probably to a certain extent by yeasts. Carini and Volonterio (1969) reported that all of the 20 strains belonging to 8 different species of Torulopsis isolated from Taleggio cheese possess intracellular milk fat hydrolyzing (lipolytic) activity. Peters and Nelson (1961) reported that

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supplementation with lipases produced by C. lipolytica could improve the quality of blue cheese. In limburger cheese production, the yeasts Debaromyces kloeckeri and C. mycoderma liberate organoleptic fatty acids (myristic, palmitic, palmitolein, stearic, and oleic acids). Baker’s yeast finds most of its utility in bread manufacture. This yeast is responsible for the fermentative synthesis of alcohol and carbon dioxide. Many bacteria are also involved in the process of dough raising. Both bacteria and yeast produce an array of chemical compounds that are responsible for the final flavor and taste. Distinguishing between the roles played by yeast and bacteria in the formation of such characteristics is difficult. However, the cumulative fermentative action of both causes chemical changes in a variety of fatty acids, organic acids, alcohols, ketones, aldehydes, and carbonyl compounds (Kohn et al., 1963).

c. ORIENTAL FOODSAND PICKLES Oriental foods like tempeh are among the richest sources of fatty acids and other organic compounds needed for human health. Herring et al. (1991) observed that when the mold Rhizopus sp. is forced to synthesize lipids de novo, an increased percentage of up to 21% of ylinolenic acid is present in fermented Indonesian tempeh. In addition to these mold species, many bacteria are also encountered during tempeh fermentation. However, in general there has been no special mention about the role of yeasts in the fermentation of oriental foods in reviews by Hesseltine (1965), Hesseltine and Wang (1967), and Saisithi et al. (1966).It is believed that yeasts have a smaller role in the development of flavor and odor than molds and bacteria. However, in miso fermentation the soybean oil (present in the lipid substrate) is digested by koji lipase to produce some fatty acids. Many osmophilic yeasts (Zygosaccharomyces major var miso, Z. sdoja var miso; spore-forming Saccharomyces zygopichia, Debaromyces, Zygosaccharomyces, Hansenula, Pseudohansenula, Pichia, and non spore-forming Torulopsis) are also involved in miso preparation (Hesseltine and Wang, 1967), but their role in flavor development is unknown. Alcohols contribute to the pleasant smell of miso. Some yeasts and bacteria also make films and affect the odor. The presence of osmotolerant yeasts in Marzipan, fermented wine, butter, and fat-based foods have been reported (Mohs, 1974),but their roles are not clear. Most of the yeasts found in margarine and butter are lipolytic in nature (Aeyraepaeae and Lindstrom, 1974). Certain yeasts (Candida citeromyces, Debaromyces, Endomycopsis, Hansenula, and Torulopsisf are also capable of oxidizing hydrocarbons (catalyzed by oxygen-dependent transformation by monoxygenase) to

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yield fatty acids or lipid-containing surfactants (Finnerty, 1984). The biotransformation of lipid-containing substrates to useful and stereospecific flavorants in the food industry has been reviewed by Welsh et al. (1989). Goldman and Perret (1965) produced 15- and 16-hydroxy palmitic acids by hydroxylating Torulopsis magnoliae and these were further cyclized to form lactones. Lactones contribute taste and flavor nuances to food. Labows et al. (1979) reported the production of lactones by the yeast Pityrosporum cultured on a lipid substrate (triolein, sebum, lecithin, oleic acid, and Tween 80) producing y-hexa-, y-hepta-, y-deca-, y-undeca-, and y-dodecalactones. Pityrosporum glaucum or P. platinis are capable of producing methyl ketones from coconut oil. So far, no ascertainable scientific description of the significance of the involvement of yeasts in pickle fermentation has been reported. It is believed that the general increase in fatty acids in pickles is attributable to the unsaponifiable fractions and the hydrolysis of lipids during fermentation. Keil and Weyrauch (1937) observed acetylcholine and lactylcholine accumulation in foods fermented by Bacterium acetylchoIini [syn. Lactobacillus plantarum, (Rowalt 1948)]. IV. Medical Importance of Yeast Lipids

Fat is a concentrated form of stored energy (9 caloriedg) supporting various metabolic activities of the body. Following the discovery of essentiality of fats in the diet by Burr and Burr (1929), it was shown that polyunsaturated fatty acids such as 0 - 3 and 0 - 6 fatty acids are essential for human health (Holman, 1968, 1982; Brown et al., 1938; Conner et al., 1992; Drevon 1992). Fatty acids are essential because the human body is incapable of synthesizing them. Long chain w-3 fatty acids (C18:4,C,o:4, C,,:,, C,,:,, and CZ2:Jare formed by the desaturation of a-linolenic acid (C18: 3 ) while linoleic acid (C18: acts as the precursor for 0 - 6 fatty acids (C,,:,, C,,:,, C,,:,, C,,:,, and CZzi5). Usually, excessive intake of diets rich in fat results in obesity and, in some caes, coronary atery diseases and certain types of cancer (Leibel, 1992). Deficiency of 0 - 3 and 0 - 6 fatty acids leads to many symptomatic features of diseases of brain, heart, retina, liver, skin, and failure of reproductive systems in man (Conner et al., 1992).This can be corrected by the administration of fat emulsions through food or intravenously in patients suffering from acute deficiency diseases (Kirby, 1992). Essential fatty acids are supplemented in our food through sources of vegetables, meat, and marine fish. Marine lipids (i.e, fish oil) are the major source of both 0-3 and 0 - 6 fatty acids. y-Linolenic acid (0-6 acid) is usually extracted from Primrose (Oenothera biennis), borage (Borago

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officinalis), and many species of Ribes. Some species of molds belonging to the genera Mortierella and Mucor are also good producers of w-3 and w-6 fatty acids. However, so far, no yeasts have been reported to produce such acids. The important w-3 fatty acids in the human plasma are alinolenic acid (C18:3) and its derivatives eicosanpentanoic and docosahexaenoic acids (Katan et al., 1991). It is already known that w-3 fatty acids serve as precursors for the synthesis of many essential compounds, thrombaxane B 3 , A3, prostaglandins E3,13, and leukotriene. Also, these substances have important physiological effects with regard to immune and platelet functions. Pharmacopeal requirements of different commercially available primary dried yeasts [S. cerevisiae and C. utilis (torula yeast)],for use in foods, mention only the proteins (- 40%) and vitamins 0.0012 mg%; riboflavin 0.0004 mg%, and (thiamine hydrochloride 0.0025 mg% nicotinic acid) and say nothing about lipid contents (Osol and Pratt, 1973). Such yeasts have not been considered a major source of essential fatty acids; however, other yeast species contain linoleic and linolenic acids in appreciable quantities (Table I). In general, linoleic acid occurs at higher levels than linolenic acid and the latter is absent in some yeasts. Leucosporidium species contain these acids as 80.23 to 87.35%of total unsaturated acids whereas some S. cerevisiae contain 35.96% (Table I ) . However, the actual utility as well as metabolism of yeast-derived linoleic and linolenic acids in the human body has not been demonstrated. It can be presumed that upon ingestion of such acids, they serve as precursors for the synthesis of elongated essential acids needed for the body metabolism.

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V. Modification of Lipids

The use of modified lipids with desirable qualities in food and beverages has much commercial as well as nutritional importance. As mentioned earlier, derivitization of such products can be achieved by biotechnological means, fermentative synthesis using wild or genetically engineered organisms, and chemical or biochemical interesterification. Lipases, in general, are reasonably stable and stereoselectively catalyze substrate modification in four different ways: (1) ester hydrolysis, (2) ester synthesis, (3) transesterification, and (4) acyl transfer (Erdman et al., 1988). Some of the important general sources of lipases, other than yeasts, are listed in Table 11. A separate list of commercially important yeast lipases and their applications are given in Table 111.

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A. FERMENTATIVE SYNTHESIS AND MODIFICATION OF LIPIDS

Fermentation is the cardinal step in which the organism multiplies and accumulates lipids or excretes lipid-containing surfactants. The metabolic aspects of yeast lipids have been described by earlier workers (Ratledge, 1989). Yeasts can be grown in large quantities under submerged conditions using cheap and available carbon sources such as agroindustrial wastes, food industry wastes, and petrochemical industry wastes like n-alkanes. Lipid production patterns depend mainly on the type of oleaginous organism and its growth conditions (Ratledge, 1982, 1989; Yoon and Rhee, 1983a). In principle, once a good range of productivity of a particular product is achieved in batch culture, it is possible to increase productivity in continuous or fed-batch systems as the next step (Iwamoto, 1972). Standardization of fed-batch culturing seems to be a difficult task, especially with regard to the nitrogen and dissolved oxygen levels in the medium. However, Yamauchi et al. (1983)achieved a productivity of 21% while working with fed-batch culturing of Lipomyces starkeyi. It was also shown that a dissolved oxygen level of 40% is required at the lipid accumulation stage of the yeast Rhodotorula gracilis (Yoon and Rhee, 1983b; Choi e t al., 1982; Pan et al., 1986). Nevertheless, attempts in our laboratory using Rhodotorula gracilis CFR-1 (fed-batch culturing trials with an economically cheap cane molasses medium) were discouraging. Such trials yielded good biomass, but low oleaginicity. Changes in external environmental conditions such as temperature, pH, substrate, and oxygen can bring about changes in intracellular lipid composition. Oxygen is required for the conversion of stearic acid to oleic acid which, in turn, is converted to linoleic and linoleinic acids. Desaturation starts with a low oxygen supply and the production of major storage lipids occurs after the start of nitrogen, phosphorus, sulfur, and iron depletion from the medium (Pandey, 1989a). Even a small change in a single parameter brings about many sequential and interdependent biochemical changes in the pathway. The lipid metabolism of oleaginous yeasts as elucidated by Evans and Ratledge (1985) and Ratledge and Evans (1988) suggested that once the nitrogen is depleted more ATP is synthesized. The ATP:citrate lyase, one of the key enzymes involved in the pathway, cleaves citrate (found in the cytosol) to oxaloacetate and then to acetyl CoA, the basic building block in the synthesis of fatty acids. The synthesis of triacylglycerols takes place in the microsomes. Of the many controlling factors involved, higher C/N ratios affect the lipid productivity. Gill e t al. (1977) achieved more lipids [(- double

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TABLE I POTENTIAL YEASTSAND THEIRLIPIDSTHATCONTAINLINOLEICAND LINOLENIC ACIDS r

04,

Fatty acids"

% Total ofb

(D

4

Organism

A

B

Debaromyces castelii D. hansenii Hansenula anomala H. anomala H. polymorpha Lipomyces kononenkoae Metschnikowia lunato Saccharornyces cerevisiae S. fibuligera Torulospora delbrueckii Yarrowa lipolytica Zygosaccharomyces rouxii

33.7 23.2 40.8 42.8 43.4 25.7 49.5 28.7

0.4 17.8 4.9 19.2 0.5 8.2

44.8 42.7 34.8 43.2

Traces

0.8

-

A&B~

C C

Reference

43.16 48.92 53.13 73.63 59.48 46.95 61.95 35.96

21 16.2 14 15.8 26.2 28.8 20.1 20.2

Moulin et al. (1975) Moul;n et al. (1975) Johnson and Brown (1972) Ng and Laneellee (1977) Dedyukhina et al. (1982) Hossack and Martins (1978) Malkhas'yan et al. (1983) Kovac et al. 11980)

59.14 47.49 59.79 51.30

33.9 20.1 47.8 15.8

Malkhas'yan et ol. (1983) Johnson and Brown (1972) Klug and Markovetz (1967) Watanabe and Takakuwa (1984)

Rhodotorula glutinis Yarrowa lipolytica Brettanomyces anomalus Candida albicans C. albicans C. guilliermondii C. humicola C. kefyr C. kefyr C. rugosa C. sake C. tropicalis C. utilis Cryptococcus ater Leucosporidiurn frigidurn L. frigidurn L. nivalis A. glutinis R. gracilis

49 51 33.7 26.8 33.5 39.1 61.3 36.1 31.9 48.5 33.8 32.8 54.1 48.7 40 25 42 53.1 34

3 1 0.4 19.8 24.7 3.6

Traces 14.9 16.7

nil 10.8 13.8 9.9 33 51 27 18

A, linoleic acid; B, linolenic acid. Total percentage of A and B, based on total unsaturated fatty acids Total percentage of saturated fatty acids.

65 60.46 43.16 60.9 77.8 51.13 71.52 62.42 57.17 57.8 51.98 50.53 79.69 74.17 84.61 87.35 80.23 62.69 65

20 14 31 23.6 25.2 16.5 14.3 19.3 15 16.1 14.2 35.1 14.8 21 9 13 14 15.3 20

Malkhas’yan et al. (1983) Malkhas’yan et al. (1983) Moulin et al. (1975) Nishi et al. (1973) Guarneri et 01. (1977) Jigami et al. (1979) Zotova et al. (1985) Moulin et al. (1975) Moulin et al. (1975) Iida et al. (1980) Kaneda and Smith (1980) Greshnykh et al. (1968) Johnson et al. (1972) Moulin et al. (1975) Watson et al. (1976) Watson et al. (1976) Watson et a]. (1976) Kaneko et al. (1976) Kessell (1968)

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Z.JACOB TABLE I1 SOURCES OF LIPASES OTHERTHANYEASTS FOR THE PURPOSE OF FATSPLITTING Sources

Animal sources Porcine pancreas Gastric Plant sources Castor bean Wheat germ Microbial sources Molds Aspergillus niger Mucor jovanicus Rhizopus arrhizus R. delamer Bacterial Chromobocterium viscosum Pseudomonos sp. Unspecified organism

Specificitya

Remarks

A A

Commercially available Commercially available

A

Commercially available Commercially available

-

Mostly nonspecific B Commercially available B Commercially available B Commercially available B Commercially available B B B

Commercially available Commercially available Commercially available, splits triacylglycerols to diacylglycerols

A, positional and chain-length specificity: B, nonspecific

the amount), 50% (w/w)]in Candida lipolytica cells when the nitrogen supply was limited to 246 mg/liter than with normal conditions of fermentation. Similarly, other nutritional conditions, especially reactor conditions, also have a tremendous effect on the productivity and composition of the lipids as well. This point has been described in many of the earlier published reviews (Ratledge, 1989; Rattray, 1988). In Rhodotorula gracilis CBS 3043, the majority of triacylglycerols are produced during the stationary phase in a nitrogen-limited medium (Rolph et a]., 1989). The phospholipid concentration of the cells is adversely affected when the cells are grown in a carbon-limited medium, although there is little effect on triacylglycerol accumulation and the quality of lipids. However, in the earlier days of yeast lipid the expensive synthesis as well as a doubtful market introduction as an alternate source to vegetable oils and fats precluded large-scale production. This opinion has now changed because of the advent of biotechnological techniques and downstream processing which lead to the conclusion that such processes are economic if efforts have been concentrated on value-added products like y-linolenic acid or a cocoa butter substitute. When considering metabolic pathways, diverse modi-

TABLE I11 INDUSTRIALLY IMPORTANT YEAST ENZYMESAND THEIRAPPLICATIONS Organism

Nature of enzyme

No positional specificity and activity on long chain triacylglycerols. Hydrolyze almost all ester bonds and liberate all sorts of acyl chains on treatment. Olive oil and cocoa butter were used as substrates in this study (Benzonana and Eposito, 1971). Used in detergents. Commercially available. Meito Sangyo Co., Japan. Nonspecific (lipase) Commercially available from Sigma (United States) or Deisenhofen Specific (lipase) (Germany). ca.600,000 units/solid, enantioselective transesterification for producing L-methyl esters (Erdman et al., 1988). Economic fermentation, 1980 units/ml. Olive oil as substrate. Chain Specific (lipase) length specificity for C14 to C18 or C2O-C22 acyl groups (Lie and Lamberstein, 1986; Noguchi and Hibino, 1984). Specific (lipase) Chain length specificity to lower chain acyl groups and poor specificity for long chain acyl groups (Noguchi and Hibino, 1984). Nonspecific (Lipase-MY) Used in improving flavor of apple wine (Tanabe Seiyaku Co., 1970). Nonspecific (lipase) Thermostable, 55-60°C, used for fat spliting (Montet et al., 1985). Nonspecific (lipase) Thermostable, 55-60°C, used for fat spliting (Muderhwa et al., 1985). Used in detergents and in blue cheese manufacture (Peters and Nelson, Nonspecific (lipase) 1961). Lipase Enzymes from both yeasts are used in limburger cheese production, liberating myristic, palmitic, palmitoleic, stearic, and oleic acids from milk fat (Seitz, 1974). Specific (lipase) Specificity for sn-1 and sn-3 positions (Macrae and Hammond, 1985; Muderhwa et al., 1986). Specific (lipase) Specificity for polyunsaturated fatty acyl groups (Baldwin, 1986). Specific (lipase) Hydrolysis of long chain fatty acids at cis 9 or cis 9, 1 2 positions. Commercially available (Macrae, 1983; Jensen, 1974; Jensen and Pitas, 1976). Specific (pectic enzyme) Used in mechanical extraction of olive oil with higher FFA and lower peroxide values. Oil with acceptable qualities. (Servilli et ol., 1989).

Candida cylindracea (C. zeylanoides) Nonspecific (lipase)

C. cylindracea C. cylindracea

C. cylindracea

Candida sp. Candida sp. C. curvata C. deformans (Yarrowa lipolytica) C. lipolytica C. mycoderma Debaraomyces mycoderma Rhodotorula pilimanae

R. rubra Geotrichum candidum (yeast-like fungus) Cryptococcus albidus

Remarks, applications if any, and reference

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fications of intracellular fats may be possible (Noguchi et al., 1982). Ratledge (1970) reported the production of shorter chain length fatty acids (Cll to CIS)by Candida 107 and discussed the possibility of producing a cocoa butter substitute from yeast lipids. Normally, yeasts do not produce these special products, but could within limits synthesize compounds that are closely related. For example, let us analyze the case of a cocoa butter substitute. Natural cocoa butter contains 35% of stearic acid in the triacylglycerol. With this in mind many attempts have been made to produce a cocoa butter substitute by adopting different protocols. Noguchi and Hibino (1984) patented a process wherein they could produce a comparable content of stearic acid in species of Rhodotorula and Candida by supplementing stearic acid or its esters in the growth medium. In another attempt, Moreton (1985) rerouted the synthesis of stearic acid in yeasts (Candida 107, Trichosporon cutaneum, and Rhodosporidium toruloides) by supplementing sterculic acid in the medium. The sterculic acid inhibits the A-9 desaturase activity which results in more stearic acid synthesis. But the technique of inhibition of A-9 desaturase activity does not work with yeasts like Lipomyces and Saccharomyces cerevisiae. Certain groups of yeasts are also capable of producing industrially important lipid-containing biosurfactants of varying chemical nature. For general information interested readers may refer to Jacob (1992), ZajiC and Saffens (1984), and KosariC et al. (1987). Modification of oils and fats is also possible by fermentative growth and subsequent lipolysis with yeasts. Glatz et al. (1984) studied the modification of fats by fermentation using C. lipolytica. A yeast like C. curvata is capable of digesting and absorbing low-grade fats and oils. It produces lipids structurally related to the substrate supplied. During the growth of the yeast, it produces modest amounts of palmitic acid while the linolenic acid content is drastically reduced. The substrate is presumably hydrolyzed and resynthesised during deposition, resulting in an oil with an altered but nonrandom glyceride structure of triacylglycerol. According to Weete (1980), one should expect only reasonable alterations of fatty acids deposited in the cells. Mathew et al. (1990) studied the possibility of producing essential fatty acids by the hydrolytic action of microbial lipases on lipids. The mono- and diacylglycerols formed in the experiment were used as emulsifiers in ice cream, cakes, and puddings. Kajs and Vanderzant (1980) used S. lipolytica and C. utilis (food yeasts) for the emulsification of tallow. Their lipolytic enzymes degraded the tallow into fatty acids which served as a carbon source for their growth. It is amazing to see the enormous presence of lipolytic (lipase and

-

~

YEAST LIPID BIOTECHNOLOGY

199

esterase) yeasts in our daily life. Alifax (1979) isolated 372 strains of yeasts from dairy, meat, bakery products, and cooked dishes. More than 70% of the strains possessed esterases, lipases, or both. The strong action of such enzymes produced by Candida curvata, C. deformans, C. humicola, C. parapsilosis, C. lipolytica, Crptococcus laurentii, Saccharomyces lipolytica, and T. cutaneum liberates fatty acids from butter oil, especially capric, caprylic, and lauric acids. Some yeasts possess isomers. Another interdifferential lipase activity against C, :, and C,: esting example is that of Endomycopsis biospora which can utilize fatty acids of rape seed and maize germ oil as the sole source of carbon. Candida lipolytica and T. cutaneum also have the capability to preferentially liberate oleic and myristic acids, respectively. As mentioned previously, many theoretical modifications are possible but only those that are physiologically acceptable will allow the organism to survive and grow. Inhibition of a particular step of metabolism may be achieved using substrate analogs and enzyme inhibitors. In this case, a number of problems need resolution. One of the constraints is the regulation of the mechanism that determines the level of intracellular accumulation of metabolites. Repressive or feedback adjustments have to be made to limit the amount of products like sterols in the cells. While doing so, however, it would be advantageous if other biosynthetic mechanisms were not disturbed.

B. GENETIC ENGINEERING ASPECTS OF YEAST LIPIDMODIFICATION Although lipids exist in various forms in nature, their commercial exploitation has been restricted because of constraints of availability and processing costs. There is then a good justification in exploring microbial sources employing gene manipulation techniques to produce many of the high value lipids. Gene manipulations are possible by mutant selection, hybridization, rare mating, spheroplast fusion, transformation, and gene cloning. Of these, mutant selection, spheroplast fusion, transformation, and gene cloning, through vectors or plasmids, are the generally employed techniques. Both plants and yeasts are mostly polyploid in nature. In contrast to the developments in plant genetic engineering, limited success has been possible with engineered industrial yeasts. In 1968 Larikova and Gal’tsova reported an almost twofold increase in total lipids and stearic acid content in yeasts treated with radiometric methyldichloroethylene and X rays. No further reports appear in the literature. Success in the development of engineered yeasts for lipid production seems to have failed because of the limiting stability of such recombinants, the lack of understanding of the metabo-

200

2. JACOB

lism of lipids under stressed conditions, and suitable techniques for the genetic characterization of various traits. One of the key enzymes in fat metabolism is fatty acid synthetase. It is a multienzyme with a molecular mass of 2.19 x lo6 Da and six protomers each with a and p subunits. The coded genes’ locations in the reported map are termed fasl and fas2 (Pandey, 1989b; Siebenlist et al., 1990). Pandey (1989b) reported a successful isolation of fas mutants from S. cerevisiae which received the fas complex from two oleaginous candidates, Rhodotorula gracilis and Candida sp. Further, a recombinant called 63a was obtained from a cross of both yeasts by protoplast fusion. The recombinant had the sugar tolerance (40%) and conversion efficiency (11-15%) of S. cerevisiae, and the lipid composition resembled that of palm oil. The production of such a microbial oil is not economical in terms of the capital cost of production or the value of palm oil in the world market. In another case, Hammond et al. (1981) produced lipids with palmitic acid similar to the cocoa butter substitute by growing mutants of C. curvata at 30°C. However, the same results were not obtained when the mutants were grown at higher and lower temperatures and at different pHs. Another serious disadvantage with such mutants is the instability of their desirable characteristics. Knowledge of stress conditions involved in turning on and off of the concerned genes to make appropriate new sets of enzymes and effector molecules is necessary in studies of oleaginous yeasts (Joseph, 1989). It seems that the stressed conditions also affect the rate of synthesis of other metabolites during lipid metabolism. According to Joseph (1989), acetyl-CoA is a key intermediate compound for the synthesis of carotenoids in Rhodotorula gracilis. This hypothesis was further strengthened when Joseph (1989, and unpublished data) observed that mutants of R. gracilis CFR-1 deficient in carotenoid pigments and ATP:citrate lyase produced only very insignificant quantities of lipids in addition to being devoid of carotenoid pigments. Sterols of yeasts are potential precursors for the chemical synthesis of hormones, growth mutators, drugs, and potent fungicides. Parks et al. (1984) described the tailoring of the yeast S. cerevisiae for sterol production by genetic manipulation. This yeast can be grown both aerobically and anaerobically and has a well-defined genetic mechanism that can be suitably manipulated to construct various combinations of markers, including many in the biosynthesis of lipids (Henry, 1982). Saccharomyces cerevisiae is not an oleaginous yeast, but recent developments may change this situation. Boulton and Ratledge (1981) observed a positive correlation between lipid accumulation in yeast and the presence of ATP:citrate lyase which, according to them, provides

YEAST LIPID BIOTECHNOLOGY

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a constant source of acetyl-CoA for fatty acid synthesis. Genetic determinants of ATP:citrate lyase could be introduced into S. cerevisiae by mutation and subsequent positive selection using resistance to polyene antifungal agents like nystatin (which make a complex with sterols in the cell membrane). However, the mutants thus obtained usually have a loss of single enzymic function in sterol biosynthesis. This indicates that many modifications may be possible, but only those that are physiologically acceptable will allow the organism to survive and grow. In addition to using sterol mutants to obtain desired end products, the inhibition of specific steps in enzymic pathways in wild types may be achieved using substrate analogs or inhibitors. In doing so, many practical problems may arise, especially while controlling repressive or feedback adjustments needed for a particular pathway.

c. CHEMICAL AND BIOCHEMICALINTERESTERIFICATIONS Modified fats have become essential food components of a myriad of sensory appealing, fast and convenient foods (mostly confectionary and baked foods). In the past, the hydrogenation of vegetable edible oils by inserting hydrogen into the double bonds of unsaturated fatty acids was found very useful in evolving plastic fats. Later it was found that the spatial rearrangement of fatty acids on the triacylglycerol by an acidolysis reaction or an ester-ester interchange yielded fats with unique properties. The products thus obtained are called speciality or modified fats. Cocoa butter fat, characterized by the high content of stearic acid (30-35%) and the predominance of l-palmitoyl-2-oleoyl-3-stearoyl glycerol, is a commercially important component of the food industry. Most soft soap and cosmetic industries rely on coconut oil for its medium chain length fatty acids (Clz and C1J. Other fats and derivatives of commercial importance are glycolipids as surfactants, carotenoids as food colorants, and poly-P-hydroxybutyrate with its unique plasticlike properties. High value fatty acids like y-linolenic acid (6,9,12octadecatrienoic acid), arachidonic acid (Cz0:J, eicosapentaenoic acid (Czo:J, and eicosahexaenoic (Czo acid are medically important. As mentioned earlier, seeds of Oenothera are the prime source of ylinolenic acid, and the total production is insufficient to meet the increasing demand. Now the question is whether these compounds can be synthesized through chemicallbiochemical means. Although the chemical interesterification reaction steps are easily performed, there is the serious disadvantage of nonspecificity yielding products with variable rheological properties. This unpredictable range of modifications can be

202

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achieved with the help of catalysts like sodium methoxide, metallic sodium, or sodium potassium alloys. These reactions do not involve costly equipment. Biochemical interesterifications are more useful in the sense that they give unique fats with rather specific and predictable rheological properties. Microorganisms use their extracellular lipases (glycerol ester hydrolase, EC 3.1.1.3) for digesting lipid substrates for metabolic activities. According to Macrae (1984), extracellular microbial lipases can be used for interesterification so as to obtain modified lipids which otherwise may not be easily obtainable by chemical interesterification. The reaction is reversible under certain conditions, and a particular organism’s own lipases may not have much significance in its own biosynthesis of modified oils and fats intracellularly (Tsujisaka et al., 1977). The natural substrates for lipases are triacylglycerols of long chain fatty acids. The enzyme acts at an interface between an insoluble substrate phase and an aqueous phase resulting in hydrolysis of a wide range of insoluble fatty acid esters. As mentioned previously, this is reversible. Consequently, hydrolysis and resynthesis of acyl glycerol groups occur when lipases are incubated with a mixture of triacylglycerols. This hydrolysis and resynthesis causes migration of fatty acyl groups between glycerol moieties and gives interesterified products. Extracellular lipases for interesterification can be grouped into two main categories according to their specificity of reaction (Brockerhoff and Jensen, 1974) (Table 11). The first group is nonspecific and the products formed have a random distribution of fatty acids as the acyl groups. This is similar to the chemical interesterification reaction. Enrichment of triacylglycerol is also possible by interesterifications. A mixture of triacylglycerol and free fatty acids as reactants are treated in the presence of lipase. The free fatty acids are exchanged with fatty acyl groups of triacylglycerol to give modified free fatty acid and triacylglycerol. Another possibility involving migration of fatty acyl glycerol and fatty acids also occurs during interesterifications. Hydrolysis and resynthesis of acyl glycerol and migration of fatty acyl groups between glycerol moieties occur when a mixture of triacylglycerols is treated with lipase. The second group is 1,3 specific. This is common with lipases of different molds such as Aspergillus niger, Mucor javanicus, and Rhizopus species (Okumura et al., 1976; Ishihara et al., 1975; Semeriva et al., 1967; Macrae, 1983). Interesterifications are performed in either stirred or packed bed reactors, depending on the type of operation (batch or continuous). For example, in a batch reaction, a mixture of palm oil mid-fraction and

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stearic acid is dissolved in petroleum ether and stirred for 16 hr at 40°C with a catalyst. Enzymatic interesterification catalysts are prepared by coating macroporous inorganic particles with lipases. For example, the A. niger lipase is precipitated with acetone onto kieselguhr particles and is activated by hydration with a small quantity of water prior to the addition to the reactor. Kimura et al. (1983) reported the use of immobilized lipases of C. cylindracea on various organic as well as inorganic supports, and the hydrophobic matrices gave the highest activity in the hydrolysis of olive oil. In the chemical interesterification process, sodium metal or sodium alkoxide promotes the migration of fatty acyl groups between glycerol molecules. In enzymatic interesterifications, it is possible to minimize the rate of hydrolysis of fats by reducing the amount of water in the reaction system and then lipase-catalyzed interesterification becomes dominant (Coleman and Macrae, 1980; Matsuo et aI., 1980,1981;Tanaka et al., 1980).Enhanced interesterifications are also possible by providing a large area of interface and a minimum quantity of water in the supports. Supports include kieselguhr, hydroxyl apatite, and alumina. Catalysts are prepared by adding a solvent such as acetone, methanol, or ethanol to a slurry of the particles in a buffered lipase solution (Macrae, 1983). The solvent precipitated enzyme coats the particles and they are collected by filtration. They can be dried and stored. These dried particles contain low activity of lipases. Hydration with up to 10% of their weight with water activates the catalyst particles. Reactivation of supports is also done by coating it with diols or triols and free fatty acids or treatment with the reactants dissolved in petroleum ether or hexane. Erdman ef al. (1988) suggested the use of enzyme immobilization carriers such as VA-epoxy Biosynth and Duolite ion-exchange resins. Activation by diols or triols such as glycerol has also been recommended (Tanaka et al., 1980). It is interesting to note that the lipase of Geotrichurn candidum has a very marked specificity for hydrolysis of a particular type of long chain fatty acids containing a cis-double bond in the ninth position (Macrae, 1983; Jensen, 1974, 1983; Jensen and Pitas, 1976). Most of the site-specific lipolysis was found to be very slow. However, this disadvantage of slow reaction can be overcome using methyl esters of oleic, palmitoleic, linoleic, and linolenic acids (all of which has a cisdouble bond in the ninth position). During the interesterification, A 4 fatty acyl groups are selectively exchanged with other A-9 acyl groups in the mixtures of triacylglycerol or triacylglycerol and free fatty acids. For example, when olive oil + linoleic + stearic is treated with the

204

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lipase, the triacylglycerols are enriched in linoleate at the expense of oleate, while the saturated fatty acyl content of triacylglycerols remains substantially unchanged. In another study a slow process of interesterification at 40°C for 400 hr using stearate and Rhizopus lipase was reported. In this case, after an initial equilibration period of a few hours, the water was removed from the hydrated catalyst particles with the formation of a larger quantity of diacylglycerol and free fatty acids until steady-state conditions were attained. To date there are no authenticated reports of extracellular lipases which catalyze reactions only at the second position of acyl glycerols. Most of the extracellular lipases have little fatty acid specificity (Macrae, 1983). Lipolytic organisms with specific reactions at the desired sites of triacylglycerol need to be isolated. The report of the development of a rapid plate procedure by Collins and co-workers (1989) for the isolation and characterization of lipolytic microorganisms is noteworthy. With this technique microbial lipases can be conveniently detected by the digestion of a target lipid (emulsified in agar medium). The specific lipolytic activity is revealed by the cleared zones due to precipitation of liberated fatty acids by Ca2+in the agar medium. Lipolytic organisms of specific properties are also isolated from different natural sources like Elaeis quineensis fruits. Interestingly, higher levels of free fatty acids in palm oil are due to the presence of yeasts and molds which are lipolytic and exist in the mesocarp of fruits from the inception of the fruit development. Isolation of such types of microorganisms also could be used for insertion and expression of desired traits for novel lipases. VI. Commercial Significance of Yeast Lipid Biotechnology

The presently available biotechniques in the area of yeast biotechnology seem to be handicapped by uneconomic productivities. Nevertheless, it would be worthwhile to mention the significance of such products in food and medicine. Future commercial potentials for the production of tailor-made as well as modified, speciality fats would be expected to increase mainly because of the ever increasing nutritional and medical awareness, modern food habits, and prolonged life span of man. The importance of quality lipids with no rancidity and a higher degree of polyunsaturated fatty acids in foods has already been accepted as important nutritional criteria. Consumption of rancid lipids may lead to peroxidation and thus damage at the cellular level. With regard to essential fatty acids of importance in food and medicine (prostaglan-

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dins) and newer or novel derivatives of such products, lipids may be helpful in resolving many of the problems in nutrition science, and probably in the curing of many diseases. For such purposes, natural sources like yeasts have not been exploited yet. Genetically engineered yeasts can offer many positive promises for the future. Figure 1 represents prospects of lipid biotechnology in a nutshell wherein the role of yeasts seems to be quite significant. Higher productivity of different kinds of vegetable oil seeds is possible by adopting the latest agricultural practices in larger areas. As previously mentioned, cloning and expressing of desired intraspecific genes to the hosts by recombinant DNA techniques and tissue culture practices may help in raising more seedlings and thus larger-scale cultivation. Alternatively, lipids similar to vegetable oils can also be produced microbially by fermenting selected candidates under appropriate conditions using suitable and cheaply available substrates (agroindustrial wastes, whey, molasses, meat industry wastes, and petrochemical carbon sources like

FIG.1. Lipid biotechnology prospects.

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n-alkanes). Chemical or biochemical alterations of the postharvest microbial, plant, or animal lipids can be facilitated in reactors to yield modified lipids. Such a product can also be produced by genetically engineered yeasts or somatic cells possessing the qualities of high oleagicity and productivity and providing the natural conditions of biomass metabolism and lipid accumulation in reactors. Tailor-made microbial lipids can also be synthesized using yeasts in which the natural metabolic pathway is rerouted in a desired manner. This method cannot be considered as a fully natural process. In normal practice the results of such techniques are discouraging and unproductive. Although both plants and yeasts are polyploids, the former has an edge in the success of cloning and expression of desired genes. The question of whether lipids obtained by the natural metabolic pathway of a particular organism can be used as a carbon source for the synthesis of modified fats by genetically engineered organisms or by other types of wild organisms is not yet fully answered. Natural lipids produced by both wild as well as engineered candidates can be cycled for chemical as well as enzymatic conversions to modified products. It may also be possible to recycle these modified lipids as carbon sources for a particular candidate’s metabolism, thus obtaining novel products. It may be quite possible in the future to incorporate all of the desired traits in specific cells (somatic/mammalian) in producing tailor-made lipids in reactors. Additionally, a modified lipid can be further modified by any of the methods previously described to produce novel lipids which may find applications in food and medkine. -~ The logic of using yeasts as enriching agents to derive desired food products and/or to degrade biological wastes is understandable. The following examples reinforce this logic. Burkholder and Gervasini (1969) reported that the growth of C. lipolytica and G. candidum can be used for reducing fats in fish meat. After fermentation the final product has an increased protein content with appealing flavor characteristics. In another instance it was suggested that inedible tallow can be utilized as a carbon source for the production of SCP and metabolites (Tan and Gill, 1984). This was studied by Kajs and Vanderzant (1980) using S. lipolytica and C. utilis. Lipolytic activity of the enzyme produced by the yeasts helps in the breakdown of the tallow into simpler fatty acids which act as carbon sources for the production of SCP (Fiorentini et a]., 1976). Anelli et al. (1975) reported the production of SCP (containing all amino acids according to FA0 standards, except methionine) from the growths of C. lipolytica, Torulopsis holmi, C. mesentrica, and Cryptococcus albidus on fat wastes containing olein. When Friesian cows were fed torula yeasts the cows produced more

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milk with a higher level of lipids (0.08 to 0.29%) (Gheorghiu et al., 1976). However, when Toprina yeasts were fed to Friesian cows, although there was no significant increase in the total lipid content, the internal tissues were enriched significantly with C,,, C,,, and C,, : 2 fatty acids (Martillotti et a]., 1977). When the Ruse strain of S. cerevisiae was fed as fodder the lipids contained a large amount of a-tocopherols (6.32 pg/g) (Roshkova and Beshkov, 1974). Similarly, when Holstein cows were fed diets with yeast there was an increase of lipids and protein in the milk (Erdman and Sharma, 1989). In another report, yeast biomasses of Toprina and Liquipron were fed to cows and pigs and subsequently the milk and adipose tissues had odd-numbered fatty acids (Boniforti et al., 1979). Rys et al. (1975) reported the feeding effects of an n-paraffin- or molasses-grown yeast in pigs wherein an increased proportion of C,,, C,,, CI6: and C,,: and a decreased proportion of CI8:, and C,,:, fatty acids in the back fat were observed. It was also demonstrated that rats fed n-paraffin-grown Candida sp. did not produce any untoward effects (Yokoyama and Kaneda, 1972). Defatting of meats by microbial action is an interesting phenomenon. Glatz et al. (1984) suggested that lipolytic organisms could be used to degrade meat cholesterol. Another possible application of lipolytic yeasts (for example, Saccharornycopsis lipolytica which has the capability to degrade meat) would be in the degradation of meat-processing wastes. However, the feasibility or success of such applications remains to be seen.

,

VII. Conclusion

Some yeasts are potential producers of lipids similar to vegetable oils and fats. Research on their applications as a dietary supplement in food or essential pharmacological components in medicine has not progressed to desirable limits. When considering the uneconomic fermentative synthesis of yeast lipids in reactors, a reorientation in the approach to develop processes for value-added lipids for use in food and medicine seems to be productive. Modification of lipids using engineered or transgenic yeasts has not been successful so far. Since yeasts are polyploid in nature, similar to plants, they may be suitably engineered so as to synthesize novel lipids which may find utility in producing value-added oils and fats for use in the food and biomedicical industries. Rerouting of the yeast’s metabolic pathway for the synthesis of biomedically important polyunsaturated fatty acids has not received much attention. The cloning and expression of specific fatty acid synthetase genes to produce specific lipids still has a long way to go, and

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J p ect in, Pect in ase , and Proto pect inase : Product io n i Properties, and Applications TAKUO SAKAI, * TATSUJI SAKAMOTO," JOHAN HALLAERT,? AND ERICKJ. VANDAMME? *Department of Agricultural Chemistry College of Agriculture University of Osaka Prefecture Osaka 593, Japan +Department of Industrial Biochemistry and Microbiology Division of General and Industrial Microbiology University of Ghent B-9000 Ghent, Belgium 1. Introduction 11. Review of Pectin A. Nomenclature B. Chemical Constituents and Structure C. Occurrence and Function D. Properties E. Determination and Characterization F. Pectin Manufacturing: Chemical Extraction and Purification G. Applications of Pectin Ill. Classification of Pectic Enzymes A. Esterases B. Hydrolases C. Lyases IV. Role of Pectic Enzymes in Phytopathogenesis V. Applications of Pectinases A. Industrial Production of Pectinases B. Fruit Juice Industry C. Other Applications VI. Protopectin-Solubilizing Enzyme (Protopectinase) A. Assay of Protopectinase activity B. A-Type Protopectinase C. B-Type Protopectinase D. Applications of Protopectinase References

I. Introduction

Pectic substances are acid polysaccharides of high molecular weight that are widespread in the plant kingdom. The size, charge density, charge distribution, and degree of substitution of pectin molecules may be changed biologically or chemically. The chemical structure of pectin has been the subject of many scientific reports for more than 50 years 213 ADVANCES IN APPLIED MICROBIOLOGY,VOLUME 39 Copyright 5 1993 by Academic Press, Inc. All rights of reproduction in any form reserved.

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(De Vries eta]., 1981). Elucidation of this structure was and is important because of the function of pectin in the cell wall as a “lubricating” or “cementing” agent (Rees and Wight, 1969), its role during ripening of fruit (Knee, 1978), its role in food processing (Rombouts and Pilnik, 1978; Van Buren, 1979), and its role as nutritional fiber. Research in the field of “the pectic substances” has been carried out by scientists and technologists from a great variety of disciplines. In the very early days of pectin chemistry, when French botanists such as Fremy were commencing to investigate the role of pectic substances in plant physiology, other chemists of the same nationality were delving into the finer details concerned with flavor in wines and ciders (Charley, 1951).The pectins in these beverages were considered to be responsible for the “veloute” or “moeilleux” (softness or silkiness) which give the fruit product its particular character. At present, interest in the pectic substances is still continuing worldwide. The pectin production process consists mainly of an acid extraction, followed by a (partial) purification of the extract and, eventually, precipitation and drying. However, this process has several disadvantages: maceration of the pulp, difficult filtration of the residue, and corrosion of equipment. The main raw materials for pectin are the peel of citrus fruits and apple pomace (Towle and Christensen, 1973). Research was initiated with the aim of developing an improved alternative microbial pectin production process, by means of a specific enzyme. A major part of this chapter is concerned with this specific enzyme, called protopectinase. II. Review of Pectin

In his classical book “The Pectic Substances’’ Kertesz (1951) stated that fruit jelly making was practiced long before pectin was discovered. The first information on water-soluble substances with a strong jellying power, occurring in fruits, was presented by Vauquelin in 1790. In the next well-known scientific publication on these substances, Braconnot (1825) related the name of these substances to their jellying properties when he derived it from the Greek work “ T ~ X T O C T , ” meaning to congeal or solidify: “. . . en attendant, je propose le nom pectique, de T ~ X T O C T , coagulum, pour distinguer ce nouvel acide de ses congeneres.” A. NOMENCLATURE

In the past, a number of confusing terms have been proposed for various pectic substances. In 1944, the Committee for the Revision of the Nomenclature of Pectic Substances, a former subdivision of the

PECTIN, PECTINASE, AND PROTOPECTINASE

215

American Chemical Society, finally accepted the following definitions (Table I): “Pectic substances” is a group designation for those complex colloidal carbohydrate derivatives which occur in, or are prepared from, plants and contain a large proportion of anhydrogalacturonic acid units which are thought to exist in a chain-like combination. The carboxyl groups may be partially esterified by methyl groups and partially or completely neutralized by one or more bases. “Pectinic acids” are the colloidal polygalacturonic acids containing more than a negligible proportion of methyl ester groups. “Pectinates” are either normal or acid salts of pectinic acids. “Pectic acids” is the group designation applied to pectic substances mostly composed of colloidal polygalacturonic acids and essentially free from methyl ester groups. “Pectates” are either normal or acid salts of pectic acids. “Protopectin” is applied to the water-insoluble parent pectic substance which occurs in plants and which upon restricted hydrolysis yields pectin or pectinic acids. Still, these definitions are rather vague. For example, there is no indication about the minimal polygalacturonic acid content required, nor about the minimal polymerization degree. Apart from the definitions based on composition and structure, there is also a definition based on composition and application of some pectic substances as gelifying agents. The general term “pectin” (or pectins) designates those water-soluble pectinic acids of varying methyl ester content and degree of neutralization which are capable of forming gels with sugar and acid under suitable conditions. Pectins with a rather high methoxyl contents show their jellying power only in the presence of a relatively high sugar and acid content, whereas gel formation by pectins with a lower methoxyl content is also possible without sugar in the presence of certain metallic ions. For this reason, two groups of jellying pectic substances are distinguished: the high-methoxyl pectins (>50%) and the low-methoxyl pectins. The term “high-methoxyl pectins’’ is often abbreviated to “pectins.” A schematic survey of the interrelationship of pectic substances is presented in Fig. 1. TABLE I NOMENCLATURE FOR PECTIC SUBSTANCES Pectic acid Polygalacturonic acid (pectate) Pectin Pectic acid partially esterified (pectinate] with methanol and containing some neutral sugars Pectic substance fixed in the plant Protopectin tissue

TAKUO SAKAI ET AL.

216

PROTOPECTIN I

I

1

+

Acid or enzymes

Warm alkali

Cold alkali

PECTATES

PECTATES

1

Nonfibrous, nonviscous Low molecular weight Insoluble due to alkaline earths from peel

PECTIN

I

Alkali, enzymes, or strong acid

Fibrous, viscous High molecular weight Insolubledue to alkaline earths from peel

I

I

Acid

PECTINATES

c

Strong acid

m enzymes

and I or pectinic acid

Mild acid

P

Nonfibrous Low molecular weight

Alkali

PECTIC ACID Fibrous High molecular weight

Cold alkali

i

Enzymes PECTATES Nonfibrous NH3 and alkali metal salts soluble

PECTATES Viscous, fibrous NH3 and alkali metal salts soluble

a-GALACTURONIC ACID

FIG.1. Interrelationship of the pectic substances (from Campbell and Palmer, 1978).

B. CHEMICAL CONSTITUENTS AND STRUCTURE 1. General Properties

Pectin is a heteropolysaccharide with galacturonic acid and methanol as the main components, with some neutral sugars attached. Pectin molecules are formed by a-l,4-glycosidic linkages between the pyranose rings of D-galacturonic acid units. The pyranose rings of D-galacturonic acid probably occur mainly in the chair L form, corresponding to the most stable conformation of D-galaCtOSe. As both hydroxyl groups of Dgalacturonic acid at carbon atoms 1 and 4 are on the axial position, the Some polymer resulting from such linkage is trans-1,4-po~ysaccharide. pectins, such as those from flax, tobacco, and sugar beet, also contain

217

PECTIN, PECTINASE, AND PROTOPECTINASE

some acetic acid. The carboxyl groups of pectin are partially esterified with methanol, and the hydroxyl groups are sometimes partially acetylated. Many pectin preparations contain other sugar units along with galacturonic acid, such as galactose, arabinose, or rhamnose, probably in accompanying polysaccharide linked as a side group to the polygalacturonic chain. Although the solubilization of pectic polysaccharides by water and their selective precipitation with alcohol was described by Vauquelin as early as 1790, the elucidation of their structure has proved particularly difficult and is still by no means completed (Cook and Stoddart, 1973). From a chemical point of view, pectin usually, and perhaps invariably, exists as a branched heteropolysaccharide in which the backbone is based on linear sequences of a-1,+linked D-galacturonate residues (Morris et a]., 1980). The carboxylic acid groups can be neutralized by mono- or divalent ions such as K + , Na+, and Ca2+ or may be partially esterified with methanol. Some of the hydroxyl groups on C, or C, may be acetylated (Fig. 2). The measure of esterification of pure galacturonic acids may be indicated by the methoxyl-(CH,O-)content or by the degree of esterification, i.e., the number of esterified carboxyl groups calculated as the percentage of the total number galacturonic acid units. When the carboxyl groups in pure polygalacturonic acids are all esterified, the methoxyl content is 16.32% and the degree of esterification is 100%. Esterification of pectinic acids, extracted from natural sources, is seldom higher than 70% (Table 11). Similarly, the acetylation can be expressed by the acetyl content or by the degree of acetylation (Table 11). The backbone, consisting of polygalacturonic acid, is also periodically interrupted by the insertion of a-L-rhamnopyranose residues. These residues are present in segments of the structure a-Dgalactopyranosyluronic acid-(l-+2)-a-~-rhamnopyranosyl-(l+4)-a-~galactopyranosyluronic acid (Lau et al., 1985; Aspinall and Cottrell, 1970; Aspinall et al., 1968a,b; Aspinall, 1981).The effect of the presence

H

OH

C4H

f-W 0

FIG.2. Hypothetical part of a pectin molecule (from Pippen et a ] . , 1950).

218

TAKUO SAKAI ET AL. TABLE I1

DEGREEOF ESTERIFICATION AND DEGREEOF ACETYLATION OF SOMEPECXICSUBSTANCES’ Source of pectin

Esterification (YO)

Acetylation (Yo)

Apple Potato Sugar beet Pear Mango Citrus fruits Sunflower

71 31

4 14 20 14 4

55 13 68 64 17

3 3

Data from Voragen et al. (1986)

of rhamnose is to cause a T-shaped “kink” in the chain (Fig. 3) (Rees and Wight, 1971; Rees, 1972; Albersheim, 1976). Neutral sugars other than L-rhamnose occur exclusively in side chains of pectins. D-Galactopyranose and L-arabinofuranose occur most frequently; D-xylopyranose, o-glucopyranose, and L-fucopyranose are less common units, while rarely found sugars like D-apiose, 2-O-methyl-Dxylose, and 2-O-methyl-~-fucoseare usually very minor, but widespread, constituents of pectins (Darvill et a]., 1978; Aspinall, 1981; Barrett and Northcote, 1965). The branching occurs through the C-2 (Ovodov et al., 1971) or C-3 atom (De Vries et a]., 1983; Aspinall et a]., 1968a) of galacturonic acid or through the C-4 (Aspinall and Fanous, 1984; Stevens and Selvendran, 1984) or C-3 atom (Darvill et al., 1978) of rhamnose. It was stated that

OH

FIG.3. The presence of rhamnose and the resulting T-shaped kinking of the pectin molecule (from Barford et al., 1986).

PECTIN, PECTINASE, AND PROTOPECTINASE

219

the rather rare sugars occur in short (one to three units) side chains, substituting the galacturonic acid skeleton, while arabinose and galactose form oligo- and polysaccharides substituting the rhamnosyl units. Also, the pectic polymers of primary cell walls have a relatively higher proportion of oligosaccharide chains on their backbone, and these side chains are much longer than those of the pectins of the middle lamellae (Selvendran, 1985). Enzymatic breakdown of pectic substances followed by analysis of the fractions revealed that these side chains were not distributed regularly along the galacturonan chain but concentrated in so-called “hairy regions,” leaving important parts of the backbone unsubstituted. Especially, the galacturonate residues in the hairy regions are esterified with methanol (Fig. 4) (De Vries et al., 1982; Rouau and Thibault, 1984; Axelos et al., 1989; Konno et a]., 1986; Thibault, 1983). Pectinic acids that lack neutral blocks will generally be referred to as “Type 11.” Detailed structure of a pectin is described in Fig. 5 (Cook and Stoddart, 1973). In the plant cell wall, the side chains link the pectin molecules to proteins, hemicelluloses, and cellulose to form the insoluble protopectin (Figs. 6 and 7).

Relatively short arabinan, galactan, or arabirogalactansidechains

-

Relatively Long arabinan, galactan or arabinogalactansidechains

Highly branched rharnnogalacturonan (primary cell wall)

FIG.4. Schematic representation of some structural aspects of pectins from the middle lamella (A) and primary cell walls (B) (from Selvendran, 1985).

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TAKUO SAKAI ET AL

FIG.5. Detailed structure of a pectin [from Cook and Stoddart, 1973).

It is also suggested that acidic and neutral pectins carry ferulic acid on the nonreducing ends of the neutral arabinose and/or galactosecontaining domains. The pectins carry approximately one feruloyl residue per 60 sugar residues. Possible roles of feruloyl pectin are in the regulation of cell expansion, in disease resistance, and in the initiation of lignification (Fry, 1983).

Rharnnogalacturonan

FIG.6. Molecular structure of the primary plant cell wall (from Adler-Nissen, 1987).

PECTIN, PECTINASE, AND PROTOPECTINASE

Extension helix

221

Cellulose microfibril

Extension nonhelical region

Xyloglucan latches

Intermolecular isodityrosine cross link

Pectin

FIG.7. Three-dimensional view of polymer arrangement in the plant cell wall (from Wilson and Fry, 1986).

The molecular mass of pectic substances from various sources has been the subject of many investigations. The reported values vary from about 10,000to 400,000 (Table 111).Apart from existing significant differences, the results are also influenced by the method of extraction and technique of measuring molecular mass.

TABLE 111 MOLECULAR MASSOF SOMEPECTICSUBSTANCES" Source

Molecular mass

Apple and lemon Pear and prune Orange Sugar beet pulp

200,000-360,000 25,000-35,000 40,000-50,000 40,000-50,000

a

Data from Fogarty and Kelly (1983).

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TAKUO SAKAI ET AL.

Finally, it must be clearly stated that there is no such thing as a uniform pectic substance. This is illustrated by the many variations in molecular mass; in degree of esterification and acetylation; and in quantity, type, and distribution of the non-uronide components. The pectic substances seem to be more heterogeneous than first assumed (Neukom et al., 1980). 2. Protopectin Protopectin was already defined as the “original” water-insoluble parent pectic substance which occurs in plants and which yields pectin or pectinic acids upon restricted hydrolysis. In most plant tissues, only this insoluble form of pectin occurs. The main exceptions to this rule are the ripe(ning) fruits. The many hypotheses about how insoluble protopectin is composed out of soluble pectic substances are summarized by Joslyn (1962).They include the reasons for the insolubility of protopectin: 1. The very large molecular weight of protopectin in comparison to that of pectin. 2. Mechanical enmeshing of the filamentous pectin macromolecules by one another and with other high polymers (cellulose, hemicellulose, lignin) of the cell wall. 3. Ester bond formation between the carboxylic acid groups of pectin and the (alcoholic) hydroxyl groups of the other cell wall constituents. 4. Lactone bond formations within the entangled pectin molecule. 5. Salt bonding between the carboxyls of pectic substances and basic groups of proteins. 6. Polyvalent ion bonding (Caz+,Mg2+,Fez+)between the carboxyls of the different cell wall constituents. 7. Secondary valence binding (i.e., H-bonds, hydration bonding, and molecular cohesion) between pectic substances.

c. OCCURRENCE AND FUNCTION Pectic substances are prominent structural constituents of (primary) cell walls in non-woody tissues, next to cellulose, several hemicelluloses, and protein (Fig. 8) (Brillouet, 1987). In addition, they are the sole polysaccharides in the middle lamella responsible for cell cohesion (Pilnik, 1981). Pectic polysaccharides occur mainly as water-insoluble protopectin. Their synthesis, beginning from UDP-D-galacturonic acid and taking place in the Golgi system (Karr, 1976), is performed mainly during the early stages of growth, in

PECTIN, PECTINASE, AND PROTOPECTINASE

223

: : Miwofibrils (organizedphase) : : ;Continuous matrix (amorphous phase)! I

,

I

,

2 a,

I

Primary wall

Secondary wall

Lumen

-5 BI U

FIG.a. Distribution of materials in the "mature" cell wall. The arrows indicate direction of increasing relative concentration (from Northcote, 1958).

young enlarging cell walls. Compared with young, actively growing tissues, lignified tissues are low in content of pectic substances. Furthermore, primary cell walls of graminaceous monocotyles have a low content of pectin compared to those of dicotyles (Jarvis et al., 1988). The average pectin content of several plant tissues is shown in Table IV. Texture of vegetables and fruits is strongly influenced by the type of pectin present. One of the most characteristic changes during the ripenTABLE IV PECTINCONTENT OF SEVERAL TISSUESO Tissue

Pectic substances (%)

Apple Bananas Peaches Strawberries Cherries Peas

(fresh) (fresh) [fresh) (fresh) (fresh) (fresh) (dry matter) Carrots Orange pulp (dry matter) (dry matter) Potatos Tomatos (dry matter) Sugar beet pulp (dry matter] Data from Chenoweth and Leveille (1975).

0.5-1.6 0.7-1.2 0.1-0.9 0.6-0.7 0.2-0.5 0.9-1.4 6.9-18.6 12.4-28.0 1.8-3.3 2.4-4.6 10.0-30.0

224

TAKUO SAKAI ET AL.

ing of fleshy fruit is softening. This change is attributed to enzymatic degradation and solubilization of the (proto) pectic substances (Labavitch, 1981; Soda et al., 1986; Pressey, 1988; Barbier and Thibault, 1982; Dick and Labavitch, 1989). However, during ripening the neutral sugar composition of the extractable pectin does not change (De Vries et al., 1981). In processing certain vegetables, for example, cauliflower, excessive softening is prevented by adding Ca salts. As a consequence, insoluble pectates and pectinates are formed, giving a firm texture to the vegetables. Similarly, the mechanical properties of cossettes cut from sugar beet can be improved by adding lime to the diffusion water. The addition of lime to sugar beet tissue at lower temperatures causes demethylation of the pectin in the cell walls of the beet tissue, allowing Ca2+to cross-link the pectin as a stable insoluble matrix. This permits alkaline diffusion with less disintegration of the pulp (Camirand et a]., 1981).

D. PROPERTIES Pectic substances are insoluble in most organic solvents. They do dissolve in water, dimethyl sulfoxide, formamide, and (warm) glycerol. The solubility in water decreases with increasing polymerization degree. Solubility is increased by all factors diminishing possibilities of intermolecular association. These factors can be of a sterical ( e g , the presence of substituents) or a chemical (e.g., charges) nature. Mostly, solubilization is proceeded by a slow swelling. Aqueous solutions of 1 to 2% (w/v) already have a relative high viscosity. This viscosity is proportional with molecular mass and is also influenced by degree of esterification (Pippen et a]., 19531, ionic strength, pH, and temperature. Depending on their degree of esterification, pectic substances are precipitated from aqueous solutions with water-miscible organic solvents or with cations. In acid solutions, the degree of esterification and/or the polymerization degree decreases. Deesterification is dominant at low temperature, whereas a high temperature enhances depolymerization. Also, the neutral sugar content decreases as these side chains are more sensitive to acid. On the average, the neutral sugar content decreases compared to the rhamnose content. The increase in the relative amount of rhamnose compared with other sugars in the heated tissue indicates possible degradation in the “hairy region.” In alkaline solutions, at low temperature, saponification of the methyl ester groups occurs readily. However, depolymerization is strongly enhanced by a rise in temperature. Such high alkali sensitivity is unique as polysaccharides are usually alkali resistant. Even more

PECTIN, PECTINASE, AND PROTOPECTINASE

225

significant is that degradation is not the result of hydrolysis of the glycosidic bonds in the classical manner but rather the result of a pelimination cleavage of glycosidic linkages (Neukom and Deuel, 1958). This reaction only occurs at glycosidic bonds adjacent to an esterified carboxyl group (Fig. 9). Pectates are indeed very much more stable at high temperature toward alkaline or neutral degradation than pectinates (Albersheim et al., 1960). Cross-linked pectin chains form insoluble polymers having ion-exchange properties. They are very selective for calcium and heavy metal ions, e.g., Zn2+,Cu2+,and Fe3+.The most unique and outstanding physical property of pectins is their ability to form gels with sugar and acid. Gels can be divided into two groups: Those containing a rather high sugar content (60-70%) and those with a lower sugar content. In the case of the former, high-methoxyl pectins (>50%) are used, while low methoxyl pectins are used in the case of the latter. Highly esterified pectin gels are obtained when, besides a sufficient high concentration, two other conditions are met: (1)Electrostatic repulsion between pectin molecules has to be decreased by repressing dissociation of carboxyl groups. Consequently, pH will play a determining role, and (2) Sugar, e.g., sucrose, or a similar carbohydrate, e.g., polyalcohols, is present in sufficient amounts. High polyol concentration decreases water activity leading to interchain interactions (Michel et a]., 1984). The hypothetical structure of a pectin-sugar gel is shown in Fig. 10. For preparations of a low ester content, gels are usually formed by the controlled introduction of calcium ions. According to Harvey (1960) the presence of calcium promotes the formation of gels by forming strong ion associations with carboxyl groups of neighboring pectin chains (“salt bridges”). However, Morris et al. (1980, 1982) have shown that the primary mechanism of this gelation involves extended chain sequences which adopt a regular twofold conformation and dimerize with specific interchain chelation of Caz+ (“egg-box” binding). Each Ca2+ ion takes part in nine coordinative links with an oxygen atom (Fig. 11).

H FIG.9. P-Elimination in a uronic acid unit in pectin [from Albersheim et al., 1960).

TAKUO SAKAI ET AL.

226

H I

H molecule

/O.

'H,

/H 0'

'H-0-

I

H H FIG.10. Hypothetical structure of a pectin-sugar gel [from Doesburg, 1965).

Apart from the degree of esterification, pH, and concentration of sugar or acid, the presence of side chains and/or groups, the degree of polymerization, the temperature, and the presence of ions also play an important role in gel formation. However, gelation of sugar beet pectins is greatly inhibited by the presence of acetyl groups. These substituents change the surface structure of the polymer (Solms and Deuel, 1951), cause sterical hindrance, and prevent the free carboxylic acid groups to form H bonds (Pippen et al., 1950). However, the presence of feruloyl

t

FIG.11. Pectins of low methoxyl content showing egg-box binding (0, non Caz+-bound oxygen atom; 0 , Caz+-bound oxygen atom) (from Thibault, 1980).

227

PECTIN, PECTINASE, AND PROTOPECTINASE

groups at the end of the neutral sugar side chains offers a third way for a gelling process, in addition to classical calcium gels of “low methoxyl” pectins and sugar acid gels of “high-methoxyl” pectins (Thibault, 1986). Indeed, sugar beet pectins can be cross-linked through their feruloyl groups and produce gels if the pectin concentration is greater than about 1%. Ammonium persulfate (Thibault et al., 1987; Thibault, 1986) and peroxide/peroxidase (Rombouts and Thibault, 1986a,b) are effective agents for this cross-linking reaction (Fig. 12). In conclusion, it can be stated that many of the unique physical properties of pectic substances are chiefly associated with the carboxyl

vH3 rvH3

a

OH

+

OH

w2- p H 3 F ___)

*

CH CH

HO-CH

HO

\

I

CH’

COOP

COOP

I

cCH

‘7’4 n

COOP

HO

OH

OH

r v H OCH, 3 @

CH \ CH

xc: -I

I

COOP

CH

AOOP

(2) FIG.12. Cross-linking of sugar beet pectins. (a) by ammonium persulfate (from Thibault et al., 1987). F, ferulate or feruloyl; P, H or pectic chain. (b) by hydrogen peroxidel peroxidase (from Markwalder and Neukom, 1976).

228

TAKUO SAKAI ET AL.

group of the galacturonic acid residues. Complete esterification of commercial pectin by chemical means totally modifies its acidic, viscosity, and gel-forming properties. For example, changes in pH have no significant effect on viscosity of solutions of totally esterified pectins. E. DETERMINATION AND CHARACTERIZATION

It was already stated earlier that pectin is a complex heteropolysaccharide with properties depending on a (varying) composition. As indeed the properties of pectin are strongly influenced by its composition, numerous techniques for analysis of pectin were developed. Pectins are usually characterized by (1)their content or uronide material, (2) their degree of esterification (DE), and (3) their degree of polymerization (DP) or some quality connected with it (viscosity, gel strength). The measurements usually made to express these characteristics give average values only (Van Deventer-Schriemer and Pilnik, 1976). On some occasions, the following analyses are also performed: (1)degree of acetylation, (2) neutral sugars, and (3) jellying power. 1. Anhydrogalacturonic Acid (AGA] Content Determinations of pectic substances in situ are qualitative, mostly by use of staining agents. For example, ruthenium red is used to obtain a molecular visualization of pectin (Hanke and Northcote, 1975). In dealing with preparations of pectic substances, the determination of polygalacturonide content is most important. In determining this AGA content, four methods have been widely used. The first method used was the decarboxylation method according to Lefevre and Tollens (1907). When uronides are boiled with hydrochloric acid (12%) they react as C,H,O$j + C,H,Oz

+ COZ + 2HzO

Then, furfural or CO, is determined gravimetrically. This method was modified by Whistler et al. (1940) and McCready et al. (1946), while Vollmert’s (1949) method, using hydroiodic acid for the decarboxylation, permits simultaneous determinations of uronide and methoxyl contents. Among colorimetric methods, the carbazole-sulfuric acid method of Dische (1950) and modified by McComb and McCready (1952), McCready and McComb (1952), and Furutani and Osajima (1965) is used most often. For estimating uronic acids in chromatographic fractions, this reaction is the most satisfactory method, but 2 hr are required for the full

PECTIN, PECTINASE, AND PROTOPECTINASE

229

development of color and, with certain compounds, the color is partially suppressed by salts (Bitter and Muir, 1962). At present, the titrimetrical method of Deuel (1943) is not frequently used any more because of interference by minerals or free acids. However, the necessary reagents and equipment are inexpensive. The most recent method [Blumenkrantz and Asboe-Hansen, 1973) is based on the appearance of a chromogen when uronic acids, heated to 100°C in concentrated sulfuric acidltetraborate, are treated with metahydroxydiphenyl. This method has been automated by Thibault (1979) and modified further by List et al. (1985). Ahmed and Labavitch (1977) used this procedure to determine uronide content of plant cell walls. 2. Degree of Esterification (Me0 content)

Here, titration (with alkali) is less problematic because the determination is carried out in two steps: titration before and after saponification. Also, methanol released on alkaline deesterification can be determined, either colorimetrically [Wood and Siddiqui, 1971; Klavons and Bennett, 1986) or gaschromatographically (McFeeters and Armstrong, 1984). The latter also used their method to measure the methoxylation of pectin in situ. The oxidation-reduction method of Laver and Wolfrom (1962) relies on the conversion of Me0 to methyl iodide. The neutralization and gas chromatographic methods have major advantages of simplicity and shortness of time of completion over the hydroiodic acid method (Walter et a]., 1983). Recent developments include 'T-NMR [ Fishman et al., 1984;Grasdalen et al., 1988) and analytical pyrolysis techniques (Barford et al., 1986). 3. Degree of Acetylation To determine the degree of acetylation by titration, the pectin sample (dissolved) has to be saponified and either steam distilled (Pippen et al., 1950) or extracted with an immiscible solvent (e.g., butanol:chloroform, 4 : 1) (Kertesz and Lavin, 1954). The reaction between esters and hydroxylamine to produce hydroxamine acids has also been applied successfully for the analysis of acetyl content in pectin (McComb and McCready, 1957) [Fig. 13). Pectin hydroxamine acid forms an insoluble complex with ferric ions and acetohydroxamic acid forms a soluble red complex. After filtration, the intensity of the red color is determined colorimetrically (at 520 nm). The Hestrin method [Downs and Pigman, 1976) is based on the same reaction. A procedure for the simultaneous quantitative analysis of methoxyl and acetate groups in pectin has been developed, using HPLC on a

1

-0

1

0 It

C-O-CH,

r

0 II

C-N

,H

+z?

+ 2n CH3CONHOH + n CH3W

0-

H

0 I O=C-CH3 _In

Acetylated pectin

n

L

Pectin hydroxamic acid

FIG.13. The hydroxamine acid reaction (from McComb and McCready, 1957).

Acetohydroxamic acid

PECTIN, PECTINASE, AND PROTOPECTINASE

231

cation-exchange resin in the protonated form and refraction index detection (Voragen et a]., 1986). 4. Neutral Sugars

Mostly, pectic substances are hydrolyzed to component sugars and, after conversion to alditol acetates of silylates, these are determined by gas chromatography (Blakeney et al., 1983; Voragen et al., 1983). 5. Molecular Mass

Generally,the estimation of the degree of polymerization or the molecular mass is the most difficult problem in the analysis of pectic substances. The results of estimations of the number of reducing end groups have been shown to be unreliable, since these results are affected by minute amounts of ballast materials. Viscosimetry has been used most frequently to determine molecular mass. However, it must be taken into account that the viscosity of a solution of pectinic acids depends on such things as molecular mass, concentration, degree of esterification, pH, and presence of electrolytes. Christensen (1954) calculated M, from viscosity measurements on commercial high-methoxyl pectins: based on the intrinsic viscosity [q],found by extrapolating qsp/c (qsp = specific viscosity, c = concentration) to c = 0.This method has been modified by Smit and Bryant (1967) to determine the M, from one measurement (of viscosity). To eliminate the complications caused by the complex colloidal properties of pectin, Schneider and Fritschi (1936) converted the pectinic acids into water-insoluble nitropectins and used acetone as the solvent for the viscosity measurements. Other techniques, although less applicable by routine, include ultracentrifugation, electron microscopy, osmometry (Jordan and Brant, 1978), light scattering (Chapman et al., 1987; Jordan and Brant, 1978; Sorochan et al., 1971), gel filtration (Anger et a]., 1977), HPSEC (Sjoberg, 1987; Fishman et al., 1989), and HPLC (Strubert and Hoverman, 1978). 6. Jellying Power

The jellying power is the most important property of pectins and, consequently, is used to grade commercial pectins. There are numerous devices for testing the consistency of jellies. Results obtained with such instruments are, however, usually expressed in arbitrary units as it is not possible to calculate the rigidities directly, i.e., in absolute units (Campbell, 1938). The jellying power of pectins is usually measured by estimating the strength of gels which have been prepared under accurately described conditions. The methods of deter-

232

TAKUO SAKAI ET AL.

minating gel strength can be divided into two large groups (IFT Committee, 1959). This was done to devise standard methods for the determination of the grade strength of pectins as commercially supplied for jam manufacture (Report of the Pectin Subcommittee, 1951). Methods belonging to a first group quantify the jellying power by measuring a controlled deformation of the jellies within their limit of elasticity. A simple apparatus, the rigidometer, has been developed for determining jelly grades of commercial pectins. It measures the modules of rigidity of gels (Owens et a]., 1947). Cox and Higby (1944) determined the percentage sag, or slump, occurring when a test jelly is removed from its supporting container and inverted upon a glass plate. The “BLOOM gelometer” measures the force required to push down the gel surface for 4 cm. In second group method of measurement, the elastic limits of the jellies (the “breaking strength”) are exceeded and the jellies ruptured. For example, the “Delaware jelly tester,” developed by Tarr (1926) and Baker (1926), measures the force required to push down a gel surface until the gel breaks. The jellying power of high-methoxyl pectins is described in relation to their sugar carrying power. The jelly grade is the number of parts (by weight) of sugar that one part of pectin will convert to a jelly under standard conditions. F. PECTIN MANUFACTURING: CHEMICAL EXTRACTION AND PURIFICATION

Data concerning pectin consumption are given in Table V. Today, the chief raw materials for the production of pectin are by-products from the manufacture of fruit juices: apple pomace (dried) and citrus residues (peel). The raw materials for pectin production are wastes from other operations, and the quality of the pectin produced is often determined by physical and chemical operations in the primary industry (Charley, 1951). TABLE V CONSUMPTION, PRICES, AND MARKETS OF PECTIN‘ Consumptionb (tondyear)

Prices

Product Low methoxyl pectin High methoxyl pectin

6000 8000

7.15-11.00 7.92-8.80

a

Data from Yalpani and Sandford (1987). For the United States, during 1983-1985

($W

Markets

(lo6 $/year) 22

PECTIN, PECTINASE, AND PROTOPECTINASE

233

Until recently, chemical extraction has been the only way to produce pectin. This extraction is performed by acid hydrolysis. Conditions vary but generally a pH in the range of 2.0-3.0 is used for 0.5-5 hr within a temperature range of 70-100°C. The solid to liquid ratio is normally about 1: 18. In some countries the use of mineral acids is prohibited and these are replaced by citric, lactic, or tartaric acids. The pectin extract is separated from the pomace using hydraulic presses and/or centrifugation. Sometimes, gelatinization of starch takes place and this necessitates an enzymatic treatment with amylases. Subsequently the extract is filtered again and then finally concentrated to a standard setting strength. In preparation of powdered pectins the concentrated liquor is treated with organic solvents or certain metallic salts to precipitate the polymers. The pectin precipitate is collected, dried, and ground. Commercial pectins are standardized products. This is done to ensure that the users always get the same gel strength. Standardization can affect the chemical structure of pectin as esters can be partly saponified or acid groups can be amidated. High-methoxyl pectin only forms gel above a soluble solids (sugar) content of about 55%. Low ester pectin forms gels in the presence of (calcium) ions, irrespective of soluble solids. A scheme of the commercial production of pectin is represented in Fig. 14.

G. APPLICATIONS OF PECTIN 1. In the Food Sector

Pectin is first and foremost a gelling agent (E440) and is used to give a gelled texture to foods, mainly fruit-based foods. About 80% of the world production of high-methoxyl pectin is used in the manufacture of jams and jellies, to make up for their “deficiency” of natural pectins. Indeed, under these conditions, i.e., a high sugar concentration and a low pH, it is the best gelifying agent available (Nelson et al., 1977). Pectin establishes a texture that retains a uniform distribution of fruit particles during transportation, gives a good flavor release, and minimizes syneresis. The pectin concentrations used vary from 0.1 to 0.4% in jams and jellies (Pectin, 1987). Low ester pectins are often used in fruit preparations for yogurt in order to create a soft, partially thixotropic gel texture, sufficiently firm to ensure uniform fruit distribution, but still allowing the fruit preparation to be easily stirred into the yogurt. The pectin may further reduce-especially when combined with other plant gums-color migration into the yogurt phase of the final product.

TAKUO SAKAI ET AL.

234

I

1

I

1 Treat with acidified isopropanol I

&

I

Rinse with is0 ro anol

in isopropanol

1-

+

(Grind to pass 60 mesh screen

1

Standardize, package, and sell as slow-setting pectin (HM)

FIG.14. Production and standardization of pectin (from Nelson et al., 1977).

Gelation (by pectin) also provides stabilization of emulsions, suspensions, and foams. This is demonstrated in the fruit drink concentrates. In recombined or instant juice products, pectin gelation restores the sensorial properties to those of the fresh juice. In dairy products, the pectin reacts with the casein, preventing the coagulation of the casein at a pH below the isoelectric pH (4.6) and allowing pasteurization of the sour milk products to extend their shelf life. Another application of pectin is in confectionary fillings. Also, the possibility of using pectin for the production of single cell protein in a modified “Symba process” was reported (Fellows and Worgan, 1986, 1987a,b).

PECTIN, PECTINASE, AND PROTOPECTINASE

235

2. In the Pharmaceutical Sector

In a number of (liquid) pharmaceutical preparations the ability of pectin to increase viscosity and stabilize emulsions and suspensions is utilized. Pectin, belonging to the chemically heterogeneous group of substances referred to as “dietary fiber” (Aspinall and Carpenter, 1984), is further reported to possess a number of valuable biological effects. It acts as a general “intestinal regulator” and a detoxifying agent, but the most well known effect is its anti-diarrhea effect (Chenoweth and Leveille, 1975). This probably explains the ancient use of the diet of scraped apples as a home remedy against diarrhea (Birnberg, 1933). Indeed, “an apple a day keeps the doctor away.” There are several hypotheses to explain these effects. It is suggested that pectin, or its degradation products, moves rapidly to the large intestine and exerts a bacteriostatic effect against several pathogens (Werch and Ivy, 1941; Campbell and Palmer, 1978). Pectin, being a colloidal carbohydrate, acts as a lubricant in the intestines, coating the mucosa with uncharged polysaccharide and promotes normal peristalsis without causing irritation. This makes it a standard additive to baby foods. The detoxifying action is probably a consequence of the binding of metal ions to pectin (fragments of) (Kohn, 1987). Pectin could also be used as a carrier for pharmaceuticals (Heinzler et al., 1987). It also decreases the toxicity of pharmaceuticals (some) and prolongs their activity without lessening their therapeutic effect (Pilnik and Voragen, 1970). More recently, pectin-gelation microglobules were developed for potential use in regional cancer chemotherapy as an intravascular biodegradable drug delivery system (Bechard and McMullen, 1986). Pectic substances also seem to show hemostatic and antifibrinolytic effects (Barth and Rumpelt, 1947; Fogarty and Kelly, 1983). Work has also been done to show that pectin administered orally is somewhat effective in reducing cholesterol levels in the blood. Probably, the absorption of bile acids is decreased and, as a consequence, more cholesterol has to be converted to bile acids (Baig et al., 1980; Kay and Truswell, 1977; Keys et al., 1960; Lin et al., 1957; Pfeffer et al., 1981). It is hoped that the use of pectin-enriched foods will aid in the prevention and treatment of arteriosclerosis (Panchev et al., 1989). 3. The Cosmetical Sector The applications in the cosmetical sector only utilize the “ordinary” properties of pectin. Examples are the numerous gels (hair) and pastes.

236

TAKUO SAKAI ET AL.

Ill. Classification of Pectic Enzymes

Basically three types of pectic enzymes exist: pectinesterase, which only removes methoxyl residues from pectin, a range of depolymerizing enzymes (pectinase), and protopectinase, which solubilizes protopectin to form pectin (Table VI). Pectinases are distinguished under three headings (Enzymic Commission, 1944) according to the following criteria: (1)Whether pectin, pectic acid, or oligo-D-galacturonate is the preferred substrate, (2) Whether they act by transelimination or hydrolysis, and (3) Whether the cleavage is random (endo-, liquefying, or depoly-

TABLE VI CLASSIFICATION OF PECTICENZYMES

Pectinesterase (PE) (Pectin methylhydrolase, EC 3.1.1.11) Catalyzes deesterification of the methoxyl group of pectin forming pectic acid. Depolymerizing enzymes Enzymes hydrolyzing glycosidic linkages: Polymethylgalacturonase (PMG) Endo-PMG, causes random cleavage of a-l,4-glycosidic linkages of pectin, preferentially highly esterified pectin. Exo-PMG causes sequential cleavage of a-1,4-glycosidic linkages of pectin from the nonreducing end of the pectin chain. Polygalacturonase (PG) glycanohydrolase], catalyzes Endo-PG [EC 3.2.1.15, poly(l,4-a-D-galacturonide) random hydrolysis of a-l,4-glycosidic linkages in pectic acid (polygalacturonic acid). Exo-PG [EC 3.2.1.67, poly(l,4-a-~-galacturonide) galacturonohydrolase], catalyzes hydrolysis in a sequential fashion of a-1,4-glycosidic linkages in pectic acid. Enzymes cleaving a-1,4-glycosidic linkages by transelimination which results in galacturonide with an unsaturated bond between C, and C, at the nonreducing end of the galacturonic acid formed. Polymethylgalacturonate lyase (PMGL) Endo-PMGL [EC 4.2.2.10, poly(methoxygalacturonide)lyase], catalyzes random cleavage of a-l,4-glycosidic linkages i n pectin. Exo-PMGL, catalyzes stepwise breakdown of pectin by transeliminative cleavage. Polygalacturonate lyase (PGL) Endo-PGL [EC 4.2.2.2, poly(l,4-a-D-galacturonide)~yase], catalyzes random cleavage of a-1,4-glycosidic linkages in pectic acid by transelimination. catalyzes sequential Exo-PGL [EC 4.2.2.9, poly(l,4-a-o-ga~acturonide)exo~yase], cleavage of a-1,4-glycosidic linkages in pectic acid by transelimination. Protopectinase The enzyme solubilizes protopectin forming highly polymerized soluble pectin.

237

PECTIN, PECTINASE, AND PROTOPECTINASE

merizing enzymes) or end-wise (exo- or saccharifying enzymes). The modes of action of the different types of pectic enzymes are illustrated in Fig. 15. However, no distinction is made between endo- and exoenzymes. The cup-plate method of Dingle et al. (1953) is a frequently used qualitative method for detecting microbial pectinase activity. A supernatant of a microbial culture is inserted into wells cut in an agar-pectin gel. If zones of activity appear around the cups, one may expect pectinase activity. A bioassay specific for polygalacturonase (PG) activity is described by Mussel1 and Moore (1969).This assay is based on fresh weight loss of cucumber pericarp tissue. Hildebrand (1971)tested many Pseudomonas sp., and also other plant pathogens, for pit formation on polypectate gels and used the results in the differentiation of the species. Another method for detecting polygalacturonase activity uses ruthenium red staining of colonies on polygalacturonate-agarose plates. Ruthenium red was shown to penetrate beneath the surface layers of the gel only in the regions surrounding a colony where degradation of polygalacturonate had occurred (McKay, 1988). A similar procedure is suitable in locating pectic enzymes in polygalacturonate-agarose overlays into which pectic enzymes diffuse from electrophoresis gels (Collmer et a]., 1988).

Protopectin

1

Protopectinase

-oo eozofi -06 Polymethylgalacturonate(Pectin)

O H H

H

I

OH

OH

OH

OHOH

O H

H O H

CoocHi

Polymethylgalacturonate lyase

H

c-3

I

-

+@

cooa4

w n

o n

cooa4

M

Polyrnethylgalacturonase

Pectinesterase

1

Polygabcturonate (Pectic acid) H

CM

-

0

H

+ m

0

OH

ic

Q

n

aa

M

o

o

-

-

-oQ

O

rm

H

u r n

O

OH

o

0

O H

Cm

H

l i b 4

Polygalacturonate lY-

=

o

O H

im

Q

+ H

OH

M

H

FIG.15. Mode of action of pectic enzymes.

o H

M

! i Q

Polygalacturonase

n

too

238

TAKUO SAKAl ET AL.

However, these methods can only be used for the quantitative determination of one specific enzyme. A. ESTERASES Pectinesterases (PE) are formed by fungi, bacteria (Table VII), yeasts, and higher plants. Pectinesterases, especially those from higher plants, are highly specific enzymes. In many cases they saponify almost exclusively the methyl ester groups of pectic substances. McDonnell et al. (1950) examined several fungal enzymes and found that rates of hydrolysis of the ethyl ester of pectic acid range between 6 and 16% of the rates obtained with the natural methyl ester. Some PEs attack pectin only at the reducing chain end while others attack the nonreducing end (Miller and Macmillan, 1971). Also, PEs seem to hydrolyze only methyl ester groups adjacent to free carboxyl groups. The enzymes then proceed in a linear fashion along the substrate. The result is a blockwise distribution of free carboxyl groups and esterified carboxyl groups (Fig. 16) (Speiser and Eddy, 1946). This also accounts for the high calcium sensitivity of enzymatically deesterified pectinic acids as compared with pectinic acids saponified in acid or alkaline milieu. In some cases, Caz+ (and also Na+) stimulate PE activity. In binding the pectic acid ( = reaction product), Ca2 prevents the PE from binding with it which would result in inhibition of the enzyme. It has also been reported that incomplete hydrolysis of methyl esters by PE can be caused by the inhibition of enzyme activity by the side chains of neutral sugars in the pectin molecules (Matsuura, 1987). The synthesis of certain pectinesterases (e.g., some Aspergillus niger strains) is repressed by glucose, even in the presence of the inducer (Maldonado et al., 1989). pH values at which PEs are active range from 4 to 8. The pH optimum for PE activity of fungal origin is generally lower than that of bacteria. Care must be taken not to confuse nonenzymatic deesterification with enzymatic activity at pH values above 7.0. The activity of these enzymes can be followed by determinating the increase of free carboxylic acid groups or by measuring the liberation of methanol into the solution. Free carboxyl groups can be measured by titration (Kertesz and Lavin, 1954). This has been simplified by the introduction of continuous, automatic titrations (Hagerman and Austin, 1986). Forster (1988) defines one unit of enzyme activity as the amount of enzyme which causes a decrease in pH of the reaction mixture of 0.1 in 30 min. The methanol may be distilled off and estimated by oxidation to formaldehyde as described by Holden (1945). Methanol can also be +

239

PECTIN, PECTINASE, AND PROTOPECTINASE TABLE VII OCCURRENCE OF PECTIC ENZYMESIN SOMEMICROORGANISMS~ ~

Source

PE

PG

Bacteria

Bacillus sp. Bacillus sp. No. RK9 Bacillus subtilis Bacillus polymyxa Bacillus purnilus Bacillus sphaericus Bacillus stearothermophifus Erwinia aroideae Erwinia carotovora Pseudomonas sp. Pseudomonos ff uorescens Pseudornonas marginalis Xanthomonas sp. Xanthomonas campestris Xanthornonas cyanopsidis Clostridiurn multiferrnentans C1ostri di urn aumntibutyricurn Clostridium felsineurn Cytophaga johnsonii Cytophaga deprimata Cytophaga albogilva Streptornyces nitrosporeus FUlgi Trichoderma koningii Trichoderrna pseudokoningii Cercospora orachidicoia Cephalosporiurn sp. Aspergillus niger Aspergillus sojae Aspergillus saita Fusarium culrnorum Fusariurn oxysporurn Fusariurn solani Penicillium expansurn Penicilliurn italicum Penicilliurn digitaturn Penicillium chrysogenum Rhizoctonia fragariae Ahizoctonia solani Ahizopus arrhizus

+ + +

+ + + +

+

+

+

+ + + + + +

+ +

+

+ + +

PGL

PMG

+ + + + + + + + + + + + + + + + + + + + + +

+

PMGL

OG

OGL

i

+

+

+ + +

+ + +

+ + +

+

+ +

+

PE, pectinesterase; PG, polygalacturonase;PGL, polygalacturonatelyase; PMG,polymethylgalacturonase; PMGL. polymethylgalacturonate lyase; OG, oligogalacturonase;OGL, oligogalacturonide lyase. (Data from Fogarty and Kelly, 1983.)

240

TAKUO SAKAI ET AL.

COOCH, COOCH, COOCH, COOCH, COOH I

1

I

I

I

COOH

COOH

COOH

I

I

I

FIG.16. Mode of saponification by pectinesterase (from Doesburg, 1965).

measured chromatographically (McFeeters and Armstrong, 1984) or colorimetrically (Wood and Siddiqui, 1971). B. HYDROLASES

1. Endopolygalacturonases (Endo-PG) These enzymes are produced by numerous fungi and bacteria (Table VII), by a few yeasts (Luh and Phaff, 1951; Ravelomanana et al., 1986), and also by higher plants and some plant-parasitic nematodes (Riedel and Mai, 1971). In general, by the action of PG, pectic acid is broken down into mono-, di-, and trigalacturonic acid. These end products may be produced by a “single chain multiple attack” mechanism, in which case they can be detected rapidly, or by a “multi-chain attack” mechanism, where the mono-, di-, and trimers accumulate only after further hydrolysis of the initial depolymerization products (higher oligogalacturonates) (Fogarty and Kelly, 1983). The former reaction is performed by the PG from Colletotrichum lindemuthianum (English et al., 1972; Albersheim, 1976) while the latter is in agreement with the action pattern of Kluyveromyces fragilis (Phaff, 1966). Endopolygalacturonases are specific for pectic acid. If the degree of methoxylation increases, the rate and extent of hydrolysis decreases. Free carboxyl groups seem to be necessary for catalytic activity (Jansen and McDonnell, 1945; Koller and Neukom, 1969). The rate of splitting of the glycosidic bonds also decreases with the shortening of the substrate chain. Still, many PGs are able to degrade the trimer, at a much lower rate (Rexova-Benkova and MarkoviC, 1976). Three different patterns of action toward low molecular substrates are known. The character of the active center constitutes the determining factor. Generally, endo-PGs are optimally active at a rather low pH (4.0 to 6.0) and at a temperature of 30-40°C. Kaji and Okada (1969) even described an endo-PG with an optimum for catalytic activity at pH 2.5. Other specific properties of the enzyme, as for example the need for coenzymes, have not been reported. Calcium ions influence the activity of polygalacturonase. However,

PECTIN, PECTINASE, AND PROTOPECTINASE

241

in some cases the activity was inhibited, while in other cases it was stimulated (Perley and Page, 1971). PGs can be produced constitutively (Wimborne and Rickard, 1978) or inducibly (Bhaskaran and Prasad, 1971; De Lorenzo et al., 1987). In other cases, the activity is only slightly enhanced (Bashan et a]., 1985). Induction or stimulation is mostly caused by low concentrations of pectins or oligo- and monomeric fragments thereof (Cooper and Wood, 1975). Brookhouser et al. (1980) reported that some microorganisms (e.g., Rhizoctonia solani) produce different froms of endo-PG and that the predominant form produced during pathogenesis differs from the single-peak form produced in culture. Some PGs are sensitive to catabolic repression (Horton and Keen, 1966; Keen and Horton, 1966; Hsu and Vaughn, 1969; Maldonado et al., 1989). Other are inhibited in vivo, mostly by a protein present in the host (Barmore and Nguyen, 1985; Collmer and Keen, 1986; Mahadevan et a]., 1965), sometimes by tannins or phenolic compounds present in the host tissues (Prasad and Gupta, 1967). The formerly mentioned inhibition by a protein is competitive. The activity of PG is also influenced by substitutions on the pectic acid molecule (Jansen and McDonnell, 1945). Over the years, mainly two methods for the determination of PG activity have been developed. Methods of a first group are based on the determination of reducing groups, released as a consequence of substrate hydrolysis by PG. The colorimetric method by Nelson (1944) or the iodometric method by Jansen and McDonnell (1945) can be used. However, viscosity measurements have also found widespread use for determinating pectinase activity (Cappellini, 1966; Bateman, 1972)

% reduction in viscosity = To - TJT,

-

T,,

where To,T,, and T, represent the flow time (in a capillary viscosimeter) in seconds for the reaction mixture without enzyme, the test mixture, and water, respectively. However, viscometric methods have met with limited success. This is due to the requirements for strictly standardized conditions, since the viscosity of solutions of pectic substances depends on pH, temperature, buffer, and ionic strength. The unit of enzyme activity is mostly selected as that amount of enzyme required for attaining a certain decrease of viscosity per unit time. 2. Exopolygalacturonases (Exo-PG)

Exopolygalacturonases occur less frequently. They are produced by fungi and some bacteria. Two types of exo-PG can be distinguished. Fungal exo-PGs (e.g., from Coniothyrium diplodiella) produce monogalacturonic acid as the main end product and have pH optima from 4.0

242

TAKUO SAKAI ET AL.

to 6.0. This enzyme is called galacturan 1,4-a-galacturonidase or exoPG 1. Bacterial enzymes, however [e.g., from Erwinia aroideae or Selenomonas ruminantium (Heinrichova and Wojciechowicz, 1989)], produce digalacturonic acid as the main end product. They are mostly designated as exo-poly-a-galacturonidase or exo-PG 2. Both enzymes, however, degrade pectic acid from the nonreducing end (Fig. 17). With respect to other, general characteristics (e.g., constitutivity, inhibition, and repression), exo-PG resembles endo-PG. Here too, the increase of the number of reducing groups can be followed to determine enzyme activity. Exopolygalacturonase activity can be detected more specifically on the basis of the release of digalacturonic acids. This can be done by spotting samples on TLC plates and developing them with a benzidine solution (Gothoskar et al., 1955). Exo- and endo-PG activity can be differentiated by measuring both increase of reducing groups and reduction of viscosity. An endo-enzyme is characterized by a strong reduction in viscosity (e.g., 50%) without a significant release of reducing groups ( 2 4 % ) .To obtain a 50% viscosity reduction, an exo-enzyme has to hydrolyze 20% of the glycosidic linkages (Nasuno and Starr, 19681. 3. Oligogalacturonases (OG) An oligogalacturonate hydrolase, free of transeliminase and unsaturated oligogalacturonate hydrolase activities, has been isolated from the cell extract of a Bacillus species by Hasegawa and Nagel (1968). The pH optimum is 6.0-6.5. The hydrolase is highly specific for saturated oligogalacturonides, attacking it from the nonreducing end of the molecule. A similar activity has been isolated from the mycelium of Aspergil-

1

Exq-PG 2 ( E )

Exo-PG 1 (C)

0-0+o-o+e-

No reaction

4 O

O

-

4

Exo:PG 1 (C)

O

+R

~

O

-

R R

ExoiPG 2 ( E )

O

f R ~

FIG.17. Action of exo-PG of Coniothyrium diplodieiia (C) and Erwinia oroideae (E) on saturated pectic acid (from Fogarty and Ward, 1974). 0,unsaturated galacturonic acid unit.

PECTIN, PECTINASE, AND PROTOPECTINASE

243

lus niger (Hatanaka and Ozawa, 1969). In addition, Nagel and Hasegawa (1968) also described an unsaturated oligogalacturonate hydrolase. 4. Polymethylgalacturonases (PMG)

Although several articles on PMGs have appeared in the literature (Perley and Page, 1971; Finkelman and ZajiC, 1978), the existence of these enzymes is still in question. Polygalacturonase preparations, contaminated with PE, can be mistaken for PMG-containing preparations. Also, if the substrate is not completely esterified, PG or PGL could hydrolyze the glycosidic bonds in these areas. Assay methods similar to those used for determining PG activity can also be applied here, with exception of course of the substrate to be used. C. LYASES

Lyases (or trans-eliminases) perform a nonhydrolytic breakdown of pectates and pectinates, characterized by a trans-eliminative split of the pectic polymer. The lyases break the glycosidic linkage at C-4 and simultaneously eliminate the H from C-5 (Ayers et al., 1966). In some aspects, e.g., induction and catabolic repression (Hubbard et al., 1978; Kurowski and Dunleavy, 1976), lyases resemble hydrolases. This can also be concluded from data in the extensive review article by Linhardt et al. (1986). The methods, used for determination of PG activity, are also suitable here. To detect lyase activity specifically, one assay has been widely and frequently used; measuring the increase in light absorption by reaction mixtures at 230 or 235 nm. At this wavelength, the double bond produced on trans-eliminative cleavage of pectin substrates absorbs maximally. The unsaturated di- and oligouronides also react with thiobarbituric acid to form red chromogens with a maximum absorption at 545-550 nm (Hasegawa and Nagel, 1962; Albersheim et al., 1960). 1. Endopolygalacturonate Lyase (Endo-PGL)

Endopolygalacturonate lyases are produced by several bacteria and fungi (Table VII). On some aspects, PGL can be distinguished clearly from PG. Their pH optima are significantly higher, ranging from 8.0 to 10.0. In addition, all endo-PGLs are activated by Ca2+(e.g.,Lyon et al., 1986) and, in some cases, also to some extent by other divalent cations like Mg2+, Co2+,and Sr2+.It is suggested that a pair of galacturonic acid chains, linked together with a salt bridge, may be the true substrate.

244

TAKUO SAKAI ET AL.

Generally, pectates are good substrates for endo-PGL, but some enzymes exert optimal activity on pectins with a specific degree of polymerization (Pilnik et al., 1973). The main end product from polygalacturonic acid is unsaturated diagalacturonic acid. Lesser amounts of unsaturated trigalacturonic acid and saturated mono- and digalacturonic acids are formed (Moran et al., 1968a). Also, PGL activity decreases as the polymerization degree decreases. 2. Exopolygalacturonate Lyases (Exo-PGL)

Exopolygalacturonate lyases release unsaturated oligogalacturonates from the reducing end of the polymer. The smallest substrate they can hydrolyze is the trimer. Just like endo-PGL, they have a high pH optimum and are activated by the addition of divalent cations (Macmillan and Phaff, 1966). 3. Oligogalacturonide Lyase (OGL)

Some phytopatogenic bacteria, e.g., Erwinia carotovora (Moran et al., 1968b) produce an OGL. Oligogalacturonide lyases are cell-bound enzymes degrading oligogalacturonates or unsaturated oligogalacturonates by removing unsaturated monomers from the reducing end of their substrates by the trans-elimination process. 4. Polymethylgalacturonate Lyase (PMGL)

The PMGLs are the only pectinases proven to be able to hydrolyze pectin. These enzymes are found in some fungi, but rarely in bacteria (Sone et al., 1988). Riedel and Mai (1971) also detected PMGL activity in aqueous extracts of a population of Ditylenchus dipsaci, a plant parasitic nematode. All PMGLs are endo-acting enzymes that cause a rapid drop in viscosity. pH optima range form 5 to 9, and Ca 2 + does not stimulate enzyme activity. The preferred substrates are highly esterified pectins; polygalacturonates are not attacked. Here too, activity decreases with decreasing chain length. IV. Role of Pectic Enzymes in Phytopathogenesis

Evidence to prove the role of pectic enzymes as a cause of (fungal) plant diseases has been accumulating (Ikotun and Balogun, 1987; Morris et al., 1980). These enzymes cause tissue maceration by degrading the pectic substances of the middle lamellae. Endopolygalacturonase and endo-PGL are considered to be the primary enzymes for maceration (Barmore and Brown, 1979). In some cases, the pectic enzymes also convert the pectic polymers of the host plant to a utilizable substrate

PECTIN, PECTINASE, AND PROTOPECTINASE

245

for pathogen growth during pathogenesis (Bateman, 1972). However, the importance of microbial PG in pathogenesis has been established not only in plant diseases characterized by rapid and extensive degradation of host cell walls, but also in some diseases where only a minimal breakdown of cell wall polysaccharides occurs during penetration and colonization of host tissue (Cervone et al., 1987). Collmer and Keen (1986) distinguished several steps during the interaction of a pectinolytic pathogen and a potential host: (1)The entering pathogen possesses structural genes encoding pectic enzymes with particular physical and catalytic properties. (2) These genes are expressed in a characteristic manner in the infected tissue. (3) The enzymes are exported from the pathogen cytoplasm to the host tissue environment. (4) In some tissues the enzymes encounter inhibitors or protected substrates. In other tissues the enzymes are active and cleave structural polymers in the primary cell wall and middle lamella, facilitating pathogen penetration and colonization. Because of the accessibility of pectic polymers in the primary cell wall to enzymatic attack and the consequent rapid release of pectic inducers, pectic enzymes are the first polysaccharidases to be induced when fungi are cultured on isolated cell walls, and the first to be produced in infected tissues. V. Applications of Pectinases

A. INDUSTRIAL PRODUCTION OF PECTINASES It is very difficult to find reliable and detailed information about the commercial production of pectinases. Probably all producer strains are Aspergillus species. Research for additional pectinase producers is hampered by the fact that only a limited number of microorganisms are approved for application in the food industry. The preparations available mostly contain mixtures of PE, PG, and PGL activity (Zetelaki, 1976). The relative amounts of the respective enzymes produced vary considerably with the particular strain used, with nutrient composition, and various environmental factors. In fact, there are three different industrial methods used to produce microbial enzymes; the surface-bran culture (Koji) method, the deep-tank (submerged) process, and the two-stage submerged process. According to Rombouts and Pilnik (1980), most pectinases are still produced by the surface method, carried out in rotating drums, although in general the submerged process is more widely used because of its easier control. A crucial factor in pectinase production is the composition of the medium. Details about such media are considered to be strictly confidential and

246

TAKUO SAKAI ET AL.

are not released by the manufacturers. In general, the medium will be a mixture of carbohydrates (glucose, molasses, CSL, and starch hydrolysates), N sources (NH,+ salts, CSL, DDS, and yeast extract), and minerals. If the enzyme is not produced constitutively, an inducer also has to be added. For reasons of economy, pectin is not used much in production media, but it is substituted by dried sugar cossettes (Zetelaki, 1976), citrus peel, or apple pomace. Control of pH is also very important. The highest enzyme production is achieved when the pH value drops from an initial value of about 4.5 to a more or less constant value of 3.5 during the course of the fermentation, which usually takes 3 to 6 days. At extreme pHs (<3 and >7) a marked inactivation occurs (Schroder and Muller-Stoll, 1962). At the end of the fermentation, the enzymes are extracted from the semisolid medium and mycelium, the dilute enzyme solution is concentrated, and the enzymes are then precipitated with organic solvents or inorganic salts. Following precipitation, the enzyme cake is centrifuged or filtered and then dried at low temperatures or spray-dried. Subsequently, it is ground to a particular particle size and used to prepare commercial enzyme formulations. Some preparations are sold as liquid concentrates. Pectinases are produced by a number of companies in Europe (NovoNORDISK, Miles Kali-Chemie, Swiss Ferment Co.), the United States (Miles laboratories, Rohm and Haas Co.), and Japan (Kikkoman Shoyu Co.). Rombouts and Pilnik (1980) estimated that the worldwide food enzyme production represents a value of about U.S. $45 million, of which perhaps one-quarter relates to pectinases. The development of this enzyme industry has been related with the fruit juice industry. Indeed, because of their low pH optima, pectinases are particularly suitable to be used in this sector. Preferably, it should also be possible to use them at elevated temperatures.

B. FRUITJUICE INDUSTRY

The majority of the pectinase preparations are used in the fruit processing industry. In the beginning of the 1930s publications in Germany and the United States reported on the application of pectinase preparations in apple juice processing in order to facilitate filtration, remove turbidity, and prevent cloud-forming. Also, when apples are of poor pressing quality because of variety or storage, the amount of juice released is low but by treating the pulp with pectinases the juice yield is increased (De Vos and Pilnik, 1973). Almost as old as the application in clarification is the application in extraction. At first, pectinases were

PECTIN, PECTINASE, AND PROTOPECTINASE

247

added to black currants to facilitate extraction by pressing and were later added to other soft fruits like raspberries and black cherries. After crushing, these crude fruit juices are often very viscous and sometimes slightly gelified. It is also very difficult to separate remaining solids from the juice. By adding pectinases, however, viscosity drops and it becomes possible to extract juice by pressing. Some preparations are used to produce juices by liquefaction. In this process, cell walls are dissolved by a combination of pectinases and cellulases and a yield (juice) of up to 100% can be obtained (Dorreich, 1983; Janda, 1983). Also, mechanical desintegration is sometimes replaced by enzymatic maceration (Rohm, 1969). Maceration results in the release of single cells, leaving intact cell walls, and thereby also releasing flavor compounds, pigments, and active ingredients in an intact state (Silley, 1986). Problems in clarification of fruit juices are caused mainly by the presence of pectic substances which suspend toward insoluble (pulp) particles. After treatment with pectinases, these particles can be separated by sedimentation or filtration. This technique is also used in wine production. An additional advantage is the improved liberation of anthocyanins as a consequence of tissue degradation and breakdown of anthocyanin-pectin complexes. As a result the “color yield” increases and the production time shortens (Pilnik, 1981). Pectinases induce clarifying, filtration improving, colour liberating, and yieldenhancing effects (Grampp, 1982). In contrast, for some fruit juices turbidity is wanted. In these cases, the native enzymes must be inactivated, mostly by heat. This prevents well-known quality defects such as cloud loss in citrus juices and gelation of concentrates, caused by citrus pectinesterases through deesterification of juice pectin which subsequently precipitates or gels as calcium pectinate or pectate.

C . OTHERAPPLICATIONS

Pectinases are also involved in the retting process (Ali, 1958).Retting is a fermentation process in which certain bacteria (e.g., Clostridium, Bacillus) and fungi (e.g., Aspergillus, Penicillium, Cladosporium) decompose pectins of the bark and release the fiber (Chaudhury, 1953). This process plays an important role in the production of many important textile fibers such as flax,hemp, and jute. Novo-NORDISK developed an enzyme preparation, “flaxzyme,” which accelerates and improves this process, Fogarty and Ward (1972) were the first to report

248

TAKUO SAKAI ET AL.

the potential application of pectinase producing organisms or their enzymes to treat commercial softwoods in order to render them more amenable to treatment with preservatives. VI. Protopectin-Solubilizing Enzyme (Protopectinase)

The enzyme that catalyzes the solubilization of protopectin was originally named protopectinase by Brinton et al. 1927. They proposed that this term should be applied to an enzyme that hydrolyzes or dissolves protopectin, causing plant cells to separate from each other, a process which is usually called maceration. The term of “protopectinase” superseded the older term “pectosinase,” with which it was synonymous. However, further research on pectic enzymes showed that the decomposition of protopectin was due to the action of a system of enzymes, including pectinesterase, endo-polygalacturonase, endo-pectate lyase, and pectin lyase. Kaji (1956, 1959) found a microbial enzyme in the culture filtrate of Clostridiurn felsineurn that catalyzes the breakdown of middle lamella pectic substance from the bark of gampi (Wikstremia sikokiana Fr. et. Sav.) to a soluble pectic substance. The enzyme activity that liberates pectin from plant tissues was also found in the culture filtrate of Aspergillus japonicus by Ishii (1976). He found that the enzyme reaction was catalyzed by the combination of endo-polygalacturonase and endo-pectin lyase. Karr and Albersheim (1970), studying Pectinol R-10, a mixture of enzymes produced by Aspergillus niger, isolated an enzyme that liberated a pectic substance from protopectin but degraded pectic acid only to a limited extent. Details about the enzyme were not studied. In these reports, the protopectin-solubilizing enzyme has been regarded as an enzyme that macerates plant tissues, and little was known about enzymes that liberate highly polymerized pectin (pectinliberating enzymes). In 1978, the first study on pectin-liberating enzymes was reported by Sakai and Okushima. A microorganism was detected that produced a protopectin-solubilizing enzyme, which liberated water-soluble and highly polymerized pectin from protopectin. They also reported that protopectin is solubilized by restricted hydrolysis and called such enzymes “protopectinase” (PPase). Since then, several PPases, which are classified into two types depending on their reaction mechanism, have been isolated (Sakai and Okushima, 1982; Sakai and Yoshitake, 1984; Sakai et a]., 1984; Sakai and Sakamoto, 1990). One type PPase reacts with the polygalacturonic acid region of protopectin (inner site) and the other on the polysaccharide chains that may connect the polygalacturonic acid chain and cell wall constitutents

PECTIN, PECTINASE, AND PROTOPECTINASE

249

(outer site), as shown in Fig. 18.Sakai and co-workers called the former A-type PPase and the latter B type PPase. These enzymes are described here in more detail. ACTIVITY A. ASSAYOF PROTOPECTINASE PPase activity is assayed by measuring the amount of pectic substance liberated from protopectin by the carbazole-sulfuric acid method (Furutani and Osajima, 1965). The reaction mixture contains 10 mg of protopectin, 100 pmol of acetate buffer [AcB) containing 50 pgiml bovine serum albumin, pH 5.0, and 50 p1 of enzyme solution, in a total volume of 1.0 ml. The reaction mixture is incubated beforehand for 1 hr at 37°C. The reaction is started by the addition of the enzyme solution, and the mixture is kept for 1 hr at 37°C. The reaction is stopped by cooling the

Neutral sugar sidechain

I reaction with A-type protopectinase

I I I

reaction with B-type protopectinase

I

M

4 % A /-

High molecular weight pectin

Low molecular weight pectin

FIG 18. Schematic illustration of structure of protopectin and reaction mode with Atype and B-type protopectinases.

250

TAKUO SAKAI ET AL.

reaction mixture in an ice bath. The control blank is run with the use of a heat-denatured enzyme solution. After the reaction, the mixture is filtered on filter paper. To a test tube containing 250 p1 of filtrate of the reaction mixture, 3 ml of chilled 32 N H2S0, solution is introduced, followed by 250 p l of 0.2% carbazole in ethanol. This step is done in an ice bath. The assay mixture is heated at 75°C for 20 min and cooled to room temperature, after which the optical density at 525 nm is measured. The pectin concentration is measured as D-galacturonic acid from a standard assay curve with D-galacturonic acid. One unit of PPase activity is defined as the activity that liberates pectic substance corresponding to 1 pmol of D-galacturonic acid per milliliter of reaction mixture at 37OC in 1 hr, and the specific activity is expressed as units per milligram of protein. Protopectin used in routine experiments can be prepared from lemon (Citrus limon Burm) peel by the following procedure. The albedo layer of the peel is scooped out, pooled, washed with distilled water until the water-soluble substances that react with carbazole-sulfuric acid are washed off, and then lyophilized. The dried protopectin preparation is stored in the refrigerator. In the experiments with B-type PPase, protopectin treated with ethylenediaminetetraacetic acid (EDTA)is used. The EDTA-treated protopectin is prepared from protopecitn as obtained earlier. The protopectin is washed with 50 mM EDTA until EDTA-soluble pectic substances are washed off completely, washed with distilled water, and lyophilized. B. A-TYPEPROTOPECTINASE

Two types of A-type PPases are known: one has polygalacturonic acid hydrolyzing activity (A, type), and the other has polygalacturonic acid transeliminase activity (A2type). 1. A,-Type PPases

a. Occurrence. Some A,-type PPases are found in the culture filtrate of yeasts and yeast-like fungus. They have been isolated as crystals form culture filtrates of Kluyveromyces fragilis IF0 0288(Sakai et al, 1984), Galactomyces reessii L. (Sakai and Yoshitake, 1984), and Trichosporon penicillatum SNO 3 (Sakai and Okushima, 1982); they are called PPase-F, -L, and -S, respectively.

b. Purification of PPases. PPases are extracellular proteins and they are purified from culture filtrates in basically the same way (Sakai,

PECTIN, PECTINASE, AND PROTOPECTINASE

251

1988). The purification procedure for PPase-F is the most complicated, and will be described here. Step 1: Production of PPase-F. Kluyveromyces fragilis IF0 0288 is used for the production of PPase-F. The yeast is maintained on agar slants of a medium containing 2% glucose, 0.6% peptone, and 0.5% yeast extract, pH 5.0. For enzyme production, the yeast is aerobically cultured in a medium (40 liters) containing 3% glucose, 0.6% peptone, 0.2% yeast extract, and 0.08% Silicone KM-70 (an antifoaming agent), pH 5.0, at 30°C. Production of the enzyme begins after about 5 hr of cultivation, and reaches a maximum at 15 hr of cultivation. The culture filtrate (37 liters) is concentrated by evaporation at reduced pressure at 30°C (to 1.5 liters) and is used for enzyme purification. Step 2: CM-Sephadex C-50 Column Chromatography. The concentrated culture filtrate is dialyzed thoroughly against 20 mM AcB, pH 5.0, and then put on a CM-Sephadex C-50 column (3 x 50 cm) equilibrated with 20 mM AcB, pH 5.0. The column is washed thoroughly with 20 mM AcB, pH 5.0, and the enzyme is then eluted with 350 ml of a linear gradient of NaCl at from 0 to 400 mM in the same buffer, at the flow rate of 20 ml/hr. The fractions containing enzyme activity are pooled and concentrated to about 5 ml by evaporation at reduced pressure at 30°C. Step 3: Sephadex G-75 Column Chromatography. The concentrated enzyme solution is chromatographed on a Sephadex G-75 column (2.2 x 80 cm) equilibrated with 20 mM AcB, pH 5.0, containing 200 mM NaC1, and elution is done at the flow rate of 6.7 ml/hr. PPase activities are recovered in two peaks (Fig. 19a). Most of the activity is recovered in fraction I1 (F-11),which is concentrated to 2 ml by evaporation at reduced pressure at 30°C. The chromatography is repeated once more and the enzyme solution obtained is concentrated to 10 ml. Step 4: Crystallization. Solid ammonium sulfate is added to the enzyme solution until faint turbidity is observed. After being left for 1 week in a refrigerator, the enzyme forms needle-like crystals (Fig. 20). The crystallization is repeated two more times. From 37 liters of culture filtrate, about 50 mg of crystalline enzyme is obtained, with a recovery of about 40%. The enzyme preparation is homogeneous on the criteria of electrophoresis and sedimentation analysis. Fraction I (F-I), which is eluted at around the void volume of the Sephadex G-75 column, is chromatographed again on a Sepahdex G-100 column (2.5 x 60 cm) with 20 mM AcB pH 5.0, containing 200 mM NaCl as the solvent and the flow rate of 2.7 ml/hr. The activities are recovered in two peaks (Fig. 19b). The molecular weight of F-1-1 is high, so Sephadex G-200

252

TAKUO SAKAI ET AL. FI-lb

FI-la

Fraction number

’ 0

30

50

K r - y

70

Fraction number (5rnVfraction) FIG.19. Chromatograms of protopectinases from K. fragilis IF0 0288 on Sephadex columns. (a] G-75; (b) G-100;(c ) G-200. -0-, protopectinase activity; ---0---, protein (Sakai, 1988).

is used instead of Sephadex G-100; the column (1.6 x 80 cm) is equilibrated with 20 mM AcB, pH 5.0, containing 200 mM NaCI. By eluting with the same buffer at a flow rate of 0.7 ml/hr, activity is recovered in two peaks (Fig. 19c). Thus, this strain seems to produce at least four PPases of different molecular weights.

c. Properties of the PPases. Some physical and biological properties of the three PPases (PPase-F, -L, and -S) are shown in Table VIII. These three PPases are similar in biological properties as well as in molecular weight (about 30,000), but not in specific activities. Table IX shows the amino acid composition and carbohydrate content of PPases. Amino acid compositions of these enzymes are different. The antiserum to PPase-S gives precipitation lines with PPase-L and -S, but it does not react with PPase-F (Sakai, 1988). Amino acid sequences at the N-terminal and the 27 residues long fragments in PPase-

PECTIN, PECTINASE, AND PROTOPECTINASE

Protopectinase-F

253

Protopectinase-L

Protopectinase-S FIG.20. Photomicrographs of crystals of protopectinases.

L and -S are identical (Fig. 21) (Sakai, 1988). PPase-F is not homologous to these two enzymes.

d. Catalytic Properties. The enzymes have pectin-releasing effects on protopectins from various origins. This is called PPase activity. The enzymes catalyze the hydrolysis of polygalacturonic acid; they decrease viscosity while slightly increasing the reducing value of reaction medium containing polygalacturonic acid (Sakai, 1988). Because of these findings, the enzymes are classified endo-polygalacturonases [EC 3.2.1.15; poly(l,4-a-~-galacturonide)glycanohydrolase].

254

TAKUO SAKAI ET AL. TABLE VIII PHYSICOCHEMICAL AND BIOLOGICAL PROPERTIES OF PROTOPECTINASES Properties

Molecular weight By electrophoresis By gel filtration By sedimentation S2O.W

E;& nm Isoelectric point N-terminal amino acid Optimum pH Optimum temperature ("C) Inhibitor pH stability Activity (U/mg) Protopectinase Polygalacturonase K , value (mgfml) For protopectin For polygalacturonic acida

PPase-F

PPase-L

PPase-S

40,000 33,000 32,800 2.99s 10.0 5.0

40,000 30,000 29,300 3.77s 11.9 8.4-8.5

40,000 30,000 29,300 3.66s 9.20 7.6-7.8

Aspartic acid 5.0 60

Glycine 5.0 55

Hgz+,Hg+, CaZ', Hgz+,Hg', Ag', Ba2+,Ca2+,PbZ+ Ba", Coz+

Glycine 5.0 50

Hg2+,Hg+,Ca2+, Ba2+,Coz+

2-8

3-7

3-7

556 2,053

3,945 16,219

5,770 21,107

90 6.6

50 7.7

30

9.0

a Polygalacturonic acid, having a mean polymerization degree of 130, was used for the determination of the K, value.

The hydrolysis of galacturonic acid oligomers is different for different enzymes. Figure 22 shows the mode of action in the hydrolysis of galacturonic acid oligomers and gives the K, and V,, values for the reaction. Three patterns of action toward galacturonic acid oligomers are known for endo-polygalacturonases (Demain and Phaff, 1954; Pate1 and Phaff, 1959, 1960; Mill and Tuttobello, 1961; Nasuno and Starr, 1966; Koller and Neukom, 1969; Rexova-Benkova, 1973; Kimura et al., 1973). PPase-S is novel in its action pattern toward oligogalacturonic acids. The K, and V, values change with the substrate chain length; the K, values tend to decrease and the V, values tend to increase with increasing chain length. V,,, is very different with trigalacturonic acid and tetragalacturonic acid. In contrast, the number of methoxyl groups in the substrate affects the molecular weight of the reaction products; the molecular weight of the reaction products increases as the number of methoxyl groups in the substrate galacturonic acid increases (Fig. 23).

PECTIN, PECTINASE, AND PROTOPECTINASE

255

TABLE IX AMINOACIDAND SUGAR COMPOSITIONS OF PROTOPECTINASES Amino acid residuesa per molecule of protopectinase ~~~~~~

~

Amino acid

PPase-F

PPase-L

PPase-S

Lysine Histidine Argin in e Tryptophan Aspartic acid Threonine Serine Glutamic acid Proline Glycine Alanine Half -cystine Valine Methionine Isoleucine Leucine Tyrosine Pheny lalanine Suaar

15 4 4 5 40 31 32 10 5 30 10 4 13.

16 6 4 9 35 19 29

13

1 13 11 4

7 1Ob

17

6 42 17 1 18 0 20 11 5 8 9C

5 6 5 37 30 39 20 6 34 16 6 16

1 21 11 4 10 3b

Calculations based on a molecular weight of 30,000 Determined as mannose. Determined as rhamnose.

e. Postulated Mechanism of Protopectinase Activity. On the basis of the kinetic properties of the enzyme action as a polygalacturonase, the mechanism of PPase activity seems to be as follows: the enzyme reacts with the pectin molecule in protopectin at sites with three nonmethoxylated galacturonic acid chains or more (actually, four or more

PPase-F PPase-L PPase-S PPase-SEl P Pase- SE2 PPase-SE3

D-S-G-T-L-S-G-K-T-AGG-G-L-S-N-?-A-T-V-T-V-N-N-V-?-V-P-AG-

G-G-A-?-V-F-K-D-AQ-S-A-I-AG-K-A-S-?-?-S-I-T-LB-N-F-A-V-PG-G-A-?-V-F-K-D-AQ-S-A-I-AG-K-A-S-S-S-S-I-?-LQ-N-FG-G-A-C-V-F4C-D-A-Q-S-A-IA-G-K-AG-G-A-?-V-F-R-D-A*-S -A-I- A-G-K-K-S G-?-A-?-V-Fg-D-A-K-S-A-I-A-G-K-K FIG.21. N-terminal amino acid sequences of A,-type PPases.

256

TAKUO SAKAI ET AL. Enzyme

Substrates

Reaction products

ooo -0-00 oooo/oooo

KmhM) >I02

'00 00

Protopectinase-F

Vmar(AM/unit/min)
1.82

1.9ox10-2

3.98

1.96X10-3

2.77

2.37X 10-2

7.09X lo-'

9.79 X lo-'

4.26

4.79X10-4

2.20

5.20X10-'

........

ooo - w o Protopectinase-L OOOO / o o o o

~0-000

00000--00QOO ....... .

ooo

-000

Protopectinase-S oooo / o o o o

FIG.22. Mode of action of protopectinases toward galacturonic acid oligomers. Reactions are performed under the optimum conditions for each enzyme. Hexagons represent D-galacturonic acid molecules.

nonmethoxylated galacturonic acid chains, considering the reaction velocities for galacturonic acid oligomers) and cleave their glycosidic linkages. In general, 50% of the galacturonic acids of pectin in protopectin are methoxylated, at random. Therefore, the pectin molecule in protopectin may be cleaved at restricted sites so as to form highly polymerized pectin. As mentioned earlier, these three PPases are type of endo-polygalacturonase. Sakai and Takaoka (1985) isolated an endo-polygalacturonase from the culture filtrate of Aureobasidium pullulans (polygalacturonase-AY), which degrades polygalacturonic acid strongly but has weak PPase activity. The enzyme has a lower affinity for protopectin from Citrus unshiu than PPase-F, -L, or -S; the K, value for protopectin is one order higher than those of the other three PPases, although they have almost the same affinity for polygalacturonic acid (Table X). The differences in affinity for protopectin seem to be one reason why different endo-polygalacturonases have different PPase activity.

PECTIN, PECTINASE, AND PROTOPECTINASE

..! 2 r v c

A Protopectinase-F 0 Protopectinase-L 0 Protopectinase-S

E l

I

I

I

1

8

t

20 30 40 50 60 70 Esterified galacturonic acid content (%)

0

257

10

I

80

FIG. 23. Relationship between the amount of methoxyl groups of the substrate and the molecular weight of the degradation product of the enzyme reaction. Reactions are performed with polygalacturonic acid (mean polymerization degree, 33) containing the various amounts of methoxyl groups indicated.

TABLE X AFFINITY OF A,-TYPE PROTOPECTINASES AND POLYGALACTURONASE-AY ON PROTOPECTIN AND POLYGALACTURONIC ACID

K, values Enzymes PPase-F PPase-L PPase-S PPase-SEl PPase-SEZ PPase-SE3 Polygalacturonase- AY

Protopectina (mg/mlJ

Polygalacturonic acid (mg/ml)

90 30 50 55

6.6 9.0 7.7

7.9

0.33 0.24 0.23 0.23

3.8

0.18 0.03

0.15 525

~~

a

Ratio of activity (protopectinasei polygaiacturonase)

Obtained from Citrus unshiu peel.

258

TAKUO SAKAI ET AL.

Thus, PPase activity is not a property of all endo-polygalacturonases, and only PPase-F, -L, and -S have been found to have this activity so far. There are differences in their amino acid compositions, although the methionine content is low in all four enzymes. PolygalacturonaseAY contains more threonine than PPase-F, -L, and -S, and the star diagram of amino acid contents of polygalacturonase-AY is somewhat different than those of three PPases (Fig. 24). The relationship between the amino acid composition and the substrate specificities is not clear.

f. Multiform of PPase-S. As mentioned earlier, K. fragilis produces at least four PPases having different molecular weights. T. Sakai et al. found that a PPase overproducing mutant of T . penicillatum (termed

Polygalacturonase-AY

Pmtopectinase-L

Protopectinme-F

Protopectinase-s

FIG. 24. Star diagrams of protopectinases and polygalacturonase-AY. The circles in the figures indicate 10 and 20 mol%.

259

PECTIN, PECTINASE, AND PROTOPECTINASE

strain B 2y3), induced by y-ray irradiation, produces three PPases (PPase-SEl, -SE2, and -SE3). The properties of these enzymes are somewhat different, although basically they have similar immunological properties (unpublished results). g. Purification and Isolation of PPases from Culture Filtrates of T. penicillatum B2y3. The ratio of protopectinase activity/polygalacturonase activity (PPaseIPGase)is not consistent and depends on the fermentation batch. The strain produces three PPases [PPase-SEl, -SE2, and -SE3) which have different PPaseIPGase values [Table X). This may be why the PPase/PGase value of the culture filtrate depends on the culture batch. These three enzymes cross-react with rabbit antiserum against PPaseSE1 (antibody-S), which is the major PPase produced by this strain, although the profiles of inactivation with the antibody are different as shown in Fig. 25. Among these PPases, PPase-SE3 is less reactive with 120

100

3 Y

.->)r CI

80

5

m

60

.-cE

2

. I

40

20

0 0

2

4

6

8

10

12

Concentration of antibody (%) FIG.25. Profiles of inactivation of PPase-SEl (O),-SE2 (0), and -SE3 (m) with treatment of antiserum against PPase-SEl.

260

TAKUO SAKAI ET AL.

antibody-& and the mobilities of the enzymes in PAGE are different (Fig. 26). These data indicate that the molecular structure of these PPases is different; PPase-SEX and PPase-SE3 may have low homology. However, homologies of the N-terminal amino acid sequences of these enzymes are high (more than 90%; Fig. 21). h. Gene Cloning of PPase-SE3 and Structure of the Gene. T. Sakai and K. Iguchi have cloned the PPase-SE3 gene (unpublished results). The gene consists of 1104 bp and encodes a polypeptide of 367 amino acids (Fig. 27). The open reading frame (ORF) for the gene is initiated by a codon whose sequence alignment ACAATGC is not very similar to the consensus sequence ACCATGG for the translation initiation sequence of eukaryotic genes. They assumed that this difference is one of the reasons for poor production of PPase-SE3. As the TATA box, the TATAAAT sequence is located at position -129 upstream to the initiation codon. In the 3’-nontranslated region of the ORF, polyadenylation consensus sequences TAGT and CAATG are located at positions + 1270 and + 1193 of the termination codon (ATT),respectively. By comparing the deduced amino acid sequence and the N-terminal amino acid sequence of the PPase-SE3, a 27 amino acid signal peptide is observed. This signal peptide is characterized by a positively charged N-terminal region, a hydrophobic core, a serine or threonine region, and a polar region. Different from the (-3, -l)-rule for the signal peptide cleavage site according to von Heijne (1983),there is a positively charged arginine at position -1 and a polar amino acid glutamic acid residue at position

FIG 2 6 . PAGE of PPase-SEl, -SE2, and -SE3.

PECTIN, PECTINASE, AND PROTOPECTINASE

261

-3. This is similar to an endo-polygalacturonase from A. niger RH5344

(Ruttkowski et a]., 1990). Thus, PPase-SE3 is probably synthesized as a protein of molecular weight 37,967 and the signal peptide is then removed to form a mature enzyme of molecular weight 35,204. 2. A,-Type PPase Two A,-type PPases have been isolated form a culture filtrate of Bacillus subtilis (T. Sakamoto and T. Sakai, unpublished results). a. Occurrence of A,-type PPases. Two PPases that split the glucoside linkage of the polygalacturonic acid (or methoxylated polygalacturonic acid) region in protopectin by trans-elimination reaction have been found in Baccilus subtilis IF0 3134. One is active on polygalacturonic acid, and the other has potent activity on methoxylated polygalacturonic acid. These were names PPase-N and PPase-R, respectively.

b. Purification of PPase-N and PPase-R. Both enzymes are purified from the culture filtrate of B. subtilis IF0 3134. For the enzyme production, the microorganism is cultured in a medium containing 1.0% soybean flour (defatted material), 2.8% K,HPO,, and 1.2% KH,PO,, pH 7.0, in a 500-ml shaking flask containing 100 ml of medium on a shaker (120 rpm) at 37OC. Purification is done by column chromatographies using Butyl-Toyopearl 650M, CM-Toyopearl 650M, and Superose 1 2 and can be isolated as a homogeneous protein. Butyl-Toyopearl gives a good separation of PPase-N from -R and -C (this strain also produced PPase-C and the details are described later). PPase-R and PPase-C are separated in CM-Toyopearl column chromotography. Ethylenediaminetetraacetic acid (EDTA) inactivates PPase-N but not PPase-R, so that measuring PPase-R activity is carried out in the presence of 4 mM EDTA. The purification procedures and some corresponding elution patterns of PPase-R are shown in Table XI and Fig. 28. c. Properties of PPase-N and PPase-R. Some physicochemical and biological properties of both PPases are summarized in Table XII. PPaseN is very susceptible to divalent cations and EDTA, but PPase-R is insensitive to EDTA. These enzymes are stable in a wide range of pH and temperature. These enzymes are different proteins by the criterion of immunological properties. The N-terminal amino acid sequences are also quite different (Fig. 29). The substrate specificities of the enzymes toward pectic acid with various degrees of esterification are summarized in Table XIII. Polygalacturonic acid is the best substrate for PPase-N,

-130 -120 -110 -100 GTTTACTATAAATATCTGTGAATCGCCTGCAATTATTTTT -90

-80

-70

-60

-50

-40

-30

-20

-10

10

TTTTTTGATAAATCTTCAAGCTCAACAACTCTTCTTCTTTTGAACTGATAAAAAGCCTTATAGACTCTCTTATTTGTTCACTCTTAACTATAACAATG CTT TTT TCT AAA TCT Met Leu Phe Ser Lys Ser 20

30

40

50

60

70

80

90

100

GCT ATC TTT GCT ATG GCT GCT CTT GCA GTT GCT GCT CCT ACT GAA GGT GAC CTT CAA GCT CGT GGC AGC GCC TGT GTT TTT AAG GAT GCC Ala Ile Phe Ala Met Ala Ala Leu Ala Val Ala Ala Pro Thr Glu Gly Asp Leu Gln Ala Arg Gly Ser Ala Cys Val Phe Lys Asp Ala 110

120

130

140

150

160

110

180

190

AAG TCT GCT ATT GCT GGC AAG AAG TCT TGT TCT TCT ATC ACT CTT GAG AAC ATT GCT GTC CCC GCT GGT CAA ACT CTT GAT CTC ACT GGA Lys Ser Ala Ile Ala Gly Lys Lys Ser Cys Ser Ser Ile Thr Leu Glu Asn Ile Ala Val Pro Ala Gly Gln Thr Leu Asp Leu Thr Gly N CE N

200

210

220

230

240

250

260

270

280

CTT GCC AAG GGC ACT GTT GTC ACC TTT GCT GGT ACC ACC ACT TTT GGC TAC AAG GAG TGG GCT GGT CCT TTG ATC TCC GTT TCT GGT GAT Leu Ala Lys Gly Thr Val Val Thr Phe Ala Gly Thr Thr Thr Phe Gly Tyr Lys Glu Trp Ala Gly Pro Leu Ile Ser Val Ser Gly Asp 290

300

310

320

330

340

350

360

370

TCT ATT ACT GTC AAG CAG GCC TCC GGC GGT AAG ATT GAC TGT GGT GGT TCT CGT TGG TGG GAC GGT AAG GGT TCC AAC TCT GGT GGT AAG Ser Ile Thr Val Lys Gln A l a Ser Gly Gly Lys Ile Asp Cys Gly Gly Ser Arg Trp Trp Asp Gly Lys Gly Ser Asn Ser Gly Gly Lys 390 400 410 4 20 430 440 450 460 CAA AAG CCC AAG TTC TTT TAC GCC CAC AAG CTC CAG AAC TCC AAC ATT CAG GGA CTC CAA GTT TAC AAC ACC CCT GTC CAG GCT TTC AGC Gln Lys Pro Lys Phe Phe Tyr Ala H i s Lys Leu Gln Asn Ser Asn Ile Gln Gly Leu Gln Val Tyr Asn Thr Pro Val Gln Ala Phe Ser

380

470

480

4 90

500

510

5 20

5 30

540

550

ATT TTG TCT GAC CAT TTG ACT TTG TCA AAC ATT CTC GTT GAC AAC AGA GCC GGT GAC AAG GCT GGT GGT CAC AAC ACC GAC GCT TTT GAT Ile Leu Ser Asp H i s Leu Thr Leu Ser Asn Ile Leu Val Asp Asn Arg Ala Gly Asp Lys Ala Gly Gly His Asn Thr Asp Ala Phe Asp 560

570

580

590

600

610

620

630

640

GTT GGT ACC AGT ACT TAC ATC ACT ATC GAC CAT GCT ACG GTC TAC AAC CAG GAC GAC TGT CTT GCT ATC AAC TCC GGT G.4C CAC ATC ACT Val Gly Thr Ser Thr Tyr Ile Thr Ile Asp H i s Ala Thr Val Tyr Asn Gln Asp Asp Cys Leu Ala Ile Asn Ser Gly Asp H i s Ile Thr

650

660

670

680

690

700

710

720

730

TTC CAG AAC GGT W C TGC TCT GGT GGA CAT GGT CTT TCT ATT GGC TCC GTT GGT GGC CGC TCT CTT AAC ACT GTT TCC AAC GTC AAT ATT Phe Gln Asn Gly Phe Cys Ser Gly Gly His Gly Leu Ser Ile Gly Ser Val Gly Gly Arg Ser Leu Asn Thr Val Ser Asn Val Asn Ile 740

750

760

770

780

I90

800

810

820

CTT AAC AGT CAG GTT GTC AAC TCT GAT AAC GGT GTC CGT ATT AAG ACC ATT TCT GGT GCT ACT GGT TCT GTC AGC GGT GTT AAG TTC CAG Leu Asn Ser Gln Val Val Asn Ser Asp Asn Gly Val Arg Ile Lys Thr Ile Ser Gly Ala Thr Gly Ser Val Ser Gly Val Lys Phe Gln 830

a40

850

860

870

880

890

900

910

GAC ATT ACT CTC TCC AAC AT" GCC AAG TAC GGT ATT GAT GTT CAG CAG GAC TAC CGT AAC GGT GGC CCC ACT GGT AAC CCC ACT AAC GGA Asp Ile Thr Leu Ser Asn Ile Ala Lys Tyr Gly Ile Asp Val Gln Gln Asp Tyr Arg Asn Gly Gly Pro Thr Gly Asn Pro Thr Asn Gly 930 940 950 960 970 980 990 1000 GTC AAG ATC ACT GGA ATC GAG 'l"'C ATC AAC ATT CAC GGT AGT GTC AAG AGC TCT GGT ACC AAC GCT TAC CTT CTC TGT GGT TCC GGC TCT Val Lys Ile Thr Gly Ile Glu Phe Ile Asn Ile His Gly Ser Val Lys Ser Ser Gly Thr Asn Ala Tyr Leu Leu Cys Gly Ser Gly Ser

920 N

m

1010

1020

1030

1040

1050

1060

1070

1080

1090

TGC TCC AAC TGG ACC TGG AGC AAG ATC AAC GTC AAG GGT GGC AAG GAC AGC GGT GCT TGC AAG AAC GTT CCT TCT GGT GCT ACT TGC AAA Cys Ser Asn Trp Thr Trp Ser Lys Ile Asn Val Lys Gly Gly Lys Asp Ser Gly Ala Cys Lys Asn Val Pro Ser Gly Ala Thr Cys Lys 1100

1110

1120

1130

1140

1150

1160

1170

1180

1190

1200

1210

CTT TAA ATAGCAACTATATATACTACCTTAAACTAAGAAACTTACATTATTTTTAAACACTATTAAATTTTCACAAATTTAATACAC~ACGTTCAATGAAGGGAAACCGTGGACAGT Leu *** 1220 1230 1240 1250 1260 1270 1280 1290 1300 1310 1320 1330 AAGMTGAGTTGAAACTTGACTGAATAC~GAGTTGTTATTTTGTACTTGTTTTAGTAAGCCTAATAAGGATCCATCATTT~GGAAAAAAAAAT~TAAT~TAAACGTTCATTTAT

1340

1350

1360

CTTCAAAATGTCATCAGTAAAGTTTCC FIG.27. Nucleotide sequence of the PPase-SE3 structural gene and flanking regions. The deduced amino acid sequence of PPase-SE3 is also given.

TAKUO SAKAI ET AL.

264

TABLE XI PURIFICATION OF PPASE-RFROM Bacillus subtilis

IF0 3134

Purification step

(ma1

PPase ( x 103 u)

Spec. act. (U/mg)

Purification (-fold)

Culture filtrate Butyl-Toyopearl 650M' Butyl-Toyopearl 650M' CM-Toyopearl650M Butyl-Toyopearl650M' Superose 12 Superose 12

31,100 2,780 411 12 6.1 0.9 0.5

1730 851 212 118 76 22 18

56 306 516 9,830 12,500 24,400 36,000

-

-

5 9

49 12

176 223 436 643

4 1 1

Protein

a

Yield

(%I

7

A linear gradient of 35-0% of ammonium sulfate was carried out

while PPase-R has potent activity on pectic acid with high degrees of esterification. In the reactions, using polygalacturonic acid (or methoxylated polygalacturonic acid) or protopectin as the substrate, the enzymes produce the substance having absorption maximum at 235 nm, which may originate from the 4,5-unsaturated galacturonide. With both enzymes, in the reaction on polygalacturonic acid or methoxylated polygalacturonic acid, relative viscosity of substrate solutions reduced over 50% when only a few percent of their glycosidic linkages are split. These data indicate that PPase-N and PPase-R split glycosidic linkage of pectic substances in at random manner. Thus, PPase-N is an enzyme belonging to the category of pectate lyase [poly(l,4-cr-~-galacturonide)lyase; EC 4.2.2.21 and PPase-R is an enzyme belonging to the category of pectin lyase [poly(methoxygalacturonide)lyase;EC 4.2.2.101, respectively.

C. B-TYPEPROTOPECTINASE As mentioned earlier, the reasons for the insolubility of the pectin in protopectin are complex, and include secondary valency bonding between pairs of pectin molecules or with other cell wall constituents such as cellulose or hemicellulose. The features of the insolubility of the pectin in protopectin suggest that other enzymes-different from A-type PPases-have PPase-like activity, with a mechanism different from that involving the restricted hydrolysis of the polygalacturonic FIG 28. Elution patterns on various column chromatographies for the purification of PPase-R.

Culture filtrate

J. 3 -

'

PPase-R

...........................

I

0.6

-

0.4

200

500 FM-TovoDearI

PPase-N

300

750

1000

PPase-R

1250

-

0 900

......

......... ..... ....

600

L

0.5

E -

30.2

(0

0

8

.-c> .-

300

l= gE

03

..-...... 0

0.09

I

750

0.06

E

0

0.03

250

jE

n.2

0

0 0.9

I

0.6

E

$

Elution volume (ml)

266

TAKUO SAKAI ET AL. TABLE XI1 PROPERTIES OF PPASE-NAND PPASE-R Properties Molecular weight SDS-PAGE Gel filtration Isoelectric point (pH) Optimum pH (at 37OC) Optimum temperature (pH 6.0) Inhibitor

pH stability (at 37°C for 16 hr) Thermostabilitv

PPase-N

PPase-R

43,000

35,000

32,000 9.4

27,000 8.2

8.0

60°C

EDTA, HgZt Mn2+,CuZt ZnZ+,BaZ+ 3-10 -60°C

8.0

60°C HgZ +

4-11 -60°C

acid region in protopectin. T. Sakai et al. have found PPases which have a different mechanism from A-type PPases. 1. Occurrence of B-Type PPase

A PPase that did not degrade polygalacturonic acid was first detected in the culture filtrate of B. subtilis IF0 3134 (Sakai and Sakamoto, 1990). The enzyme, called PPase-C, has potent activity on the protopectin from sugar beet pulp but has much less on lemon peel protopectin. The same type of PPase (PPase-T) is found in the culture filtrate of microorganisms belonging to Trametes (T. Sakai et a]., unpublished results). 2. Purification of PPase-C and PPase-T PPase-C is purified from the culture filtrate of B. subtilis I F 0 3134. The microorganism is cultured in a medium containing 0.5% dextrin, 0.05% yeast extract, 1% NH,H,PO,, 0.05% CaCl2.2H,O, 0.02% MgS04-7H,0, 0.02% KC1, and 0.5% sugar beet extract at 37°C for 20 hr. The purification procedure consists of treating the culture filtrate with EDTA followed by chromotography on CM-Cellulofine CH, butyl-

PPase-N A-D-L-G+Q-T-L-G-S-N-DG-D-G-A-Y-SPPase-R A-V-D-F-P-N-T-K-K-N-G-L-L-G-F-A-G-N--4-K-N-E-K-G1PPase-C S-F-WG-A-S4-E-L-L-H-D-R-T-MIK-E-G-S-S- W-Y-A-LG-T-N-L-NFIG.29. Comparison of N-terminal amino acid sequences of PPases from B. subtilis I F0 3134. * Italic means uncertain.

PECTIN, PECTINASE, AND PROTOPECTINASE

267

TABLE XI11 ACTIVITYOFPPASE-N AND PPASE-RTOWARD PECTIC ACIDWITH VARYING DEGREES OF METHYL ESTEFUFICATIOV Relative activity (YO) Substrate

PPase-N

PPase-R

Pectic acid Methoxylated pectic acid (13% esterified) Methoxylated pectic acid (26% esterified) Methoxylated pectic acid (35% esterified) Methoxylated pectic acid (56% esterified) Methoxylated pectic acid 75% esterified)

100 97

1 6

aa

14

80

25

53

58

14

100

Methyl esterification of pectic acid was carried out according to the method of Jansen and Jang (1946). The methoxyl group in pectic acid is measured by the method of Wood and Siddiqui (1971).

Toyopearl 650, and Toyopearl HW-55s. The enzyme is isolated as a homogeneous protein. PPase-T is purified from the culture filtrate of T . sanguinea IF0 6490. The organism is maintained on agar slants containing 2% glucose, 0.5% yeast extract, and 0.5% peptone, pH 5.5. For the study of enzyme production, the organism is cultivated aerobically in 10 liters of a liquid medium containing 1.5% soybean flour, 3.25% KH,PO,, and 0.75% K,HPO, in a 14-liter fermentor at 35°C for 72 hr with aeration (at a flow rate of 5 literdmin), and occasionally Silicone KM70 is added as an antifoaming agent. The pH is adjusted to 5.7 during cultivation. The enzyme is purified by column chromotography on DEAE-toyopearl650, CM-Toyopearl 650, Butyl-Toyopearl 650, and Superose-12, and is isolated as a homogeneous preparation. 3. Properties of PPase-C and PPase-T

Although PPase-C and -T are B-type PPases, they have different molecular properties and substrate specificities. PPase-C has an apparent molecular weight of 30,000 (by SDS-PAGE) with an isoelectric point of around pH 9.0. Contrary to this, PPase-T is a protein with a molecular

2 6%

TAKUO SAKAI ET AL.

weight of 55,000 (by SDS-PAGE) and an isoelectric point of around pH 8.1.. Amino acid compositions of the enzymes are also different as shown in Table XIV. PPase-T are rich in aspartic acid, serine, and glycineI, whereas PPase-C are rich in glutamic acid, serine, and glycine. Some properties of PPase-C and -T are compared in Table XV. 4. Activity of PPase-C and PPase-T on Various Protopectins

PPase-C and -T act on protopectin from various citrus fruit peels and other plant tissues, releasing pectin (Table XVI]. However, the pattern of pectin-releasing activity is different. PPase-C has high activity toward protopectins from sugar beet pulp and apple, but PPase-T has potent activity on the protopectin of peels of hassaku and Valencia orange, and carrot tuber, but less activity toward sugar beet and apple. These results indicate that these two enzymes react with different sites in protopectin. TABLE XIV AMINOACIDCOMPOSITIONS OF PPASE-CAND PPASE-T Amino acid residuesa per molecule of protopectinase Amino acid

PPase-C

PPase -T

Lysine Histidine Arginine Tryptophan Aspartic acid Threonine Serine Glutamic acid Pro1ine Glycine Alanine Half-cystine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine

7 7 2 2 24 14 71

13 6 16 11 69

36 50

48

38

8

21

60 24

61 44

2

4

10

34

3 6 7

8 34

5

19 15

4

35

Calculations based on a molecular weight of 30,000 for PPase-C and 55.000 for PPase-T.

269

PECTIN, PECTINASE, AND PROTOPECTINASE TABLE XV PROPERTIES OF PPASE-CAND PPASE-T Properties

PPase-C

PPase-T

30,000 9.0

55,000 8.1 4.0 50

~

Molecular weight Isoelectric point (pH) Optimum pH (at 37°C) Optimum temperature (“C) Inhibitor pH stability Thermostability

6.0 60

Agt, Fe3+,Hgzt, MnZt, SnZt

Agt, CaZ+

5-9

2-6

-60°C

-60°C

The properties of the pectin extracted from lemon peel with PPaseT are shown in Table XVII. Its molecular mass is higher than that of pectin extracted by acid treatment. The mechanism of the release of pectin from protopectin by the action of PPase-C and -T has been studied by T. Sakamoto and T. Sakai (unpublished results). In that study, the polysaccharides that can be degraded by PPases have been isolated from sugar beet pulp.

TABLE XVI OF ACTIVITIES OF PPASE-CAND PPAsE-TO COMPARISON ~~

Relative activity (YO1

Origin of protopectin

PPase-C

PPase-T

Lemon peel (Citrus limon) Hassaku peel Satsuma mandarin orange peel (C. unshiu) Valencia orange peel (C. sinensis) Sugar beet pulp Apple Burdock (gobou) root Carrot root Radish tuber Watermelon peel

100 71 294 550

100 254 64 172 30 49 53 118 74 87

a

2200 1300 94 13 419 0

Reactions were done at 37°C.pH 6.0 (PPase-C) and pH 5.0 (PPase-T).

270

TAKUO SAKAI ET AL. TABLE XVII SOMEPROPERTIES OF PECTINFROM LEMONPEEL BY PPASE-T LIBERATED Relative viscosity of 0.1% solution Methoxyl group (%) Esterified carboxyl group (%) Galacturonic acid (%) Molecular weight Gel strength (“)b

1.80 59.7 79.4 74.1 132

208

Determined by the method of Christensen (1954). Determined by the SAG method of the IFT Committee (1959).

5. Reaction Mechanism of PPase-C

a. Preparation of Polysaccharides to Act as Substrates of PPase-C. Polysaccharides to be degraded by PPase-C are prepared as follows. First, a mixture containing 100 g of sugar beet pulp and 1 liter of 2.5% Ca(OH), solution is heated at 100°C for 1 2 hr. Then the pulp is removed by filtration through cloth and the filtrate is centrifuged. Next, the pH of the resultant clear solution is brought to 4.0 with acetic acid, and the precipitate that forms is removed by centrifugation. The supernatant is concentrated, and crude polysaccharides are obtained as a precipitate by the addition of 10 vol of ethanol. The precipitate is washed with ethanol three times. By dehydration under reduced pressure, polysaccharides are obtained as sticky yellow fibers. The crude polysaccharide preparation that dissolved in distilled water is put on a DEAE-Cellulofine AH column (3.2 x 34 cm) equilibrated with distilled water, and eluted with distilled water at a flow rate of 40 ml/hr. The fractions abundant in polysaccharides are collected and concentrated under reduced pressure. The concentrated solution is chromatographed with a Toyopearl HW-55s column (2.5 x 92 cm) at the flow rate of 18 ml/hr. The fractions containing polysaccharides that can be degraded by PPase-C are pooled, concentrated, and collected by precipitation with ethanol, giving PO-I. For the isolation of the smallest polysaccharide that can be degraded by PPase-C, PO-I is digested with purified a-L-arabinofuranosidase from Aspergillus niger K1 (Tagawa and Kaji, 1988b) in 20 mM citratephosphate buffer, pH 4.0, at 4OoC. The reaction is monitored for the increase in the reducing end of PO-I. When the rate of hydrolysis slows (when about 30% of the PO-I is hydrolyzed), the mixture is cooled to 20°C and incubated at this temperature for 1 2 hr more. The reaction

PECTIN, PECTINASE, AND PROTOPECTINASE

271

mixture is then heated for 5 min at 100°C to inactivate the enzyme, and ethanol is added to the final concentration of 80%. The precipitate is collected and dissolved in hot water, and the solution is left at 2°C for 24 hr. for reprecipitation. The polysaccharide PO-I1 is obtained by lyophilization. The properties of PO-I and -11 are listed in Table XVIII. PO-I is very sticky and soluble. PO-I1 is a white powder sparingly soluble in cold water; it is soluble in boiling water. Both PO-I and -11 are readily hydrolyzed in a hot, weakly acidic solution.

b. Identification of PO-II. The chemical structure of PO-I1is identified by HPLC analysis of its hydrolysate, GLC and GC-MS with methylation, and NMR. Methylation experiments with GLC are: PO-I1 is methylated by the use of dimethyl sulfinyl carbanion (Me,SC) in dry dimethyl sulfoxide (DMSO) and CH,C1, according to the Hakomori procedure (1964). DMSO is prepared by the method of Rauvala (1979). The permethylated product is hydrolyzed, and the sugars released are converted into their alditol acetates and analyzed by GLC. GLC is done with a Hewlett Packard Component HP 5890A chromatograph (equipped with a flame ionization detector) with a capillary column (SP 2330, 0.32 mm x 30 m). The column temperature is programmed to increase from 160 to 220°C at the rate of 4"C/min, and the injector and detector temperature is 180°C. The flow rate of the carrier gas, He, is 25 ml/min. In the GLC analysis, two sharp peaks are detected at around the retention times of 8 and 12.5 min. The relative retention times (Table XIX) of these compounds and authentic samples to the retention time of 1,5-di-O-acetyl-2,3,4,6-tetra-O-methyl-~-glucitol are calculated by RI

=

(R2 - R1)

X

(T - Tl)/(T2 - T1)

+ R1,

where RI is the retention index of the sample; T is the retention time of the sample; R 1 and R2 are retention indices of the solvent (acetone)

TABLE XVIII PROPERTIES OF Po-1 AND P0-11 Properties

Sugar components Solubility in water [a]?(10 mg/ml in water) Intrinsic viscosity [ T ]

PO-I

PO-I1

L-Arabinose (96%) o-Galactose (4%) 195 mg/ml -164' 19.8 g/cm

L-Arabinose (100%) 3.2 mg/ml at 25°C - 146' 23.2 g/cm

2 72

TAKUO SAKAI ET AL. TABLE XIX

RELATIVERETENTIONTIMESON GAS CHROMATOGRAPHY OF PARTIALLY METHYLATED ALDITOL ACETATE Sample

Relative retention time"

1,4-Ac-2,3,5-Me-~-arabinitol~ 1,5-Ac-2,3,4-Me-~-arabinitol~ 1,4,5-Ac-2,3-Me-~-arabinitol~ Major peak Minor peak

0.69 0.79 1.13 1.13 0.69

ORelative to 1,5-di-0-acetyl-2,3,4,6-tetra-O-methyl-o-glucitol. Ac, acetyl; Me, methyl. Authentic samples were synthesized chemically.

and 1,5-di-O-acetyl-2,3,4,6-tetra-O-methyl-~-glucitol, respectively; and T1 and T2 are retention times of the solvent (acetone) and 1,5-di-0acety~-2,3,4,6-tetra-O-methyl-~-glucito~, respectively. As shown in Table XIX, the minor peak in GLC is identified as auand the major thentic 1,4-di-0-acetyl-2,3,5-tri-O-methyl-~-arabinitol, peak corresponds to authentic 1,4,5-tri-O-acety1-2,3-di-O-methyl-~arabinitol. Analysis of these peaks by GC-MS supports the determination above (data not shown). The major compound may be formed from the 1,s-linkage of L-arabinose, and the minor compound may be derived from the nonreducing terminal arabinose of PO-11. The molar ratio of these compounds is about 1 : 25, and PO-I1 is found to be a homopolymer composed of about 2 5 residues of L-arabinofuranose. Spectral data of the 'H- and I3C-NMRof PO-I1 are given in Table XX. The signal for H-1 is assigned to 6 4.76 because of the position of the resonance of H-2 and H-4 (at 6 3.90 3.78 ppm), H-3 (at 3.70 3.56 ppm), and H-5 (at 6 3.56 3.44 ppm). The same results are obtained by 'H-I3C correlation spectroscopy (COSY; Fig. 30). The coupling constant of the doublet for H-1, ] = 1.9 Hz, suggests that the glycosidic linkage is in the a-configuration. The assignment of the resonances for H-1, -2, -3,-4, and -5 can be obtained in a straightforward way from the 'H-'H COSY spectrum. There is only one cross-peak of H-1 with H-2 at 6 3.8 ppm, and no additional peak of H-1 is found. Doublet signals at 6 5.38 and 5.21 ppm make cross-peaks with H-2 and H-3, respectively. These findings indicate that the doublet signals at 6 5.38 and 5.21 are for the -OH of H-2 and H-3. From data obtained by 13CNMR, C, does not seem to have any -OH groups. These results show and PO-I seems to be that PO-I1 is identified as being a-l,5-~-arabinan,

-

-

-

PECTIN, PECTINASE, AND PROTOPECTINASE

273

TABLE XX 'H- AND 13C-NMR CHEMICAL SHIFTDATAFOR PO-I1 [6, ppm, in (CD,),SO]

8

Atom

'H-NMR 5.38 5.21 4.76 3.90-3.78 3.70-3.56 3.56-3.44

0h-2 0h-3 H-1 H-4, H-2 H-3, H-5, H-5b

I3C-NMR 109.46 83.02 82.79 78.60 68.38

c-2 c-4

c-1

110

c-1 c-2 c-4 c-3 c-5

100

90 6 l3C (PPR)

80

70

60

FIG.30. I3C-lH COSY spectrum of PO-I1 in (CD,),SO, at a 'H frequency of 300 MHz.

274

TAKUO SAKAI ET AL.

L-arabinan. L-Arabinofuranose units are attached to a-l,s-~-arabinan, from its properties in Table XVIII (Tagawa and Kaji, 1988a). c. Reaction of PPase-C with L-Arabinan. Arabinan seems to be in the form of L-arabinan in sugar beet protopectin. To study the reaction pattern of the enzyme with L-arabinan, samples from a reaction mixture containing 1 mg of substrate, the enzyme solution, and 150 pl of 100 mM AcB, pH 6.0, are withdrawn for the identification of intermediate and end products by HPLC at from 1 min to 4 hr of incubation at 60°C. The reaction is stopped by addition of 25 mM NaOH and the reducing-end sugar product is labeled with 3Hby treatment with NaB3H, in 25 mM NaOH, according to the method of Takasaki and Kobata (1974). In an early stage of the reaction, saccharides with high molecular mass appear; they are then degraded to arabinose. But arabinose is converted to arabinobiose and to arabinotriose upon further reaction. HPLC shows that the enzyme hydrolyzes L-arabinan at random to form arabinose and that the enzyme also has arabinosyltransferase activity to produce arabinobiose or arabinotriose (Fig. 31). d. Postulated Mechanism of the Protopectin Reaction of PPaseC. PPase-C catalyzes the degradation of a-I,5-~-arabinanand Larabinan, which seems to have a main structure consisting of a-1,5-~arabinan, to which L-arabinofuranose units are attached at position 3 in the a-configuration to form one-unit side chains from sugar beet pulp in a random manner. It is therefore an enzyme in the same category as arabinan endo-l,!j-a-~-arabinase (EC 3.2.1.99). However, endo-1,5-aL-arabinase from B. subtilis (the enzyme catalyzes the hydrolysis of arabinooligomer to form L-arabinose and arabinobiose) (Kaji and Saheki, 1975) shows low PPase activity. Thus, PPase-C is novel in the aspect of having potent protopectinase activity. Albersheim (1975) has mentioned that pectin (rhamnogalacturonan) is attached at the reducing terminals of the arabinogalactan molecule in sycamore cells; and arabinogalactan seems to consist of a chain of arabinose coupled to another chain of galactose (Fig. 32). In sugar beet pulp, rhamnogalacturonan connected to neutral sugar side chains (hairy region) in pectin is degraded by p-l,4-~-galactanase and forms rhamnogalacturonan and arabinan with a small amount of galactose residues. This may indicate that the structure of the hairy regions of pectin in sugar beet protopectin is substantially the same as in sycamore cells, with rhamnogalacturonan attached to cell wall constituents with arabinogalactan interposing, as shown in Fig. 32. PPase-C splits the a-1,5L-arabinofuranoside linkage of the arabinan region in arabinogalactan,

2 min

~

0 D rnin

5

-

10

15

,

20

Retention time (min) FIG.31. Analysis of the mode of reaction of PPase-C toward L-arabinan by HPLC.

276

TAKUO SAKAI ET AL.

Rhamnogalacturonan (Pectin

0 0 FIG.32. Postulated structure of arabinogalactan in sycamore cell [from Albersheim, 1975). G , D-gahCtOSe; A, L-arabinose.

which attaches pectin to the cell wall constituents, so that PPase-C releases pectin. 6. Reaction Mechanism of PPase-T PPase-T can degrade rhamnogalacturonan in sugar beet protopectin, releasing pectic substance. The smallest polysaccharide (called as SPS) that can be the substrate of PPase-T is prepared from sugar beet pulp by the extraction with NaOH and digestion with a-L-arabinofuranosidase, a-L-arabinase, and @1,4-~-galactanase. The reaction products of SPS with PPase-T are isolated by chromatography on DEAE-Toyopearl 650M and Toyopearl HW40-S columns and are analyzed by labeling of reducing ends with NaB3H, and 13C-NMR spectroscopy. The results indicate that PPase-T cleaves galactopyranosyluronic-rhamnopyranosyl linkages in SPS. The outline of the isolation of the polysaccharide serving as the substrate for the PPase-T reaction is given here. a. Enzyme Activity toward Pectic Substances from Sugar Beet Pulp. Pectic substances prepared from sugar beet pulp have been studied in detail (Rombouts and Thibault, 1986a,b; Guillon and Thibault, 1989; Guillon et al., 1989),and its hairy regions were found to be mainly galacturonic acid, arabinose, galactose, and rhamnose and a small amount of fucose, glucose, mannose, and xylose. To investigate the distribution of the reaction site of PPase-T in plant tissue, pectic substances are extracted sequentially by water, sodium hexametaphosphate, hot HC1, and cold NaOH from sugar beet pulp, according to the methods of Pilnik and Voragen (1970),Barbier and Thibault (1982), and Rombouts and Thibault (1986a). (These extracts are tentatively called WSP, HMP, HP, and OHP, respectively.) The activities of PPaseT on these pectic substances are summarized in Table XXI. The enzyme catalyzes the hydrolysis of HP and OHP, but not that of WSP and HMP.

277

PECTIN, PECTINASE, AND PROTOPECTINASE TABLE XXI SUGAR

COMPOSITION OF PECTINS FROM SUGAR BEETPULP AND HYDROLYSIS ACTIVITY BY PPAsE-TO

Pectins

GalA

Rha

WSP HMP HP OHP

68 80 54

2 1

Enzyme activity (U/ml)

Gal

Ara

7 4 15

22

0

13

24

10

0 17 45

(mol%)

56

6 8

22

Small amounts of sugars are neglected.

b. Preparation of Alkali-Soluble Pectin (ASP). Polysaccharides that can be degraded by PPase-T are prepared from sugar beet pulp by being heated in an alkaline condition (the enzyme is most active on OHP). Sugar beet pulp (50 g) is mixed with 3 liters of 0.1 N NaOH, and the mixture is heated at 100°C for 1hr. The slurry is filtered through gauze and the filtrate is centrifuged. Then 3 vol of acetone is added to the resultant clear solution. The precipitate formed is dissolved in distilled water. The solution is treated batchwise with Dowex-50W (H'-form) to remove cationic ions associated with pectic substances and passed through a Cellulofine column equilibrated with distilled water to remove pigments. The effluent is placed on a DEAE-Cellulofine AH column (6 x 28 cm), equilibrated with 50 mM AcB, pH 5.0, and washed with the same buffer. The bound polysaccharides are eluted with 1M AcB, pH 5.0. The fractions containing polysaccharides, which served as substrates of PPase-T, are collected, concentrated, dialyzed against distilled water, precipitated by the addition of 3 vol of ethanol, and finally lyophilized, giving ASP. The ASP is assumed to contain the hairy region as its main component because the homogalacturonan region is unstable in alkaline treatment, decomposing into oligogalacturonic acid. c. Preparation of SPS. The ASP is completely digested by a mixture of a-L-arabinofuranosidase and a-L-arabinase in 20 mM AcB, pH 5.0, and the reaction mixture is chromatographed by gel filtration with a Superose-12 column (Fig. 33). The fractions containing polysaccharides that can be degraded by PPase-T are collected, giving ASP-A. ASP-A is treated by &1,4-~-galactanasein 20 mM AcB, pH 5.0, and purified

TAKUO SAKAI ET AL.

278 16

1

-

12

a

- 0.75 - 0.5 II - 0.25 I

8 -

I

4 -

I

! I'

EF

-

a m

E . cS

I

I

O

b

- 0.75

9 6 -

- 0.5

3 -

-

0.25

I I C

8

n 0,

-.-->

>.

,

CI

0

m

SPST

6 -

4 2 -

I 10

20

Elution volume (ml) FIG.33. Elution patterns of pectic substances degraded by various enzymes on Superose 12 for the isolation of SPS. (a) ASP; (b) reaction products of ASP with arabinofuranosidase and arabinase; (c)reaction products of ASP-A with galactanase; (d) reaction products of SPS with PPase-T.

by chromatography on the same column (Fig. 33), giving SPS. PPaseT cleaves SPS to reduce its molecular size, giving SPST (Fig. 33). SPS is used as the substrate of PPase-T. d. Identification of Reducing Ends of Reaction Products of SPS and PPase-T. For identification of the reducing end sugar residues formed in the PPase-T reaction, the following experiment is performed. The

PECTIN, PECTINASE, AND PROTOPECTINASE

279

reaction mixture, containing SPS and PPase-T, is incubated at 45°C for 1 hr, and the newly formed reducing groups are labeled with tritium by treatment with NaB3H, in 25 mM NaOH according to the method of Takasaki and Kobata (1974). The poly- and oligosaccharides labeled with tritium are hydrolyzed with 1 N HC1 at 110°C for 2 hr, and the resulting monosaccharide labeled with tritium is identified by HPLC. HPLC of the reducing-end sugars is shown in Fig. 34. The polysaccharides formed by PPase-T have galacturonic acid at their reducing ends. PPase-T hydrolyzed neither poly- nor oligogalacturonic acid. e. Isolation and Characterization of Reaction Products of SPS and PPase-T. For checking of the nonreducing-end sugar residues in products of the PPase-T reaction, the reaction products of SPS and PPase-

C

I

I

1

I

Retention tlme (min) FIG.34. Analysis of the hydrolyzed product of SPS with PPase-T. a, galacturonate (alditol); b, rhamnitol; c, galactitol; d, decarboxylated galacturonate (alditol).

280

TAKUO SAKAI ET AL.

T are isolated. SPS is completely hydrolyzed by PPase-T, and the hydrolysates are chromatographed on a DEAE-Toyopearl column (Fig. 35). The unbound and bound fractions, which are called SPST-1 and SPST2, respectively, are collected and subsequently chromatographed on a Toyopearl HW40-S column. The results show that the molecular size of SPST-1 is smaller than that of SPST-2. The I3C-NMR spectra of SPS and SPST-1 are compared using the results published by Colquhoun et al. (1990) (Fig. 36 and Table XXII). The 13C-NMRspectrum of SPST1 is almost identical with that of oligosaccharide fraction D [a oligosaccharide obtained by enzymatic degradation of the modified hairy regions of apple pectin with a rhamnogalacturonase (Schols et al., 1990a)], and the structure of SPST-1 seem to be that shown in Fig. 37. The resonances at 94.88 ppm (aC-l), 98.78 ppm (PC-l), 76.45 ppm (PC-3), 79.92 ppm (aC-4), and 77.33 ppm (PC-5) are assigned to the galacturonic acid residues on the reducing ends of SPST-1 (residue A in Fig. 37). These findings are evidence that PPase-T produces saccharides with galacturonic acid as the reducing end residues in the experiment with radioisotopes. The two peaks at 103.07 and 103.22 ppm arise in

15

30

45

60

Fraction number FIG.35. Chromatography of reaction products of SPS with PPase-Ton DEAE-Toyopearl. (a] Before the reaction; (b) after the reaction.

-

L-

e - C - 4 D

/

C-5A0 C-3A0

P

A

-u U -3

00

282

TAKUO SAKAI ET AL. TABLE XXII CHEMICAL

Unit GalApO ffA PA a-Rhap B B'b

a-GalAp C a-Rhap D D'" P-Galp E,F

SHIFTS (ppm) OF l3C RESONANCES FOR SPST-1

c-1

C-2

94.88 98.78

70.64

101.08 101.08

78.70 79.30

100.06 100.42

70.64

103.22 103.07

73.10 72.88

106.03

74.35

c-3

c-4

c-5

C-6

79.92

73.39 77.33

177.5 176.85

74.64 83.11

71.70 70.33

19.28 19.53

78.96 79.05

74.20

72.77 72.59

74.89 83.51

71.41 69.89

19.37 19.59

75.37 75.44

71.32

77.80

63.59

76.45 72.07 72.07

177.5

Reducing end. Gal-substituted Rha at position 4. Gal-substituted terminal Rha at position 4.

the region from 94 to 108 ppm for C-1 resonances in SPST-1. These peaks are identical to those of a-rhamnopyranose on the nonreducing ends of oligosaccharide fraction D reported by Colquhoun et al. (1990). The other peaks, at 73.10, 72.88, 74.89, 83.51, and 69.89 ppm, which are assigned to C-2D, C-ZD', C-4D, C-4D', and C-5D', are also present in SPST-1. From these results, the nonreducing-end sugar residue of SPST-1 is identified as a-rhamnopyranose. PPase-T hydrolyzes galactopyranosyluronic-rhamnopyranosyl linkages in protopectin to release

Reaction site of PPase-T a

D, D'

ByB'

A

a-Rhap(l-*4)-a-GalpA-(li 2)-a-Rhap(l--+ 4)-a-GalpA

r"

F (P-Galp)

C

+

r" E (B-GaW

FIG.37. The postulated structure of SPST-1.

PECTIN, PECTINASE, AND PROTOPECTINASE

283

pectin. The results shows that PPase-T has a reaction mechanism similar to that of the rhamnogalacturonase reported by Schols et al. (1990a). The resonances of C-1 and -4 in the (1+4)-linked a-GalA units in SPST-1 are 1.5 ppm lower than those of the same unit in oligogalacturonate. This indicates that the two neighboring sugars of the 1,+linked GalA residues in SPST-1 are not a-D-GalA but a-L-Rhap residues (Colquhoun et a]., 1990). This result is consistent with the structure shown in Fig. 37. Rhamnogalacturonan I (McNeil et a]., 1982; Lau et al., 1985) isolated from sycamore cells and modified hairy regions (Schols et a]., 1990b) from apple have repeats -4)-a-~-GalpA-(1+2)-a-~-Rhap-(l+. The ASP prepared form sugar beet pulp seems to have a backbone of repeats of +4)-a-~-GalpA-(1+2)-a-~-Rhap-(l+. SPST-2, which is separated from SPST-1 by DEAE chromatography, is larger than SPST1 in molecular size and galactose content. SPST-2 is not hydrolyzed by PPase-T and may resist the enzyme because it is rich in galactan side chains. The enzyme activity of PPase-T toward ASP, ASP-A, and SPS is 24, 33, and 36 U/ml, respectively. These results suggest that PPaseT hydrolyzes -(GalA-Pha)- units without long side chains. The reaction product that corresponded to SPST-1 is produced in the reaction of ASP with the enzyme. This indicates that ASP has -(GalA-Rha)- units without long side chains.

D. APPLICATIONS OF PROTOPECTINASE 1. In Pectin Production

Pectin is useful in the manufacture of food, cosmetics, and medicine; it is an industrially important substance, produced on an industrial scale. Citrus peel, a by-product of the citrus processing industry, is a suitable source of pectin. In industrial production, pectin is extracted by placing the peel in vats of water, bringing the mixture to a boil as a slurry, and adding concentrated hydrochloric, sulfuric, nitric, or other acids to adjust the pH to about 2.0. Filtration of the extract is a tedious process because the extract, containing pectin and disintegrated peel, is both corrosive and viscous. The peel of mandarin orange (including Citrus unshiu, which accounts for more than 80% of the citrus fruit produced in Japan) is not suitable as a raw material in this process because the peel is fragile and becomes pasty when heated, which prevents the separation of pectin from the residues (Miyazaki and Terada, 1974). Thus, in Japan, pectin is not manufactured chemically, although nearly

284

TAKUO SAKAI ET AL.

5 x i05 tons of citrus peel, containing about 5% pectin on a freshweight basis, is produced each year. Sakai and Okushima (1980) have tried pectin production with C. unshiu peel as the raw material by developing a new microbial method by which pectin can be enzymatically extracted form citrus peel without maceration of the peel. The outline of the process is: T. penicillatum SNO 3, which is a PPase-S producer, grows well in an extract of citrus peel as the sole nutrient source. The organism must assimilate the water-soluble carbon and nitrogen compounds in the peel, since nothing is present other than citrus peel. The amount of pectin extracted depends on the concentration of peel in the medium. The peel/water ratio of 1 : 2 1 : 3 is suitable. Pectin is extracted effectively between 25 and 30°C, although the microorganism grows well between 25 and 37°C. At 30°C, pectin begin to appear after 5 hr and the amount increases with fermentation time; after 20 to 25 hr, the amount of pectin extracted reaches a maximum (Fig. 38). During fermentation, a small amount of pectin is extracted by water without action of the microorganism; however, a much greater amount of pectin can be extracted upon inoculation. The increased amount of pectin extracted by the microorganism is termed microbial-soluble pectin in Fig. 38. By this method, almost all of the pectin in the peel can be extracted. Physical and chemical properties of isolated pectin are compared in Table XXIII with those of acid-extracted and commercial pectin. The

-

I 1

/--P

0

*

Y

Y

Y

0

4

8

12

16 20

24

Fermentation period (hour) FIG.38. Time course of pectin extraction during fermentation.0 .water-soluble pectin; A , microbial-soluble pectin; A, total pectin extracted; 0, relative viscosity of pectin (0.3% solution at 37°C).

PECTIN, PECTINASE, AND PROTOPECTINASE

285

TABLE XXIII

SOME PROPERTIES OF PECTIN Pectin type From C. unshiu peel ~

Properties

~~~

Commercial (from lemon)

Extracted by fermentation

Extracted by acid-heat

1.53 9.24 63.1 85.0 5.7 3.96 102

1.46 8.58 73.8 68.2 23.2 3.24 105

1.23 9.13 66.1 80.3 10.5 4.34 50

40.86 5.76 0.80

40.27 5.77 0.61

38.27 5.40 0.41

Relative viscosity of 0.1% solution Methoxyl group" (%) Esterified carboxyl group" Galacturonic acid" (%] Neutral sugara (yo) pH of 0.5% solution Molecular weightb ( x - 3 ) Element analysiso C H N

Values expressed on an ash- and moisture-free basis. Calculated by the equation of Smit and Bryant (1967).

pectin produced by this method is not very different from the two other kinds of pectin, except that this pectin contains more neutral sugar. Peels or segment covers of various citrus fruits are good raw materials; vegetables such as carrots, wax gourds, and radishes ar poor as raw materials (Table XXIV). The new pectin bio-production system established by Sakai and Okushima (1980) is illustrated in Fig. 39. In this system, peels are washed, if necessary, and suspended in sterilized water in a fermentor, to which a seed culture of the microorganism is introduced (corresponding to from 3 to 5% of the volume of the fermentation broth) from a seed tank. After 15 to 20 hr of fermentation at 25 to 30°C, the residual peels are filtered off, and the resultant filtrate is passed through another filter to remove the microbial cells. The filtrate is concentrated and the pectin is precipitated in ethyl alcohol, collected, and dried. Using this procedure, 20 to 25 kg of pectin is obtained per ton of mandarin peels, which are a by-product of the citrus processing industry. 2. In Other Sectors a. Isolation of Plant Protoplasts.

PPases can be used for the isolation of plant mesophyll protoplasts. Mitsui et al. (1990) have shown that

286

TAKUO SAKAI ET AL. TABLE XXIV

YIELDSOF PECTIN EXTRACTED BY T. penicillaturn SNO 3

FROM VARIOUSORIGINS'

Pectin extracted (g/100 g) Origin Novel orange (Citrus sinesis) Peel Segment cover Grape fruit Peel Segment cover Mandarin (Citrus unshiu) Peel Segment cover Mandarin (Citrus natsudaidai) Peel Segment cover

Water solubleb

Microbial solubleC

Total

0.4 0

2.5 2.7

2.9 2.7

0.2 0

2.5 2.1

2.1

0.3 0

2.6 1.8

2.9

0.1 0

2.1

2.2

1.9

1.9

0.9

1.7 1.6 0.1

2.6

2.7

1.8

Lemon Peel Segment cover Apple (Kogyoku) Radish Carrot Wax gourd

0 0.1

0 0 0.4

0.2

1.6 0.2 0.2

0.1

0.1

0.3

0.7

Fermentation was carried out at 30°C for 12 hr. Extracted without inoculation. Extracted by fermentation.

PPase-S can be used for the isolation of mesophyll tissues of shoot such as Tagetes minuta, Brassica napa, Raphanuys sativus, Lactuca sativa, Triticum aestivum, SecaJe cereale, Hordum vulgare, Panicum crusgalli, Avena sativa, Zea mays, and Oryza sativa. These plant cells often survive and regrow to give a plant. PPase-S is outstanding for the preparation of protoplast of monocotyledonous cells.

b. Production of Single-Cell Foods. PPases are used for processing single-cell preparations from vegetables for use in foods. Vegatables processed to give a single-cell preparation might provide a new kind of ingredient for use in foods. Vegetable cells isolated by enzymatic methods often survive with intact cell walls. Such individual cells can be regrown to give a plant if incubated under suitable conditions, because their biological functions are undamaged by such isolation

PECTIN, PECTINASE, AND PROTOPECTINASE

287

Evaporator

Citrus peel

Dried yeast

cell

Pectin

FIG.39. Schematic diagram of the biochemical manufacturing process of pectin from citrus peel.

procedure. The flavors, pigments, and nutrients such as vitamins present in the original vegetable are likely to be preserved in single cells prepared in this way. T. Nakamura and T. Sakai (unpublished results) found that PPases can be used to produce single cells from vegetable tissues. The PPases from T. penicillatum, A. awamori, and B. subtilis are useful for this purpose. The constituents of the single cells are more stable than those in the preparation obtained by mechanical tissue disruption. ACKNOWLEDGMENTS

Tatsuji Sakamoto was a researcher in the laboratory of Professor Erick. J. Vandamme during 1990-1991. E. J. Vandamme thanks the Belgium National Science Foundation (WFWO) and the University of Gent for grants related to his research on fermentation, biochemistry, and genetics on useful microbial enzymes and bio-active metabolites.

288

TAKUO SAKAI ET AL.

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Physicochemical and Biological Treatments for Enzymatic/Microbial Conversion of LignocelIulosic Biomass PURNENDU

GHOSHAND AJAYSINGH

Biochemical Engineering Research Centre Indian Institute of Technology New Delhi 110 016, India

I. Introduction 11. Structure of Lignocellulosic Biomass 111. Physical Treatment

IV.

V.

VI.

MI.

A. Mechanical B. Radiation Thermal Treatment A. Autohydrolysis B. Steam Explosion C. Hydrothermolysis Chemical Treatment A. Alkali B. Acid C. Oxidizing Agents D. Gases E. Cellulose Solvents F. Solvent Delignification Biological Treatment A. Potential Microorganisms B. Biodelignification C. Biopulping D. Biobleaching E. Kraft Effluent Treatment Epilogue References

I. Introduction

Microbial utilization of the inexhaustible lignocellulosic biomass for the production of industrial chemicals, liquid fuels, protein-rich food and feed, and preparation of cellulose polymers (Fig. 1)is an attractive approach to help meet energy and food demands. The feasibility of several lignocellulosic materials for such purposes has been studied around the world, depending on their availability. Lignocelluloses are the most abundant natural materials present on the earth. Regardless of source, lignocelluloses contain cellulose, hemicellulose, and lignin 295 ADVANCES IN APPLIED MICROBIOLOGY. VOLUME 39 Copyright 0 1993 by Academic Press, Inc. All rights of reproduction in any form reserved.

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Dehydration

I

c c

Cellulose-Glucose

Hydrogenation

Iserization-

Polyesters Res i ns Drugs

So1 vents Organic acids SCP

Gums Sorbi to1 Glycerol Sweeteners Vitamins Fructose syrup

So1 vents Organic acids Fermentation

Xyl i to1 Sweeteners

BIOHASS -

.Hemicellulose-Xylose

Dehydration

-Li gni n

[

-

Binders Activated carbon Solid fuels Chemicals Fuels Pheno1 s Rubber Vani 1 1 ine

c

Furfural Amino acids Drugs Polyurethene Foams Resins

Phenolic Res i ns Polyesters Adhes i ves

FIG.1. Useful products derived from lignocellulosic biomass

as major components (Table I). Cellulose consists of a p-1,4-linked glucose polymer; hemicellulose contains complex polymers of pentoses and hexoses; and lignin is a complex heterogenous polymer of phenylpropanoid units. Natural cellulose is present in association with hemicellulose, lignin, and extractives. The enzymatic saccharification of native cellulose is very slow. The characteristics of the cellulosic substrate such as crystallinity, lignin content, and specific surface area are related to cellulose saccharification. Cellulase enzymes readily degrade easily accessible,

297

LIGNOCELLULOSIC BIOMASS TABLE 1

COMPOSITION OF SELECTED LIGNOCELLULOSIC BIOMASSMATERIALS Percentage on a dry weight basis ~~

Substrate Agricultural residue Bagasse Corn cobs Groundnut shells Oat straw Rice straw Wheat straw Cotton straw Forest residue Aspen Birch Pine Red maple

~

Cellulose

Hemicellulose

Lignin

33 42 38

30 39

29 14 16

32 30

36 16 24 24

42

12

50

28 33

41

40 41 39

10

33

11

13 18 15

Ash

4

2 5 12 12 10 6

15

1

21 27 23

4 8 2

amorphous cellulose as compared to crystalline cellulose due to enzyme transport limitation imposed by the closely ordered lattice of the cellulose molecule (Norkrans, 1950). 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 cellulase (Klyosov et al., 1986).Hemicellulose present in lignocellulosic biomass appears to shield cellulose from enzymatic attack. Decreased particle size and increased available surface area result in an increase in cellulose digestibility. The extractives (resins, waxes, etc.) also interfere with cellulose hydrolysis because of their hydrophobic nature (Lipinsky, 1979). Pretreatment is, therefore, necessary for the effective utilization of lignocellulosic material. This can be achieved by several different methods, which in principle should cause disintegration, thereby creating a large surface area on which enzymes can work. Thus the intention of pretreatment is to open the structure of the lignocellulosic materials, making it accessible to the cellulolytic enzymes (Ghose and Ghosh, 1979). This is accomplished by (1) increasing the specific surface area, (2) removing lignin, or (3) solubilizing hemicellulose. Several approaches, including physical, thermal, chemical, biological or a combination of these, have been explored to meet the specific requirements which can be described as follows: 1. To obtain each of the polymeric components of the lignocellulosic material in maximum yield and purity;

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2. To increase the accessibility of cellulosic carbohydrates to cellulase enzymes with the goal of obtaining glucose for fermentations; 3. To produce low-cost sugars (hexoses and pentoses) for use in biotechnological routes to fuels and chemicals; 4. To yield lignin components which can be transformed further to several other chemicals; 5. To use low-cost lignocellulosic biomass in processes to produce protein-rich feed for ruminants; 6. To improve pulping and bleaching processes in the pulp and paper industry.

Considerable developments in various pretreatment techniques have been made during the last decade, and in using these processes substantial amounts of lignin and hemicelluloses can be degraded to products extractable with water, solvents, or alkaline solutions. Recent advances in various pretreatment techniques and their effects on enzymatic hydrolysis and bioconversion are discussed in this article. II. Structure of Lignocellulosic Biomass

The wood cell is a multilayered structure consisting of an external microfibril primary layer and a secondary wall containing three sublayers: S1, S2, and S3 (Cowling and Kirk, 1976). The concentration of cellulose is highest in sublayer S2 and diminishes toward the middle lamella. The concentration of hemicellulose and lignin is maximum in the middle lamella and decreases toward the lumen. The concentration of lignin in the middle lamella is about 70% in softwood and 90% in hardwood, with the major part (70-80%) of the lignin being distributed within the secondary wall flange, 1954; Latham et a ] . , 1978). Wood cells may contain up to 90% fiber whereas only about 35-3996 of the cells in straw are fiber (Fan et a]., 1981). 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. The capillary void in wood includes cell wall capillary and gross capillary. Most of the cell wall capillaries are closed when lignocellulosics are free of water but open up when moisture is absorbed. When fully saturated with water they expand to about 200 A in diameter. The total surface area exposed in gross capillaries (2 x lo3 cm2 * g-’) is several orders of magnitude smaller than the total surface area exposed within the cell wall capillaries (3 x lo6 cm2 . gl). The penetration of cellulase enzymes to cell wall capillaries substantially increases the saccharification rate than if

LIGNOCELLULOSIC BIOMASS

299

cellulase is confined to the surface of the gross capillaries. A correlation between the porosity and the accessibility to cellulase enzymes has been demonstrated by Stone et al. (1969).They observed a linear relationship between the initial hydrolysis rate and the surface area that was accessible to the molecule with a diameter of 40 A (this corresponds to the diameter of the cellulolytic enzyme molecule). Cellulose contains crystalline and amorphous regions. The proportion of the crystalline fraction ranges between 50 and go%,and cellulose disruption operates both at intercrystalline and intracrystalline levels. Also, cellulose consists of regions of high and relatively low crystallinity so that different regions exhibit different susceptibilities during the progress of the enzymatic reaction (Nisizawa, 1973; Cowling, 1975; Fan et al., 1982). 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. The present concept indicates that lignin and cellulose form a mutually interpenetrating system that is largely physical in nature (Kaushik and Bisaria, 1989). The nature of the lignin-hemicellulose bond is suggested to be ether and 4-0-methylglucuronic acid ester to the a carbon of the lignin unit (Crawford, R. L., 1981). Through conventional means of isolation and cleavage it is very difficult to study the structure of lignins (Kaushik and Bisaria, 1989). There are several reasons for this complication: (1) lignins, unlike proteins, nucleic acids, and polysaccharides, do not have identical readily hydrolyzable linkages repeating at regular intervals; (2) they are irregular and have no precise chemica€ structure, but have a series of chemical groupings; (3) lignins are difficult to extract in an unmodified state, and (4) lignins cannot be degraded efficiently into monomer units because of C-C and diary1 ether type bonds which are difficult to cleave. The hydrolyzable linkages in lignin are suggested to be of two types: @-arylether and aaryl ether (Adler, 1977). Among these, the predominant @-aryl ether type bond is more resistant to cleavage (Sarkanen and Ludwig, 1977). Under mild hydrolytic conditions, the cleavage of the ether bond is exclusively restricted to those of the a-aryl ether type. The inter- and intramolecular bonds that exist between various constituents of lignocellulose are vital factors in the proper selection of a pretreatment method (Leisola and Fiechter, 1985; Kirk and Farrell, 1987). In addition to cellulose, the hemicellulose and lignins plant cell wall contains extraneous materials, including extractives and nonextractives (Ladisch et a]., 1983). Wood contains 0.4-8.3% extractives on a dry weight basis; agricultural residues contains even greater amounts (Janes, 1969).Extractives consist of fats, waxes, tannins, resins, essential

300

PURNENDU GHOSH AND AJAY SINGH

oils, alkaloids, starches, gums, and various other cytoplasmic constituents (Cowling and Merill, 1954; Hillis, 1962). Extractives can be removed by treating the substrate with water or neutral organic solvents like ethyl ether, acetone, ethanol, and benzene. The nonextractives make up 0.2-0.8% of the dry weight and include inorganic components such as silica, carbonates, oxalates, and noncellular substances (Janes, 1969). In agricultural residues, the nonextractives make up 10% of the dry weight (Ladisch et al., 1983). The separation of lignocellulose components is possible only when hydrogen bonds between the constituents and the hemicellulose-lignin ester cross-linking 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 (Millet et al., 1976) or bases (Lora and Wayman, 1978) are used. The breaking of lignin association may require the use of solvents with a different solubility parameter from those used as swelling agents for the carbohydrate (Lipinsky, 1981). Using lignolytic microorganisms, lignin can also be removed from lignocellulosic biomass. Ill. Physical Treatment

A. MECHANICAL Mechanical pretreatment methods include grinding, milling, and extrusion. These methods utilize shearing and impacting forces to yield a fine substrate possessing a low crystallinity index, thus enhancing its susceptibility to enzyme action (Fan et al., 1981). Using a fine substrate allows a higher slurry concentration, thus reducing the reactor volume. Reducing the crystallinity of cellulosic substances by milling was introduced by Krupanova (1963). Milling helps in the distribution of reactants throughout the material as well as reducing crystallinity. Milling at elevated temperatures is preferred. It has also been proposed that an increase in cellulose digestibility is because of decreased particle size and increased available surface area rather than a result of reduced crystallinity (Browning, 1963; Caulfield and Moore, 1974). Ball milling is an effective means of pretreatment (Mandels et a]., 1974; Millet et al., 1975). In addition to reducing particle size, ball milling disrupts the crystalline structure and breaks down the chemical bonding of long chain molecules. Millet et al. (1975) observed that the effectiveness of ball milling depends on the material to be processed; softwoods are the least responsive. Vibratory ball milling appears to be more effective than ordinary ball milling in translating energy input into

LIGNOCELLULOSIC BIOMASS

301

size reduction and alteration of cellulose crystalline structure (Millet et al., 1976). Vibratory milling enhances the susceptibility of spruce and aspen wood toward enzymatic hydrolysis (Pew, 1957). The differential speed two roller mill has been used in a variety of cellulosic substrates (Tassinari and Macey, 1977; Tassinari et al.,1980). It resulted in higher cellulose susceptibility toward enzymatic hydrolysis and utilized less energy when compared to ball-milled equivalents. Rollers operating at equal speeds have been used by Tassinari et al. (1982) and need low power consumption (due to decreased friction) and less roll wear as compared to a differential speed roller. The effect of compression milling on cellulose structure indicates that it reduces the crystalline index and crystallite size, slightly changes specific surface area, significantly varies the degree of polymerization, and increases the accessibility of cellulose to enzymatic hydrolysis (Ryu et al., 1982). Roller milling of wheat straw led to a considerable amplification of the enzymatic hydrolysis rate (Fan et a]., 1981). The relative extent of roller milling was 2.5 for 15 min and 3.1 for 30 min. Drying steam-treated aspen wood has a negative effect on enzyme accessibility (Saddler et al., 1982; Vallander and Eriksson, 1990). However, milling the dried sample in a Willey mill results in a positive effect on enzymatic hydrolysis. In contrast, Fitz milling provides only minor improvements in the hydrolysis rate (Fan et al., 1981). Fine Fitz milling, although substantially reducing the size of the substrate, affects the crystallinity index only slightly. Hammer milling gives good size reduction and increases bulk density but only promotes an insignificant gain in hydrolysis susceptibility (Ghose and Kostick, 1969; Mandels et al., 1974). Prolonged hammer milling reduces the susceptibility of cellulose to enzymatic hydrolysis (Mandels et al., 1974). Fluid energy milling or wet colloid milling reduces particle size and increases susceptibility but only after extensive treatment at a higher energy input (Ghose, 1979). Wet milling creates fibrillation and delamination of cellulose (McIntosh, 1967), but crystallinity and chain length are unaffected. It is either ineffective or much less effective than dry milling (Scallan, 1971). However, Neilson et 01. (1982) observed enhanced hydrolysis by simultaneous attrition (wet milling) and enzymatic hydrolysis of cellulosic substrate. This process allows continuous formation of new reactive sites. Milling is considered a good way of increasing cellulose reactivity toward enzymatic saccharification; however, these methods are unattractive energy wise (Datta, 1980). The major factor contributing to increased energy input is the fibrous nature of lignocellulose. The extru-

302

PURNENDU GHOSH AND AJAY SINGH

sion of a substrate with or without pressure slightly reduces the size of the substrate, but a pretreated substrate appears fibrous with an improved hydrolysis rate over an untreated substrate (Fan eta]., 1981). Results on the effect of mechanical treatment on the enzymatic hydrolysis of cellulosic substrates are shown in Table 11. B. RADIATION

In irradiation treatment, cellulose undergoes extensive depolymerization, thus increasing specific surface area for enzymatic/microbial attack. Radiation causes oxidative degradation of the cellulose molecule and cleavage of the cellulose chain. 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 (Han and Ciegler, 1983). The solubility of irradiated cellulose is increased in water and in alkaline solutions. The radiation effect is generally noticeable at or above 1 Mrad and is proportional to the radiation dose. While studying electron beam irradiation of rice straw, chaff, and

TABLE 11 EFFECTOF CHEMICAL PRETREATMENT ON ENZYMATIC HYDROLYSIS OF LIGNOCELLULOSIC MATERIALS Pretreatment

Substrate

Enzyme source

(%I

Reference

P.f." A.nb

70 76

T.r.C C30

80 25

Rao et al. (1983) Singh et al. (1990) Saddler et 01. 11982) Rolz et al. (1987) Rolz et al. (1987) Singh et al. (1990) Singh et al. (1990) Ghose et al. (1983) Ghose et al. (1983) Bonn et al. (1987) Vallander and Eriksson (1985) Fan et al. (1981) Dale and Moreira (1982)

NaC10, Peracetic acid Butanol Ethanol Organosolv H,O,

Bagasse Bagasse Aspen Bagasse Bagasse Corn cobs Wheat straw Rice straw Rice straw Poplar Wheat straw

Onozuka Onozuka An. A.n. T.r. T.r. T.v.~ T.r.

Ethylene glycol Freez explosion

Wheat straw Alfalfa

T.r. Novo SP122

NaOH NaOH HNO, NH,

so2

Penicillium funiculosum.

* Aspergillus niger.

Trichoderma reesei. Trichoderma viride.

Saccharification

45

36 34

85 80 90 56

54

94

LIGNOCELLULOSIC BIOMASS

303

sawdust, Kumakura and Kaetsu (1978) observed that reduced sugar formation increases slowly with a dose above 1 x lo8 rad and then increased more rapidly when the dose was between 1 and 5 x lo8 rad. The fiber strength of sugarcane bagasse changes considerably; as the irradiation dose increases, the fiber loses its strength and becomes mushy (Han and Ciegler, 1983). Radiation also appears to affect the lignin of lignocellulosic biomass as was evident from the increased presence of phenolic groups in irradiated wood fiber (Lipinsky, 1980). It has been suggested that irradiation causes an increase in the molecular weight of lignin thereby creating coalescence of the lignin layer and its detachment from much of the cellulose, increasing cellulose saccharification. Crystallinity of cellulose is one of the deterrents in effectively converting lignocelluloses to fermentable sugars. Radiation causes an apparent decrease in crystallinity and increases the digestibility of lignocelluloses. High energy electron irradiation provides an effective means of enhancing the digestibility of the carbohydrates in wood by rumen microorganism (Lawton et a]., 1951; Saeman et a]., 1952; Pritchard et al., 1962; Millet et al., 1976; Han and Ciegler, 1982,1983). For instance, the crystallinity index of 66.6% for bagasse is reduced to 44.4% by 100 Mrad gamma radiation. Samples of barley straw, pea straw, sugarcane bagasse, sunflower hulls, and pine sawdust irradiated up to 10 Mrad show little change whereas doses of 100 Mrad or more cause substantial losses of the fiber components and result in an increased rumen digestibility (Ibrahim and Pearce, 1980). Rice straw has maximum digestibility when treated at 5 x lo8 rad (Kumakura and Kaetsu, 1979), and the optimum dose for wheat straw digestibility is 2.5 x lo8 rad (Pritchard et al., 1962). Fan et al. (1980) observed that a 10 Mrad dose of gamma irradiation slightly reduces the further hydrolysis of wheat straw. The hydrolysis rate increases only when the radiation dose reaches a certain level (50 Mrad). These results suggest that the increase might have occurred because of the depolymerization of cellulose caused by an intensive input of energy. Further, inhibitory products formed at a low dosage might have also decomposed on extended irradiation treatment. The degree of polymerization decreases with an increased irradiation dose of gamma rays (from “Co) on waste paper (Kumakura and Kaetsu, 1982). These authors did observe any difference in degradation rate per unit dose between gamma and electron beam irradiation of filter paper. In addition to depolymerization, irradiation produces changes in the susceptibility of cellulose to subsequent acid/enzymatic hydrolysis. When bagasse was irradiated and subsequently hydrolyzed with 1 N H,SO,, a three times larger sugar yield was obtained compared to an untreated substrate (Han et al., 1981).

304

PURNENDU GHOSH AND AJAY SINGH

Irradiation treatment, despite its ability to enhance cellulose saccharification, holds little promise for commercial application because of high investment costs. However, the dosage requirement to solubilize or modify the substrate structure can be reduced considerably with a combination of chemical and irradiation pretreatment. IV. Thermal Treatment

Thermal treatment with or without steam has been used for upgrading the digestibility of various lignocellulosic materials (Bender et a]., 1970; Nesse etal., 1975; Wayman and Lora, 1978; Overend and Chornet, 1987; Bonn etal., 1987; Tanahashi et a]., 1988; Bobleter et a]., 1991; Dekker, 1991). Steam treatment may be of two types, viz. autohydrolysis and steam explosion, depending on pretreatment conditions. Autohydrolysis uses temperatures in the range of 170 to 200"C, whereas in steam explosion the temperature range extends to 250°C and the pretreatment ends with a sudden release of pressure (Vallander and Eriksson, 1990). A. AUTOHYDROLYSIS

Lignocellulosic substrate, when subjected to high-pressure steam for a specific period followed by a sudden release of pressure, results in extensive disintegration (Dietrichs et a]., 1978). The hemicellulose fraction can then be extracted with water, alkali, or solvents. The added advantage of extraction is the removal of inhibitory products, if any, generated in the process. The autohydrolysis reaction involves the formation of acetic acid from acetyl groups located in the hemicellulose fraction. The acid formed catalyzes the hydrolysis of hemicellulose and also the breakdown of the lignin-cellulose matrix (Bouchard et a]., 1989). Steam treatment causes substantial quantities of water to get into the intercrystalline region of cellulose (Wayman and Lora, 1978). The release of pressure after the treatment causes lignin coalescence and mechanical abrasion of the fiber (Lipinsky, 1981). The autohydrolysis process is effective in the case of hardwoods but not in softwoods (Clark and Mackie, 1987). Hardwoods containing high amounts of acylated xylan have long been successfully treated (Mammers and Menz, 1984; Clark and Mackie, 1987). Xylan solubilized by steam treatment gives rise mainly to oligosaccharides and only to relatively minor amounts of xylose monomers (Puls et al., 1985). Autohydrolyzed bagasse produces a substrate wherein the cellulose is rapidly saccharified enzymatically (Dekker and Wallis, 1982,1983).The in vitro

LIGNOCELLULOSIC BIOMASS

305

digestibility of crop by-products increased from 40 to 55% as a result of the steam pretreatment technique (Ibrahim and Pearce, 1983).

B. STEAM EXPLOSION Hemicelluloses and a highly depolymerized lignin fraction are then extracted. The residence time at higher temperatures should be kept low to minimize the formation of inhibitory by-products (Bouchard et al., 1986; Dekker, 1991). The remaining exploded cellulose fraction is an excellent substrate for enzymatic hydrolysis (Table 11).Pretreatment using the steam explosion technique results in the 59% conversion of sugarcane bagasse to sugars in 48 hr of enzymatic hydrolysis (Rao et a]., 1983). Other workers have also observed a positive effect of the explosion technique on glucose yield during enzymatic hydrolysis (Rolz et al., 1987; Bouchard et al., 1990). However, with increasing temperature, material becomes progressively darker because of the production of toxic phenolic compounds from lignin pyrolysis (Morjanoff and Gray, 1987).

Both batch and continuous high-pressure explosion systems have been developed for the pretreatment of lignocellulosic biomass (Brown, 1983). A continuous process is considered more effective because of the possibility of exercising precise control of the operating conditions and the efficient utilization of steam (Hasnain, 1985). A typical continuous process may consume 0.7 kg of steam per kg of dry matter processed. A modification of the process incorporating removal of hemicellulose prior to steam explosion is likely to give higher yields of reducing sugars on saccharification. Rolz et al. (1987) studied the pretreatment of sugarcane chips using the steam explosion technique. Steam explosion solubilized the hemicellulose completely and it also promoted the greatest enzymatic saccharification when compared to alkali, organosolv, or aqueous phenol treatment. Grouse et al. (1986) showed that steam explosion increases the pore volume accessible to the enzyme molecules, whereas substrate drying results in a reduction of pore volume. Similarly, Wong et al. (1988) found that steam explosion increases cellulose digestibility by increasing fiber porosity, i.e., pore volume accessible to the enzymes. C. HYDROTHERMOLYSIS

Hydrothermolysis is pretreatment in which the raw material is subjected to liquid water at a high temperature and pressure but no steam appears in the process (Bonn et al., 1987). When the influence of hydro-

306

PURNENDU GHOSH AND AJAY SINGH

thermal and organosolv pretreatment of poplar wood and wheat straw on enzymatic hydrolysis is compared, hydrothermolysis is a more suitable treatment of wheat straw, while organosolv treatment is better for poplar. Both treatments give glucose yields of 80-90% of the theoretical value in 70 hr of enzymatic hydrolysis. The more recent objective of hydrothermolysis pretreatment has been the fractionation of lignocellulosics in obtaining each of the polymeric components in maximum yield and purity (Rubio et al., 1986; Bouchard et al., 1990; Bobleter et al., 1991). V. Chemical Treatment

Chemical pretreatment methods have been extensively used for removing lignin and structurally modifying lignocellulosic biomass. Although most of the methods are effective, waste chemicals are often difficult to recycle or dispose of. Table I11 presents the effect of chemical treatment on the enzymatic hydrolysis of lignocellulosic materials. A. ALKALI Alkalies such as NaOH and ammonia increase the biodegradability of lignocellulosic materials (Feist et al., 1970; Han and Callihan, 1970; Toyama and Ogawa, 1975; Toyama, 1976; Pannir Selvam and Ghose, TABLE 111 EFFECTOF PHYSICAL, THERMAL, AND RADIATION PRETREATMENT ON ENZYMATIC OF LIGNOCELLULOSIC MATERIALS SACCHARIFICATION Pretreatment

Substrate

Ball milling Fitz milling Roller milling Extrusion

Wheat Wheat Wheat Wheat

straw straw straw straw

Autohydrolysis Steam explosion Steam explosion

Bagasse Bagasse Aspen

Enzyme source'

Saccharification

1%)

Reference

T.r." QM9414 T.r. QM9414 T.r. QM9414 T.r. QM9414

23 9 14 6

T.r. C30 Onozuka T.r. C30

83 85 70

Fan et al. (1981) Fan et al. (1981) Fan et al. (1981) Fan et al. (1981) Dekker and Wallis (1983) Rolz et al. (1987) Saddler et al. (1982) Bonn et al. (1987) Fan et al. (1981)

Hydrothermolysis Poplar T.v.~ Gamma Wheat straw T.r. QM9414 irradiation Trichodermo reesei. Trichoderma viride.

75 19

LIGNOCELLULOSIC BIOMASS

307

1980; Fan et al., 1981; Saida et al., 1982; Rao et al., 1983; Rolz et al., 1987). The treatment of lignocellulose by dilute alkali causes swelling, decreases the degree of polymerization and crystallinity, separates lignin, and disrupts lignin structure (Kleinert, 1966). The main consequence of alkali treatment is the saponification of intermolecular ester bonds, thus promoting the swelling of cellulose and favoring enzyme penetration into the cell wall (Feist et al., 1974; Fan et al., 1982). Cellulose digestibility depends on the amount of alkali per unit amount of cellulosic solid. For maximum cellulose digestibility with various cellulosic substances, the amount of alkali ranges between 0.1 and 0.15 g NaOH per g solid (Dunlap et al., 1976; Pearce et al., 1979). However, the optimum level of NaOH in treating substrates is the subject of much controversy as different workers have indicated different optimum levels of alkali concentration. Treating corn cobs, rice, and wheat straws with 2% NaOH was found optimum by Gupta et al. (1981) and Singh et al. (1989a), while Souza and Furtado (1977) reported higher levels of NaOH (4%) in the pretreatment of rice straw. In contrast, Tewari et al. (1987) reported 0.5% NaOH treatment as optimum for corn cobs. NaOH pretreatment of lignocellulosic materials increases the susceptibility to microbial degradation (Singh et a]., 1988a,b, 1989b). It is possible to decrease the requirement of alkali by means of presoaking. Pannir Selvam and Ghose (1980) found a significant increase in sugar production by presoaking followed by heat treatment of cellulosic substrates. Pretreatment of barley and pea straws with NaOH and NH,OH decreases the neutral detergent fiber content of the materials to varying degrees (Ibrahim and Pearce, 1983). In sugarcane bagasse, the addition of NaOH during steaming reduces the lignin content by 4 units. Alkali treatment also results in improved in vitro digestibility (Ibrahim and Pearce, 1983) and fermentation (Abdul-Halim et al., 1988). Combined chemical and thermal treatment increases the in vitro organic matter digestibility of agricultural residues by 60-80y0 (Table IV). While comparing different chemical pretreatments of various agricultural residues, it was observed that alkali treatment was the best for enzymaticdegradation (Singh et al., 1990) as well as for biodegradation (Singh et al., 1988a,b) of cellulosic substrates. Alkali-treated rice husks form an excellent substrate for solid-state fermentation (Kuhad and Singh, 1992). Hardwood hemicelluloses are more soluble and hence can be removed by cold alkali treatment. This treatment increases the average pore size in the cell wall structure of lignocellulose (Fan et al., 1981). About 8.6% lignin can be removed from wheat straw using this method. However, autoclaving the material along with alkali removes 43.5% lignin. Singh et al. (1988a,b) have shown that alkali treatment of corn

308

PURNENDU GHOSH AND AJAY SINGH TABLE IV

EFFECTOF COMBINED CHEMICAL AND THERMAL TREATMENT ON in Vitro ORGANIC MATTER DICESTIBILITY~ Concentration Pretreatmentb

(g/100 g dry wt)

Untreated NaOH NaClO, NH,OH NaHSO, a

-

6.0 3.0 5.2 1.0

Barley straw

Pea straw

Sugarcane bagasse

37.6 65.0 65.4 66.7 50.1

40.0 48.6 50.2 43.7 42.4

27.2 66.5 53.4 59.0 59.4

Adapted from Ibrahim and Pearce (1983). Steaming preformed in 500-ml capacity steel bombs at 170°C for 60 min

cobs removes 82% lignin (Fig. 2). The substrate after pretreatment in the former case appeared light yellow, and light brown and fluffy after autoclaving. Liquid ammonia has also been used, in a new technique called "freeze explosion," to increase cellulose reactivity toward enzymatic hydrolysis (Dale and Moreira, 1982). This method relies on the treatment of lignocellulose with volatile liquid under pressure followed by pressure re-

'"1

Pretreatment Wheat straw

Bagasse

Corn cobs

Groundnut shells

FIG 2. Effect of chemical treatment on delignification of lignocellulosic substrates. For alkali (SH) treatment, substrate was treated with 2% NaOH solution for 1 hr at 121°C; for sodium chlorite (SC) treatment, substrate was treated with 2% NaClO, solution for 2 hr at 80-90"C; for peracetic acid (PA) treatment, substrate was treated with 20% peracetic acid solution for 1 hr at 80-90°C.

LIGNOCELLULOSIC BIOMASS

309

lease to evaporate the liquid and reduce the temperature. Liquid ammonia is volatile and is known to swell cellulose (Lewin and Roldan, 1971; Schleicher et a]., 1974).Using this technique, more than 90% conversion of cellulose to glucose was achieved by enzymatic hydrolysis of alfalfa and rice straw (Dale and Moreira, 1982). B. ACID

Acids such as sulfuric, hydrochloric, nitric, and phosphoric are generally used for the hydrolysis of cellulose. Dilute acid has been used as an effective pretreatment agent for the further enzymatic hydrolysis of substrates such as newsprint, corn stover, and oak (Fan et a]., 1982). Mild pretreatment conditions (temperature less than 220°C, sulfuric acid concentration less than 1%,and reaction time 1 2 sec) are inadequate for acid hydrolysis but the treated substrate showed an increased response toward enzymatic hydrolysis. The acid-treated poplar on subsequent hydrolysis is reported to yield more than 90% of the theoretical amount of glucose in 24 hr (Knappert et a]., 1981).The increased susceptibility to enzyme attack is mainly attributed to hemicellulose removal and the reduction in the degree of polymerization of cellulose during the pretreatment. Datta (1981) fractionated corn stover using sulfuric acid treatment as outlined in Fig. 3. The weight loss during each fractionation step gave the weight of each of the major components: water solubles, hemicellulose, cellulose, and lignin in raw material. Pretreatment of wheat straw with dilute H,SO, (4.4%) at 98°C for 1 hr gave approximately a fourfold increase in the hydrolysis rate (Fan et al., 1981). Dilute acid treatment does not remove lignin from the substrate but modifies the lignin-carbohydrate linkage to a great extent. A two-step acid hydrolysis followed by a fermentation scheme has been suggested for biomass processing (Fig. 41 by several groups (Ghose and Ghosh, 1978; Ladisch et al., 1983; Cahela et al., 1983). Morjanoff and Gray (1987) achieved an improved hydrolysability of steam-exploded bagasse if 1 g HzSO,/lOO g dry substrate was added prior to the steam treatment. Abdul-Halim et al. (1988) examined the rates and percentage solubilization of wood components, mediated by an increasing concentration of H,SO, (from 0.1 to 72% v h ) at 25, 100, and 121°C. Maximum rates and solubilization were recorded with 72% H,SO, at 121°C. Brownell and Nakas (1991) studied the acid hydrolysis of poplar hemicellulose for further bioconversion to acetic acid. Hydrolysis with dilute sulfuric acid (4%, v/v) at 100°C for 60 min released 14.7 g/liter of fermentable substrate (mainly pentoses).

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PURNENDU GHOSH AND AJAY SINGH

Hemicellulose

1

PLANT BIOMASS Acid

Hydrolysis +

Acid recycle

A

A

Lignocellulose

Pentoses,-

I

Hexoses Enzyme

Glucose

+

I

Acid recovery

Microorganisms

Fermentation

Residualsubstrate

I

Microorganisms

Fermentation FIG.4. A two-step acid hydrolysis followed by a fermentation scheme for biomass processing.

LIGNOCELLULOSIC BIOMASS

311

A combination of acid (H,SO, or HCl) prehydrolysis with alkali [NaOH) delignification was attempted by Koukios and Valkanas (1982). However, this method leads to significant losses of polysaccharides. Pretreatment of holocellulose isolated from bagasse with a solution of ZnC1, and 0.5% HC1 (145°C for 10 min), then subjected to enzymatic hydrolysis, resulted in a 93% conversion as compared to 37% with untreated substrate (Azzam, 1987). Delignifying aspen wood with xylene sulfonic acid at 100°Cproduced a residue containing only 5.6% lignin (Springer and Zoch, 1971). Since the reaction takes place at only lOO"C, a pressure vessel is not required in this method. Hydrogen fluoride (HF) solvolysis, either in vapor or in solution, effectively hydrolyzes wood [Murray and Aster, 1984). Cellulose and hemicellulose are attacked by HF to give sugar fluorides. Removal of HF results in repolymerization with the formation of watersoluble oligosaccharides composed of up to 10 units. These oligomers can be converted by weak acid hydrolysis to monomers and can be further used for fermentation. C. OXIDIZING AGENTS Peracetic acid, sodium hypochlorite (NaOCl), sodium chlorite (NaClO,), and hydrogen peroxide are strong oxidizing agents that carry out chemical oxidation of lignin, form water-soluble compounds, and liberate cellulose for enzymatic/microbial degradation (Toyama and Ogawa, 1975; Toyama, 1976; Fan et al., 1981; Singh et al., 1 9 8 9 ~ )Under . extended periods of reaction time, sodium chlorite and hypochlorite also react with cellulose. The pretreatment with these reagents structurally modifies the substrate. Impressive enzymatic hydrolysis [Fan et al., 1981; Singh et al., 1990) and bioconversion [Singh et a]., 1988a, 1989c) rates have been obtained using these oxidizing agents. Pretreatment of lignocellulosic biomass with peracetic acid results in a substrate that is highly susceptible to enzymatic hydrolysis as well as biodegradation (Fan et a]., 1981; Singh eta]., 1988a,b, 1989c, 1990). Approximately a 5- to 10-fold increase in the hydrolysis rate over the untreated substrate is observed. This pretreatment can remove lignin extensively and the substrate appears white and fibrous. Alkaline peroxide treatment of agricultural residues increases enzymatic hydrolysis (Gould, 1984). Wheat straw and corn stover, when treated with an alkaline (pH 11.5) solution of hydrogen peroxide (HzOz : substrate ratio, 0.25, w/w), releases approximately half of the lignin and most of the hemicellulose. It is possible to recycle the supernatant, after adding make-up peroxide and readjusting pH, for further

312

PURNENDU GHOSH AND AJAY SINGH

treatment. Vallander and Eriksson (1985) studied the effect of alkaline peroxide (5% H,O,, 1 hr, pH 11.0-12.3) pretreatment of wheat straw for its further hydrolysis to reducing sugars. This method resulted in an efficient saccharification after 25 hr of enzymatic hydrolysis. Because of the mild conditions involved and nontoxic final products, the H,O, system appears to be significantly important as a potential substrate pretreatment method for the subsequent enzymatic hydrolysis from both an engineering and ecological point of view (Ghose, 1977).

D. GASES Several gases such as chlorine (Koukios and Valkanas, 1982), nitrous oxide (Wilke, 1978), ozone, and sulfur dioxide (Dunlap et a]., 1976; Millet et al., 1976; Ibrahim and Pearce, 1983; Rolz et al., 1987) have been used as pretreatment agents. In most cases, gases cause solubilization of lignin. Ozone attacks both lignin and carbohydrates, although the rate of carbohydrate decomposition is much slower and 50% delignification can be achieved. Using anhydrous ammonia as a water-free gas (3.5 g NH,/100 g dry matter) results in considerable improvement in vitro dry matter digestibility of barley straw, pea straw, and sugarcane bagasse (Ibrahim and Pearce, 1983). The efficiency of fractionation of lignocellulosic biomass can be increased by gaseous chlorination (Koukios and Valkanas, 1982). Sugar losses are minimal with this pretreatment method. The degradation of cellulose during delignification could also be avoided while recovering lignin quantitatively from pulping liquor. Clarke and Mackie (1987) showed that softwoods can be effectively hydrolyzed enzymatically if they are impregnated with SO, prior to steam treatment. With such a treatment of Pinus, 82% saccharification of cellulose is achieved in 72 hr. Pollution is a big problem with SO, treatment. The soluble and unfermentable lignosulfonates have to be purged from the system. This creates an effluent problem similar to those in the sulfite pulping industry. Handling and recovery of gases are certainly more difficult than liquids.

E. CELLULOSE SOLVENTS Certain solvents possess the ability to dissolve cellulose. The dissolution of cellulose by these solvents and its subsequent regeneration is one method of cellulose pretreatment. Methods of cellulose dissolution by a wide variety of solvents are available in the literature (Turbak et a]., 1980). The applicability of cellulose so obtained toward enzymatic

LIGNOCELLULOSIC BIOMASS

313

hydrolysis has been studied by the Purdue group (Tsao, 1978). The solvents used were CMCS, cadoxen, and sulfuric acid. CMCS, a nontoxic aqueous alkaline complex of iron and sodium tartrate, dissolves up to 4% cellulose at higher temperatures. Cellulose can be regenerated by adding excess water. The hemicellulose-free CMCS-treated residue has increased susceptibility to enzymatic hydrolysis. Cadoxen, an aqueous solution of ethylene diamine and cadmium oxide, readily dissolves cellulose. At room temperature cadoxen dissolves 10% by weight of cellulose. The addition of NaOH to cadoxen increases the solubility of the cellulose. Cellulose-cadoxen in the presence of excess water forms a soft floc which is easily hydrolyzed. A three-step process consisting of hemicellulose separation (by acid or alkali), cellulose dissolution, and cellulose regeneration has been developed (Tsao, 1978). The commercial attractiveness of the cadoxen process is limited because of the toxicity of cadmium compounds. According to Tsao (1978), the most practical solvent for cellulose dissolution is concentrated sulfuric acid (70-80%). Dissolved cellulose can be reprecipitated by adding methanol. This reprecipitated cellulose is amorphous and can be easily hydrolyzed by cellulase. However, the viability of the cellulose solvent process depends on the recovery of solvent.

F. SOLVENT DELIGNIFICATION Considerable interest in an "organosolv" process for the removal of lignin from lignocellulose has been generated in recent years. This interest stemmed from the original claim of Kleinert and Tayenthal (1931) that the lignin and hemicellulose sugars are not significantly altered in the solvent pulping process. Several solvent systems have been evaluated for their effectiveness in removing lignin from wood and agricultural residues (Katzen et a]., 1980) and include ethanol (Kleinert, 1966, 1974; April et al., 1979; April and Hansen, 1981; Ghose et a]., 19831, butanol (April et al., 1979, 1982; Hansen and April, 1981; Ghose et al., 1983), ethylamine (Pannir Selvam and Ghose, 1980), phenol (April et a]., 1979; Hansen and April, 1981), acetone (Nguyen, 1980), ethylene glycol (Fan et al., 1981), triethylene glycol (Han et a]., 1981), and dimethyl sulfoxide (Clermont and Bender, 1972). Ethylene glycol is an excellent solvent for modifying the structure of wheat straw. The relative extent of enzymatic hydrolysis, upon pretreatment with boiling (170°C for 30 min), was 7.3 and that with autoclaving was 8.6 (Fan et al., 1981).A similar significant increase in the hydrolysis rate of bagasse and rice straw upon ethylene glycol pretreatment was

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PURNENDU GHOSH AND AJAY SINGH

observed by Pannir Selvam and Ghose (1980).Solvents like triethylene glycol and ethylene glycol are effective delignification agents, but because of their high boiling point, recovery of these solvents is difficult. Dimethyl sulfoxide reacts with lignin (Koura et a]., 1972).Phenol is also considered a good solvent for lignin (Hansen et a]., 1981).However, under acidic conditions phenol is lost through copolymerization with lignin and possibly with the furfural formed from the hemicellulose (Schweers, 1974;April et a]., 1979). Among the solvent systems, aqueous ethanol and butanol are the most effective toward delignification (Ghoseet a]., 1983).Figure 5 shows the effect of butanol pretreatment on delignification of various agricultural residues. Although butanol may be a better lignin solvent because of its polarity, it is not as effective as ethanol in decrystallizing cellulose (Ghose et al., 1983).A process involving autohydrolysis followed by solvent treatment has been developed by the BERC group at I.I.T. Delhi (Fig. 6). This pretreatment process separates cellulose from hemicellulose and lignin present in rice straw and is carried out in two batch cycle operations. After autohydrolysis, the water extract containing pentose sugars is separated and used for single cell protein (SCP) production. Residual substrate is further treated with aqueous ethanol to obtain cellulose, for its subsequent use in bioconversion, and lignin as a by-product. Experimental evidence suggests that the mechanism of lignin removal is independent of the solvent system employed (Hansen and April, 1981).Thus changing the solvent system does not change the reaction

FIG.5. Effect of solvent (aqueous butanol, 1 : 1,vlv)treatment (120'C for 1 hr)on delignification of different agricultural residues.

LIGNOCELLULOSIC BIOMASS

315

RICE STRAW. 214

Steam.245 Water, 1017.6

1

--+ AUTOHYDROLYSIS --+

Flash steam, 174

t

903.1 (sugars, 4.5%)

L

Flash condensate

Catalyst, 2.5

I

Ethanol (65%) 546.7 Steam, 148 Steam, 425 I

I

SOLVENT

FERMENTOR

4 4 Condensate, 148 ]FILTER

PEG$

1' -I

Steam, 150 Solvent, 180.4

Solvent, 241.6

Condensate, 150 Condensate, 425 (Moisture, 10%)

I

ETHANOL FERMENTATION

V LIGNIN, 25.7 (Moisture, 10%)

FIG. 6. A two-stage pretreatment process (autccydrolysis followed by sc- 'ent treatment] for bioconversion of lignocellulosic material.

steps in the delignification process. However, the choice of catalyst is important for lignin removal. Various organic and inorganic compounds such as mineral acids, aluminum chloride, aluminum sulfate, ferric chloride, ferrous sulfate, maleic acid, oxalic acid, salicylic acid, and aromatic acids are good catalysts in the delignification process (Kleinert, 1966; Sarkanen, 1980; Ghose et a]., 1983; Park and Phillips, 1984). The effective role of acid catalysis may be due to the rapid hydrolysis of hemicellulose resulting in increased porosity and accessibility of the

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PURNENDU GHOSH AND AJAY SINGH

solvent to lignin. A higher acid concentration favors lignin condensation and reprecipitation of dissolved lignin (Park and Phillips, 1984). In the depolymerization reactions, especially in the breaking of P-aryl ether linkages, carbonium ions are formed as intermediates (Wayman and Lora, 1978). The role of aromatic compounds as catalysts can be explained by considering the view that these compounds prevent lignin self condensation. However, a higher temperature increases degradation of hemicellulose sugars into products which are likely to condense with lignin rendering it less useful (O’Neil et a]., 1979). The high temperature process also leads to increased cellulose depolymerization (Ghose et al., 1983);therefore, a comparatively low temperature process is preferred. The catalyst effective at low temperature should possess the following properties: 1. Sufficient polarity to break the lignin-carbohydrate bond selectively without depolymerizing cellulose 2. Rapid decrystallization and swelling of cellulose so that the susceptibility of treated residue to enzymatic hydrolysis is improved 3. Resistance to lignin depolymerization and condensation 4. High operational stability.

The solvent process brings native lignin into the solvent phase. It is uncertain whether native lignin enters the solution phase or whether molecular weight reduction by pretreatment causes lignin solubility. The removal of lignin is considered to be mass transfer controlled and not a chemical reaction (Sarkanen, 1980; April et al., 1982; Ghose et al., 1983). The physical behavior of lignocellulose, such as preferential swelling of the secondary wall of the fiber or different diffusivities of soluble lignin in different alcohol-water mixtures, probably plays an important role in the delignification process (April et al., 1982). The comparison of activation energy for lignin separation by different pretreatment agents indicates minimum activation energy with aqueous solvents (April et al., 1982). Although a precise comparison is not possible because of different materials and process conditions used, the lower value of activation energy obtained in the presence of aromatic acids is certainly due to its role as a selective catalyst in the delignification process. VI. Biological Treatment

Biological processing is based on the catalysis of reactions by whole cells or by isolated enzymes. Among the potential advantages over conventional chemical processing are: (1)greater substrate and reaction

LIGNOCELLULOSIC BIOMASS

317

specificity, (2) lower energy requirements, (3) lower risk of pollution, (4) higher yields of desired products, and (5) opportunities for transformations not feasible with chemical reagents [Kirk and Chang, 1981; Eriksson, 1991). A. POTENTIAL MICROORGANISMS The continuous biodegradation of lignocellulosic plant biomass by saprophytic microorganisms in nature is one of the most important parts of the biospheric carbon-oxygen cycle. The complete degradation of complex biomass is believed to be a result of the cooperative action among fungi, bacteria, and microflora in the soil. Degradation of woody materials is mainly accomplished by higher fungi. Fungi infect wood by the aid of spores or hyphae which grow into sound wood from infected material [Eriksson and Kirk, 1985). The decay of wood involves specific enzymes secreted from fungal hyphae that attack the cell wall of wood fibers. The structural features of lignin in wood and other lignocellulosic materials dictate unusual constraints on biodegradation systems. These constraints must be extracellular, nonspecific, and nonhydrolytic unlike other biopolymer degrading systems, which are hydrolytic and specific in nature (Kaushik and Bisaria, 1989). Based on the type of decay, lignin-degrading fungi are classified as white rot, brown rot, and soft rot [Table V). White rot and brown rot fungi are Basidiomycetes, whereas soft rot fungi are Ascomycetes or Fungi Imperfecti. Soft rot fungi slowly degrade lignin and are more successful in degrading polysaccharides than lignin. The hyphae of brown rot and white rot fungi penetrate from one cell to another through openings or by producing bore holes through the cell walls (Eriksson and Kirk, 1985). Brown rot fungi mainly degrade cellulose and hemicellulose in wood. B. BIODELIGNIFICATION 1. White Rot Fungi

The most extensive biodegraders are white rot fungi that degrade lignocellulosic materials in two distinctive ways: (1) the simultaneous degradation of cellulose, hemicellulose, and lignin; and (2) the selective degradation of lignin and hemicellulose. Phanerochaete chrysosporium (Sporotrichum pulveruientum, imperfect stage) is the most potent and best characterized biolignolytic organism available that possesses characteristic features of rapid growth, extensive degradation of lignin,

318

PURNENDU GHOSH AND AJAY SINGH TABLE V

POTENTIAL BIOLIGNOLYTIC SYSTEMS Fungi

Bacteria

White rot Phanerochaete chrysosporium (Sporotrichum pulveruientum) Coriolus versicolor Cyathus stercoreus Fomes annosus Ganoderma ostraie Panus conchatus Pleurotus ostreatus Phlebia radiata P. giganta Trametes versicolor Brown rot Lenzites trabea Poria placenta Serpula lacrymans Soft rot Aspergillus flavus Chaetomium globosum Fusarium oxysporum Paecilomyces sp.

Actinomycetes Arthrobacter sp. Micromonospora sp. Microbispora sp. Nocardia sp. Rhodococcus sp. Streptomyces badius S. cyaneus S . flavovirens S. setonii S. viridosporus Thermomonospora mesophila Other bacteria Acinetobacter sp. Bacillus sp. Ciostridium xyianolyticum Pseudomonas sp. Xanthomonas sp.

relatively high temperature optima, formation of asexual spores (conidiospores) in abundance, and rapid completion of the sexual cycle (Kirk and Eriksson, 1985; Kaushik and Bisaria, 1989). For lignin metabolism, the lignolytic system of this fungus is (1)produced constitutively, (2) expressed only during secondary metabolism, (3) triggered by carbon, sulfate, and nitrogen limitation, and (4) markedly affected by oxygen concentration (Kirk, 1983; Kirk and Shimada, 1985).Eriksson and co-workers have developed cellulase-less mutants of S. pulverulentum (Cel44), Phlebia radiata (Cel 26), and Phlebia giganta (Cel 50) which no longer degrade cellulose (Eriksson et al., 1980a,b; Eriksson and Vallander, 1982); degradation of wood is limited to lignin and hemicellulose (Table VI). When birch wood is treated with a cellulaseless mutant of P. chrysosporium, about 30% lignin removal is possible in 4 weeks (Table VII). Scanning and electron microscopic studies on the growth of wild-type and cellulase-less mutants have revealed that branching of fungal hypha within the cells is common for both (Eriksson and Kirk, 1985). In birch, the mutant Cel 44 grows from one cell to another by penetrating the perforated plates between vessels. In pine, the mutant spreads through the pits between ray cells and adjacent

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319

TABLE VI POTENTIAL BIOCHEMICAL MUTANTS OF WHITE ROT FUNGI Organism/Strain No. Phanerochaete chrysosporium Cel44 63-2 3113 LMT 26 der 8-2 der 8-5 PSBL-1 Phlebia giganta Cel26 Phlebia radiata Cel 50 a

Biochemical characteristicsa

Reference Ander and Eriksson (1976) Eriksson et al. (1983) Johnsrud and Eriksson (1985) Liwicki et al. (1985) Boominathan et al. (1990) Boominathan et al. (1990) Tien and Meyer (1990)

Cel-I, LI+

Eriksson and Vallander (1982) Eriksson and Vallander (1982)

Cel, cellulase; Xyl, xylanase; LI, lignin degradation;PO, phenol oxidase; DN, nitrogen deregulated;

+, positive; - , negative.

TABLE VII WEIGHT AND LIGNIN LOSSESFROM VARIOUS LIGNOCELLULOSIC SUBSTRATES BY LIGNOLYTIC ORGANISMS

yo of initial amount

Organism Phanerochaete chrysosporium Strain K-3 Strain 3113 Strain 85118 Streptomyces viridosporus

Streptornyces setonii

Substrate

Time (weeks)

Weight loss (%)

Lignin loss (oh)

4

24.4

4

19.0

19.9 29.5

4

16.6

26.6

Spruce

12

18.8

30.9

Maple

12

23.0

32.0

Grass

12

56.7

44.2

Spruce

12

20.8

34.1

Maple

12

19.3

29.5

Grass

12

49.2

39.0

Birch Birch Birch

Reference

Eriksson (1985) Eriksson (1985) Eriksson (1985) Antai and Crawford (1981) Antai and Crawford (1981) Antai and Crawford (1981) Antai and Crawford (1981) Antai and Crawford (1981) Antai and Crawford (1981)

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PURNENDU GHOSH AND AJAY SINGH

tracheids. One interesting feature of the Cel 26 mutant of P. radiata is that, unlike the wild type, it does not cause thinning of the cell wall. Instead it produces cavities and fissures in the fiber walls, particularly in connection with the middle lamella (Eriksson et al., 1980b). Straw treatment with Pleurotus ostreatus was first studied by Zadrazil (1977) and Lindenfelser et al. 1979). The preferential removal of lignin and increased in vitro digestibility have been studied by various workers (Zadrazil, 1985; Kamra and Zadrazil, 1986; Valmaseda et al., 1990, 1991).Although many of these studies were concerned with improving straw for animal feeding, the use of Trametes versicolor and P. ostreatus in the poplar pulp product has also been suggested (Kirpatrick et a]., 1989; Schiesser et al., 1989). Trametes versicolor degrades lignin and polysaccharides simultaneously, but under certain growth conditions it can be used for straw delignification (Zafar et a]., 1989). When P. ostreatus is used for partial delignification of wheat straw, 22 and 40% lignin can be removed in 30 and 70 days, respectively (Datta, 1981).During this period cellulose losses are 14 and 32%, respectively. The wheat straw residue after 10-20 days of biodelignification does not show any improvement in saccharification yield. However, a four- to fivefold improvement in saccharification was observed with residue treated for 50 days. Lignin degradation by white rot fungi is accompanied by cellulose and hemicellulose degradation. These easily metabolizable co-substrates are required for the growth of lignolytic organisms (Kirk and Moore, 1972). Thus valuable energy-rich fermentable material is lost in the process. For wheat straw delignification with P. ostreatus, the total weight loss is about 25%, and about 25% of the cellulose and 80% of hemicellulose are degraded before enough lignin (35%) is degraded to substantially increase saccharification yields (Datta, 1981).Although it is a low energy-consuming process, the energy losses due to lignin and saccharide degradation are substantial (Table VIII). TABLE

vm

ESTIMATEDENERGY AND FERMENTATION SACCHARIDE LOSSESON BIOLOGICAL TREATMENT'

Lignolytic organism

Substrate

Approx. loss of energy content (Yo)

Pleurotus ostreatus Sporotrichum pulverulentum

Wheat straw Birch wood

25 20

Adapted from Datta (1981).

Loss of fermentable saccharides (%) 40

15

LIGNOCELLULOSIC BIOMASS

321

Important progress in the knowledge of the biodelignification has been made during the last decade (Agosin et al., 1985; Dodson et al., 1987; Muheim et al., 1990; Mishra and Leatham, 1990; Valmaseda et al., 1991). However, the structural changes in the straw lignin during straw delignification and the mechanism of lignin depolymerization and biodegradation have only been partially established. The low molecular size lignin fractions disappear after fungal treatment (Hammerli et al., 1986). The degradation of cinnamic acid, the reduction in syringyl, and an increase in the p-hydroxyphenyl content in lignin were observed during fungal treatment (Lapierre and Monties, 1989). Cinnamic acids are involved in lignin-polysaccharide linkages (Iiyama et al., 1990) and their degradation could increase lignin extractibility. The fungal breakdown of p-0-4 and p-1 dimers has been extensively investigated, and the simultaneous degradation of condensed and uncondensed lignin and biphenyl structures have been demonstrated (Katayama et al., 1989). Fungal degradation of lignin side chains is responsible for the increased carboxyl content in lignin from wood decayed by Ganoderma australes (Martinez et al., 1990) and increased vanillic acid by P. tremellosus (Hedges et al., 1988). As discussed earlier, low molecular weight sugars are necessary to provide energy for growth and metabolism. In addition these sugars are also required to produce H,Oz which has an important role in lignin degradation [Eriksson, 1985). Enzymatic sources for extracellular H,O, have been investigated in several lignolytic fungi (Bourbonnais and Paice, 1988; Guillen et a]., 1990; Muheim et al., 1990). Valmaseda et al. (1991) studied the kinetics of straw solid state fermentation with P. ostreatus and T. versicolor to characterize the biodelignification process by these two white rot fungi. A decrease in syringyl/guaicyl and syringyl/p-hydroxy phenyl ratios and cinnamic acid content was noticed during fungal treatment. In contrast, an increase in the phenolic acid yield revealed fungal degradation of the side chains in lignin. Two successive phases were defined during the biodelignification process, characterized by a strong increase in respiratory activity and a decrease in free sugars, extractives, and in vitro digestibility (Table IX). 2. Bacteria

Actinomycetes are a diverse group of gram-positive bacteria commonly involved in degradation and humification of organic matter in soils. Because of the growth of actinomycetes as branching hyphae similar to filamentous fungi, they are well adapted to penetrate insoluble substrates like lignocellulose. Species belonging to genera Streptomyces, Arthrobacter, Micromonospora, and Nocardia degrade lignocellu-

322

PURNENDU GHOSH AND AJAY SINGH TABLE IX CHANGES IN WHEAT STRAW COMPONENTS AFTER A 6 0 - D A Y

TREATMENT WITH

LIGNOLYTICORGANISMS" Microorganism Parametersb

Control

Trametes versicolor

PIeurotus ostreatus

Weight loss Klason lignin Soluble lignin Extractives G1u cose

0 15.2

54.3

28.3 10.2 1.8

1.5

9.3 37.6

Xylose

10.5

Dinestibilitv

46.8

a

8.4 2.1

12.1 22.5 2.8 70.5

6.3 40.1 4.4

61.5

Adapted from Valmaseda et al. (1991). Values are represented in percentages.

losic biomass (Eriksson et al., 1990). Crawford and co-workers (1977; Crawford, 1978; Crawford and Sutherland, 1981; Antai and Crawford, 1981) have done considerable work on lignocellulose degradation using different Streptomyces species Streptomyces viridosporus and S. setonii degrade substantial amounts of lignin (Table VII) in hardwood, softwood, and grasses (Antai and Crawford, 1981). Lignin degradation by Streptomyces is a primary metabolic activity in contrast to P. chrysosporium (McCarthy and Broda, 1984). The lignin degradation by S. viridosporus is oxidative and involves demethylations, aromatic ring cleavage, and oxidative attacks on side chains (Crawford et al., 1983). Actinomycetes mainly solubilize lignin, whereas mineralization to CO, is much less than that of white rot fungi. Both S. viridosporus and T. mesophila produce an acid-precipitable polymeric lignin from corn and straw lignocellulose (McCarthy et al., 1986; Adhi et al., 1988). A number of Streptomyces species metabolize phenols and lignin-related aromatic acids, Douglas-fir lignin, and dehydrovanillin (Crawford et al., 1981). A crude extracellular enzyme preparation from S. cyaneus solubilized 20-30% of a [14C]ligninstraw (Mason et al., 1988).Nocardia and Xanthomonas species also degrade many lignin-related compounds. Many other bacterial species of Pseudomonas, Acinetobacter, Bacillus, and Clostridium degrade lignin (Janshekar and Fiechter, 1982; Kerr et al., 1983; Haider et al., 1985; Ammar et al., 1986; Kern and Kirk, 1987; Katayama et a]., 1988; Rogers et al., 1992). Substantial degradation (20-40%) of poplar dioxane lignin by bacteria was achieved in 7 days

LIGNOCELLULOSIC BIOMASS

323

by Odier et al. (1981).Low molecular weight fractions of lignosulfonates and kraft lignin may also be metabolized by bacteria (Ammar et al., 1986); however, natural lignin is more susceptible to degradation by bacteria. Anaerobic bacterium Clostridium xylanolyticum has shown an ability to degrade lignin (Rogers et al., 1992).It produces low molecular weight aromatics by inter-ring cleavage and ring modification. However, the major role of this bacterium in wood degradation is xylan degradation. Ultrastructural studies have shown that certain tunneling bacteria (belonging to Myxobacteriales and Cytophagales) are able to degrade lignified cell walls (Nilsson and Daniels, 1986). These bacteria release about 10% of the 14C02from ring- and side-chain-labeled DHP (synthetic lignin) in 16 days. Although certain strains of actinomycetes and bacteria metabolize intact lignin, it is at a much slower rate than with white rot fungi.

C. BIOPULPING The possible areas in the pulp and paper industry for using lignindegrading fungi include primary pulp manufacture, pulp modification, recycled fiber treatment, by-product conversion, and waste treatment. Kraft pulping is the major pulping method in which about 90% of the lignin is removed. Sporotrichum pulverulentum has been used to pretreat wood chips (Ander and Eriksson, 1978; Eriksson, 1981, 1985; Eriksson and Kirk, 1985; Jurasek and Paice, 1986). Biological pulping has been successfully carried out using Cel44 and CelZ6 strains (Eriksson et a]., 1980a; Eriksson and Vallander, 1982). In larger scale experiments, spruce and pine pulp were impregnated with 17% glucose and then inoculated with Cel 44. Chips treated for 2 weeks require less energy than untreated chips. Treatment with white rot fungi may be very useful for the production of thermomechanical pulp (Eriksson and Kirk, 1985; Jurasek and Paice, 1986). In industrial applications wood chips can be sterilized or pasteurized, inoculated with fungus, and placed in a series of silos from which the chips can be continuously removed. Studies have shown that pretreating aspen wood chips with P. brevispora decreases the energy requirements by 47% compared to untreated chips (Boominathan and Reddy, 1992). The application of this fungus also increases the tensile strength of pulp. Considering the high cost and uncertainty of energy supplies, especially in developing countries, an efficient biopulping process would be greatly advantageous from the standpoints of both economy and ensuring a continuous availability of good pulp. Biopulping would essentially eliminate the pollution hazards associated with the chemical

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PURNENDU GHOSH AND AIAY SINGH

pulping processes. However, more detailed studies are required on the physiological requirements during lignin degradation and the enzyme mechanism to successfully develop and run a large-scale biotechnological process using the fungal pretreatment of wood chips for reducing energy in mechanical pulping. An examination of pulp qualities and the energy-saving combined with an economic evaluation suggest that the microbial pretreatment of wood chips for pulp production has a viable future. This should lead to a lowered energy requirement and greater increased selectivity yielding a high-quality pulp product. Thermomechanical treatment dissolves and partly hydrolyzes hemicellulose. The hemicellulose degradation products can be enzymatically hydrolyzed to monosaccharides and used in various biotechnical processes as feedstocks. The remaining lignocellulose can be used in various biotechnical processes as feedstocks. The remaining lignocellulose is enzymatically delignified and lignin degradation products can be used in chemical processes for the production of speciality chemicals. The cellulose produced by this process is used as fiber or if required after hydrolysis, for biotechnical processes.

D. BIOBLEACHING The kraft pulping process removes about of the lignin. The remaining 10% or so in the pulp consists of various conjugated structures, including quinones, catechols, and chalcones which are responsible for the brown color characteristics of kraft pulp. Chlorination of the pulp removes most of the lignin and lignin-derived products. Many of the different compounds released during chemical bleaching are toxic and carcinogenic. A number of studies have focused on replacing the chlorination step with other methods (Eriksson and Kirk, 1985). The discovery that white rot fungi metabolize h a f t lignin prompted investigations into biobleaching (Kirk and Yang, 1979; Hiroi and Eriksson, 1976; Kirk and Shimada, 1985). White rot fungi are effective in removing residual lignin and in causing bleaching without damaging polysaccharides (Hiroi and Eriksson, 1976; Lindquist et al., 1977). Phanerochaete chrysosporium and T. versicolor are excellent biological agents for this purpose (Kirk and Yang, 1979). Various experiments with P. chrysosporium have shown that the kappa number, a measure of lignin content, could be reduced by 50-75% in 6-8 weeks (Eriksson and Kirk, 1985). Up to 50% of pulp cellulose is depleted in 7 days, but this can be retarded by adding glucose (malt extract, cane molasses, corn syrup, and starch) to repress the cellulase system. The biological bleaching of kraft pulp using white rot fungi in aerated

LIGNOCELLULOSIC BIOMASS

325

agitated cultures significantly increases the pulp brightness and reduces the lignin content (Kirpatrick et al., 1990).Reid et al. (1990)have shown that delignification with T. versicolor followed by an alkali extraction could remove up to two-thirds of the residual lignin from softwood kraft pulp. This delignification also increases biobleachability of the pulp and pulp sheet strength. Immobilization of fungi, P. chrysosporium and T. versicolor, during biological bleaching results in the production of biologically bleached pulp free from the fungal mycelium (Kirpatrick et al., 1989, 1990).

E. KRAFTEFFLUENTTREATMENT The pulp and paper industry generates over 700 billion gallons of toxic and intensely colored waste effluent that contains high molecular weight, modified, and chlorinated organic compounds that are of environmental concern. The effluents are treated by biological oxidation in aerated lagoons andlor activated sludge systems before being discharged into streams. Such treatments reduce the BOD and COD of the effluent but the color of the effluent persists. Several investigators have shown that kraft waste effluent can be decolorized by the white rot group of fungi, in particular, P. chrysosporium (Eaton et al., 1989; Huhnh et al., 1985; Eriksson and Kolar, 1985). Fungal decolorization, dechlorination, and detoxification of pulp and paper mill effluents has been achieved using P. chrysosporium (Sundman et al., 1981; Livernoche et al., 1983; Fukui et al., 1992). Color reduction up to 80% is achieved with P. chrysosporium in 1 week of treatment of bleach plant effluent. Up to 60% decolorization of E l stage liquor (first alkali extraction after chlorination) was achieved within 4 days by Eaton et al. (1989). Using a thin film reactor, 80% color removal is possible (Linko and Zhong, 1987). Immobilized cells of Coriolus versicolor have also been used for color removal from kraft mill wastewater (Livernoche et al., 1983).A continuous process, known as MyCoR (mycelial color removal), was developed by Messner et al. (1989) using P. chrysosporium. This process involves the immobilization of fungus in a rotary biological contactor (RBC) reactor. This process not only reduces the color significantly but also reduces the COD. Both high and low molecular weight compounds are removed by this process (Jones and Briedis, 1992). Yin et al. (1989) suggested that treating the E l effluent with ultrafiltration before RBC treatment is economically attractive. Prouty (1990) examined the use of a reactor for the treatment of bleach plant effluents. Several advantages of the aerated reactor are identified

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PURNENDU GHOSH AND AJAY SINGH

over the MyCoR RBC process; (1)the reactor uses air instead of pure oxygen, which reduces the cost of and is less hazardous to operate; and (2) the life time of fungus is longer in the aerated culture than in the MyCoR process. However, the color removal rate is lower in the aerated reactor than that of the MyCoR process, but the results obtained in a bench scale process provide a reasonable basis for the evaluation of an industrial fungal color removal process. VII. Epilogue

Lignocellulosic raw materials by virtue of their structure are relatively refractory to direct bioconversion. Cellulose in lignocellulosic substrates has regions of a highly resistant crystalline structure and the lignin surrounding cellulose forms a physical barrier which allows only limited sites available for enzymatic attack. Pretreatment is therefore necessary for the effective utilization of lignocellulosic material. Several approaches, including physical, thermal, chemical, biological, or combinations of these, have been explored in order to obtain each of the polymeric components of the lignocellulosic material in maximum yield and purity; to produce low-cost sugars (hexoses and pentoses) for use in biotechnological routes to fuels and chemicals; to use low-cost lignocellulosic biomass as protein-rich feed for ruminants; and to improve pulping and bleaching processes in the pulp and paper industry. Irradiation pretreatment holds little promise for commercial application because of high investments. Chemical pretreatment is considered more effective than physical treatment in producing a substrate for enzymatic hydrolysis and bioconversion. However, key factors appear to be the cost of chemical recovery and the effluent problem. Steam pretreatment has gained wide general acceptance because a high degree of enzymic digestibility can be achieved using this method. Because of the excellent ability of the white rot group of fungi in degrading lignin in wood, they have been used successfully for biodelignification, upgrading waste plant materials and low quality forages for animal feed, and for better quality pulp in the pulp and paper industry. The use of whole P. chrysosporium cells and lignolytic enzymes in the pulp and paper industry appears to hold promise in processes designed for biopulping, biobleaching, and in decolorizing bleach plant effluents. This will have a favorable impact on the world economy, sparing scarce energy sources and alleviating the current environmental problems with waste effluents. Biological treatment of wood and agricultural residues for their upgrading and subsequent use as a feed resource is another perspective.

LIGNOCELLULOSIC BIOMASS

327

Considerable developments in various pretreatment techniques have been made during the last decade. On the basis of what is already known, it can be considered that even though further developments may be necessary, many existing techniques can be presently used on a commercial scale.

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INDEX

Algae, fatty acid composition, 56 Alkali, lignocellulose treatment with, 306-309 Alkaline peroxide, lignocellulose treatment with, 311-312 Alkane monooxygenase system, yeast, 48 Alkane oxidation mechanisms, 43 yeast site, 47-48 Alkane utilization biomass correlation with, 57 induced cytological changes, 33-35 microbes associated with, 31-35 Alkane-utilizing microorganisms acetate incorporation, 39 acylglycerols, 59-61 alkane oxidation mechanisms, 43 biolipid extract, 76-78 bioremediation using, 78-79 biotechnological considerations, 41-42, 56-59 dicarboxylic acid production, 57-58 environmental considerations, 78-80 epoxides, 65-66 fat sources, 41-42 fatty acid accumulation, 55-56 fatty acid composition, 47-53 fatty alcohols, 64-65 genera containing, 31-35 glycolipids, 73-75, 79-80 hydrocarbons, 66 ketones, 64-65 lipid characteristics, 30 lipid classes, 37-41 lipid content, 35-37 mycolic acids, 79-80 oil recovery enhancement using, 79 peptidolipids, 75-76 phospholipids composition, 68-72 environmental effects, 72 fatty acid composition, 72-73 photosynthetic microorganisms, 56

A Acetic acid production aeration effects, 136 pentose concentration and, 136 pentose fermentation and, 99, 123-125 toxicity, pentose fermentation and, 138-139 Acetone production nutritional factors affecting, 133 oxygenation affecting, 133-134 pH effects, 132 temperature effects, 132-133 from xylose fermentation, 119-121 Acquired immune deficiency syndrome, associated virus, shiitake mushroom effects, 172 Actinomycetes delignification, 321-322 glycolipid toxicity, 79-80 produced glycolipids, 74-75 Acute-phase transport protein-inducing factor, lentinan activity and, 161 Acylglycerols from alkane-utilizing microorganisms, 59-61 yeast, 60, 193 Aeration effects acetone and butanol production, 133-134 butanediol production, 135 ethanol production, 129-131 organic acid production, 136 AIDS, see Acquired immune deficiency syndrome Air\liquid transfer, bioreactor contamination, 6 Air system, bioreactor, 18-19 Alcohols, fatty, alkane-utilizing microorganisms, 64-65 Aldose reductase, yeast, 104-105 335

336

INDEX

sterols, 61-63 waxes, 66-67 Antimicrobials, bioreactor asepsis, 22 Ammonia anhydrous, lignocellulose treatment with, 312 liquid, lignocellulose treatment with, 308-309

Ammonium hydroxide, lignocellulose treatment with, 307 Amylase, pectin extraction, 233 Anhydrogalacturonic acid, determination methods, 228-229 Anthocyanin, fruit juice clarification and, 247

APPIF, see Acute-phase transport protein-inducing factor L-Arabinan, protopectinase-C reaction with, 274 Arabinogalactan, structure, 274, 275 Arabinose fermentation, acetone-butanol from, 120 Arachidonic acid, production, 59 Aseptic bioreactor system agitator shaft and seal, 13-14 air system, 18-19 antimicrobials, 22 bioreactor internals, 12-13 liquid transfer system, 19-20 maintenance, 20-22 overcautious approaches, 24 piping and valves, 17-18 ports, 14-17 product changeover, 23 schedules and procedures, 23-24 Ash, in lignocellulosic biomass, 297 ATF':citrate lyase, yeast lipid synthesis, 201 Autoclave, bioreactor asepsis, 9-10 Azidothymidine, lentinan therapy, 169 AZT, see Azidothymidine

B Bacteria, see also specific types of bacteria bioreactor contamination, 3 butanediol production, 121-123 cytological changes, hydrocarbon-induced, 33-35

ethanol production, 114-116 hydrocarbon-utilizing genera, 30-35 lentinan effects, 168 pentose fermentation, aeration effects, 131

produced organic acids, 124-127 use in delignification, 321-323 Baked products, flavoring by yeast, 189-190

Beer fermentation, yeast lipids in, 187-189

Beverages, yeast lipids in, 187-189 Biobleaching, delignification, 324-325 Biochemical interesterification, modification of lipids, 201-204 Biolipid extracts alkane-utilizing microorganisms, 76-78 applications, 77-78 fatty acid composition, 76 Biomass, conversion techniques, 92-93 Biopulping, delignification, 323-324 Bioreactor agitator shaft and seal, 13-14 air system, 18-19 asepsis autoclavable bioreactor and, 10 definition, 11 design development, 6-7 inoculum preparation, 8-10 laboratory design, 8-10 medium preparation, 10-12 sterility assessment methodology, 7-8

contamination bioreactor system in, 5-6 filter device, 6 inoculum, 4 microbial mutation, 6 nutrient medium, 5 preventive measures, 6-7 probability, 2 foam sensors, 20 internals, 12-13 liquid transfer system, 19-20 maintenance medium, sterilization, 10-12 nonsterility, 2 consequences of, 3-4 microbes associated with, 3 ports, 14-17

337

INDEX slime, 3 sterility concept, 1-2 system, contamination sources in, 5-6 Bioremediation, with alkane-utilizing microorganisms, 78-79 Biotransformation, sterol, 62-63 Bleaching, delignification by, 324-325 Bread, yeast fermentation, 190 Butanediol fermentation, 121-123 pentose fermentation product, 99-100 production nutritional factors affecting, 134-135 oxygenation affecting, 135 pH effects, 134 temperature effects, 134 water activity affecting, 135 xylose fermentation product, 109 Butanol production nutritional factors affecting, 133 oxygenation affecting, 133-134 pH effects, 132 temperature effects, 132-133 toxicity, pentose fermentation and, 138 use in delignification, 314 xylose fermentation product, 119-121 Butyric acid effect on acetone and butanol production, 133 toxicity, pentose fermentation and, 138-139

C Cadoxen, lignocellulose treatment with, 313

Calcium pectin gels and, 225-226 polygalacturonase activity and, 240-241 Cancer, see also Tumors lentinan effects, 163 Cell culture, contamination probability, 2 Cellobiose, surfactant, 75 Cellulase, cellulose susceptibilities to, 298-299 Cellulose acetone-butanol fermentation, 1 2 1

chemical composition, 296 hydrolysis, 296-297 chemical, 306-312 radiation, 302-304 thermal, 304-306 in lignocellulosic biomass, 297 solvents, 312 wood concentration, 298 Cetyl palmitate, alkane-utilizing microorganisms, 67 Cheese flavoring, yeasts, 189-190 Chlorhexdecane, oxidation, lipids from, 6 7 Chlorine, lignocellulose treatment, 312 Cholesterol biotransformation, 63 reduction pectin in, 235 shiitake mushroom in, 154-155, 171-172 Cinnamic acids, delignification and, 321 Citrus peel, production methods, 283-285,287 Clostridia, acetone-butanol production, 119-121

CMCS, lignocellulose treatment with, 313 Coca butter, yeast lipid substitute, 198, 201 Complement pathway, lentinan effects, 158 Cord factor, 75 Corn stover, sulfuric acid treatment, 309 Cortexolone, biotransformation, 63 Cyclic AMP, lentinan effects, 165 Cyclophosphamide, lentinan therapy and, 166 Cytotoxic T lymphocytes, lentinan effects, 165

D Dairy products, yeast flavoring, 189-190 Dehydrogenation, alkane degradation, 43 Delignification alkali treatment, 307-308 biological bacteria, 321-323 biobleaching, 324-325 biopulping, 323-324

338

INDEX

cinnamic acids and, 321 energy losses from, 320 potential systems, 317 saccharide losses from, 320 white rot fungi, 317-321 catalyst choice in, 315-316 temperature effects, 316 solvent butanol, 314 ethanol, 314 ethylene glycol, 313-314 phenol, 314 wheat straw, component changes after, 321,322 Diabetes, lentinan effects, 170 Diacylglycerophosphoinositol,68 Diacylglycerophosphoinositolmannoside,47 Diacylglyerophosphoglycerol,68 Dicarboxylic acid production, alkane-utilizing microorganisms, 57-58

from lignocellulosic hydrolysate, 118-119 lipid effects, 131 metabolic inhibitor effects, 132 nutritional factors affecting, 128-129 oxygenation effects, 129-131 pH effects, 127-128 strain improvement, 139-141 temperature effects, 128 xylose isomerase and yeast, 116-118 yeast xylose fermentation and, 112-113 tolerance, pentose fermentation, 136-139 use in delignification, 314 Ethylene glycol, use in delignification, 313-314 Exopolygalacturonase activity, 241-242 microbial sources, 239 Exopolygalacturonate lyase, as pectic enzyme, 244

E F Eicosapentaenoic acid, production, 59 Emitanin, neoplasm inhibitor, 173 Endopolygalacturonate lyase characteristics, 243-244 microbial, plant disease and, 244-245 Endopolygalacturonase activity, 240-241 microbial plant diseases and, 244-245 sources, 239, 240 Endotoxin, see also Lipopolysaccharide assay, lentinan, 175 Epoxide production, alkane-utilizing microorganisms, 65-66 Ergosterol, alkane-utilizing microorganisms, 61-62 Eritadenine, hypolipidemic activity, 154-155 Esterases, as pectic enzyme, 238-240 Ethanol production bacteria fermentation and, 114-116 comparative microbial kinetics, 116 filamentous fungi fermentation and, 113-114

Fat alkane-utilizing microorganisms as sources, 41-42 modification by yeast, 198 modified, 201 Fatty acids beer and wine fermentation and, 187-188 biolipid extract, 76 composition acyl-CoA synthetase mutants, 57 algae, 56 alkenes as substrates, 47 decane as substrate, 55 heptadecane as substrate, 47, 52, 53,55 hexadecane as substrate, 45-47, 52, 53, 55, 60 octadecane as substrate, 47, 53 odd-chain alkane substrates, 43-45, 49,53 pentadecane as substrate, 47, 52, 53 tetradecane as substrate, 49 undecane as substrate, 55

INDEX content, biomass correlation, 57 functions, 42 interesterification, 202-204 lipid lipase activity and, 198 medical importance, 191 production, biotechnology, 57-59 short chain, production, 58-59 Fatty acid synthetase, recombinant, lipid modification, 200 Fatty alcohol production, alkane-utilizing microorganisms, 64-65 Fatty ketones, alkane-utilizing microorganisms, 65 Fermentor, see Bioreactor Ferredoxin, pentose metabolism limitation, 109 Ferulic acid, pectin, 220 Filamentous fungi, see Mold Filtration, sterilization by, bioreactor medium, 11-12 Flaxzyme, retting process and, 247 5-Fluorouracil, lentinan therapy and, 165

Food, yeast lipids in, 187-189 Fruit juice industry, pectinases in, 246-247

Fumaric acid production aeration effects, 136 nutrition effects, 136 pentose fermentation and, 126 Fungi, see also Mold; Yeast alkane utilization, 31-35 bioreactor contamination, 3 brown rot, use in delignification, 318 cytological changes, hydrocarbon-induced, 33-35 fatty acid composition, alkane oxidation, 53-56 lentinan effects, 168-169 lipid content, alkane utilization and, 41 organic acids, 124-127 pentose fermentation, 98-99 soft rot, use in delignification, 318 triacylglycerol accumulation, 61 white rot biobleaching using, 324-325 biochemical mutants, 318, 319 biopulping using, 323 delignification using, 317-321 Kraft effluent treatment with, 325

339 G

Galacturonic acid, pectin esterification, 217

Gels, pectin, 225-227, 231-232 Genetic engineering, use in yeast lipid modification, 199-201 Glucomannan, residue distribution, 94 Glucose fermentation, itaconic acid from, 127

Glycolipids actinomycete, toxicity, 79-80 alkane-utilizing microorganisms, 73-75 mycobacterial, toxicity, 80

H Hemicellulose acetone-butanol from, 121 chemical composition, 296 feedstocks, conversion techniques, 93 hydrolysate ethanol inhibitors in, 132 xylose fermentation, 118-119 in lignocellulosic biomass, 297 removal, alkali treatment, 307 residue distribution, 93 sugar composition, 93 thermal extraction, 304-305 wood concentration, 298 Hexadecane, oxidation, fatty acids from, 52-53

Hexane, oxidation, lipids from, 47-49 HIV, see Human immunodeficiency virus Human immunodeficiency virus lentinan effects, 169-170 shiitake mushroom effects, 172-173 Hydrocarbon inclusions, alkane utilization, 33 Hydrocarbon production, alkane-utilizing microorganisms, 66 Hydrocarbon-utilizing microorganisms, see Alkane-utilizing microorganisms Hydrocortisone, biotransformation, 63 Hydrochloric acid, lignocellulose treatment, 311 Hydrogen fluoride, lignocellulose treatment, 311

340

INDEX

Hydrogen peroxide, lignocellulose treatment, 311-31 2 Hydrolases, as pectic enzyme, 240-243 Hydroperoxidation, alkane degradation, 43 Hydroxamine acid reaction, pectin, 229 Hydroxylation, alkane degradation, 43 3-Hydroxymyristic acid, sterility assessment, 8 p-Hydroxyphenyl, delignification, 3 2 1 Hypertension, shiitake mushroom effects, 171-172

I Inclusions, hydrocarbon, 33 Interesterification, chemical, use in lipid modification, 201-203 Interferon, induction by shiitake mushroom, 155-156 Interleukin-2, lentinan effects, 165 Interleukins, lentinan effects, 159 Intraplasmic membrane, alkane utilization, 33 Itaconic acid production, pentose fermentation and, 127

K Ketone production, alkane-utilizing microorganisms, 64-65 Killer T cells, effect of lentinan, 159-160, 163

Kraft effluent treatment, lignocellulose hydrolysis and, 325-326

anti-parasite activity, 168 antitumor effects, 158 age factor and, 163 antiviral adtivity, 169-170 cell-mediated immune response to, 159 chemical structure, 157-158 combination therapies, 164- 168 enzyme activity affected by, 161-162 fertility effects, 163 immunoassay, 175 immunotherapeutic agent, 173-174 low natural killer syndrome effects, 170 medicinal benefits, 171 mode of action, 158 postoperative treatment, 175 radioprotection, 170 serum protein effects, 161 T-cell effect on activity, 158-160 toxicity, 163-164 treatment with, combination therapies, 164-168 tumor-host system effects, 162-163 tumor necrosis factor induction, 165 Lignin cellulose hydrolysis affect, 297 chemical composition, 296 function in wood, 299 in lignocellulosic biomass, 297 radiation effects, 303 removal alkali treatment, 307-308 microbial, 300 solvent, 313-31 7 structure, 299 thermal extraction, 305 wood concentration, 298 Lignocellulose, see also Hemicellulose feedstocks, conversion techniques, 92-93

L Lactic acid production, pentose fermentation and, 124 Lactone, lipid biotransformation, 190 Lentinacin, hydolipidemic activity, 154 Lectinan, see also Shiitake mushroom antibacterial activity, 168 anti-diabetic activity, 170 antifungal activity, 168-169 anti-inflammatory agents, 174

pretreatment, 94 Lignocellulosic biomass biological treatment bacteria, 321-323 biodelignification, 317-323 hiopulping, 323-324 Kraft effluent treatment, 325-326 potential microorganisms, 317 white rot fungi, 317-321 chemical treatment acid, 309-311

34 1

INDEX alkali, 306-309 cellulose solvent, 312 gases, 312 oxidizing agents, 311-312 solvent delignification, 313-316 enzymatic hydrolysis, effect of chemical pretreatment on, 302 mechanical treatment ball milling, 300-301 fluid energy, 301 hammer milling, 301 roller milling, 301 wet colloid, 301 pretreatment methods, 297-298 products derived from, 295-296 radiation effects, 302-304 structure, 298-300 selected, composition, 296-297 thermal treatment autohydrolysis, 304 hydrothermolysis, 305-306 steam explosion, 305 Linoleic acid, yeast lipid content, 192 Linolenic acid, yeast lipid content, 192 Lipase cheese flavoring, 189-190 detection, 204 sources, 192 use in interesterification, 202-204 wine flavoring, 188 yeast, use in lipid modification, 192 Lipids classes alkane utilization and, 37-38 alkane-utilizing microorganisms, 37-41

composition, hexadecane-grown cells, 38 content acetate-grown cells and, 39 alkane chain length and, 40 alkane-utilizing microorganism, 35-37

yeast, 39-41 ethanol production affected by, 131 modification fermentative synthesis and, 193-199

genetic engineering aspects, 199-201 interesterification, 201-204

stress effects, 200 yeast metabolism and, 187 sophorose, alkane-utilizing microorganisms, 74 sterols, yeast, 200 yeast, see Yeast lipids Lipopolysaccharide, lentinan therapy with, 164 Liquid transfer system, bioreactor, 19-20 Lyase endopolygalacturonate, 243-244 exopolygalacturonate, 244 oligogalacturonide, 244 polymethylgalacturonate, 244 Lymphokine-activated killer cell, lentinan effects. 165

M Macrophages, lentinan effects, 160 Maltose monocorynomycolate, surfactant, 75 Margaric acid, alkane-utilizing microorganisms, 67 Marinactan, lentinan therapy, 165 Medium preparation, bioreactor asepsis and, 10-12 Membrane filter bioreactor air system, 18-19 use in sterility assessment, 8 Mesophilic bacteria pentose fermentation, 99-100 physiological characteristics, 100 Metabolic inhibitors, ethanol production, 132

Methyldichloroethylene, use in lipid modification, 199 Methyl ketones, alkane-utilizing microorganisms, 65 Methyltrophic bacteria, alcohol and ketone production, 64 Milling, lignocellulose pretreatment, 300-302

ball, 300-301 hammer, 301 roller, 301 Miso production, yeasts in, 190 Mold, see also Fungi alkane utilizing, 31-35

342

INDEX

chlorinated alkane utilization, 55-56 ethanol production, 113-114 lipid content, alkane utilization and, 41 produced fatty acids, 192 xylose fermentation, aeration effects, 129-131 Mutations bioreactor contamination by, 6 ethanol production and, 141 Mycobacteria alkane utilization, 32,33 lipid classes in, 38 lipid content, 36 Mycolic acids, environmental effects, 79-80 Mycoplasma, bioreactor contamination by, 3

Oligogalacturonide lyase, characteristics, 244 Organic acid production factors affecting, 136 pentose fermentation and, 123-127 Oriental food, yeast fermentation, 189-190 Oxygenation effects acetone and butanol production, 133-134 butanediol production, 135 ethanol production, 129-131 fermentation process and, 188 lipid modification and, 193 organic acid production, 136 Ozone, lignocellulose treatment with, 312

P N Naphthalene, oxidation, 33 Neutrophils, lentinan effects, 160 Nitrogen, lipid modification and, 193, 196

Nitropectin, pectin polymerization and, 231 Norbornanone, biotransformation, 63 Nutrient medium, nonsterility, bioreactor, 5 Nutrition effects acetone and butanol production, 133 butanediol production, 134 ethanol production, 128-129 organic acid production, 136

0 Oil modification by yeast, 198 recovery, alkane-utilizing microorganisms, 79 sterilization, bioreactor, 11 Oleic acid, yeast wine fermentation, 187-188 Oligogalacturonase activity, 242 microbial sources, 239

Parasites, lentinan effects, 168 Pectic acid exopolygalacturonase action on, 242 nomenclature, 215 Pectic enzyme applications, 197 classification, 236 endopolygalacturonase as, 240-241 esterases as, 238-240 exopolygalacturonase as, 241-242 hydrolases, 240-243 lyases, 243-244 mode of action, 237 oligogalacturonase as, 242-243 in plant disease, 244-245 polymethylgalacturonase as, 243 Pectic substances acetylation, 217-218 depolymerization, 224-225 esterification, 2 17-2 18,2 24 history of, 213-214 interrelationship, 215-216 molecular mass, 221, 231 nomenclature, 214-215 occurrence and function, 222-224 properties, 224-228 solubility, 224 viscosity, 224 Pectin alkali-soluble, preparation, 277

INDEX anti-diarrhea effects, 235 applications cosmetic, 235 food, 233-234 pharmaceutical, 23 5 in arabinogalactan molecule, 274, 275 cell wall distribution, 222-223 characterization acetyl content, 229-231 anhydrogalacturonic acid content, 228 ester content, 229 jelling power, 231-232 methoxyl content, 229 molecular mass, 231 neutral sugar content, 231 chemical extraction, 233 chemistry, 216-222 citrus peel, production methods, 283-285,287 consumptin, prices and markets, 232 content, plant tissue, 223 cosmetic uses, 235 cross-linking, 227 drug delivery use for, 235 ferulic acid, 220 food uses, 233-234 gel formation, 225-227 gel strength, 231-232 hydroxamine acid reaction, 229 isolated, chemical and physical properties, 284-285 lemon peel, properties, 269, 270 manufacturing, 232-233 nomenclature, 215 pharmaceutical uses, 235 standardization, 233 structure detailed, 219-220 hypothetical, 217 middle lamellae, 219 plant cell wall, 219-220 polymer arrangement, 219,221 rhamnose effects, 217-218 side chains, 218-219 sugar beet composition, 276-277 gelation, 226-227 yields, various sources and, 285, 286 Pectinase applications, 245-248

343

classification, 236 detection method for, 237 in fruit juice industry, 246-247 production enzyme source, 245 factors affecting, 246 medium used, 245-246 retting process, 247 Pectinesterase characteristics, 238-240 classification, 236 microbial sources, 238-239 mode of action, 237, 238 saponification, 238 Pectinic acids nomenclature, 215 type 11, 219 Pentose, natural sources, 93-94 Pentose fermentation, see also specific pentose alcohol inhibition mechanisms, 137 butanol inhibition, 138 future prospects, 142-143 lipid effects, 131 metabolic inhibitor effects, 132 nutrition effects, 128-129, 133, 134-135 oxygenation effects, 129-131, 133-134, 135 pH effects, 127-128, 132, 134 product tolerance, 136-139 strain improvement, 139-142 temperature effects, 128, 132-133, 134 water activity effects, 135 weak acid inhibition, 138-139 Pentose-fermenting organisms acetic acid from, 125-126 acetone and butanol from, 119-121 butanediol from, 109, 121-123 ethanol from, 112-119 filamentous fungi, 98-99 growth characteristics, 95 mesophilic bacteria, 99-100 organic acids from, 123-127 yeast, 96-98 Pentose metabolism, see also specific sugar anaerobic scheme for, 109 bacteria, 107-112 transport mechanisms in, 101, 108 yeast, 102-107

344

INDEX

Peptidolipids, alkane-utilizing microorganisms, 75-76 Peptidomannan, shiitake mushroom, 156 Peracetic acid, lignocellulose treatment, 311 Phage, bioreactor contamination, 3 pH effects acetone and butanol production, 132 butanediol production, 134 ethanol production, 127-128 on organic acids, 136 Phenol, use in delignification, 314 Phosphoketolase bacteria, 109 yeast, 107 Phospholipids composition, alkane-utilizing microorganisms, 68-72 fatty acid composition, 72-73 production, alkane-utilizing microorganisms, 67-73 yeast, 68, 196 Pickle, yeast fermentation, 190 Platelet aggregation, shiitake mushroom and, 155 Polyploid, construction, ethanol production and, 141-142 Polygalacturonase bioassay, 237, 241 classification, 236 microbial, plant disease and, 244-245 mode of action, 237, 240-242 Polygalacturonase-AY, star diagram, 258 Polygalacturonate lyase classification, 236 mode of action, 237 Polygalacturonic acid, A,-type protopectinase affinity, 256, 257 Polymethylgalacturonase activity, 243 classification, 236 microbial source, 239 mode of action, 237 Polymethylgalacturonate lyase characteristics, 244 classification, 236 mode of action, 237 Pregnenolone, biotransformation, 63 Procaryotes, alkane utilization, 31-32

Protopectin A,-type protopectinase affinity, 256, 257 insolubility, reasons for, 222 nomenclature, 215 protopectinase action on, 249 source, 250 structure, 249 Protopectinase, see also specific protopectinase activity assay, 249-250 applications pectin production, 283-285 plant protoplast isolation, 285-286 single-cell foods, 286-287 A,-type affinity on protopectin and polygalacturonic acid, 256, 257 amino acid composition, 252, 255 biological properties, 252, 254 catalytic properties, 253-254 classification, 253 crystals, 251, 253 galacturonic acid hydrolysis by, 254-257 mechanism of activity, 255-258 N-terminal amino acid sequences, 252,255 occurrence, 250 physicochemical properties, 252, 254 purification, 250-252 star diagrams, 258 sugar composition, 252, 255 &-type biological properties, 261-264, 266 N-terminal amino acid sequence, 261,266 occurrence, 261 physicochemical properties, 261-264,266 purification, 261 substrate specificities, 261, 267 B-type amino acid composition, 268 occurrence, 266 pectin-releasing activity, 268, 269 properties, 267-268 purification, 266 classification, 236

345

INDEX history, 248-249 mode of action, 237 Trichosporon penicillotum, purification and isolation, 259-260 types of, 248 Protopectinase-C, see also Propectinase, B-type L-arabinan reaction with, 274 product identification, 271 reaction mechanism, 274 substrate characteristics, 2 71 identification, 271 preparation, 270 properties, 271 spectral data, 272-274 Protopectinase-F, see Protopectinase, &-type Protopectinase-L, see Protopectinase, &-type Protopectinase-N, see also Protopectinase, ArtYPe classification, 264 Protopectinase-R, see also Protopectinase, &-type classification, 264 elution profile, 261, 265 purification, 261 Protopectinase-S, 250; see also Protopectinase, A,-type gene cloning, 260 gene structure, 260 multiform, 258-259 PAGE, 260 profiles of inactivation, 259 purification and isolation, 259 Protopectinase-T, see also Protopectinase, B-type reaction products chromatography, 280 identification, 278-279 NMR,280-281 structure, 280, 282 substrate for alkali-soluble, 277 smallest, 277-278 specificity, 276 sugar composition, 276, 277 Protopectin-solubilizing enzyme, see Protopectinase

Protoplast isolation, protopectinase in plant, 285-286 Pulping delignification in, 323-324 Kraft, 323

R Radiation, Iignocellulose hydrolysis affected by, 302-304 Radioactive carbon, use in sterility assessment, 8 Radioprotection, lentinan, 170 Recombinant tumor necrosis factor, lentinan enhancement, 166 Retting process, pectinase in, 247 Rhamnolipids, alkane-utilizing microorganisms, 74 Rhamnose, pectin, 217-218 Rigidometer, gel strength, 232 Rotary biological contactor, Kraft effluent treatment, 325-326

S Serum proteins, effects of lentinan, 161 Shiitake mushroom, see also Lentinan anti-clotting, 174 antibiotics, 172 asthma prevention, 174 bone formation accelerator, 174 cosmetic industry, 175 history, 153-154 immunoregulatory substances, 173 medicinal benefits, 1 7 1 neoplasm inhibitor, 173 patented products and processes, 171-175,176 properties antibiotic, 155 anti-cancdanti-tumor, 156 anti-thrombotic, 155 antiviral, 155-156 hypolipidemic, 154-155 ulcer suppression, 174 viricides, 172-173 Slime, bioreactor, 3

346

INDEX

Single cell protein pectin, 234 yeast, 206-207 Sitosterol beer fermentation, 188 biotransformation, 63 Sodium, use in interesterification, 203 Sodium alkoxide, use in interesterification, 203 Sodium azide, ethanol production, 132 Sodium hydroxide, lignocellulose treatment with, 307 Solvent production, yeast, pentose metabolism, 112-113 Sophorose lipids, alkane-utilizing microorganisms, 74 Steam, lignocellulose pretreatment, 304-305

Stearic acid, yeast production, 198 Sterility, bioreactor, see Bioreactor, asepsis Sterility assessment methodology, bioreactor, 7-8 Sterilization bioreactor batch, 10-11 external continuous, 11 filtration, 11-12 Sterol production alkane-utilizing microorganisms, 61-63 biotransformation and production,

T Tallow, breakdown by yeast, 206 Tegafur, lentinan therapy and, 166 Tempeh, yeast lipids, 190 Temperature effects acetone and butanol production, 132-133

butanediol production, 134 delignification and, 316 ethanol production, 128 lignocellulose hydrolysis and, 304-306 Tempilistiks, bioreactor sterilization, 22 Tetradecane, oxidation, lipids from, 49 Thermophilic bacteria pentose fermentation, 100-101 properties, 101 sterilization time, 101 Thioglycolate, use in sterility assessment, 7 Thymus, lentinan activity, 158-160 Trehalose lipids, alkane-utilizing microoganisms, 74-75 Triacylglycerols, see Acylglycerols Tumor necrosis factor, recombinant, see Recombinant tumor necrosis factor Tumors, see also Cancer shiitake mushroom effects, 156 lentinan effects, 158 Turbosep, bioreactor asepsis, 16

62-63

Stigmasterol, biotransformation, 63 Stirrer shaft seal, bioreactor, 13-14 Succinic acid production, pentose fermentation, 124 Sulfite liquors, acetone-butanol, 120-121 Sulfur dioxide, lignocellulose treatment with, 312 Sulfuric acid lignocellulose treatment with, 309-311 use in cellulose hydrolysis, 313 Surfactants alkane oxidation, 73-74 environment effects, 78-79 Surfactin, 75 SurgicaUendocrine therapy, lentinan and, 167

Syringyl, delignification and, 321

U UFT, lentinan therapy with, 166

V Viruses lentinan effects, 169-170 shiitake mushroom effects, 155-156 Volutin, inclusions, 33

W

Water activity, butanediol production, 135

347

INDEX Wax production, alkane-utilizing microorganisms, 66-67 Wine fermentation, yeast lipids in, 187-189 Wood chemical composition, 298-300 extractive and nonextractive components, 299-300 Wort production, lipids in, 188

Xylose reductase bacteria, 108 yeast, 104-105 Xylulokinase bacteria, 108 yeast, 106 Xylulose, yeast fermentation, 96-98

Y X X-rays lentinan therapy and, 167 use in lipid modification, 199 Xylan, residue distribution, 93 Xylene sulfonic acid, 1ignocelluIose treatment with, 311 Xylitol production, xylose fermentation, 130 yeast production, 97 Xylitol dehydrogenase, yeast, 105-106 Xylose dehydrogenase, bacteria, 108 Xylose fermentation acetone and butanol from, 119-121 aeration effects, 129-131 butanediol from, 109,121 ethanol from, 112-119 ethanol tolerance, 137-138 fumaric acid from, 126 itaconic acid from, 127 lignocellulosic hydrolysate, 118-1 19 organic acids from, 123-127 strain improvement, 139-141 xylose isomerase, 116-118 yeast, 96-98 Xylose isomerase bacteria, 107-108 genetic transformation, 140 xylose fermentation with, 116-118 yeast, 104 Xylose metabolism anaerobic scheme, 109 bacteria, 107-112 early enzymes in yeast, 102 transport, 101-102 yeast, 102-107

Yeast, see O ~ S OFungi acetone-butanol production, 119-121 alkane oxidation site, 47-48 chlorinated alkane assimilation, 55-56 as enriching agents, 206 enzymes, applications, 192, 197 ethanol production, 112- 113, 1 18-1 19 ethanol tolerance, 137 fatty acid composition, alkanes as substrates, 48-53 hydrocarbon-utilizing, 31-35 lipase detection, 204 use in lipid modification, 192 lipid biotechnology, 204-207 lipid content, alkane utilization and, 39-41 oil and fat modification by, 198 pentose fermentation, 96-98 pentose metabolism, 102-107 phospholipid composition, 68, 196 single cell protein, 206-207 sterols, alkane utilization and, 61-63 triacylglycerol accumulation, 60, 193 xylose fermentation aeration effects, 129-131 with xylose isomerase, 116-118 xylose transport, 101-102 Yeast lipids beverage and food importance, 187-189 biotechnology, commercial significance, 204-207 cocoa butter substitute for, 198 composition, factors affecting, 193 dairy and baked products, 189-190 linoleic and linolenic content, 192 as lipid source, 186-187 medical importance, 190-192

348

INDEX

metabolism, lipid modification, 187 modification biological, significance, 204-207 fermentative synthesis and, 193-199 genetic engineering aspects, 199-201

interesterification, 201-204 yeast metabolism and, 187 oriental foods and pickles, 190-191 stress effects, 200 vegetable oils and, 185-186

CONTENTS OF PREVIOUS VOLUMES

Volume 29

Stabilization of Enzymes against Thermal Inactivation Alexander M. Klibanov Production of Flavor compounds by Microorganisms G. M. Kernpler New Perspective on Aflatoxin Biosynthesis J. W. Bennett and Siegfried B. Christensen Biofilms and Microbial Fouling W. G. Characklis and K. E. Cooksey Microbial Influences: Fermentation Process, Properties, and Applications Erick J. Vandamme and Dirk G. Derycke

Microbial Metabolism of Polycyclic Aromatic-Hydrocarbons Carl E. Cerniglia Microbiology of Potable Water Betty H. Olson and Laslo A. Nagy Applied and Theoretical Aspects of Virus Adsorption to Surfaces Charles P . Gerba Computer Applications in Applied Genetic Engineering Joseph L. Modelevsky Reduction of Fading of Fluorescent Reaction Product for Microphotometric Quantitation G. L. Picciolo and D. S. Kaplan INDEX

Volume 31

Enumeration of Indicator Bacteria Exposed to Chlorine Gordon A. McFeters and Anne K. Camper Toxicity of Nickel to Microbes: Environmental Aspects H . Babich and G . Stotzky

Genetics and Biochemistry of Clostridium Relevant to Development of Fermentation processes Palmer Rogers The Acetone Butanol Fermentation B. McNeil and B. Kristiansen

Volume 30

Survival of, and Genetic Transfer by, Genetically Engineered Bacteria in Natural Environments G. Stotzky and H. Babich

Interactions of Bacteriophages with Lactic Streptococci Todd B. Klaenhammer

Apparatus and Methodology for Microcarrier Cell Culture S. Reuveny and A. W. Thoma

INDEX

349

350

CONTENTS OF PREVIOUS VOLUMES

Naturally Occurring Monobactams William L. Parker, Joseph O’Sullivan, and Richard B. Sykes

Antitumor Anthracyclines Produced by Streptomyces peucetius A. Grein

New Frontiers in Applied Sediment Microbiology Douglas Gunnison

INDEX

Ecology and Metabolism of Thermomatrix thiopara Daniel K. Brannan and Douglas E. Caldwell Enzyme-Linked Immunoassays for the Detection of Microbial Antigens and Their Antibodies John E. Herrmann The Identification of Gram-Negative, Nonfermentative Bacteria from Water: Problems and Alternative Approaches to Identification N. Robert Ward, Roy L. Wolfe, Carol A. Justice, and Betty H. Olson INDEX

Volume 32

Microbial Corrosion of Metals Warren P. Iverson Economics of the Bioconversion of Biomass to Methane and Other Vendable Products Rudy 1. Wodzinski, Robert N. Gennaro, and Michael H. Scholla The Microbial Production of 2,3Butanediol Robert J. Magee and Nain Kosaric Microbial Sucrose Phosphorylase: Fermentation Process, Properties, and Biotechnical Applications Erick J. Vandamme, Jan Van Loo, Lieve Machtelinckx, and Andre De Laports

Volume 33

The Cellulosome of Clostridium thermocellum Raphael Lamed and Edward A. Buyer Clonal Populations with Special Refer. ence to Bacillus sphaericus Samuel Singer Molecular Mechanisms of Viral Inactivation by Water Disinfectants R. B. Thurman and C. P. Gerba Microbial Ecology of the Terrestrial Subsurface William C. Ghiorse and John T. Wilson Foam Control in Submerged Fermentation: State of the Art N. P. Ghildyal, B. K. Lonsane, and N. G. Karonth Applications and Mode of Action of Formaldehyde Condensate Biocides H. W. Rossmoore and M. Sondossi Occurrence and Mechanisms of Microbial Oxidation of Manganese Kenneth H. Nealson, Bradley M. Tebo, and Reinhardt A. Rosson Recovery of Biaproducts in China: A General View Xiong Zhenping INDEX

Volume 34

What’s in a Name?-Microbial Secondary Metabolism J. W. Bennett and Ronald Bentley

CONTENTS OF PREVIOUS VOLUMES Microbial Production of Gibberellins: State of the Art P. K. R. Kumar and B. K. Lonsane Microbial Dehydrogenations of Monosaccharides MiloS Kulhanek Antitumor and Antiviral Substances from Fungi Shung-Chang Jong and Richard Donovick Biotechnology-The Golden Age V. S. Malik INDEX

Volume 35

Production of Bacterial Thermostable a-Amylase by Solid-state Fermentation: A Potential Tool for Achieving Economy in Enzyme Production and Starch Hydrolysis B. K. Lonsane and M. V. Ramesh Methods for Studying Bacterial Gene Transfer in Soil by Conjugation and Transduction G. Stotzky, Monica A. Devanas, and Lawrence R. Zeph Microbial Levan Youn W. Han Review and Evaluation of the Effects of Xenobiotic Chemicals on Microorganisms in Soil R. J. Hicks, G. Stotzky, and P. Van Voris Disclosure Requirements for Biological Materials in Patent Law Shung-Chang Jong and Jeannette M. Birmingham INDEX

351

Volume 36

Microbial Transformations of Herbicides and Pesticides Douglas J. Cork and James P. Krueger An Environmental Assessment of Biotechnological Processes M. S. Thakur, M. J. Kennedy, and N. G. Karanth Fate of Recombinant Escherichia coli K-12 Strains in the Environment Gregg Bogosian and lames F. Kane Microbial Cytochromes P-450 and Xenobiotic Metabolism F. Sima Sariaslani Foodborne Yeasts T. De6k High-Resolution Electrophoretic Purification and Structural Microanalysis of Peptides and Proteins Erik P. Lillehoj and Vedpal S. Malik INDEX

Volume 37

Microbial Degradation of the Nitroaromatic Compounds Frank K. Higson An Evaluation of Bacterial Standards and Disinfection Practices Used for the Assessment and Treatment of Stormwater Marie L. O’Shea and Richard Field Haloperoxidases: Their Properties and Their Use in Organic Synthesis M. C. R. Franssen and H. C. van der Plas Medicinal Benefits of the Mushroom Ganoderma S. C. Jong and J. M. Birmingham

352

CONTENTS OF PREVIOUS VOLUMES

Microbial Degradation of Biphenyl and Its Derivatives Frank K. Higson The Sensitivities of Biocatalysts to Hydrodynamic Shear Stress Ales Prokop and Rakesh K. Bajpai Biopotentialities of the Basidiomacromycetes Somasundararn Rajarathnarn, Mysore Nanjarajurs Shashirekha, and Zakia Bano INDEX

Volume 38

Selected Methods for the Detection and Assessment of Ecological Effects Re-

ISBN 0-12-002639-2

sulting from the Release of Genetically Engineered Microorganisms to the Terrestrial Environment G. Stotzky, M. W. Broder, J. D. Doyle, and R. A. Jones Biochemical Engineering Aspects of Solid-state Fermentation M. V. Ramona Murthy, N. G. Karanth, and K. S. M. S. Raghava Rao The New Antibody Technologies Erik P. Lillehoj and Vedpal S. Malik Anoxygenic Phototrophic Bacteria: Physi ology and Advances in Hydrogen Production Technology K. Sasikala, Ch. V. Ramana, P. Raghuveer Rao, and K. L. Kovacs INDEX