Advances in Applied Microbiology, Volume 43

ADVANCES IN

Applied Microbiology VOLUME 43

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ADVANCES IN

Applied Microbiology Edited by

SAUL L. NEIDLEMAN Oakland, California

ALLEN I. LASKIN Somerset, New Jersey

VOLUME 43

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This book is printed on acid-free paper. @ Copyright 0 1997 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. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher's consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (222 Rosewood Drive, Danvers, Massachusetts 01923), for copying beyond that permitted by Sections 107 or 108 of the U.S. Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-1997 chapters are as shown on the title pages, if no fee code appears on the title page, the copy fee is the same as for current chapters. 0065-2164/97 $25.00

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4

3

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1

CONTENTS

Production of Acetic Acid by Clostridium thermoaceticum

MUMRCHERYAN, S A W PAREKH, MINISHSHAH, AND KUSUMAWITJITRA I. 11. 111. IV. V. VI. VII.

Historical Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acetic Acid Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cfostridiuin fhermoaceticum . . . . . . . . . , . . . , . . . . . . . . . . . . . . . . . . . . . . . . Strain Improvement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Low-Cost Media . . . . . . . . . . . . . . . . . . . . , . , . . . . . . . . . . . . . . . . . . . . . . . . Bioreactors for Improving Productivity . . , . . . . . . . . . . . . . . . . . . . . . . . . . . Downstream Processing of Acetate Fermentation Broths. . . . . . . . * . . . . . . . ...,.*........ References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

.

.

1 3 5 13 16 24 29 31

Contact Lenses, Disinfectants, and Acanthamoeba Keratitis

DONALDG. AHEARNAND MANALM. GABRIEL ..................

I. Introduction and Taxonomy 11. Biology. ... ... . . . . . . . .. .. . .. .. . . . .

V. VI.

................. ................. . . .. . .... ... , . .. .. .. .. . ................ Disinfection. . . . . . . . . . . . . . . Adherence to Lenses. . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . .

35 3a 40 42 44

47 51

Marine Microorganisms as a Source of New Natural Products

V. S. BERNAN,M. GREENSTEIN, AND W. M. MAIESE I. 11. 111. IV V.

Introduction .................................................... Natural Products from Marine Microorganisms . , . . . . . . . . . . . . . . . . . . . Overview of Wyeth-Ayerst (W-AR) Marine Natural Products Program . . . . Marine Biotechnology. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Summ......................................................... References .....................................................

.

..

V

.

.

.

57

58 70 86 a7 a7

vi

CONTENTS

Stereoselective Biotransformations in Synthesis of Some Pharmaceutical Intermediates &MESH

N . PATEL

I . Introduction ....................................................

I1. Tax01 Semisynthesis ............................................. EI. Thromboxane A 2 Antagonist ......................................

N . ACE Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

.

V Anticholesterol Drugs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

VI. Antiinfective Drugs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VII . Calcium Channel Blocking Drugs .................................. Vm . Antipsychotic Agents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IX . X. XI. XII.

Antiarrhythmic Agents. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Potassium Channel Openers., . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Antiinflammatory Drugs .......................................... Antiviral Agents ................................................ XI11 Prostaglandin Synthesis .......................................... References .....................................................

.

91 92 98

101 107 113 120 121 125 127 129 130 132 133

Microbial Xylanolytic Enzyme System: Properties and Applications

P R ABAJPAI ~ I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

II. Structure of Xylan and Its Interaction with Plant Cell Walls. . . . . . . . . . . . . III. Properties of Xylanolytic Enzymes ................................. N . Production of Xylanolytic Enzymes ................................. V. Application of Xylanases .........................................

VI. Conclusions .................................................... References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

141 142 149 163 167 184 185

Oleaginous Microorganisms: An Assessment of the Potential JACEK LEMAN

.

I Introduction .................................................... I1. Microbial Oil ...................................................

Single Cell Oil .................................................. Specialty Fats and Oils ........................................... Valuable Metabolites ............................................. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

195 197 203 206 225 234 235

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

245

C~NTENTSOFPREVIOUSVOLUMES ..........................................

251

III. N. V. VI.

INDEX

Production of Acetic Acid by Clostridium thermoaceticum MUNIRCHERYAN, AND

S A W PAREKH, MINISHSHAH, KUSUMAWITJITRA

Agricultural Bioprocess Laboratory University of Illinois Urbana, Illinois 61802

I. Historical Background 11. Acetic Acid Production A. Aerobic Process B. Anaerobic Process 111. Clostridium thermoaceticum A. Substrates for Fermentation B. Mineral Requirements IV. Strain Improvement V. Low-Cost Media VI. Bioreactors for Improving Productivity VII. Downstream Processing of Acetate Fermentation Broths References

I. Historical Background

Acetic acid (ethanoic acid, methyl carboxylic acid) has been produced as long as wine making has been practiced and therefore dates back to at least 10,000 BC (Nickol, 1979; Agreda and Zoeller, 1993), although it could be predated by certain fermented foods made from milk (Allgeier et al., 1974).It is assumed that the first vinegar, which is an aqueous solution of acetic acid, resulted from spoiled wine (Ghose and Bhadra, 1985), given that the Latin word acetum means sour or sharp wine. It initially functioned as a medicinal agent and was most likely the first known antibiotic. For most of human history, all acetic acid was derived by the same age-old process of sugar fermentation to ethyl alcohol and subsequent oxidation to acetic acid by rnicroorganisms to produce vinegar. This was the sole source of acetic acid. Late in the nineteenth century, this process was supplemented by the advent of wood distillation, which provided an additional source of acetic acid (Agreda and Zoeller, 1993). In 1995, annual production of acetic acid by the petrochemical route in the United States was 4.68 billion pounds, 1 ADVANCES IN APPLIED MICROBIOLOGY,VOLUME 43 Copyright 0 1997 by Academic Press, Inc. All rights of reproduction in any form reserved. 0065-2164/97$25.00

2

M. CHERYAN et 01.

ranking 35th among all chemicals produced (Anonymous, 1996). Production increased at an annual rate of 18% in 1993-95. Vinyl acetate ranked 41st, averaging 3 billion lb in 1993-95. In 1916, the first dedicated plant for the production of acetic acid by chemical rather than biological means became commercial (LeMonnier, 1965). This method was based on acetylene-derived acetaldehyde, and it marks the advent of inexpensive, industrial-grade acetic acid and the birth of a viable industry based on its use (Agreda and Zoeller, 1993). The advantages to chemical synthetic routes include high acetate concentrations (35-45 YOby weight), high production rates, and acetic acid generated in the free-acid form. The major disadvantages are the need for high temperatures and high pressures, good agitation, the threat of explosion, the high cost of catalysts, and the dependence on nonrenewable uncertain sources of raw materials (crude oil). Fermentation production routes have traditionally been aimed at the food market. Vinegar production usually requires lower capital investment, has shorter start-up times, and can generate different types and flavors of vinegar when different carbohydrate sources are used. Furthermore, the raw material (e.g., corn) is a renewable resource. The cost of acetic acid from chemical synthesis has ranged from 25 to 35 a/lb on a 100% basis, while it is 35-45 allb from aerobic fermentation. Clearly the latter value must be decreased if fermentation production is to supply the demand for nonfood uses. The industrial importance of acetic acid can be understood from Fig. 1. The major outlet today is for vinyl acetate, which is used for vinyl plastics, adhesives, textile finishes, and latex paints. This market has grown rapidly during the past few years due to the demand for synthetic fibers. In 1979 calcium magnesium acetate (CMA) was identified as a noncorrosive environmental-friendly alternative to chloride salts for deicing roads (Marynowski et al., 1985). Road salt use is 10-12 million tons per year in the United States, and CMA in solid form could supply about 5-10% of that market within the next decade (Wise, 1992). Liquid potassium acetate is being used now as a deicer for airport runways and as a heat exchange fluid; in the latter role it could serve as a partial replacement for ethylene glycol (1995 annual production of 5.23 billion lb). In addition, there are reports that CMA or calcium acetate can also be used as an additive to coal-fired combustion units, for example, boilers used by electrical utilities (Levendis, 1991; Manivanan and Wise, 1991; Sharma, 1991). Here calcium acts as a “grabber” for sulfur in the coal, reduces sulfur dioxide emissions, and partially relieves the problem of acid-rain pollution. If these environment-related substitu-

PRODUCTION OF ACETIC ACID BY C. thermoaceticum

3

ACETIC ACID

I

1

Acetic Anhydride

1

r

1

Pharmaceuticals

I Textile Finishes Vinyl Plastics Latex Paints Adhesives

1

Plasticizers

Cellulose Acetate

1

Pharrnaceut.

/Solvents1

Transparent Sheets Textile Fibers Photo Film

Heat Transfer Liquids Meat Preservative Neutralizer Fungicide De-icers

FIG.1. Uses of acetic acid.

tions take place, the demand for acetic acid would increase tremendously. II. Acetic Acid Production

Acetic acid as an industrial chemical is presently produced from fossil fuels and chemicals by three processes: acetaldehyde oxidation, hydrocarbon oxidation, and methanol carbonylation. It can also be produced by biological routes, which forms the basis of this review. A. AEROBICPROCESS Food-grade acetic acid is produced by the two-step vinegar process (Allgeier et a]., 1974; Ebner and Follman, 1983, Crueger and Crueger, 1990; Shreve and Brink, 1977). The first step is the production of ethanol from a carbohydrate source such as glucose. This is carried out at 30°C using the anaerobic yeast Saccharomyces cerevisiae. C6H12O6

+ 2C02 + 2CH3CH20H.

The second step is the oxidation of ethanol to acetic acid. Although a variety of bacteria can produce acetic acid, only members of Acetobacter are used commercially, typically the aerobic bacterium Acetobac-

4

M. CHERYAN et al.

ter aceti at 27-37°C. This fermentation is an incomplete oxidation because the reducing equivalents generated are transferred to oxygen and not to carbon dioxide: 2CH3CHzOH + 0

2

-+ ZCHsCOOH + 2Hz0.

The overall theoretical yield is 0.67 g acetic acid per gram glucose. At the more realistic yield of 76% (of 0.67, i.e., 0.51 g per gram glucose), this process requires 2.0 pounds of sugar or 0.9 pounds of ethyl alcohol per pound of acetic acid produced (Busche et al., 1982). Complete aeration and strict control of the oxygen concentration during fermentation are important to maximize yields and keep the bacteria viable (Muraoka et d.,1980; Osuga et a]., 1984). Submerged fermentation has almost completely replaced surface fermentation methods. The drawand-fill mode of operation can produce acetic acid at concentrations up to 10% wt/wt in continuous culture at pH 4.5 in about 35 hours (Crueger and Crueger, 1990; Ebner and Follman, 1983; Nickol, 1979). B. ANAEROBIC PROCESS

In the 1980%another process for production of acetic acid emerged based on anaerobic fermentation using Clostridia. These organisms can convert glucose, xylose, and some other hexoses and pentoses almost quantitatively to acetate according to the following reactions: CsHlZOe + 3CH3COOH, + 5CH3COOH.

ZCSH1005

Typical acidogenic bacteria are Clostridium aceticum (Braun et al., 1981), Clostridium thermoaceticum (Fontaine et al., 1942; Andreesen et al., 1973), Clostridium formicoaceticum (Andreesen et d., 1970) and

Acetobacterium woodii (Balch et d., 1977). Many can also reduce carbon dioxide and other one-carbon compounds to acetate (Ljungdahl, 1983).

This fermentation route has several advantages. It is anaerobic and thus should have lower fermentation cost. The theoretical yields are higher than the aerobic fermentation: 3 moles of acetic acid are produced per mole of glucose consumed, that is, 1 g acetic acid/g glucose (Brownell and Nakas, 1991; Brumm, 1988; Parekh and Cheryan, 1990a,b, 1991; Schwartz and Keller, 1982b; Wise et al., 1991). Actual yields with C. thermoaceticum have ranged from 0.85 (Fontaine et al., 1942; Ljungdahl et al., 1986; Wang et al., 1978) to 0.90 g acetic acid per gram glucose and greater (Parekh and Cheryan, 1990a, 1994a; Shah and Cheryan, 1995b). Until 1967, C. thermoaceticum was the only acetogen

PRODUCTION OF ACETIC ACID BY C. thermoaceticurn

5

Glucose

t

Pyruvate

CO,

Formate

--+

-7-

Formyl-THF -CH,-THF

2 Acetyl CoA

/--

CoA

Lzz-

Corrinoid enzyme

CoA Acetyl CoA

FIG.2. Embden-Myerhoff pathway for production of acetic acid.

easily available for study (Ljungdahl, 19861. Consequently, the most detailed studies of acetate synthesis have been performed with this organism. III. Clostridium thermoaceticum C. thermoaceticum was isolated from horse manure. It is an obligate anaerobe, Gram-positive, spore-forming, rod-shaped, thermophilic organism with an optimum growth temperature of 55-60°C and an optimum pH of 6.6-6.8 (Ljungdahl et al., 1985). The wild strains produce 2.55 moles of acetic acid per mole of glucose fermented (actual yield) and only 13-20 g/liter acetic acid in batch fermentation (Fontaine et al., 1942; Wang et a)., 1978; Sugaya et al., 1986).The fermentation of sugars to acetate is a complex process. As shown in Fig. 2, one mole of hexose is metabolized by the Embden-Meyerhof pathway to yield 2 moles of pyruvate, which are further metabolized to 2 moles of acetate (formed from carbons 2 and 3 of the pyruvate] and to 2 moles of COz (formed from the carboxyl groups) (Wood, 1952a). The two moles of COz serve as electron acceptors, and one mole COz is finally reduced to methyltetrahydrofolate (CH,THF). The CH3THF

6

M. CHERYAN et al.

then combines with the second C 0 2 and coenzyme-A (CoA), forming acetyl-CoA, and finally the formation of the third mole of acetate (Barker and Kamen, 1945; Wood, 1952b). The overall reaction can be written as follows: C6HI2O6+ 2H20 -+ ZCHBCOOH+ 2 C 0 2 + 8H’ + 8e-, 2C02 + 8H’ + Be- -+ CH3COOH + 2H20.

Enzymes involved in formation of the third mole of acetate are tetrahydrofolate enzymes, carbon monoxide dehydrogenase (CODH),NADPdependent formate dehydrogenase (FDH), and a corrinoid enzyme (Ljungdahl, 1986). These enzymes are metalloproteins, for example, CODH contains nickel, iron and sulfur (Drake et al., 1980; Ragsdale et al., 1983), and FDH contains iron, selenium, tungsten, and a small quantity of molybdenum (Yamamoto et al., 1983), while the corrinoid enzyme (vitamin BIZ compound) contains cobalt (Hu et al., 1984). A. SUBSTRATES FOR FERMENTATION

In most physiological studies conducted on C. thermoaceticum, cells were cultivated in a complex undefined growth medium containing substantial quantities of yeast extract and tryptone. Thus, the true anabolic capabilities of C. thermoaceticum remained unclear for a long time. Lundie and Drake (1984) attempted to establish the basic nutritional requirements of C. thermoaceticum and thus define its anabolic potential and limitations. In developing a minimally defined growth medium, after a series of deletion experiments they found that C. thermoaceticum did not display any specific amino-acid requirement and required nicotinic acid as the sole essential vitamin. Koesnandar et al. (1990) determined the optimum concentration of five metals in the glucose-minimal medium as iron 100 pM, cobalt 400 pM, molybdateGO0 pM, selenite 1pM, and nickel 0.15 pM. C. thermoaceticum is now routinely grown in a rich medium containing glucose, a complex nitrogen source such as yeast extract or corn steep liquor, tryptone, reducing agent, bicarbonate, phosphate, and minerals (Andreesen et al., 1973; Schwartz and Keller, 1982a). This is shown in Table I as the “base-standard” medium. Tryptone was excluded from the culture media by Ljungdahl et al. (1985),although yeast extract was used. However, yeast extract is an expensive ingredient; Ljungdahl et al. (1986)suggested that yeast extract could be replaced by such cheaper materials as yeast autolysates and corn steep liquor. This . has been confirmed by Cheryan and Shah (1996), Witjitra et ~ l(1996), and Shah and Cheryan (1995a), as discussed later.

7

PRODUCTION OF ACETIC ACID BY C. thermoaceticum

TABLE I NUTRIENT MEDIUM FOR CLOSTRIDIUM THERMOACETICUM (g/liter)

Component Glucose Buffering components: KHCO, KZHPO, KHZPO, Yeast extract Salts: (NH,I,SO, MgSO,. 7H2O Fe(NH,),(SO,), . 6H,O COSO,. 7.5HzO Na,WO, 2H,O Na,MoO, 2H,O NiC1, 6H,O ZnSO, . 7H,O Na,SeO, Cysteine . HC1 H,O

Base (standard] medium (1x1

Medium at 2X level

As requireda

As required

9.0

1.8

1.40 1.10

1.40 1.10 10.0

5.00

1.0

2.0

0.25

0.50

0.04 0.03 0.0033 0.0024 0.00024 0.00029

0.08 0.06

0.0066 0.0048

0.000017

0.00048 0.0005 0.000034

0.25

0.25

Vsually 20 g/liter for maintenance and initial growth,

Typical fermentation patterns are shown in Figs. 3 and 4. There is usually a lag period before growth and acetate production commences. This is followed by a sharp drop in cell numbers as indicated by OD measurements, and a simultaneous decrease in the rate of acetate production. Growth and acetate production by C. thermoaceticum is critically dependent on nutrient type and levels. When nutrients are supplied at the “normal” level ( I X concentration shown in Table I), the final acetate concentration and yield are low (22.5 g/liter and 0.77 g acetate/g glucose consumed), as shown in the upper graph of Fig. 3. A large portion of the substrate (glucose) is unutilized. When the nutrient supply was increased to 2X in Experiment 2 (Fig. 3, bottom), the final acetate concentration increased significantly to 35 g/liter. Additional fermentation parameters are shown in Table 11. Increasing the nutrients to 3X and supplying additional glucose during the fermentation resulted in a higher acetate concentration of 47.8 g/liter (Expt. 3; Fig. 4). The concentration of cells also increased with

8

M. CHERYAN et al.

4

Acetate

. 3

2

f X \ I.,

Cells

1

L I

20

40

60

00

100

128

Time (hours) FIG.3. Typical fermentation patterns of C. thermoaceticum using yeast extract and ammonium sulfate as nutrients. Temperature of all fermentations was 6OOC. TOP: Experiment 1 with 1X level of yeast extract (5 g/liter), ammonium sulfate (1g/liter), and salts (see Table I1 for concentrations). BOTTOM: Experiment 2 with 2X concentration of yeast extract, ammonium sulfate and salts (Cheryan and Shah, 1996).

increased supply of nutrients. Maximum cell concentration increased from 1.8 g/liter in Experiment 1to 4.8 g/liter in Experiment 3. Fructose, an intermediate product associated with nutrient depletion or microorganism stress (Witjitra et al., 1996), was also produced when glucose was added the second time in Experiment 3 (Fig. 4) but was gradually metabolized by the end of the experiment. In subsequent experiments (listed in Table 11) the yeast extract concentration was reduced and ammonium sulfate concentration was increased while keeping the concentration of salts fixed at 2X. In Experiment 4 the yeast extract concentration was reduced to 5 g/liter, while other nutrients were supplied in the same concentration as in Experi-

9

PRODUCTION OF ACETIC ACID BY C. thermoaceticum 5

4

i 39

v)

*6 1

0

Time (hours) FIG.4. Fermentation of dextrose by C. thermoaceticum using yeast extract and ammonium sulfate, each at 3X level (see Table 11).Additional glucose was added as shown after 28 hours of fermentation [Cheryan and Shah, 1996).

ment 2. This resulted in acetate concentration and yields of 31.5 g/liter and 0.77 g/g glucose, respectively; both these values were lower than Experiment 2. In addition, fructose was produced as a by-product and some glucose remained unutilized at the end. Increasing the ammonium sulfate concentration to 3 g/liter (3X) caused the acetate concentration to increase significantly to 38 g/liter, improving the yield to 0.86 g/g, with no by-product formation. However, further reduction of yeast extract to 0.5X reduced the acetate concentration to 33 g/liter and the yield to 0.78 g/g; in addition, fructose was produced toward the end of the experiment (Expt. 6, Table 11). Increasing the ammonium sulfate to 3.5X restored performance (Expt. 7: 36 g/liter acetate produced in 71 h, with an acetate yield of 0.86 g/g and no fructose). Thus, it appears that ammonium sulfate could reduce yeast extract requirements by 75% with minimal effects on acetate production and yield. In fact, the productivity was higher, increasing from 0.39 g/liter-h in Experiment 2 to 0.5 g/liter-h in Experiment 7 (Table 11). The fermentation is clearly growth-associated. As shown in Figs. 3 and 4, cell concentration increased exponentially initially and then decreased until the end of the fermentation. Acetate concentration also increased and then leveled off. Acetate productivity was more than 1 g/liter-h initially and declined with the time of fermentation, parallel to the decrease in OD. This emphasizes the necessity for keeping the culture viable and active. A by-product, which appears to be fructose, appeared after the OD started to decrease in almost all cases, especially

TABLE I1 EFFECT OF NUTRIENTLEW ON FERMENTATIONP.m.o.mnsa Experiment no.

Nitrogen source and level*

YE,1x YE,2X YE,3x YE,1x YE,1x YE,0.5X YE,0.5X CSL,1x CSL,2x

SaltsC level 1x 2x 3x 2x 2x

2x 2X

2X 2x

(NHJLQ level

'Erne (hl

1x 2x 3x 2x 3x 3x 3.5x

115 90 125 72 65 100 71 90 95

3x 2X

Acetate (glliter) 22.5 35.0 47.8 31.5 38.0 33.1 36.1 31.1 30.8

Yield of acetate

Yield of fructose

(g/d

(!A4

0.77 0.83

-

0.19

0.01 0.05 -

0.39 0.38 0.43

n

0.58 0.33 0.51 0.35 0.32

z z

0.85 0.77 0.86 0.78 0.84

0.03

0.78 0.82

0.08 0.08

Wtial glucose concentration = 46 goiter. Source:Cheryan and Shah (1996). %E = yeast extract (see Table 1 for IX concentrations). CSL = corn steep liquor (X= 10 g solidsfliter medium). Q o e s not include ammonium sulfate.

-

Productivity (g acetate/ litedh)

5

E

z 4

Y

PRODUCTION OF ACETIC ACID BY C. therrnoaceticurn

11

when the nutrients were inadequate either in quality and/or quantity. High glucose concentration inhibits the initial growth of C. thermoaceticum (Witjitra et al., 1996). However, after adaptation (which takes time and energy, and results in slow initial growth rate and some glucose utilization without acetate production), the fermentation proceeds rapidly. There appears to be a minimum ratio of nutrient concentration to glucose concentration to produce acetic acid. If glucose was still available but not the nutrient, the microorganism produced fructose instead of acetate. Acetate production from glucose by C. thermoaceticum generates five moles of ATP per mole of glucose consumed (Fuchs, 1986). This results in high levels of cell mass per mole of glucose consumed (Andreesen et al., 1973). To maintain productivity, the cells must balance their ATP supply and demand. Since growth consumes more ATP than maintenance, most of the acetic acid produced by C. thermoacetjcum occurs during the growth phase. In all our work, especially those with suboptimal levels of nutrients, a decrease in cell density is observed 24-48 hours into the fermentation, which is followed by a decrease in the rate of production of acetate. Thus, to increase acetic acid production, the cells must be continuously growing. This requires a continuous supply of nutrients, as shown in Experiments 1-3 (Table 11). This can also explain why a large excess of nutrients was necessary to obtain very high (>80 glliter) acetate concentrations (Parekh and Cheryan, 1994a). It also explains the limitations of a continuous cell-recycle bioreactor (see later): merely maintaining higher cell concentrations or using low dilution rates could not produce high acetate concentrations without also proportionally increasing the nutrient concentration. When cells use yeast extract as a source of amino acids, nucleotides, and fatty acids, they will need less ATP than if they have to synthesize these compounds using ammonium ions as the starting material. Thus, assimilation of ammonium ions is important if cells are to recycle the ATP generated during production of acetic acid. Therefore, much smaller amounts of ammonium sulfate could replace yeast extract (e.g., compare levels in Expts. 2 and 7, Table 11). However, since yeast extract contains growth factors in addition to nitrogen, it could not be completely substituted by ammonium sulfate. It appears that acetate production could be sustained only with sufficient ATP-consuming reactions taking place in the cells. If sufficient nutrients are not supplied, fructose was produced (e.g., Expts. 4, 6, 8, and 9). By increasing ammonium sulfate (e.g., Expts. 5 and 7), by-product formation was avoided and acetate yields were good (0.84-0.86).

12

M. CHERYAN et a].

Thus, by substituting the much cheaper ammonium sulfate for yeast extract, the ratio of acetate produced to yeast extract needed can be substantially increased. As will be seen later, the medium cost could be lowered further by substituting corn steep liquor for yeast extract. C. thermoaceticum grows well on pyruvate as the primary substrate (Barker, 1944; Andreesen et a]., 1973). Parent (wild) strains of C. thermoaceticum did not grow on DL-lactate but an adapted strain grew on both D- and L-lactate with a yield of acetate of 0.95-1.00. Both strains grew on and consumed lactate when pyruvate, glucose, fructose, or xylose was used as the second substrate (Brumm, 1988). It is important to maintain a low level of the carbon source in the fermenter at all times to ensure viability of the cells (Shah and Cheryan, 1995b). By using different concentrations of externally added sodium salts to the growth media, the relative growth inhibition caused by the anion was found to be in the order of acetate > chloride > sulfate. Various externally added cations of acetate were also examined, and the relative magnitude of inhibition on the growth rate was found to be ammonium > potassium > sodium (Wang and Wang, 1984). This could be specific to a particular strain (e.g., DSM 521 used by these investigators), since recent work has shown that strain ATCC 47907 could successfully produce ammonium acetate up to 50 glliter acetic acid (Cheryan and Shah, 1996). B . MINERAL REQUIREMENTS

The fact that metal is involved in acetate production explains why metal is needed in the culture media. Andreesen et al. (1973) found positive benefits of metals (ferrous, molybdate, and calcium) on growth yield, enzymes, and synthesis of acetate from CO,. Formate dehydrogenase was the only enzyme of those assayed that was affected by the addition of metals to the growth medium. Addition of selenite and molybdate or tungstate to the growth medium stimulated the formation of formate dehydrogenase during growth of C. thermoaceticum (Andreesen and Ljungdahl, 1973). Later, Ljungdahl and Andreesen (1978) demonstrated that, although their strain of C. thermoaceticum could grow without including selenite, tungstate, or molybdate in the medium, high formate dehydrogenase activity was obtained only when these metals were present in the medium. Shoaf et al. (1974) found that NH: or K+ but not Na+ increased the thermostability of formyltetrahydrofolate synthetase from C. thermoaceticum, but that phosphate ions inhibited the enzyme, and this inhibition was stronger in the presence of NH:. Nickel (Ni2+)has no effect

PRODUCTION OF ACETIC ACID BY C. fhermoaceticum

13

on the growth of C. thermoaceticum, but less carbon monoxide dehydrogenase was formed when Ni2+was omitted from the medium (Diekert and Thauer, 1980). Increasing cobalt by fourfold (to 400 pl4) did not affect cell growth, glucose consumed, acetate produced, or its molar yield, but it did increase corrinoid production (Koesnandar et al., 1990). Cysteine is used in the medium as a source of sulfur (Koesnandar et a]., 1990) and as a reducing agent. Since cysteine is expensive, studies have been conducted with alternate low-cost sulfur-containing reducing agents (Shah et al., 1996). It appears that the recommended dosage of 0.25 g/liter of cysteine . HC1. H,O is probably too much, at least for strain ATCC 47907 with yeast extract as the complex nitrogen source. Decreasing it to 0.05 g/liter actually improved the final acetate concentration in the broth while simultaneously reducing the maximum OD in the fermentation broth by 50%. Thus, the increase in acetate yield may be a result of a decrease in cell mass yield that allowed more carbon to be channeled into acetate production. Cysteine could also be successfully replaced with sodium thioglycolate and sodium sulfide (Na2S). The sulfur source should not be eliminated completely: with no sulfurcontaining reducing agent, acetate and cell concentrations were lower (Shah et al., 1996). IV. Strain Improvement

Better strains of C. thermoaceticum were developed by Schwartz and Keller (1982a,b) that could grow and produce acetic acid at pH 4.5 (available as ATCC 31490). Reed et al. (1987) developed strains tolerant to high acid-low pH conditions to allow recovery of product in the acid or undissociated form rather than the salt form. An improved strain of ATCC 39289, an acid-tolerant strain, produced 29 g/liter acetate in 140 hours at pH 6.65 and could grow in the presence of 70 g/liter sodium acetate (Parekh and Cheryan, 1990a). In a fed-batch mode, this particular improved strain performed well (Parekh and Cheryan, 1990b). Several other strains of C. thermoaceticum have also been screened (Ljungdahl, 1983; Ljungdahl et al., 1985; Wang and Wang, 1984). Ljungdahl et al. (1986) reported on a particular mutant strain that could apparently produce high levels of acetate (up to 100 g/liter acetate or more). However, no further studies with that strain have been reported. Parekh and Cheryan (1991) developed several strains by treatment with chemical mutagenic agents (NTG and nitrous acid) and selective enrichment procedures. Mutagenesis yielded several mutants, some of which exhibited growth at pH 5.6 and above, as well as acetate yields

14

M. CHERYAN et al.

above 0.8 g acetate per gram pyruvate, when 10-20-mM pyruvate was supplied as the sole energy source. Table 111 summarizes the behavior of a few mutants that showed consistent growth when screened in selective media. All strains exhibited growth in media containing up to 5% acetate at pH 6.6 while producing an additional 1.0% acetate with yields (Yp,s) of 0.8 or better. Prior to mutagenesis, the parent strain (C. thermoaceticum ATCC 39289, a spontaneous mutant isolated by Schwartz and Keller (1982b)by repeated culturing at low pH) could be grown and cultivated in media at pH 6.4 and above containing 5% acetate or 10-mM pyruvate or formate. However, media containing 1% acetate at pH below 5.3 inhibited growth and repressed acetate production. In contrast, two mutant cultures, G-10 and 5-40, showed visible signs of growth after 140 h of cultivation in media containing 1% acetate at pH 5.3. However, these cultures could not be subcultured regularly in the same 4% acetate media at pH 5.3 and attain good growth. Mutant strain G-20 was the only one that appeared to grow when cultivated at pH 5.6 in 50-mMpyruvate as the sole energy source (Table 111). Subsequent experiments showed it could grow and attain acetate yields of 0.81 in media containing 80-mMpyruvate controlled at pH 5.5 and incubated for 4 days. The surviving cells could be readily retransferred to an 80-mMpyruvate medium and showed visible growth in 72 h. Cells from this culture could be subcultured regularly in 80-mM pyruvate without difficulty when the pH was controlled at pH 5.6. In addition, culture G-20 could also tolerate dolime concentrations as high as 6% when cultured at pH 6. Wiegel et al. (1991) also reported that their C. thermoaceticum mutants derived after EMS treatment showed excellent tolerance to dolime. A similar procedure with the other mutants did not result in any growth when the media contained over 20-mM pyruvate (Table 111). The rationale of using pyruvate- and formate-containing media was that these compounds play a fundamental role in the metabolism of acetogenic bacteria (Ljungdahl, 1983). Pyruvate is the precursor of acetate in the pyruvate dehydrogenase reaction, and it is also the source of the “CO” unit in the CO-Ni-E complex. This unit also plays an important role in synthesis of acetate from COz. Similarly, formate is the precursor of the methyl group in acetate synthesis via the autotrophic pathway. This approach has yielded several mutants with better acetate producing capabilities (Brumm, 1988; Wiegel et al., 1991). Pyruvate and formate inhibited growth of some mutants, especially at pH 5.5 and below (Parekh and Cheryan, 1991; Wiegel et al., 1991). While some strains could grow in the presence of 10-mM pyruvate at pH 5.6 (Table 111), they lacked tolerance to 50-mM pyruvate and dolime

z0

TABLE 111

aL

SELECTION CONDlTIONS'

Glucose/pyruvate

Pyruvate (pH 6.5)

Strains

pH 5.5

pH 6.5

10 mM

Parent strain

-

+

+

50mM

-

2

Pyruvate (pH 5.61 1OmM

20 mM

+

-

50mM

2 80mM

-

-

$ > n

Mutant strainb G-10 G Z O 640 N-10

N-40 5-40 s-100 0

- - no growth or acetate production; + = growth and acetate production. Source: Parekh and Cheryan (1991).Reprinted with permission

( I -

from Springer-Verlag. bc = NTG treatment;N = nitrous acid treatment; S = EMS treatment. Numbers refer to the time (minutes) of exposure of cells to t h e reagent.

a

16

M. CHERYAN et

01.

and did not show any improvement in acetate production (Parekh and Cheryan, 1991). Only G-20 could grow in 80-mM pyruvate and 50-mM formate media when used as the sole energy source (Table111). Attempts to grow this mutant in higher-pyruvate media and at low pH (below 5.4) were unsuccessful. Growth resumed in media with pH above 5.6. In addition, even though the parent strain was unable to grow in 10-mM pyruvate media at pH 5.3, it metabolized pyruvate in the presence of glucose when cultivated above pH 6.5. Some C. thermoaceticum strains utilize mixed substrates (glucose and lactate or formate in a 1:l ratio) above pH 6.5 independently of the starting substrate (Brumm, 1988). The ability of this mutant to grow and produce acetate below pH 6 makes it useful in production of industrial acetates such as CMA, since it would assist in the soluhilization of dolime and in the recovery of soluble CMA. When evaluated in a fed-batch fermentation system, this mutant strain performed better than the parent strain (Fig. 5): 18% higher concentration of acetate and 25% faster rate compared to the parent strain (Parekh and Cheryan, 1991). Experiments with this mutant strain have been repeated and carried out for several years in laboratory and pilot scale (up to 40 liters) in a variety of bioreactors (batch, fed-batch, multiple-stages, cell-recycle, and membrane) operated up to 54 days continuously and producing several salts of acetate (ammonium, calcium, calcium-magnesium, potassium, and sodium). No alterations in stability or performance were observed. This strain has produced as much as 10% CMA in a fed-batch reactor in 140 hours, with 93% substrate utilization and acetate yields of 0.80 g/g glucose (Parekh and Cheryan, 1994a). Such robustness is important for industrial production of bulk chemicals. This mutant culture is available as ATCC 49707 and DSM 6867. V. Low-Cost Media

The superior performance of mutant strains of C. thermoaceticum comes at a price: substantial quantities of high-quality yeast extract are needed and fermentation times are long, resulting in low productivity. This leads to high fermentation cost, For example, with a yield of 0.85 g/g and a glucose cost of $0.16 per kilogram, the cost of CMA would be $2.49 per kilogram if even the lowest-cost commercial yeast extract ($8.50 per kilogram) is used. This far exceeds today’s price of petroleum-derived CMA of $0.70-0.80 per kilogram. Thus, the cost of nutrients must be reduced, while maintaining a reasonably high concentration of acetate and good productivity.

17

PRODUCTION OF ACETIC ACID BY C. thermoaceticum 70

i

Calcium-mepesium acetate

2 20

70 h

20

;

10

y

0

c

z a

10

(I)

0

0

0

50

100

150

0

50

100

150

200

Fermentation time (hours) FIG.5. Fed-batch production of acetate by improved and mutant strain G-20 of C. therrnonceficurn.pH was maintained at 6.3 f 0.5 by the addition of 20% high-magnesium lime, LEFT: Calcium/magnesium acetate produced (expressed as acetic acid concentration). RIGHT: Glucose consumed during the fed-batch fermentation (Parekh and Cheryan, 1991). Reprinted with permission from Springer-Verlag.

Table IV is a list of the various nutrients studied by Witjitra et al. (1966). Bacto yeast extract from Difco is universally recognized as an

excellent stimulator of bacterial growth. With C. thermoaceticurn fermentation, it results in excellent growth and acetogenesis, especially at concentrations of 15-30 g/liter or more (Parekh and Cheryan, 1991, 1994a). It is not clear which components of yeast extract are important. Lundie and Drake (1984) suggested that nicotinic acid is the sole essential vitamin for C. thermoaceticum: yeast extract is rich in water-soluble vitamins, including nicotinic acid (Bridson and Brecker, 1970; Difco Laboratories, 1984). Although hydrolyzed cottonseed is an excellent nutrient source for Lactobacillus amylovorus (Cheng et al., 1991), C. thermoaceticum showed no growth or acetate production even after 100 h of incubation. It might lack the component[s) required for the growth of C, therrnoaceticum, or it might have contained a component from the cottonseed that inhibited its growth (e.g., gossypol). The fermentation pattern with hydrolyzed corn gluten meal was similar to that observed with yeast extract except that much less acetate ( 4 5 g/liter) was produced. It is possible that corn gluten meal contains a growth inhibitor for C. thermoaceticum that might explain the lag phase. For example, phytic acid, a component of corn, binds strongly with minerals, making them biologically unavailable as nutrients (Cheryan, 1980).

TABLE TV

C o ~ p o m OF o ~NUTRIEN~ SOURCE(% wtlwt)"

Hydrolyzed pratein nutrients Heavy

Corn gluten

N-&Soy

stillagd

3.4

3 .O

4.4

2.9

2.6

2.4

8.7

10.1

13.7

93.0 na 0.39

na

na na

na

0.28

na

0.14

CottonsMd

Moistum a-amino nitrogen Total nitrogen Fat Fiber Lactic acid

Ash

na

-

-

corn

Uefatted

Difco

steep liquo$

SOY

yeast

flow

exkdct

50.0

7.0

5.5

na

7.0

8.3 na

na 1.84 1.0 na

1.47

11.3

-

9.5 na nn na

7.3

6.0

10.0

0.11

0.01

na

4.3

6.1

6.3

3.4 0.36

5.7

Sodium

8.1. 0.69

0.04

0.56 0.03

PH

6.3

6.0

6.8

4.3

"Source:Witjitra (1994). "Received in a concenhted liquid form. All others were in solid powdered form. na = data no1 available.

3.0

g

Fi 5z a

n

!-

PRODUCTION OF ACETIC ACID BY C. thermoaceticum

19

The performance of the corn refining by-products was also good, although some pretreatment may have to be given to them to optimize their utilization. “Stillage” is the liquid residue after the ethanol has been stripped away in the beer well of a yeast-based ethanol fermentation plant. The solids consisted mainly of dead yeast cells, traces of ethanol, low-molecular-weight sugars, organic acids (lactic, succinic), glycerol, and amino acids. With untreated stillage instead of yeast extract at 10 g soliddliter (Fig. 6), there was a lag period at the beginning of the fermentation and fructose was produced instead of acetate. When growth began, fructose levels dropped and acetate was produced. When cell concentration started to decrease, fructose started accumulating again. Fermentation was better when the stillage was filtered before being added to the media (Fig. 6). Once again a clear correlation between decreased cell concentration, decreased acetate production rate, and appearance of fructose can be observed in the data. Filtration appeared to eliminate a growth inhibitor in stillage. When the filtered stillage was used in the medium without dilution, that is, at a level of 64 g solids/liter, the fermentation profile was also very good (Fig. 6). There was slow growth initially but no fructose was produced during this period. After adapting itself, the culture continued to grow and produce acetate. When the cells started lysing and decreasing in numbers (after 160 h), the same phenomenon occurred, that is, subsequent production of fructose. The high concentration of nutrients in the undiluted stillage was obviously sufficient for extended cell growth and production of acetate, but perhaps too concentrated to be used in the initial stages, since there appeared to be some inhibition initially. A better method of utilizing stillage would be in a fed-batch mode, where the stillage is added with the glucose as needed to sustain viability and productivity. The yield with the undiluted filtered stillage was 1.07 g acetate per gram glucose (Table V). This is because there was lactic acid and sugars in the stillage that were probably utilized by the organism to produce acetate (Brumm, 1988). Corn steep liquor (CSL) contains soluble components leeched out from the corn during soaking. It is rich in organic nitrogen (4446% protein on a dry matter basis), with about half the nitrogen as free amino acids; the balance is small peptides with very little intact protein (Christianson et al., 1965). It contains relatively high levels of several important vitamins, trace elements, and lactic acid (10-30%, dry basis); the latter can be utilized by C. fhermoaceficum, thus increasing the yield of acetate. It has been industrially used as a nutrient in ethanol fermentation and for production of pharmaceuticals. In acetate fermen-

20

M. CHERYAN et al. 40

thewnoaceticum

35

.

Stillage, g/L Unfiltered.1 C Filtered, 10

b

._ € 3 1

U '

0

'

'

'

50

100

"

'

,

150

200

'

.

250

Time (hours) FIG.6. Fermentation of glucose by C. thermoaceticum using corn-refining by-products as nutrients. Open points = with corn steep liquor (CSL)at 10 g soliddliter. Closed points = with stillage at concentrations shown on dry solids basis. (Adapted from Witjitra et al., 1996.)

tation, yield of acetate was similar to fermentation using yeast extract, but less acetate was produced with corn steep liquor, probably due to lower concentration of essential nutrients. Fructose began to accumulate as soon as acetate production stopped (Fig. 6). Increasing the concentration of CSL improves the fermentation, as shown in Fig. 7. Pretreatment of the CSL with dolime and filtering out the precipitate also is beneficial, possibly by removing the phytic acid (Shah and Cheryan, 1995a). Figure 7 shows the effect of different levels of pretreated and filtered CSL on the fermentation pattern. The 1 X experiment was conducted with corn steep liquor at 5 g/liter (dry basis), salts at the concentration shown in Table I, and added vitamins. The final acetate concentration was only 19 g/liter, and cell concentration

TABLE V FERMENTATION s-P FOR C. THERMOACETICUMWITH VARIOUS NUTRIENTS AT A CONCENTRATION OF 10 g Soms/LITER OF F~RMENTATION BROTH,UNLESS OTHERWISE STATED'

Glucose utilizationb Nubient Yeast extract, Difco Hydrolyzed corn gluten N-Z-Soy@BL Hydrolyzed soy flour Corn steep liquor Stillage, unfiltered Stillage, filtered Stillage, filtered (64g/liter)

(%I 91 46 55 90

aa 57 63 74

Acetate produced (g/liter) 34.6 14.5 14.1 33.9 31.1 17.6 19.9 36.0

Fructose produced (g/liter)

0 U

5

g

Yield (S/d

Productivity (g/liter-h)

-

0.80 0.72

0.47 0.15

0.7

0.57

0.29

>

0.79 0.83 0.68 0.71 1.07

0.39 0.32 0.15 0.21 0.15

.c 9

-

1.5

4.5 4.9 2.7

2

> R

=! n

8m

"Source: Witjitra (1994). bGlucose in media initially was 50 glliter.

N Y

22

M. CHERYAN et al.

Time (hours)

FIG.7. Effect of CSL concentration on fermentation of glucose (initial glucose = 46 g/liter). Open points = dolime-treated and filtered. 1 X is CSL at 5 g dry solids/liter. Closed points = No treatment (2x1. Medium was supplemented with thiamine (0.15 glliter), riboflavin (0.35 glliter), pyridoxine (0.175 g/liter), nicotinic acid (3 g/liter), and pantothenic acid (1g/liter). (Adapted from Shah and Cheryan, 1995a).

reached only 0.9 g/liter before decreasing. Doubling the CSL and salts doubled the maximum cell density and increased acetate level to 3 1 g/liter. At 2.7X concentration of CSL and salts, the final acetate level was 38.5 g/liter and maximum cell density was also three times the cell density at 1X. The importance of proper pretreatment of the CSL can be seen in Fig. 7. At the 2X level, untreated (i.e., no dolime treatment and no prefiltration) CSL resulted in only 26 g/liter of acetate and a much lower productivity than with pretreated CSL.

PRODUCTION OF ACETIC ACID BY C. thermooceticum

23

Table I1 shows the interactions when CSL and ammonium sulfate in the medium were varied. Experiments 2 and 9 are similar except that CSL was substituted for the yeast extract. Acetate concentration was lower, but yields were similar. Experiments 4 and 8 were also similar in design and results. Fructose was produced in all CSL experiments. However, as will be seen later, adding additional CSL during fermentation (e.g., by fed-batch operation) caused the fructose to disappear and be replaced with more acetate. Low-hydrolysis soy flour is also a promising low-cost nutrient. Soybean meal is the residue from soybeans after the extraction of the oil. It is a complex mixture of protein (50% dry basis), carbohydrates (oligosaccharides and fiber 30%), fat (l%), and lecithin (1.8%).It is a good source of nutrients for industrial fermentation, especially for antibiotics (Crueger and Crueger, 1990). Enzymatically hydrolyzing the protein, even to a limited extent, has a beneficial effect on fermentation by making the nitrogedprotein in soy meal more available, and perhaps releasing other bound nutrients. It is possible that the residual lipids and/or lecithin in the soy flour is the key nutrient. Lipids are essential components of the membranes of Clostridia. Baumann et al. (1965) reported that about 4.5% of the dry weight of log phase cells of C. butyricum is lipid; out of this, 70% is phospholipid. The lipid material in the soy meal could contribute to the stability of the cell wall during fermentation, reducing the rate of cell lysis and allowing better fermentation. It is worth noting that no fructose was detected with low-hydrolysis soy flour as the nutrient. On the other hand, a high degree of hydrolysis [with the resultant high levels of free amino acids) yielded less product than the less extensively hydrolyzed soy flour. Similar results were observed with L. amylovorus (Cheng et al., 1991) and L. bulgaricus (Leh and Charles, 1989). Apparently, certain microorganism may be stimulated by peptides of a certain length or of a certain amino-acid composition. Since hydrolyzed soy flour, corn steep liquor, and ethanol stillage cost less than $0.50/kg each, their use will significantly reduce the overall nutrient cost and the cost of fermentation-derived acetate. The economic significance of using stillage as a nutrient source goes beyond the acetate industry. At present, the economic value of stillage, after evaporation and drying, is marginal at best (moisture removal of stillage requires substantial amounts of energy). If, on the other hand, stillage is used as the nutrient source for acetate fermentation, there will be little or no stillage handling costs. This will improve ethanol economics while simultaneously improving acetate economics by making available a good nutrient source at almost no cost, except for the filtration (which

24

M. CHERYAN et al. 60

1

2

3

4

5

4

67

-

C

.-4-0 m

-2 -

-1

b C

0 0 C

u"

Time (hours) FIG.8. Fed-batch fermentation with 3X levels of CSL and salts. Arrows indicate addition of nutrient solutions in the following amounts: 1,2, and 3 = 50 ml of solution A (CSL and salts 6.6X, glucose 260 g/liter); 4 = 100 ml of solution B (CSL and salts 10x1; 5 and 6 = 50 ml of solution B; 7 = 20 ml of solution C (glucose 500 g/liter). Initial volume = 1.0 liter, final volume = 1.47 liter. From Shah and Cheryan (1995a). Reprinted with permission from The Macmillan Press Ltd.

removes the growth inhibitors and suspended solids and simultaneously sterilizes the stillage). VI. Bioreactors for Improving Productivity

Much of the work done with this fermentation has been done with batch fermenters, where all the carbohydrate and nutrients are added at the start of fermentation. With fermentations that are substrate-inhibited, a better method is to use the fed-batch mode of operation. This significantly improves the performance of C. thermoaceticum fermentation (Parekh and Cheryan, 1990b, 1994a). Figure 8 shows a typical fed-batch fermentation using CSL as the complex nitrogen source. Since it is substrate-inhibited, the initial glucose concentration was low (20 g/liter), while CSL and salts were at the normal 1 X concentration shown in Table I. Subsequently, about 24 hours into the fermentation, when the OD decreased, indicating a decrease in viable cell numbers, more nutrients and glucose were added (in concentrated form to minimize volume changes). Eventually, after 70 hours, cell numbers went up again, As before, fructose was produced during the death phase of the cells. But when cell viability improved the fructose disappeared, apparently being converted into acetic acid. At the end of the fermentation, there was no fructose in the broth. This pattern has been repeatedly observed in the fed-batch mode (Shah and Cheryan, 1995a).

PRODUCTION OF ACETIC ACID BY C. thermoaceticum

25

Alkali PH controller

1-1

1 cr”----i - -*El&

Prod&

(Permme)

FIG.9. Schematic of a continuous bioreactor using membrane separations for cell recycle.

Continuous fermentation (Sugaya et al., 1986) and immobilized whole cells have been used to increase the productivity of homoacetogenic fermentations (Reed and Bogdan, 1985; Wang and Wang, 1983). However, with Ca-alginate-immobilized cells of C. thermoaceticum, steady-state performance was not achieved when cultured at high acetate concentrations (Wang and Wang, 1983). On the other hand, cell-recycle bioreactors using a membrane module as the separation device have been shown to vastly increase the productivity of several anaerobic fermentations, such as ethanol and lactic acid (Cheryan and Mehaia, 1986) and may have some advantages over immobilized cells, such as higher concentration of free cells, no diffusion limitation, excellent mixing in the bioreactor, and a cell-free product stream. The biggest advantage is that cell concentrations far in excess of normal levels can be used with no danger of cell washout. For example, yeast concentrations of 100 g/liter can be used for production of ethanol from glucose (equivalent to 1011-1012 cells/ml). This is far greater than the 10-20 g/liter ( lo8 celldml) normally used in industrial fermentations. These higher cell concentrations result in much faster fermentations, thus vastly improving productivity (Cheryan, 1986). Figure 9 is a schematic of a continuous cell-recyle membrane bioreactor. The reaction vessel of the fermentation system is coupled in a semi-closed loop configuration to an ultrafiltration or microfiltration module of the appropriate chemical nature and physical configuration (Cheryan, 1986). The reaction vessel is initially charged with the cells

26

M. CHERYAN et al. 1 .o

2.0

=:

\ 1.5 (r

v

.-> .+d

c

0.9

0.8

2\

0.7

1.0

0

4-l

0.6

0

: a

-2

0.5

.E >-

0.5 0.4

0.0

1 301

--. -A

5 20

0

GI u c 0 s e,*A

,

l 0 L 4 0 0.005 0.010 0.015 0.020 0.025 0.030

Dilution r a t e (h-')

FIG.10. Fermentation of glucose by C. thermonceficum in a membrane recycle bioreactor. Cell concentration (X) = 17 g/liter, glucose concentration (S) = 58 g/liter. Yeast extract and salts were at the 2X level. Data taken after at least 5 volume changes. From Parekh and Cheryan (1994b). Reprinted with permission from Elsevier Science Inc.

and adjusted to the required cell concentration. Feed is then pumped into the reaction vessel and product is removed as permeate at the same rate, thus keeping the volume in the bioreactor constant. For growth-associated fermentation, some provision has to be made to bleed excess cells to avoid pumping problems. Figure 10 shows typical results obtained with such a bioreactor with C. thermoaceticum at a cell concentration of 1 7 g/liter (normal cell concentrations in batch and fed-batch fermentations are 2-4 g/liter). An increase in dilution rate (i.e., a decrease in residence time) resulted in a decrease in acetate concentration, a decrease in acetate yield, and an increase in unutilized glucose (Parekh and Cheryan, 1994b). High dilution rates also resulted in the appearance of a by-product (not shown in Fig. 10; the authors suggested the by-product was lactate, but it could have been fructose). The long-term stability of this bioreactor was very good with this strain of C. thermoaceticum. Interestingly, the usual bell shape of the cell-density curve observed in batch operations was not present: no loss

PRODUCTION OF ACETIC ACID BY C. thermoaceticum

27

Cells 0

50

100

150

200

260

300

350

400

Time (hours)

FIG.11. Performance of a two-stage continuous-membrane bioreactor. The membrane was attached only to the second stage. The feed was pumped into the first stage reaction vessel. Broth from the first stage was pumped continuously into the second vessel at a rate to keep fermentation volumes constant. Fermentation broth from second stage was recycled through the membrane module. The cell-free permeate was removed at the same rate as the feed flow rate to the first stage. The retentate containing the cells was recycled back to the first stage at 25% of the feed flow rate to the first stage, and the remaining was recycled to the second stage. Dilution rate is based on the total volume used in the two stages. From Shah and Cheryan (1995h). Reprinted with permission from Humana Press Inc.

of viability or vitality was noted during the 54 days of continuous operation. Microscopy examination did not indicate any dramatic change in the external morphology (cell size or shape). The recycled cells were isolated and preserved in 50% glycerol broth at -20°C. On retrieving, the culture did not demonstrate any change in the rate or efficiency of acetate/CMA production, suggesting that the mutant culture is stable (Parekh and Cheryan, 1994b). Similar microbial stability was shown with a 2-stage membrane bioreactor in which the membrane was attached only to the second stage and a part of the retentate containing the cells was recycled to the first stage. As shown in Fig. 11, with a cell concentration of 4.5 g/liter in the first stage and 12.5 g/liter in the second stage, a dilution rate of 0.02 h-l resulted in acetate concentrations of 25 and 37.5 g/liter in the first and second stages, respectively. This outlet concentration was slightly better than a one-stage membrane reactor (Shah and Cheryan, 1995b). When the dilution rate was increased to 0.033 h-l, the two-stage bioreactor resulted in lower acetate concentration, but productivity was higher. In addition, the cell concentration increased to 15.5 g/liter. These data

28

M. CHERYAN et a].

Ok

56

160

160

260

’

260

300

’

361

Time (hours) FIG.12. Performance of a draw-and-fill bioreactor. At the end of the fermentation, 80% of the fermentation volume was withdrawn and clarified with a cross-flow microfiltration membrane. The retentate containing the cells was recycled to the fermenter, which was then recharged with fresh substrate and fermentation allowed to continue. “Acetate” is expressed in terms of acetic acid. pH was maintained at 6.2 with ION NaOH. From Shah and Cheryan (1995b). Reprinted with permission from Humana Press Inc.

confirmed previous reports that high productivity and high product concentration are mutually exclusive in such high-rate fermenters (Cheryan, 1986; Parekh and Cheryan, 1994b). The yield of acetate was 0.85-0.9 g/g glucose consumed. A “draw-and-fill” bioreactor in combination with a membrane appeared to be the optimum design (Shah and Cheryan, 1995b). In this design, the reaction vessel is operated as a batch fermenter. At the end of the fermentation, a portion of the fermentation broth is withdrawn through the membrane module. The cells are recycled and the reaction vessel charged with fresh substrate. Figure 1 2 shows a series of such draw-and-fill operations with 80% removal of the volume in each cycle. With feed glucose concentrations of 50 g/liter, fermentation times were usually about 34 hours, acetate concentration (expressed as acetic acid) was 38 g/liter, and overall productivity was 0.93 g/liter. There was a lag in acetic acid production at the beginning of each cycle. Also, in each cycle of fed-batch operation, the cells passed through a growth phase, followed by a significant decrease in OD, probably due to cell lysis, caused perhaps by nutrient depletion. These characteristics were similar to those observed in conventional batch fermentations. However, the average cell concentration increased in later cycles, which allowed the fermentation to be completed faster (every 24 hours).

PRODUCTION OF ACETIC ACID BY C. thermoaceticum

I I

Sugars

'

1

29

Nutrients ,

Nutrient recycle

Water recyle

f

Concentrated Organic Acid

Nutrient r e c y c l e d Alkali recycle

FIG.13. Possible downstream processing of acetate fermentation broths by membrane technology. MF is cross-flow microfiltration, NF is nanofiltration, ED is electrodialysis, LR is low-rejection membranes, and HR is high-rejection membranes.

In batch fermentation without cell recycle, acetic acid production is proportional to the amounts of yeast extract and trace salts supplied in the medium. For all types of bioreactors studied, increasing dilution rate increases volumetric productivity but decreases specific productivity (g acetate produced per gram cells). Thus, in cell-recycle bioreactors the nutrient supply should be increased in proportion to cell concentration to realize the full potential of the microorganism. VII. Downstream Processing of Acetate Fermentation Broths

Downstream processing refers to the series of unit operations used to isolate, purify, and concentrate the product. Downstream processing often determines the economic feasibility of the process. One possible downstream processing scheme for acetate utilizes membrane separations technology, as shown in Fig. 13. The first operation is cell separation, which can be done by cross-flow microfiltration. Cell harvesting by membranes is rapidly replacing conventional filtration and centrihgation techniques. When combined in a semiclosed loop configuration to the bioreactor or fermenter, it becomes a powerful tool to dramatically improve the productivity of the fermentation, while simultaneously

30

M. CHERYAN st al.

providing a cell-free broth for subsequent downstream processing (Cheryan, 1986). Depending on the physical and chemical nature of the fermentation products, the cell-free broth is subjected to chromatography, electrophoresis, crystallization, precipitation, extraction, distillation, and/or membranes. Solvent extraction with azeotropic distillation is the preferred method for chemically derived acetic acid, while freeze concentration is used for vinegar. Both require substantial amounts of energy since a change in phase of the solvent is required. Furthermore, if the acetate is required in the free-acid form, there will be additional cost to convert the salt form produced in the anaerobic fermentation to the free-acid form. Among the membrane techniques, electrodialysis and nanofiltration are particularly useful for separating and partially concentrating acetates. A relatively new membrane technology, nanofiltration (NF) can separate charged compounds from noncharged ones and from each other, depending on the relative sizes of the ions and the degree of dissociation (Raman et d., 1994). In Fig. 13 the cell-free broth is first passed through a low-rejection (LR) NF membrane, which separates the acetate from most of the rest of the broth components, including the sugars and many of the nutrients. The substantially purified, but dilute, acetate solution then has to be concentrated. If the acetate is in the salt form, it can be either evaporated or possibly processed through high-rejection (HR) membranes to partially concentrate it prior to evaporation and/or drying. Several NF membranes have recently been screened for the separation of acetic acid (Han and Cheryan, 1995), and preliminary economic calculations suggest it is an attractive technique for this purpose (Han and Cheryan, 1996). Electrodialysis (ED) is a membrane-separation process that separates and concentrates charged compounds from liquid feed solutions by transport under the application of electrical energy and through anionor cation-selective membranes. Ultimately, the ED unit could be coupled to a high-performance membrane bioreactor that would integrate fermentation and separation steps for continuous production of acetates, as shown in Fig. 13. The first stage of the ED could be done with conventional anion- and cation-exchange membranes as a prepurification step, followed by a second stage with bipolar membranes that would generate the alkali and the free acid form of acetate. This would substantially reduce the cost of alkali needed for fermentation as well as reducing waste treatment costs. ED has been found to be particularly useful in concentrating vinegar, for example, to save on transportation costs (Chukwu and Cheryan, 1996).

PRODUCTION OF ACETIC ACID BY C. thermoaceticum

31

In summary, the industrial production of acetic acid by fermentation using Clostridium thermoaceticum appears to be feasible. The mutant strain ATCC 47907 is especially promising. It is a robust organism that can be adapted to a variety of nutrient sources. Considerable research has been done to lower costs by adapting the culture to corn steep liquor and ammonium sulfate as nitrogen sources instead of yeast extract, by reducing the level of reducing agent (cysteine) or by substituting it with a cheaper source such as sodium sulfide, and by using fed-batch fermentation systems in combination with cell recycle by cross-flow microfiltration. Of the material costs, the sugar (e.g., dextrose) will cost about $150/ton of acetic acid and the nutrients should account for only $20/ton. Considering that acetic acid from petroleum sources sells for $550-660/ton in 1996, it indicates good potential for the fermentation process to provide a significant share of the market. ACKNOWLEDGMENTS

Research on acetate production by Clostridium thermoaceticum in the first author’s laboratory has been supported by the Illinois Corn Marketing Board, the Minnesota Corn Promotion and Research Council, the U S . Department of Agriculture through the NRICGP program, and the Illinois Agricultural Experiment Station at Urbana-Champaign. REFERENCES Agreda, V. H., and Zoeller, J. R. (1993). “Acetic Acid and Its Derivatives.” Dekker, New York. Allgeier, R. J., Nickol, G. B., and Connor, H. A. (1974). Food Prod. Dev. 8(6),50-56. Andreesen, J. R., and Ljungdahl, L. G. (1973). f. Bacteriol. 116, 867-873. Andreesen, J. R., Gottschalk, G., and Schlegel, H. G. (1970). Arch. Mikrobiol. 72,154-174. Andreesen, J. R., Schaupp, A,, Neurauter, C., Brown, A., and Ljungdahl, L. G. (1973). J. Bacteriol. 114, 743-751. Anonymous. 1996. Chem. Eng. News 74(15), 15-19. Balch, W. E., Schoberth, S., Tanner, R. S., and Wolfe, R. S. (1977). Int. f . Syst. Bacteriol. 27, 355-361.

Barker, H. A. (1944). Proc. Natl. Acad. Sci. U.S.A. 30,88-90. Barker, H. A., and Kamen, M. D. (1945). Proc. Nut. Acad. Sci. U.S.A. 31,219-225. Baumann, N. A., Hagen, P.-O., and Goldfine, H. (1965). f . Biol. Chem. 240, 1559-1567. Braun, M.,Mayer, F., and Gottschalk, G. (1981). Arch. Microbiol. 128, 288-293. Bridson, E.Y., and Brecker, A. (1970). In “Methods in Microbiology” (J. R. Norris and D. W. Ribbons, eds.), Vol. 3A, pp. 229-304. Academic Press, New York. Brownell, J. E., and Nakas, J. P. (1991). J. Ind. Microbiol. 7, 1-6. Brumm, P. J. (1988). Biotechnol. Bioeng. 32, 444-450. Busche, R. M., Shimshick, E. J., and Yates, R. A. (1982). Biotechnol. Bioeng. Syrnp. Ser. 12, 249-262.

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Cheng, P., Mueller, R. E., Jaeger, S., Bajpai, R., and Iannotti, E.L. (1991). 1.Ind. Microbiol. 7, 27-34. Cheryan, M. (1980). CRC Crit. Rev. Food Sci. Nutr. 13, 297-335. Cheryan, M. (1986). “Ultrafiltration Handbook.” Technomic, Lancaster, PA. Cheryan, M., and Mehia, M. A. (1986). Chemtech 16(11), 676-681. Cheryan, M., and Shah, M. M. (1996). Unpublished data. Christianson, D. D., Cavins, J. F., and Wall, J. S. (1965). J. Agric. Food Chem. 13, 277-280. Chukwu, U. N., and Cheryan, M. (1996). Concentration of vinegar by electrodialysis. J. Food Sci. 61. In press. Crueger, W., and Crueger, A. (1990). In “Biotechnology: A Textbook of Industrial Microbiology,” 2nd ed., pp. 143-147. Sinauer Associates, Inc., Sunderland, MA. Diekert, G., and Thauer, R. K. (1980). FEMS Microbiol. Lett. 7, 187-189. Difco Laboratories (1984). Difco Manual: Dehydrated Culture Media a n d Reagents for Microbiology.Difco Laboratories Inc., Detroit. Drake, H. L., Hu, S. I., and Wood, H. G. (1980).1.Bid. Chem. 255, 7174-7180. Ebner, H., and Follmann, H. (1983). In “Biotechnology” (H. J. Rehm and G. Reed, eds.), pp. 387-407. Verlag Chemie, Weinheim. Fontaine, F. E., Peterson, W. H., McCoy, E., Johnson, M. J., and Ritter, G. J. (1942). I. Bacteriology 43, 701-715. Fucbs, G. (198fi).FEMS Microbiol. Rev. 39, 181-213. Ghose, T. K., and Bhadra, A. (1985). In “Comprehensive Biotechnology” (M. Moo-Young, ed.), Vol. 3 , pp. 701-729. Pergamon, New York. Han, I. S., and Cheryan, M. (1995). J. Membrane Science 107, 107-113. Han, I. S., and Cheryan, M. (1996). Appl. Biochem. Biotechnol. 57/58, 19-28. Hu, S. I., Pezacka, E., and Wood, H. G. (1984). J. Biol. Chem. 259, 8892-8897. Koesnandar, Nishio, N., and Nagai, S. (1990). 1. Ferment. Bioeng. 71, 181-185. Leh, M. B., and Charles, M. (1989).J. Ind. Microbiol. 4,77-80. LeMonnier, E. (1965). In “Kirk-Othmer Encyclopedia of Chemical Technology,” Vol. 8, pp. 386-404. Wiley-Interscience, New York. Levendis, Y. A. (1991). In “Calcium Magnesium Acetate (CMA)” (D. Wise, Y. Levendis, and M. Metghalchi, eds.), pp. 211-223. Elsevier, Amsterdam. Ljungdahl, L. G. (1983). In “Organic Chemicals from Biomass” (D. L. Wise, ed.), pp. 219-248. Benjamin/Cummings Publishing Co, Inc., Menlo Park, CA. Ljungdahl, L. G. (1986). Ann. Rev. Microbrol.40, 415-450. Ljungdahl, L. G., and Andreesen, J. R. (1978). Meth. Enzymol. 53, 360-372. Ljungdahl, L. G., Carreira, L. H., Garrison, R. J., Rahek, N. E., Gunter, L. F., and Wiegel, J, (1985). Biotecbnol. Bioeng. Symp. 15, 207-223. T.jungdah1, L. G., Carreira, L. H., Garrison, R. J., Rahek, N. E., Gunter, L. F.,and Wiegel, J. (1986). CMA Manufacture (11):Improved Bacterial Strain for Acetate Production. U.S. Department of Transportation Report No. FHWA/RD-86/117. Lundie, L. L., and Drake, H. L. (1984). J. Bacteriol. 159, 700-703. Manivanan, S., and Wise, D. (1991). In “Calcium Magnesium Acetate (CMA)” (D. Wise, Y. Levendis, and M. Metghalchi, eds.), pp. 257-272. Elsevier, Amsterdam. Marynowksi, C. W., Jones, J. L., Tuse, D., and Boughton, R. L. (1985). Ind. Eng. Chem. Prod. Res. Dev. 24(3), 457-465. Muraoka, H., Watabe, Y., and Ogasawara, N. (1980). 1. Ferment. Techno!. 60, 171-180. Nickol, G. R. (1979). In “Microbial Technology” (H. J. Peppler and D. Perlnian, eds.), Vol. 2, pp, 155-172. Academic Press, New York, NY. Osuga, J., Mori, A,, and Kato, J. (1984). 1.Ferment. Techno]. 62, 139-149. Parekh, S. R., and Cheryan, M. (199Oa). Process Biochem. 25(4), 117-121.

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Parekh, S . R., and Cheryan, M. (199Ob). Biotechnol. Lett. 12, 861-864. Parekh, S . R., and Cheryan, M. (1991). Appl. Microbiol. Biotechnol. 36,384-387. Parekh, S.R., and Cheryan, M. (1994a). Biotechnol. Lett. 16,139-142. Parekh, S . R., and Cheryan, M. (1994b). Enzyme Microb. Technol. 16, 104-109. Ragsdale, S. W., Clark, J. E., Ljungdahl, L. G., Lundie, L. L., and Drake, H. L. (1983). J. Biol. Chem. 258, 2364-2369. Raman, L. P., Cheryan, M., and Rajagopalan, N.(1994). Chem. Eng. Progr. 90(3), 68-74. Reed, W. M., and Bogdan, M. E. (1985). Biotechnol. Bioeng. Symp. Ser. 15, 641-647. Reed, W. M., Keller, F. A., Kite, F. E., Bogdam, M. E., Ganoung, J. S . (1987). Enzyme Microb. Technol. 9, 117-120. Schwartz, R. D., and Keller, F. A. (1982a). Appl. Environ. Microbiol. 43,1385-1392. Schwartz, R. D., and Keller, F. A. (1982b). Appl. Environ. Microbial. 43,117-123. Shah, M. M., and Cheryan, M. (1995a). J. Ind. Microbiol. 15, 424-428. Shah, M. M., and Cheryan, M. (1995h). AppI. Biochem. Biotechnol. 51/52, 413-422. Shah, M. M., Akanbi, F., and Cheryan, M. (1996). Appl. Biochem. Biotechnol. In press. Sharma, P. K. (1991). In “Calcium Magnesium Acetate (CMA)” (D. Wise, Y. Levendis, and M. Metghalchi, eds.), pp. 273-298. Elsevier, Amsterdam. Shoaf, W. T., Neece, S. H., and Ljungdahl, L. G. (1974). Biochim. Biophys. Acta. 334, 448-458. Shreve, R. N., and Brink, J. A,, Jr. (1977). “Chemical Process Industries.” McGraw-Hill, New York. Sugaya, K., Tuse, D., and Jones, J. L. (1986). Biotechnol. Bioeng. 28, 678-683. Wang, D. 1. C., Fleishchaker, R. J., and Wang, G. Y. (1978). AIChESymp. Ser, 182,105-110. Wang, G., and Wang, D. I. C. (1983). Appl. Biochem. Biotechnol. 8, 491-503. Wang, G., and Wang, D. I. C. (1984). Appl. Environ. Microbiol. 47,294-298. Wiegel, J., Carreira, L. H., Garrison, R. J., Robek, N. E., and Ljungdahl, L. G . (1991). In “Calcium Magnesium Acetate (CMA)” (D. Wise, Y. Levendis, and M. Metghalchi, eds.), pp. 359-416. Elsevier, Amsterdam. Wise, D. L. (1992). In ”Biochemical Engineering for 2001’’ (S. Furusaki, I. Endo, and R. Matsuno, eds.), pp. 723-726. Springer-Verlag, Tokyo. Wise, D. L., Levendis, Y. A., and Metghalchi, M., eds. (1991). “Calcium-Magnesium Acetate (CMA).” Elsevier Science Publishers, New York. Witjitra, K. (1994). M.S. Thesis, University of Illinois, Urbana. Witjitra, K., Shah, M. M., and Cheryan, M. (1996). Enzyme Microb. Technol. 19(7), 322-327. Wood, H. G. (1952a). J. Bid. Chem. 199, 579-583. Wood, H. G. (1952b). J. Biol. Chem. 199, 905-931. Yamamoto, I., Saiki, T., Liu, S. M., and Ljungdahl, L. G. (1983). J. Bid. Chem. 258, 1826-1832.

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Contact Lenses, Disinfectants, and Acantharnoeba Keratitis DONALDG. AHEARNAND MANALM. GABRIEL Department of Biology Georgia State University Atlanta, Georgia 30302-4010

I. Introduction and Taxonomy 11. Biology 111. Infections

IV. Ecology V. Disinfection VI. Adherence to Lenses References

I. Introduction and Taxonomy

Small filose, free-living amoebae with a cyst stage with ostioles are classified in the genus Acanthamoeba, family Acanthamoebidae, order Amoebida, phylum Rhizopoda. Acanthamoeba are characterized by typically uninucleate trophozoites with fine protoplasmic projections (acanthopodia) arising anteriorly or laterally from the clear hyaline ectoplasm and by a prominent cyst stage (see Page, 1967; Sawyer and Griffin, 1975). The trophozoite divides by binary fission with the early disappearance of the nucleolus and nuclear membrane. In axenic broth cultures the trophozoites are somewhat globose to slightly irregular and of variable size (12 to 25 pm in diameter). In the presence of bacteria and particularly when traversing a surface, the trophozoites are more irregular in form and usually larger (15 to over 60 pm for some species) in one dimension. A large central or posterior vacuole is present. The trophozoite may become coated with bacteria or fungal cells that ultimately are ensnared in capsular material; a pincer-like pseudopod may be present in some species. In broth cultures with low densities of bacteria, trophozoites may agglutinate in clumps of 10 to 30 globose cells. A cellulosic cyst stage (endocyst wall reacts most strongly for cellulose) that progresses from a round precyst to a double layered wall form is prominent. The cysts contain a variable number of ostioles with closed apertures. Under scanning electron microscopy, the round to 35 ADVANCES IN APPLIED MICROBIOLOGY, VOLUME 4 3 Copyright 0 1997 by Academic Press, Inc All rights of reproduction in any form reserved. 0065-2164197 1625.00

36

D. G. AHEARN AND M. M. GABRIEL

FK:.1. Cysts and trophozoites of Acanfhnmoeba castellonii on surfaces in contact lens cases contaminated with bacteria and fungi. Note that bacteria adhered to surface of cyst (top right) and variable (round to stellate-like) cysts (top left); characteristic acanthapodia of trophozoite (bottom left]. Trophozoites “capped” with ensriared cells of Pseudoinonas aeruginnsa in PBS (bottom right).

stellate cysts appear polyhedral with ridges defining the facets. Cysts may have bacteria or fungi adhering to their surfaces (Fig. 1). Pussard and Pons (1977) divided the genus into three subgroups on the basis of cyst morphology: Group I-deeply scalloped stellate endocysts with a rounded, slightly rippled or smooth wall, ostioles at tips of the rays, ectocyst and endocyst walls usually separated; Group 11-cysts generally smaller than those in Group I, globular, ovoid, polygonal or triangular endocysts with ectocyst wrinkled or rippled, ostioles usually at the angles of the rays; Group 111-cysts round to slightly angular to irregular with ectocyst wall thin and delicate, single or obscure ostioles. A listing of species as to their probable group on the basis of cyst morphology is presented in Table I. These groupings are helpful in distinguishing some species, but variabilities in cyst morphology between strains and with culture conditions make some species assignments difficult or arbitrary. Species in Group I are generally larger than those in the other two groups. Supplemental information such as isoen-

CONTACT LENSES AND ACANTHAMOEBA KERATITIS

37

TABLE I PROVISIONAL GROWINCS OF ACANTHAMOEBA SPECIES ON THE BASISOF CYST MORPHOLOGY" Group I Mostly smooth ectocyst, stellate scalloped endocyst

A. astronyxis (Ray and Hayes, 1954) Page, 1967

Group I1 Mostly wrinkled ectocyst, variable endocyst

Group I11 Ectocyst thinly rippled to smooth, round to slightly angular endocyst

A. castellanii (Douglas, 1930) Page, 1967' (9-14, 11)

A. culbertsoni Singh and Das, 1970' (13-28, 18)

A. hatchetti Sawyer, Visvesvara, and Harke,

A. jacobsi Sawyer, Thomas, Nerad, and Visvesvara, 1992 cysts spherical (12.5-1 7.5, 15)

(16.0-28.0, 19)h

A. comandoni Pussard, 1964 (21-30, 22)

1977' (11.5-21, 16)

A. echinulata Pussard and Pons, 1977 (10-32, 25)

A. polyphaga (Puschkarew, 1917) Page, 19fi7" (9-16, 13)

A. palestinensis (Reich, 1933) Page, Page, 1967 (13-22, 18)

A. griffini Sawyer, 1971r (14-25, 20)

A. rhysodes (Singh, 1652) 1979' (10-26, 18)

A. lenticulata Molet and Ermolieff-Braun, 1976 (n/a)

A. pearcei Nerad, Sawyer, Lewis, and McLaughlin,

A. stevensoni Sawyer, Nerad, Lewis, and McLaughlin, 1993

A. royreba Willaert, Stevens, and Tyndall, 1978 (n/a)

1995 (17.5-25, 21)

Singh and Hanumaiah,

(10-23, 17)

A. tubiashi Lewis and Sawyer, 1979 (18-30, 23) "Based on Pussard and Pons (1977) and Sawyer rt al. (1992: 1993). bRanges and mean diameters of cysts in @m in parenthesis ( 1 are approximated from literature reports and observations of representative cultures: nla = data not available. "Species associated with human eye infections.

zyme profiles (propionyl esterase, leucine aminopeptidase, and acid phosphatase) and temperature tolerance for growth are necessary for distinguishing species (Stratford and Griffiths, 1978; Sawyer et al., 1993; Nerad et al., 1995). Molecular studies have further demonstrated the classification complexities within Acantharnoeba. Restriction-fragment-length polymorphism (RFLP) analyses of mitochondria1 DNA (mtDNA) of isolates of A. castellanii and A. polyphaga indicate that the two species may represent a single species complex (Byers et al., 1983; Yagita and Endo, 1990). Digestion with BgIII, EcoRI, and Hind111 of whole-cell DNA of 33 clinical isolates ostensibly representing these two species produced RFLPs that differentiated the strains into seven multiple-strain and three sin-

38

D. G. AHEARN AND M.M. GABRIEL

gle-strain groups (Kilvington et al., 1991). The study included four strains of A. polyphaga (identified on the basis of cyst morphology) that were placed among three RFLP groups. The authors indicate that the number and positions of the EcoRI RFLPs were identical to those obtained by Byers et al. (1983) in their study of mtDNA from A. castellanii. This fact and the similarity in sizes (about 40 kb) of the RFLPs indicated that their RFLPs originated from mtDNA. Ledee et al. (1996) studied PCR-amplified nuclear 18s rDNA genes from three isolates of A. griffini. These genes, approximately 2800 bp in length in A. griffini and A. Zenticulatu, contain a Group I Intron (Gast et al., 1994). Comparison of the sequences of the RFLPs from isolates of A. griffini, one from an eye, one from tap water, and one from a contact lens associated with the same individual, showed that the isolates were identical and differed by less than 1%in bp sequences from the type culture (Ledee et al., 1996). The possible presence of endosymbionts in Acanthamoeba spp. presents an added difficulty in comparisons of mtDNA and in PCR-based studies. Byers (1986) in his extensive review expresses the need for sequence data on more conservative regions of DNA because of high interstrain variability in the mtDNA. Additional data on strains of varied environmental and clinical origin showed no overall correlation between mtDNA fingerprint groups and environmental source and further demonstrated the diversity of mtDNA fingerprint groups (Gautom et a]., 1994). 11. Biology

Acanthamoeba species are voracious predators of various Gram-negative bacteria, cyanobacteria, and fungi (Nero et al., 1964; Wright et al., 1981; Schuster et a]., 1993). Species are cannibalistic or pathogenic and may be grown in axenic enrichment culture and on defined media. An enrichment broth of peptone (2.0%), glucose (l.80/0),and yeast extract (0.2%)in a basic salts mixture is frequently employed for axenic culture of Acanthamoeba, but peptose-glucose broths fortified with salts, or skim milk broth, tissue culture fluids, etc., will support growth of many strains (Neff, 1957). Acanthamoeba spp. may be cultivated also in defined media composed of basic salts, various mixtures of amino acids, biotin, and thiamine and fortified with acetate or glucose (Adam, 1959, 1964). The generation times for Acanthamoeba species vary with the strain and culture conditions but in axenic culture frequently range from 13 to 18 h at 25°C.

CONTACT LENSES AND ACANTHAMOEBA KERATITIS

39

Neff (1957) demonstrated that live or dead Aerobacter aerogenes and Escherichia coli and dead cells of Saccharomyces cerevisiae served as excellent food sources for the growth of an isolate of Acanthamoeba. Live yeasts, including Candida famata, Cryptococcus neoformans, and Rhodotorula rubra, are known to support the growth of various strains of Acanthamoeba (Castellani, 1930; Nero et al., 1964; Bunting et al., 1979). Schuster et al. (1993) found that A. castellanii migrated in a wateragar medium to both Gram-negative and Gram-positive bacteria but preferably ingested Enterobacter cloacae, Klebsiella pneumoniae, Shigella boydii, and Bacillus cereus. A weak feeding response was observed to Serratia marcescens and the response to Staphylococcus aureus was moderate; Pseudomonas aeruginosa was not ingested. Stenotrophom o n m (Xanthonomonas) maltophilia cocultivated in saline with A. castellanii or A. polyhaga for 24 h supported more luxuriant growth of the amoebae than Escherichia coli (Bottone et a]., 1994). Pseudomonas aeruginosa, a species that may be lethal for Acanthamoeba species (Qureshi et al., 1993), E. coli, Staphylococcus aureus, and S. epidermidis were only sparsely internalized. Bottone et al. (1994) also noted that a “capping” phenomenon occurred with Stenotrophomonas maltophilia and F! aeruginosa where the bacterial cells seemed to be ensnared in mass in a sticky-surface substance secreted by the amoebae. Capping was not observed with E. coli nor the nonmotile S. epidermidis. Most Acanthamoeba species will grow on live or dead E. coli (the use of E. coli for the isolation and enumeration of Acanthamoeba is a standard practice), but growth on staphylococci and particularly pigmented Gram-negative bacilli is frequently reported as negative. The inhibition of Acanthamoeba species by bacteria may be associated with toxic pigments or other metabolic by-products (Singh, 1945). Nevertheless, inhibitory bacteria such as r! aeruginosa and S. marcescens have been found to support the growth of A. castellanii in saline (Wang and Ahearn, in press). Various investigators have indicated that Acanthamoeba migrate to bacteria that are not internalized. The food preferences demonstrated by strains of Acanthamoeba in laboratory studies are most likely influenced by nutrient carryover with the bacterial inocula, the metabolic state of the bacteria, and the density ratios and strains of amoebae to bacteria. Certain bacteria and yeasts ingested by strains of Acanthamoeba may be maintained as endosymbionts for varied time periods (Drozanski and Chmielwski, 1979; Fritsche et al., 1993). Bacillus spp. and Saccharomyces spp. may be maintained in amoebae cysts, whereas Legionella spp. may proliferate in the trophozoites (Moffat and Tompkins, 1992).

40

D. G. AHEARN AND M. M. GABRIEL

The cysts of Acanthamoeba species are produced under varied conditions that include low temperature and low nutrients, age in culture, and the presence of inhibitors. A. castellanii and A. palestinensis may produce over 70% cysts within 48 h when shifted from enrichment media to 0.5 to 0.8M MgC12, or to NaCl or KC1 in the case of the latter species (Griffiths and Hughes, 1968; Lasman and Shafran, 1978). An encystment factor produced by transforming cells induces or enhances cyst formation by both these species (Akins et al., 1985; Lasman, 1987).

I l l . Infections Acanthamoeba species are the cause of rare human infections. Visvesvara (1995) includes A. culbertsoni, A. castellanii, A. polyphaga, A. astronyxis, A. healy, and A. divionensis as agents of chronic granulomatous amoebic encephalitis. This disease of the central nervous system occurs primarily among the immunosuppressed or chronically ill, particularly AIDS patients. Only about 100 cases have been reported worldwide (Visvesvara, 1995). The first reported case of Acanthamoeba keratitis developed in 1973 in a Texan farmer who had rinsed an eye with water from a horse trough following trauma to the eye from a shaft of straw (Jones et a)., 1975). Both trophozoites and cysts of A. polyphaga were isolated repeatedly from corneal scrapings. Shortly afterwards, two eye infections caused by Acantharnoeba were recognized in the United Kingdom. One occurred in a teacher of 32 years who had mild unilateral keratoconjunctivitis and uveitis that did not respond to treatment. Within 6 months, the eye deteriorated, with corneal ulceration, pain, and loss of vision. A. polyphaga was isolated frequently from the affected eye. A corneal graft was eventually rejected. The second case involved a farmer of 59 years with very similar eye conditions (unilateral central or paracentral corneal infiltrate) who eventually lost his eye. Acanthamoeba was isolated from his eye tissues (Nagington et al., 1974). During the next few years, a series of reports associated amoebic keratitis with contact lens wear (CDC, 1986; Kyle and Noblet, 1987; Auran et al., 1987; Stehr-Green et al., 1989). In contrast to the mostly fatal granulomatous encephalitis, Acanthamoeba keratitis (also a rare disease) occurs most commonly among healthy normal individuals and seems associated with the minor trauma to the eye that accompanies contact lens wear. The species diagnosed from eye infections, in order of their decreasing frequency, are A. castellanii, A . polyphaga, A. rhysodes, A. culbertsoni, A. hatchetti, and A. griffini.

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The symptoms of Acanthamoeba keratitis include a foreign body sensation followed by severe pain, tearing, photophobia, and blepharospasm. The syndrome emerges over a period ranging from only a few days to months. Initial corneal changes may range from opaque streaks to epithelial opacities, or corneal epithelial stippling and microcystic edema. The syndrome worsens to dense central infiltrates with thinning, circumferential peripheral guttering, and peripheral vascularization. In most cases, the recurrent cycle of healing and breakdown in epithelium overlying the stromal infiltrate is apparent (Cohen et al., 1987; Lindquist eta]., 1988).The disease responds poorly to chemotherapy and has frequently resulted in loss of vision. Initially, amoebic keratitis associated with contact lens wear was frequently misdiagnosed; the disorder and its lesions were often confused with fungal, bacterial, or herpetic keratitis. It is not clear whether cysts or trophozoites or both are the infectious entities in Acanthamoeba keratitis, nor is there information available on an infectious dose. It is clear, however, that Acanthamoeba spp. are common in the environment and that Acanthamoeba keratitis is of rare occurrence, much less frequent than contact-lens-related Gram-negative bacterial keratitis. In turn, contamination of contact lens cases with Gram-negative bacteria is not uncommon, and Acanthamoeba may occur in storage cases of wearers with no symptoms of infection. Failure to comply with proper hygienic procedures for cleaning and disinfecting lenses appears to be the major factor in most contact-lens-related eye infections, although some disinfectant systems are less forgiving of deviations from recommended regimen. Bacterial and fungal colonization of the lens case appears to be the primary step in most Acanthamoeba keratitis related to contact lens wear. Proliferation of the amoebae in the case is followed by transport of the amoeba to the eye via the lens. Trophozoites of A. castellanii and A. polyphaga will bind to the cornea and seem to share a mannose-type receptor with l? aeruginosa (Morton et a]., 1991). Predisposition of the cornea of the eye from the minor trauma of contact lens wear or by a coinfecting agent follows binding of the trophozoite. Cysts do not adhere to submerged surfaces, but they do become entrapped in biofilms and the cyst may potentially be embedded in the lens. The cyst itself could traumatize the cornea. Recently, Mathers et al. (1996) suggested that Acanthamoeba keratitis is more prevalent in the United States than suspected. In a tandem-scanning-confocal-microscopy screening of 2 1 7 keratitis patients where 5 1 patients were suspected to be infected with Acanthamoeba, 36 of these demonstrated highly refractile ovoid bodies typical of Acanthamoeba.

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None of the patients were culture-positive for Acanthamoeba; although epithelial smears demonstrated trophozoite- and cyst-like cells. After antiamoebic therapy was commenced (polyhexamethylene or, preferably, chlorhexidine biguanide, each at 0.02% and in combination with propamidine), 43 patients had fewer symptoms, clinical signs, and refractile bodies indicated by tandem-scanning-confocal microscopy. Only 1 7 of the 43 patients were wearing contact lenses when symptoms developed. The investigators suggest that their apparent mini-epidemic of amoebic keratitis could indicate a much more common infection, possibly caused by a new species of amoebae. Detection of Acanthamoeba keratitis by confocal microscopy, a rapid, noninvasive in vivo procedure, is of value for early diagnosis of the disease. The diagnosis is accurate even when culture results are negative (Pfister et al., 1996). Various chemotherapeutic agents have been used in vitro: propamidine, dibromopropamidine (Wright et al., 1985), hydroxystilbamidine, paromomycin, 5-fluorocytosine (Jones et a]., 1975), and clotrimazole (Borochovitz et al., 1981; Stevens and Willaert, 1980) for the treatment of Acanthamoeba keratitis. Kilvington and White (1994) suggested that propamidine is the drug of choice in the treatment of Acanthamoeba. At the present time, propamidine combined with polyhexamethylene biguanide (PHMB) or propamidine combined with chlorhexidine digluconate have been employed successfully (Hay et al., 1994; Seal et al., 1995b; Ledee et al., 1996). The biguanides chlorhexidine and PHMB are used topically at concentrations of 0.02%. They have broad-spectrum antimicrobial activity against bacteria and fungi as well as amoebae and are included as disinfectants or preservatives in various products. Their broad antimicrobial activity may be important in treatment of Acanthamoeba keratitis because there is a possibility that at least with some strains of amoeba virulence is related to the presence of coinfecting bacteria or endosymbionts (see Badenoch, 1991). IV. Ecology

Species of Acanthamoeba dwell in damp soils and upper zones of mud in rivers and lakes and in anthropogenic water reservoirs (Daggett et al., 1982; Sawyer et al., 1987; Bhattacharya et al., 1987; Martinez, 1985; Visvesvara, 1986). Cysts and trophozoites are carried by humans and distributed by wind, water, and animals. Extensive studies by Sawyer and his colleagues have demonstrated that distributions of Acanthamoeba in soils, fresh waters, and marine coastal habitats may coincide with sewage contamination (see Sawyer, 1989; Sawyer et al., 1993; Sawyer et al., 1996). Even species initially described from true

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marine habitats probably are of terrestrial or estuarine origin (Bhattacharya et al., 1987; Nerad et al., 1995; Sawyer et al., 1996). Species of Acanthamoeba readily colonize surface biofilms in swimming pools, hot tubs, heating-, ventilation-, and air-conditioning systems, and domestic water taps and eye-wash stations (Martinez, 1985; Bier and Sawyer, 1990; Paszko-Kolva et al., 1991; Mergeryan, 1991). The association of eye infection with contact lens wear in the United States coincided with the use of home-prepared saline solutions (CDC, 1986, 1987). The amoebae grow on microbial contaminants in nonpreserved saline or inadequately preserved storage solutions and are transferred on the contact lens to the eye (Ubelaker, 1991). As many as 50% of normal contact lens wearers have contaminating microorganisms in their contact lens cases at some time during their use of lenses (Wilson et al., 1990). Donzis et al. (1989) reported that Acanthamoeba spp. contaminated lens cases and solutions only when bacteria or fungi were present. No Acanthamoeba spp. were found in the parent bottles of disinfectant solutions. Of ten individuals with Acanthamoeba spp. in their lens care systems, Gram-negative bacteria, mainly Pseudomonas spp., were also present; in six instances in conjunction with Bacillus spp. More than 70% of the cornea tissues and lens cases of patients with Acanthamoeba keratitis were found to be cocontaminated with bacteria and amoebae (Bacon et d.,1993). Giovannini et al. (1994) suggested that the early onset of bacterial conjunctivitis may enrich the natural conjunctival microbiota, as well as the contact lens case, with a n abundant bacterial supply, thus favoring the growth and the development of virulent Acanthamoeba spp. in the lens case and eye. In vivo experiments with rats have shown that the virulence of Acanthamoebae spp. in keratitis is enhanced by coinoculation with bacteria (Lawin-Brussel et al., 1993). Bacterial numbers in a system might decrease from phagocytosis by amoebae or bacterial numbers may increase because of their higher growth rate (Bamforth, 1985; Griffiths, 1990). Larkin et al. (1990) proposed that bacteria in the external eye or in the lens system could stimulate cysts of Acanthamoeba spp. to excyst to the infectious vegetative stage (trophozoites). Trophozoites have not been observed to grow in unadulterated contact lens solutions, and, though encystment may result, the cysts show poor viability. Cysts, however, particularly those from cultures grown on Stenotrophomonas and Escherichia, will persist for at least up to several weeks in otherwise microbe-free disinfectant solutions. In the United States, the withdrawal of salt tablets for home-prepared saline from the market in the mid-1980s resulted in a decreased incidence of Acanthamoeba keratitis. In England and Asia during the

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199Os, reports of Acanthamoeba keratitis seem to be increasing (see Kilvington and White, 1994). Rinsing lenses in tap water (particularly if the water supply is from a roof-storage facility deficient in disinfectant) may also be implicated in Acanthamoeba keratitis (Talamo and Larkin, 1993; Ledee et al., 1996). In the report by Ledee et al. (1996), the same strain of A. griffini was isolated from the cornea, the contact lens storage case, and the domestic water supply. Contact-lens-associated Acanthamoeba keratitis in Hong Kong and England in the mid-1990s may be related to disinfectant deficiencies in reservoir-type water systems. In general, primary contamination of the lens care system, particularly with Gram-negative bacteria, including Pseudomonas spp., seems to be a prerequisite for the establishment of infectious populations of Acanthamoeba. Studies that indicate that Acanthamoeba spp. and P aeruginosa are mutually exclusive in lens cases may not recognize that survival or presence of both bacterium and amoeba in the lens case varies with time and population dynamics.

V. Disinfection

Various reports have indicated that cysts of Acanthamoeba species may resist desiccation from months to years, tolerate 2.0% concentrated HC1 (0.25 M) and resist exposure to chlorine, chlorides, ozone, and peroxides (Sawyer, 1970; Cursons et al., 1980; Bryant et a]., 1982). Penley et al. (1989) found that trophozoites and cysts of two corneal isolates of Acanthamoeba survived in most contact lens solutions beyond the manufacturer’s recommended disinfectant times. Only an isopropyl alcohol cleaning system (now approved for disinfection), and an “02 conditioner” killed both (2 x lo3) cysts and trophozoites of A. polyphaga and A. castellanii within 30 min of exposure. Solutions with chlorhexidine (0.005 and 0.006%) and chlorhexidine and alkyl triethanol ammonium chloride (0.013%) plus thimerosal (0.002%) appeared to be the next most effective against A. polyphaga; no viable amoebae were recovered at 18 h. An isolate of A. castellanii was more resistant, surviving in all tested solutions (except the O2 conditioner and isopropyl alcohol). A similar pattern of relative efficacies for contact lens solutions was reported by Connor et al. (1989) for cysts of A. culbertsoni; the isopropyl alcohol solution and solutions containing thimerosal appeared more effective than hydrogen peroxide or solutions with PHMB.

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Ludwig et al. (1986) indicated that heat disinfection (80°C for 10 min) in a preserved saline solution (NaCl 0.7% with sodium borate, boric acid and thimerosal O.OOl%, and EDTA 0.1%) killed pellet inocula of 5 x lo2 cysts and 5 x lo3 trophozoites of A . castellanii and A. polyphaga suspended in 2 ml of disinfectant solution. A chlorhexidine (0.005°/~)/thimerosal(O.OO1°/o) solution after 4 h at room temperature killed the trophozoites and cysts of A. castellanii but not of A. polyphaga. Both species of Acanthamoeba survived 4-hour exposure to alkyl triethanol ammonium chloride (0.013%)/thimerosal 0.002% and chlorhexidine (0.005%), and 10 and 30 min exposure to hydrogen peroxide (3%). Davies et al. (1990) found that strains of A. polyphaga and A. castellanii (lo3 to lo4 cysts per milliliter) were killed by the triethanol ammonium chloride-thimerosal solution within 4-6 h. Hydrogen peroxide (3%) was effective at 4 h, whereas solutions with 0.001% polyquaternium and 0.00005% polyaminopropyl biguanide (equals polyhexamethylene biguanide or PHMB) yielded viable amoebae after 24-h exposure. Zanetti et al. (1995) studied disinfectant solutions diluted in a peptone-yeast extract-glucose medium. Hydrogen peroxide (1.5%) killed 100% of the trophozoites (1.5 x 104/ml) of a clinical strain of A. castellanii within 30 min. This solution was more effective than a diluted chlorhexidine-thimerosal solution (100% kill at 3 h). A mixed cyst-trophozoite suspension (over 90% trophozoites) yielded viable amoeba after 6 h but not after 9 h of exposure to these same preparations. Other diluted commercial disinfection solutions containing benzalkonium chloride, PHMB, or polyquaternium yielded viable amoeba after 9 h exposure to the disinfectants. Connor et al. (1989) compared the effectiveness of chlorhexidine digluconate at concentrations from 0.0001% to O . O l % , a commercial solution with PHMB (0.0005%), hydrogen peroxide YO), thimerosal (O.OOZ%), and heat (80°C for 10 min) against cysts (1.5 x lo4 per ml) of A. culbertsoni. Chlorhexidine at 0.005% appeared equivalent to the PHMB solution; 42% survival of the cyst inocula was reported after 24-h exposure. Hydrogen peroxide for 60 min and thimerosal after 24 h appeared least effective, whereas heat was most effective, but still 6% of the cyst inocula was reported to survive 80°C for 10 min. Viability of cysts after exposure to the disinfectants was based on hemocytometer counts of calcofluor white-stained cysts that showed a decrease in cyst number. The surviving cysts were incubated in thioglycolate broth, and cells were enumerated 1 week later to determine if any growth had occurred. Growth occurred in all systems. When the concentration of chlorhexidine was increased (0.09%),the cysts were destroyed.

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D. G. AHEARN AND M.M.GABRIEL

The biguanides chlorhexidine and PHMB have demonstrated clinical efficacy at concentrations of 0.02% (Larkin et al., 1992). Burger et al. (1994) found that PHMB at 0.009% (90 ppm) in a borate buffer killed over 98% of cysts (1O5-1O6/ml) of A. castellanii and A. polyphaga within 30 s. Hay et al. (1994), in an extensive comparison and pairing of 1 2 antimicrobials against 20 isolates of Acanthamoeba, found that susceptibilities varied with the strain, but chlorhexidine and PHMB were the most active single compounds after 48-h exposure. Concentrations below 0.005% were active against 2 x lo4 cysts and trophozoites of most strains. The two biguanides each showed slight in vitro synergy when in combination with pentamidine. In a peptone-yeast extractglucose medium, concentrations of 0.005 to 0.01% of either chlorhexidine or PHMB were biocidal at 24 h for 105 trophozoites or cysts per ml of four Acanthamoeba spp. (Tirado-Angel et al., 1996). The biocidal effect was dependent upon exposure time, concentration of inocula and inhibitor, and strain. Some synergism between the two biguanides was noted. Nearly all the above investigations noted that trophozoites were more susceptible than cysts to inhibitors. Most indicated that biguanides, particularly chlorhexidine, were the more active antiamoebal compounds. The antimicrobial activities usually showed a concentration (both of inocula and inhibitor) and time dependence. The various discrepancies in the data on the relative efficacies of commercial systems have been attributed to strain differences and procedural differences, particularly the quantitation of surviving cells. Trophozoites and some precysts lyse in the presence of preservatives, particularly biguanides. Dependent upon the strain of amoeba and other test conditions, some inhibitors (particularly at sublethal concentrations) may induce encystment. Some of the differences in the results also may be attributed to failure to quantitate the viability of “surviving” cysts and failure to use neutralizers in the recovery process (or the use of an inappropriate neutralizer). Also, cysts of some strains may have relatively poor viability or demonstrate highly variable degrees of recalcitrance (see Schuster and Jacob, 1992). We have made similar observations, particularly with cysts that have been produced in axenically maintained cultures. Such cultures may need to be grown on selected bacteria to regain the more typical resistant cysts. In particular, bacterial contamination may be of some importance in determining the resistance of cysts. Outgrowth of bacteria or yeasts (endosymbionts or exogenous contaminants) in the test system can stimulate trophozoites to feed and cysts to excyst, resulting in increased susceptibility of the amoeba to the disinfectant.

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VI. Adherence to Lenses

Various reports have indicated that trophozoites adhere preferentially over cysts to contact lenses and that various species and strains differentially adhere to the four FDA hydrogel lens groups (Table 11). In a comparison of two nonionic lens types, a polymacon 38.6%-water-content lens and a lidofilcon 70%-water-content lens, Kilvington and Larkin (1990) found that both cysts and trophozoites showed greater adherence to the higher-water-content lenses. In contrast, John et al. (1991) reported lower adherence of A. castellanii to ionic, 58% watercontent etafilcon A lenses than to nonionic, 38% water-content polymacon lenses and nonionic, 70Y0 water-content lidofilcon lenses. Kilvington (1993) found different degrees of adherence to the four FDA contact lens groups for two strains of Acanthamoeba. Trophozoites of both strains showed greater adherence than cysts. Inocula of 1 x 106 washed cells suspended in normal saline were placed into wells of a microtiter plate, each well containing a single lens. After 4 h of incubation at room temperature, each lens was removed and gently submerged in two separate 10-ml volumes of saline. The lens was then placed in 1 ml of ice-cold saline for 15 min. The tube was vortexed for 10 s and the cells in suspension were counted. Trophozoites of one strain had greater adherence to a Group I lens (38% water content), with no significant differences in adherence to the other lenses (water contents ranging from 45 to 70%). The other strain showed equivalent adherence to Groups I, 111, and IV lenses and reduced adherence to the Group I1 lens (70% water content). Cleansing and rinsing procedures of all four lens types with commercial systems as per the manufacturer’s instructions removed all amoeba from the lenses. Kelley and Xu (1995) examined the effect of increasing concentrations of trophozoites of A. polyphaga on adherence to lens fragments for four different lenses: polymacon, nonionic, 38% water content; bufilcon A, ionic, 45% water content; etafilcon A, ionic, 58% water content; and lidofilcon A, nonionic, 70% water content. After 2 h of incubation of the lens materials in amoeba suspensions with agitation at room temperature, followed by three washes in phosphate-buffered saline (PBS) (volumes not given), the higher concentrations of trophozoites ( 105/lens fragment) adhered more to the nonionic 70%-watercontent lens than to the nonionic 45%-water-content lens. Adherence to the nonionic lenses was significantly greater than to the ionic lenses. With inocula concentrations per lens fragment of about 250-500 trophozoites, no significant difference in adherence to lidofilcon and bufilcon materials was detected. The etafilcon A lens at all inocula concentra-

TABLE I1 FDA GROUPINGS OF HYDROGEL CONTACTLENSES

Group 1 low water (<50% H,O) nonionic polymers

Group 2 high water (>50% H,O) nonionic polymers

Group 3 low water (<50% H,O) ionic polymers

Group 4 high water (>50% H,O) ionic polymers

Tefilcon (38%) Tetrafilcon A (43%) Crofilcon (38%) Nefilcon A&B (45%) Isofikon (36%) Mafilcon (33%) Polymacon (38%)

Lidofilcon B (79%) Surfilcon (74%) Lidofilcon A (70%) Netrafilcon A (65%)

Bufilcon A (45%) Deltafilcon A (43%) Droxifilcon A (47%) Phemfilcon A (38%) Ocufilcon (44%)

Bufilcon A (55%) Perfikon (71%) Etafilcon A (58%) Ocufilcon B (53%) Ocufilcon C (55%) Phemfilcon A (55%) Methafilcon (55%) Vifilcon A (55%)

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tions appeared to have the lowest levels of adherence. Later, Kelly et al. (1995) submerged segments of polymacon, nonionic, 38%-water-content, etafilcon A, ionic, 58%-water-content, siloxane acrylate, and polymethylmethacrylate (PMMA) lenses in cell suspensions of A. CQStellanii (105/ml, trophoz0ite:cyst ratio, 90:lO) in a peptone-yeast extract-glucose medium. After incubation with agitation for varying periods u p to 2 h, the lens segments were submerged for 3 s in PBS and than examined under a microscope. The number of trophozoites adherent to the PMMA and RGP segments increased with time of incubation. This phenomenon was not observed for cysts. More trophozoites adhered to these lenses (mean adherence: 50 to 75 per mmz) than to the hydrogel lenses, which retained no more than 25 cells per mm2 after 2 h. No statistically significant differences in adherence existed between the two types of rigid lenses or between the two hydrogel lenses. In this latter study, where lens segments were incubated for 2 h with cells in a peptone-yeast extract-glucose medium, the mean number of adherent trophozoites, about 10 to 15 mm2, was considerably higher than those adhering from PBS (<1.0/mm2)(Kelly and Xu, 1995). Seal et al. (1995a) reported a highly significant difference for the number of trophozoites and cysts of A. castellanii and A. polyphaga adhering to quadrant sections of the four FDA lens groups incubated in a glucose-salts medium. The sections were placed directly into bottles from their sterile packaging solution into 1.5 ml of amoeba1 suspension (1.0 x lo6 cysts or trophozoites/ml). The sealed bottles were agitated for 90 min at 25°C. Each lens section was removed and passed into new bottles containing 1.5 ml of sterile PBS and shaken for 5 min. The lens sections were removed, shaken gently, and examined microscopically. The mean number of adhering trophozoites was at least twice the number of cysts. Ionicity (as the principal factor) and water content had significant effects: greater numbers of amoeba adhered to ionic higher-water-content lenses than to nonionic low-water-content lenses. The mean number of amoeba of both species observed per section ranged from about 61 to 66 for high-water ionic lenses, from 37 to 45 for low-water ionic lenses, versus 22 to 26 for high-water nonionic and 10 for low-water ionic lenses. These investigators did not find any adherence to silicone acrylate, fluorosilicone, and PMMA lenses. Studies in our laboratories (Gorlin et al., 1996a) are in general agreement with those of Seal et al. (1995a). Gorlin et al. (1996a) exposed whole lenses to about 2 x lo6 amoeba in 1.0 ml of saline for 2 h at 25OC. The number of amoeba retained on entire hydrogel lenses were determined with radiolabeled cells and a cell detachment procedure after

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D. G. AHEARN AND M. M. GABRIEL

different rinsing regimens. Trophozoites of A. castellanii, A. polyphaga, A. culbertsoni, and A. hatchetti retained on nonionic and ionic lenses following a single rinsing procedure were at higher densities on the higher-water-content lenses along with a tendency for greater retention on the ionic (negatively charged) high-water-content lenses. After several rinses (five repeated immersions each in two successive volumes of 100 ml chilled saline or in successive 50 ml volumes on a shaker), nearly all amoeba were removed from the lenses of all water contents. High water content and surface tension appeared more significant than ionicity in binding of the amoeba to unused contact lenses. Cysts showed little or no adherence to lenses. Pseudomonas aeruginosa has been shown to adhere irreversibly to hydrogel lenses, generally for nonionic lenses in an inverse relationship to water content (Miller and Ahearn, 1987). In this latter study, the least adherence of I? aeruginosa was to an etafilcon A lens (high-water ionic). Preston and King (1984) found that the polar flagella of I? fluorescens bound avidly to A. castellanii. Gorlin et al. (1996b) found that when I? aeruginosa was adhered to lenses prior to incubation with amoebae the binding to lenses of strains of A. castellanii, A. hatchetti, and A. polyphagu was significantly increased. In the presence of adhered I? aeruginosa, the association of amoeba with lens water content was the inverse of that with pure cultures of amoebae. Exposure of lenses to Staphylococcus epidermidis prior to exposure to amoebae did not enhance retention of amoebae on lenses. Differences between strains and inocula, in preparation and processing of lenses, in incubation times, in methods of quantitation, and particularly in the rinsing procedure (the volume and sequential volumes are significant), and whether amoeba were suspended in saline or more viscous growth media, explain some of the disparities in the results on the “adherence,” “association,” and “binding” of amoebae to contact lenses in laboratory studies. There is a general consensus that more amoebae are associated with higher-water-content lenses than to lower-water-content lenses. Amoebae on the lens surface may be maintained in the hydrogel-bound water, which may not freely exchange with water in short-term rinsing procedures. Cysts and precysts that are not producing protoplasmic extensions and thus have reduced surface area are readily flushed from unused contact lenses or from worn lenses. Whether ionicity of the lens or surface tension is most active in retention of an amoeba on a hydrogel surface probably depends upon the metabolic activity of the trophozoite and deposits on the hydrogel surface. The overall charge of the trophozoite is probably negative, but charges on pseudopodia probably vary and relate to passage over a

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surface. Rinsing conditions, including degree of agitation, may cause retraction of pseudopodia and rounding of the trophozoite with resultant alteration of surface tension. It is not clear that all studies reviewed above involved pure cultures. If Gram-negative flagellated bacteria were present in the test system (with their irreversible adherence to the lens surface), amoebae would be more firmly bound to the lens (Gorlin et al., 1996b). Acanthamoeba species have attained increased importance because of the recognition of their rare involvement in sight-threatening to grave infections of the central nervous system. Whether the increasing populations of compromised host will result in an increased incidence of these serious infections is unknown, but such an event should not be unexpected. In any regard, fastidious cleaning and disinfection of contact lenses, with particular care in lens storage, are necessary in the control of Acanthamoeba keratitis. Acanthamoeba spp. show broad interspecific and intraspecies diversity. Additional basic studies on the physiology, classical and molecular systematics, and virulence factors of these amoebae are necessary. More standardized laboratory procedures would provide obvious benefits, and these procedures would be best developed with an extensive range of characterized strains. ACKNOWLEDGMENTS

We thank Robert B. Simmons for his technical assistance in the preparation of scanning electron micrographs. Thomas K. Sawyer is thanked for his generous and beneficial comments during the preparation phase and in his review of the manuscript. REFERENCES Adam, K. M. G. (1959). The growth of Acanthamoeba sp. in a chemically defined medium. J. Gen. Microbiol. 21, 519-529. Adam, K. M. G. (1964). The amino acid requirements of Acanthamoeba sp. Neff. J. Protozool. 11, 98-100. Akins, R. A,, Gozs, S. M., and Byers, T. J. (1985). Factors regulating the encystment enhancing activity (EEA) of Acanthamoeba castellanii. J. Gen. Microbiol. 131,26092617. Auran, J. D., Star , M. B., and Jakobiec, F. A. (1987). Acanthamoeba keratitis. A review of the literature. Cornea 6, 2-26. Bacon, A. S., Frazer, D. G., Dart, J. K. G., Matheson, M., Ficker, L. A., and Wright, P. (1993). A review of 72 consecutive cases of Acanthamoeba keratitis, 1984-1992. Eye 7, 7 19-7 25. Badenoch, P. R. (1991). The pathogenesis of Acanthamoeba keratitis. Australian and New Zealand J. Ophthalmol. 19,9-20. Bamforth, S . S . (1985). The role of protozoa in litters and soil. J. Protozool. 32, 404-409.

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Bhattacharya, A., Ghosh, M., and Choudhury, A. (19871. Seasonal abundance of A c m thamoeba rhysodes (Singh, 1952) (Protozoa:Gymnamoebia) in a mangrove litter-soil ecosystem of Gangetic-estuary, India. J. Protozool. 34, 403-405. Bier, J. W., and Sawyer, T. K. (1990). Amoeba isolated from laboratory eyewash stations. Curr. Microbiol. 2 0 , 349-350. Borochovitz, D., Martinez, A. J., and Patterson, G. T. (1981). Osteomyelitis of a bone graft of the mandible with Acanthamoeba castellanii infection. Hum. Pathol. 12,573-576. Bottone, E. J., Perez, A. A., Gordon, R. E., and Qureshi, M. N. (1994). Differential binding capacity and internalisation of bacterial substrates as factors in growth rate of Acanthamoeba spp. J. Med. Microbiol. 40, 148-154. Bryant, R. J., Woods, L. E., Coleman, D. C., Fairbanks, B. C., McClellan, J. F., and Cole, C. V. (1982). Interactions of bacterial and amoeba1 populations in soil microcosms with fluctuating moisture content. Appl. Environ. Microbiol. 43, 747-752. Bunting, L. A., Neilson, J. B., and Bulmer, G. S. (1979). Cryptococcus neoformans: Gastronomic delight of a soil ameba. Sabouraudia 17, 225-232. Burger, R. M., Franco, R. J., and Drlica, K. (1994). Killing Acanthamoebae with polyaniinopropyl biguanide: Quantitation and kinetics. Antimicrob. Agents Chemother. 38, 886-888. Byers, T. J . , Bogler, S. A . , and Burianeck, L. L. (1983). Analysis of mitochondria1 DNA polymorphism detected in Acanthamoeba by restriction endonuclease analysis. Mol. Biochem. Parasitol. 8, 145-163. Byers, T. J. (1986). Molecular biology of DNA in Acanthamoebae, Amoeba, Enfamoeba, and Naegleria. Int. Rev. Cytol. 99, 311-340. Castellani, A. (1930). An amoeba found in cultures of yeast: Preliminary note. J. Trop. Med. Hyg. 33, 160. Centers for Disease Control (CDC) (1986). Acanthamoeba keratitis associated with contact lenses-United States. Morbid. Mortal. Weekly Rep. 35, 405-408. Centers for Disease Control (CDC) (1987). Acanthamoeba keratitis in soft-contact-lens wearers. Morbid. Mortal. Weekly Rep. 36, 397-404. Cohen, E. J., Parlato, C. J., Arentsen, J. J., Genvert, G. I., Eagle, R. C., Wieland, M. R., and Laibson, P. R. (1987). Medical and surgical treatment of Acanthamoeba keratitis. Am. J. Ophthalmol. 103, 615-625. Connor, C. G., Blocker, Y., and Pitts, D. G. (1989). Acanthamoeba culbertsoni and contact lens disinfection systems. Optom. Vis. Sci. 66, 690-693. Cursons, R. T. M., Brown, T. J . , and Keys, E. A. (1980). Effect of disinfectants on pathogenic free-living amoebae in axenic conditions. Appl. Environ. Microbiol. 40,62-66. Daggett, P.-M., Sawyer, T. K., and Nerad, T. A. (1982). Distribution and possible interrelationships of pathogenic and nonpathogenic Acanthamoeba from aquatic environments. Microb. Ecol. 8 , 371-386. Davies, D. J. G., Anthony, Y., Meakin, B. J , , Kilvington, S., and Anger, C. B. (1990). Evaluation of the anti-acanthamoebal activity of five contact lens disinfectants. ICLC 17, 14-20. Donzis, P. B., Mondino, B. J . , Weissman B. A,, and Bruckner, D. A. (1989). Analysis of contact lens care systems contaminated with Acanthamoeba. Am. I. Ophthalmol. 108, 53-56. Drozanski, W., and Chmielwski, T. (1979). Electron microscopic studies of Acanthamoeba castellanii infected with obligate intracellular bacterial parasite. Acta Microbiol. Pol. 28. 123-133.

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Fritsche, T. R., Gautom, R. K., Seyedreza, S., Bergeron, D. L., and Lindquist, T. D. (1993). Occurrence of bacterial endosymbionts in Acanfhamoeba spp. isolated from corneal and environmental specimens and contact lenses. J. Clin. Microbiol. 31, 1122-1126. Cast, R. J., Fuerst, P. A., and Byers, T. J. (1994). Discovery of group I introns in the nuclear small subunit ribosomal RNA genes of Acanthamoeba. Nucleic Acids Res. 22, 592596. Gautom, R. K., Lory, S., Seyedirashti, S., Bergeron, D. L., and Fritsche, T. R. (1994). Mitochondria1 DNA fingerprinting of Acanthamoeba spp. isolated from clinical and environmental sources. J. Clin.Microbiol. 32,1070-1073. Giovannini, A., Tittarelli, R., Bertelli, E., Frongia, G. B., Mariotti, C., Manso, E., and Biavasco, F. (1994). Bilateral Acanthamoeba keratitis in a gas-permeable contact lens wearer. Ophthalmologia 208, 321-324. Gorlin, A. I., Gabriel, M. M., Wilson, L. A,, and Ahearn, D. G. (1996a). Binding of Acanthamoeba to hydrogel contact lenses. Curr. Eye Res. 15,151-155. Gorlin, A. I., Gabriel, M. M., Wilson, L. A,, and Ahearn, D. G. (1996b). Effect of adhered bacteria on the binding of Acanthamoeba to hydrogel lenses. Arch. Ophthalmol. 114, 576-580. Griffiths, A. J., and Hughes, D. E. (1968). Starvation and encystment of a soil amoeba Hartmannella castellanii. J. Protozool. 15,673-677. Griffiths, B. S . (1990). A comparison of microbial-feeding, nematodes and protozoa in the rhizosphere of different plants. Bid. Fertil. Soils 9,83-89. Hay, J., Kirkness, C. M., Seal, D. V., and Wright, P. (1994). Drug resistance and Acanthamoeba keratitis: The quest for alternative antiprotozoal chemotherapy. Eye 8 , 555-563. John, T., Desai, D., and Sahm, D. (1991). Adherence of Acanfhamoeba castellanii to new daily wear, extended wear, and disposable soft contact lenses. CLAO J. 17,109-113. Jones, D. B., Visvesvara, G. S., and Robinson, N. M. (1975). Acanfhamoeba polyphaga keratitis and Acanthamoeba uveitis associated with fatal meningoencephalitis. Trans. Ophthalmol. Soc. UK 95, 221-232. Kelly, L. D., and Xu, L. (1995). The effect of Acanthamoeba concentration on adherence to four types of unworn soft contact lenses. CLAO J. 21, 27-30. Kelly, L. D., Long, D., and Mitra, D. (1995). Quantitative comparison of Acanfhamoeba castellanii adherence to rigid versus soft contact lenses. CLAO J. 24, 111-113. Kilvington, S . (1993). Acanthamoeba trophozoite and cyst adherence to four types of soft contact lens and removal by cleaning agents. Eye 7,535-538. Kilvington, S.,and Larkin, D. F. P. (1990). Acanthamoeba adherence to contact lenses and removal by cleaning agents. Eye 4,589-593. Kilvington, S.,and White, D. G. (1994). Acanfhamoeba: Biology, ecology and human disease. Rev. Med. Microbiol. 5, 12-20. Kilvington, S., Beeching, J. R., and White, D. G. (1991). Differentiation of Acanthamoeba strains from infected corneas and the environment by using restriction endonuclease digestion of whole-cell DNA. J. Clin. Microbiol. 29, 310-314. Kyle, D. E., and Noblet, G. P. (1987). Seasonal distribution of thermotolerant free-living amoebae. 11: Lake Issaqueena. J. Protozool. 34,10-15. Larkin, D. F. P., Kilvington, S., and Easty, D. L. (1990). Contamination of contact lens storage cases by Acanthamoeba and bacteria. Br. J. Ophthalmol. 74,133-135. Larkin, D. F. P., Kilvington, S., and Dart,J. K. G. (1992). Treatment of Acanthamoeba keratitis with polyhexamethylbiguanide. Ophthalmology 99,185-191.

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Lasman, M. (1987). Encystment-inducing factor from cultures of Acanthamoeba palestinensis. Dev. Growth and Differ. 29, 547-552. Lasman, M., and Shafran, A. (1978). Induction of encystment in Acanthamoeba palestinensis. Factors influencing cyst formation. I. Protozool. 25, 489-491. Lawin-Brussel, C. A., Refojo, M. F., Leong, F, Hanrinen, L., and Kenyon, K. R. (1993). Effect of I! aeruginosa concentration in experimental contact lens-related microbial keratitis. Cornea 12, 10-18. Ledee, D. R., Hay, J., Byers, T. J., Seal, D. V., and Kirkness, C. M. (1996). Aconthamoeba griffini. Molecular characterization of a new corneal pathogen. Invest. Ophthalmol. Ws. Sci. 37, 544-550. Lindquist, T. D., Slier, N. A., and Doughman, D. J. (1988). Clinical signs and medical therapy of early Acanthamoeba keratitis. Arch. Ophthalmol. 106, 73-77. Ludwig, I. H., Meisler, D. M., Rutherford, I., Bican, F. E., Langston, R. H., and Visvesvara, G. S. (1986). Susceptibility of Acantharnoeba to soft contact lens disinfection systems. Invest. Ophthalmol. Vis. Sci. 27, 626-628. Martinez, A. J. (1985). In “Free-Living Amebas: Natural History, Prevention, Diagnosis, Pathology and Treatment of Disease,” p. 156. CRC Press, Boca Raton, FL. Mathers, W. D., Sutphin, J. E., Folberg, R., Meier, P. A., Wenzel, R. P., and Elgin, R. G. (1996). Outbreak of keratitis presumed to be caused by Acanthamoeba. Am. J. Ophthalmol. 121, 129-142. Mergeryan, H. (1991). The prevalence of Acanthamoeba in the human environment. Rev. Infect. Dis. 13, 410-412. Miller, M. J., and Ahearn, D. G. (1987). Adherence of Pseudomonas aeruginosa to hydrophilic contact lenses and other substrata. ]. Clin. Microbiol. 25, 1392-1397. Moffat, J. F., and Tompkins, S. L. (1992). A quantitative model of intracellular growth of Legionella pneumophila in Acanthamoeba castellanii. Infect. Immun. 60, 296-301. Morton, L. D., McLauglin, G. L., and Whiteley, H. E. (1991). Effects of temperature, amebic strain, and carbohydrates on Acanthamoeba adherence to corneal epithelium in vitro. Infect. Immun. 59, 3819-3822. Nagington, J., Watson, P. G., Playfair, T. 7.. McGill, J., and Jones, B. R. (1974). Amoebic infection of the eye. Lancet 2, 1537-1540. Neff, R. J. (1957). Purification, axenic cultivation, and description of a soil amoeba, Acanthamoeba sp. 1.Protozool. 4, 176-182. Nerad, T. A., Sawyer, T. K., Lewis, E. J., and McLaughlin, S. M. (1995). Aconthamoeba pearcei n. sp. (Protozoa:Amoebida) from sewage contaminated sediments. 1. Euk. Microbiol. 42, 702-705. Nero, L. C., Tarver, M. G., and Hederick, L. R. (1964). Growth of Acanthamoeba castellanii with the yeast Torulopsis farnata. J. Bacteriol. 87, 220-225. Page, F. C. (1967). Taxonomic criteria for limax amoebae with descriptions of three new species of Hartmannella and three of Vahlkampfia. 1.Protozool. 14, 499-521. Paszko-Kolva, C., Yamamoto, H., Shahamat, M., Sawyer, T. K., Morris, G., and Colwell, R. R. (1991). Isolation of amoebae, Pseudomonas and Legionella spp. from eyewash stations. Appl. Environ. Microbiol. 57, 163-167. Penley, C. A., Willis, S. W., and Sickler, S. G . (1989). Comparative antimicrobial efficacy of soft and rigid gas permeable contact lens solutions against Acanthamoeba. CLAO

I. 15, 257-260. Pfister, D. R., Cameron, J, D., Krachmer, J. H., and Holland, E. J. (1996). Confocal microscopy findings of Acanthamoeba keratitis. Am. ]. Ophthalmol. 121, 119-128.

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Preston, T. M., and King, C. A. (1984). Binding sites for bacterial flagella at the surface of the soil amoeba Acanthamoeba. J, Gen. Microbiol. 130,1449-1458. Pussard, M., and Pons, R. (1977). Morphologie de la paroi kystique et taxonomie du genre Acanthamoeba (Protozoa, Amoebida). Protistologica 13,557-598. Qureshi, M. N., Perez, A. A., 11, Madayag, R. M., and Bottone, E. J. (1993). Inhibition of Acanthamoeba species by Pseudomonas aeruginosa: Rationale for their selective exclusion in corneal ulcers and contact lens care systems. J. Clin. Microbiol. 31, 1908-1910. Sawyer, T. K. (1970). The influence of seawater media on growth and encystment of Acanthamoeba polyphaga. Proc. Helminthological SOC. Wash. 37,182-189. Sawyer, T. K. (1989). Free-living pathogenic and nonpathogenic amoebae in Maryland soils. Appl. Environ. Microbiol. 55,1074-1077. Sawyer, T.K., and Griffin, J. L. (1975). A proposed new family, Acanthamoebidae n. fam. (order Amoebida), for certain cyst-forming filose amoebae. Trans. A m . Microsc. SOC. 94,93-98. Sawyer, T. K., Nerad, T. A., Daggett, l?-M., and Bodammer, S. M. (1987). Potentially pathogenic protozoa in sediments from oceanic sewage disposal sites. In “Oceanic Processes in Marine Pollution, Vol. 1: Biological Processes and Wastes i n the Ocean” (J. M. Capuzzo and D. R. Kester, eds.), pp. 184-194. Robert E. Kruger Publishing Co., Malahar, FL. Sawyer, T. K., Nerad, T. A., and Visvesvara, G. S. (1992). Acanthamoeba jacobsi sp. n. (Protozoa:Acantbamoebidae) from sewage contaminated ocean sediments. Proc. Helminthological SOC. Wash. 39,223-226. Sawyer, T. K., Nerad, T.A., Lewis, E. J., and McLaughlin, S. M. (1993). Acanthamoeba stevensoni n. sp. (Protozoa:Amoebida) from sewage contaminated shellfish beds in Raritan Bay, New York. J. Euk. Microbiol. 40, 742-746. Sawyer, T. K., Nerad, T. A., Gaines, C. J., Small, E. B., and Cooper, R. A. (1996). Bacterial and protozoan indicators of sewage contamination of deepwater sediments at Hudson Canyon. J. Marine Env. Eng. In press. Schuster, F. L., and Jacob, S. (1992). Effects of magainins on amoeba and cyst stages of Acanthamoeba polyphaga. Antimicrob. Agents Chemother. 36, 1263-1271. Schuster, F. L., Rahman, M., and Griffith, S. (1993). Chemotactic responses of Acanthamoeba castellanii to bacteria, bacterial components, and chemotactic peptides. Trans. A m . Microsc. SOC.112,43-61. Seal, D. V., Bennet, E. S., McFadyen, A. K., Todds, E., and Tomlinson, A. (1995a). Differential adherence of Acanthamoeba to contact lenses: effects of material characteristics. Optom. Vis. Sci. 72,23-28. Seal, D. V., Hay, J., and Kirkness, C. M. (1995b). Chlorhexidine or polyhexamethylene biguanide for Acanthamoeba keratitis. Lancet 345,136. Singh, B. N. (1945). The selection of bacterial food by soil amoebae, and the toxic effects of bacterial pigments and other products on soil protozoa. Br. J. Exp. Pathol. 26, 316-325. Stehr-Green, J. K., Bailey, T. M., and Visvesvara, G. S. (1989). The epidemiology of Acanthamoeba keratitis in the United States. Am. J, Ophthalmol. 107,331-336. Stevens, A. R., and Willaert, E. (1980). Drug sensitivity and resistance of four Acanthamoeba species. Trans. Roy. SOC. Trop. Med. Hyg. 74,806-808. Stratford, M. l?, and Griffiths, A. J. (1978). Variations in the properties and morphology of cysts of Acanthamoeba castellanii. J. Gen. Microbiol. 108,33-37.

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‘I’alamo, J. D., and Larkin, D. S. (1993). Bilateral Acanthamoeba keratitis and gas-permeable contact lenses. Am. 1.Ophthalmol. 116, 651-651. Tirado-Angel, J., Gabriel, M. M., Wilson, L. A,, and Ahearn, D. G. (1996). Effects of polyhexamethylene biguanide and chlorhexidine on four species of Acanthamoeba in vitro. C u m Eye Res. 15, 225-228. Uhelaker, J. E. (1991). Acanthnmoeba spp.: Opportunistic pathogens. Trans. Am. Microsc. sac. 110,289-299. Visvesvara, G. S. (1986). Susceptibility of Acanthamoeba to soft contact lens disinfection systems. Invest. Ophtbalmol. Vis. Sci. 27, 626-628. Visvesvara, G. S. (1995). Pathogenic and opportunistic free-living amoebae. In “Manual of Clinical Microbiology” (E. H. Lennette, A. Balows, W. J. Hausler, Jr., and J. P. Truant, eds.), 6th ed., pp. 1196-1203. Am. SOC. Microbiol., Washington, DC. Wang, X., and Ahearn, D. G. (1997). Effect of bacteria on survival and growth of Acanthamoeba castellanii. Curr. Microbial. 34.In press. Wilson, L. A., Sawant, A. D., Simmons, R. B., and Ahearn, D. G. (1990). Microbial contamination of contact lens storage cases and solutions. Am. J. Ophtbolmol. 110, 193-198. Wright, P., Warhurst, D. C., and Jones, B. R. (1985). Acanthamoeba keratitis successfully treated medically. Br. 1.Opbthalmol. 69,778-782. Wright, S. J. L., Redhead, K., and Maudsley, H. (1981). Acantharnoeba castellanii, a predator of cyanobacteria. J. Gen. Microbiol.125, 293-300. Yagita, K., and Endo, T. (1990). Restriction enzyme analysis of mitochondria1 DNA of Acanthamoeba strains in Japan. J. Protozool. 37,570-575. Zanetti, S., Fiori, P. L., Pinna, A,, Usai, S., Carta, F., and Fadda, G. (1995). Susceptibility of Acarithnmneba castellanii to contact lens disinfecting solutions. Antimicrob. Agents Chemother. 39,1596-1598.

Marine Microorganisms as a Source of New Natural Products V. S. BERNAN, M. GREENSTEIN, AND W. M. MAIESE Wyeth-Ayerst Research Natural Products Research Pearl River, New York 10965

I. Introduction 11. Natural Products from Marine Microorganisms A. Free-Living Microorganisms B. Commensal Bacteria C. Symbiotic Bacteria D. Bioactive Metabolites from Fungi 111. Overview of Wyeth-Ayerst (W-AR) Marine Natural Products Program A. Microbial Isolation and Taxonomy B. Fermentation Studies C. Biological Activities D. Marine Bacteria E. Marine Fungi IV. Marine Biotechnology V. Summary References

I . Introduction Natural products have proven to be rich sources of novel compounds exhibiting many different biological activities. Their chemical structures have been diverse and complex, and they have often demonstrated selective activities in various biological systems. Historically, most bioactive products of microbial origin have come from terrestrial bacteria belonging to one taxonomic group, the Actinomycetales. Although this group continues to be a prolific source of bioactive compounds, it is clear that the yield of novel metabolites is decreasing, and new sources of bioactive natural products must be investigated (Iwai and Takahashi, 1992). In the 1960s, exploration of the marine environment began as an exciting resource for the isolation of new compounds. Initially, the marine natural products field focused on the chemical characterization of metabolites from macroscopic marine algae and invertebrate animals, and to date there are more than 3700 new natural products reported from these groups (Faulkner, 1995). Although macroorganisms continue to supply us with new metabolites of unprecedented complexity and 57 ADVANCES IN APPLIED MICROBIOLOGY. VOLUME 43 Copyright 0 1997 by Ar ademic Press, Inc A l l rights of reproduction in any form reaerved 0065-2164/97 $25 00

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variety, the search has now been expanded to include marine microorganisms, which are a readily renewable resource. The many habitats in the sea have created niches for the evolution of diverse life forms. The marine environment ranges from nutrient-rich regions to nutritionally sparse locations where only a few organisms can survive. As a result, marine microorganisms encompass a complex and diverse assemblage of microscopic life forms, of which less than 5 % have been cultured or identified. The metabolic and physiological capabilities of marine microorganisms that allow them to survive in their unique habitats also provide great potential for the production of metabolites not found in terrestrial environments. It is estimated that less than 1%of potentially useful chemicals from the marine environment have been screened thus far, with microbial products representing approximately 1%of that total number (Colwell, 1993). Even though comparatively little research has been directed toward the study of natural products from marine microorganisms, results from preliminary studies are encouraging. The data from these investigations demonstrate complex chemical interactions between marine bacteria and their hosts, including systems of signalling and territorial marking. Initial investigations have shown that the compounds isolated from marine microorganisms have demonstrated antibiotic, antitumor, and other pharmacological activities (Fenical and Jensen, 1993). Thus, it is clear that the marine environment represents a relatively untapped resource for new bioactive compounds. The need for new therapeutic compounds is always expanding because of the evolving resistance of microorganisms to existing antibiotics, the emergence of new viral diseases, and the appearance of drug-resistant tumors. Microorganisms have traditionally met the demand for finding novel, useful compounds, and in this review we will address the tremendous potential that the marine environment holds for the discovery of additional new bioactive agents. This article will provide a brief review of natural products isolated from marine microorganisms, information obtained from our marine natural products program, and lastly a discussion of new directions in marine biotechnology and their impact on the drug discovery process. II. Natural Products from Marine Microorganisms

A. FREE-LIVING MICROORGANISMS Early investigations of marine microorganisms focused on the discovery of antimicrobial agents. One of these marine metabolites, isolated by Burkholder, was the highly brominated antibiotic 2,3,4-tribromo-5-

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(l’-hydroxy-2’,4’-dibromophenyl)pyrrole. This compound was isolated from an Alteromonas species obtained from the surface of the Caribbean seagrass Thalassia collected near La Paguera, Puerto Rico. This compound showed in vitro activity against Gram-positive bacteria, with minimum inhibitory concentrations (MICs) ranging from 0.0063 to 0.2 pg/ml (Burkholder et al., 1966). This initial report was followed by the characterization of several additional antimicrobial brominated pyrroles: tetrabromopyrrole, hexabromo-Z,2’-bispyrrole, and 2-(2’-hydroxy3’,5’-dibromophenyl)-3,4,5-tribromopyrrole.These compounds were purified by Andersen from a purple-pigmented Alteromonas sp. isolated from a seawater sample collected at the North Pacific Gyre. Tetrabromopyrrole showed moderate antimicrobial activity in vitro against Staphylococcus aureus, Escherichia coli, Pseudomonas aeruginosa, and Candida albicans (Andersen et al., 1974). In 1977 this same group isolated an antibiotic-producing Pseudomonas from a tidepool in La Jolla, California. Isolation and purification of the metabolites yielded 6-bromo-indole carboxaldehyde, its debromo analogue, and a mixture of 2-n-pentyl- and 2-n-heptylquinolinol. The most interesting of these was 2-n-pentylquinolinol, which had potent activity against S. aureus. The unique metabolite 6-bromo-indole carboxaldehyde lacked antimicrobial properties (Wratten et al., 1977). The isolation of these highly brominated compounds illustrates a unique property that distinguishes marine from terrestrial bacteria. Marine bacteria have a common mechanism to incorporate bromine or other halogens into organic compounds that can potentially lead to enhanced bioactivities. Studies on unicellular bacteria continued with the work of Holland, who isolated a series of C16 aromatic acids from the marine bacterium Alteromonas rubra. These acids, derived from fatty acid synthesis, showed pharmacological properties in bronchodilator and neuromuscular assays (Holland eta]., 1984). More recently, Needham et al. (1991) reported a novel metabolite, oncorhycolide, isolated from a surface seawater sample taken near a chinook salmon farm. Oncorhycolide is a unique lactone that may have a polyketide origin. However, the oxygenation and methylation patterns may be derived by unique reactions not previously observed in nature. Fenical’s group from Scripps Institute of Oceanography isolated a deep-sea sediment bacterium that produced a series of novel cytotoxic and antiviral macrolides, the macrolactins A-F. This bacterium was an unidentified Gram-positive organism that produced six macrolides and two open-chain hydroxy acids only when fermented in the presence of salt at atmospheric pressure. Macrolactin A was the predominant com-

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pound produced. It showed moderate antibacterial activity but was quite active against B16-Fl0 murine melanoma in vitro with an IC5,-, value of 3.5 pg/ml. Of greater significance, the compound inhibited several viruses, including Herpes simplex (IC50= 5.0 pg/ml) and HIV, the human immunodeficiency virus (IC5,, = 10.0 pg/ml) (Fenical and Jensen, 1993). Two new caprolactams were isolated by Davidson and Schumacher (1993) from an unidentified Gram-positive bacterium cultured from a deep-ocean sediment. The caprolactams A and B were obtained as an inseparable mixture and are composed of cyclic-L-lysine linked to 7methyloctanoic acid and 6-methyloctanoic acid, respectively. Natural products containing a lysine are uncommon and have only been reported in several sponges and one fungus. Thus, one may speculate that these compounds isolated from sponges may in fact be produced by a symbiotic microorganism, The compounds are mildly cytotoxic toward human epidermoid carcinoma and colorectal adenocarcinoma cells with MIC values of 10 and 5 pg/ml, respectively, and exhibit antiviral activity toward Herpes simplex type I1 virus at a concentration of 100 pg/ml. Since terrestrial actinomycetes have been such prolific producers of bioactive molecules, it was natural to investigate marine species of the same group. Beginning in the early 1970s, scientists at the Institute of Microbial Chemistry, Tokyo, investigated metabolites from marine actinomycetes. They, as well as others, have discovered that streptomycetes inhabit shallow-sea areas while Micromonospora and nocardioform actinomycetes predominate in the deep sea (Weyland and Helmke, 1988; Jensen et al., 1991). One of the first compounds described by Okami (1993) was a benzanthraquinone antibiotic isolated from the actinomycete Chainia purpurgensa SS-228. This antibiotic selectively inhibited Gram-positive bacteria with MIC values between 1 and 2 pg/ml and was active against Ehrlich ascites tumor cells in mice. It also produced a hypotensive effect in mice, probably due to its inhibition of dopamine hydroxylase in the pathway of epinephrine biosynthesis. Interestingly, this culture only produced the antibiotic when it was fermented in diluted yeast extract medium containing “Kobu Cha” (the brown seaweed Laminaria) and with the addition of 3% NaC1. Istamycins A and B were also isolated by this Tokyo group from a marine streptomycete collected near the shore of Tenjin Island, Japan. The culture produced bioactivity only when it was fermented in a seawater-containing medium. The istamyciiis strongly inhibited both Gram-positive and -negative bacteria with MIC values between 0.10 and

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3.0 pg/ml against various species of Staphylococcus, Bacillus, Corynebacterium, and Escherichia. These compounds are related to the fortimicin-sporaricin group of aminoglycoside antibiotics but are more interesting because of their resistance to aminoglycoside-inactivating enzymes. In addition, the Tokyo group carried out fusions of protoplasts of the istamycin-producing strain with protoplasts from a streptomycinproducing streptomycete. One of the resulting clones produced the novel antibiotic indolizamycin, which is entirely unrelated to the parent compounds (Okami, 1993; Hotta et al., 1980). Several of the more unusual compounds isolated from marine actinomycetes are the aplasmomycins A-C. These were obtained from the fermentation of Streptomyces griseus SS-20 isolated from shallow-sea mud in Sagami Bay, Japan. Again, the antibiotic was produced only under low-nutrient conditions containing “Kobu Cha” and 3% NaC1. The aplasmomycins are inhibitory against Gram-positive bacteria such as S. aureus, B. subtilis, B. anthracis, and Corynebacterium smegmatis with MIC values between 0.8 and 3.0 pg/ml. When administered orally to mice infected with Plasmodium berghei (the causative agent of malaria), the number of red cells containing the protozoan decreased and all treated mice survived, hence the name aplasmomycin. X-ray crystallography revealed that the structure contained a symmetric ring in which boron was placed in the center and bound with a crown etherlike structure (Okami et al., 1976; Sat0 et al., 1978). Takahashi in Japan detected a monoterpene alkaloid, altemicidin, by screening for antitumor activity using the common brine shrimp Artemia salina. Reports have shown that brine shrimp toxicity correlates well with antitumor activity. This novel alkaloid was purified from a marine strain of Streptomyces sioyaensis SA-1758 isolated from a sea mud sample collected in the northern part of Japan. It produced potent antitumor activity in vitro against L1210 murine leukemia and IMC carcinoma cell lines with IC50 values of 0.84 and 0.82 pg/ml, respectively. Unfortunately, the compound showed weak antibacterial activity and was relatively toxic in mice (LD5, = 0.3 mg/kg iv) [Takahashi et al., 1989a,b). A study from the shallow sediments of Bodega Bay, California, resulted in the isolation of an unknown Streptomyces sp. that was found to produce compounds with broad antibacterial activity. Subsequent fermentation of this isolate in a saltwater-based medium produced four new alkaloid esters of the phenazine class which contained the sugar L-quinovose at the 2’ and 3’ positions. These compounds were found to exhibit antibacterial activity against Gram-negative and -positive bacteria with MIC values in the 1-4 pg/ml range (Pathirana et al., 1992).

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In a similar study, the same group isolated marinone and its debromo analogue debromomarinone from an estuarine actinomycete. Both compounds possess a common naphthoquinone with rare sesquiterpenoid structural components. These new molecules are among a rare group of bacterial metabolites produced via mixed biosynthesis involving both acetate and terpene pathways. Marinone inhibits Bacillus subtilis at 1 pg/ml, and debromomarinone has MIC values of 1-2 pg/ml against S. aureus, S. epidermis, S. pneumoniae, and S . pyogenes (Pathirana et al., 1993).

Investigations on marine actinomycetes that were non-Streptomyces led Fenical’s group to isolate a Maduramycete from the shallow sediments of Bodega Bay. Fermentation of this actinomycete resulted in the isolation of maduralide, a new member of a rare class of macrolides. This compound is a member of a rare %-membered ring lactone group represented by rectilavendomycin (Pathirana et al., 1991). More recently, the same group isolated an unidentified marine actinomycete from sediment collected in the Agua Hedionda Lagoon in Carlsbad, California. The lagunapyrones A-C were isolated from this culture when fermented in the presence of a seawater-based medium. The lagunapyrones are closely related to the a-pyrones functionalized by a highly methyl-branched C19 side chain and showed modest in vitro cytotoxicity (EDso = 3.5 pg/ml) against the human colon cancer cell line HCT-116. The carbon skeleton of the lagunapyrones has not been previously described. These compounds could be biosynthetically derived by the condensation of acetate and/or propionate units, or by a combination of these that includes the methylation of the carbon chain through the methionine pathway (Lindel et al., 1996). Screening with a novel macrophage effector bioassay, the Tokyo group isolated a bioactive siderophore from a strain of Alteromonas haloplanktis SB-1123 isolated from a deep-sea mud sample collected off the Aomori Prefecture coast at a depth of 3300 m. This compound was only produced when the strain was fermented in a substrate containing seawater, sardine and cuttlefish powders, and maltose as the carbon source. Even though the strain was isolated at 700 atm pressure, it appeared to grow well at surface pressure and temperature. This compound was shown by X-ray crystallography to contain a 22-membered ring siderophore, and due to its structural relation to cataberin it was named bisucaberin. Bisucaberin has the unique biological property of rendering tumor cells susceptible to the cytolytic action of murine peritoneal macrophages. This approach of promoting natural immu-

MICROORGANISMS AS SOURCE OF NEW NATURAL PRODUCTS

63

nological actions may serve as an alternative to potent cytotoxic cancer chemotherapy (Kameyama et al., 1987; Takahashi et al., 1987). Some other examples of the potential of marine bacteria to produce diverse bioactive compounds include the discovery of a new enzyme that degrades the glucan of Streptococcus mutans, which is the cause of dental caries. The enzyme was isolated from a marine Bacillus and showed optimum activity at 37"C, which makes it favorable for use in the oral cavity (Okami et al., 1980). Another example is marinostatin, a protease inhibitor produced by an Alteromonas species, which is significantly different from existing inhibitors of this class of enzymes (Imada et a]., 1986).

B. COMMENSAL BACTERIA Commensal marine bacteria inhabit surfaces, tissues, and internal spaces of other organisms, but often the exact associations and interactions are not well understood. Documentation of these coexistences is increasing, but evidence supporting true symbiosis is still a matter of question. However, these associations are significant in the quest for new natural products because of their potential role in the production of novel metabolites. Umezawa and co-workers screened fermentation broths containing various bacterial species, isolated from the surface of seaweeds, for the production of polysaccharides. One genus, Flavobacterium ugliginosum, isolated from the surface of seaweed, was found to produce marinactan, which is capable of suppressing sarcoma-180 tumors in mice. At daily doses of 10-50 mg/kg in mice, marinactan inhibited 75-95% of the growth of these solid tumors. Marinactan is a neutral heteroglycan consisting of fucose, mannose, and glucose (Umezawa et al., 1983). Fenical's group has also investigated actinomycetes living on the surfaces of marine invertebrates. He and co-workers isolated a streptomycete from the surface of a gorgonian coral (Pacifigorgia sp.) collected from the Gulf of California, Mexico. When fermented in marine media, the isolate produced several metabolites, including the 20-hydroxy derivative of oligomycin A, the 5-deoxy derivative of enterocin, and the octalactins A and B. The octalactins belong to a new structure class, which are C19 ketones possessing rare eight-membered ring lactone functionalities. Octalactin A demonstrates potent in vitro cytotoxicity against B16-Fl0 murine melanoma and HCT-116 human carcinoma cell lines. Conversely, the surface of a jellyfish yielded a Streptomyces sp. that produced two new bicyclic peptides, salinamides A and B, which

64

V. S. BERNAN at 01.

have novel depsipeptide backbones. The salinamides exhibit activity against all Gram-positive microorganisms tested (Fenical and Jensen 1993).

Commensal bacteria have also been isolated from fish. Anguibactin, another interesting siderophore, was isolated from the fish pathogen Vibrio anguillarum. It is a novel catechol with thiazoline and imidazole rings, the biosynthesis of which has been genetically linked to a 65 kb plasmid. Anguibactin and its required plasmid were demonstrated to be essential in the virulence of this pathogen. One can speculate that, since iron is present in limited quantities in the sea, marine bacteria may have evolved this ability to produce siderophores (Jalal et al., 1989).More recently, Takahashi isolated a strain of Streptomyces hygroscopicus from the gastrointestinal tract of the marine fish Halichoeres bleekeri. When this culture was fermented in a medium containing artificial seawater, it produced a novel class of macrolide, the halichomycins. Halichomycins exhibited potent cytotoxicity (EDs0 = 0.13 yg/ml) in a P388 lymphocytic leukemia test system (Takahashi et al., 1994).

Additional examples of commensal bacteria are described in studies from Andersen’s group. They purified andrimid and moiramides A-C from a marine Pseudornonas fluorescens grown on agar. This culture was isolated from an unidentified tunicate collected on Prince of Wales Island in Moira Sound, Alaska. Andrimid and moiramide B exhibited potent in vitro antibacterial activity, especially against methicillin-resistant S. aureus. Andrimid labeling with stable isotopes demonstrated that the acylsuccinimide fragment, which is required for bioactivity, was derived from a combination of acetate and amino-acid building blocks that proceed through a putative dipeptide-like intermediate generated from y-amino-P-keto acids (Needham et al., 1994). Lastly, the same group also isolated a Bacillus sp. from the tissues of a marine worm collected in Papua, New Guinea, that produced the novel cyclic decapeptide antibiotic loloatin B. This antibiotic has structural similarities to the tyrocidines and inhibits the growth of methicillin-resistant S . aui-eus, vancomycin-resistant Enterococcus sp., and penicillin-resistant Streptococcuspneumoniae with MICs of 1-2 pg/ml (Gerard et al., 1996). Recently, a group at Pharma Mar isolated a new species of Micromonospora from an unidentified marine soft coral collected off the coast of Mozambique. This culture produced a novel depsipeptide designated PM-93135 when fermented in a nutrient-rich medium. PM-93135 exhibited antibacterial activity against S. aureus, B. subtilis, and Micrococcus luteus and inhibited RNA synthesis in P388 cells with an IC50of 0.008

MICROORGANISMS AS SOURCE OF NEW NATURAL PRODUCTS

65

pg/ml. In addition, this compound demonstrated significant antitumor activity against P388, human lung carcinoma A-549, human colon carcinoma HT-29, and human melanoma MEL-28 with IC,,,s of 0.0002, 0.002, 0.01, and 0.0025 yglml, respectively (Baz et al., 1995).

BACTERIA C. SYMBIOTIC Even though it is a widespread phenomenon, little is known about the symbiosis of prokaryotic organisms with eukaryotic taxa. These symbioses can range from relatively loose coexistence to highly interdependent intracellular associations. One of the first reports of marine symbiosis described relationships between chemoautotrophic bacteria and marine invertebrates in deep-sea hydrothermal vents. Hydrothermal vents emit sulfide, which, in turn, provides the necessary energy and reducing power for chemoautotrophic bacteria. Many of these bacteria inhabit the large populations of animals living around these vents, sometimes directly providing their sole source of food. Furthermore, the invertebrates living there either contain a greatly reduced digestive tract or have no digestive tract at all, which forces them to rely upon these endosymbionts for survival (Distel et al., 1988). Other examples of symbionts have been described, but unfortunately pure viable cultures of these bacteria have not been attainable and therefore cannot be used for the production of bioactive compounds. Bacterial symbionts that are chemoheterotrophic have been described and isolated in pure culture. Fenical studied the resistance of the estuarine shrimp Palaemon macrodactylus to pathogens and observed that the eggs of this animal harbored bacterial epibionts. When these epibionts were removed from the eggs by antibiotic treatment, rapid infestation by pathogenic fungi occurred. A symbiotic bacterium, an Alteromonas species, was isolated, fermented, and found to produce a potent antifungal agent, 2,3-indolinedion, also known as isatin. Isatin has been known for many years as a synthetic intermediate in the production of indigo dyes but not as an antifungal agent (Gil-Turnes et al., 1989). Gil-Turnes found a similar case when the eggs of the American lobster Hornarus americanus were found to be totally covered with an unidentified unicellular bacterium. Upon culturing, the phenolic compound tyrosol (2-p-hydroxyphenyl ethanol) was isolated. The bacterium produces this phenol in vast quantities, which are sufficient to control pathogenic microorganisms (Gil-Turnes and Fenical, 1992). GilTurnes also investigated the tropical cyanobacterium Microcoleus lyngbyaceus recovered from waters off of Puerto Rico. These studies in-

66

V. S. BERNAN et al.

volved the sampling of 65 sites around the island for the isolation of surface bacteria associated with this filamentous microorganism. Only four specific strains of highly colored bacteria were consistently isolated from the surface of the cyanobacterium. Since all sampling sites were distinct, each bacterial species isolated was obtained from an individual source, thus eliminating the possibility of cross-contamination. Fermentation of the bacteria revealed that all four strains produced the same antifungal and antibacterial quinone at a level of 20 pg/ml. Interestingly, this compound had never been isolated as a natural product. It appears to be synthesized by the bacteria through an oxidative cleavage of a ubiquinone-type precursor (Fenical and Jensen, 1993). Marine invertebrates are considered to be particularly good sources of novel metabolites and symbiotic microorganisms. Sponges, for example, can have 40% of their cellular volume occupied by associated bacteria. However, identification of the symbiotic or parasitic microorganism(s) responsible for the production of a specific metabolite can be difficult. Often only circumstantial evidence obtained from the marine invertebrate is used to indicate that the metabolite originated from a symbiotic microorganism. This evidence usually consists of extremely low or variable metabolite yields, isolation of the same metabolite from diverse sources, or similar structure to a metabolite already described from a microorganism. One of the earlier studies that described the bacterial production of metabolites previously isolated from a sponge include the work of Stierle et al. (1988). A bright orange-pigmented Micrococcus sp. was isolated from the sponge Tedania ignis, which produced three diketopiperazines previously reported from the sponge. However, this evidence is not conclusive since a majority of Gram-negative bacteria produce diketopiperazines when fermented in rich media (Fenical, 1993). In another study, Elyakov and co-workers reported that two strains of a Vibrio sp. produced brominated diphenyl ethers that had originally been isolated from the sponge Dysidea sp. collected i n the Indian Ocean (Elyakov et al., 1991). However, no additional data have been reported. Shigemori et al. (1992) examined extracts of symbiotic bacteria associated with various marine invertebrates and isolated a new cyclic alkaloid, alteramide A. This compound was produced by an Alteromonas species isolated from the marine sponge Halichondria okadai collected off Nagai, Kanagawa, Japan. Alteramide A is a macrocyclic lactam containing dienone and dienoyltetramic acid functionalities that can undergo a [4 + 4) cycloaddition to generate a hexacyclic derivative. Alteramide A exhibited cytotoxicity against murine leukemia P388

MICROORGANISMS AS SOURCE OF NEW NATURAL PRODUCTS

67

cells, murine lymphoma LIZ10 cells, and human epidermoid carcinoma KB cells (Shigemori et al., 1992). Related macrocyclic lactams previously were isolated from terrestrial actinomycetes and a different marine sponge, but not from symbiotic bacteria. Although this bacterium was described as a symbiont of the sponge, the compound was never conclusively identified from extracts of the marine animal. Lastly, Oclarit and co-workers isolated a Vibrio sp. from the homogenate of the sponge Hyatella sp. collected along the coast of Oshima Island, Miyazaki, Japan. When this bacterium was cultured on marine agar, it produced the peptide antibiotic andrimid. This same bioactive substance was also found in the sponge extract, suggesting that the active component may be synthesized by the associated microorganism. Andrimid has been previously isolated from cultures of an Enterobacter sp. that is an intracellular symbiont of the Brown Planthopper Nilaparvatu lugens and was found to exhibit potent activity against Xanthomonas campestris pv. oryzae. Needham et al. (1994) also isolated andrimid, but from a marine Pseudomonas fluorescens, which was active against methicillin-resistant S. aureus. Due to the diversity of the microorganisms producing this toxin, one can speculate that the production of this compound can be encoded by genes transferable on a plasmid (Oclarit et a]., 1994). Probably the best example of marine symbionts are those responsible for producing many of the marine toxins that pose human health hazards. Surugatoxin, which specifically blocks nicotinic receptors and was a causative agent of shellfish poisoning in Japan, was initially isolated from the gut of the Japanese Ivory Shell mollusc Babylonia japonica. Subsequently, neosurugatoxin and prosurugatoxin, precursors of surugatoxin, were isolated from a Gram-positive Coryneform sp. obtained from the mid-gut of B. japonica (Kosuge et al., 1985). Following the isolation of surugatoxin, tetrodotoxin and anhydrotetrodotoxin were isolated from the fermentation media of three different species of bacteria. Before this discovery, the origin of these potent marine neurotoxins was considered to be the pufferfish. They have been isolated recently from crabs, octopus, and even an amphibian, all suggesting a microbial source. Some of the microbial sources include a Pseudomonas sp. from the surface of red algae, an Alteromonas sp. from deep-sea sediments, and a Vibrio sp. that originated from the skin of the pufferfish. It has been speculated that the diversity of the microorganisms producing this toxin may be due to one or more genes being transferable on a plasmid. More studies will have to be undertaken to fully explain the source of these toxins (Yasumoto e f al., 1986; Noguchi et a]., 1986; Yotsu et a]., 1987).

68

V. S. BERNAN et al.

The toxin involved in paralytic shellfish poisoning is saxitoxin, which is a metabolite known to be produced in cold-water habitats by the marine dinoflagellate Protogonyaulax. There have been inconsistencies in this concept since toxic shellfish have been observed in areas devoid of Protogonyaulax. Recently, Kodama et al. (1988, 1990) have shown evidence that a marine Moraxella sp. isolated from Protogonyaulas tamnrensis is responsible for the production of this toxin. FROM FUNGI D. BIOACTIVEMETABOLITES

Since the discovery of penicillin G in 1929, terrestrial fungi have been a major source of medicinals. However, there have been comparatively few attempts to explore the potential of marine fungi. As of 1991, only 321 marine fungi had been described, compared to 69,000 terrestrial fungi. One of the earliest reports in this area was the isolation of a small lactone, leptosphaerin, from cultures of a lignicolous Ascomycete, Leptosphaeria oraemaris. The lignicolous forms of marine fungi, which reside in submerged wood surfaces, are relatively common. Strongman et al. (1987) compared the antifungal activities of 27 lignicolous fungi to assess the degree of interspecies chemical similarity within an environmental group. Four isolates were found to produce inhibitory substances, and all were shown to produce the sesquiterpene diol culmorin in addition to the pigments obioninene and oreamarin. This work suggests that chemical defense is an important adaptation of lignicolous fungi competing for substrates. In the late 1970s, Okutani described the isolation of the common bacterial toxin gliotoxin from a marine Aspergillus sp. (Okutani, 1977). Ten years later, Shin and Fenical reported the structurally related metabolite gliovictin from the marine fungus Asferomyces cruciatus (Shin and Fenical, 1987). Kupka and co-workers detected antimicrobial activity from the basidiomycete Halocyphina villosn. Chemical analysis revealed that the antibiotic was derived from siccayne, an acetylenic hydroquinone that was also produced by the terrestrial fungus Helminthosporium siccans. Siccayne was weakly active against Gram-positive and -negative bacteria and also inhibited DNA and RNA synthesis in vitro (Kupka et nl., 1981). A trinor-eremophilane, dendryphiellin, was isolated by Guerriero et al. (1988) while searching for antibiotics produced by marine deuteromycetes. Dendryphiellin A, the first fungal trinor-eremophilane, was isolated from the fermentations of the marine deuteromycete

MICROORGANISMS AS SOURCE OF NEW NATURAL PRODUCTS

69

Dendryphiella salina. Closer examination of these fermentation broths revealed the related metabolites dendryphiellin B-D, which only vary in the positions of hydroxylation on the fatty-acid side chains. Lastly, dendryphiellin E was isolated from this culture, which proved that these compounds are derivatives of the ermophilane sesquiterpene class. Although no bioactivity was discovered, these compounds represent a new type of fungal natural product and again highlights the potential of marine fungi as a source of novel molecules (Guerriero et a]., 1989). Recent work from the Gloer group at the University of Iowa has lead to the isolation of new bioactive compounds from fungi. Poch et al. (1991) isolated a series of chemically similar naphthoquinones, dimer kirschsteinin and two new chlorinated diphenyl esters from a previously undescribed Kirschsteiniothelia species. Kirschsteinin is unusual because of its ethyliden bridge between two unsymmetrically substituted monomers, This compound demonstrated antibiotic effects against S. aureus and B. subtilis and cytotoxicity on several human tumor cell lines. A new polyketide named obionin, from the marine fungus Leptosphaeria obiones, was also reported from this group. L. obiones is a halotolerant ascomycete isolated from the surface of the salt marsh grass Spartina alterniflora. Obionin A inhibited the binding of dopamine D-1-selective ligands to membranes found in the central nervous system. Poch and Gloer (1991) also reported recovering the interesting lactones heliascolides A and B from the marine fungus Helicascus kanaloanus isolated from a mangrove swamp in Hawaii. The helicascolides are epimeric secondary alcohols at the C-3 position. The fungus Prussia aurantaica, also found in the mangroves, produces auranticin A, which has antibiotic activity against B. subtilis and S. aureus at concentrations of 5-50 pg/disk. Lastly, they have isolated a diacid, hymenoscyphin A, from the salt marsh fungus Hymenoscyphus sp. Hymenoscyphin consists of an unusual repeating alpha-hydroxy isobutyric acid group and exhibits toxicity against brine shrimp at 10 pg/ml (Poch et al., 1989a,b; Fenical and Jensen, 1993). In recent work reported from Japan, a number of fungi isolated from marine animals have been shown to produce novel compounds. The phomactins have been reported from a marine Phoma sp. that was isolated from the crab Chinoecetes opilio. These compounds are platelet-activating-factor agonists (Sugano et al., 1991). The terrestrial fungus Penicillium fellutanum has been isolated from the gut of the fish Apogon endekataenia. This strain of l? fellutanum was shown to produce the lipopeptides fellutamides A and B, which had cytotoxic as well as

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nerve-growth-factor-synthesis stimulatory action (Shigemori et al., 1991). Three cytotoxic quinazoline derivatives-fumiquinazolines A, B, and C-were obtained from Aspergillus fumigatus, which had been isolated from the fish gut of Pseudolabrus jabonicus (Numata et al., 1991). Ten novel leptosins, which are chaetocin derivatives, were isolated and characterized by Takahashi et al. (1995). They were isolated from the mycelium of a strain of Leptosphaeria sp. attached to the marine alga Sargassum tortile. The related leptosins are dimeric epipolysulfanyldiketopiperazines, where the sulfuric linkage may consist of two to four members. The leptosins are cytotoxic against P388 leukemia cells: however, leptosins A and C also exhibited antitumor activity against Sarcoma 180 ascites. A marine fungus of the class Hyphomycetes was isolated from the Indo-Pacific sponge Jaspis aff. johnstoni. Fermentations of this marine culture led to the isolation of the tricyclic sesquiterpenes coriolin B and dihydrocoriolin C as well as the novel chloriolines A and C. The chloriolines are chlorinated bi- or tricyclic sesquiterpenes. Coriolin B and dihydrocoriolin C were previously isolated from the terrestrial wood-rotting basidiomycete Coriolus consors. Coriolin B exhibited strong inhibition of human breast and CNS cell lines with ICsOvalues of 0.7 and 0.5 pM, respectively (Cheng et al., 1994). Lastly, in screening for new inhibitors of the mammalian cell cycle, Cui et al. (1996) isolated a new phthalide carrying an acetonyl group at the C-3 position called acetophthalidin. Acetophthalidin was purified from a Penicillium that was isolated from a sea-sediment sample collected from Miho, Sizuoka Prefecture, Japan. This compound was unstable in aqueous solution because of the presence of the acetonyl group at the C-3 position and was first isolated as a proto-inhibitor. This proto-inhibitor form was then converted into the active inhibitor by heating a water solution of the sample at pH 1 for 20 minutes in a boiling water bath. The compound completely inhibited the cell cycle progression of tsFT210 cells in the M phase at a final concentration of 6.25 pg/ml. Ill. Overview of Wyeth-Ayerst (W-AR) Marine Natural Products Program

A. MICROBIAL ISOLATION AND TAXONOMY

The search for novel bioactive compounds with therapeutic potential has become increasingly more difficult, and it is apparent that the discovery of unique natural products depends upon the development of effective strategies for the isolation and cultivation of novel microor-

MICROORGANISMS AS SOURCE OF NEW NATURAL PRODUCTS

71

TABLE I CHARACTERIZATION AND DISTRIBUTION OF MARINE ISOLATES Sample type

Grampositive

Gramnegative

Gramvariable

Fungi

3 04 300

56

202 189

15 6

19 59

12

Fiji Vancouver

115 54

18

72

35

5 1

2 6

18

2

Fiji Vancouver

138 84

32

99

5

0

7

70

2

5

2 0

Fiji Vancouver

40 293

12 25

23 199

0 6

1 54

4 9

79

4

18

2

8

Collection

Total

Fiji Vancouver

Actinomycete

Invertebrate 18

28

Core 10

Water

Vegetative

Shoreline Fiji

47

ganisms (Bull et al., 1992). We have expanded our search for biodiversity to include the isolation of marine microorganisms from the warm waters surrounding Fiji and the cold waters around Vancouver Island, Canada. In addition, collections have been conducted off the coasts of Florida and New England. The majority of these expeditions have been accomplished through collaborations with Dr. Chris Ireland, University of Utah, and Dr. Raymond Andersen, University of British Columbia. Microorganisms have been isolated from diverse samples including core sediments, invertebrate animals, water columns, marine vegetation, and shoreline sediment. This effort has resulted in a collection of over 650 microorganisms from the South Pacific Ocean, over 700 isolates from the North Pacific Ocean, and over 100 microbes from collections in the United States. The types and sources of microorganisms from the North and South Pacific Oceans are summarized in Table I. An analysis of the isolates revealed that the majority were Gram-negative bacteria (71%). The remaining isolates were identified as follows: Gram-positive bacteria (17%), Gram-variable bacteria (3%), fungi (lloh),and actinomycetes (9%). It is not surprising that the majority of these isolates were Gram-

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et al.

negative bacteria since they are reported to represent the largest and most diverse group of marine chemoheterotrophic bacteria. However, this representation may be inaccurate due to our lack of knowledge of the nutritional requirements of marine microorganisms and, therefore, their lack of culturability. In fact, Jensen et al. (1996) recently reported a study comparing the recovery of culturable bacteria from the surfaces of marine algae using high- and low-nutrient media. The low-nutrient medium comprised of seawater and agar supported the growth of significantly more bacteria than a high-nutrient medium composed of peptone, glycerol, yeast, agar, and seawater. To classify the types of microorganisms obtained from the various marine environments, taxonomic characterization was carried out by capillary GC analysis of fatty-acid methyl esters using the MIDI'" system. This chromatographic method determines the fatty-acid composition of unknown isolates and then searches an extensive database of compositions of known strains for accurate species identification. The MIDI'" analysis is based on a similarity index that expresses how closely the unknown organism compares to reference strains in our extensive marine database, which is continuously updated. Data from fatty-acid analyses revealed that Pseudomonas and Bacillus species predominated among both the warm- and cold-water isolates; however, the isolation of specific genera could be assigned to particular marine samples (Table 11). For example, Psychrobacter was only isolated in the North Pacific from invertebrate and core sediment samples, and Sphingornonas was only isolated in the South Pacific from vegetative marine samples. Thirteen percent of the isolates from Fiji and 16% of the isolates from Vancouver could not be identified by the MIDI'" system and were subsequently analyzed by a second identification system, BIOLOG'". BIOLOG'" is a bacterial identification system based on 95 biochemical tests, the results of which are used to search an extensive database for comparison to known metabolic patterns. This approach was also unsuccessful in taxonomically identifying these isolates. Using a dendrogram to analyze the fatty-acid profiles, 85% of these isolates were found to represent distinct genera. A further analysis as to the source of these unidentified microorganisms revealed that the majority of unknown cultures from Fiji and Vancouver were isolated from invertebrate samples; however, a significant number of cold water unidentified cultures originated from the water column (Table 111).This was in direct contrast to the results from the water column in Fiji, where all the cultures could be taxonomically identified. These data further demonstrate the vast differences in marine microbial populations in different parts of the

73

MICROORGANISMS AS SOURCE OF NEW NATURAL PRODUCTS TABLE I1 MARINE MICROORGANISMS AND SOURCES

Microorganism

Invertebrate

Aeromonas Agrobacteriurn Altermonos Arthrobacter Aureobacterium Cellulomonas Chryseomonas Clavibacter Cornamonas Corynebacterium Cytophago Enterobocter Erwinio Esch erichia Flavimon as Flavobacterium Gordon0 Hydrogen oph ago Klebsiello Kluyvera Methylobacterium Micrococcu s Ocean ospirillu m Ochroboctrum Paracoccus Phenylobacterium Psychrobacter Rhodococcus Sphingomonas Vibrio Xon thomonas

F F FV F FV F F F V FV V F F F F V F V F F FV FV F V V F V F F F F

Vegetative

Core

Water

F V V V

FV F V

F

Shoreline

F

F

FV V F

V

F

F F

V F

V

V

V

FV -

F = Fiji Isolates: V = Vancouver Isolates. FV = Fiji and Vancouver.

world. These results are encouraging and indicate that the marine environment is indeed a rich source of new and unique microorganisms; therefore, the likelihood is high that they will produce new bioactive compounds, B. FERMENTATION STUDIES

Since it has been well established that the assimilation of nutrients in a culture medium controls the production of secondary metabolites,

74

V. S. BERNAN et 01.

TABLE I11 FRZQIJENCY OF UNIDENTIFIED

MARINE

BACTERIA BY

SAMPLE ORIGIN

Sample type

Fiji

Vancouver

Core sediment Vegetative Invertebrate Water column

23% 20%

3% 35 %

57%

41 yo

0

21%

we have chosen media with very different nutritional compositions to evaluate the capacity of marine isolates to produce bioactive compounds. These media included formulations specific either for marine or terrestrial microorganisms. The marine media contained 3 % NaC1, glucose as a carbon source, and additional nutrients, such as Laminaria. The terrestrial media contained little or no NaCl and complex carbon and nitrogen sources, such as starch and NZ-Amine. Two terrestrial and two marine media producing the highest percentage of antibacterial activity were selected for routine fermentation studies. Fermentation studies of 342 isolates from the Fijian collections showed that the majority of cultures (70%) produced activity at both 3 and 5 days of growth. Since 17% of the cultures were active only after fermentation for 3 days, while slightly fewer organisms (13%) produced bioactivity only at 5 days, we chose a harvest period of 4 days to screen our marine isolates for the production of bioactive compounds. In addition to whole fermentation broths, concentrated extracts of these broths were tested to increase the probability of detecting bioactive metabolites. Centrifuged cell pellets were extracted with 80% methanol, while supernatants were processed over a C18 column and eluted with methanol. The distribution of activities among the various sample types is described in Table IV. Since the highest percentage of active cultures was found in the eluate and pellet fractions, all subsequent fermentations were processed in this manner.

C. BIOLOGICAL ACTIVITIES

To determine which cultures might be the most productive, all isolates from the Fijian collections were fermented and assayed for activity in our antitumor and antibacterial screening panels. Antibacterial activities were confirmed for 95 isolates (22%) from the Fijian collections

MICROORGANISMS AS SOURCE OF NEW NATURAL PRODUCTS

75

TABLE IV

BIOLOGICAL ACTIVITIES VERSUS SAMPLE PROCESS Sample process

Percentage of active cultures

Eluate only Eluate and pellet Effluent, eluate, and pellet Pellet only Eftluent only Pellet and effluent Effluent and eluate

39 30

17 7

4 2

1

(Table V). An analysis of these active cultures revealed that 48% were isolated from invertebrate tissues, 27% from core sediment, 15% from shoreline, and 10% from water column samples (Fig. 1).Although 66% of these isolates were Gram-negative rods, they accounted for only 36% of the active cultures (Fig. 2). As expected, a high frequency of antimicrobial activity was detected among the actinomycetes (29%) even though they represented only 13% of the total isolates. Bioactive compounds from several of these cultures have been identified. As expected, the compounds isolated represent a mixture of novel marine metabolites and known terrestrial products (Table VI). D. MARINEBACTERIA

Our interest in symbiotic marine microorganisms has led us to conduct microbial isolations from an orange ascidian, Polysyncruton lithostrotum, which when extracted with organic solvents yields potent cytotoxic DNA-damaging compound(s). Ten grams of orange ascidian were macerated and plated onto 8 different selective agar media for the isolation of microorganisms. Forty-one eubacteria (Table VII) and 18 actinomycetes were isolated and taxonomically analyzed by the MIDI'" system. All isolates were fermented in 8 different media and tested for DNA-damaging activity using the Biochemical Induction Assay (BIA) (Greenstein et al., 1993). Four of the isolates, three actinomycetes and one eubacterium, were active in the BIA. MIDI'" analysis of the halophilic eubacterium indicated it was an unidentified Gram-negative species, and the three halophilic actinomycetes were different Micro-

TABLE V NUMBER OF lMARINE MICROORGANISMS ACTIVE IN Vmous ASSAY SYSTEMS Assay systems

Microorganism type

CWlO

cw2

cw3

cw4

cw5

Gm+

Gm+ Gm-

Gm+

Gm+

23

1

3

2

9

2 0 0 0 0 1

1 1

0 0 1 0 0 3

CW6

Gm+b

Antitumor

< v

m Gram- rods Gram+ rods Gram variable Gram+ budding Gram- cocci Gram+ sheathed Actinomycetes

4

2 2 0 11

"CW1-6 = actives in different cell wall assays. bGm+= Gram-positive. cGm- = Gram-negative.

0 0 1

0

4

0 0

0 1 0 0 0

0 1

0 1

0 0 1

1 1 0

2 0 1

10 3 1

El

9

z

0

B

0 0

!-

4

0

MICROORGANISMS AS SOURCE OF NEW NATURAL PRODUCTS

77

27%

10%

48 %

15%

FIG.1. Distribution of antibacterial activity by microorganism source.

Gram Negative Rods EActinomycetes

Gram Positive Rods 0 Gram Variable

.1

Gram Positive Sheathedl

36%

FIG.2. Distribution of antibacterial activity by microorganism type.

7%

V. S. RERNAN et ol TABLE VI

BIOACI‘IVECOMPOUNDS IDENTIFIED FROM W-AR MARINE h‘flCRCX1RGANISM SCREENING PROGRAM

Organism

Identification

Pseudoinonas aeruginosa Bacillus purnilus Bacillus rnaceraris Bacillus subtilis Bacillus subtilis Bacillus rnacerans Bacillus rnacerans Streptornyces viridodiastaticus “litornlis” Agrobacterium sp. Actinomycete sp. Ifypoxylon oceanicurn Actinomycete sp. LL-141352

Tubermycin Bacilysin Bacilysin Bacilysin TL119 Lipopeptides Bacillomycin D Bioxalomycin (Novel) Thiotropocin Chartreusin LIr15G25601 Antimycin LL-l41352a,p (Novel)

TABLE VII TY PBS OF O R G A N E M S ISOLATED FROM

Organism type Fungi Mycobocterium Oceanospirillum Bacillus Pseudonionas

Agrobacterium Rhodococcus Micromonosporo Unidentified

Total no. of isolates

POLYSYNCRATON LITHVSTROTUM

Number of unique isolates

2

2 genera

2

1 species

5 4 3 5 1

2 species 3 species 2 species 1 species 1 species 3 species

16 24

6 genera; 10 species

monospora species. In addition, another morphological type of bacterium produced a potent compound with activity against Gram-negative and -positive bacteria. The microbiology of the eubacteria and the chemistry of their products are described later. Studies of the other microorganisms are ongoing.

MICROORGANISMS AS SOURCE OF NEW NATURAL PRODUCTS

FIG.3. Structures of LL-141352~~ and

I . Antibiotics 141352a and

79

p produced by strain LL-141352.

p

Strain LL-141352 was isolated from a tissue sample of the ascidian Polysyncruton lithostroturn collected near Namenalala Island in Fiji. Strain LL-141352 is a halophilic, nonmotile, pleomorphic, Gram-negative eubacterium. This strain grows as a shiny, flat, beige colony with an olive-green center. In resting stage, the culture consists predominantly of thin-walled coccoid bodies. When transferred into a nutrientrich liquid medium, the strain undergoes a morphologically complex life cycle. Large-scale fermentations were carried out to provide material for isolation and structure elucidation. The antibacterial and DNA-damaging activities were found in the supernatant after centrifugation of the fermentations. Preparative HPLC yielded two BIA-active components, LL-141352~and p (Fig. 3). LL-141352a contains a phenazine moiety linked to an uncommon P-hydroxy valine residue through an ester bond. The structure of the p component was established based on its molecular weight and the fact that hydrolysis of a yields p. Compound LL-141352a exhibited antitumor activity with a mean ICs0 of 0.48 pg/ml in cell culture-based cytotoxicity assays and showed excellent Gram-positive activity against S. aureus (MIC = 0.12-0.25 pg/ml) and moderate activity against Gram-negative microorganisms. 141352p (lacking the amino-acid residue) is much less active than a (-40-fold less against S . aureus), with a mean IC,, of 1.8 pg/ml in cell-based cytotoxicity assays. Mechanistic studies in bacteria suggested DNA synthesis to be the primary macromolecular target. In animal experiments, both LL-141352a and p were inactive in a murine P388 leukemia model and in an S. aureus infection model at doses up to 8 mg/kg. This lack of in vivo activity may be due to the fact that LL141352a is chemically unstable and converts to the less active p.

80

V. S. BERNAN et al. TABLE VIII CULTURAI. CHAKACTERISTICS OF STRAIN LL-211457 ~

Result

Characteristic Gram stain

Negative

Motility

Positive

Cell morphology

Rods i n star-shaped aggregates

Colony morphology

Brown, smooth mucoid

Salt tolerance

25%

Anaerobic growth

Negative

Whole-cell sugars

Ribose, mannose, glucose

Ubiquinones

Q10

Fatty acids

C18:1 and 3-OH-C14:0

2. Antibiotic LL-21I457

Another taxonomically distinct marine eubacterium, LL-211457, was isolated from the middle section of the tunicate Polysyncraton lithostrotum. Fermentation broths of this strain exhibited activity against antibiotic-resistant Gram-positive isolates. Taxonomic identification revealed that this strain was a halophilic Agrobacterium species. This finding was unusual since there are very few marine Agrobacterium species reported, and these have been isolated from the brackish waters of the Baltic Sea and the northeastern Atlantic Ocean (Riiger and Hofle, 1992). Table VIII describes the cultural characteristics of LL-211457. The production of both the a and components, monitored every 8 h by HPLC assay, peaked at 25 h (Fig. 4). The active component was identified as thiotropocin (Fig. 5), which had previously been isolated from a terrestrial Pseudomonas strain (Kintaka et al., 1984). Thiotropocin was the first example of a naturally occurring tropothione derivative and has a unique structure containing an S-0-CO moiety. 3.

Antibiotic U-31F508

A more in-depth evaluation was conducted on a marine actinomycete designated LL-31F508, isolated from an intertidal sample collected in Key West, Florida. Interest in this microorganism emanated from its activity in our pharmacological screening programs for cancer chemo-

81

MICROORGANISMS AS SOURCE OF NEW NATURAL PRODUCTS

.--

__-

fl

Pcv

---a- Alpha --t-

Gamma

10

CI

6

I

s

Y

>

P

“ 2 0

n

2

a

*O?

E

W

E

.-

Q)

I

Fermentation (hours) FIG.4. Time course of growth and antibiotic production for strain LL-211457. Thiotropocin (mg/liter) was determined by HPLC. Packed cell volume (PCV) was determined for a 10-ml sample. Production of thiotropocin peaked approximately 25 h after inoculation.

therapy. This isoIate was taxonomically identified and named Streptomyces viridodiastaticus “litoralis.” Fermentation samples exhibited both antibacterial and DNA-damaging activities. A new series of compounds, the bioxalomycins, were identified from this culture. Largescale fermentations were carried out to provide material for isolation and structure elucidation. Production of the bioxalomycins peaked approximately 36 h after inoculation (Fig. 6). Medium optimization studies further enhanced the production of the bioxalomycins. Addition

82

V. S. BERNAN et al.

THlOTRO P 0 C IN 211457y FIG.5. Structure of thiotropocin produced by strain LL-211457.

of 2% NaCl to medium A-1 increased biomass by 33% and activity fourfold. This observation is not surprising since this culture was isolated from a marine environment and NaCl may be required for optimum growth and production of the bioxalomycins (Table IX). The purification of the novel bioxalomycins was accomplished utilizing preparative reverse phase HPLC. The structures were elucidated using a variety of NMR techniques and mass spectral data (Fig. 7). The p species were found to be the quinone forms of the corresponding a components. The p components are distinguished from the antibiotic naphthyridinomycin by the presence of a second oxazolidine ring in a region of the molecule analogous to quinocarcin. Bioxalomycin showed MIC values ranging from <0.015 to 0.5 pg/ml against Gram-positive bacteria and from 0.12 to 2.0 pg/ml against most Gram-negative bacteria. An evaluation of bioxalomycin a against S. aureus infections in mice showed protection at one-tenth the maximum tolerated dose. Mechanistic studies were performed using exponentialphase cells of E. coli to determine the effect of bioxalomycin a on the incorporation of radiolabeled precursors into DNA, RNA, and protein. After ten minutes, bioxalomycin a inhibited DNA synthesis by 94%, while RNA and protein synthesis were inhibited by only 33% and 30%, respectively (Bernan et al., 1994; Singh et al., 1994; Zaccardi et a]., 1994).

Bioxalomycin a2 also demonstrated excellent in vitro activity against a tumor cell panel. Mechanistic studies determined that it inhibited RNA synthesis preferentially in HeLa cells. Because of bioxalomycin a2

83

MICROORGANISMS AS SOURCE OF NEW NATURAL PRODUCTS

10 Glucose & Bioxalomycin

10

1

.1 0

10

20

30

40

50

Hours FIG.6. Production of bioxalomycin by Streptomyces viridodiastaticus "litoralis" LL31F508. Fermentations were carried out in 300-liter tanks at 28OC. Bioxalomycin CL (mg/liter) was determined by HPLC. Packed cell volume (PCV) was determined for a 10-ml sample. Dextrin and glucose are reported in mg/ml. Production of the bioxalomycins peaked approximately 36 h after inoculation during a defined idiophase after cessation of exponential growth. TABLE IX EFFECTSOF SALTCONCENTRATION UPON BIOACTIVITY PRODUCED BY STRAIN LL-31F508

Fermentation medium"

Relative activity b

Packed cell volume (%)

A-1 A-l + 1 % NaCl A-1 + 2% NaCl A-1 + 3% NaCl

1:8 1:32

20

1:64 1:32

30

30 30

"Organisms were grown in 50 ml of medium in 250-ml flasks for 5 days at 28"C, 200 rpm. A-1 medium contained 1%dextrose, 0.5% NZ-amine A, 0.1% calcium carbonate, 2% soluble starch, 0.5% yeast extract, and 0.04% agar. bSerialtwofold dilutions of fermentation samples were tested.

84

V. S. BERNAN et

QI.

p l R=H p2 R=CH3

FIG.7 . Structure of bioxalomycins

c t l , ~ 2 Pl, , and p2.

activity in our in vifro anticancer screening panel, it was evaluated in vivo in a mouse P388 leukemia model (Fig. 8). This compound exhibited an increase in life survival (ILS) of 80% in this model.

E. MAIUNE FUNGI Marine fungi are also evaluated in our screening programs as a source of novel bioactive metabolites. In screening fermentation samples for inhibitors of fungal cell wall biosynthesis, culture LL-15G256, a strain of Hypoxylon oceanicum, produced a positive response. The optimum temperature range was 22-28"C, and salinity was not an important variable in the fermentation. Fermentations of this culture were readily processed by extraction with ethyl acetate to isolate the active compounds. Chromatography of the organic extract by preparative HPLC produced one major compound referred to as LL-l5G256a, which was associated with the biological activity. LL-15G256a was found to be identical with BK223-A and NG-012, which was evident from comparisons of UV spectra, 13C-NMR data, and molecular weight (Fig. 9) (Breinholt et a]., 1993; Ito ef al., 1992a,b). Although these compounds are known, their production by a

MICROORGANISMS AS SOURCE OF NEW NATURAL PRODUCTS

85

80 60

40

20

0 -20 -4 0 -60 0

0.2

0.4

0.6

0.8

1

BIOXALOMY C IN (mg/kg) FIG.8. Activity of bioxalomycin a2 against P388 leukemia in mice. On Days 1,5, and

(lo6 per animal) were implanted intraperitoneally into BDFl mice, bioxalomycin a2 was administered at various doses by the same route. At 0.6 mg/kg, an 80% increase in median life span, well above the 25% cutoff for activity in this test system, was observed. 9 after P388 murine leukemia cells

OH

HO

OH

0

15G256a Hypo xylo n ocean icu m

A) JeBK223n icill iu m v errucu lo su m ( N E 01 2 ) Pen icillium verruculosum F-4542 FIG.9. Structure of antibiotic LL-15G256a produced by Hypoxylon oceonicum.

86

V. S. BERNAN et al.

marine fungus and their mechanism of action have not been previously reported (Schlingmann, 1994). IV. Marine Biotechnology

It is estimated that less than 5% of the bacteria observed by microscopy are culturable under standard laboratory conditions. This fact has greatly limited our ability to isolate highly unusual marine microorganisms. This is particularly true for many symbionts such as the marine bacteria found in the deep-sea hydrothermal vents or those found in extreme environments. Due to the great potential for isolating new natural products from these bacteria, biotechnology companies and other investigators are now concentrating on recombinant methods to clone the genetic information involved in the production of targeted metabolites into common industrial microbes. One company, ChromaXome, utilizes a unique application of genetic engineering called “combinatorial biology,” a strategy that randomly mixes and matches multiple genes from more than one species to produce novel metabolites. This new technology has been enabled by the development of very large vectors for transferring foreign DNA into bacteria. ChromaXome uses cosmids to transfer up to 30 to 40 kb of DNA and newly developed bacterial artificial chromosomes to transfer u p to 300 to 400 kb of DNA. This permits the genetic information for 30 to 400 enzymes to be transferred at one time into a commercial vector. The DNA is then transcribed using synthetic promoters to create a novel biosynthetic pathway. By transferring genes from marine microorganisms into one clone, this technique could yield biologically active compounds never before observed in nature. Another biotechnology company, Recombinant BioCatalysis’“Inc., is concentrating on creating new generations of uniquely stable biocatalysts by directly cloning genes from organisms found in extreme environments, including marine niches. These extremophiles are isolated from terrestrial hot springs at temperatures above 100°C and in deep-sea thermal vents at temperatures below 0°C in Arctic waters. Others are present beneath several miles of water under immense pressure or in the saturated salt environment of the Dead Sea. One of the more unique collection sites is whale falls, which are dead whales that have sunk to the bottom of the ocean and have decayed over a period of 10-15 years. During this process, they provide a home to microbial mats that contain unique microflora. By using these extremophiles as a resource, they have been able to discover new enzymes with exceptional stability.

MICROORGANISMS AS SOURCE OF NEW NATURAL PRODUCTS

87

Using these unique microbial sources, Recombinant BioCatalysis'" Inc. has discovered more than 150 new enzymes called CloneZymes'". Based on functional groupings, these CloneZymes'" are organized into tiers that can be screened for properties appropriate for a particular process. Currently, four CloneZyme'" libraries are available and consist of aminotransferases, esterases/lipases, glycosidases, and phosphatases. Marine bacteria have been recognized as a new source of therapeutic leads and have been shown to be capable of producing a large number of diverse biochemicals. However, out of the large number of species estimated, only a fraction of the marine bacteria have been isolated and cultured. The future of this technology is promising as a way of accessing new natural products from the marine environment that would not be obtainable by traditional methods. V. Summary

Over the past decade, marine microorganisms have become recognized as an important and untapped resource for novel bioactive compounds. The oceans cover greater than 70% of the earth's surface and, taking this into account by volume, represent better than 95% of the biosphere. Given this fact, the oceans present themselves as an unexplored area of opportunity for the discovery of pharmacologically active compounds. In this review, data have been presented to illustrate the diversity of microorganisms living in the sea and the plethora of chemical compounds that have been discovered from them. However, it is important to pursue basic research on the marine environment in order to permit the continued isolation of unique microorganisms. There is still limited knowledge of the physiological requirements of most marine microorganisms, and a greater understanding of their conditions for growth will offer new insights into the complex world of marine microbiology. Clearly, a greater investment in the development of marine biotechnology will produce novel compounds that may contribute significantly toward drug development over the next decade. REFERENCES Andersen, R. J., Wolfe, S. M., and Faulkner, D. J. (1974). Mar. B i d . 24, 281-285. Baz, J. P., Millan, F. R., De Quesada, T. G., and Gravalos, D. G. (1995). International Patent WO 95127730. Bernan, V. B., Montenegro, D. A,, Korshalla, J. D., Maiese, W. M., Steinberg, D. A., and Greenstein, M. (1994). J. Antibiot. 47, 1417-1424. Breinholt, J., Jensen, G. W., Nielsen, R. I., Olsen, C. E., and Frisvad, J, C. (1993). J. Antibiot. 46(7), 1101-1108.

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Bull, A. T., Goodfellow, M., and Slater, J. H. (1992). Ann. Rev. Microbiol. 46,219-252. Burkholder, P. R., Pfister, R. M., and Leitz, F. P. (1966). App. Microbiol. 14, 649-653. Cheng, X. C., Varoglu, M., Abrell, L., Crews, P., Lohkovsky, E., and Clardy, J. (1994). J. Org. Chem. 59,6344-6348. Colwell, R. R. (1993). In “World Bank Discussion Papers” (R. A. Zilinskas and C. G. Lundin, eds.), No. 210, pp. IX-X. World Bank Press, Washington, DC. Cui, C., Uhukata, M., Kakeya, H., Onose, R., Okada, G., Takahashi, I., Isono, K., and Osada, H. (1996). J. Antibiot. 49,216-219. Davidson, B. S., and Schumacher, R. W. (1993). Tetrahedron Lett. 49,6569-6574. Distel, D. L., Lane, D. J., Olsen, G. J., Giovannoni, S. J. Pace, B., Pace, N. R., Stahl, D. A., and Felbeck, H. (1988). J. Bacteriol. 170,2506-2510. Elyakov, G.B., Kuznetsova, T., Mikhailov, V. V., Mal’tsevi, I., Voiniv, V. G., and Fedoreyev, S. A. (1991). Expedentia 47,632-633. Faulkner, D.J. (1995). Natural Products Reports 12,223-269. Fenical, W. (1993). Chem. Rev. 93, 1673-1683. Fenical, W., and Jensen, P. R. (1993). In “Marine Biotechnology” (D. H. Attaway and 0. R. Zahorsky, eds.), Vol. 1, pp 419-458. Pharmaceutical and Bioactive Natural Products, Plenum, New York. Gerard, J.. Haden, P., Kelly, M. Y., and Andersen, R. J. (1996). Tetrahedron Lett. In press. Gil-Turnes, M. S., and Fenical, W. (1992). B i d . Bull. 182,105-108. Gil-Turnes, M. S., Hay, M. E., and Fenical, W. (1989). Science 246,117-118. Greenstein, M.,Wildey, M. J., and Maiese, W. M. (1993). In “Enediyne Antibiotics as Antitumor Agents” (D. B. Borders and T. W. Doyle, eds.), pp 17-27. Dekker, New York. Guerriero, A., D’Ambrosio, M., Cuomo, V., Vanzanella, F., and Pietra, F. (1988). Helv. Chim. Acta. 71, 57-61. Guerriero, A., D’Ambrosio, M., Cuomo, V., Vanzanella, F., and Pietra, F. (1989). Helv. Chim. Acta. 72, 57-61. Holland, G. S., Jamieson, D. D., Reicheldt, J. L., Viset, G., and Wells, R. J. (1984). Chemical Industry, 3 December. Hotta, K., Yoshida, M., Hamada, M., and Okami, Y. (1980). J. Antibiot. 33, 1515-1520. Imada, C., Hara, S. Maeda, M., and Shimidu, U. (1986). Bull. Jpn. Soc. Sci. Fish 52, 1455-1459. Ito, M., Maruhashi, M., Sakai, N., Mizoue, K., and Hanada, K. (1992a). J. Antibiot. 45(10), 1559-1565. Ito, M., Maruhashi, M., Sakai, N., Mizoue, K., and Hanada, K. (1992h). J. Antibiot. 45(10), 1566-1572. Iwai, Y.,and Takahashi, Y. (1992). In “The Search for Bioactive Compounds from Microorganisms” (S. Omura, ed.), pp. 281-302. Springer, New York. Jalal, M. A. F, Hossain, M. B., van der Helm, D., Sanders-Loehr, J., Actis, L. A., and Crosa, J. H. (1989).J. Am. Chem. Soc. 111,292-296. Jensen, P. R., Dwight, R., and Fenical, W. (1991). Appl. Environ. Microbiol. 57,1102-1108. Jensen, P. R., Kauffman, C. A., and Fenical, W. (1996). Mar. Biol. In press. Kameyama, T., Takahashi, A., Kurasawa, S., Ishizuka, M., and Okami, Y. (1987). J. Antibiot. 40,1664-1670. Kintaka, K., Ono, H., Tsuhotani, S., Harada, S., and Okazaki, H. (1984). J. Antibiot. 11, 1294-1300. Kodama, M., Ogata, T., and Sato, S. (1988). Agric. Biol. Chem. 52,1075-1077. Kodama, M., Ogata, T., Sakamoto, S., Sato, S., Honda, T., and Miwatani, T. (1990). Toxicon. 28. 707-714.

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Kosuge, T., Tsuji, K., Harai, K., and Fukuyama, T. (1985). Chem. Phar. Bull. 33, 30593061. Kupka, J., Anke, T., Steglich, W., and Zechlin, L. (1981). J. Antibiot. 34,298-304. Lindel, T., Jensen, P. R., and Fenical, W. (1996). Tetrahedron Lett. 37,1327-1329. Needham, J., Andersen, R. J., and Kelly, M. T. (1991). Tetrahedron Lett. 32, 315-318. Needham, J., Kelly, M. T., Ishige, M., and Andersen, R. J. (1994). J. Org. Chem. 59, 2058-2063. Noguchi, T., Jeon, J.-K., Arakawa, O., Sugita, H., and Deguchi, Y. (1986). J. Biochem. 99, 311-314. Numata, A,, Takahashi, C., Matsushita, T., Miyamoto, T., Kawai, K., Usami, T., Matsumum, E., Inoue, M., Ohishi, H., and Shingu, T. (1991). Terahedron Lett. 33, 16211624. Oclarit, J.. Okada, H., Ohta, S., Kaminura, K., Yamaoka, Y., Lizuka, T., Miyashiro, S., and Ikegami, S. (1994). Microbios 78,7-16. Okami, Y. (1993). J. Mar. Biotech. 1, 59-65. Okami, Y., Okazaki, T., Kitahara, T., and Umezawa, H. (1976). J. Antibiot. 29,1019-1025. Okarni, Y., Kurasawa, S., and Hirose, Y. (1980). Agric. Bid. Chem. 44, 1191-1192. Okutani, K. (1977). Bull. Jpn. SOC.Sci. Fish 43,995. Pathirana, C., Tapioloa, D., Jensen, P. R., Dwight, R., and Fenical, W. (1991). Tetrahedron Lett. 32,2323-2326. Pathirana, C., Jensen, P. R., Dwight, R., and Fenical, W. (1992).J. Org. Chem, 57,740-742. Pathirana, C., Jensen, P. R., and Fenical, W. (1993). Tetrahedron Lett. 33, 7663-7666. Poch, G. K., and Gloer, J. B. (198%). Tetrahedron Lett. 30, 3483-3486. Poch, G. K., and Gloer, J. B. (1989b). J. Nat. Prod. 52,257-260. Poch, G. K., and Gloer, J. B. (1991). J. Nat. Prod. 54, 213-217. Poch, G. K., Gloer, J. B., and Shearer, C. A. (1991).J. Nat. Prod. 55,1093. Sato, K., Okazaki, T., Maeda, K., and Okami, Y. (1978). J. Antibiot. 31,632-635. Riiger, H.-J., and Hofle (1992). Int. J. Sys. Bact. 42,133-143. Schlingmann, G. (1994). Int. Conf. Chem. Synth. Antibiotics and Related Microbial Products, 4th, 11-16 September, Nashville, TN. Shigemori, H., Wakuri, S., Yazawa, K., Nakamura, T., Sasaki, T., and Kobayashi, J. (1991). Tetrahedron Lett. 47,8529-8534. Shigemori, H., Bae, M., Yazawa, K., Sasaki, T., and Kobayashi, J. (1992).J. Org. Chem. 57, 43174320. Shin, J., and Fenical, W. (1987). Phytochemisty 26,3347. Singh, M. P., Petersen, P. J., Jacobus, N. V., Maiese, W. M., and Greenstein, M. (1994). Antimicrob. Agents Chemother. 38,1808-1 812. Stierle, A. C., and Cardellina, J. H., I1 (1988). Tedania ignis. Experientia 44, 1021. Sugano, M., Sato, A., Iijima, Y., Oshirna, T., Furuya, K., Kuwano, H., Hata, T., and Hanzawa, H. (1991). J. Am. Chem. SOC.113, 5463-5464. Strongman, D. B., Miller, J. D., Calhoun, L., Findlay, J. A., and Whitney, N. J. (1987). Bot. Mar. 30, 21-26. Takahashi, A., Nakamura, H., Kameyama, T., Kurasawa, S., and Naganawa, H. (1987). J. Antibiot. 40,1671-1676. Takahashi, A., Ikeda, D., Nakamura, H., Naganawa, H., and Kurasawa, S . (1989a). J. Antjibjot. 42,1562-1566. Takahashi, A., Kurosawa, S . , Ikeda, D., Okami, Y., and Takeuchi, T. (1989b).J. Antibiot. 42,1556-1561. Takahashi, C., Takada, T., Yamada, T., Minoura, K., Uchida, K., Matsumura, E., and Numata, A. (1994). Tetrahedron Lett. 35,5013-5014.

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Stereoselective Biotransformations in Synthesis of Some Pharmaceutical Intermediates RAMESHN. PATEL Deparimeni of Microbial Technology Bristol-Myers Squibb Pharmaceutical Research Institute New Brunswick, New Jersey 08903

I. 11. 111. IV. V. VI. VII. VIII. IX. X. XI. XII. XIII.

Introduction Taxol Semisynthesis Thromhoxane A2 Antagonist ACE Inhibitors Anticholesterol Drugs Antiinfective Drugs Calcium Channel Blocking Drugs Antipsychotic Agents Antiarrhythmic Agents Potassium Channel Openers Antiinflammatory Drugs Antiviral Agents Prostaglandin Synthesis References

I. Introduction

Much attention is being focused on the interaction of small molecules with biological macromolecules. The search for selective enzyme inhibitors and receptor agonists or antagonists is one of the keys for target-oriented research in the pharmaceutical industry. Increasing understanding of the mechanism of drug interaction on a molecular level has led to an increasing awareness of the importance of chirality as the key to the efficacy of many drug products. It is now known that in many cases only one stereoisomer of a drug substance is required for efficacy and the other stereoisomer is either inactive or exhibits considerably reduced activity. Pharmaceutical companies are aware that, where appropriate, new drugs for the clinic should be homochiral to avoid the possibility of unnecessary side effects due to an undesirable stereoisomer. In many cases where the switch from racemate drug substance to enantiomerically pure compound is feasible, there is the opportunity to double the use of an industrial process. The physical characteristic of enantiomers versus racemates may confer processing or formulation advantages. 91 ADVANCES IN APPLIF,D MICROBIOLOGY.VOLUME 43 Copyright 0 7997 by Academic: Prcss. Inc. All rights of rcproductiou in any form reaerved. 0065-2164/97 $25.00

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R. N. PATEI,

Chiral drug intermediates can be prepared by different routes. One is to obtain them from naturally occurring chiral synthons mainly produced by fermentation processes. The chiral pool primarily refers to inexpensive, readily available optically active natural products. Second is to carry out resolution of racemic compounds. This can be achieved by preferential crystallization of stereoisomers or diastereoisomers and by kinetic resolution of racemic compounds by chemical or biocatalytic methods. Finally, chiral synthons can also be prepared by asymmetric synthesis by either chemical or biocatalytic processes using microbial cells or enzymes derived therefrom. The advantages of microbial or enzyme-catalyzed reactions over chemical reactions are that they are stereoselective and that they can be carried out at ambient temperature and atmospheric pressure. This minimizes problems of isomerization, racemization, epimerization, and rearrangement that generally occur during chemical processes. Biocatalytic processes are generally carried out in aqueous solution. This avoids the use of environmentally harmful chemicals used in the chemical processes and solvent waste disposal. Furthermore, microbial cells or enzymes derived therefrom can be immobilized and reused during many cycles. A number of review articles (Sih and Chen, 1984a; Jones, 1986; Crout et al., 1994; Davies et al., 1990; Csuk and Glanzer, 1991; Crosby, 1991; Kamphuis et al., 1990a; Sih et al., 1992; Stinson, 1992; Santaniello, 1992; Margolin, 1993; Cole, 1994; Wong and Whitesides, 1994; Mori, 1995; Feng et al., 1995) have been published on the use of enzymes in organic synthesis. This report provides some specific examples of stereoselective biotransformations to prepare chiral intermediates required for the synthesis of pharmaceutical drugs. II. Taxol Semisynthesis

Among the antimitotic agents, taxol (paclitaxel) 1 (Wani et al., 1971; Kingston, 1991), a complex polycyclic diterpene, exhibits a unique mode of action on microtubule proteins responsible for the formation of the spindle during cell division. In contrast to such other “spindle formation inhibitors” as vinblastine or colchicine, both of which prevent the assembly of tubuline, taxol is the only compound known to inhibit the depolymerization process of microtubulin (Schiff et a]., 1979). Because of its biological activity and unusual chemical structure, taxol represents the prototype of a new series of chemotherapeutic agents. Various types of cancers have been treated with taxol, and the results in treatment of ovarian cancer are very promising. In collabora-

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

93

tion with the National Cancer Institute, Bristol-Myers Squibb developed taxol for treatment of refractory ovarian cancer. Taxol was originally isolated from the bark of the yew Taxus brevifolia (Wani et al. 1971) and has also been found in other Taxus species in relatively low yield. Taxol was initially obtained from T brevifolia bark in about 0.07% yield. It required cumbersome purification of taxol from the other related taxanes. It is estimated that about 20,000 pounds of yew bark (equivalent to about 3000 trees) are needed to produce 1 kg of purified taxol (Kingston, 1991). This created a concern among the environmentalists about mass destruction of trees to produce the required amount of taxol. Alternative methods for production of taxol by cell suspension cultures and by semisynthetic processes are being evaluated by various groups (Holton et al., 1988; Denis et al., 1986; Christen et al., 1991). The development of a semisynthetic process for the production of taxol from baccatin I11 3 or 10-deacetylbaccatin 111 (10-DAB)3 and C-13 taxol side chain was a very promising approach. Taxol, related taxanes, baccatin 111, and 10-DAB can be derived from such renewable resources as extract of needles and shoots and young Taxus cultivars. By using selective enrichment techniques, Hanson et al. (1994a) isolated two strains of Nocardioides that contained novel enzymes C-13 taxolase and C-10 deacetylase. The extracellular C-13 taxolase derived from filtrate of the fermentation broth of Nocardioides albus SC-13911 catalyzed cleavage of C-13 side chain from taxol and such related taxanes as taxol C, cephalomannine, 7-P-xylosyltaxol, 7-P-xylosyl-10deacetyltaxol, and 10-deacetyltaxol (Fig. 1). The intracellular C-10 deacetylase derived from fermentation of Nocardioides luteus SC-13912 catalyzed the cleavage of C-10 acetate from taxol, related taxanes, and baccatin 111 to yield 10-DAB (Fig. 2). Fermentation processes were developed for growth of N. a l b SC-13911 ~ and N. luteus SC-13912 to produce C-13 taxolase and C-10 deactylase, respectively, in 5000-liter batches (Nanduri et al., 1995). A bioconversion process was demonstrated for the conversion of taxol and related taxanes in extracts of Taxus plant cultivars to single-compound 10-DAB using both enzymes (Figs. 1 and 2). In the bioconversion process, ethanolic extracts of the whole young plant of five different cultivars of Taxus were first treated with a crude preparation of the C-13 taxolase to give complete conversion of measured taxanes to baccatin I11 and 10-DAB within 6 h. Nocardioides luteus SC-13192 whole cells were then added to the reaction mixture to give complete conversion of baccatin I11 to 10-DAB. The concentration of 10-DAB was increased by 5.5- to 24-fold in the extracts

94

R. N. PATEL 0

0

+

Taxanea

Iarpl

ROH

2 EssaUm

mQlG

-

FIG.1. Hydrolysis of the C-13 side chain of taxanes by C-13 taxolase from Nocardioides albus SC13911 (Hanson et d., 1994a). Reprinted with permission from the American Society for Biochemistry arid Molecular Biology.

treated with the two enzymes. The bioconversion process was also applied to extracts of the bark of 7: bravifolia to give a 12-fold increase in 10-DAB concentration (Table I). The enhancement of 10-DAB concentration in yew extracts was useful in increasing the amount and purification of this key precursor for the taxol semisynthetic process using renewable resources. Another key precursor for the taxol semisynthetic process is the preparation of chiral C-13 taxol side chain &. Two different stereoselective enzymatic processes were developed for the preparation of chi1993a, 1994a). In one ral c-13 taxol side-chain synthon (Pate1 et d., process, the stereoselective microbial reduction of Z-keto-3-(N-benzoylamino)+phenyl propionic acid ethyl ester 5 to yield (2R,SS)-(-]-N-benzoyl-3-phenyl isoserine ethyl ester was demonstrated. The reduction of compound 5 could result in the formation of four possible alcohol diastereomers (2a-2d) (Fig. 3). Remarkably, conditions were found under which predominantly only the single (2R,3S) isomer was obtained by the biotransformation. After an extensive microbial screen, two strains of Hansenula were identified that catalyzed the stereoselective reduction of ketone 5 to the desired product @ in >800/0 reaction yield and >94% optical purity. Preparative-scale bioreduction of ketone 5 was

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

IiaxQl

95

JhxQLc

FIG.2. Hydrolysis of the C-10 acetate of taxanes by C-10 deacetylase from Nocordioides luteus SC-13912 (Hanson et d., 1994a). Reprinted with permission from the American Society for Biochemistry and Molecular Biology.

demonstrated using 20% cell suspensions of Hansenula polymorpha SC-13865 and Hansenula fabianii SC-13894 in independent experiments. In both batches, a reaction yield of >8O% and an optical purity of >94% were obtained for the desired alcohol isomer &. A 20% yield of undesired antidiastereomers (4c, 4d) was obtained with H. polymorpha SC-13865 compared to a 10% yield with H. fabianii sc-13894.A 99% optical purity of desired alcohol isomer & was obtained with H. polymorpha SC-13865 compared to a 94% optical purity with H. fabianii SC-13894. From one batch, 5.2 g of compound & was isolated in a 65% overall yield. The isolated compound gave 99.6% optical purity and 99% chemical purity. A single-stage fermentation-bioreduction process was also developed for the preparation of c-13 taxol side chain 4a. Cells of H. fabianii were grown in a 15-liter fermenter for 48 h, the bioreduction process was then initiated by addition of 30 g of substrate and 250 g of glucose and continued for 72 h. A reaction yield of 88% and an optical purity of 95% were obtained for the desired product &. In an alternate process for the preparation of C-13 taxol side chain, the stereoselective enzymatic hydrolysis of racemic cis-3-(acetyloxy)-4phenyl-2-azetidinone 6 to the corresponding (S)-(-)-alcohol 2 was carried out (Pate1 ef al., 1994a). Lipase PS-30 from Pseudomonas cepacia

TABLE I

co

BIOCONVERSION OF TAXUS PLANT EXTRACTS BY TAXOLASES~

Source (extract used)

Reaction time (hours)

Reaction volume (mll

T. adams (120 ml)

0 6 22 0 6 22 0 6 22 0 6 22 0 6 22 0 6 22

876

T. runyani (200 ml)

T. dark green spreader (150 ml) T. hicksii (120 ml) T. caspidata (150 ml) T. brevifolia (150 ml)

Cephalomannine (mgl 6.1

10-desaacetyl tax01 (mgl

0

17.5 0 0

1460

4.4 0 0

10.2 0 0

1170

9.3

8.2

0

0 0

0

0 936

1170

16.3 0 0 11.2 0 0

1220

15.8 0 0

15.9 0

0 10.5 0 0 31.7 0 0

Tax01 (mgl

Baccatin 111

lo-desacetyl baccatin 111

Increase in

(mgl

(mp)

X

10-DAB

14.0 0 0 13.2 0 0

7.0 27.1 0.0 2.3 21.9 0

6.1 20.1 41.1 1.4 13.1 33.8

12.8 0 0

3.51 28.1 0

3.5 44.4

12.6

23.4 0 0 10.6 0 0

2.8 42.1 2.4

12.2 27.1 66.4

5.5

8.2

12.2 0 0

8.5 82.9

5.8 18.7 41.0 10.8 41.4 128.0

19.2 0

0

6.7

24.0

7 ?:

8.2

7.0

11.8

qeaction mixture contained 25 mMphosphate buffer (pH 7), designated amount of plant extract (see source), and 10 units of C-13 taxolase (batch TEZ0693, 40% ammonium sulfate ppt, and lyophilized enzyme). After 6 h reaction period at 25°C with 150 rpm agitation, C-10 acetylase enzyme (10units, frozen cells, batch T10Z0893) was added and reaction was continued overnight (16 h). HPLC assay was used to determine taxane content of reaction mixture.

;I?3

SYNTHESIS OF SOME PHARMACEIJTICAL INTERMEDIATES

Microbial Reduction C0,El

~

-

H.DbhrmorahaSC 13865 H.f&kQUSC13894

97

miH 6H

5

wo iH

COIEt

FIG.3. Synthesis of taxol side-chain synthon. Stereoselective reduction of Z-keto+(Nbenzoylamino)-3-phenyl propionic acid ethyl ester 5 (Pate1 et al., 1993a). Reprinted with permission from Elsevier Science Ltd.

(Amano International Enzyme Company) and BMS lipase (extracellular lipase derived from the fermentation of Pseudomonas sp. sc-13856) catalyzed hydrolysis of the undesired enantiomer of racemic 5, producing S-(-)-alcohol 7 and the desired R-(+)-acetate 8 (Fig. 4). Reaction yields of >96% and optical purities of >99.5% were obtained. For a very efficient enzyme source (BMS lipase), a lipase fermentation using Pseudomonas sp. sc-13865 was developed. In a fed-batch process using soybean oil, the fermentation resulted in 1500 unitdm1 of extracellular lipase activity. Crude BMS lipase (1.7 kg containing 140,000 units/g) was recovered from the filtrate by ethanol precipitation. BMS lipase and lipase PS-30 were immobilized on Accurel polypropylene (PP). These immobilized lipases were reused (10 cycles) without loss of enzyme activity, productivity, or the optical purity of the product in the resolution process. The enzymatic process for the resolution of racemic acetate 6 was scaled up to 75 and 150 liter at a lO-g/liter substrate concentration using immobilized BMS lipase and lipase PS-30, respectively. From each reaction batch, 3-(R)-acetate 8 was isolated in 88-90 M% yield and 99.5% optical purity. 3-(R)-acetate 8 was chemically converted to 3-(R)-alcohol9. The C-13 taxol side chain (@ or 9) produced either by the reductive or resolution process could be coupled to baccatin I11 or 10-DAB after protection and deprotection of each compound to prepare taxol by semisynthetic process (Fig. 5).

98

R. N. PATEL

4 Rncemic Acetate

s (3R)-A~etate

s

2

(3R)-Acetate

(~s)-Alcohol

2 (3R)-Aleohol [C-13 T a d Side-chain]

FIG.4. Synthesis of taxol side-chain synthon. Stereoselective hydrolysis of cis-3-(acetyloxy)-4-phenyl-2-azetidinone6 (Pate1 et al., 1994a). Reprinted with permission from Portland Press Ltd.

Preparation of taxol side-chain precursors by the lipase-catalyzed enantioselective esterification of methyl trans-j3-phenylglycidate has been demonstrated (Gou et al., 1993). The preparation of enantiomerically pure 3-hydroxy-4-phenyl p-lactam by lipase-catalyzed enantioselective hydrolysis and transesterification of racemic esters and alcohols, respectively, have also been described by Brieva et al. (1993).

Ill. Thromboxane A2 Antagonist

Thromboxane A2 (TxAZ) is an exceptionally potent proaggregatory and vasoconstrictor substance produced by the metabolism of arachidonic acid in blood platelets and other tissues. Together with the potent antiaggregatory and vasodilator, it is thought to play a role in the maintenance of vascular homeostasis and to contribute to the pathogenesis of a variety of vascular disorders. Approaches towards limiting

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

99

+ Taxd Sldechain h

J

Baccatln-ill2 or 1&DAB 9

Taxol Sldechain g

Taxol 1 FIG.5. Semisynthesis of taxol 1. Coupling of baccatin I11 2 and taxol side-chain or 9. synthon

the effect of TxA2 have focused on either inhibiting its synthesis or blocking its action at its receptor sites by means of an antagonist (Nakane, 1987; Ford-Hutchinson, 1991). The lactol [3aS-(3aa,4a,7a, 7aa)]-hexahydro-4,7-epoxyisobenzo-furan-l-( 3H)-one or corresponding chiral lactone 11 are key chiral intermediates for the total synthesis of [~S-[~a,~a(Z),3a,4a[[-~-[3-[[[[~-oxoheptyl)-amine]acetyllmethyl]-~-oxabicyclo-[~.~.~]-hept-2-yl]-~-heptanoic acid 12,a new cardiovascular agent useful in the treatment of thrombotic disease (Das et al., 1987; Hamanka et al., 1990). Horse liver alcohol dehydrogenase (HLADH) catalyzes the oxidoreduction of a variety of compounds (Jones, 1986; Yamada and Shimizu, 1988). It has been demonstrated that HLADH catalyzes the stereospeci-

100

R. N. PATEL

Dlol

Thromboxane A2 Antagonisifl

FIG.6 . Synthesis of chiral synthon for thromboxane A2 antagonist. Stereoselective oxidation of (exo,exo)-~-oxabicyclo[~.~.~]heptane-~,~-dimethanol 13 to the corresponding lactol and lactone 11 (Patel st nl., 1992a). Reprinted with permission from Elsevier Science Inc.

fic oxidation of only one of the enantiopic hydroxyl groups of acyclic and monocyclic meso-diols (Lok et al., 1985, Jones and Francis, 1984). They demonstrated the oxidation of meso exo- and endo-7-oxabicyclo[2.2.l]heptane dimethanol to the corresponding enantiomerically pure y-lactones by HLADH. Nicotinamide adenine dinucleotide (NAD+) and flavin adenine dinucleotide (FAD) at concentrations of 1 and 20 mM, respectively, were required for the stereoselective oxidation of 12.7 mM of substrate. Due to the high cost of enzyme and required cofactors, this process for preparing chiral lactones was economically not feasible for scale-up. Patel et al. (1992a) described the stereoselective oxidation of (exo,exo)-7-oxabicyclo[2.2.~]heptane-2,3-dimethanol~ to the corresponding chiral lactol and lactone 11 (Fig. 6) by cell suspensions (10% wtlvol, wet cells) of Nocardia globerula ATCC-15592 and Rohdococcus sp. ATCC-15592. The reaction yield of 70 M% and optical purity of 96% were for chiral lactone 11 after a 96-h biotransformation process at a 5-glliter substrate concentration using cell suspensions of N. globerula ATCC-15592. An overall reaction yield of 46 M% (lactol and lactone combined) and optical purities of 96.7 and 98.4% were obtained for lactol and lactone 12, respectively, using cell suspensions of Rhodococcus sp. ATCC-15592. Substrate 13 was used at a concentration of 5 glliter. The stereoselective asymmetric hydrolysis of (exo,exo)-7-oxabicyclo[2.2.l]heptane-2,3-dimethanol diacetate ester 14 to the corresponding chiral monoacetate ester 15 has been demonstrated with lipases (Patel

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

hilonoacetate Ester 15

Dlacetate Ester I4

ThromboxaneA2 antagonlst

101

J

--

Oxidation & Hydrolysis

Lactol 1p Frc;. 7. Synthesis of chiral synthon for thromboxane A2 antagonist. Asymmetric hydrolysis of ( e x o , e x o ) - ~ - o x a b i c y c l o [ ~ . ~ . ~ ] h e p t a n ediacetate - ~ , 3 o l ester 14 (Pate1 et a/., 1992b). Reprinted with permission from Springer-Verlag.

et al., 1992b). Lipase PS-30 from Pseudomonas cepacia was most effective in asymmetric hydrolysis to obtain the desired enantiomer of monoacetate ester. The reaction yield of 75 M% and optical purity of >99% were obtained when the reaction was conducted in a biphasic system with 10% toluene at 5 g/liter of substrate. Lipase PS-30 was immobilized on Accurel polypropylene (PP),and the immobilized enzyme was reused ( 5 cycles) without loss of enzyme activity, productivity, or optical purity. The reaction process was scaled u p to 80 liter (400 g of substrate), and monoacetate ester 15 was isolated in 80 M% yield with 99.3% optical purity, as determined by chiral HPLC and NMR analysis. The isolated product gave a GC HI of 99.5%. The chiral monoacetate ester 15 was oxidized to its corresponding aldehyde and subsequently obtained hydrolyzed to give chiral lactol lo (Fig. 7). The chiral lactol by this enzymatic process was used in chemoenzymatic synthesis of thromboxane A2 antagonist 12. IV. ACE Inhibitors

Captopril is designated chemically as 1-[(2S)-3-mercapto-Z-methylpropionyll-,-proline (16).It is used as an antihypertensive agent through suppression of the renin-angiotensin-aldosterone system (Ondetti and Cushman, 1981; Ondetti et al., 1977; Cushman and Ondetti,

102

R. N. PATEL

FIG.8. Synthesis of chiral captopril side chain. Stereoselective hydrolysis of thioester of 3-acylthio-2-methyl propanoic acid 17.

1980).Captopril and such other compounds as enalapril and lisinopril (Fig. 8) prevent the conversion of angiotensin I to angiotensin I1 by inhibition of ACE (angiotensin converting enzyme). The potency of captopril Is as an inhibitor of ACE depends critically on the configuration of the mercaptoalkanoyl moiety; the compound with the S configuration is about 100 times more active than its corresponding R-enantiomer (Cushman et al., 1977). The required 3-mercapto-(ZS)methylpropionic acid moiety has been prepared from the microbially derived chiral3-hydroxy-(2R)-methylpropionic acid, which is obtained by the hydroxylation of isobutyric acid (Goodhue and Schaeffer, 1971; Schimazaki eta]., 1991; Hasegawa et al., 1982). The use of extracellular lipases of microbial origin to catalyze the stereoselective hydrolysis of 3-acylthio-2-methylpropanoic acid ester in an aqueous system has been demonstrated to produce optically active 3-acylthio-2-methyl propanoic acid (Sih, 1987; Gu et al., 1986a; Sakimas et al., 1986). The synthesis of the chiral side chain of captopril by lipase-catalyzed enantioselective hydrolysis of the thioester bond of racemic 3-acetylthio-2-methyl propanoic acid Lz to yield S-(-)-Uhas . various lipases evalubeen demonstrated (Pate1 et al., 1 9 9 2 ~ )Among ated, that from Rhizopus oryzae ATCC-24563 (heat-dried cells) and lipase PS-30 in an organic solvent system (1,1,2-trichloro-1,2,Z-trifluoroethane or toluene) catalyzed the hydrolysis of the thioester bond of an undesired enantiomer of racemic c7 to yield desired S - ( - ) - u , R-(+)3-mercapto-2-methylpropanoicacid 18,and acetic acid 19 (Fig. 8). The

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

103

reaction yield of >24% (theoretical maximum 50%) and optical purity of >95% were obtained for S-(-)-Q using each lipase in an independent experiment. In an alternate approach to prepare the chiral side chain of captopril and zofenopril, the lipase-catalyzed stereoselective esterification of racemic 3-benzoylthio-2-methylpropanoic acid 20 in an organic solvent system was demonstrated to yield R-(+)-methyl ester 21 and unreacted acid enriched in the desired S-(-) enantiomer 20 (Patel et a]., 1991a). Using lipase PS-30 with toluene as solvent and methanol as nucleophile, the desired S-(-)-20 was obtained in 37% reaction yield and 97% optical purity. Substrate was used at 22 g/liter concentration. The amount of water and concentration of methanol supplied in the reaction mixture were very critical. Water was used at 0.1% concentration in the reaction mixture. Higher than 1%water led to aggregation of enzyme in the organic solvent along with a decrease in the rate of reaction due to mass transfer limitation. The rate of esterification decreased as the methanol-to-substrate ratio was increased from 1:1 to 4:l. Higher methanol concentrations probably inhibited the esterification reaction by stripping the essential water from the enzyme. Crude lipase PS-30 was immobilized on three different resins (XAD-7, XAD-2, and Accurel PP) in absorption efficiencies of about 68, 71, and 98.5%, respectively. These immobilized lipases were evaluated for the ability to stereoselectively esterify racemic 20. Enzyme immobilized on Accurel PP catalyzed efficient esterification, giving a reaction yield of 36-45% and 97.7% optical purity of The immobilized enzyme under identical conditions gave similar optical purity and yield of product in 23 additional reaction cycles without any loss of activity and productivity. S-(-)-= is a key chiral intermediate for the synthesis of captopril (Moniot, 1988) or zofenopril 22 (Ondetti et al., 1982); both are antihypertensive drugs (Fig. 9). The S-(-)-a-[(acetylthio)methyl]benzenepropanoicacid 23 is a key chiral intermediate for the neutral endopeptidase inhibitor 24 (Delaney et a]., 1988; Seymour et al., 1991). The lipase-catalyzed stereoselective hydrolysis of thioester bond of racemic a-[(acetylthio)methyllbenzenepropanoic acid 23 was carried out in organic solvent to yield R-(+)-a[(mercapto)methyl]benzenepropanoicacid 25 and S-(-)-= (Patel e f al., 1 9 9 2 ~ )Using . lipase PS-30, the S-(-)-= was obtained in 40% reaction yield (theoretical maximum 50%) and 98% optical purity (Fig. 10). The S-(-)-2-cyclohexyl-l,3-propanediolmonoacetate ZS and the S-(-)-2-phenyl-l,3-propanediol monoacetate 22 are key chiral intermediates for the chemoenzymatic synthesis of Monopril 28 (Fig. ll),a new

,%(-)-a.

FIG.9. Synthesis of chiral zofenopril side chain. Stereoselective esterification of 3-benzylthio-2-methyl propanoic acid (Pate1 et al., 1991a). Reprinted with permission from Springer-Verlag.

3

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

0

I

f

0

+ I

\

)=o

0 111.

d ' +

105

106

~A O, , J , ,O CA

R. N.PATEL

0 Dlacetate 29

ACO-OH

Biphasic System PPL

0

*

Chromobacterlumvlacosum Upase

Dlacetatr 3Q

ACO-OH

S-(-)-Monoacetate Zz

0 FIC.11. Synthesis of chiral synthon for monopril. Asymmetric hydrolysis of Z-cyclohexyl-2 and 2-phenyi-1,3-propanediol diacetate 30 (Patel et a]., 1990). Reprinted with permission from Springer-Verlag.

hypertensive drug which acts as an ACE inhibitor. The asymmetric hydrolysis of 2-cyclohexyl-1,3-propanedioldiacetate 29 and 2-phenyl1,3-propanediol diacetate 30 to the corresponding S-(-) monoacetate S and S-(-) monoacetate 22 by porcine pancreatic lipase (PPL) and Chromubacterium viscosum lipase have been demonstrated by Patel et al. (1990). In a biphasic system using 10% toluene, a reaction yield of >65% and optical purity of 99% were obtained for S-(-)-2J using each enzyme. S - ( - ) - B was obtained in 90% reaction yield and 99.8% optical purity using C. viscosum lipase under similar conditions. The stereoselective hydrolysis of dimethylesters of symmetrical dicarboxylic acids including meso-diacids such as cis-l,2-cycloalkane dicarboxylic acids and diacids with prochiral centers has been demonstrated by Mohr et al. (1983) using pig liver esterase (PLE). The product of these stereoselective hydrolyses, chiral monoacetate of dicarboxylic acids, were obtained with an enantiomeric excess (e.e.) from 10 to 90% depending upon the substrate. Enantioselective hydrolysis of cis-1,2-

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

107 NHZ

I

coa

HCOOH

Formnte Dehydrogsnass CBZ-L-lysine

33

L-z-Oxylyslne 32

Keto d d 34

i"; 0

C.nnoprll3l FIG.12. Synthesis of chiral synthon for ceranopril. Conversion of CBZ-L-lysine 33 to L-Z-oxylysine 32 (Pate1 et al., '1992b). Reprinted with permission from Springer-Verlag.

diacetoxycycloalkane and 2-substituted 1,3-propanediol diacetate have been demonstrated using PPL by Laumen and Schneider (1985) and Tombo et al. (1986), respectively. Ceranopril 31 is another ACE inhibitor (Karenewsky et al., 1988) that requires chiral intermediate carbobenzoxy (CBZ)-L-oxylysine 32 (Fig. 12). A biotransformation process was developed to prepare CBZ-Loxylysine (Hanson et al., 1992). NE-CBZ-L-lysine 33 was first converted to the corresponding keto acid 34 by oxidative deamination using cells of Providencia alculifaciens SC-9036 that contained L-amino acid oxidase and catalase. The keto acid 34 was subsequently converted to CBZ-L-oxylysine 32 using L-2-hydroxyisocaproate dehydrogenase from Lactobacillus confisus. The NADH required for this reaction was regenerated using formate dehydrogenase from Candidu boidinii (Fig. 12). The reaction yield of 95% along with 98.5% optical purity was obtained in overall process. V. Anticholesterol Drugs

Chiral P-hydroxy esters are versatile synthons in organic synthesis specifically in the preparation of natural products (Mori and Tanida,

108

R. N. PATEL

1984; Hirama and Uei, 1982; Gopalan and Sih, 1984). The asymmetric reduction of carbonyl compounds using Baker’s yeast has been demonstrated and reviewed (Ward and Young, 1990; Csuk and Glanzer, 1991; Sih et a]., 1984). In the stereoselective reduction of P-ketoester of 4chloro and 4-bromo-3-oxobutanoic acid, specifically 4-chloro-3-oxobutanoic acid methylester, Sih and Chen (1984b) demonstrated that the sterteoselectivity of yeast-catalyzed reductions may be altered by manipulating the size of the ester group using y-chloroacetoacetate as substrate. They also indicated that the e.e. of the alcohol produced depended upon the concentration of the substrate used. Nakamura et al. (1989) demonstrated the reduction of P-keto ester with Baker’s yeast and controlled stereoselectivity by the addition of a$-unsaturated carbony1 compounds. The additive tended to shift the stereoselectivity of the reduction reaction toward the production of R-hydroxy ester. The shift in stereoselectivity was accounted for based on inhibition of the competitive enzyme that produced S-hydroxy ester. They also used immobilized Baker’s yeast to improve the stereoselectivity of the reduction reactions (Nakamura et a]., 1990). The enantiomeric excess of alcohols produced was improved to 90% by using immobilized cells compared to 31% obtained with free cells when methyl 4-chloroacetoacetate was use as the substrate. Pate1 et al. (1992d) have described the reduction of 4-chloro-3-oxobutanoic acid methyl ester 35 to S-(-)-4-chloro-3-hydroxybutanoicacid methyl ester 36 (Fig. 13) by cell suspensions of Geotrichurn candidurn SC-5469. S-(-)-E is a key chiral intermediate in the total chemical synthesis of 37, a cholesterol antagonist that acts by inhibiting hydroxymethyl glutaryl CoA (HMG-CoA)reductase. In the biotransformation process, a reaction yield of 95% and optical purity of 96% were obtained for &6 by glucose-, acetate-, or glycerol-grown cells (10% wthol) of G. candidurn SC-5469. Substrate was used at a concentration of 10 g/liter. The optical purity of S was increased to 99% by heat treatment of cell suspensions (55% for 30 min) prior to conducting bioreduction of 35. Glucose-grown cells of G. candidurn SC-5469 have also catalyzed the stereoselective reduction of ethyl-, isopropyl-, and tertiary-butyl esters of 4-chloro-3-oxobutanoic acid and methyl and ethyl esters of 4-bromo3-oxobutanoic acid. A reaction yield of ~ 8 5 %and an optical purity of >94% were obtained. NAD+-dependent oxidoreductase responsible for the stereoselective reduction of P-keto esters of 4-chloro- and 4-bromo3-oxobutanoic acid was purified 100-fold. The molecular weight of purified enzyme was 950,000 Da. The purified oxidoreductase was

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

109

Q F

HMGCoA Reductme InhibitorX FIG.13. Synthesis of chiral synthon for anticholesterol drug. Stereoselective reduction of 4-chloro-3-oxobutanoic acid methyl ester 35 (Pate1 et al., 1992a). Reprinted with permission from Elsevier Science Inc.

immobilized on a Eupergit C and used to catalyze the reduction of 35 to S. The cofactor NADH required for the reduction reaction was regenerated by glucose dehydrogenase. Nakamura e f al. (1991) have isolated four different oxidoreductases from Baker's yeast that catalyzed the reduction of P-keto esters to P-hydroxy esters. Two oxidoreductases, namely, D-enzyme-1 (molecular weight 25,000) and D-enzyme 2 (MW 1,600,000 Da), catalyzed the reduction of P-keto ester stereoselectivity to D-P-hydroxy ester. In contrast, one other enzyme, L-enzyme (MW 321,000 Da), reduced substrates to L-J3-hydroxy esters. The NADP-oxidoreductases, designated the Sand R-enzymes, have been purified and characterized from cell extracts of Saccharomyces cerevisiae, and catalyze the enantioselective reductions of ~ - o x o - 4-oxo-, , and 5-0x0-esters (Heidlas et al., 1988). The S-enzyme had a molecular weight of 48,000 and reduced 3-oxo-esters, ~ - o x o -and , 5-0x0-acids and esters enantioselectively to S-hydroxy compounds in the presence of NADPH. This enzyme may be located in the mitochondria1 fraction. The R-enzyme, which had a molecular weight of 800,000 and contained subunits having molecular weights of 200,000 and 210,000, specifically reduced 3-0x0-esters to R-hydroxy esters using NADPH as coenzyme. The R-enzyme, which occurs in the cytosol, was considered to be identical to a subunit of the fatty acid synthetase complex.

110

R. N. PATEL

Most microorganisms and enzymes derived therefrom have been used in reduction P-keto or a-keto compounds involved reduction of singleketo groups (Jones and Beck, 1986; Keinan et al., 1986; Patel et d.,1981; Bradshaw et al., 1992a; Christen and Crout, 1988). Patel et al. (1993b) have demonstrated the stereoselective reduction of a diketone 3,5-dioxo-6-(benzyloxy) hexanoic acid ethyl ester 38 to (3S,5R)-dihydroxy-6(benzyloxy) hexanoic acid ethyl ester (Fig. 14). The compound is a key chiral intermediate required for the chemical synthesis of [4-[4a,6~(E)ll-6-[4,4-bis-[4-fluorophenyl)-3-(l-methyl-lH-tetrazol-5-yl) -1,3-butadienyl]-tetrahydro-4-hydroxy-2H-pyren-2-one. Compound Ra new anticholesterol drug, acts by inhibition of HMG-CoA reductase (Sit et al., 1990). Among various microbial cultures evaluated for the stereoselective reduction of diketone 38, cell suspensions of Acinetobacter calcoaceticus SC-13876 reduced 38 to -. The reaction yield of 85% and optical purity of 97% were obtained using glycerolgrown cells. The substrate was used at 2 g/liter and cells at 20% (wt/vol, wet cells) concentration. Cell extracts of A. calcoaceticus SC-13876 in the presence of NAD+, glucose, and glucose dehydrogenase reduced 38 to the corresponding monohydroxy compounds 41 and 42 (3-hydroxy-5-oxo-6-benzyloxy hexanoic acid ethyl ester 41, and 5-hydroxy-3-oxo-6-benzyloxy hexanoic acid ethyl ester 4 2 ) . Both 41 and 112 were further reduced to (3S,4R)-dihydroxy compound by cell extracts (Fig. 14). A reaction yield of 9 2 % and an optical purity of 99Y0 were obtained when the reaction was carried out in a 1-liter batch using cell extracts. The substrate was used at 10 g/liter. Product 39a was isolated from the reaction mixture in 72% overall yield. The GC and HPLC area purity of the isolated product was 99% and the optical purity was 99.5%. Reductase that converted 38 to was purified about 200-fold from cell extracts of A. calcoaceticus SC-13876. The purified enzyme gave a single protein band on SDS-PAGE corresponding to 33,000 Da. Using a resolution process, chiral alcohol R-(+)-@ was also prepared by the lipase-catalyzed stereoselective acetylation of racemic 40 in organic solvent (Patel et al., 1992e). They evaluated various lipases, among which lipase PS-30 (Amano International Enzyme Co.) and BMS lipase efficiently catalyzed acetylation of the undesired enantiomer of racemic 40 to yield S-(-)-acetylated product 3 and unreacted desired R-(+)-* (Fig. 15). A reaction yield of 49 MY" (theoretical maximum 50 MYv) and an optical purity of 98.5% were obtained for R - ( + ) - a when the reaction was conducted in toluene as solvent in the presence of isopropenyl acetate as acyl donor. Substrate was used at a concentration

-

(+)-a,

-

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

0

111

FIG.14. Synthesis of chiral synthon for anticholesterol drug. Stereoselective reduction of 3,5-dioxo-6-~enzyloxy)hexanoic acid ethyl ester 38 (Pate1 et al., 1993b). Reprinted with permission from Elsevier Science Inc.

+ +

N

F F

+ ____)

F F

FIG.15. Stereoselective acetylation of racemic 40 to S-(-)-acetate 43 and R-(+)-m, an anticholesterol drug (Pate1 et al., 199Ze). Reprinted with permission from Springer-Verlag.

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

113

of 4 g/liter. In methyl ethyl ketone at a 50-g/liter substrate concentration, a reaction yield of 46 M% and optical purity of 96.4% were obtained for R-(+)-@. Lipase PS-30 was immobilized on Accurel PP and the immobilized enzyme reused five times without any loss of activity or productivity in the resolution process to prepare R-(+)-@. The enzymatic process was scaled up to a 640-liter preparative batch using immobilized lipase PS-30 at 4 g/liter and racemic substrate 40 in toluene as solvent. From the reaction mixture R - ( + ) - a was isolated in 35 M*h overall yield with 98.5% optical purity and 99.5% chemical purity. The undesired S-(-)-acetate 43 produced by this process was enzymatically hydrolyzed by lipase PS-30 in a biphasic system to prepare the corresponding S-(-)-alcohol @. Thus, both enantiomers of alcohol 40 were produced by the enzymatic process. Pravastatin 44 and Mevastatin 45 are anticholesterol drugs that act by competitively inhibiting HMG-CoA reductase (Endo et al., 1976a). Pravastatin sodium is produced by two fermentation steps. The first step is production of compound ML-236B 46 by Penicillium citrinum (Endo et al., 1976a,b; Hosobuchi et al., 1993a,b). Purified 46 was converted to its sodium salt with sodium hydroxide and in the second step was hydroxylated to Pravastatin sodium 44 (Fig. 16) by Streptomyces carbophilus (Serizawa et al., 1983). A cytochrome P,,,-containing enzyme system has been demonstrated from S. carbophilus that catalyzed the hydroxylation reaction (Matsuoka et al., 1989). The chiral intermediate 2,4-didoexyhexose derivative required for HMG-CoA reductase inhibitors has also been prepared using 2-deoxyribose-5-phosphate aldolase (DERA).This enzyme accepts a wide variety of acceptor substrates and has been useful in organic synthesis (Barbas et a]., 1990; Chen et al., 1992). As shown in Fig. 1 7 , the reactions start with stereospecific addition of acetaldehyde 47 to a substituted acetaldehyde to form a 3-hydroxy-4-substituted butyraldehyde 48, which subsequently reacts with another acetaldehyde to form a 2,4-dideoxyhexose derivative 49 (Gijsen and Wong, 1994). DERA has been overexpressed in Escherichia coli.

VI. Antiinfective Drugs

During the past several years, synthesis of a-amino acids has been pursued intensely (Williams, 1989; O’Donnell, 1988; O’Donnell et al., 1989, 1970; Evans et al., 1987; Schmidt et al., 1989; Imada et a]., 1981) because of their importance as building blocks of compounds of medicinal interest, particularly antiinfective drugs. The asymmetric synthesis

114 R. N.PATEL

0

c

FIG. 16. Stereoselective hydroxylation of ML-236B to Pravastatin 44.

0

"i a' B

a

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

'

L 2 - J

a

FIG.17. Preparation of 2,4-dideoxyhexosederivatives 49 by aldolase (Gijsen and Wong, 1994). Reprinted with permission from the American Chemical Society.

115

R. N.PATEL

116

NADH

NAD*

\

H 2 N A o H

I

NH4*

Coda

a-Keto-&hydroxylsovalerate 52

L-&hydroxyvallneB

\COa

[(CH3)3N*CH2CH20H]a

Tigemonam 51 FIG.18. Synthesis of chiral synthon for tigemonam. Stereoselective reductive amination of a-keto-P-hydroxyisovalerate52 (Hanson et ol., 1990).

of P-hydroxy-a-amino acids by various methods has been demonstrated (Bold et al., 1989;Evans et al., 1987;Ito et al., 1988;Saito et al., 1985; Gordon et al., 1982)because of their utility as starting materials for the total synthesis of monobactam antiobiotics. L-P-hydroxyvaline So (Sykes e f a]., 1981) is a key chiral intermediate required for the total synthesis of an orally active monobactam, Tigemonam 51 (Fig. 18).The resolution of CBZ-0-hydroxyvaline by chemical methods has been demonstrated (Godfiey et al., 1986;Shanzer et al., 1979;Berse and Bessette, 1971). Leucine dehydrogenase from strains of Bacillus (Schutte et al., 1985;Monot et al., 1987)has been used for the synthesis of branchedchain amino acids. Hanson et al. (1990)have described the synthesis of ~-P-hydroxyvaline50 from a-keto-P-hydroxyisovalerate52 by reductive amination using leucine dehydrogenase from B. sphaericus ATCC-4525 (Fig. 18). NADH required for this reaction was regenerated by either formate dehydrogenase from Candida boidinii or glucose dehydrogenase from B. megaferium. Such immobilized cofactors as polyethylene glycol-NADH and dextrans-NAD were effective in the biocatalytic proc-

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

O HO Amoxicillin 99

'COpH

H

0

117

CHS

@-CON NH2 Cefadroxyl

a

Cephalexin51

FIG.19. Structure of antiinfective agents amoxicillin, cefadroxil, and cephalexin.

ess. The required substrate 52 was generated either from a-keto-P-bromoisovalerate or its ethyl esters by hydrolysis with sodium hydroxide in situ. In an alternate approach, the substrate 52 was also generated from methyl-2-chloro-3,3-dimethyloxiran-carboxylate and the corresponding isopropyl and 1,I-dimethyl ethyl ester. These glycidic esters are converted to substrate 52 by treatment with sodium bicarbonate and sodium hydroxide. In this process, an overall reaction yield of 98% and an optical purity of 99.8% were obtained for the L-P-hydroxyvaline So. D-phenylglycine is required for the semisynthetic antibiotic ampicillin, and D-hydroxyphenylglycine (Fig. 19) is used in the production of amoxicillin and cefadroxyl (Aida et al., 1986; Syldatk et al., 1987). The use of D-p-hydroxy phenylglycine will significantly increase because such new drugs as aspoxicillin, cefbuperazine, and cepyramide are expected to be marketed. Currently, D-amino acids are commercially produced by a chemoenzymatic route using D-hydantoinase. In this process, chemically synthesized DL-5-substituted hydantoin 53 is hydrolyzed to N-carbamoyl-D-amino acid 54 by microbially derived D-hydantoinase (Fig. 20). The latter compound undergoes rapid and spontaneous racemization under the reaction conditions; therefore, theoretically a 100% yield of 54 can be obtained. The compound 54 is further chemically converted to the corresponding D-amino acid 55 (Moller et al., 1988; Morin et al., 1986; Takahashi et al., 1979; Yamada et al., 1978; Kamphuis et a]., 1990b). Microbial N-carbamoylases have

118

R. N. PATEL

DL-5-8ubstltutd Hydantdn

Dsmlno acid I

1

D-HydantolnsM,

PN-carbmoyl acld 54

FIG.20. Synthesis of D-amino acids by D-hydantoinase and carbamoylase.

been demonstrated that catalyzed the conversion of N-carbamoyl-Damino acid to the corresponding D-amino acid. Some organisms contained both D-hydantoinase and N-carbamoylase activity (Olivieri et al., 1979; Runser et a]., 1990; Yokozeki et al., 1987; Kim and Kim, 1995). L-hydantoinase has also been described from a microbial source that catalyzes the conversion of DL-5-substitutedhydantoin to N-carbamoylL-amino acid. This process has been used in the production of L-amino acids (Yokozeki et al., 1987; Syldatk et al., 1987; Nishida et al., 1987; Tsugawa et al., 1966). D-amino acids and L-amino acids have also been prepared by D-Specific or L-specific acylases derived from microbial sources. In this process, DL-N-acetyl amino acid (Fig. 21) is resolved by hydrolytic reaction to yield the D- or L-amino acid 57 depending upon D- or L-selective acylase used in the reaction (Sugi and Suzuki, 1980; Chibata et al., 1988; Chenault et a]., 1989; Sakai et al., 1991). Sandoz commercialized the production of cyclosporin A, which is used in the treatment of transplant rejection and autoimmune disease. L-valine is an essential component of cyclosporin A (Ice and Agathos, 1989; Kobe1 and Traber, 1982). L-a-amino acids have been prepared by the resolution of racemic a-amino acid amide by the L-specific aminopeptidase from Pseudomonas putida ATCC-12633 (Kamphuis et a]., 1990a; Vriesema et al., 1986).Enzyme from Pseudomonas putida ATCC12633 cannot be used to resolve a-alkyl-substituted amino acid amides. Amino amidase from Mycobacterium neoaurum ATCC-25795 has been

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

)r'

R-C-COOH

I

NH

I

o=c

-

I

D-Acylsse

R

I

H,

D&-N-pcetyl Pmlao edd

COOH Hn I

HN &OOH

I o=c

4ch

HZN

I

CHI

119

CH3

L-N-pcetyl -no

d d

D-smino add 9

FIG.21. Synthesis of D-amino acids by o-acylase.

FIG.22. Synthesis of L-a-alkylamino acids by amidase.

used in the preparation of L-acid 58 and D-amide of a-alkyl-substituted amino acids 59 (Fig. 22) by an enzymatic resolution process using racemic a-alkylamino acid amide as a substrate (Kamphuis et al., 1990b, 1992).Amidase from Ochrobactrum anthropi catalyzed the resolution of a,a-disubsituted amino acids, N-hydroxyamino acids, and a-hydroxy acid amides. The resolution process could lead to the production of chiral amino acids or amides in 50% yield. Amino-acid racemases have been used to get 100% yield of chiral amino acids (Kamphuis et al., 1992). Aminotransferases have been used in the production of chiral amino acids (Primrose, 1984; Rozzell, 1987, 1989; Carlton et al., 1986). Aminotransferases catalyze the transfer of an amino group, a proton, and a pair of electrons from a primary amine substrate to the carbonyl group of an acceptor molecule such as oxaloacetate, a-ketoglutarate, or pyruvate (Fig. 23). o-aminotransferases from Pseudomonas sp. F-126 have also been used to produce homochiral amine products (Burnett et al., 1979). The stereoselective removal of either the pro-(S) or the pro-(R) proton has been demonstrated (Bouclier et al., 1979; Tanizawa et al., 1982). w-aminotransferase specific for the secondary amines has also been demonstrated from Bacillus megaterium and Pseudomonas aerugiRosa (Stirling, 1992).

120

R. N. PATEL

HOOCHsC

L-Aspartic acid

SKeto a d d Amlnotransferase COOH

0 Oxaloacatate FIG. 23.

L-Amino acid

Synthesis of L-amino acids by aminotransferase.

The bioconversion of nitriles and primary amides have been used in the production of such optically active a-hydroxy or a-amino acids as L-phenylalanine, L-lactic acid, and L-phenylglycine, which are a chiral synthon in many pharmaceutical syntheses. Nitrilases have been isolated from organisms belonging to genus Brevibacterium, Rhodococcus, and Pseudornonas (Asano et al., 1982; Bui et al., 1982; Kobayashi et al., 1988; Nagasawa et al., 1986, 1987). After the discovery of the antibiotic thienamycin, compounds that contain the carbapenem and penem ring systems have attracted much attention. The importance of the stereochemistry of the hydroxyethyl group is demonstrated by the fact that this group must be in the R configuration for antimicrobial activity. Previously, synthesis of carbapenem and penem compounds have often utilized the optically active p-lactam intermediates [Fujimoto et al., 1986; Alpegiani et al., 1985; Shibata et al., 1985). ~-(-)-3-hydroxybutyric acid prepared by the microbial hydroxylation of butyric acid have been used in p-lactam synthesis (Iimori and Shibazaki, 1985; Ohashi and Hasegawa, 1992). VII. Calcium Channel Blocking Drugs

a,

Dilthiazem a benzothiazepinone calcium channel blocking agent that inhibits influx of extracellular calcium through L-type voltage-operated calcium channels, has been widely used clinically in the treatment of hypertension and angina [Chaffman and Brogden, 1985). Since dilthiazem has a relatively short duration of action (Kawai et al., 1981), an 8-chloroderivative has been introduced in the clinic as a more potent

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

121

analogue of dilthiazem (Isshiki et al., 1988). Lack of extended duration of action and little information on the structure-activity relationship in this class of compounds led Floyd et al. (1990) and Das et al. (1992a,b) to prepare isosteric 1-benzazepin-2-ones and led to the identification of the 6-trifluoro-methyl-2-benzazepin-Zone derivative as a longer-lasting and more potent antihypertensive agent. A key chiral intermediate 52 ((3R-cis)-l,3,4,5-tetrahydro-3-hydroxy-4-(4-methoxyphenyl)-6-(trifluoromethyl)-2H-l-benzazepin-Z-one) was required for the total chemical synthesis of the new calcium channel blocking agent @ ((cis)-3-(acetoxy)~-[2-(dimethylamino)ethyl]-~,3,4,5-tetrahydro-4-(4-methoxyphenyl)-6-triflutomethy1)-2H-1-benzazepin-2-one). A stereoselective microbial process (Fig. 24) was developed for the reduction of 4,5-dihydro-4-(4methoxyphenyl)-6-(trifluoromethyl)-1H-l-benzazepin-2,3-dione 64 to chiral g (Pate1 et al., 1991b). Compound % exits predominantly in the achiral enol form, which is in rapid equilibrium with the 2-keto form enantiomers. Reduction of @ could give rise to formation of four possible alcohol stereoisomers. Remarkably, conditions were found under which only the single-alcohol isomer 52 was obtained by microbial reduction. Among various cultures evaluated, microorganisms from the genera Nocardia, Rhodococcus, Corynebacterium, and Arthobacter reduced compound @ to compound g with a 60-70% conversion yield at 1g/liter substrate concentration. The most effective culture, Nocardia salmonicolor SC-6310, catalyzed the bioconversion of to s2 in a 96% reaction yield with 99.9% optical purity at a Z-g/liter substrate concentration. Product g was isolated and identified by NMR and MS. A preparative-scale fermentation process for growth of N.salmonicolor and a bioreduction process using cell suspensions of the organism were demonstrated. VIII. Antipsychotic Agents

Much effort has been directed towards the understanding of the Sigma receptor system in the brain and endocrine tissue. This effort has been motivated by the hope that the Sigma site may be a target for a new class of antipsychotic drugs (Ferris et al., 1991; Junien and Leonard, 1989; Walker et al., 1990). Characterization of the Sigma system helped clarify the biochemical properties of the distinct haloperidol-sensitive Sigma binding site, the pharmacological effects of Sigma drugs in several assay systems, and the transmitter properties of a putative endogenous ligand for the Sigma site (Steinfels et a]., 1989; Massamiri and Duckles, 1990; Martinez and Bueno, 1991; Taylor et al., 1991). R-(+) compound @ (BMY-14802)is a Sigma ligand and has a high affinity for

sc 6310 &-

Dilth&uem_BL

Calcium Channel B l o c k P

I SO 32191 P

FIG.24. Synthesis of chiral synthon for calcium channel blocker. Stereoselective reduction of 4,5-dihydro methoxyphen~~l)-6-~trifluoromethyl~-lH-benzazepin-2,3-dione @ (Patel et al., 1991b). Reprinted with permission from Els Science Inc.

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

Mlcrobydtlon

Ketone p

123

~

Antlpsychotlc agent R-(+)-BMY 14002 @

FIG.25. Preparation of R-(+)-BMY-14802, an antipsychotic agent. Stereoselective reduction of ~-(~-fluorophenyl)-~-[~-(5-fluoro-~-pyrimidinyl)-l-piperazinyl]-l-butanone 66 (Patel et al., 1993c). Reprinted with permission from Portland Press Ltd.

Sigma binding sites and antipsychotic efficacy. The stereoselective microbial reduction of keto compound l-(4-fluorophenyl)-4-[4-(5-fluoro-2pyrimidiny1)-1-piperazinyll-1-butanone @ to yield the corresponding hydroxy compound R-(+)-BMY-14802@ (Fig. 25) has been developed by Patel et al. (1993~). Among various microorganisms evaluated for the reduction of ketone 66, Mortierella ramanniana ATCC-38191 predominantly reduced compound &6 to R-(+)-BMY-14802and Pullularia pullulans ATCC-16623 reduced compound @ to S-(-)-BMY-14802. An optical purity of >98% was obtained in each reaction. In a two-stage process for reduction of compound @, cells of M . ramanniana ATCC-38191 were grown in a 380-liter fermenter containing 250 liter of medium and harvested after 31 hours of growth. Cells harvested from a 380-liter fermenter were used for the reduction of ketone 66 in a 15-liter fermenter using 10 liter of cell suspensions (20% wt/vol, wet cells). Ketone @ was used at a 2-g/liter concentration and glucose was supplemented at a 20-g/liter concentration during the biotransformation process to generate NADH required for reduction. After a 24-h biotransformation period, about a 90% yield (99.0% optical purity) of R-(+)-BMY-14802was obtained. The R-(+)-BMY-14802 was isolated from the 10-liter fermentation broth in overall 70 MY0 yield and 99% GC and HPLC purity. Isolated R-(+)-BMY-14802 gave an optical purity of 99.5”/0as analyzed by chiral HPLC. A single-stage fermentation-biotransformation process was demonstrated for reduction of ketone 66 to R-(+)-BMY-14802by cells of M . ramanniana ATCC-38191. Cells were grown in a 20-liter fermenter containing 15 liter of medium. After 40 h of growth in a fermenter, when the residual glucose was depleted (0.1”/0)and the pH of the medium

124

R. N. PATEL

dropped to 4.5, the biotransformation process was initiated by addition of 30 g of ketone 54 and 300 g of glucose. The biotransformation process was completed within a 24-h period, with a reaction yield of 100% and an optical purity of 98.9% for R-(+)-BMY-14802.At the end of the biotransformation process, cells were removed by filtration and product was recovered from the filtrate in overall 80% recovery. The isolated product had 99% HPLC purity and 98.8% optical purity. A reductase with a molecular weight of 29,000 Da has been purified to homogeneity which catalyzed the conversion of ketone 66 to R-(+)-BMY-14802, For the production of optically active alcohols, reduction of the inexpensive prochiral ketones is a promising method. Commercially available alcohol dehydrogenases derived from horse liver and Thermoanaerobium brockii (Jones, 1986; Yevich et al., 1986) have been used in the preparation of chiral alcohols. Alcohol dehydrogenase from T brockii is heat stable and has broad substrate specificity toward aliphatic ketones. Substrates with bulky side chains (such as acetophenone) are poor substrates. Alcohol dehydrogenase from yeast, horse liver, and T brockii transfer the pro-R hydride to the reface of the carbonyl to give (S) alcohols, a process described by Prelog's rule (Jones and Beck, 1986; Keinan et al., 1986). Alcohol dehydrogenases from Pseudomonas sp. strain PED and Lactobacillus kefir and Mortierella isabellina have been shown to catalyze the enantioselective reduction of aromatic, cyclic, and aliphatic ketones to the corresponding chiral alcohols (Bradshaw et al., 1992a,b). Both enzymes exhibit anti-Prelog specificity, transferring the pro-R hydride to form (R) alcohols. Most oxidoreductases used for the preparation of optically active alcohols involve the use of NADH as cofactor. Simon (1990) demonstrated the use of reductases from anaerobic Clostridium strains that catalyzed the reduction of a variety of compounds to optically active alcohols using methyl or benzyl viologen as an electron donor. 2-enoate reductase and 2-oxocarboxylate reductase have been used in the stereoselective reduction of carbon-carbon- and carbon-oxygen-containing compounds R-(+)-BMY-14802@ has also been prepared by lipase-catalyzed resolution of racemic BMY-14802 acetate ester fl (Hanson et al., 1994b). Lipase from Geotrichum candidum (GC-20 from Amano Enzyme Co.) catalyzed the hydrolysis of acetate @ t o R-(+)-BMY-14802(Fig. 26) in a biphasic solvent system in 48% reaction yield and 98% optical purity. The rate and enantioselectivity of the hydrolytic reaction was dependent upon the organic solvent used. The enantioselectivity ( E values) ranged from 1in the absence of solvent to >loo in dichloromethane and

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

1

GC-20 Toluene /H20

125

t

R-(+)-BMY-14802 45

FIG.26. Preparation of R-(+)-BMY-14802,an antipsychotic agent. Stereoselective hydrolysis of BMY-14802 acetate 67 (Patel et al., 1995). Reprinted with permission from Elsevier Science t t d .

toluene. S-(-)-BMY-l4802 was also prepared by chemical hydrolysis of undesired BMY-14802 acetate obtained during the enzymatic resolution process. IX. Antiarrhythmic Agents Larsen and Lish (1964) reported the biological activity of a series of phenethanolamine-bearing alkyl sulfonamido groups on the benzene ring. Within this series, some compounds possessed adrenergic and antiadrenergic actions. D-(+)-SOtalOlis a beta-blocker (Uloth et al., 1966) that, unlike other beta-blockers, has antiarrhythmic properties and has no other peripheral actions (Lish et al., 1965). The P-adrenergic-blocking drugs such as propranolol and sotalol have been separated chemically into the dextro- and levorotatory optical isomers, and it has been demonstrated that the activity of the lev0 isomer is 50 times that of the corresponding dextro isomer (Somani and Bachand, 1969). Chiral alcohol X fJ is a key intermediate for the chemical synthesis of D-(+)-sotalol m. The stereoselective microbial reduction of N-(4-(2-chloroacety1)phenyl)methanesulfonamide 70 to the corresponding (+)-alcohol@ (Fig. 2 7 ) has been demonstrated (Patel el al., 1993d). Among numbers of microorganisms screened for the transformation of compound 70 to compound @, Rhodococcus spp. ATCC-29675, ATCC-21243, Nocardia salmonicolor SC-6310, and Hansenula pofymorpha ATCC-26012 gave the desired chiral alcohol @ in >90% optical purity. H. polymorpha ATCC-26012 catalyzed the efficient conversion of compound 70 to

126

R. N. PATEL

___T

H Ketone LQ

H. Potymorpha ATCC 26012

CHaOY\N 0 1 H Chlral Alcohol

FIG.27. Synthesis of chiral synthon for ~-(+)-sotalol.Stereoselective reduction of N-(4-(2-chloroacetyllphenyl)methanesulfonamide 70 (Pate1et al., 1 ~ 3 d )Reprinted . with permission from Springer-Verlag.

compound @ in 95"/0 reaction yield and >99% optical purity. Growth of 13. polymorpha ATCC-26012 culture was carried out in a 380-liter fermenter, and cells harvested from the fermenter were used to conduct transformation in a 3-liter preparative batch. Cell suspensions (20% wet cells in 3 liter of 10 mM potassium phosphate buffer, pH 7.0) were supplemented with 1 2 g of compound 70 and 2 2 5 g of glucose, and the reduction reaction was carried out at 25"C, 200 rpm, pH 7. Complete conversion of compound 211 to chiral alcohol SS was obtained in a 20-h reaction period. Using preparative HPLC, 8.2 g of compound @ were isolated from the reaction mixture in overall 68% yield with >99% optical purity. Both enantiomers of solketal (2,2-dimethyl-l,3-dioxolane-4-methanol) and their corresponding aldehydes are attractive building blocks for the preparation of enantiomerically pure and biologically active compounds (Hirth and Kindler, 1982; Peters et a]., 1987), specifically S-P-blocking agents. Since solketal is relatively inexpensive and commercially available, enantioselective oxidation of its alcohol function has provided an economically feasible process for the production of R-(-)-solketal by preferentially oxidizing the S-enantiomer to its corresponding acid (Bertola et al., 1987). Quino-haemoprotein ethanol dehy-

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

127

drogenase from Comamonas testosteroni has been used in the preparation of S-(+)-solketal (Geerlof et al., 1994). X. Potassium Channel Openers

The study of potassium (K) channel biochemistry, physiology, and medicinal chemistry has flourished, and numerous papers and reviews have been published (Edwards and Weston, 1990; Robertson and Steinberg, 1989,1990).It has long been known that K channels play a major role in neuronal excitability, and it is now clear that K channels play a critical role in the basic electrical and mechanical function of a wide variety of tissues, including smooth muscle, cardiac muscle, and glands (Hamilton and Weston, 1989). A new class of highly specific pharmacological compounds has been developed that either open or block K channels (Robertson and Steinberg, 1990; Ashwood et al., 1986). K channel openers are powerful smooth muscle relaxants with in vivo hypotensive and bronchodilator activity (Hamilton and Weston, 1989). The synthesis and antihypertensive activity of a series of novel K-channel openers (Bergmann et al., 1990; Jacobsen et al., 1991; Evans et al., 1983; Atwal et al., 1991) based on monosubstituted trans-4-amino-3,4dihydro-2,2-dirnethyl-2H-l-benzopyran-3-o1 71 have been demonstrated. Chiral epoxide 22 and diol 73 are potential intermediates for the synthesis of K channel activators, important as an antihypertensive and bronchodilator agents. The stereoselective microbial oxygenation of 2,2-dimethyl-2H-l-benzopyran-6-carbonitrile 74 to the corresponding chiral epoxide 2 and chiral d i o l B (Fig. 281 has been demonstrated (Pate1 et al., 1994b). Among microbial cultures evaluated, the best culture, Mortierella ramanniana SC-13840, gave reaction yields of 67.5 MY0 and optical purities of 96% for (+)-trans diol 73. A single-stage process (fermentation-epoxidation) for the biotransformation of 74 was developed using Mortierella ramanniana SC-13840. In a 25-liter fermenter, the (+)-transd i o l B was obtained in the reaction yield of 60.7 MY0 and optical purity of 92.5%. In the two-stage process using a %liter cell suspension (10% wt/vol, wet cells) of Mortierella ramanniana SC-13840, the (+)-trans diol 73 was obtained in 76 MY0 yield with an optical purity of 96%. The reaction was carried out in a 5-liter Bioflo fermenter with a 2 g/liter substrate and a 10 g/liter glucose concentration. Glucose was supplied to regenerate NADH required for this reaction. From the reaction mixture, (+)-transd i o l 3 was isolated in 65 M% 14.6 g) overall yield. An optical purity of 97% and a chemical purity of 98% were obtained for the isolated (+)-trans diol 3.

CH:,

Potassium Channel Opener 7r

FIG.28. Oxygenation of 2,2-dimethyl-ZH-l-benzopyran-6-carbonitrile 74 to the corresponding chiral epoxide 22 and (+ d i o l B by Morfierella ramanniana SC-13840 (Patel et al., 1994b). Reprinted with permission from Elsevier Science Ltd.

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

129

In an enzymatic resolution approach, chiral diol 73 was prepared by the stereoselective acetylation of racemic diol with lipases from Candida cylindraceae and Pseudomonas cepacia. Both enzymes catalyzed the acetylation of the undesired enantiomer of racemic diol to yield monoacetylated product and unreacted desired (+)-trans diol 73. A reaction yield of >40% and an optical purity >go%were obtained using each lipase (Pate1 et al., 1995).

XI. Antiinflammatory Drugs Naproxen, (S)-2-(6-methoxy-2-naphthyl)propanoic acid 75, is a nonsteroidal antiinflammatory and analgesic agent first developed by Syntex (Fried and Harrison, 1967; Harrison et al., 1970). Biologically active desired S-naproxen has been prepared by enantioselective hydrolysis of the methyl ester of naproxen by esterase derived from Bacillus subtilis Thai 1-8 (Giordano et al., 1992). The esterase was subsequently cloned in Escherichia coli with over an 800-fold increase in enzyme activity. The resolution of racemic naproxen amide and ketoprofen amides have been demonstrated by amidases from Rhodococcus erythropolis MP50 and Rhodococcus sp. C311 (Yamamoto et al., 1990; Gu et al., 1986b; Battistel et al., 1991; Layh et al., 1994). S-naproxen 75 and S-ketoprofen 76 (Fig. 29) were obtained in 40% yields (theoretical maximum 50%) and 97% e.e. The enantioselective esterification of naproxen has been demonstrated using lipase from Candida cylindraceae in isooctane as solvent and trimethylsilyl as alcohol. The undesired isomer of naproxen was esterified, leaving desired S-isomer unreacted (Tsai and Wei, 1994) Ibuprofen 22 is another well-known analgesic antiinflammatory drug, and it is believed that it will be marketed as a single-isomer drug. The kinetic enzymatic resolution of racemic ibuprofen has been reported (Trani et al., 1995). The reaction for resolution has been scaled up to make gram quantities of S-ibuprofen. This was accomplished by two enantioselective reactions each catalyzed by Novozyme 435. In the first reaction, 300 g of racemic ibuprofen were esterified with 1-dodecanol to yield the R-ester and S-ibuprofen to produce 89 g of S-ibuprofen in 85% enantiomeric excess. In the second reaction, 75 g of the 85% e.e. material were used to prepare 39 g of S-ibuprofen with a 97.5% e.e. Another approach for the enzymatic preparation of S-ibuprofen has been demonstrated by de Zoete et al. (1994). The enantioselective ammonolysis of ibuprofen-2-chloroethyl ester by Candida antarctica lipase (lipase SP435) gave the remaining ester S-(+)-enantiomerin 44% yield and 96% e.e.

130

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0

H

S-(-)-Ketoprofen26 FIG. 29. Structure of antiinflanlatury drugs naproxen 25, ketoprofcn

X ,and ibuprofen 22.

S-2-chloropropionic acid used as a chiral synthon €or pharmaceutical synthesis of various nonsteroidal antiinflammatory drugs has also been prepared by stereoselective dehalogenase reactions. Pseudomonas p u tida contained two dehalogenases, one a low-molecular-weight enzyme showing 100% specificity for S-Z-chloropropionate and the other enzyme with a higher molecular weight with specificity towards R-2-chloropropionate (Hardman and Slater, 1981; Hardman, 1991; Barth et al., 1992; Fetzner and Lingens, 1994). Future use of‘ R- and S-specific dehalogenases in enzymatic resolution processes will be very promising. XII. Antiviral Agents

Purine nucleoside analogues have been used as antiviral agents (Mansuri and Martin, 1987). Lamivudine, Zidovudine, and Didanosine are effective antiviral agents. Lamivudine 28, a highly promising drug candidate for HIV2 and HIV3 infection, provides a challenge to the synthetic chemist due to the presence of two acetal chiral centers, both sharing the same oxygen atom. The use of cytidine deaminase from Escherichia coli (Mahmoudian et a]., 1993) has been demonstrated to deaminate 2’-deoxy-3’-thiacytidine enantioselectively to prepare optically pure (2’-R-cis)-2’-deoxy-3’-thiacytidine (3TC, Lamivudine, Fig. 30).

131

SYNTHESIS OF SOME PHARMACEUTICAL INTERMEDIATES

-

Cytidine Deaminase

Racsmlc 24

Ha(>

(4-B 3TC 0

I

Didanosim (Vldex)

Zldovudins (Retrovir)

FIG.30. Preparation of (-)-lamivudine 28, an antiviral agent, by cytidine deaminase (Mahmoudian et al., 1993). Reprinted with permission from Elsevier Science Inc.

A novel enzymatic resolution process has been developed for the preparation of chiral intermediate for lamivudine synthesis. An enzymatic resolution of a-acetoxysulfides by Pseudomonas fluorescens lipase has been demonstrated to give a chiral intermediate in >45% yield and 97% optical purity (Milton et al., 1995). Toluene cis-dihydrodiols has been used as a synthetic building block for the preparation of a lactam that was converted into a new antiviral drug. cis-dihydrodiols are synthetic building blocks for pharmaceutical synthesis (Sheldrake, 1992). The enzymatic cis-dihydroxylation of aromatic compounds to give cis-dihydrodiols offers potential as raw materials for the preparation of chiral synthon needed for medicinal usage. Gibson et al. (1975) first demonstrated this reaction using benzene as a substrate. Subsequently, cis-dihydroxylation of numerous aromatic compounds was demonstrated using a bacterial system. In contrast to the bacterial system, a mammalian enzyme system produced trans-dihydrodiols as major products (Gibson and Subramanian, 1984). Imperial Chemical Industries has developed industrial processes to prepare kilogram quantities of arene-cis-dihydrodiols (Taylor, 1987).

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HO'

FIG. 31. Synthesis of chiral synthons for prostaglandins and leukotrienss. Stereoselective reduction of bicyclo[3.2.0lhept-2-en-6-one 81 and 7,7-dimethylbicyclo[3.2.O]l~ept-2-ene-6-one @ (Roberts, 1985).Reprinted with permission from Kluwer Academic Publishers.

XI1I. Prostaglandin Synthesis

Optically active epoxides are useful chiral synthons in the pharmaceutical synthesis of prostaglandins. Microbial epoxidation of olefinic compounds was first demonstrated by van der Linden (1963). Subsequently, May et al. (1975) demonstrated the epoxidation of alkenes in addition to hydroxylation of alkanes by an w-hydroxylase system. Oxidation of alk-1-enes in the range C6-Cl2, a,w-dienes from C6-C12, alkyl benzene, and ally1 ethers was demonstrated using an w-hydroxylase enzyme system from Pseudomonas oleovorans. R-epoxy compounds in greater than 75% e x . were produced by epoxidation reactions using the w-hydroxylase system (Abbott and Hou, 1973; May et al., 1975, de Smet et a]., 19831. The epoxidation system from Nocardia corallina is very versatile, has broad substrate specificity, and reacts with unfunctionalized aliphatics as well as aromatic olefins to produce R-epoxides (Takagi et al., 1990; Furuhashi, 1992).

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133

Chiral bicyclo[3.2.0]heptanone has been recognized as a chiral precursor for (+)-prostaglandinA2 79 and (+)-prostaglandin-F2a 80 synthesis. The reduction of racemic bicyclo[3.2.0]hept-2-en-6-one 81 in high optical purity by cells of Mortierella ramanniana has been demonstrated (Fig. 31). The same organism has been used for the stereoselective reduction of 7,7-dimethylbicyclo[3.2.0]hept-2-ene-6-one 82 to prepare the chiral synthon for (+)-leukotriene-B4 synthesis 83 (Roberts, 1985). Cycloalkonone oxygenase from Pseudomonas putida AS1 and Acinetobacter sp. NCIMB9871 has been used to catalyze the regio- and stereoselective Baeyer-Villiger type oxidation of [3.2.01hept-Z-en-6-one. The enantiomerically pure lactones prepared by this enzymatic reaction are chiral synthons for prostaglandin syntheses (Shipston et al., 1992; Lenn and Knowles, 1994). Asymmetric hydrolysis of diethyl-3-hydroxy-3-methylglutyrate to its corresponding monoester in high optical purity by pig liver esterase has been demonstrated (Haung et al., 1975; Chen et ~ l . 1981; , Gais and Lukas, 1984). Chiral monoesters are synthons for chemoenzymatic preparation of prostaglandins. REFERENCES

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Microbial Xylanolytic Enzyme System: Properties and Applications PRATIMA BAJPAI Chemical Engineering Division Thapar Corporate Research and Development Centre Patiala 147 001, India

I. Introduction 11. Structure of Xylan and Its Interaction with Plant Cell Walls 111. Properties of Xylanolytic Enzymes A. Xylanases B. 0-Xylosidases C. a-Arabinosidases D. a-Glucuronidases E. Esterases IV. Production of Xylanolytic Enzymes V. Application of Xylanases A. In Pulp and Paper Making 1. Prebleaching of Kraft Pulps 2. Enzymatic Debarking 3. Fiber Modification 4. Production of Dissolving Pulp 5. Removal of Shives 6. Retting of Flax Fibers B. Other Applications VI. Conclusions References

I. Introduction

Xylan is a major component of plant hemicellulose. After cellulose, it is the most abundant renewable polysaccharide in nature. Xylan and cellulose are the predominant hemicellulosic polysaccharides found in the cell walls of land plants, in which they may constitute more than 30% of the dry weight. Aside from terrestrial plants, in which xylans are based on a P-1,4-linked D-xylosyl backbone, marine algae also synthesize xylans of different chemical structure based on a P-1,3-linked D-xylosyl backbone. In some species of Chlorophyceae and the Rhodophyceae where cellulose is absent, xylans form a highly crystalline fibrillar material. 141 ADVANCES IN APPLIED MICROBIOLOGY. VOLUME 4 3 Copyright 0 1997 by Academic Preas, Inc. All rights of reproduction i n any form reserved. 0065-2164197 $25.00

142

P. BAJPAI

Xylanolytic enzymes of microorganisms have received a great deal of attention in the last 10 years. These enzyme systems are of interest for several reasons. On the one hand, they are clearly involved in providing sources of carbon and energy for the organisms that produce them. They act in the same fashion for hosts harboring xylanase-producing organisms, and they are involved in the growth, maturation, and ripening of cereals and fruits. Moreover, xylanases appear to be involved in the invasion of plants and fruits by pathogens. On the other hand, xylandegrading enzyme systems have considerable potential in several biotechnological applications. The present article reviews the properties and application of microbial xylanolytic enzyme systems. Attention is focused on recent advances, and particularly on several aspects that were not covered in earlier reviews.

I I . Structure of Xylan and Its Interaction with Plant Cell Walls Xylan structure is variable, ranging from linear 1,4-P-linked polyxylose chains to highly branched heteropolysaccharides. The prefix “hetero” denotes the presence of sugars other than D-xylose. Some major structural features are summarized in Fig. 1. The main chain of xylan is analogous to that of cellulose but composed of D-xylose instead of D-glucose. Branches consist of L-arabinofuranose linked to the 0-3 positions of D-xylose residues and of D-glucuronic acid or 4-0-methyl-D-ghcuronic acid linked to the 0-2 position. Both side-chain sugars are linked a-glycosidically. The degree of branching varies depending on the source. Xylans of several wood species, particularly of hardwoods, are acetylated: for example, birch xylan contain >1mol of acetic acid per 2 mol of D-xylose. Acetylation occurs more frequently at the 0-3 than the 0-2 position, and double acetylation of a D-XyloSe unit has also been reported. There is a relationship between the chemical structure of xylans and their botanical origin and also in their cytological localization. This results in a certain degree of complexity of xylan-containing materials that may possess several different xylan polymers of related structures but differ by more or less important features. Xylans of terrestrial plants are present in various proportions in the cell wall of all lignified tissues, but many may also be found in plant species as diverse as mosses and ferns (Aspinall, 1959; Joseleau, 1980). They usually are constituents of the secondary walls of tissues having structural functions, but they are also present to some extent in the primary walls of growing cells (Joseleau and Barnoud, 1974; McNeil et a]., 1979), as well as in the

143

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

H

H

H OH

H

H

H

H

H OH

COOH

A

OAC

0

n a y Hc@H

"C H3O @H

Araf

a

-

***

lo

Ac I* 3 3 4xylgl-4xyl~l-1xyl~1-~xylpI- Lxylbl-4xylfll-4xyl~1-4xyl~l- lxylgl-4Xylpl-

2

7 0 a 1

IC

Ac

Me G l c A

f10 U

Me GicA

endo-1,bp-xylanase (EC 3.2.1.8) P-xylosidase (EC 3.2.1.37) acetyl esterase (EC 3.1.1.6) or acetyl xylan esterase

0 a-glucuronidase (EC 3.2.1) c) a-L-arabinofuranosidase (EC 3.2.1.55) FIG.1. Structure of hardwood xylan and the site of attack by xylanolytic enzymes.

primary walls of seeds and bulbs (Shaw and Stephen, 1966a,b) of certain plant species in which they have reserve functions. In all the various ultrastructural localizations that they have in plant walls, xylans interact with other structural components, in particular with cellulose microfibrils, with other noncellulosic polymers, and in most cases with lignin. Noncovalent interactions of xylans with other polysaccharides essentially involve hydrogen bonding whereas covalent bonds interconnect xylans, lignins, and some phenolic acids. Although relatively little is known about the conformation of xylan chains within the cell wall lattice, conformational analysis of xylans has been carried out by crystallographic studies after purification of the isolated hemicelluloses. The development of techniques that permit the observation and analysis of polymers in their natural location in the cell wall-like

144

P. BAJPAI

electron microscopy, solid-state NMR, and Fourier transform infrared spectroscopy-are now being employed. Like most polysaccharides of plant origin, xylans display a large polydiversity and polymolecularity (Joseleau et al., 1992). This corresponds to their being present in a variety of plant species and to their distribution in several types of tissues and cells. All land plant xylans are characterized by a p-1,4-linked D-xylopyranosyl main chain that carries a variable number of neutral or uronic monosaccharide substituents or short oligosaccharide side chains. Very few unbranched linear xylan homopolysaccharides from land plants have been isolated. The best known is the xylan from esparto grass (Chanda et al., 1950), which, because of the peculiarity of its structure, served as a model for chemical and physical studies (Marchessault et a]., 1961). Other unsuhstituted linear xylans have been isolated from tobacco stalks (Eda et al., 1976) and guar seed husks (Montgomery et al., 1956). From the simple p-1,4-D-xylopyranosyl chain, the structural complexity of xylan rises with the number of substituting mono- and oligosaccharides attached to the P-1,4-linked xylosyl main chain. Timell reviewed the chemistry of hemicelluloses from angiosperms (Timell, 1964) and gymnosperms (Timell, 1965), and Wilkie described the structural characteristics of the different xylans from monocotyledonous species (Wilkie, 1979). The general features of all of these xylans from hardwoods, softwood, and grasses are given in Fig. 2. Much has been known for a rather long time about the chemical structure of alkali-soluble xylans. Most of the chemical structures reported in the literature were acquired by chemical, enzymatic, or spectroscopic analytical methods used separately or in combination. This led to description of the most characteristic features of polysaccharides belonging to the xylan family and corresponded to averaged structures with no or only little information about minor structural elements that were considered insignificant or due to impurities. The structural features of complex heteroxylans were obtained with more powerful investigative tools principally involving the use of the numerous glycohydrolases, esterases, and glycanases, of HPLC, and of 2D NMR spectroscopy. However, a lot less is known about their true primary structure, that is, if the arabinosyl, uronic acid, or acetyl substituents are attached to the xylosyl backbone randomly or as regular repeating sequences. The main structural elements commonly found in land plant cell wall xylans are shown in Table I. On the basis of the nature of the substituents, four main families can be considered where the complexity increases from linear to highly

( O IA c )

-

~

o

~

-

o

~

o

~

-

HOo

w0 o

~

L

~

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OR

OH

X

5z

R: a-~-GIcpA(1+2)Xyl ... 4-OMe-a-~-GlcpA(1+2)Xyl ... a-L-Araf(l+3)Xyl ... a-L-Araf(l+2)Xyl ... P-D-Galp(1+S)a-L-Araf(l+3)Xyl.. p-D-xylp(l-+2)a-~-Araf(l+3)XyI.. a-L-Araf(l-3-2, 1+3 and 1+2,3 Araf),(l-+3)Xyl Feruloyl

0

s2

n m

5

v)

...

p. coumaroyl Lignin FIG.2. Principal side-chain substitution on xylan backbone.

3 4 z

~

146

P. BAJPAI TABLE I PRINCIPAL STRUCTU TYPES ~ L FOUND IN THE XYLANFAMILY

Structural tY Pe

Nature of side chains

Source

Reference'

Linear homoxylan

= none

Esparto grass Tobacco stalk

Chanda et al., 1950 Eda et a]., 1976

Arahinoxy Ian Low branching

Terminal a-L-Araf

Common harhery monocots Primary walls Flours Gramineae pericarp

Henderson & Hay, 1972

High branching Complex side chain

a-L-arabinan oligomers

McNeil et al., 1979 Ewold & Perlin, 1959 Brillouet & Joseleau, 1987

Glucuronoxylan

CY-D-G~C~A a-4-0-Me-o-GlcpA

Soybean hull Hardwood Gramineae Legumes

Aspinall et a]., 1966 Timell, 1965 Wilkie, 1979 Reicher eta]., 1989

Glucuronoarabinoxylan

Terminal a - ~- Glc pA a-4-Me-o-GlcpA a-L-Araf &D-GalO

Softwoods Gramineae Dicot primary wall

Xmell, 1965 Wilkie, 1979 McNeil e t a ] . . 1979

T h e references cited here illustrate examples of typical xylans.

substituted xylans. A broad distinction may thus be made between the arabinoxylans having only side chains of single terminal units of a-Larabinofuranosyl substituents, the true glucuronoxylans in which a-Dglucuronic acid and/or its 4-0-methyl ether derivative represent the only substituent, and the more complex glucuronoarabinoxylan in which a-L-arabinose and a-D-glucuronic acid and 4-0-methyl-a-D-glucuronic acid are present at the same time. In addition to these three main families, one may distinguish arabinoxylans having a high degree of substitution by more or less short side chains of 2,3,5- and 2,3-linked arabinofuranosyl oligosaccharides attached to the 0-3 position of the xylosyl main chain, and galactoglucurono-arabinoxylans,characterized by the presence of terminal P-D-galactopyranosyl residues on complex oligosaccharide side chains of xylans from several perennial plants (Buchala and Meirer, 1972; Wilkie, 1979). Table I1 gives some of the most commonly encountered side chains that substitute heteroxylans of vari-

147

MICROBIAL XYLANOLYTIC ENZYME SYSTEM TABLE I1 VARIABILITY IN SIDE-CHAIN SUBSTITUTION I N THE HETEROXYLAN FAMILY ~

__

Side chains

~

~

Usual position of attachment to main chain

Source

References

Terminal single unit U-D-GICPA a-4-0-Me-D-GlcpA a-L-Araf

Angiosperms Gymnosperms

Timell, 1964 Timell, 1965

Corn-cob Barnhoo leaves

Dekker & Richard, 1975 Wilkie & Woo, 1977 Buchala & Meirer, 1972

-3

Wheat kernel

Brillouet

+2.+3,+2,3

Angiosperms

Karacsonyi et a]., 1983 Saavedra et ol.,1988 Reicher eta]., 1989 Kato & Nevins. 1985

-13

Complex oligosaccharidcs P-D-xylp(lj2)a-L-Araf P-D-Galp(l+5]a-L-Araf P-D-Galp(l-141-D-Xylp (+Z)-a-L-Araf 4-Me-a-D-GlcpA(1+4)-DXylp(1 +4)-D-Galp

-13 +3 +3

Arahinan side chains +Z)-L-Araf -+3)-L-Araf +5)-L-Araf +2,3)-~-Araf

1

&

Joseleau, 1987

Nonsaccharide side chains Acetyl

L-Araf Feruloyl

+BL-Araf

Monocots

ous organics. Substitution by nonsaccharide groups or molecules that are covalently attached to the xylan backbone or side chains are also listed in Table 11. Aside from the typical structures reported in Tables I and 11, several polymers having unusual characteristics have been reported. For example, an acidic xylan substituted only with a-D-glucuronosyl residues with the exclusion of the 4-0-methyl ether derivative was purified from the husk of a legume (Swamy and Salimaih, 1990). A neutral arabinoxylan containing an average structural unit corresponding to a xylosyl residue doubly substituted with a single arabinofuranosyl residue at the 0-3 position and two (1-3)linked L-arabinofuranosy1 residues at 0-2 was isolated from the bark of Litsea glutinosa

148

P. BAJPAI

(Herath et al., 1990). This polysaccharide had the very unusual feature of having both a - ~and - P-1,-arabinofuranosylresidues, as demonstrated by lH and 13CNMR and by a homonuclear lHlH 2D-COSY experiment. Similarly, a 2D heteronuclear correlated spectrum was the determinant for assigning signals of C-1 and H-1 of arabinosyl substituents attached to the same xylose residue of the backbone rather than to different xylose residues of the main chain (Ebrigerova et al., 1990). All of the structural data describing the ratio of substituents and their various positions on the main chain are insufficient for a complete description of the primary structure of xylan, since only little information is provided about the sequence of the distribution of the side chains along with the P-1,4-linked xylan backbone. Yet this information is needed for an understanding of the conformation of xylan in solution or in the cell walls and to understand their interaction with other cell wall polymers. Both the relative distribution of the side chains and the physical conformation of the polysaccharide are of great importance for the action of enzymes and may influence their mode of attack and hydrolysis yield. Only a few techniques and approaches provide information about the distribution of the side chains and substituents. In this area, specific degradation involving chemical reagents can provide useful results (Aspinall, 1982), but certainly the best tools are enzymes. There is a great diversity of available xylan-degrading enzymes, glycohydrolases, and glycanhydrolases that when used in conjunction with modern techniques for analysis of oligosaccharides (like HPLC, NMR and FAB mass spectroscopy) provide the best results for a detailed description of the true primary structures of xylans (Comtat and Joseleau, 1981; Debeirre e f al., 1990; Kovac et al., 1982). After exhaustive hydrolysis of the highly acidic arabinoxylan from Sequoia sempervirens (Dutton and Joseleau, 1977) by a purified fraction of endoxylanase, the analysis of the released oligosaccharides and of the undegraded polymer residue suggested that the uronic acid substituents were irregularly distributed on the xylosyl backbone (Comtat and Joseleau, 1981). The results also constituted evidence for the existence on this xylan chain of open hydrolyzable regions of unsubstituted or less substituted portions where endoxylanases could readily attack, and of nonhydrolyzable blocks, immune to endoxylanase attack because of the higher density of substituents. Similar results had been found for the distribution of uronic acid in the xylan from another softwood (Shimizu et al., 1978), whereas specific alkaline-catalyzed p-elimination demonstrated that glucuronic acid groups were randomly dispersed on the backbone of hardwood xylan (Rose11 and Svensson, 1975).

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

149

Ill. Properties of Xylanolytic Enzyme Systems

Due to the complex structure of xylans, several different enzymes are needed for their enzymatic degradation and modification. The two main glycanases that depolymerize the hemicellulose backbone are endo-l,4P-xylanase (EC 3.2.1.8) and endo-l&P-mannanase (EC 3.2.1.78). Small oligosaccharides are further hydrolyzed by 1,4-p-D-xylosidase, 1,4-p-Dmannosidase, and 1,4-P-D-glucosidase. The side groups are split off by a-L-arabinosidase, a-D-glucuronidase and a-D-galactosidase. Esterified side groups are liberated by acetyl xylan esterase and acetyl galactoglucomannan esterases. A. XYLANASES

Nonspecific xylanases from Trichoderma spp. can attack cellulose (Beldman et al., 1985), carboxymethylcellulose, p-nitrophenyl-p-glucoside (Beldman et al., 1985; Shikata and Nisizawa, 1975), cello-oligomers (Biely et al., 1991; Claeyssens et al., 1990; Shikata and Nisizawa, 1975), cellobiose (Beldman et al., 1985), laminarin (Lappalainen, 1986), and p-nitrophenyl-0-cellobioside (Shikata and Nisizawa, 1975). Carboxymethyl-cellulose, p-nitrophenyl-glucoside, and xylan apparently compete for the same active site on an enzyme from T viride that does not attack insoluble cellulose (Shikata and Nisizawa, 1975). A number of nonspecific glycanases have been characterized in Trichoderma spp. With one exception, they have relatively larger MWs (32-76 kDa) and more acidic PI values (3.5-5.3). This PI range excludes all of the apparently specific xylanases except for the 21-kDa xylanase from 'I: lignorum. These observations corroborate the hypothesis that there is a cluster of nonspecific glycanases with acidic PIS (Hrmova et al., 1986). The observation that these glycanases were induced by sophorose but not xylobiose suggests that they could be considered cellulases rather than xylanases. A multiplicity of xylanases has been documented in numerous organisms (Wong et al., 1988), with evidence for the occurrence of three to five xylanases in bacteria and fungi (Berenger et al., 1985; Fournier et al., 1985; Fredrick et al., 1981, 1985;John et al., 1979; Marui et al., 1985; Mitsuishi et al., 1987; Okhoshi et al., 1985; Shei et a]., 1985; Sreenath and Joseph, 1982; Takenishi and Tsujisaka, 1973, 1975; Tsujibo et al., 1990; Yoshioka et al., 1981). Recent analyses at the molecular genetic level have verified the occurrence of multiple xylanases in bacterial species (Flint et al., 1989; Gilbert et al., 1988; Mondou et al., 1986;

150

P. BAJPAI

Sakka et al., 1990; Vats-Mehta et al., 1990; Yang et al., 1989). Five xylanases have been detected in Trichoderma spp. (Dekker, 1983; Wong et al., 1986a) and three have been purified and characterized in T harzianum E58. The functional and genetic basis of these multiple enzymes has not been completely elucidated. Electrophoretically distinct xylanases may arise from posttranslational modifications of the same gene product, such as differential glycosylation or proteolysis. Trichoderma xylanases have been reported to be glycosylated in some cases (Lappalainen, 1986; Toda et al., 1971) but not in others (John and Schmidt, 1988; Ujiie et al., 1991; Wood and McCrae, 1986). The latter group of enzymes includes one pair of xylanases isolated from IT: koningii and another from 'I:lignorum. It therefore appears that differential glycosylation cannot explain the occurrence of multiple xylanases in these cases. Furthermore, a comparison of aminoacid compositions suggests that the three xylanases purified from IT: harzianum are distinct gene products (Wong et al., 1986a). The amino-acid composition of seven different Trichoderma xylanases has been reported. These data suggest high similarity between xylanase A from T harzianum and low-MW xylanases from ?: koningii and 7: viride. The other two xylanases from IT: harzianum appear to be distinct because of their relatively high alanine content. Furthermore, xylanase B is distinguished by a lack of tryptophan and a high cysteine content. These observations would suggest that there is a class of similar low-MW xylanases that occurs among Trichoderma spp. Amino-acid sequence comparisons have already suggested that xylanase A from T harzianum and a low-MW xylanase from T viride have over 90% homology (Roy et al., 1991; Yaguchi et al., 1992). Their similarity to corresponding enzymes from Bacillus pumilus, B. subtilis, and Schizophyllum commune is over 40% (Roy et al., 1991; Yaguchi et al., 1992). Preliminary X-ray diffraction analyses have been reported for xylanase A (Rose et al., 1987) and a 22.5-kDa xylanase from B. pumilus (Katsube et al., 1990; Moriyama et al., 1987). Completion of these studies would provide a means of directly comparing the structure of related xylanases from different sources. Amino-acid sequence comparisons of the catalytic domains of numerous glycanases have suggested that two classes of xylanase occur in microorganisms: the low-MW xylanases containing 182-234 aminoacid residues and the high-MW xylanases containing 269-809 residues (Gilkes et al., 1991). The low-MW xylanase class includes enzymes from B. pumilus, B. subtilis, and Clostridium acetobutylicum. Species of these two bacterial genera have most clearly illustrated the consistent occurrence of two forms of xylanases in one organism (Wong et al.,

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

151

1988). It is not known whether these two genetic units correspond to genetic units in Trichoderma spp. Historically, the functional division of xylanases has been related to the ability of certain xylanases to release arabinosyl substituents from arabinoxylan (Dekker, 1985). The work of Takenishi and Tsujisaka (1975) had shown a functional importance for the occurrence of an arabinose-releasing xylanase in Aspergillus niger. This enzyme was apparently responsible for the removal of arabinosyl substituents while another was responsible for the hydrolysis of xylotriose; the two together cooperated to increase the hydrolysis yield from arabinoxylan. However, such cooperation was not observed between two xylanases isolated from T. koningii: IM173022 (Wood and McCrae, 1986) or among xylanases from ?: harzianum E58 (Wong et al., 1986b),probably because their arabinose-releasing and xylotriose-cleaving activities were not complimentary. Nevertheless, the three cellulase-free xylanases from T. harzianum cooperated in the hydrolysis of aspen xylans. The degree of cooperativity apparently increased with increasing complexity of the substrate, from the form of the deacetylated polymer to that of the acetylated polymer and, most significantly, to the form of aspen holocellulose. Furthermore, the three xylanases isolated from Talaromyces byssochlarnydoides YH-50 also cooperated to maximize the hydrolysis of hardwood xylan (Yoshioka et al., 1981). The functional contributions of the individual xylanases have not yet been elucidated. Other characteristics proposed for classifying xylanases include the distinction between xylan and xylodextrin-hydrolyzing enzymes (Bailey and Gaillard, 1965), between enzymes with endo- and exo-type hydrolytic mechanisms (Reilly, 1981), and between enzymes with and without preference for the substitution sites on the xylan backbone (Fredrick et al., 1985; Nishitani and Nevins, 1991). The characterization of a 43-kDa xylanase from B. subtilis suggested that its hydrolysis sites are specifically oriented to the glucuronosyl rather than the arabinosyl substituents (Nishitani and Nevins, 1991). The optimal conditions for activity of Trichoderma xylanases range from 45 to 65OC and from pH 3.5 to 6.5. As might be expected, the xylanases with higher temperature optima are relatively more thermally stable (hie et al., 1990; Tan et al., 1985a; Wood and McCrae, 1986) than those with lower temperature optima (Hashimoto et a]., 1971; John and Schmidt, 1988; Tan et al., 1985a; Wood and McCrae, 1986). Two xylanases have been reported to be stable at 50°C for 1 h (hie et al., 1990; Tan et al., 1985a) and one at 60°C for 20 min (Wood and McCrae, 1986). These properties are relatively moderate when compared to xylanases isolated from thermophilic microorganisms. For example, a xylanase

152

P. BAJPAI

isolated from Thermostoga sp. strain FjSS3-B.1 has a temperature optimum of 105°C at pH 5.5 and a half-life of 90 min at 95°C (Simpson et al., 1991). Furthermore, alkaline-tolerant xylanases have been isolated from Bacillus spp. that have a broad range of pH optima and stabilities, ranging up to pH 10 (Honda et al., 1985b; Horikoshi and Atsukawa, 1973; Okazaki et al., 1985). Mercury ions have been found to be inhibitory to the activity of Trichoderma xylanase at concentrations ranging from 0.1 to 10 mM (Hashimoto et al., 1971; Huang et al., 1991: John and Schmidt, 1988). One exception is a partially purified xylanase from 7: viride that was not inhibited by 1 mM Hgz+(Gibson and McCleary, 1987). Of the other ions tested (Gibson and McCleary, 1987; Hashimoto et al., 1971; Huang et al., 1991; John and Schmidt, 1988; Tan et al., 1985b), 1mM Ca2+was found to be inhibitory in one case (Huang et al., 1991) and 1 mM Cu2+ in another (John and Schmidt, 1988). Trichoderma xylanases have been found to be active on xylans from different sources, usually producing xylooligomers (xylobiose and xylose). Xylose is not usually the major product, and it is typically produced after an accumulation of xylooligomers. Of the xylanases characterized, one isolated from T pseudokoningii (Baker et al., 1977) and two isolated from T. viride (Dean et al., 1991) were reported to be unable to produce xylose. Two nonspecific glycanases from T viride were also found to be unable to produce xylose during xylan hydrolysis (Beldman et a]., 1988; Shikata and Nisizawa, 1975). One of these glycanases produced xylobiose as an initial product, indicating that it acts like an exoxylanase (Shikata and Nisizawa, 1975). The hydrolysis patterns of Trichoderma xylanases, however, have suggested that most are endoxylanases. Xylans are not completely hydrolyzed by crude culture filtrates (Poutanen et al., 1987, 1990a) or purified xylanases from Trichoderma spp. (Poutanen and Puls, 1989; Wong et al., 1986b; Wood and McCrae, 1986). However, hydrolysis yields from certain xylans could be improved by using mixtures of three different xylanases purified from T harzianum (Wong et al., 1986b). Xylose yields obtained using a purified xylanase from T reesei were increased when a purified 0-xylosidase was added (Poutanen and Puls, 1989). They were further increased where the relevant debranching enzymes were added to the hydrolysis reaction. When acetylated xylooligomers were partially deactivated by freezedrying over ammonia, they became more accessible to hydrolysis by xylanase. All these observations suggest that the substituents on xylans can restrict their hydrolysis by xylanases.

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

153

The occurrence of debranching activities in xylanases appears to be variable. The four xylanases isolated from ?: reesei (Dekker, 1985) were reported to be arabinose-releasing enzymes as were the 18-kDa xylanases from 7: koningii and T viride. However, six other characterized Trichoderma xylanases lack this activity for removing the arabinosyl substituents from arabinoxylans. On the other hand, the removal of glucuronosyl residues is not an activity that has been associated with xylanases. Although this activity has seldom been examined in Trichoderma xylanases, it has been reported to be absent from the 20-kDa xylanase from T reesei (Poutanen and Puls, 1989),from the 18-kDa xylanase from T viride (Sinner and Dietrichs, 19761, and from a crude preparation of 20-kDa xylanase from T harzianum (Clark et al., 1990, 1991). This debranching activity has generally been attributed to other enzymes, namely, a-glucuronidases. There is, however, some evidence for the release of some acetic acid during the hydrolysis of an acetylxylan by a purified xylanase from ?: reesei (Poutanen and Puls, 1989). Xylotriose is the smallest oligomer hydrolyzed by most of the characterized xylanases. Of the three reported exceptions, the 20-kDa xylanase from ?: harzianum hydrolyzed xylotetrose using a pathway initiated by transglycosylation (Tan et al., 1985b). Transferase activity was also reported in the 20-kDa xylanase from 'I: lignorum (John and Schmidt, 1988)and in two nonspecific glycanases (Beldman et a]., 1988; Biely et al., 1991). This type of reaction may enable a nonspecific glycanase to split xylobiose after it has been incorporated into cello-oligomers (Biely et al., 1991).Transglycosylation also appears to be an important reaction in the hydrolysis of xylobiose by a P-xylosidase from T viride (Matsuo and Yasui, 1984). The activity of this xylobiase apparently decreases with increasing chain length of xylooligomers. However, other xylobiases have higher activity on xylooligomers than on xylobiose (Beldman et al., 1988; John and Schmidt, 1988) and can have substantial hydrolytic activity on xylans (Beldman et al., 1988). Trichoderma xylanases are known to solubilize carbohydrates from cell wall preparations, holocellulose substrates, and kraft pulps. Only 20% of the xylosyl residues from corn shoot cell walls and 10% of those from bean shoot cell walls were solubilized by a xylanase purified from T pseudokoningii (Baker et al., 1977). Much lower levels of other sugar residues were solubilized, suggesting the high degree of selectivity observed in certain Trichoderma xylanases (Clark et al., 1990, 1991; Senior et a!., 1988; Sinner et al., 1976, 1979; Tan et a].,1985b; Viikari et al., 1990; Wong et al., 198fia). Although xylose was not detected in

154

P. BAJPAI

the cell wall hydrolysates obtained using the 'T: pseudokoningii xylanase, it was found in holocellulose and pulp hydrolysates obtained using xylanases from 7: harzianum, 'T: reesei, and 'T: viride. These latter enzymes could solubilize 11-71% of the xylan in hardwood and softwood holocellulose (Sinner et a]., 1976, 1979), 9-25% of that in kraft pulps (Clark et a]., 1990; Senior et al., 1988; Viikari et a]., 1990), and 54% of that in a bleached hardwood h a f t pulp (Senior et al., 1988). There was a decrease in the degree of polymerization (DPJ of the xylan remaining in h a f t pulps after xylanase treatment (Miller et al., 1991). In beechwood holocellulose, the percentage of xylan solubilized by a xylanase from 'T: viride increased with decreasing particle size of the substrate (Sinner eta]., 1976). At the smallest particle size tested (0.070.1 mm), xylan accessibility could not be increased in beechwood holocellulose using a cellulase treatment (Sinner et al., 1976) or in spruce holocellulose using a mannanase treatment (Sinner et al., 1979). These observations suggest that xylan accessibility in solid substrates is not dependent on the other carbohydrates. In intact h a f t pulp fibers, however, xylan accessibility to a xylanase from Aspergillus foetidus could be increased using a cellulase (Puls et a]., 1990) and that to a xylanase from T harzianum using an extraction with sodium hydroxide (Clark et al., 1991). Xylan accessibility in solid substrates, therefore, appears to be dependent on the nature of these substrates. This is also illustrated by the unexpected observation that xylan accessibility increases in a series of radiata pine kraft pulps containing increasing levels of residual lignin, even though the percentage of xylan content decreases (Clark et al., 1990). In addition, the nature of solid substrates could have an inhibitory effect on xylanases from I: harzianum (Senior et a]., 1990, 1991). Furthermore, the hydrolytic properties of the enzymes is important because different xylanases from this fungus provided different hydrolysis yields from aspen holocellulose, with the maximum achieved using a mixture of all three enzymes (Wong et a]., 1986b).

B. P-XYLOSIDASES Exo-1,4-P-D-xylosidases (EC 3.2.1.37) hydrolyze xylooligosaccharides and xylobiose to xylose by removing successive D-xylose residues from the nonreducing termini. P-xylosidase is part of most microbial xylanolytic systems, but the highest extracellular production levels have been reported for fungi. P-xylosidases are rather large enzymes with molecular weights exceeding 100 kDa and are often reported to consist of two or more

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

155

subunits (Kersters-Hilderson et al., 1982; Matsuo and Yasui, 1984; Poutanen and Puls, 1988; Rodionova et al., 1983; Ujiie et al., 1985). Most purified P-xylosidases show highest activity toward xylobiose and no activity toward xylan. The activity toward xylooligosaccharides generally decreases rapidly with increasing chain length (Rodionova et al., 1983; Van Doorslaer et al., 1985). In addition to formation of xylose, many 0-xylosidases produce transfer products with higher molecular weights than that of the substrate (Conrad and Nothen, 1984; Rodionova et al., 1983). Some P-xylosidases have also been reported to possess P-glucosidase activity (Rodionova et a]., 1983; Ujiie et al., 1985). An important characteristic of P-xylosidases is their susceptibility to inhibition by xylose, which may significantly affect the yield under process conditions (Dekker, 1983; Poutanen and Puls, 1988; Rodionova et a]., 1983). P-xylosidase is the key enzyme for production of monomeric xylose from soluhilized xylan fragments, such as those obtained from a steaming process (Poutanen and Puls, 1988). They have been shown to act in synergy with suhstitutent-cleaving enzymes in the hydrolysis of suhstituted xylooligosaccharides (Poutanen, 1988a; Puls et al., 1987). The P-xylosidase of Trichoderma reesei was not able to hydrolyze xylobiose bearing an acetyl substituent at the nonreducing end without the presence of acetyl xylan esterase (Poutanen et d., 1990h). C. ~-ARABINOSIDASES

a-L-arabinofuranosidases(EC 3.2.1.55) hydrolyze nonreducing a-Larahinofuranosyl groups of arabinans, arabinoxylans, and arabinogalactans, as reviewed by Kaji (1984). The production of arabinosidases in microorganisms is often associated with the production of pectinolytic or hemicellulolytic enzymes, for example, in Corticiuni rolfsii (Kaji and Yoshihara, 1970), Sclerotina fructigena (Feilding and Byrde, 1969), T reesei (Poutanen et al., 1987; Poutanen, 1988b), and different Streptomyces species (Johnson et al., 1988a,h; Kaji et al., 1981). Some reported molecular characteristics of a-arabinosidases are presented in Table 111. The purified a-arabinosidase of Aspergillus niger (Tagava and Kaji, 1969), as well as that partially purified from a commercial pectinase preparation (Neukom et al., 1967), was able to release L-arahinose from wheat L-arabino-D-xylan. As the reaction proceeded, an amorphous precipitate consisting mainly of D-xylan with only traces of arahinose was formed. Adrewartha et 01. (1979) prepared a series of arabinoxylans

156

P. BAJPAI TABLE 111 CX-ARABINOSIDASES'

Molecular weight (kDa)

Microorganism

Aspergillus niger Trichoderma reesei Streptomyces spp. S trep tomyces p urp urescens Ruminococcus nlhus

53h

PI 3.6

53"

7.5

92'

4.4

495,h 62'

3.9

305,b 75'

6.8

"Sources: Poutanen et a].,1991;Kaji et al., 1969;Komae et al., 1982:Tagava and Kaji, 1988. 'Gel chromatography. 'SDS-PAGE.

from purified wheat-flour arabinoxylan by partial removal of arabinosyl side branches using an a-L-arabinosidase. They suggested that the solublizing effect of the arabinosyl substituents was not a result of increased hydration, but was due to their ability to prevent intermolecular aggregation of unsubstituted xylose residues. Cereal endospermic arabinoxylans especially are known to form viscous solutions and gels. It is obvious that suitable a-arabinosidases could be used to control the degree of substitution and hence the water-binding capacity of these pentosans. In a similar way, a-galactosidases have been used in adjusting the degree of a-galactosyl substitution and hence the gelling properties of galactomannans (Fujita and Nakamura, 1986; Overbeeke et al., 1987).

D. a-GLUCURONIDASES a-glucuronidases are required for hydrolysis of the a-l,2-glycosidic linkage between xylose and D-glucuronic acid or its 4-0-methyl ether. The presence of acidic oligosaccharides in xylan hydrolysates produced by hemicellulolytic enzyme preparations indicates the absence or inadequacy of this enzyme (Poutanen et al., 1987; Sinner and Dietrichs, 1976). 4-0-methylglucuronic acid was first detected in the enzymatic hydrolysates of glucuronoxylan by Sinner et al. (1972). The presence of a uronic acid-liberating enzyme, together with P-xylosidase, was

157

MICROBIAL XYLANOLYTIC ENZYME SYSTEM TABLE IV CX-GLIJCIJRONUIASES AND ITS SUBSTRATES’

Organisms

Substrate

A. bisporus T reesei

4-O-MeGlcAXz

S. olivochromogenes

Wheat bran

M.paranaguensis

Larchwoodxylan, wheat bran xylanase, p-nitorphenyl-uglucuronoside

S . olivochromogenes D. dendroides T palustris Trichoderma spp. l? versicolor L. sulphureus A. bisporus l? ostreatus Streptomyces spp. T reesei T aurantiacus

+ xylanase t

(4-0-MeGfcAJ-xylitol

Larchwoodxylan

+ xylanase

4-O-MeGlc AXz 4-0-MeGlcAX3

“Sources: Puls, 1992; Puls et al., 1986; Mackenzie et a]., 1987; Ishihara and Shimizu, 1988: Fontana ef ol., 1988.

claimed to increase the xylose yield in the enzymatic hydrolysis of hardwood xylan (Puls et al., 1976). The presence of a-glucuronidase in the hemicellulolytic system of T reesei was demonstrated in 1983 by Dekker. The production of a-glucuronidase by many fungi and bacteria (Table IV) has been reported (Puls, 1992). Only a few a-glucuronidases have been totally or even partially purified and characterized (Table V). The a-glucuronidase isolated from a culture filtrate of Agaricus bisporus by gel chromatography is a very large protein (450 kDa) (Puls et al., 1987). The enzyme had a very low isoelectric point and a pH optimum of about 3.3. A series of 4-0-methylglucuronosubstituted xylooligosaccharides with a DP up to 5 tested as substrates showed highest activity against 4-0-methylglucuronoxylobiose (Korte, 1980). The a-glucuronidase of A . bisporus had no activity toward polymeric xylan. The a-glucuronidase of ?: reesei also had an acidic isoelectric point (Poutanen, 1988a). It had a molecular weight of about 70 kDa as estimated by gel chromatography and a pH optimum at 6 with 4-0-methylgucurono-xylobiose as substrate. The a-glucuronidase of the thermo-

158

P. BAJPAI TABLE V PURIFIED CL-GLLJCURONIDASES" Molecular

PH

weight

Organism

Trichoderma reesei Rut C-30 Trichoderma reesei Thermoascus aurantiacus Agaricus bisporus

(kDa)

PI

optimum

>lo0 100

<4 nd nd

6.0 5.0 4.5 3.3-3.8

118 160

2.6-2.9

"Sources: Puls, 1992; Khandke ef nl., 1989. nd = not determined.

TABLE VI SUBSTRATE SPECIFICITY OF

CL-GI .SJCURONIDASESPRODUCED FROM A . BISPORUSa

AIJRANTIACUS AND

Degradation (%) Substrate

'I: aurantiacus

4-0-MeGlcAX 4-0-MeGlcAX2 4-0-MeGlcAX3 4-0-MeGlcAX4 4-0-MeGlcAX5 4-0-MeGIcAX6 4-0-MeGlcAXylan

96 100 96 96

A . bisporus 0 100 99

96

nd nd

50

52

0

53

"Source: Puls, 1992. nd = not determined.

philic fungus Thermoascus aurantiacus (Khandke et a]., 1989) was a single polypeptide chain with an MW of 118 kDa. The enzyme had a pH optimum at 4.5, and it hydrolyzed 4-0-methylglucurono-substituted xylooligomers from XI to X7 at rates comparable with that of xylan. The substrate specificities of T aurantiacus and A. bisporus a-glucuronidases are presented in Table VI (see Puls, 1992).

E. ESTERASES In addition to enzymes hydrolyzing the glycosidic linkages of xylans, the requirement of esterases to remove esterified acids from xylans has

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

159

been discovered. Acetyl xylan esterases remove 0-acetyl groups from the C-2 and C-3 positions of xylose residues in both xylan and xylooligomers. Feruloyl esterases liberate ferulic acid from arabinoxylans of monocotyledons. Esterases are classified according to their substrate specificity. This classification, however, is not very clear due to the rather wide substrate specificities of many esterases. Acetyl esterases (EC 3.1.1.6), a group of enzymes having highest activity against esters of acetic acid, are widely distributed in nature. They are produced by many animals, plants, and microorganisms (Krisch, 1971). The occurrence of microbial esterases acting on various synthetic acetyl derivatives of mono- and disaccharides was demonstrated in several early studies (Fink and Hay, 1969; Frohwein et al., 1963; Mandels and Reese, 1960; Reuter and Huttner, 1977), but the role of esterases in the hydrolysis of native acetylated xylans was emphasized only after 1980 (Biely et al., 1985; Poutanen et a]., 1986). Biely et al. (1985) first reported the presence of acetyl xylan esterases in fungal cellulolytic and hemicellulolytic systems: Trichoderma reesei, Aspergillus niger, Schizophyllum commune, and Aureobasidium pullulans. As compared with plant and animal esterases, these fungal esterases exhibited high specific activities toward acetylated glucuronoxylan and were therefore named acetylxylan esterases. The production of esterases that deacetylate xylan from a number of hemicellulolytic microorganisms has been reported (Table VII). Hitherto, only the acetyl xylan esterases of T reesei and A. oryzae have been purified and characterized (see Table VIII) (Poutanen and Puls, 1988; Sundberg and Poutanen, 1991). The T reesei enzymes had neutral isoelectric points but differed in their native molecular weights as assayed by gel chromatography. Both enzymes showed optimal activity at pH values between 5 and 6. One enzyme released only a little acetic acid from acetylated xylooligomers but showed high activity toward acetyl xylobiose. The other occurred as multiple isoenzymes and showed high activity toward acetylated xylan fragments and polymeric acetyl-4-0-methylglucuronoxylan (Poutanen et al., 1990b). Enzymatic deacetylation of beechwood xylan caused precipitation of the polymer. This was shown to be due to molecular aggregation analogous to the behavior of arabinoxylan after a-arabinosidase treatment. Feruloyl esterase activity was first detected in culture filtrates of Streptomyces olivochromogenes (Mackenzie et al., 1987) and has thereafter also been reported for some hemicelluloyltic fungi. A partially purified feruloyl esterase from S. commune liberated hardly any ferulic

160

P. BAJPAI

TABLE VII OCCURRENCE OF ESTERASES’

Esterase activity6

Microorganism

Butyrivibrio fibrisolvens S treptomyces fla vogriseu s Streptomyces olivochromogenes Streptomyces rubiginosus Aspergillus uwumori Aspergillus japonicus Aspergillus niger Aspergillus versicolor Aspergill us oryzae Fusarium oxysporum Schizopliyllitm commune Trichoderma viride Trichoderma reesei

Acetyl xylan esterase

Feruloyl esterase

+

nd

+

-

+, 3 different

+, 3 different

+, 5 different

+

+ + + + + +

nd nd

+, 1

+, 3 different +, 2 different

+ nd +, 1 purified nd +, 2 different nd -

types purified “Source: Poutanen et nl., 1991. = activity detected, - = no activity detected, nd = activity not determined

’+

acid without the presence of xylanase (Mackenzie and Bilous, 1988). Tenkanen et al. (1991) have purified a feruloyl esterase from Aspergillus oryzae. The enzyme is an acidic monomeric protein having an isoelectric point of 3.6 and a molecular weight of 30 kDa. It has wide substrate specificity, liberating ferulic, p-coumaric, and acetic acids from steamextracted wheat-straw arabinoxylan. The late discovery of acetyl xylan and feruloyl esterases has been partly due to the lack of suitable substrates. Xylans are often isolated by alkaline extraction, in which ester groups are saponified. Treatment of plant materials under mildly acidic conditions as in steaming or aqueous-phase thermomechanical treatment leaves most of the ester groups intact. These methods, however, partly hydrolyze xylan to shorter fragments (Khan et al., 1990; Puls et al., 1985). Polymeric acetylated xylan can be isolated from delignified materials by dimethyl sulfoxide extraction (Hagglund et al., 1956).

161

MICROBIAL XYLANOLYTIC ENZYME SYSTEM TABLE VIII PROPERTIES OF PURIFIED MICROUIAI. ESTERASES ACTING ON XYLANSU

PI MW" ( m a ) pH optimum pH stability" T stability" I"') Substrates

Acetyl esterase

Acetyl xylan esterase

6.6, 6.0 45 5.5 4-5 45

7.0, 6.8 34 5.O-6.0 3-7 60

Acetylated short-chain xylooligomers

Acetylated xylan

A. oryzae

Esterase 3.6 30 4.5-6.0 3-7 45

Phenolic and acetic acidsubstituted xylan

'Source: Tenkanen and Poutanen, 1992. "By SDS-PAGE. "The stability values represent the pH range and the highest temperature at which the xylanase retained full activity for 24 h.

The choice of substrate is especially important in studies of esterases for deacetylation of xylans. The use of small chromophoric substrates (p-nitrophenyl acetate, a-naphthyl acetate, and methylumbelliferyl acetate) analogously to the assays of disaccharides may lead to the monitoring of esterases unable to deacetylate xylan (Johnson et al., 1988b; Khan et al., 1990; Puls and Poutanen, 1989). It is obvious that some of the accessory enzymes are capable of releasing substituents from polymeric xylan and they consequently could be used to change the viscosity and solubility of xylans (Adrewartha et d., 1979; Khandke et d.,1989; Poutanen et al., 1990b). On the other hand, some substituent-cleaving enzymes have been reported to prefer short oligosaccharides as substrates (Mackenzie and Bilous, 1988; Poutanen, 1988b; Puls et al., 1987). In nature, however, all these enzymes usually occur in mixtures with other biomass-degrading enzymes and act synergistically with them to break down plant cell walls. Synergism between a-arabinosidase, xylanase, and P-xylosidase has been demonstrated in the hydrolysis of wheat-straw arabinoxylan with

162

P. BAJPAI

purified enzymes of T. reesei (Poutanen and Puls, 1989). When only xylanase and P-xylosidase were used in the hydrolysis, the xylose yield was only 66% of that produced by the whole culture filtrate at the same activity levels of these two enzymes and no arabinose was produced. Addition of a-arabinosidase increased the yields of both xylose and arabinose. Enhanced hydrolytic action of hemicellulolytic or pectinolytic enzymes in the hydrolysis of alfalfa cell wall polymers by addition of Ruminococus albus a-arabinosidase has also been reported (Greve et al., 1984). The synergistic action of depolymerizing and side-group-cleaving enzymes has most clearly been demonstrated using acetylated xylans as substrates. Due to the high degree of acetylation, xylanases have only limited access to the polymer backbone in the absence of esterases (Poutanen and Puls, 1989). Deacetylation by acetyl xylan esterase prior to the action of xylanases, however, resulted in a lower yield than that obtained by the simultaneous action of xylanase, P-xylosidase, and esterase (Poutanen et al., 1990b). The sequence of enzyme application not only influenced the extent of hydrolysis but also the nature of the oligomeric end-products. The production of acetyl xylan esterases by several cellulolytic and hemicellulytic fungi was first reported by Biely et al. (1985). These enzymes showed high specific activities toward acetylated glucuronoxylan as compared to acetyl esterases from plant and animal origin and were therefore named acetyl xylan esterases. Fractionation of T.reesei and T. viride culture filtrates revealed the multiplicity of esterases with different relative activities against acetylated xylan and 4-nitrophenyl acetate (Biely et al., 1987). The acetyl xylan esterases were thus also concluded to be at least partly different from more unspecific acetyl esterases. Several other fungi have been reported to produce acetyl xylan esterases (Johnson, 1990; Smith et al., 1991). Some bacteria, mainly different Streptomyces and some anaerobic rumen bacteria, have also been reported to secrete acetyl xylan esterases (Borneman et al., 1990; McDermid et al., 1990; Zimmermann, 1989). Xylan-deacetylating enzymes were also produced by the nonxylanolytic yeast Rhodot o r d a mucilaginosa (Lee et a]., 1987). The presence of an esterase acting on phenyl side groups in xylans was first detected in Streptomyces olivochromogenes culture filtrate (Mackenzie et al., 1987). This esterase hydrolyzed ester linkages of ferulic acid from the hemicellulose preparation of wheat bran. Feruloyl esterases have also been detected in the culture filtrates of other Streptomyces species, several Aspergillus species, S. commune, and

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

163

anaerobic rumen fungi and bacteria (Borneman et al., 1990; Johnson et al., 1988b, 1989; Mackenzie and Bilous, 1988; McDermid et al., 1990; Smith et al., 1991; Tenkanen et al., 1991). Esterases that liberate p-coumaroyl side groups have been produced by Streptomyces viridisporus and several mesophilic, thermophilic, and anaerobic rumen fungi (Borneman et al., 1990; Donnelly and Crawford, 1988; Smith et a]., 1991). IV. Production of Xylanolytic Enzymes

The cost of enzymes is one of the factors determining the economics of a biocatalytic process and can be reduced by finding optimum conditions for their production, by the isolation of hyper-producing mutants, and (possibly) by the construction of efficient producers using genetic engineering. A rational approach to these goals requires knowledge of the regulatory mechanisms governing enzyme production. Studies on the regulation of xylanolytic enzymes have largely focused on induction of enzyme activities under different conditions rather than on gene regulation. In addition, the studies have mainly focused on xylanase and xylosidase. Xylanolytic enzymes appear to be inducible. Xylanase and xylosidase are produced in high amounts during growth on xylan, and synthesis of the enzyme is catabolite repressed by easily metabolized carbon sources such as glucose or xylose. Xylan cannot enter the cells, so that the signal for accelerated synthesis of xylanolytic enzymes must involve lower-molecular-weight fragments, namely, xylobiose and xylotriose. The oligosaccharides are formed by hydrolysis of xylan in the medium by tiny amounts of enzymes produced constitutively. Induction can also be achieved by various synthetic alkyl and aryl P-D-xylosides in a Streptomyces sp. (Nakanishi et al., 1976) and by methyl P-D-xyloside in yeast (Biely and Petrakova, 1984a; Hrmova et al., 1984; Yasui et al., 1984). These compounds enable the production of xylanolytic enzymes in the absence of xylan and xylooligosaccharides. In the yeast C. albidus (Biely and Petrakova, 1984a), only methyl 0-D-xyloside induced the xylanolytic system. Other alkyl and aryl P-D-xylosides were unable to do so; they could induce a nonspecific P-glucosidase that hydrolyzed aryl P-D-xylosides but not xylooligosaccharides (Pecirova and Biely, 1982). This emphasizes the need to test xylosidase activity on natural substrates. Xylanolytic systems of yeasts can also be induced by positional isomers of xylobiose (Biely and Petrakova, 1984a; Hrmova et al., 1984).

164

P. BAJPAI

Induction with 1,Z-P-xylobiose is analogous to induction of cellulases in filamentous fungi by sophorose (1,2-P-glucobiose) (Zhu et al., 1982). However, the slow response of cells to 1,Z-P-xylobiose (relative to 1,4P-xylobiose) as well as the evidence for its transformation into 1,4-P-xylobiose (Biely and Petrakova, 1984b), the natural inducer, suggested that, in yeast, the isomeric disaccharide is a precursor of the natural inducer. The nature of the regulation in filamentous fungi has not been established. Generalization will perhaps never be possible due to the diversity of cellular control mechanisms. Fungal xylanases appear to be inducible or under derepression control, which includes enzyme production on carbon sources that are used slowly. Regulatory studies in fungi are often complicated by concurrent production of xylanase and cellulase, and by substrate cross-specificity of cellulases and xylanases. There are different types of cellulases and xylanases, the substrate specificities of which range from absolute for one polymer to about equal affinity for both polymers. For example, the cellulolytic systems of T reesei QM9414 contain two groups of glycanases exhibiting endoxylanase activity: specific endo-1,4-P-xylanases (one species with isoelectric point PI 4.8 and several with PI 8.5-9.0) and nonspecific endo-l,4-P-glucanases (PI 3.2-4.2) hydrolyzing both cellulose and xylan. The xylanolytic and cellulolytic systems in some filamentous fungi are likely to be under separate regulatory control. During growth on xylan, several species produce specific xylanases with little or no cellulase. However, when grown on cellulose, cellulases are produced together with xylanases (specific xylanases included) (Eriksson and Rzedowski, 1969; Gong et al., 1981; Honda et al., 1985a; Saddler and Mes-Hartee, 1983; Stewart et a]., 1983). The reason for production on cellulose of specific xylanases is unclear. Perhaps it results from the presence of xylan remnants in cellulose or depression on cellulose, a carbon source that is used slowly. Experiments with defined low-molecular-weight inducers in T reesei afforded similar results (Honda et al., 1985a): sophorose induced both specific and nonspecific endo-1,4P-glucanases, cellobiohydrolase I (exo-1,4-P-glucanase),and very little xylanase. Induction with xylobiose produced only specific xylanases. Therefore, the strategy for xylanolytic systems free of cellulases might simply be to grow cells on xylan uncontaminated by cellulose. However, this strategy could not be applied to all fungi. In Schizophyllurn conimune high xylanase production is linked strictly to cellulase production. The fungus grows poorly on xylan in the absence of

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

165

cellulose. The possibility of producing xylanolytic systems free of cellulase should be clarified in strains of Aspergillus, since they belong to the best xylanase and xylosidase producers. An alternative (and promising) approach to production of xylanolytic systems free of cellulases is the isolation of cellulase-deficient mutants, as outlined with Polyporus adustus (Eriksson and Goodell, 1974). Another possibility is the construction of appropriate recombinants by genetic engineering. It is noteworthy that xylan exerts a stimulatory effect on cellulase production by Streptomyces flavogriseus (Kluepfel and Ishaque, 1982), 'T: reesei (Honda et al., 1985a), and an acidophilic fungus (Cauchon and LeDuy, 1985). It remains to be seen whether this results from a specific effect of some xylan fragments or is simply a stimulation of cells by a carbon source more easily metabolized than cellulose. Recombinant DNA techniques offer opportunities for the construction of microbial strains with selected enzyme machinery. In xylan bioconversion, the main objectives for recombinant DNA technology are: (1) the construction of producers of xylanolytic systems free of cellulolytic enzymes, and (2) the improvement of the fermentation characteristics of industrially important xylose-fermenting organisms by introducing genes for xylanase and xylosidase so that the direct fermentation of xylan would be possible. All the cloning work published so far has been restricted to bacterial genes. Three groups have reported the isolation of xylanase genes from Bacillus species and their expression in Escherichia coli (Bernier et al., 1983; Honda et al., 1985a; Panbangred et al., 1983, 1985). In only one case (Honda et al., 1985a) the expressed enzyme appears to be secreted from the host cells. Further biochemical studies of xylanase-secreting and nonsecreting transformants could lead to better understanding of the secretory process and to the development of cloning strategies that would guarantee secretion of the desired products. Further difficulties can be expected with cloning genes from eukaryotic microorganisms. In addition to permitting the introduction of novel genes, cloning techniques could enable amplification of the expression genes already present. For instance, the production of xylanase in B. subtilis was enhanced successfully using a plasmid vector carrying the B. pumilus gene (Panbangred et al., 1985). The transformant produced approximately three times more extracellular xylanase than the donor strain. Moreover, the enzyme was produced constitutively, suggesting that regulatory elements of the donor organism were absent in the vector used for the transformation (Panbangred et al., 1985). A gene for intracellular xylosidase of B. pumilus has also been cloned.

166

P. BAJPAI

Detailed studies on the production and regulation of acetyl and aryl esterases acting on xylans have not been carried out. Production of these esterases can be enhanced by using different lignocellulosic materials as inducers (Biely et al., 1988; Khan et al., 1990). These experiments have mainly been carried out using cellulose, xylan or wheat bran as growth supplements. However, in most studies commercial alkali-extracted xylans have been used instead of natural esterified preparations. The microorganisms capable of secreting high levels of acetyl xylan esterase activity (>300 nkat/ml) include 7: reesei, S. commune, and several Streptomyces species. However, activities reported by different authors can be only approximately compared due to the different conditions and substrates used in activity assays. Biely et al. (1988) reported that acetyl xylan esterases were co-produced with endoglucanase and xylanase by T reesei and S. commune. The production of acetyl xylan esterases by these microorganisms was somewhat increased when the deacetylated xylan in the growth medium was replaced by acetylated xylooligomers. However, supplementation with cellulose increased the production even more. By contrast, the production of acetyl xylan esterases by different Aspergillus and Streptomyces species was enhanced more by the presence of xylan than by cellulose in the growth medium (Khan et al., 1990; Mackenzie et a]., 1987). The highest level of feruloyl esterase activity was produced by S. olivochromogenes and S. commune, analogously to the production of acetyl xylan esterases on the medium supplemented by xylan or cellulose, respectively (Mackenzie and Bilous, 1988; Mackenzie et al., 1987). A . niger is reported to produce almost tenfold activity levels of feruloyl esterase as compared with S. olivochromogenes or S. commune (Johnson et al., 1989). The ferulic acid content of the lignocellulosic material added to the cultivation medium correlated with the feruloyl esterase activity levels produced by anaerobic rumen fungi but not in those produced by several strains of Aspergillus (Borneman et al., 1990; Tenkanen et al., 1991). a-glucuronidase was found in culture filtrates of Agaricus bisporus, Trichoderma reesei, and Aspergillus awamori. The substrate used was 4-O-Me-Glc AXZ. a-glucuronidase was also reported in culture filtrates of Pleurotus astreatus (Puls et al., 1987), Streptomyces olivochromogenes (Mackenzie et al., 1987), Dactylium dendroides (Fontana et al., 1988), Tyromyces palustris, Polyporus versicolor, Laetiporus sulphureus (Ishihara and Shimizu, 1988), and Thermoascus aurantacus (Khandke et al., 1989). Agaricus bisporus was found to be the most powerful a-glucuronidase producer on the basis of specific enzyme

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

167

activity, using 4-0-MeGlcAX2 as a substrate for screening (Puls et al., 1987). In another screening of a-glucuronidase-producing fungi including Agaricus bisporus, (4-O-MeGlcA)-xylitolwas used as a substrate and Tyromyces polustris was found to be the most potent enzyme producer (Ishihara and Shimizu, 1988). However, from another study it became apparent that the analogous 4-O-MeGlc AX substrate was not degraded by the Agaricus bisporus enzymes. Aside from the different substrates for testing, a variety of carbon sources were used for induction of a-glucuronidase activity, including aqueous extracts from birchwood, obtained after steaming pretreatment (Korte et al., 1991), oat spells xylan or wheat bran (Mackenzie et al., 1987), cellulose, and brewer’s grain (Poutanen, 1988a). V. Application of Xylanases

A. IN PULP AND PAPER MAKING Potential biotechnological applications in the pulp and paper industry have been reviewed by several authors (Trotter, 1990; Eriksson, 1990; Viikari et al., 1994). Currently, the most promising application for xylanase is in the prebleaching of kraft pulps. Other applications proposed for xylanases include debarking, modification of fiber properties for improving drainage and beatability, preparation of dissolving pulp, and shive removal.

Prebleaching of Krafl Pulps The use of xylanase enzymes in bleaching can be traced from the work of the late 1970s to the early 1980s, a period of rapid growth for biotechnology programs worldwide. Much of the attention was placed on the conversion of waste biomass (e.g., decaying woodstocks, municipal waste, and food processing and pulp mill wastes) to value-added chemicals and fuels. Such conversions require that the biomass be hydrolyzed into component sugars prior to fermentation, for example, to alcohol. To accomplish hydrolysis, researchers applied living cultures of wood-inhabiting fungi and bacteria. In nature, these microbes secrete enzymes that decay biomass, thus providing soluble sugars for their growth. Screening programs provided microbial culture collections with dozens of organisms capable of secreting hydrolytic enzymes that could degrade biomass. Wood-inhabiting microbes produce a variety of enzymes that degrade all components of wood. These enzymes include ligninases, hemicellulases, and cellulases. Ligninases attack and solubilize lignin, and cellu1.

168

P. BAJPAI

lases and hemicellulases degrade cellulose and hemicellulose, respectively. The early 1980s saw attempts to separate and isolate the enzymes produced by these microbes. One type of hemicellulase that has been isolated is called xylanase. Xylanases are xylan-degrading enzymes that are also secreted in large quantities by many of these microbes and are the first enzymes involved in the decay of wood. In 1984, Paice and Jurasek at the Pulp and Paper Research Institute of Canada (PAPRICAN) attempted to prepare kraft pulp suitable for rayon manufacture by applying a crude mixture of cellulases and xylanases to remove undesirable xylan (Paice and Jurasek, 1984). In 1986, Finnish workers at VTT treated kraft pulps with a similar enzyme preparation and looked at the impact of the treatment on bleaching. They found that a significant reduction in bleaching chemicals could be achieved while maintaining high brightness (Jurasek and Paice, 1992; Viikari et a]., 1987). Thus, biobleaching was born. Soon thereafter, the PAPRICAN group demonstrated that the xylanases were predominantly responsible for the biobleaching effect by applying highly purified xylanases to pulps. Since then, various papers have described the benefits of xylanase treatments in pulp bleaching. Mill trials began as early as 1989 in Finland, and trials continue to be run with remarkable frequency. Since 1991, commercial use of xylanase has become a reality. As reported by Jurasek and Paice (1992) at the International Symposium on Pollution Prevention in the Manufacture of Pulp and Paper, 10 mills were said to use xylanase prebleaching on a commercial basis and more than 80 mill trials were carried out. In 1993, the process gained even wider acceptance especially in Canada and Europe (Lavielle, 1992). The factors explaining this rapid development are many but can be summarized as follows: 0 Xylanase prebleaching belongs to the soft technologies that require no or very little capital investment to operate. Process changes are minimal in most cases (neutralization of brown stock). Mill trials are very simple, inexpensive, and involve minimal risk. Xylanase helps reduce pollution from bleaching. 0 Savings on chemicals can pay for the process. Xylanase may help to increase mill capacity where there are chlorine dioxide limitations. 0 The process is easily combined with many bleaching sequences for ECF and TCF pulps.

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

169

In enzymatic pretreatment for bleaching, the hydrolysis of hemicellulose is restricted to a minimum by using only small amounts of enzymes in order to maintain a high pulp yield and the advantageous properties of hemicellulose in pulp (Viikari et al., 1986, 1987). Hemicellulose in pulp plays an important role both in fiber morphology and fiber physics. Retention of hemicellulose increases the pulp yield, improves pulp strength, and affects fiber quality.

Production of Xylanases for Bleaching. Several criteria are essential for choosing a microorganism to produce xylanases. In addition to giving the desired biobleaching effect, enzyme preparation must yield a sufficiently large quantity and the xylanase technology must be compatible with the technology of a pulp mill. Also, it is essential that enzyme preparation be completely free of cellulase side activity. Any cellulase activity will have serious economic implications in terms of cellulose loss, degraded pulp quality, and increased effluent treatment cost. Noncellulolytic preparations have been produced by genetic engineering, selective inactivation, or bulk scale purification (Barnard et al., 1986; Pedersen et al., 1992). High productivity has been achieved by exhaustive screening, genetic engineering, and growth optimization programs. To produce xylanases, the selected organism is grown for several days in fermentation vessels containing nutrients and oxygen under specific conditions of pH, temperature, and agitation. During this time, it secretes enzymes into the growth medium. The living cell mass is then removed, leaving a xylanase-rich liquid. This is then concentrated, assayed to determine its activity, and packaged for shipment to pulp mills. With the addition of bacteriostatic preservatives, the xylanase preparation remains stable for months. Excessive temperature and freezing can cause loss of activity and should be avoided. Xylanase preparation is not corrosive nor reactive and does not require resistant materials for handling. Numerous studies have been published on the production and properties of hemicellulases, especially xylanases. The productivity has been increased both by development of more efficient production strains and by optimizing the production methods (Baile et al., 1993; Nevalainen et al., 1990; Suominen et al., 1992). Xylanases have also been cloned in new noncellulolytic production hosts (Bertrand et al., 1989). Several xylanases, although only a few of them purified, have been tested for their properties in bleaching experiments with various testing methods (Viikari et al., 1994). The strains reported to be used for commercial production of xylanases include Trichoderma reesei, Ther-

170

P. BAJPAI

momyces lanuginosus, Aureobasidium pullulans, and Streptomyces lividans (Bajpai and Bajpai, 1996a; Jager et al., 1992; Senior et al., 1992b). In practical process conditions, such properties of the enzymes as substrate specificity and the pH and temperature optima are of utmost importance. Enzymes with high pH and temperature optima have been isolated and tested for improving the bleachability of pulps (Davis et al., 1992; Hogman et al., 1992; Jager et a]., 1992; Pedersen e t d., 1991; Senior et al., 1992b). Several alkali-tolerant strains of Bacillus sp. have been used for production of xylanases with pH optima around 9. The most thermostable xylanases, with a half-life of 90 min at 9SoC, have been produced by a Thermotoga strain (Simpson et ol., 1991). The pH range has also been increased up to about 8 and the temperature range up to 75°C in some commercial preparations. Table IX presents a list of some of the commercial xylanases used for prebleaching. The cost of the enzyme preparations is in the range of $2-7 per ton of pulp processed depending on specific bleaching conditions. Xylanase Treatment in Mills. Xylanase is typically added to pulp as an aqueous solution at the final brown stock washer. Because the enzymes are extremely potent catalysts, the desired effects are produced with small amounts of enzymes. The brown stock, which (though washed) is highly alkaline (pH 9-12), must be neutralized with acid (usually sulfuric), to be compatible with enzyme treatment. The pulp is pumped to the high-density storage tower, where the enzyme acts. From the high-density storage tower, the enzyme-treated pulp is then pumped into the first bleaching tower, where first contact with the oxidizing chemicals destroys the enzyme. Unlike other bleaching chemicals, xylanases do not brighten or delignify the pulp. They modify the pulp to make the lignin more accessible to removal by other bleaching chemicals. The enzyme-treated pulp then passes through the bleach plant with decreased chemical requirement for bleaching. Factors Affecting Treatment Efficiency. The major factors affecting the xylanase treatment efficiency include pH, temperature, enzyme dosage and dispersion, consistency, and reaction time. The optimum pH for xylanase treatment varies among enzymes. Generally, the xylanases derived from strains of bacterial origin are most effective between pH 6 and 9, while those derived from strains of fungal origin should be used within the pH range of 4-6. The optimum temperature ranges from 35 to 60°C. To obtain the best results from enzyme use, enzyme dosage must be optimized in each case. In general, the optimal dosage lies

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

171

TABLE IX COMMERCIAL XYLANASES Product Cartazyme HS Cartazyme HT Irgazyme 40-4X Albazyme 40-4X Irgazyme-lo/ Albazyme 10 VAI Xylanase Pulpzyme HA Pulpzyme HB Pulpzyme HC Bleachzyme F Ecopulp X-200 Ecopulp XM Xylanase Xylanase I.-8000 Ecozyme

Supplier Sandoz Chemicals, UK Sandoz Chemicals, UK Genencor, Finland Ciba Geigy, Switzerland Genencor, Finland/ Ciba Geigy, Switzerland Voest Alpine, Austria Nova Nordisk, Denmark Novo Nordisk, Denmark Novo Nordisk, Denmark Biocon India, Bangalore Alko Ltd. Biotechnology, Finland Alko Ltd. Biotechnology, Finland Iogen Corporation, Canada Solvay Interox, USA Zeneca BioproductdICI, Canada

within the range of 2 to 5 international units (IU) of enzyme per gram of dry pulp. In addition, pulp consistency must be optimized to obtain effective dispersion of the enzyme used to improve the efficiency of enzyme treatment. Screw conveyors and static mixers are examples of efficient mixing systems (Skerker et al., 1992). Most of the beneficial effect of the bleaching can be obtained after only 1 h of treatment, although the enzyme effect does slowly continue throughout an 18-h treatment period. Usually, the reaction time is set at 2 h. However, long retention times must be avoided if cellulases are present. Bajpai and Bajpai (1996b) found that pulp washing after enzyme treatment was not necessary to improve bleaching efficiency. Current research efforts in the area of xylanase production have been focused on developing new xylanase preparations that are able to work at very high temperatures and pH values, permitting xylanase to be used effectively under various conditions and at different stages of the bleaching process.

Effect of Xylanase Treatment on Pulp Bleaching. Xylanase treatment has been shown to reduce the requirement of chlorine for bleaching while still achieving high brightness and good pulp properties. Results

172

P. BAJPAI

from laboratory studies and mill trials show about 3 5 4 1YO reduction in active chlorine at the chlorination stage for hardwoods and lo-20% for softwoods, whereas savings in total active chlorine were found to be 20-250!0 for hardwoods and 10-15% for softwoods (Buchert et al., 1992; Clark et al., 1991; Perrolaz et a]., 1991; Sinner et a]., 1991; Skerker and Farell, 1991; Viikari et al., 1986, 1987, 1994). In the elementary chlorine-free bleaching sequences, the use of enzymes increases the productivity of the bleaching plant when the production capacity of CIOz is a limiting factor. This is often the case when the utilization of chlorine gas has been abandoned. In totally chlorine-free bleaching sequences, the addition of enzymes increases the final brightness value, which is a key parameter in marketing of chlorine-free pulps. In addition, savings in TCF bleaching are important with respect both to costs and to the strength properties of the pulp. The results of some published mill trials are presented in Table X. The production of TCF pulp has increased dramatically during the past 2 years. Several alternative new bleaching techniques have been developed. TCF bleaching methods are based on different oxygen chemicals like oxygen, ozone, and peroxide. An oxygen stage has already been installed at many haft mills. Enzymes are frequently being used on oxygen-delignified pulps to increase brightness in both laboratory and mill scale trials. Enzymes have also been combined with ozone bleaching (Yang and Eriksson, 1992). The TCF technologies applied today are usually based on bleaching of oxygen-delignified pulps with enzymes and hydrogen peroxide. Xylanase pretreatment has led to reductions in effluent AOX and dioxin concentrations due to reduced chlorine requirement to achieve a given brightness (Senior and Hamilton, 1991; Senior and Hamilton, 1992; Vaheri et a]., 1989). The level of AOX in effluent was significantly lower for xylanase-pretreated pulps as compared to conventionally bleached control pulps (Tables XI-XIV). When softwood h a f t pulp pretreated with xylanase was bleached to 90% IS0 using a (CD)EpDED sequence, the required kappa factor was reduced from 0.22 to 0.15, which was below the kappa factor of 0.18-0.19 required for the formation of chlorinated dioxins and furans (Berry et al., 1989). The AOX concentration in a combined effluent stream was reduced by 33% compared to the control (Senior and Hamilton, 1992).The effluent BOD doubled, and there were increases in effluent COD and total organic carbon (TOC). The BOD/COD ratio also increased, indicating that the effluent was more amenable to biological degradation in a secondary treatment plant. Effluent toxicity remained essentially the same (Table XIII). In the same study, xylanase pretreatment of a hardwood h a f t pulp

TABLE X MILLSCALE TRIALRESULTSw m COMMERCIAL XYLANASES"

Mill Enso Gutzeit, Finland Metsa Botnia, Finland Canfor's International Mill Brit. Col., Canada Tasman Pulp and Paper Company, New Zealand Bukoza Pulp Mill. Czechoslovakia Scott Paper Mill, Spain Munksjo, Sweden Crest Brook Forest Industries, Canada

Bleaching sequence

Reduct ion in chemical consumption

AOX reduction

(%I

Brightness increase (%ISO)

Pulp properties

ze

29

Unchanged

M

nd

Unchanged

F

nd

Unchanged

x DEopDED

15.6% ClO,

i z 0

2 (DC)EoDED

20% ClOZ

20

Unchanged

(CDIEDEHD

30% c12

nd

Unchanged

=!

n

8 30% hypochlorite

EopDEPD OQElPEzP DEopDED

7-17% reduction

4-6

-

2-4

nd

-

No attack on cellulose Unchanged Unchanged

$

m

;c,;;3

z

in kappa factor, increased productivity CI

"Source: Bajpai and Bajpai, 1996a. nd = not determined.

v W

174

P. BAJPAI TABLE XI EFFECT OF XYLANASE TREATMENT ON EFFLUENT PROPERTIES' Control

Enzyme

Pretreatinent PH Xylanase charge (unit/kg pulp) Time (h) Temperature ("C) Kappa number Viscosity (mPa s)

32 25.1

3 37 31.3 30.7

( C d d stage Clz (kg/mt pulp) ClOz (kg/mt pulp) Artive CI, (kg/mt pulp)

19.2 17.0 64.0

15.0 13.3 49.9

35.2 5.0

27.0 5.0

4.8

4.4

2.02 155 15 74

1.68 118

Yes 4.8 5000

-

( E d stage NaOH (kg/mt pulp) 0 2 (kg/mt pulp) E" kappa number Effluent properties AOX (kg/mt pulp) Color (kglmt pulp] BOD (kg/mt pulp) COD (kg/mt pulp)

11

62

"Source: Viikari et ol., 1986. Unlr1eai:Iietl kappa no. 32.0; viscosity 30.7 mPa . s

TABLE XI1 EFFECT0 1 7 XYLANASE TREATMENT ON THE AOX CONTENT OF (C8"D2,)EpDEDEFFLUENT" Chlorine charge (kg/mt pulp)

AOX (kglmt pulp)

Brightness (YOISO)

Control

52 63

4.35 6.00

91.2

Xylanase treated

43 52 57

3.00 3.75 4.50

91.0

Pulp

TABLE XI11 EFFLUENTPROPERTIES OF SOFTWOOD PULPFOLLOWING XYLANASE TREATMENT^ ~

~~~~~

Control Parameter Kappa factor C-stage residual (% on pulp) (CD)EpDEDbrightness (Yo ISO) BOD (mg/liter) COD (mg/liter) BODlCOD TOC (mglliter) Toxicity (YO solution) AOX (mg/liter)

Xylanase treated

WICDIEp

W(m)Ep

X(CD)Ep

X(CD)Ep

X(O)EP

0.15

0.22

0.10

0.15

0.18

-

0.04

-

-

0.32

88.1

92.8

85.8

91.6

94.1

W control stage

X xylanase stage

51

50

102

113

120

4 0

78

600

800

756

948

926

58

370

0.085

0.063

0.14

0.12

0.13

0.86

0.21

503 33.2

668 24.3

585 53.9

787 25.6

752 22.7

54 100

291 100

42.7

65.1

22.7

43.5

68.4

-

-

%ource: Senior and Hamilton, 1992. Substitution of chlorine dioxide in chlorination stage = 20%; kappa no. of control pulp = 28.3; kappa no. of xylanase treated pulp = 28.3; effluent determinationsmade on combined W or X((=D)Ep stage.

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P. BAJPAI

under the same conditions led to a reduction of 35-40% chlorine in chlorination charge (Table XIV). The AOX level in El stage effluent was 24% less than in the control, and the BOD/COD ratio was increased. The organochlorine content of the pulp was found to be reduced by 41% at a chlorine dioxide substitution level of 40% (Senior and Hamilton, 1992). Bajpai and co-workers (Bajpai and Bajpai, 1996b; Bajpai et al. 1993; Bajpai et al., 1994) reported that the total organochlorine (TOC1) content in the extraction stage effluent was reduced by 30% when the pulp was first pretreated with xylanase and then subjected to CEH bleaching. The enzyme-treated pulps show unchanged or improved strength properties (Bajpai and Bajpai, 1996b; Bajpai et al., 1994; Dunlop and Gronberg, 1994; Viikari et al., 1986). Also, these pulps are easier to refine than the reference pulps. Improved viscosity of the pulp has been noted as a result of xylanase treatment (Bajpai and Bajpai, 1996b; Bajpai et al., 1993, 1994; Senior et al., 1992a; Yang et al., 1992). This is probably caused by the selective removal of xylan as determined by the pentosan values. Xylan with lower DP than cellulose can be expected to lower the average viscosity of h a f t hemicellulose. However, the viscosity of the pulp was adversely affected when cellulase activity was present (Allison and Clark, 1992; Bajpai et al., 1993; Clark et al., 1990; Puls et al., 1990). Therefore, the presence of cellulase activity in the enzyme preparation is not desirable, In a few cases, lower mechanical strength has been obtained on xylanase-treated pulp, probably due to the presence of cellulase in the enzyme preparation (Chauvet et al., 1987).

Benefits from Xylanase Treatment. Xylanase pretreatment of pulps prior to bleaching reduces bleach chemical requirements and permits higher brightness to be reached. A reduction in chemical charges can translate into significant cost savings when high levels of chlorine dioxide and hydrogen peroxide are being used. Reduction in the use of chlorine chemicals clearly reduces the formation and release of chlorinated organic compounds in the effluent and in the pulps. The ability of xylanases to activate pulps and increase the effectiveness of bleaching chemicals may allow new bleaching technologies to become more effective. This means that, for expensive chlorine-free alternatives such as ozone and hydrogen peroxide, xylanase pretreatment may eventually permit them to become cost-effective. Traditional bleaching technologies also serve to benefit from xylanase treatments. Xylanases are easily applied and require essentially no capital expenditure. Because chlorine dioxide charges can be reduced,

TABLE XIV CHARACTERISTICS OF ElSTAGE EFFLUENT FROM HARDWOOD PULP' Control Parameter Kappa factor (DC)E1DE2D brightness [%IS01 BOD (%/liter) COD (mg/liter) BODlCOD TOC (mg/liter)

Toxicity [% solution) AOX [mg/liter)

W(DC)E

Xylanase treated

W(DC)E

X(DC)E

X(C+D)E

X(CD)E

0.15

0.20

0.10

0.10

0.10

89.3

90.0

89.5

89.1

89.3

261

259

212

283

280

1100

1160

956

906

988

0.24

0.22

0.22

0.31

0.28

429

4 64

385

379

402

8.9

8.8

4.9

6.6

4.8

17.6

25.3

13.3

7.5

7.5

'?Source: Senior and Hamilton, 1992. Substitution of chlorine dioxide in chlorination stage = 40%; kappa no. of control pulp = 12.2;kappa no. ofxylanase treated pulp = 11.6.

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P. BAJF’AI

xylanase may help eliminate the need for increased chlorine dioxide generation capacity. Similarly, installation of expensive oxygen delignification facilities may be avoided. The xylanase bleach-boosting stage can also shift the degree of substitution toward higher chlorine dioxide levels while maintaining the total dosage of active chlorine. Use of high chlorine dioxide substitution dramatically reduces the formation of TOC1. Xylanases may also be the ticket to success for a chlorine-free bleaching alternative. Table XV shows the various benefits obtained by the use of xylanase enzymes. Xylanase technology has been catapulted from biotechnology labs to pulp mills in just a few years. The main driving factors have been the economic and environmental advantages that the enzyme brings to the bleach plant. Such intense demand for the enzyme has pushed enzyme producers to develop an entirely new industry in a remarkably short time. The increasing competition among manufacturers will continue to improve products and reduce price. Development Prospects in Bleaching. Xylanase treatment represents a successful new technology for reducing chlorine use. Research aimed at understanding the mechanism of xylanase action in bleaching kraft pulp, finding new niches for xylanascs, and optimizing various combinations of xylanase treatments with the other bleaching stages are currently underway. Future developments will focus on producing enzymes with better resistance to high pH and temperatures. This is guided by the fact that the optimal conditions for an enzyme depend on both parameters, which means that high temperatures lower the pH optima of the enzyme and high pH values similarly lower the optimum temperature. The pH and temperature optima need to match the conditions occurring in the pulping process. 2. Enzymatic Debarking

Removal of the bark is the first step in all processing of wood. This step consumes substantial amounts of energy. Extensive debarking is needed for high-quality mechanical and chemical pulps because even small amounts of bark residues cause darkening of the product. In addition to its high energy demand, complete debarking leads to losses of raw material due to prolonged treatment in the mechanical drums. The border between wood and bark is cambium, which consists of only one layer of cells. This living cell layer produces xylem cells toward the inside of the stem and phloem cells toward the outside. In all the wood species studied, common characteristics of the cambium include a high content of pectins and the absence or a low content of lignin (Dey and

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

179

TABLE XV

BENEFITSOF XYLANASE PREBLEACHINC Bleaching

Sequence

Benefits

Conventional

X(CD)ElDEzD

Reduction of elemental chlorine and AOX Reduction of chlorine consumption and AOX; increase in productivity when C102 is limited Increase in brightness, Reduction in chemical consumption; unchanged strength properties

ECF

XDElDEzD XDElPDEzD OXDElDEzD

TCF

OXQPP OXQPZP QXZP

Brensen, 1984; Kato, 1981). The content of pectins in cambium cells varies among the wood species studied but may be as high as 40% of the dry weight. The content of pectic and hemicellulosic compounds is also high in the phloem (Fu and Timell, 1972). Pectinases are found to be key enzymes in the process, but xylanases may also play a role because of the high hemicellulose content in the phloem of the cambium (Viikari et al., 1989). One of the major difficulties with enzymatic debarking is the poor infiltration of enzymes in the cambium of whole logs. 3 . Fiber Modification

Enzymatic modification of fibers aims at decreased energy consumption in the production of thermomechanical pulps and increased beatability of chemical pulps or improvement of fiber properties. In highyield mechanical pulps, most of the lignin and hemicellulose remains in the pulps. According to determinations of the medium pore width and immunolabelings of untreated wood, it is evident that enzymatic modifications to the composition of mechanical pulps can be achieved only on the outer surface of the fiber. This was verified when xylanases were applied to thermomechanical pulp (Jeffries and Lins, 1989). Even when using rather high enzyme dosages, only about 1%of the pulp was dissolved. When combined with an alkaline pretreatment, the enzymatic treatment was substantially improved and the amount of energy required for refining thermomechanical pulp was decreased. In 1942, a patent claimed that microbial hemicellulases from Bacillus and various Aspergillus species could aid refining and hydration of

180

P. BAJPAI

pulp fibers (Diehm, 1942). In 1959, Bolaski et al. patented the use of cellulases from Aspergillus niger to separate and fibrillate pulp (Bollaski et al., 1959). The technology was principally applied to cotton linters and other nonwood pulp. A process patented in 1968 used cellulases from a white rot fungus to reduce beating or refining time (Yerkes, 1968). The desired structural changes in the fiber created during beating and refining are external fibrillation and fiber swelling, which improve the flexibility and bonding ability of the fibers. The role of xylans in fiber properties was studied using xylanases from Sporotrichium dimorphosporum in the treatment of fully bleached spruce sulphite and birch kraft pulps. Electron microscopic examination revealed external fibrillation and good flexibility of fibers, implying internal modification (Mora et al., 1986). The water retention value, which describes fiber swelling, was considerably increased. The conclusion was that xylan was hydrolyzed in the whole delignified cell walls. The enzymatically treated pulps were comparable to slightly beaten pulps. Beatability was generally enhanced, and the energy demand was reduced about threefold (Noe et al., 1986). Recently Bhardwaj et al. (1996) examined the effectiveness of several commercial xylanase enzymes for energy savings in beating and refining. A number of unbleached kraft pulpsnamely, softwood, bamboo, and mixed pulp (60”/0 waste corrugated kraft cuttings and 40% unbleached softwood pulp)-were treated with the enzymes. With softwood pulp, the maximum reduction in beating time was found to be 25%, while treatment with commercial enzymes with bamboo pulp and mixed pulp reduced the beating time by about 18 and 1 5 % , respectively. The strength properties of the pulp were not found to be affected. Water removal on the paper machine has been shown to improve as a result of limited hydrolysis of the fibers in recycled paper. A mixture of xylanase and cellulase enzymes at low concentrations has been found to markedly increase the freeness of recycled fibers without substantially reducing yield (Fuentus and Robert, 1988). The lower the initial freeness, the greater the gain following treatment. Many different cellulases and hemicellulases have been found to improve freeness (Bhardwaj et al., 1995; Pommier et al., 1989; Pommier et al., 1990). Freeness shows a rapid initial increase, with over half of the observed effect occurring in the first 3 0 minutes. A relatively small amount of enzyme is required. While the initial effects are largely beneficial, extending the reaction time with large concentrations of enzyme is detrimental. Unfortunately, crude enzyme mixtures also reduce strength properties. Mill trials have been carried out successfully using a commercial T reesei enzyme called Pergalase A40 (Pommier et al., 1990).

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

181

Mixed xylanases and cellulases peel the surface of fibers. If treatment is limited, the enzymes remove only elements that have a great affinity for water but which contribute little to the interfiber bonding potential. By selectively removing these surface components, pulp-water interactions are reduced and drainage increases without noticeable changes in the final mechanical strength properties of the pulp. If the treatment is extended, however, fibrillation becomes pronounced and drainage decreases. If large quantities of crude enzymes are used, the average fiber length is reduced, fines disappear, and the strength properties of the fibers are lost. Therefore, an optimum level of enzyme treatment is required. It has been reported that the drainability of mechanical pulp can also be enhanced by the addition of hemicellulases (Karsila et al., 1990).The authors claim that xylanase improves the freeness of deinked recycled pulp while having no detrimental effect on fiber tensile strength properties. By comparison, the tear indices of recycled pulps treated with cellulase decreased (Karsila et al., 1990). These findings suggested that xylanases might be much more effective than cellulases or crude xylanase-cellulase mixtures. Xylanases, however, remove hemicellulases that promote interfiber bonding. This effect can also lead to poor paper properties. The degree of polymerization of pulp treated with a cellulase-free xylanase was found to increase, apparently due to the selective removal of xylan that has a lower DP (Clark et al., 1989; Puls and Poutanen, 1989; Sharma, 1987).Thus, treatment of pulps withxylanases has been shown to increase their viscosities (Senior et al., 1989). However, even low cellulase activities in the enzyme preparation result in decreased viscosity. 4 . Production of Dissolving Pulp

Dissolving pulps are used to produce such cellulosic materials as acetates, cellophanes, and rayons (Hinki et al., 1985). Their manufacture is characterized by the derivatization and thus solubilization of highly purified cellulose. Hemicellulosic contaminants lead to color and haze in the product as well as insolubles that hamper the manufacturing process. Their extraction from pulps requires the use of high caustic loadings and appropriate pulping conditions, the latter restricted to sulfite pulping and acid-pretreated h a f t pulping. The use of xylanase for purifying cellulase was first proposed by Paice and Jurasek (1984). It was found that complete enzymatic hydrolysis of hemicellulose within the pulp is difficult to achieve. Even with very high loadings of enzymes, only a relatively small amount of xylan could be removed. The xylan content in delignified mechanical aspen pulp was reduced

182

P. BAJPAI

from approximately 20 to lo%, whereas in bleached hardwood sulphite pulp the xylan content was decreased from 4 to only 3.5% (Paice and Jurasek, 1984). The complete removal of residual hemicellulose thus seems unattainable, probably due to the inaccessible location of the substrate. Nevertheless, xylanase treatment may reduce the chemical loading required during caustic extraction or facilitate xylan extraction from haf t pulps. 5. Removal of Shives

Shives are small fiber bundles of fibers that have not been separated into individual fibers during the pulping process. They appear as splinters that are darker than the pulp. One of the most important quality criteria for bleached ha ft pulp is the shive count. Tolan et al. (1994) have studied the effect of xylanases on shive removal. It was found that a novel enzyme formulation called Shivex can be used to increase the efficiency of shive removal by bleaching. By treating brownstock with Shivex, mills can increase the degree of shive removal in the subsequent bleaching by 55%. Depending upon the shive level in the incoming brownstock and the desired shive level of the bleached pulp, this allows a mill to decrease its actual shive count or to increase its margin of safety against shives. The increase in shive removal is accompanied by an increased efficiency in the bleaching of pulp. Therefore, mills can decrease chlorine use in a bleach plant without compromising on shive counts. Shive is a multicomponent mixture of proteins, some of which are xylanases, but the degree of shive removal by the enzyme is not directly related to the enzyme’s xylanase activity or bleach-boosting effectiveness. 6. Retting of Flax Fibers

Another potential application of cellulase-free xylanolytic systems might be processing plant fiber sources such as flax and hemp (Sharma, 1987). At present, fiber liberation is affected by retting, that is, the removal of binding material present in plant tissues using enzymes produced in situ by microorganisms. Pectinases are believed to play the main role in this process, but xylanases may also be involved (Sharma, 1987). Replacement of slow natural retting by treatment with artificial mixtures of enzymes could become a new fiber liberation technology. B. OTHERAPPLICATIONS

The use of xylanases has also been proposed for clarifying juices and wines (Beck and Scott, 1974; Biely, 1985, 1991; Woodward, 1984;

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

183

Zeikus et al., 1991); for extracting coffee, plant oils, and starch (Biely, 1991; McCleary, 1986; Woodward, 1984); for improving the nutritional properties of agriculture silage and grain feed (Bedford and Classen, 1992; Campbell et al., 1991; Fengler and Marquardt, 1988; Grahm and Inborr, 1992; Groot Wassink et al., 1989; Linko et al., 1989; Poutanen and Puls, 1988; Zeikus et al., 1991; Pettersson and Aman, 1989); for macerating plant cell walls (Beck and Scott, 1974); for producing food thickeners (Zeikus et al., 1991); and for providing different textures to bakery products (Maat et al., 1992; McCleary, 1986). Maat et al. (1992) have identified a particular endo-l,4-Pxylanase produced by an Aspergillus niger as being very effective in increasing the specific volume of breads without giving rise to a negative side effect on dough handling similar to that observed with xylanases derived from other fungal or bacterial sources. The effect of xylanase on bread volume improvement can be ascribed to redistribution of water from the pentosan phase to the gluten phase. The increase in gluten volume fraction gives the gluten more extensibility, which eventually results in better ovenspring. Xylanase-based enzyme products are now widely used throughout the world to supplement pig and poultry diet based on wheat, triticale, and rye. Other enzyme activities in such products normally include P-glucanase, pectinase, amylase, and protease. Addition of these products can improve feed utilization and live weight gain primarily by improving digestion of such nutrients as starch, proteins, and fats in the intestine. It would appear that enzymes reduce variation in live weight, essentially eliminate litter problems, and help alleviate digestive disorders. There has also been an interest in the production of xylose, xylobiose, and xylooligomers (Biely, 1991; Puls et al., 1988; Woodward, 1984). More recent reports again show that such sugars can be prepared by the enzymatic hydrolysis of xylan (Irie et al., 1991; Korte et al., 1991; Pellerin et a]., 1991), whereas other sugar residues can be added using the transglycosylation activity of such enzymes as 0-xylosidase (Kizawa et al., 1991; Shinoyama et al., 1991). These xylose-containing sugars may be useful for research as well as for their rheological properties. With the exception of xylose conversion to xylitol, the bioconversion of lignocellulosic materials to fermentable sugars does not yet appear to be an economic prospect because of other more competitive sources of sugars such as starch and sucrose (Linko et al., 1989). However, the massive accumulation of agricultural, forestry, and municipal solidwaste residues are creating a large volume of low-value feedstock (Eriksson et al., 1990; Lynd et al., 1991; Trotter, 1990). Alternative

184

P. BAJPAI

technologies are desirable for dealing with all these materials, even if they are from the perspective of waste management. One alternative being considered is bioconversion to produce fuel ethanol, single-cell protein, xylanases, and other chemicals from xylan-rich material (Biely, 1985; Eriksson et a]., 1990; Gamerith and Strutzenberger, 1992; Shaw and Stephen, 1966a). Effective enzymatic hydrolysis of lignocellulosic substrates is dependent on the nature of both the enzyme system and the substrate. The characteristics of the xylan component depend on the pretreatment used to make the carbohydrate component of the substrate more accessible to enzymatic hydrolysis. In steam pretreatment processes, a stream rich in xylooligomers can be separated (Nguyen and Saddler, 1991), and these sugars can still have branches such as acetyl substituents. On the other hand, alkaline treatment tends to deacetylate xylans. If the feedstock is variable, a complete xylanolytic system would appear desirable to ensure maximum hydrolysis. Such an enzyme system would include xylanases, 0-xylosidases, and the various debranching enzymes. The prospects for the use of xylanolytic enzymes in bioconversion are low because of the high efficiency with which acid hydrolyzes xylans. VI. Conclusions

The new major large-scale application area of xylanases is clearly in the pulp and paper industry in order to increase the bleachability of kraft pulps. The improved bleachability is based mainly on the action of endo-P-xylanases, which can be produced efficiently on an industrial scale. Partial hydrolysis of xylan facilitates the extraction of lignin from the pulp in higher amounts and with higher molecular masses. The enzymatic pretreatment method is applicable to any traditional or modern bleaching sequence at existing plants without significant investment. The primary goals of the enzymatic treatment have been to decrease chemical consumption, reduce environmental loading, and/or to increase the final brightness of pulp. In the TCF bleaching sequences, enzymes can improve brightness, which may otherwise remain below an acceptable level without loss of viscosity. Enzyme-aided bleaching is thus both environmentally and economically advantageous. The potential for use of xylanolytic enzymes in the food and feed industries is also high. The main aim in the application of xylanolytic enzymes had been the hydrolysis of hemicellulosic substrate for production of fermentable sugars. The knowledge gathered on the hydrolysis mechanism of hemicelluloses, especially xylans, has greatly promoted the rapid application of these enzymes in new areas. The general

MICROBIAL XYLANOLYTIC ENZYME SYSTEM

185

information already available on individual xylanolytic enzymes will help in the development of future applications. ACKNOWLEDGMENTS

We are thankful to Shri S. S. Gill for excellent typing of the manuscript and to Shri S. P. Mishra for skillful assistance in the preparation of the manuscript. ABBREVIATIONS

Adsorbable organic halogens Biological oxygen demand Chlorination Treatment by mixing chlorine and chlorine dioxide simultaneously (Cl proportion is higher than D) Sequential addition of chlorine followed by chlorine dioxide Chemical oxygen demand Chlorine dioxide treatment Treatment by mixing chlorine dioxide and chlorine simultaneously (D proportion is higher than Cl) Sequential addition of chlorine dioxide followed by chlorine Degree of polymerization Alkaline extraction First and second alkaline extraction respectively Elemental chlorine free Alkaline extraction in presence of hypochlorite Alkaline extraction in presence of oxygen Alkaline extraction in presence of oxygen and hydrogen peroxide Alkaline extraction in presence of hydrogen peroxide Hypochlorite treatment International Standards Organization Oxygen delignificatiodbleaching Hydrogen peroxide treatment Chelation of metal Totally chlorine free Total organochlorine Xylanase enzyme treatment REFERENCES

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Tenkanen, M., and Poutanen, K. (1992). “Xylans and Xylanases” (J. Visser, G. Beldman, M. A. Kusters-van Someren, and A. G. J. Voragen, eds.), Progress in Biotechnology, Vol. 7, p. 203. Elsevier Science, Amsterdam. Tenkanen, M., Schuseil, J., Puls, J., and Poutanen, K. (1991). J. Biotechnol. 18, 69. Timell, T. E. (1964). A d z Carb. Chem. 19, 247. Timell, T. E. (1965). Adv. Carb. Chem. 20,409. Toda, S.,Suzuki, H., and Nisizawa, K. (1971). J. Ferment. Tech. 49,499. Tolan, J. S., Guenette, M., Thibault, L., and Winstanley, C. (1994). Pulp and Paper Can. 9512,T493. Trotter, P. C. (1990). Tappi]. 73(5), 201. Tsujibo, H., Sakamoto, T., Nishino, N., Hasegawa, T., and Inamori, Y. (1990). J. Appl. Bacteriol. 69,398. Ujiie, M., Matsuo, M., and Yasui, T. (1985). Agric. B i d . Chem. 49,1167. Ujiie, M.,Roy, C., and Yaguchi, M. (1991). Appl. Environ. Microbiol. 57,1860. Miiki, K., Tokela, V., Kitunen, V., and Salkinoja-salonen, M. (1989). Int. Symp. Vaheri, M., Chlorinated Dioxins and Related Compounds, gth, Toronto, Canada. Van Doorslaer, E., Kersters-Hilderson, H., and de Bruyne, C. K. (1985). Carbohydr. Res. 140,342. Vats-Mehta, S., Bouvrette, P., Shareck, F., Morosoli, R., and Kluepfel, D. (1990). Gene 8 6 , 119. Viikari, L., Ranua, M., Kantelinen, A., Sundquist, I., and Linko, M. (1986). Proc. Int. Conf. Biotechnol. Pulp and Paper Industq, 3rd, Stockholm, p. 67. Viikari, L., Ranua, M., Kantelinen, A., Linko, M., and Sundquist, J. (1987). Proc. fnt. Symp. Wood and Pulping Chemistq, 4th, 27-30 April, Paris, Vol. 1, p. 151. Viikari, L., Ratto, M., and Kantelinen, A. (1989). Finnish Pat. Appl. 896,291. Viikari, L., Kantelinen, A., Poutanen, K., and Ranua, M. (1990). In “Biotechnology in Pulp and Paper Manufacture” (T. K. Kirk and H.-M. Chang, eds.), p. 145. Butterworth-Heinemann, Boston. Viikari, L., Kantelinen, A,, Sundquist, J., and Linko, M. (1994). FEMS Mircobiol. Rev. 13, 335. Wilkie, K. C. B. (1979). Adv. Carb. Chem. 36,215. Wilkie, K. C. B., and Woo, S. L. (1977). Carbohydr. Res. 57,145. Wong, K. K. Y., and Saddler, J. N. (1992). “Xylans and Xylanases” (1. Visser, G. Beldman, M. A. Kusters-van Someren, and A. G. J. Voragen, eds.), Progress in Biotechnology, Vol. 7, p. 171. Elsevier Science, Amsterdam. Wong, K. K. Y., Tan, L. U. L., Saddler, J. N., and Yaguchi, M. (1986a). Can. J. Microbiol. 32, 570. Wong, K. K. Y., Tan, L. U. L., and Saddler, 7. N. (1986b). Enzyme Microb. Technol. 8 , 617. Wong, K. K. Y., Tan, L. U. L., and Saddler, J. N. (1988). Microbiol. Rev. 52,305. Wood, T. M., and McCrae, S. I. (1986). Carbohydr. Res. 148,321. Woodward, J. (1984). Top. Enzyme Ferment. Biotechnol. 8, 9. Yaguchi, M., Roy, C., Ujiie, M., Watson, D. C., and Wakar Chuk, W. (1992). “Xylans and Xylanases” (J. Visser, G. Beldman, M. A. Kusters-van Someren, and A. G. J. Voragen, eds.), Progress in Biotechnology, Vol. 7, p. 149. Elsevier Science, Amsterdam. Yang, J. L., and Eriksson, K. E. L. (1992). In “Biotechnology in the Pulp and Paper Industry” (M. Kuwahara and M. Shimada, eds.), p. 151. UM Publishers, Tokyo. Yang, J. L., Lou, G., and Eriksson, K. E. (1992). Tappi J. 75(12), 95. Yang, R. C. A., Mackenzie, C. R., Bilous, D., and Narang, S. A. (1989). Appl. Environ. Microbiol. 55,568.

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Yasui, T., Nguyen, 8. T., and Nakanishi, K. 1. (1984). Ferment. Technol. 62, 353. Yerkes, W. D. (1968). U.S. Pat. 3,406,089. Yoshioka, H., Nagato, N., Chavanich, S., Niluhol, N., and Hayashida, S. (1981). Agric. Biol.Chem. 45,2425. Zeikus, J. G., Lee, C., Lee, Y. E., and Saha, B. C. (1991). ACS Symp. Ser. 460,36. Zimmermann, W. (1989). In “Enzyme Systems for Lignocellulose Degradation” (M. P. Coughlan, ed.], p. 167. Elsevier Applied Science, London. Zhu, Y., Wu, Y. Q., Chan, W., Tan, C., Gao, J. H., Fei, J. X., and Shih, C. N. (1982). Enzyme Micrab. Technol. 4.3.

Oleaginous Microorganisms: An Assessment of the Potential JACEK

LEMAN

Znstitute of Food Biotechnology University of Agriculture and Technology 10-724 Olsztyn, Poland

I. Introduction 11. Microbial Oil A. Oleaginous Microorganisms B. Lipid Accumulation C. Lipid Characteristics 111. Single Cell Oil IV. Specialty Fats and Oils A. Cocoa Butter Equivalents (CBEs) B. Polyunsaturated Fatty Acids (PUFAs) V. Valuable Metabolites A. Fat-Soluble Vitamins and Pigments B. Other Lipid Classes VI. Conclusions References

I. Introduction

Yeasts were the first microorganisms recognized over a century ago to accumulate lipids. Since then, other lipid-accumulating species have been identified among bacteria, algae, and molds. The articles by Woodbine (1959) and more recently by Ratledge (1982, 1987, 1989, 1991, 1992a,b, 1993) provide a broad outline of the achievements in the study of microbial lipids. Interest in oleaginous microorganisms has been ongoing because microbial lipids and plant oils share some important characteristics with regard to fatty acid distribution, triacylglycerol types, and secondary metabolites. Thus, oleaginous microorganisms are considered as an alternative source of lipids, attractive because of their enormous growth rates on a variety of substrates, their ability to synthesize an array of products, and their amenability to genetic manipulation. Their competence in carrying out numerous transformation reactions, such as oxidation, desaturation, and hydrogenation, enables the upgrading of both lipid and carbohydrate products. The production of yeast oil-rich biomass on different waste materials for oil or fodder (Slater, 1988; 195 ADVAN( ES IN APPLIED MICROBIOLOGY VOLUME 43 Copyright B 1997 by ALademic lnc All righis of repruduction in any form rrsrrved 0065 2164197 $25 00

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Davies, 1988; Rydin et al., 1990; Hassan et a].,1994a; Johnson et a]., 1995) is the simplest example of such transformations. However, when only considered as a means of waste treatment, this strategy does not exhaust the possibilities that the oleaginous microorganisms offer. Modifications of this strategy have attempted to improve the quality of plant and animal oils and fats (Hassan et al., 1995) or to increase the yield from a single fermentation step, for example, by simultaneously producing a lipid-rich biomass and biosurfactant (Zhou and Kosaric, 1993). The first biotransformations of plant or animal lipids carried out with whole yeast cells (Hammond et al., 1981; Glatz et al., 1984; Bati et al., 1984) proved that the composition of lipids recovered from yeasts was similar to the substrate lipid, depending on the substrate (Johnson et a]., 1992a). After initial trials with fat biomodification (Koritala et al., 1987) two factors became obvious: either the mutants were lacking in enzyme activity, thus preventing further reactions of the substrate, or organisms had inefficient enzyme systems. The first factor benefitted the cocoa butter equivalent process, whereas the latter may be employed for biological refining of mixtures of triacylglycerols and fatty acids (Cho et al., 1990). Current interest in lipid biomodification with oleaginous microorganisms seems to focus mainly on concentrating nutritionally important polyunsaturated fatty acids present in plant and fish oils, and on transforming steroids and fatty acids. Except for a few minor products, the interest in oleaginous organisms has never resulted in industrial production. The problem is one of economics. The prices of plant oils and animal fats are relatively low. This is especially true for palm oil-derived cocoa butter (Ratledge, 1993), forcing the microbial production of cocoa butter-like oil out of the competition. Opinions regarding the economics vary, however. On the one hand, the instability of the global cocoa butter market may stimulate efforts to develop new and improved cocoa butter equivalents (CBEs) at more stable and lower prices. On the other hand, the cost of microbial oil may be a small factor compared to the health benefits derived. Thus, the opportunities for using oleaginous microorganisms now exist for such highest-value products as dietarily important nutrients (polyunsaturated fatty acids of the n-6 and n-3 series), pharmaceuticals (steroid hormones and eicosanoids), or industrial products (e.g., polymers, wax esters, hydroxy fatty acids). Attempts to reduce the cost of microbial lipids are continuing with emphasis on increasing the product value, using inexpensive substrates, screening for more efficient strains, and reducing the processing steps necessary for oil recovery from the cells.

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This review considers the potential of oleaginous microorganisms in producing microbial lipids with an emphasis on functionally superior products such as specialty fats and oils, and important metabolites that have not yet been extensively studied. The first major compilation of information on microbial oils in industrial applications has been published (Kyle and Ratledge, 1992). II. Microbial Oil

A. OLEAGINOUS MICROORGANISMS Oleaginous microorganisms were pragmatically defined by Ratledge (1989) as those accumulating lipid at more than 20% of their biomass. As this definition is somewhat arbitrary, microorganisms with a lower content of lipid can be included provided that they warrant commercial consideration (Ratledge, 1991). No distinct biochemical criterion of oleaginicity exists, although in general the presence of the enzyme ATP citrate lyase indicates the ability of a cell to accumulate over 20% of its biomass as lipid (Boulton and Ratledge, 1981). Oil-forming organisms are found among bacteria, yeasts, molds, and algae. Some examples of the principal species accumulating more than 40% lipid in their biomass are shown in Table I (a complete listing is given by Ratledge, 1989). According to an estimation by Ratledge (1991), about 125 species of molds and yeasts are oleaginous. Bacteria as a source of oil have been dismissed by most of the reviewers because the amount of lipid produced is small, typically less than 10% of dry biomass. Although some Arthrobacter sp. can accumulate as much as 80% and Mycobacterium, Corynebacterium, and Rhodococcus (Nocardia) species up to 30-40% lipid, they either grow slowly, produce lipids that are hard to extract, or are associated with toxic and allergic factors (Ratledge, 1984~). The lipid content in microalgae is typically 20 to 40%, but it may be as high as 85% dry biomass (Shifrin and Chrisholm, 1981). Using microalgae is not always economically feasible, as they require special growth conditions and harvestinghecovery techniques. However, they are considered a potential source of polyunsaturated fatty acids (PUFAs) and can now be grown heterotrophically (Radwan, 1991; Kyle, 1992). B. LIPID ACCUMULATION

Accumulation of lipid in the cell is typically a biphasic process requiring an excess of carbon over other nutrients, particularly nitrogen

198

J. LEMAN TABLE I SEI.ECTED

MICROORGANISMS ACCUMULATING FROM 40 TO 80% LIPID IN DRYBIOMASS

Bacteria Arthrobacter sp. Yeasts Cryptococcus curvatus C. terricolus Candida NCYC 911 Lipomyces lipofer L. starkey L. tetrasporus Rhodospiridium toruloides Rho do torula glu tinis Trichosporon cutaneum Endomycopsis vernalis Ti-igonopsis variabilis

Molds Entomophthorn conica Cunninghamella elegans Mortierella isabelina M. pusilla M. vinacea Mucor circinelloides M. mucedo M. plumbeus Pythium ultimum A s p e r g i h s fisch eri A. oryzae Chaetomium globosum Fusarium bulbigenum Oidium lactis Gibberella fujikoroi Humicola lanuginosa Penicillium lilacin um F! spinulosum Cladosporium herbarum Claviceps purpurea Ustilago zene

Algae Botryococcus braunii Dunaliella salina Nannochloris sp. Monalanthus salina Chlorella pyrenoidosa

(Ratledge, 1982; Boulton and Ratledge, 1984; Jacob, 1992; Brown et a]., 1989). Usually a C:N ratio of 40:l is applied, although it may vary greatly depending on the strain (Davies and Holdsworth, 1992).The first phase is characterized by rapid cell growth until the nitrogen is consumed, which is followed by the second phase, wherein the excess carbon in the medium is converted to lipid. The rate of lipid synthesis in both phases is similar, but the lipid accumulates in the cells because the synthesis of biomass ceases after the nitrogen is exhausted. A typical pattern of lipid accumulation in batch-cultured cells is shown in Fig. 1A. In continuous culture (Fig. lB),lipid accumulation is achieved by providing the nitrogen-limiting conditions at a dilution rate of about 0.03-0.15 h-l (Ratledge and Hall, 1979). The biochemical mechanism of lipid accumulation was determined by Colin Ratledge and his research group at the University of Hull, UK (Ratledge, 1984b; Ratledge and Evans, 1989) and can be summarized as follows.

U

I

0 zy

I

0

I

N

0

K

c

W

-8

- w 0

N

- 0

J 0

POTENTIAL OF OLEAGINOUS MICROORGANISMS

L

% ‘Pldll

W

FIG.1. Pattern of lipid accumulation in oleaginous organisms grown in batch (A) and continuous (B) cultures.

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FIG.2. Mechanism of fatty acid biosynthesis in oleaginous yeasts and molds. PFK = phosphofructokinase; PK = pyruvate kinase; PD = pyruvate dehydrogenase; PC = pyruvate carboxylase; CS = citrate synthetase; CT = citrate translocase; ACL = ATP citrate lyase; MD = malate dehydrogenase; ME = malic enzyme. *Mitochondria1 citric acid cannot be metabolized through the reaction of the tricarboxylic acid cycle because under N-limiling conditions the conversion of isocitrate to a-ketoglutarate is prevented with a low intracellular concentration of AMP under these conditions, specifically required for isocitrate dehydrogenase activity. Thus, provided a supply of carbon is still available, the cells begin to accumulate citrate when they have exhausted their supply of nitrogen.

An oleaginous microorganism metabolizing excess glucose under nitrogen-limiting conditions increases the ATP:AMP ratio in the cell to which isocitrate dehydrogenase in the mitochondria1 tricarboxylic acid cycle responds with an increased level of citrate (Fig. 2). The citrate is then transported out of the mitochondrion by the citrate-malate translocase system to the cytoplasm, where ATP citrate lyase cleaves citrate to acetyl-CoA and oxalacetate. This is the rate-controlling step for lipid biosynthesis and is feedback-inhibited by long-chain fatty acyl-CoA esters. The acetyl-CoA is the basic building block for fatty-acid biosynthesis, and the oxalacetate is converted to malate, which is transported back to the mitochondrion by the translocase system and converted

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201

there to oxalacetate, which is then used for citrate formation. Other control points in the glycolysis pathway are phosphofructokinase and pyruvate kinase. Both are inhibited by citrate, but this inhibition is removed by ammonium ions accumulating rapidly or slowly intracellularly when an organic or inorganic source of nitrogen, respectively, is used. The fatty acid synthesis from acetyl-CoA is catalyzed by cytosolic acetyl-CoA carboxylase, which is thought to be the rate-limiting step in fatty-acid biosynthesis, and fatty acid synthetase multienzyme complex. The immediate precursors for biosynthesis of triacylglycerols, the major storage lipid in oleaginous microorganisms, are fatty acyl-CoA esters and glycerol phosphate. This pathway is shown in Fig. 3. The enzyme phosphatidic acid phosphatase is the rate-limiting step of this pathway. The entire pathway of lipid biosynthesis from citrate translocation across the mitochondria1 membrane and cleavage of citrate by ATP citrate lyase to the final steps of lipid synthesis seems to be under strong regulatory control (Holdsworth and Ratledge, 1991). The yields of lipid from carbohydrate are generally from 15 to 20%, with the theoretical value being about 33% (Ratledge, 1991). For this reason the production of microbial oil is uneconomical. Many factors affect lipid biosynthesis, including the growth medium and its composition, pH, temperature, oxygen level, and growth rate. The amount of lipid and its composition are affected by these factors to different degrees, depending on the strain. Most organisms are able to grow over a wide range of pH values, although pH effects may be important depending on the strain (Johnson et al., 1992b) or when combined with the effects of temperature (Turcotte and Kosaric, 1989). A number of carbon sources including hydrocarbons, and most commonly available fermentation substrates except for methanol and cellulose, can be utilized by oleaginous microorganisms. The degree of lipid unsaturation, one lipid quality factor, has generally been shown to increase with an increase in: (i) unsaturation of fatty acyl-supplement in the medium, (ii) the saturation level of n-alkanes serving as the carbon source, and (iii)a decrease in the growth temperature. Increased aeration or higher pH values have also been reported to increase lipid unsaturation (Turcotte and Kosaric, 1989; Suutari et al., 1993). Only temperature and nitrogen limitation appear to have a definite effect on the lipid content, whereas the carbon source and temperature have a definite effect on the lipid and fatty-acid compositions. General reviews covering the aspect of lipid accumulation physiology have been compiled by Moreton (1988) and Turcotte and Kosaric (1989). As the control of fatty-acid and lipid biosynthesis is economically important, current approaches aim at determining by mathematical modeling the relation-

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202

R1-C- S-COA

6

CH20P03H2 H-&-OH kH2OH

-

FATTY ACYL CoA

sn-GLYCEROL 3-PHOSPHORIC ACID GLYCEROLPHOSPHATE ACYL TRANSFERASE

YH20POaH2 LYSOPHOSPHATIDICACID H-7-OH CH207- R i FATW ACYL GLYCEROLPHOSPHATE ACYL TRANSFERASE

CoA

CH20P03H2 H-k-OC-RP

I 6

CH205 -Ri

PHOSPHATIDICACID

I 0

H20 PI

7 a

PHOSPHATIDICACID PHOSPHATASE

y42OH H-C-o?-R2

I 0

FATTY ACYL

CoA

DIACYLGLYCEROL

ACYL TRANSFERASE

CH20C-R3

I 6 l o

H-C-o$-R2

TRIACYLGLYCEROL

FIG.3. Biosynthesis of lipids via the a-glycerol phosphate pathway. R,, Rz, and R3 are fatty-acid alkyl chains.

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203

ships between the availability of various nutrients and cell fatty-acid content (Granger et a]., 1993a,b), cell oil and oil fatty-acid contents 1993), and cell concentration and lipid content (Aggelis (Kennedy et d., et a]., 1995).

C. LIPIDCHARACTERISTICS

The lipid classes that occur in oleaginous microorganisms are many and varied depending on the species. The structures of typical classes of lipids and fatty acids in microbial oil are presented in Fig. 4. Triacylglycerols account for up to 95% of the lipids (Rolph et a]., 1989). Other common lipid classes (i.e., glycolipids, and mono- and diacylglycerols), usually contribute up to 10% of the total lipids (Ratledge, 1982). Many unusual lipid classes, such as sulfo- and peptidolipids, hydrocarbons, sterols, polyhydroxyalkanoates, wax esters (Ratledge and Boulton, 1985), glycerosulfates, and ether lipids (Poralla et al., 1980), are found in bacteria including archaebacteria. Yeasts and molds produce a variety of carotenoids, steroids, sphingolipids of the ceramide type, and glycolipids (Ratledge and Boulton, 19851. Branched-chain fatty acids and hydroxy fatty acids occur in bacterial lipids (Ratledge and Boulton, 1985). Algal lipids exhibit an exceptionally high proportion of PUFAs associated with polar lipids (Radwan, 1991). The fatty acids of yeasts are similar to those of plant oils, with oleic, palmitic, linoleic, and stearic acids being predominant (Ratledge, 1982). The fatty acids of molds show a greater range and diversity than yeasts, with short-chain fatty acids (ClO-C14), PUFAs, and hydroxy fatty acids being accumulated (Ratledge and Boulton, 1985). Ill. Single Cell Oil

Single cell oil (SCO) is a triacylglycerol type of oil from yeasts, analogous to plant and animal edible oils and fats (Ratledge, 1993). The use of microorganisms for this purpose has attracted attention since the early 1980s, although the idea is not new, having its beginnings in Germany during World Wars I and 11. Interest in using lipid-accumulating microorganisms for the commercial production of edible fats declined, however, when the fat shortage disappeared, and remained low until the processes for producing palm oil- and cocoa butter-type lipids were patented in the 1980s (Moon and Hammond, 1980; Tatsumi et al., 1977; Matsuo et al., 1981; Gierhart, 1982, 1984). The extensive research and development into SCO production carried out over the past several years was basically aimed at improving economic competitiveness as

204

J. LEMAN LIPID CLASSES

'kiacylglycerols

Phospholipids

CHzCOORl

CHZCOORl

CHCOORZ

CHCOOR2

CHzCOOR3

CHzCHzOP03-X

Rl, RP, R3 = fatty acid alkyl chain

X = choline, ethanolamine, serine, inositol

1

1

I

Glycolipids

Sphingolipids

CHzCOOR1

CH20-Y

CHCOORz

CHNHCORl

I

I

I

I

CH20-S

R,CH=CH-CHOH

S = sugar

Y = H (in ceramides), galactose or glucose (in cerebrosides)

Rz = fatty acid alkyl chain Rz = C13 alkyl chain

Hydroxy alkanoates

Sterols

FATTY ACIDS Saturated

CH,(CHzJ,COO€I

Unsaturated

CH,(CH~).I(CH=CH-CH~),~(CH~),COOH n l = 1 (n-3), 4 (n-61,7 (n-9), or 10 (n-12) n2 = 1, 2, 3, 4, 5, or 6

Hydroxy acids

CH,(CH2),,CH(OH)(CH2),,,COOH no = 0 (a),1 (P), 2 Ir),3 (6)

FIG.4. Selected lipid classes and fatty acids occurring in oleaginous organisms.

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205

TABLE I1 COMPARISON OF PRICES IUR PLANT AND M~CROBIAL OILS

Oils Bulk plant oils palm-, soya-, rape-, cottonseed

Price per ton (USD) 40OU-700

Reference Ratledge, 1993 (after Mielke, 1993)

Single cell oil

>loo0

Davies & Holdsworth, 1992 (based on 1985 costs)

Cocoa butter

3000

Ratledge, 1993

Palm oil-derived CBE"

2200

Ratledge, 1993

Microbial CBE

2400

Davies & Holdsworth, 1992 (tentative value)

"Palm oil; forecast to be 415-450 USD/ton in the year 2000 (Lubis ct a]., 1993). bCBE = cocoa butter equivalents.

compared to the plant-derived oils (Table 11). To achieve this goal, substrate costs for fermentation must be very low since the efficiency of carbohydrate-to-lipid conversion is not greater than 20%, which means that 5 kg of fermentable substrate is required to produce 1kg of oil. The only substrates that can offer cost benefits are waste materials of the food industry, preferably when the oil is intended for human consumption. The SCO processes based on molasses (Slater, 1988) and cheese whey (Davies, 1988) have been developed simultaneously. Although they were claimed to be economical, neither process is in operation commercially (Davies and Holdsworth, 1992). The technology of SCO (Davies, 1984; Slater, 1988; Davies and Holdsworth, 1992) consists of the production of oil-rich biomass by fermentation of a carbohydrate substrate by an oleaginous yeast, followed by downstream processing that includes the cell and oil recoveries (Fig. 5). A variety of yeasts are known to efficiently accumulate oil (Ratledge, 1989); however, Cryptococcus curvatus (formerly Apiotrichum curvatum ATCC20509 and Candida curvata D) isolated by Moon et al. (1978) has been shown to have the greatest economic potential (Davies and Holdsworth, 1992; Ratledge, 1991). After appropriate pretreatment, the substrate is either batch fermented or continuously fermented at 30°C and pH 5.5 with a sufficient amount of yeast inoculum

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Biomass production

FIG.5. Scheme of single cell oil process

to yield a biomass with 30-35% (wt/wt) oil. The cells are then recovered by centrifugation, disintegrated or dried, and solvent extracted, yielding a crude oil that is refined using conventional techniques. The technological details of SCO production from cheese whey, the most economical at present, has been reviewed by Davies and Holdsworth (1992). The oil is close in composition and properties to palm oil, and it may be converted to cocoa butter-like oil when fermentation is carried out under low-oxygen conditions (Davies et ul., 1990). The lack of commercial success for the SCO process, of primary interest for a long time for the exploitation of oleaginous microorganisms, however, was a driving force behind modifying fats to produce either plant oil equivalents of higher value (for example, cocoa butter) or products unavailable from agricultural sources. IV. Specialty Fats and Oils

The microbial specialty fats and oils denote lipids structured by environmental or genetic modification of an oleaginous species for functional, dietetic, and therapeutic purposes. The interest in microbial lipids as alternatives to unique edible plant oils resulted in the development of cocoa butter-like lipids. Also, the trend to construct healthy fats to meet nutritional needs has created a great interest in oils rich in PUFAs because of their unique physiological activities in the human body. This resulted in processes for the production of such products. The future for oleaginous microorganisms is linked to such lipid hormones as sterols and eicosanoids, and valuable microbial metabolites not found in plants, for example, hydroxyalkanoates or biosurfactants. A. COCOA BUTTEREQIJIVALENTS (CBEs) Cocoa butter, the storage lipid obtained from the mature bean of Theobromu cucuo, consists mainly of 1,3-disaturated-2-unsaturated

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triacylglycerols containing 23-30% palmitic acid, 32-37% stearic acid, and 30-37% oleic acid as fatty-acid components (Fincke, 1965; Padley et al., 1986). Thus, a cocoa butter substitute should consist of two-thirds saturated fatty acids and one-third monounsaturated fatty acids. Oleaginous species accumulate lipids generally characterized by a high content of oleic acid (40-50%) and palmitic acid (20-30%) but a relatively low content of stearic acid (5-10%). The percentage of oleic acid in the lipids of oleaginous microorganisms is therefore too high and the content of stearic acid too low to use these lipids as CBEs (Moon and Hammond, 1978; Davies, 1984). Although microorganisms capable of producing oil with a relatively high content of stearic acid (11-25%) in their triacylglycerols have actually been selected for use in fermentation (Davies and Holdsworth, 1992), they exhibit poor oil yields in the fermenter. Thus, the structuring of microbial lipids into CBEs relies on strategies that allow for an increase in the amount of stearic acid at the expense of oleic acid. Such strategies were first attempted by growing microorganisms on such lipid substrates as hydrocarbons and free or derivatized fatty acids. In contrast to carbohydrate substrates, lipid substrates influence the fatty-acid composition of the oil formed (Hammond et al., 1981), also benefiting by supplying a higher carbon content on a per-mole basis. The fatty-acid content of a lipid medium should contribute from 10 to 40% or even higher of the total carbon source in order to alter the metabolism of the cell (Gierhart, 1982). The composition of fatty acid employed as the carbon source is important, for each fatty acid affects a specific type of shift in the oil-synthesizing metabolism of a given species. When a mixture of fatty acids is used, their combined effect is an interaction yielding a mixture of triacylglycerols containing the various fatty acids present in the medium. Although the oil composition produced from any particular carbon source cannot be accurately predicted, some generalizations are possible and experimental attempts have been undertaken (Lee et al., 1993). The presence of a fatty acid of any given carbon length in the carbon source results in an increase in its proportion in the triacylglycerol. Thus, the use of palmitic, oleic, and stearic acids in the form of an emulsion as a carbon source promotes the formation of oils that are close to cocoa butter (Picataggio and Smittle, 1979; Noguchi et al., 1982; Gierhart, 1982, 1984; Beavan et al., 1992). Another strategy for increasing the stearic acid content in microbial oil is to inhibit the stearate-to-oleate conversion. Microorganisms do not synthesize unsaturated fatty acids directly. They are produced from a saturated “parent” by sequential introduction of double bonds first at

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the Ag position and then at the A12 and A15 positions. The enzyme responsible for inserting a double bond into stearic acid to convert it into oleic acid is Ag-desaturase (Baker and Lynen, 1971). To block the Ag-desaturase, three approaches have been attempted: the use of inhibitors, mutants, and hybrid strains. The naturally occurring cyclopropene compound sterculic acid, a specific Ag-desaturase inhibitor, was found to increase the stearic acid content up to 40% in selected oleaginous species (Gierhart, 1984; Moreton, 1985).However, a disadvantage of methods relying on the use of lipid substrates or desaturase inhibitors is that they need significant amounts of expensive components. Therefore, these methods are rather impractical from an economic point of view. The mutation strategy seems to offer greater advantages for structuring microbial oils (Ykema et al., 1989; Hassan et a]., 1993, 1994b). One such approach was the development of unsaturated fatty acid auxotrophs of Cryptococcus curvatus (formerly Apiotrichum curvaturn and Candida curvata D) with functionally inactive Ag-desaturase,referred to as Ufa mutants (Verwoert et al., 1989; Ykema et al., 1989, 1990; Hassan et al., 1993, 1994a). The selection of these mutants is based on their inability to grow in the absence of either oleic acid or oleic acid emulsified with Tween 80 after mutagenic treatment with ethylmethane sulfonate. For the efficient production of oil from oleic acid-requiring mutants, the growth medium must be supplemented with 0.6 or 0.2 g/liter of oleic acid as the essential component for a vital membrane system. Mutants cultivated in batch and single-state continuous culture with 0.2 g oleic acid per liter were reported to produce lipids with a fatty-acid composition comparable to cocoa butter (Table 111) and when cultivated in a liquid medium were sufficiently stable without significant reversion to nonfatty acid-requiring cultures for at least 50 generations (Hassan et al., 1993, 1994a). Although this seems a promising approach, the dependence on oleic acid still has an unfavorable effect on production costs. The hybrid strains of C. curvatus with a partially active Ag-desaturase and grown without oleic acid in a glucose medium or whey permeate seemed to offer a better solution. These strains were produced by spheroplast fusion between a methionine auxotroph mutant (with wildtype desaturase activity) and an unsaturated fatty-acid mutant (with blocked desaturase activity) (Verwoert et al., 19891. The Ufa strains used as a parent in the fusion were reported to affect the fatty-acid composition supposedly due to gene dose effects and/or nuclear DNA rearrangement in the hybrid cells. Yet, in general, lipids isolated from the hybrids

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TABLE 111 COMPARATIVE COMPOSITION OF MAJOR FATTYACIDS[FA) OF LWDS FROM

Relative % (wtlwt) of FA

CRYPTOCOCUS CUR VAT US^

Ufa Wild-type"."

M3b

Hybrid strain F33.10C

Cocoa butterd

26.0

23.8

23-30

Palmitic

34.0b

Stearic

10.2

13.9

36.5

31.2

32-37

42.8

43.7

22.0

29.6

30-37

7.0

9.6

8.0

5.6

2-4

46.6

44.3

65.2

62.6

60-64

Oleic Linoleic SFA

28.1'

OWild-type, unsaturated fatty acid (Ufa) mutant M3, Ufa hybrid strain F33.10 and cocoa butter. SFAs are saturated fatty acids. *Data from Hassan et al. (1994). CData from Verwoert et af. (1989). dData from Fincke (1965).

contained a higher level of saturated fatty acids, ranging from 33 to 64% of the total lipids, than wild-type C. curvatus. Some of the hybrid strains produced lipids with a fatty-acid composition close to cocoa butter (Table 111), showing growth rates and lipid production rates similar to the wild-type (Verwoert et a]., 1989). While these methods represent distinct improvements in tailoring the lipids of oleaginous microorganisms, Davies et al. (1990) showed that an increase of up to 23% by weight in the stearic acid content of C. curvatus wild-type cells can be achieved by limiting the oxygen uptake rate of the culture during the lipid-accumulating phase of cultivation. Economical production of CBEs requires then that the microorganism or stable mutant strain accumulates large amounts of stearic acid within the oil and utilizes an inexpensive carbon source in the growth medium. Among the microorganisms employed for CBE production, yeast species predominate, and C. curvatus has the greatest potential. The main attribute of this yeast lies in its excellent growth performance in fermenters and its susceptibility to genetic manipulation. Certain mold strains of the Mucorgenus, capable of accumulating a high percentage of stearic acid when grown on glucose (50 glliter) or acetic acid (2 g/liter) as a sole carbon source, have been reported (Kock and Botha, 1993; Roux et al., 1994).Of the 28 Mucorstrains grown on carbon sources, some produced

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h5, 2 0 : 4

18:2 a y - 1 8 : 3 - % 2 0 : 3 linoleic GIA DHGLA acid

n-6

AA

I

n-3

a - linolenic

EPA

DHA

acid

FIG.6. Pathways for the biosynthesis of PUFAs of the n-6 (linoleic) and n-3 (a-linolenic) series. A', A', AG, A", and A1' are fatty-acid desaturases, and E is elongase. Desaturation occurs toward the carboxyl end of the molecule and chain elongation takes place at the carboxyl end, leaving the methyl end unaltered. GLA is y-linolenic acid [6,9,12-cis-octadecatrienoic acid, 18:3, n-6); DHGLA is dihomo-y-linolenic acid (8,11,14cis-eicosatrienoic acid, 20:3, n-6); AA is arachidonic acid (5,8,11,14-cis-eicosatetraenoic acid, 20:4,17-6);EPA is eicosapentaenoic acid (5,8,11,14,17-cis-eicosapentaenoic acid, 20:5, n-3); and DHA is docosahexaenoic acid (4,7,10,13,16,19-cis-docosahexaenoic acid, 2233, n-3).

neutral lipids with a content of stearic acid as high as 27%. Changing the carbon source from glucose to acetic acid modified the metabolism of the strains, producing a 4.5-fold increase in the stearic acid content of the neutral lipids of M. circinefloides f . circinefloides CBS108.16 (Roux ef al., 1994).This may be due to an inhibitory effect of acetic acid on the generation of NADPH, fatty acid desaturation, and elongation cofactor (Ratledge, 1989).This observation suggests that there is still a need for further improvements and developments, including screening for suitable organisms and studying their responses to environmental modifications. The substrates used for CBE production are low-cost waste materials such as sweet whey (Davies, 1988),prickly-pear juice (Hassan et al., 1995), or monocarboxylic acids contained in wastes from the petrochemical process (Kock and Botha, 1993).

B. POLYUNSA'TURATED FATTYACIDS(PUFAS) Current information indicates that certain PUFAs are essential in human nutrition. These include the PUFAs of the n-3 and 17-6 series (Fig. 6),which differ in the position of the double bond closest to the methyl-terminal group of the fatty-acid chain. The nutritional role of other PUFAs, namely, the n-9 (oleic) and n-7 (palmitooleic) series, is

POTENTIAL OF OLEAGINOUS MICROORGANISMS

211

obscure, even though they have been detected in living organisms (Shimizu and Yamada, 1989; Ratledge, 1993). The n-6 and n-3 series of PUFAs play important roles in the structure and function of biological membranes and have other unique biomedical activities, such as lowering plasma cholesterol and triacylglycerol levels and preventing atherosclerosis and cardiovascular disease. These fatty acids are also precursors of various lipid hormones such as prostaglandins, leukotrienes, and thromboxanes (eicosanoids) in humans. Several articles on PUFAs provide excellent background information on a variety of topics (Dyerberg, 1986; Needleman e f al., 1986; Horrobin and Huang, 1987; Hansen, 1990; Sardesai, 1992; Hodge et a]., 1993; Stilwell et al., 1993).

Higher plants in which the PUFAs are synthesized de novo and animal tissues where they occur as the metabolites of linoleic or a-linolenic acid are the traditional sources of the n-3 and n-6 series of PUFAs (Table IV). However, these sources are rather unsatisfactory for several reasons, mostly because of their low oil productivity and the PUFA content in the resultant product as well as the presence of other fatty acids and flavor compounds with less desirable properties. Microorganisms, namely, some molds belonging to the order Mucorales, were suggested as promising PUFA sources in the mid-1960s (Shaw, 1965). However, only since the early 1980s have numerous research groups started to screen extensively for microorganisms capable of accumulating PUFA-rich lipids in order to obtain a suitable organism for largescale production. These efforts resulted in the successful commercial production of y-linolenic acid (GLA, C18:3, n-6) with a Mortierella strain in Japan and a Mucor strain in the United Kingdom (Ratledge, 1991; Suzuki, 1988). Strains of Mucorales, among others, are also being explored for the production of dihomo-y-linolenic acid (DHGLA,C20:3, n-6), arachidonic acid (AA, C20:4, n-6), and the fatty acids of the n-3 series: eicosapentaenoic acid (EPA, C20:5, n-3) and docosahexaenoic acid (DHA, C22:6, n-3) (Yamada et al., 1988; Kendrick and Ratledge, 1990). The production of PUFAs of the n-3 series from microalgae is being reconsidered and is expected to proceed rapidly (Kyle, 1992; Ratledge, 1994). The prospect for efficiently producing PUFAs requires a knowledge of the mechanism by which biosynthesis of PUFAs is accomplished by the cell. This, however, is still poorly understood, although the main aspects of PUFA accumulation are known. The first step of the biosynthesis of PUFAs is desaturation at the A6 position of linoleic acid or a-linolenic acid to yield GLA or octadecatetraenoic acid (C18:4, n-3),

212

J. LEMAN TABLE IV

MAJORSULlKCRS OF THE n-6 AND n-3 SERIES OF PUFAS

Source

PUFA content % of total fatty acids

Plants Evening primrose Borage B lackcurran t

6-10 19-25 17

Fungi Mortierella isabelina Mucor circinelloides

15-18

Fungi Mortierella alpina 1s-4

PlJFA

Productivity mg/g

Reference

n-6

GLA

DHGLA

AA

Ratledge, 1993 Ratledge, 1993 Ratledge, 1993 55

Shimizu & Yamada, 1989 Ratledge, 1993

7-23

28-123

Shimizu & Yamada, 1989; Jareonkitmongkol eta]., 1992, 1993

Animals Porcine liver

2-8

0.5

Shimizu & Yamada, 1989

Algae Porphyridium cruentum

5-36

20-80

Shimizu & Yamada, 1989

Fungi Mortierella alpina 1s-4

58

275

Shimizu & Yamada, 1989

Animals Fish oil

10-15

20

Shimizu & Yamada, 1989

Algae Chlorella minutissima

3540

10

11-3

EPA

Fungi Mortierella alpina 1.54

Pythium irregulare DHA

Animals Fish oil

Shimizu & Yamada, 1989

20

64

25

25

7-14

Jareonkitmongkol ct a]., 1993 O’Brien & Senske, 1993 Yongmanitchai & Ward. 1989

Fungi

Thraustochytriurn aureurn

49

70

Bajpai et al., 1991a,b,c,d

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213

respectively. This is followed by successive reactions of chain elongation and desaturation to yield the respective C20 and C22 PUFAs (Fig. 6). It is now well recognized that the desaturation and elongation of fatty acids occurs with membrane-associated activity in eukaryotic cells (Ratledge, 1989; Kendrick and Ratledge, 1992b). Complex enzyme systems, including enzymes necessary for supplying NADPH required for desaturation, and acetyl-CoA and NADPH for elongation, are now being studied (Chen et al., 1990; Kendrick and Ratledge, 1992a, 1996). The production of mutants blocked in some of the key desaturases (Fig. 6) and/or knowledge of how some of the desaturases obtain the required reducing power to drive the reaction would thus open up opportunities for increasing the yields of particular fatty acids and oil. Because PUFAs are mainly associated with membrane constituents such as phospholipids, the factors that promote the biosynthesis of polar lipids in the cell and/or increase membrane fluidity would probably optimize the synthesis of PUFAs. In addition to basic nutrients, temperature, and pH, lipophilic compounds known to stimulate the production of cell membranes and promote phospholipid biosynthesis as well as micronutrients should be studied in order to optimize the medium. Temperature is the principal regulating factor for the degree of unsaturation in the lipids of the fungus Entomophthora exitalis, whereas growth of the fungus at a constant dilution rate and temperature over a range of dissolved oxygen tension values has failed to influence lipid unsaturation (Kendrick and Ratledge, 1 9 9 2 ~ ) . Industrial use of fungi for PUFA production would require design of a process that would include fermentation, mycelium separation from the fermentation broth, and lipid extraction from the fungal mycelium. Several techniques have been developed for producing highly concentrated PUFAs. They all include solvent extraction (O’Brien and Senske, 1994) or supercritical fluid extraction (Sakaki et al., 1990) followed by isolation of the specific PUFA of interest, most efficiently accomplished using urea fractionation, low-temperature fractional crystallization (winterization) (Gunstone, 1992), supercritical C 0 2 extraction (Mishra et a]., 1993), or lipase-catalyzed alcoholysis, interesterification, and esterification (Li and Ward, 1993a,b,c, 1994b; Haraldsson et al., 1989). Other fractionation procedures such as preparative gas chromatography, fractional distillation, or preparative high-performance liquid chromatography would be necessary for producing highly purified PUFAs for medical purposes (Sajiki et a]., 1993; Wille et al., 1987). The study of PUFA production, discussed in more detail in the following paragraphs, has yielded potent microorganisms and improved our knowledge of their biosynthesis. However, more work is

214

J. LEMAN

needed to optimize C 2 W 2 2 PUFA production and to provide information that allows the initiation of commercial processes. 1. n-6 PUFAs

GLA was the first PUFA to be studied extensively. Initial research on the microbial production of GLA centered on screening a large number of microorganisms (Abraham and Srinivasan, 1984; Sumner et al., 1969; Suzuki et al., 1981;Tyrell, 1967; Shaw, 1965), and such studies are still continuing (Kristofikova et al., 1991; Cohen et al., 1992; Kendrick and Ratledge, 1992d; Roux et al., 1994; Hung-Chang-Chen and Chi-ChiaChang, 1994). Table V presents the species in which GLA has been identified as a lipid component. The most suitable species appeared to be mold strains of the order Mucorales that produce up to 30% GLA in the oil (Sumner et al., 1969). Strains producing high GLA levels generally have low oil content, whereas those with high oil content have low GLA levels (Ratledge, 1993; Kennedy et al., 1993). Two strains, Mucor circinelloides and Mortierella isabelina, were selected as those giving a satisfactory overall GLA content in the biomass (about 4%), thus ensuring a relatively high yield of oil (up to 20% (wt/wt)) with a high percentage (20%) of GLA (Ratledge, 1993). Commercial processes were developed in the United Kingdom in 1985 and in Japan in 1988 for both these strains (Ratledge, 1993;Suzuki, 1988). Although production in the United Kingdom was suspended in 1990 due to marketing problems, in Japan the production of GLA-containing oil has continued using a newly selected strain, Mortierella rarnrnaniana var. angulispora, which produces 25% GLA in the neutral lipids (Nakahara et al., 1992).Because the optimum conditions for lipid accumulation by this strain were found to be different from those for cell growth, a two-stage continuous fermenter system has been employed in which the first fermenter is for cell growth under a high glucose concentration (up to 100 g/liter), and the second, larger fermenter is for lipid accumulation at lower pH and temperature (Nakahara et al., 1992). Attempts to develop an industrial process for GLA-rich oil using Mucor hiemalis IPD51 are continuing in New Zealand (Kennedy et al., 1993,1994).To optimize the process, this group has developed a model for maximum GLA production, concluding that a number of medium compositions needed to be evaluated to determine the tradeoff between oil content of the cell and GLA content of the oil (Kennedy et al., 1993). Shake flask culture has provided a quick and inexpensive way of optimizing the medium composition (Kennedy et al., 1994). Solid-state fermentation (Lonsane et al., 1985) on moist cereal substrates has also been employed successfully for GLA production by

POTENTIAL OF OLEAGINOUS MICROORGANISMS

215

TABLE V SELECTED SPECIES IDENTIFIED FOR Y-LINOLENIC Acm (GLA) ACCUMULATION Species

Absidia Blastocla diella e m ursoni Blakeslees trispora Chaonephora cucurbitarum Dactylaria ampulliforne Echin osph orangium tran sversolis Gilbertella persicara Mortierella Mucor Paramecium tetraurelia Rhizopus Synceph olostum Tatrohymena Thamnidium elegans Spirulina

Reference Iwasaki etal., 1989: Nishimura et al., 1988 Sumner, 1970 Dedyukhina & Bekhtereva, 1969 Deven & Manocha, 1975 Sumner & Evans, 1971 Akimoto et a/., 19gOa,b Seto, 1986a,b Nishimura et al., 1988 Mumma etal., 1970; Nishimura etal., 1988 Hennessey & Nelson, 1983 Nishimura et al., 1988; Herbert & Heith, 1985 Nishimura et al., 1988 Gosselini et a]., 1989 Seto, 1986b; Manocha & Campbell, 1978 Gunstone, 1992; Hudson & Karis, 1974; Materassi et al., 1980

Cunninghamella japonica (Emelyanova, 1996). Using this technique, the GLA content of the biomass ranged from 7 to 8% (wt/wt) after growing the fungus on rice and millet, and the lipid obtained had 20% GLA in the total fatty acids, thus being comparable with commercially produced GLA oils (Ratledge, 1993). Because solid-state fermentation is useful when demand for a product is limited (Lonsane et a]., 1985), it is an interesting method for a newly developing market, as the risk to a producer and the investment costs may be expected to be less dramatic. The use of immobilized Mucor ambiguus IF06742 cells to enhance GLA productivity has also been considered. The mold is immobilized on porous support particles and excretes lipids containing GLA into the culture broth and/or the surface of the cell wall in the presence of nonionic surfactants (Fukuda and Morikawa, 1987). Mortierella MM151 mutants that have an enhanced GLA productivity, that is, 16.5% in the oil compared to 9.7% in wild-type cultures, have been reported as well. Industrial processes typically use a glucose-based medium that is a relatively expensive substrate. To reduce cost, the carbohydrate sub-

216

J. LEMAN TABLE VI FATTY ACIDPROFILES OF GLA-RICHOILS Content (%wt/wt)

Planta Oil Fatty acids Palmitic Palmitoleic Stearic Oleic 1,inoleic y-linolenic (GLA) Other

19

Commercial strain'

Mucor hiemalis IDD51C

Cunninghamelln japonid

20

41

37

23.5

25.2

12.6

1.0

0.8

1.0

2.5

6.5

9.6

5.3

8.0

9.0

39.5

32.6

43.2

70.0

11.0

11.9

16.3

10.0 0.4

16.5

15.4"

21.2

0.2

3.0

0.4

"Evcning primrose (Ratledge, 10031. *MMucor circinnelloides (Ratledgc, 1993). 'Shake flask culture on whey-based medium (Kennedy ef a!., 1993). dSolid-state culture on soaked rice (Emelyanova. 1996). "Maximum oil ConteIil 31.3% at rcduced oil conlent (Kennedy et al., l'J93)

strates were replaced with monocarboxylic acids, such as acetic acid, contained in waste materials from petrochemical processes (Kock and Botha 1993, 1994; Roux et al., 1994; Preez et al., 1995, 1996) for GLA production by Mucor strains. Using subtoxic levels of acetic acid (2 g/liter), M. circinelloides CBS203.28 was reported to accumulate up to about 40 mg GLA/g biomass based on a crude oil content of 28% that contained 91% neutral lipids, of which GLA was about 16% (Preez et al., 1995). This strain was also shown to utilize n-butyric and n-valeric acids as rapidly as acetic acid, whereas their isoforms and propionic acid were assimilated at slower rates (Preez et a].,1996). Monocarboxylic acids yielded about 27% crude oil containing about 84% neutral lipids in which GLA was about 14% (Preez et al., 1996). The fatty-acid profiles of some GLA-rich oils (Table VI) are of importance in view of stability and/or refining. The methods for producing highly purified GLA oils (up to 90%) include urea adduct formation (Traitler et al., 1984, 1988), separation on zeolites (Arai et al., 1987), differential crystallization (Yokochi et d., 1990), supercritical fluid

POTENTIAL OF OLEAGINOUS MICROORGANISMS

217

chromatography (Sakaki et al., 1988), and lipase-catalyzed selective esterification and hydrolysis of the microbial oils (Mukherjee and Kiewitt, 1991). According to Ratledge (1993), however, purified GLA microbial oils do not have any advantage over the original oils at present. Fungi belonging to the order Mucorales are also an attractive potential source of arachidonic acid (AA, C20:4, n-6). Because AA is produced via dihomo-y-linolenic acid (DHGLA, C20:3, n - 6 ) in the n-6 route (Fig. 6 ) , all the AA-producing fungi potentially have the ability to produce DHGLA (Shimizu et al., 1988a,b, 1989a; Yamada et d., 1987a,b, 1988; Shimizu and Yamada, 1989). The most active AA-producing fungi belong to the genus Mortierella, and the soil isolate M . alpinu 1s-4has been identified as accumulating a large amount of AA (Shimizu et al., 1988b, 1989b). In contrast to AA, the fungal content of DHGLA is, however, low, with the ratio of DHGLA to AA usually ranging from 0.1 to 0.3 (Shimizu and Yamada, 1989; Eroshin et al., 1996). The strains with moderate AA content (10-20% of the total lipids) were found to produce larger amounts of GLA and DHGLA when grown on a potatoglucose-agar medium with 0.84 g acetylsalicylic acid per liter (Eroshin et a]., 1996). Efforts were made to increase the mycelial content of DHGLA in M. alpina 1s-4 by inhibiting the A5-desaturase responsible for the conversion of DHGLA to AA (Fig. 6 ) . This was first attempted with natural inhibitors of the desaturase such as sesamin, contained in sesame oil, or curcumin, the major component of turmeric (Shimizu et al., 1989a, 1991, 1992). Although effective, this method had little practical value when compared to the As-desaturase-defective mutants of M. alpina 1s-4(Nakajima and Shimauchi, 1992; Jareonkitmongkol et al., 1992a,b, 1993b). In submerged culture under optimal conditions for 6-7 days at 28°C in 5-10-liter fermenters, the mutants produced lipids in which DHGLA amounted to about 23-43%, whereas AA was only 1-4% (Jareonkitmongkol et al., 1992b, 1993b). Although the ability to produce AA is widely distributed in Mortierella, high AA production is rather rare. The AA content of fungal lipids ranges from 2 to 15% (Shimizu et al., 1989b), although values as high as about 50% of total lipids have been reported in Mortierella species (Totani and Oba, 1987; Totani et al., 1987; Bajpai et al., 1991c; Shinmen et al., 1993). Only 3 strains out of 66 screened by Eroshin et al. (1996) were found to produce 42 to 55% AA, with an inverse relationship between AA and oleic acid content being observed. The production of AA by fungi can be substantially enhanced by altering the

218

J. LEMAN

cultivation conditions (Table VII). Early Japanese studies, confirmed later by Bajpai et al. (1991c), reported that the proportion of AA in M. alpina 1s-4total fatty acids was nearly doubled on storing the mycelia at 18°C for 6 days after growth at 28°C (Shinmen et al., 1989). At the lower temperature, the fungus grown in batch culture produced lipid with a greater degree of unsaturation at 12°C than at 28°C (Shimizu et al., 1988a,b). This effect of lower growth temperature on the higher accumulation of AA was also evidenced for Entomophthora exitalis grown in continuous culture at temperatures ranging from 20 to 30°C (Kendrick and Ratledge, 1 9 9 2 ~ )This . mold produced PUFAs of both the n-3 and n-6 series, of which the latter predominates, with AA as the major fatty acid (Kendrick and Ratledge, 1 9 9 2 ~ )As . the growth temperature decreased, the proportion of AA increased in the phospholipids, and in the sphingo- and glycolipids, all having a structural role in the cell, whereas the triacylglycerol fraction, being a reserve storage lipid, showed little change in the degree of unsaturation. This phenomenon may result from an adaptation mechanism to coldness that helps to maintain membrane fluidity that is regulated by phospholipid unsatuPythium ultimum 144 ration (Kendrick and Ratledge, 1 9 9 2 ~ )However, . did not respond to lower growth temperature by increasing the degree of fatty-acid unsaturation, behaving quite the opposite (Gandhi and Weete, 1991). Apart from temperature, olive and soybean oils were also reported to stimulate the accumulation of AA in Mortierella fungi (Shinmen et al., 1989), supporting the findings on the positive effects of myristic and palmitic acids on the AA level in the Aspergillus strains (Radwan and Soliman, 1988). Manganese ions in a concentration range of 2-500 mg/liter have been reported to favor the accumulation of AA, whereas iron ions at a concentration above 40 mg/liter had a strong inhibitory effect on AA accumulation in Mortierella sp. S-17 (Sajbidor et al., 1992). Sajbidor et al. (1994) have developed a method for producing highly pure (up to 90%) AA concentrate using M. alpina S-17 grown in shaker flasks with 3% glucose medium at 28°C for 14 days. After the urea adduct formation, the AA content in the resultant lipid increased from 50 to 78%, or to 91% when the fatty acids were fractionated as methyl esters. Despite many scale-up problems with solid-state fermentations (Lonsane et al., 1992), solid substrates such as barley, rice, or wheat were studied for improving the fungal production of AA-rich oils. M. alpina CCF185 grown on millet or barley at 28-30"C for 14 days under aeration yielded up to 16% lipids in the biomass with 50% AA in the total fatty acids (Stredanska et al., 1993).

POTENTIAL OF OLEAGINOUS MICROORGANISMS

219

TABLE VII EFFECT OF SOME GROWTH CONDITIONS ON THE AA PERCENTAGE IN TOTAL FATTY ACIDSOF FUNGAL OILS Variable

%AA

Comments and reference

Carbon Glucose n-Alkane Olive oil

36.9 19.3 28.6

Shake flask cultures of Mortierella alpina CCF185 on glucose medium; Stredanska et al., 1993

11.6 34.5 36.4

Stredanska eta]., 1993

Temperature 30°C 20%

28.8 51.4

Continuous culture of Entomophthora exitalis on glucose medium; Kendrick & Ratledge, 1992a

Air flow rate 1000 ml/min 150 ml/min

37.5 36.9

Kendrick & Ratledge, 199Za

Nitrogen (M412S04

NaN03 Yeast extract

A market for n-6 PUFAs, and for GLA oils in particular, seems to be developing along with the production technology. Although pharmacological applications are now limited to the treatment of marginal disorders (Ratledge, 1993), dietary applications are being considered for both humans and animals. GLA oil has been suggested as a food supplement or health food (Nakahara et a]., 1992). In animals, GLA oil has been reported to protect against rumen biohydrogenation as both AA and DHGLA contents in cow milk increased on supplying a cow with GLA instead of linoleic acid (Hermansen et al., 1995). 2. n-3 PUFAs

In contrast to the n-6 PUFAs, the production of eicosapentaenoic (EPA, C20:5, n-3) and docosahexaenoic (DHA, C22:6, 12-3) acids of the n-3 PUFAs had received little attention. Studies suggesting the essentiality of these fatty acids in the diet, in addition to the requirement for n-6 PUFAs (Sardesai, 1992), however, have led to increased research activity. Growth in the market for functional food enriched with n-3 PUFAs (Lauritzen, 1994) can be expected. The idea of producing EPA from fungi was first reported by Yamada et al. (1987a). When screening for molds capable of accumulating de-

220

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tectable amounts of EPA, they found that lowering the cultivation temperature increased the accumulation of EPA by all the AA-producing Mortierella strains they examined. Subsequently, these authors suggested that EPA was formed by a unique A17-desaturase(Fox et al., 1983) activated on cold adaptation and acting on AA from the n-6 series of PUFAs (Shimizu et al., 1988a,b) (Fig. 6). This temperature-dependent activity of Mortierella strains has been reported by other authors, although the optimum temperature for EPA production was found to differ depending on the strain or the medium composition (Bajpai et al., 1991a,b,c,d, 1992; Kotula and Yi, 1994). Several other fungi belonging to the genera Saprolegnia (Shirasaka and Shimizu, 1995) and Pythium (Wessinger et al., 1990; Gandhi and Weete, 1991; O’Brien et al., 1993) have also been reported to produce relatively large amounts of oil with EPA. Yazawa et al. (1992)identified a strain of marine bacteria, Shewanella putrefaciens, having 10 to 15% lipid, 25 to 40% of which was EPA. Other EPA-producing bacteria isolated from marine and freshwater sources include species of Altermonas, Flexibacter, and Vibrio (Yazawa et al., 1992; Ringo et al., 1992). However, better sources of EPA are marine microalgae such as the red alga Porphyridiurn cruentum that produces over 40% EPA in its total fatty acids (Cohen and Heimer, 1992) and Phaeodactylum tricornutum, the marine diatom having a daily EPA productivity of 6 mg/liter (Reis et al., 1996). Microbial sources of n-3 PUFAs have been reviewed by Radwan (1991) and discussed in detail in a multiauthor treatise published by the American Oil Chemists’ Society (Kyle and Ratledge, 1992). However, research efforts aiming at utilizing microorganisms for the biotechnological production of n-3 fatty acids are just beginning. Little is known about the responses of the microorganisms to environmental variables. For EPA production, the isolation of microorganisms with low levels of AA is essential because of the opposite effects these acids could produce as precursors of differently active eicosanoids (Sardesai, 1992), along with the optimization of growth conditions. Interestingly, the effects of cultivation variables are quite different for Mortierella and Pythium strains. The differences in cultivation methods and growth medium make any comparisons difficult. The EPA production by eight strains of Mortierella grown in shaker flasks has been studied by Kotula and Yi (1994) to obtain some definite information on the process. The effects of temperature, temperature shifting, media, and supplements were evaluated for EPA production by several species, M. elongata ATCC16271 was found to be the most effective strain when grown in yeast malt broth for 7 days at 18OC and

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221

pH 5.1, yielding 66 mg EPA/g dry mycelial cells. This yield was more than two times higher than that obtained from temperature-shifting cultures, although temperature shifting was shown to increase the production of EPA most effectively the earlier the shifting time occurred. A potato dextrose medium supplemented with sucrose, glucose, soluble starch, raffinose, glutamic acid, alanine, inositol, panthotenic acid, or antibiotics (penicillin G plus streptomycin) did not increase the content of EPA. The production of EPA was completely inhibited by ammonium sulfate and nitrate. Shimizu et al. (1989b) demonstrated that the Murtierella strains accumulate EPA in their mycelia when grown in media containing a-linolenic acid or its natural source such as linseed oil. This was further demonstrated in the A12-desaturase-deficientmutant of M. alpina 1s-4 (Jareonkitmongkolet al., 1993a). When cultured in a 5-liter fermenter at 24°C for 2 days and then at 20°C for 8 days on 1%glucose, 1% yeast extract, and 3% (v/v) linseed oil, the fungus produced about 64 mg EPA/g dry mycelium, which accounted for about 20% EPA of the total fatty acids, mostly in the triacylglycerol form. However, the mycelial EPA:AA ratio was low (2.5), which was unsatisfactory from a nutritional point of view. Two freshwater fungal strains, Saprolegnia spp. 28YTF-1 and 28GTF, were studied by Shirasaka and Shimizu (1995). The EPA content of these strains ranged from 13 to 1 5 mg/g dry mycelium, and the EPA:AA ratio was 5.5. Optimization of the culture conditions for Saprolegnia sp. 28YFT-1 showed that olive oil was a better carbon source for EPA production than starch, dextrin, saccharose, and glucose. The fungus produced about 1 7 mg EPA/g dry mycelium when grown with shaking on 2.5% olive oil and 0.5% yeast extract at 28°C and pH 6 for 6 days. Shifting the temperature from 28 to 6°C markedly increased the EPA:AA ratio in the fungus, but this was mostly due to a decrease in AA production at an EPA content ranging from about 19 to 23 mg/g dry mycelia. An additional nitrogen source (unspecified) in the medium decreased the production of EPA, but stimulated the growth of the fungus. The EPA was found to be distributed among the neutral (about 60%) and phospholipid fractions, and its proportion in the total fatty acids was only 3.5%, indicating the rather poor potentiality of the fungus. In contrast to EPA production by filamentous fungi grown in shake flasks (Gandhi and Weete, 1991; Shirasaka and Shimizu, 1995; Akimoto et al., 1990b; Kotula and Yi, 1994),O'Brien et al. (1993) has investigated EPA production by Pythium irregulare in a fermenter operated in airlift mode at 22 or 14°C using sweet whey permeate medium or glucose medium as the carbon source and yeast extract or corn steepwater as a

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complex nitrogen source. Sweet whey permeate (lactose) and yeast extract were found to be the preferred carbon and nitrogen sources, respectively, over glucose and corn steepwater for the production of EPA-rich oil, and lower growth temperature stimulated EPA accumulation in the fungus. The productivity obtained was about 25 mg EPAIg dry biomass, and the mycelial lipid had about 25% EPA contained mostly (about 90%) in the neutral fraction, which is important for the feasibility of EPA recovery from the lipid. A bacterial strain, possibly Shewanella putrefaciens, isolated from mackerel intestines was grown in natural or artificial seawater containing peptone and yeast extract. The cellular lipids contained about 9 mg EPAIg dry biomass when grown in this medium at 25°C and pH 7 for 8 h (Akimoto et al., 1990b). Commercial production of EPA with this marine bacterium is being considered using the novel isolate (Yazawa et a]., 1992). There is not much information regarding the responses of microalgae to environmental variables, with the exception of the green alga Chlorella minutissima (Seto et al., 1984), the red alga Porphyridium cruentum (Cohen and Heimer, 1992), and the marine diatom Phaeodactylum tricornutum (Arao et al., 1987; Kaixian and Borowitzka, 1993; Moreno et al., 1979; Velosa et al,, 1991). The effects of light intensity, temperature, pH, salinity, and nutrient starvation on the growth rate and EPA productivity by these microalgae have been studied (Cohen et al., 1988, 1992; Cohen, 1990; Set0 et a!., 1984). These variables have been shown to greatly influence the growth rate, EPA production, and fattyacid composition, as well as the distribution of EPA and fatty acids among particular lipid classes. Chlorella and Porphyridium differ in their responses to cultivation conditions. The optimal temperature (25OC) for Porphyridiurn growth was also found to be optimal for EPA production, whereas Chlorella produced higher amounts of this fatty acid at a nonoptimal growth temperature. Similarly, high salinity reduced the total fatty-acid content and EPA production in Porphyridiurn, whereas in ChlorelIa the EPA content increased using sodium chloride supplements. The mass production of Phaeodactylum tricornutum as a source for lipids and EPA has been studied using low temperatures and various light intensities (Yongmanitchai and Ward, 1991, 1992; Moreno et al., 1979; Velosa et al., 1991). The continuous production of EPA by this diatom at 24OC in indoor light has been reported (Reis et al., 1996). The EPA:AA ratio for this organism is extremely high (Table VIII), making EPA purification relatively easy.

223

POTENTIAL OF OLEAGINOUS MICROORGANISMS TABLE VIII

COMPARISON OF FATTY ACIDPERCENTAGES IN THE TOTAL FATTY Acms OF FUNGAL AND ALGALO m Fatty adds“ (% total)

Source

SFA

MUFA

PUFA

AA

nd 22 nd

nd 17.2 71.ge

nd 15.6

7.8 10.4

5.6

36.0

3.7

19.5

28.7

11.3 20.1

EPA

Other

EPAI AA

Fungi

Mortierella alpino 15-4 Mut 48‘

Pythium imgula& Saprolegnia sp. 28YTF-ld

72.7 39.5 18.1

2.5 2.4

1.0

19.5 25.2 3.5

20.7 0.4

28.7 25.8

0 5.5

1.4 64.5

3.5

Algae

Porphyridium cruentum 113.809

Phaeodactylum tricornutud

OSFA = saturated fatty acids: MUFA = monounsaturated fatty acids: PUFA = polyunsaturated fatty acids, except arachidonic = AA and eicosapentaenoic = EPA acids. ’Jareonkitmongkolet al., 1993. “O’Brienet al., 1993. dShirasaka and Shimizu 1995. eOleic acid. kinoleic acid. Kohen, 1990. *Reis et ol., 1996.

The growth of microalga MK8909, which accumulates 50% of its weight as an oil consisting mainly of myristic, palmitic, and oleic acids and 2 4 % EPA, has been optimized (Kyle et al., 1991; Boswell et al., 1992) for heterotrophic conditions under which it produced over 8 g of oil per liter of broth per day. Compared to EPA, dihomohexaenoic acid (DHA) has received relatively little research attention. Potential producers of this fatty acid are now being sought among phytoplankton, algae, fungi, and bacteria (Bimbo, 1987; Yongmanitchai and Ward, 1989; Kendrick and Ratledge, 1992d; Kyle et al. 1992; Molina Grima et a]., 1993; Kashiwakura et al., 1994; Yano et al., 1994). Fungal species belonging to the orders Saprolegniales and Entomophthorales, and particularly Thraustochytrjales, which display light-stimulating growth, appear to be the most promising sources of DHA (Yongmanitchai and Ward, 1989; Bajpai et al., 1991a,b, 1992; Li and Ward, 1994a; Singh et al., 1996). The production of DHA by fungal strains of Traustochytrium aureum has been reported

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by Ward’s group at the University of Waterloo, Canada (Bajpai et al., 1991a,b),and studies on the screening of strains of Traustochytrium and Schizochytrium have been reported by Li and Ward (1994a) and Singh et al. (1996). The effects of medium composition, temperature, and temperature shifting on the fungal growth, lipid and DHA production, and fatty-acid profile were studied in shaking cultures under light. The biomass yield and oil content of these molds are small, u p to 10 g/liter and 15%, respectively: however, the DHA content can range from 50 to 80% of the total fatty acids (Singh et al., 1996). The optimum strategy for the development of a process using these fungi consisted of optimizing the biomass production. The optimum conditions for growth and DHA production by the selected strain Thraustochytrium sp. ATCC 20892 were pH 7 at 25°C with 40 g glucose per liter for 4 days. The temperature shifting enhanced the yield and productivity of UHA but not biomass yield under these conditions. Even less is known about the production of DHA by microalgae. The strain MK88505 optimized to grow under heterotrophic conditions was reported to produce only 35% of its biomass as triacylglycerols but with DHA making up 30% of the total fatty acids (Kyle et al., 1992). An American firm (Martek Corporation, Columbia, MD) has patented a process using marine dinoflagellates to produce single cell edible oil containing DHA. In the process, marine dinoflagellates are cultured in fermenters and induced to produce the oil, which is subsequently recovered by extraction with solvents (Kyle et al., 1995). Efficient production of DHA by heterotrophic semicontinuous fermentation of the marine alga Crypthecodinium cohnii and its subsequent use for aquaculture of marine juvenile fish has been reported by Kashiwakura et al. (1994). The maximum yield of mycelial biomass after 100-150 h of culturing was 100 g/liter medium, and the lipid and DHA contents were 25% of dry biomass and 39% of the total fatty acids, respectively, with a DHA productivity of 8 g/liter medium. In experimentation on nutritional enrichment of shrimp followed by brining for preservation, the final content of brine shrimp was 30% of the total fatty acids, or 3% of dry bodies, with the brining process not causing any loss in DHA biological activity. The marine microalga Isochrysis galbana is another potential producer of DHA and EPA (Lopez Alonso et d . ,1992; Molina Grima et al., 1992, 1993, 1994). After being fractionated by the urea and HPLC methods, the oil of this microalga yielded EPA and DHA concentrates of 92-96”/0 purity (Robles Medina et al., 1995). Incorporation of EPA and DHA into plant and fish oils is being studied with both commercially available lipases and whole cells. The

POTENTIAL OF OLEAGINOUS MICROORGANISMS

225

strategy with the whole cells includes growing both oleaginous and nonoleaginous organisms in a medium with an EPA and/or DHA source (e.g., fish oil) in order to enhance the ability of the microorganisms to incorporate the substrate fatty acids into their lipids. An increase of up to 29% and 20%, respectively, in the EPA and DHA content in microbial oil was reported using such an approach (Shinmen et al., 1992; Watanabe et al., 1994). Isolated enzyme systems used for the concentration of n-3 PUFAs in plant and fish oils by hydrolysis (Yadward et al., 1991; Li and Ward, 1993a,b; Shimada et al., 1994) and for synthesis of n-3 PUFA-rich glycerols (Li and Ward, 1994a; Tanaka et al., 1994; Lee and Akoh, 1996), while more efficient, depend greatly on lipase specificities. Lipases of oleaginous microorganisms have not been extensively studied except in a few cases (Asmer et al., 1987; Hara and Nakashima, 1996; Lee and Akoh, 1996), although high levels of extracellular lipase have occasionally been reported for organisms grown in oil. V. Valuable Metabolites

Apart from the basic activity of oleaginous microorganisms to produce oil, they have the ability to synthesize a wide range of different lipid compounds with various structures and properties. These compounds, however, have not been studied as extensively as the productivity and composition of oil. No regular studies, except in a few cases, have been made to optimize the process for their production. Only since it was realized in the early 1990s that this group of compounds might offset the high cost of biotechnological production has some research work been undertaken. This holds true particularly for hydroxy longchain fatty acids and the studies on their biosynthesis in yeasts and fungi (Van Dyk et al., 1994). Much more work is needed to examine the use of oleaginous microorganisms for the biosynthesis of unusual lipids such as wax esters and hydroxyalkanoates and for catalyzing the formation of medically important products such as steroid hormones. Current knowledge about valuable metabolites of oleaginous microorganisms other than storage lipid is now presented in light of the previously mentioned potentials. A. FAT-SOLUBLE VITAMINS AND PIGMENTS The lipid classes synthesized by microorganisms include sterols, carotenoids, quinones, and other terpenoid lipids, some of which are known to have vitamin activities. However, relatively little is known about the microbial production of these compounds. After the initial

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interest in the 1970s and 1980s, biotechnoiogical processes for the production of such fat-soluble vitamins as @-carotene,vitamin A provitamin (Fig. 71, vitamin D (Fig. 8 ) ,vitamin E (Fig. 91, and vitamin K [Fig. 10)no longer occupied the same prominence. This might be due to poor strain productivity or difficulties with the harvesting of biomass and recovering the compound. Industrial processes for the production of p-carotene and vitamin D include a process for oil rich in @-carotene using the mucoral Blakeslea trispora or green alga Duniella salina. These processes compete successfully with chemical synthesis of this compound (Cerda-Olmedo and Alvalos, 1994; Cerda-Olmedo, 1989; Borowitzka and Borowitzka, 1989). A great potential appears to be associated with the mucoral Phycomyces blakesleeanus, which can also accumulate as much as 41% lipid in dry mycelium (Cerda-Olmedo and Alvalos, 1994). The fatty-acid composition and the ratio of free to esterified fatty acids in the lipids of this mold depend on the age of the culture and the growth conditions. Among the most abundant fatty acids (e.g., palmitic, linoleic, linolenic, and stearic acid), GLA is also present, the importance of which has already been discussed. The p-carotene content of the I? blakesleeanus mycelium varies with the growth conditions and the strain, and optimum conditions have not yet been determined. Phycomyces has also been the subject of genetic research (Cerda-Olmedo and Alvalos, 1994). As the genetics, biochemistry, and regulation of p-carotene production in Phycomyces are relatively well known, l? blakesleeanus offers great promise in production of GLA- and P-carotene-rich oil. Commercial production of the alga D. salina began in the mid-1980s (Borowitzka and Borowitzka, 1989), but the producers keep their processes confidential. D. salina cultures may be grown either in open ponds or in batch and continuous fermenters under light and at temperatures from 21 to 40°C and at pH 7-9, respectively, in media with appropriately adjusted salinity and a deficiency of nitrogen. Normal salinities used for growth vary from 20 to 30% (wt/vol) NaC1. As a photoautroph it can use only CO, as a carbon source. The cell densities in large cultures are about 1 g/liter. p-carotene represents 10% of the D.salina biomass and is accumulated as droplets in the chloroplast stroma. Ergosterol,vitamin D2 provitamin, is a major sterol in yeast and fungi, where its level is about 0.5% of dry cells, except in some of the Saccharomyces and Rhodotorula strains, in which it can account for 3 4 % dry weight (Margalith, 1989). Organisms capable of producing high levels of sterols also include some species of Candida, Fusarium, and Trichoderma and are described primarily in patents (Margalith,

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227

2

9

a

228 J. LEMAN

8

a

-a"B

POTENTIAL OF OLEAGINOUS MICROORGANISMS

CH2-[CH2-CH2-

CH -CH3I3-H

CH3

CH3

229

I

a-tocopherol, vitamin E FIG.9. Chemical structure of a-tocopherol (vitamin E].

1989). The production of sterols comprising 18-19% of the total lipids may be enhanced by a high concentration of carbohydrates, nitrogen starvation, precursors (e.g., isopentenol) in the medium, hydrocarbons, exposure to ethanol vapors, and aeration (Margelith, 1989). Ergosterol isolated from spent brewer’s yeast was used for the commercial production of ergocalciferol (vitamin D,), an analogue of cholecalciferol (vitamin DJ. Dehydrocholesterol is a precursor of cholecalciferol. Because vitamin D3 is more active than vitamin D,, some efforts have been made to produce vitamin D3 with yeast mutants deficient in specific desaturases (Grunwald-Raji and Margerith, 1990; Sekula, 1988). No optimized processes have so far been developed for vitamin E and K (Tani, 1989). The accumulation of tocopherols (vitamin E) has been demonstrated in chlorophyll-containing organisms, especially in the alga Euglena gracilis Z . Extensive screening studies for quinone (vitamin K)-producing microorganisms conducted in the 1980s in Japan succeeded in the selection of only Flavobacterium species as potent organisms. The culture conditions for vitamin K production by F: meningosepticum and its mutants are discussed i n detail by Tani (1989). Taxonomic studies have revealed new species of marine and soil microorganisms capable of quinone production. A new genus of marine budding phototrophic bacteria, Rhodobium gen. nov., including R. orientalis sp. nov. and R. marinum comb., produces menaquinone 10 (vitamin K,) and ubiquinone 10 (CoA), a substitute for vitamin K, as major quinones (Hiraishi et al., 1995). Menaquinone MK-9 was also found in four new species of the genus Actinokineospora isolated from soil (Tamura et al., 1995). Biosynthesis of the first aromatic intermediate in the vitamin K,, pathway, o-succinylbenzoic acid, has been demonstrated for the first time in Bacillus subtilis (Palaniappan et al., 1994). The biosynthesis of this intermediate has also been attempted genetically (Sharma et a]., 1993).

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0

I

&CH3

CH3

II 0

CH2CH=C(CH2),

I

CH3

I

CH3

I

CH3

CH(Ct 3 3 CH(CH2)a CHCHQ

vitamin K,

vitamin

K2

FIG.10. Chemical structure of phylloquinone (K, vitamin K,) and menaquinone (MKn, vitamin KJ.

B. OTHERLIPIDCLASSES

Several lipids produced by oleaginous microorganisms grown on hydrocarbons exhibit surface-active properties that arise from the presence of a hydrophilic moiety in their molecule. The hydrophilic moiety may consist of a variety of groups, including sugars, amino acids, or fatty acids. These surface-active lipids of oleaginous microorganisms are extracellular products of selected strains of Arthrobacter and Candida (Torufopsis)and include trehalose lipids, rhamnolipids, sophorose lipids, polysaccharide-lipid complexes, and neutral lipids. The ability of microorganisms to take up water-immiscible substrates such as alkanes indicates that a specific mechanism exists for the process. Since the discovery of this phenomenon in the 1970s, the mechanism has not been explained despite being intensively studied, employing both mutants and genetic manipulations (Eggink et d . ,1987; Koch et a]., 1988, 1991; Mulligan and Gibbs, 1989; Witholt et a]., 1990). Thus, two proposed hypotheses for the mechanism, that is, direct contact between cell surfaces and alkane droplets and interaction with pseudosolubilized droplets (Sing and Desai, 1986), await verification. Although originally considered agents for the cleanup of oil spills and for microbial-enhanced oil recovery (Shennan and Levi, 1987),biosurfactants from oleaginous microorganisms have fallen into disfavor be-

POTENTIAL OF OLEAGINOUS MICROORGANISMS

231

FIG.11. Chemical structure of sophorolipids

cause they are considered incompatible with nitrogen requirements for regulation of lipogenesis (Kosaric et al., 1984). Only two biosurfactants are produced commercially: one from Candida bombicola (formerly Torulopsis bombicola) for use in cosmetic products (Davies and Holdsworth, 1992), and the other from Acinetobacter calcoaceticus RAG-1 under the trade name Emulsan for use in cleaning oil-contaminated vessels and spills and for microbial-enhanced oil recovery (Fiechter, 1992). Further studies on Emulsan production with A. calcoaceticus RAG-l mutants are continuing to yield interesting results on properties (Pines and Gutnick, 1986; Shabtai and Gutnick, 1986), biosynthesis (Leahy, 1994), and culturing methods (Brown and Cooper, 1992). Numerous papers still appear on sophorose lipids (Fig. 11)produced by Candida bombicola (Krivobok et al., 1994; Hommel et a]., 1994; Hommel and Huse, 1993; Zhou and Kosaric, 1993; Walsum and Cooper, 1993; Lee and Kim, 1993; Klekner et a]., 1991). Studies on their production, antibiotic properties, and cytotoxicity revealed a 60-70% yield on olive oil, an antibacterial activity against Staphylococcus Q U reus, and potential acute toxicity to human fibroblasts (Krivobok et ~ l . , 1994). Interfacial properties equivalent to those of other biosurfactants, and antibacterial activity particularly against Gram-positive bacteria, of two types of mannosylerythritol lipids produced from soybean oil by Candida antarctica have also been reported (Kitamoto et al., 1993).

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The discovery of surface-active lipids in oleaginous microorganisms has led to rapid development of biosurfactants from species considered nonoleaginous such as Bacillus, thus advancing significantly both product technology and knowledge of microbial physiology (Fiechter, 1992; Hommel and Ratledge, 1993). An interesting bacterial product is poly-P-hydroxybutyrate (PHB), which, although it is not a fatty acid-containing lipid, is classified as a lipid because of its solubility characteristics. Like the lipids of microorganisms, PHB is produced in an increased amount when nitrogen is exhausted from the medium. The best producing microorganism is Alcafigenes eutrophus, which accumulates PHB up to 80% of its biomass when grown on glucose medium (King, 1982).This compound has excellent emulation and thermoplastic properties and is fully biodegradable. A. eutropus is used for the commercial production of a copolyester of 3-hydroxybutyrate and 3-hydroxyvalerate (Byrom, 1987). Further studies on PHB and its copolymers have been carried out in conjunction with an investigation of the technological aspects of production (Eggink et al., 1993; Seung-Yoo and Woo-Sik-Kim, 1994; Sheppard et al., 1994; Boom-Soo-Kim et al., 1994; Tanaka et al., 1993; Linko et al., 1993), recombinant bacteria (Hong-Zhang et af., 1994; Lee et af., 1994; Kim et a]., 1992), biosynthesis (In-Young-Lee et a]., 1993; Steinbuchel and Schlegel, 19911, new substrates (Eggink et a]., 1993; Alderete et al., 1993), and culture conditions (Volova et al., 1992). Rhodococcus ruber NCI MB40126 has been found to produce similar copolymers (Haywood e f al., 1991) with a higher proportion of 3-hydroxyvalerate (Anderson et al., 1992). The production of waxes, phospholipids, sphingolipids, and unusual fatty acids using oleaginous microorganisms needs to be explored. Wax esters based on long-chain fatty acids and alcohols have industrial application in cosmetics, pharmaceuticals, paints, and lubricants because of their wetting properties at interfaces. Significant amounts of waxes, up to 15% of the cell biomass, are produced by the bacterium Acinetobacter grown on hydrocarbons (Finnerty, 1984). These waxes are similar to sperm whale oil and jojoba oil (Dewitt et al., 1982; Ervin et a f . , 1984). The modification of wax composition can be achieved by varying the nature of the carbon source in the medium and the growth temperature. In general, C32-C42 waxes can be produced for varying the chain length of the carbon source and the temperature (Rattray, 1984).Wax ester production by Euglena gracifis has been reported (Ono et nf., 1995). When aerobically grown, the algal cells synthesized and intracellularly accumulated the wax esters having a carbon chain length of 24-32 from paramylon, the reserve polysaccharide of this organism.

POTENTIAL OF OLEAGINOUS MICROORGANISMS

233

C20 PUFA

I 20:3 n-6

DHGLA

I

1

20:4 n - 6

20:s n - 3

AA

EPA

I Eicosanoids

Cell function regulation FIG. 12.

Formation of hydroxy polyunsaturated fatty acids (eicosanoids) from C20

PUFAs.

Phospholipids and glycosphingolipids occur in all living organisms and can be incorporated to a large extent in biological membranes, where they influence such membrane functions as the regulation of temperature sensitivity, substrate uptake, or membrane protein activity by a subtle choice of lipid type. Phospholipids may reach up to 30% of the cell biomass in yeast grown on hydrocarbons (Voigt et al., 1979). Because of their surface-active properties due to the presence of a large hydrophilic moiety such as choline, they may also offer a variety of applications, including the production of artificial membranes (liposomes). The accumulation of cerebrosides in some n-alkane-assirnilating yeasts has been reported (Ratledge and Evans, 1989; Mineki et al., 19941, but generally not much information is available. The possibility of oleaginous microorganisms converting n-alkanes to industrially important hydroxy or dicarboxylic acids is well known (Ratledge, 1984a). Hydroxy fatty acids formed via oxygenase reactions in the cell are used in cosmetics, paints and coatings, lubricants, and

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the food industry. They are useful chemical intermediates in the lipasecatalyzed synthesis of fine chemicals and pharmaceuticals, in which they occur as estolides, lactones, or polyolides. The oleaginous mold Claviceps, which accumulates u p to 60% lipid, of which 12-hydroxystearic acid (ricinoleic acid) makes up to SO%, would be an attractive source of this acid if not for difficulties in the culturing organism and lipid recovery (Ratledge, 1984a). Other organisms known to accumulate hydroxy fatty acids are some species of Candida (Torulopsis),Rhodotorula, Ustilago, and Yarowia, which produce di- and trihydroxy fatty acids and o-hydroxy fatty acids. Hydroxy polyunsaturated fatty acids (Fig. 1 2 ) are important because of their potential medical use. The occurrence, metabolic pathways, and biological activity of hydroxy polyunsaturated fatty acids has been reviewed by Van Dyk et al. (1994). Conversion of alkanes to a,o-dicarboxylic acids in yields of up to 70% and with amounts up to 60 g/liter have been reported with mutants of Candida cloacae and C. tropicalis (Hill et al., 1986). The latter organism is used for commercial production of brassylic (13-dicarboxylic) acid (Davies and Holdsworth, 1992). Short- and medium-chain fatty acids (C6-C12) produced by some Entomorphthora species (Boulton and Ratledge, 1984) deserve attention for their possible use in structuring the medium-chain triacylglycerols (MCTs), known for their nutritional value (Latta, 1990). Fatty acids and fatty alcohols (C8-Cl8) can be readily oxidized by selected bacteria and yeasts or their enzymes, yielding dicarboxylic acids and hydroxy fatty acids (Ratledge, 1992b; ElSharkawy et al., 1992; Esaki et al., 1994). Biotransformations of other lipid substrates, and particularly of steroids, carried out with immobilized cells of Rhodococcus, producing extra- and intracellular cholesterol oxidases (Kreit et al., 1994; Wilmanska et al., 1995), enable the formation of value-added products with potential for industrial (ChungYoung-Wu et al., 1995) and medical (Warhurst and Fewson, 1994) applications. VI. Conclusions

The knowledge acquired so far about oleaginous microorganisms and the biochemistry of lipid accumulation in particular provides for sharing some optimism about their future prospects. Current achievements reflected in the development of processes for SCO, CBEs, and GLA-rich oils only confirm the potential of the microorganisms and create new perspectives that show the importance of economic requirements. Further opportunities for utilizing oleaginous microorganisms include

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INDEX

A Acanthamoeba spp. biological aspects, 38-40 disinfection, 44-46 ecology, 42-44 eye infections from, 40-44 taxonomy, 35-38 ACE inhibitors, 101-107 Acetic acid aerobic process, 3-4 anaerobic process, 4-5 bioreactor production for, 24-29 downstream processing, 29-30 fermentation broths, 29-30 production, history, 1-3 Acetophthalidin, 70 Acetyl xylan esterases, 162 Acinetobacter calcoaceticus, 110 Acinetobacter spp., 232 Actinomycetes, 60-63, 62 Aerobic process, 3-4 Agrobacteriurn spp., 80 Alcaligenes eutrophus, 232 Alcohol dehydrogenases, 124 Alk-1-enes, 132 Altemicidin, 61 Alteramide A, 66-67 Alteromonas haloplanktis, 62-63 Alteromonas rubra, 59-60 Alteromonas spp., 65, 66-67 Amidase, 119 Amino acid synthesis, 113-120 Aminotransferases, 119 Anaerobic process, 4-5 Andrimid, 64 Angiotensin converting enzyme, see ACE inhibitors Anguibactin, 64 Antiarrhythmic agents, 125-127 Antibiotics, see specific antibiotic Anticholesterol drugs, 107-113 Antiinfective drugs, 113-120 Antiinflammatory drugs, 129-130 Antipsychotic agents, 121-125

Antiviral agents, 130-131 Aplasmomycins A-C, 61 a-arabinosidases, 155-156 Aromatic acids (C16), 59 Aspergillus oryzae, 160

B Bacillus spp., 165 Bacteria, see specific bacterium BIOLOG system, 72 Bioreactors, 24-29 Biotechnology, 86-87 Bioxalomycins, 81-84 Bisucaberine, 62-63 Bleaching benefits, 176-178 brown stock stage, 170 factors affecting, 170-171 preparation, 167-169 shive removal, 182 xylanase production for, 169-17 BMS lipase Pseudomonas cepacia, 95-98 racemic acetylation by, 110-113 Brain, see specific regions Brominated compounds, 59 Brominated diphenyl ethers, 66 Broths, acetate, 29-30

C Calcium channel blockers, 120-121 Calcium magnesium acetate, 2 Candida spp., 231 Caprolactams A-B, 60 Captopril, 101-103, 102 p-carotene, 226 CBEs, see Cocoa butter equivalents Ceranopril, 107 Chiral diol, 129 Chiral P-hydroxy, 107-108 Chiral synthons, 92, 94 245

246

INDEX

S-2-chloropropionic acid, 130 ChromaXome, 86 CloneZyme libraries, 87 Clostridium thermoaceticum fermentation mineral requirements, 12-13 process, 4-5 substrates, 6-12 low-cost media, 16-24 strain media, 13-16 CMA, see Calcium magnesium acetate Cocoa butter equivalents, 206-210 Combinatorial biology, 86 Contact lens, 43-51 Coriolin B, 70 Cnriolus coiisors, 70 Corn gluten, 17 Corn steep liquor (CSL), 19-24 Cyclosporin A, 118-119 Cysteine, 13

Ethanoic acid, see Acetic acid Euglena gracilis, 232 Eyes contact lens, 44-51 diseases, see specific disease infections, 40-44

F

Fats, see specific fats Fat-soluble vitamins, 225-229 Fed-batch reactors, 24-29 Fellutamides A-B, 69-70 Fermentation acetate broths, 29-30 acetic acid, 1-3 aerobic process, 3-4 anaerobic process, 4-5 Closfridjjum th erm oa ceticu m bioreactors for, 24-29 low-cost media, 16-24 D mineral requirements, 12-13 process, 4-5 Deacetylase (ClO), 93-94 strain media, 13-16 substrates, 6-12 Debarking, 178-179 Debromomarinone, 62 corn products for, 17-24 marine microorganisms, 73-75 Dendryphiellin A-E, 68-69 pharmaceutical intermediates, 91-92 Dicarboxylic acids, 106 soybean for, 23-24 Didanosine, 130-131 Fiber modification, 179-181 Dihydrocoriolin C, 70 Filamentous fungi, 164 Diketopiperazines, 66 Flavo ba cterium ugligin m u m , 63 Dilthiazem, 120-121 2,~-dimethyl-~,3-dioxolane-4-methanol,Flax fiber retting, 182 Food production, 183-184 126 Formate, 14-16 DNA libraries, see specific libraries Fungi, see also specific fungus Downstream processing, 29-30 Draw-and-fill bioreactor, 28-29 bioactive metabolites from, 68-70 filamentous. 164 Drugs, see specific drug classifications

E Electrodialysis (ED), 30 Encephalitis, see Granulomatous amoebic encephalitis D-enzyme 1-2,109 Esterases, 158-163

G Geotrich uni carzdidum, 108-109 a-glucuronidases, 156-157, 166-167 Glycosphingolipids, 233 Granulomatous amoebic encephalitis, 40-42

247

INDEX H Halichomycin, 64 Halocyphina villosa, 68 Hansenula polymorpha, 125-126 Hansenula spp., 94-95 Heliascolides A-B, 69 Helicascus kanaloanus, 69 Horse liver alcohol dehydrogenase, 99-100 Hymenoscyphus spp., 69 Hyphomycetes, 70 Hypoxylon oceanicum, 84-86

I Ibuprofen, 129-130 2,3-indolinedion,65 Indolizamycin, 61 Infections, see Eye infections Isatin, 65 Isochrysis galbana, 224 Istamycins A-B, 60-61

7 Juice clarification, 182-183

K Keratitis, 4 0 4 4 Keratoconjunctivitis, 40 Kirschsteinin, 69 Kirschsteiniothelia spp,, 69 Kraft pulp, see Pulp

L Lactam, 131 Lagunapyrones A-C, 62 Lamivudine, 130-131 Lens, see Contact lens Leptosins, 70 Leptosphaeria obiones, 69 Leptosphaeria oraemaris, 68 Leptosphaeria spp., 70

Leptosphaerin, 68 Lipase PS-30 Pseudomonas cepaciu, 95-98, 101 racemic acetylation by, 110-113 Lipids in oleaginous microorganism production characteristics, 203 classification, 230-234 history, 195-197 process, 197-203 LL-31F508, 80-81 LL-l5G2561~, 84-86 LL-141352, 79 LL-211457, 80 LL-l41352~t,79 Loloatin B, 64

M Macrolactins A-F, 51-60 Maduralide, 62 Marinactan, 63 Marine microorganisms, see also specific microorganisms biological activities, 74-75 BIOLOG system, 72 biotechnology, 86-87 classification, 72-73 fermentation, 73-75 isolation, 70-73 MIDI system, 72 natural products free-living metabolites, 58-63 fungi, 68-70 symbiotic bacteria, 65-68 production, history, 57-58 symbiotic, 75-84 taxonomy, 70-73 Wyeth-Ayerst program, 70-73 Marinone, 62 Methyl carboxylic acid, see Acetic acid Mevastatin, 113 Microbial oil, 197 Micrococcus spp., 66 Microcoleus lyngbyuceus, 65-66 Microorganisms, see Marine microorganisms; Oleaginous microorganisms MIDI system, 72

248

INDEX

Moiramides A-C. 64 Moraxella spp., 68 Mortierella mmanniana, 123-124, 1 2 7 Mortierella spp., 220-221

N

Nanofiltration, 30 Naphthoquinones, 69 Naproxen, 129 Neutral endopeptidase inhibitor, 103-106 Nocardia corallina, 132 Nocardioides spp., 93-94

0 Obionin, 69 Obioninene, 68 Octalactins, 63 Oils, see specific oils Oleaginous microorganisms cocoa butter equivalents, 206-210 fats, specialty, 206 fat-soluble vitamins from, 225-229 lipid production, 230-234 characteristics, 103 history, 195-197 process, 197-203 oils from microbial, 197 specialty, 206 pigments from, 225-229 polyunsaturated fatty acids, 210-214 17-3PUFA, 219-225 n-6 PUFA, 214-219 single cell oil, 203-206 Oreamarin, 68 (exo,exo)-7-oxahicyclo[2,2 ,l]heptane-2,3dimethanol, 100 Oxidoreductases, 108-109

Penicillium spp., 70 Phaeodactylum tricornutum, 222 PIIB, see Poly-P-hydroxybutyrate Phenazines, 61-62 Phomactins, 69 PhoJna spp., 69 Phospholipids, 233 Phycornyces blakesleean us, 2 26 Pigments, 2 2 5-2 29 Plant cell walls, 142-149 PM-93135, 64-65 Poly-P-hydroxybutyrate, 232 Polysyncration lithostroturn, 75, 79-80 Polyunsaturated fatty acids, 210-214 Potassium channel openers, 127-129 Pravastatin, 113 Prostaglandin synthesis, 132-133 Protoplasts, 61 Pseudornonas cepacia, 101 Pseudomonas spp., 95-97, 119 PUFAs, see Polyunsaturated fatty acids Pulp bleaching debarking, 178-179 dissolving, 181-182 shive removal, 1 8 2 with xylanases benefits, 176-178 brown stock stage, 170 factors affecting, 170-171 preparation, 167-170 Purine nucleoside analogues, 130 Pyruvate, 14-16

Q Quinone, 66

R R-[+)-BMY-14802,124 Recombinant BioCatalysis, 86-87 Retting, 182

P Paclitaxel, see Taxol Paper, see Pulp Penicillium citrinum, 113 Penicilliurn fellutan urn, 69-70

S

Saccharomyces cerevisiae, 109 Salinamides A-B, 63-64

249

INDEX Saxitoxin, 68 Schizophyllum commune, 164-165 SCO, see Single cell oil Sesquiterpene diol culmorin, 68 Shewanella pufrefaciens,222 Shive removal, 182 Siccayne, 68 Sigma receptor system, 121,123 Single cell oil, 203-206 Solketal, 126 Soybean meal, 23-24 Sponges, 66 Stillage, filtered, 19 Streptomycetes, 60-61 Streptomyces carbophilus, 113 Streptomyces sioyaensis, 61 Streptomyces spp., 159-160,166 Streptomyces viridodiastaticus, 80-81 Surugatoxin, 67 Symbiotic bacteria, 65-68

T Taxolase (C13),93-94 Tax01 semisynthesis, 92-98 T ~ X USPP., S 93-94 Thiotropocin, 80 Thromboxane A2 antagonists, 98-101 2,3,4-tribromo-5-(1‘-hydroxy-2’,4’dibromophenyl)pyrrole, 58-59 Trichoderma reesei, 159,166 Trichoderma spp., 149-154 Trophozoites, 43-44,46-50 Two-stage membrane bioreactor, 27-28 Tyrosol (2-p-hydroxyphenyl ethanol), 65

U

Uveitis, 40

Vitamin K, 229 Vitamins, fat-soluble, 225-229

W Waxes, 232 Wyeth-Ayerst (W-AR) Marine Natural Products Program, 70-73

X Xylan biological aspects, 141-142 enzymatic systems, properties, 149 plant cell wall interaction, 142-149 structure, 142-149 Xylanases, 149-154 debarking with, 178-179 fiber modification with, 179-181 flax fiber retting, 182 food production, 183-184 juice clarification, 182-183 pulp bleaching benefits, 176-178 brown stock stage, 170 factors affecting, 170-171 microorganisms for, 169-1 70 preparation, 167-169 pulp dissolving, 181-182 shive removal, 182 Xylanolytic enzymes, 163-167 P-xylosidases, 154-155

Y Yeasts, lipid accumulation, 195-197

V Z

Vibrio sp., 67 Vitamin E, 229

Zidovudine, 130-131

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CONTENTS OF PREVIOUS VOLUMES

Volume 33

The Cellulosome of Clostridium thermocellum Raphael Lamed and Edward A. Bayer Clonal Populations with Special Reference to Bacillus sphaericus Samuel Singer Molecular Mechanisms of Viral Inactivation by Water Disinfectants R. B. Thurman and C. P Gerba

Microbial Production of Gibberellins: State of the Art I? K. R. Kumar and B. K. Lonsane Microbial Dehydrogenations of Monosaccharides MiloS Kulhdnek Antitumor and Antiviral Substances from Fungi Shung-Chang Jong and Richard Donovick Biotechnology-The Golden Age I.! S. Malik

Microbial Ecology of the Terrestrial Subsurface William C. Ghiorse and John ?: Wilson

INDEX

Volume 35

Foam Control in Submerged Fermentation: State of the Art N. I? Ghildyal, B. K. Lonsane, and N. G. Karanth Applications and Mode of Action of Formaldehyde Condensate Biocides H. I% Rossmoore and M. Sondossi Occurrence and Mechanisms of Microbial Oxidation of Manganese Kenneth H. Nealson, Bradley M. Teho, and Reinhardt A . Rosson Recovery of Bioproducts in China: A General View Xiong Zhenping INDEX

Volume 34

What’s in a Name?-Microbial Secondary Metabolism J. I% Bennett and Ronald Bentley

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 I% Han Review and Evaluation of the Effects of Xenobiotic Chemicals on Microorganisms in Soil R. J. Hicks, G. Stotzky, and I? Van Voris Disclosure Requirements for Biological Materials in Patent Law Shung-Chang Jong and Jeannette M. Birmingham INDEX

251

252

CONTENTS OF PREVIOUS VOLUMES

Volume 36

Microbial Transformations of Herbicides and Pesticides Douglas J. Cork and James E! 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 James F. Kane

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 Somasundaram Rajarathnam, Mysore Nanjarajurs Shashirekha, a n d Zakia Bano INDEX

Microbial Cytochromes P-450 and Xenobiotic Metabolism F: Sima Sariaslani Foodborne Yeasts Z Decik High-Resolution Electrophoretic Purification and Structural Microanalysis of Peptides and Proteins Erik l? 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 a n d H. C. van der Plas Medicinal Benefits of the Mushroom Ganoderma S. C. Jong and J. M. Birmingham

Volume 38

Selected Methods for the Detection and Assessment of Ecological Effects Resulting from the Release of Genetically Engineered Microorganisms to the Terrestrial Environment G. Stotzky, M. W Broder, J. D. Doyle, a n d R. A . Jones Biochemical Engineering Aspects of Solid-state Fermentation M. V Ramana Murthy, N . G. Karanth, and K. S. M. S . Raghava Rao The New Antibody Technologies Erik l? Lillehoj and Vedpal S. Malik Anoxygenic Phototrophic Bacteria: Physiology and Advances in Hydrogen Production Technology K. Sasikala, Ch. V Ramana, l? Rahuveer Rao, and K. L. Kovacs INDEX

Volume 39

Asepsis in Bioreactors M. C. Sharma and A . K. Gurtu

CONTENTS OF PREVIOUS VOLUMES Lipids of n-Alkane-Utilizing Microorganisms and Their Application Potential Samir S . Radwan and Naser A. Sorkhoh Microbial Pentose Utilization Prashant Mishra and A j a y Singh Medicinal and Therapeutic Value of the Shiitake Mushroom S. C. Jong and J. M. Birmingham Yeast Lipid Biotechnology Z. Jacob Pectin, Pectinase, and Protopectinase: Production, Properties, and Applications Takuo Sakai, Tatsuji Sakamoto, Johan Hallaetf, and Erick J. Vandamme Physicochemical and Biological Treatments for Enzymatic/Microbial Conversion of Lignocellulosic Biomass Purnendu Ghosh and Ajay Singh INDEX

Volume 40

Microbial Cellulases: Protein Architecture, Molecular Properties, and Biosynthesis Ajay Singh and Kiyoshi Hayashi Factors Inhibiting and Stimulating Bacterial Growth in Milk: An Historical Perspective D. K. O’Toole Challenges in Commercial Biotechnology. Part I. Product, Process, and Market Discovery AleS Prokop

253

Challenges in Commercial Biotechnology. Part 11. Product, Process, and Market Development AleS Prokop Effects of Genetically Engineered Microorganisms on Microbial Populations and Processes in Natural Habitats Jack D. Doyle, Guenther Stotzky, Gwendolyn McClung, and Charles W Hendricks Detection, Isolation, and Stability of Megaplasmic-Encoded Chloroaromatic Herbicide-Degrading Genes within Pseudornonas Species Douglas J. Cork and Amjad Khalil INDEX

Volume 41

Microbial Oxidation of Unsaturated Fatty Acids Ching ?: Hou Improving Productivity of Heterologous Proteins in Recombinant Saccharomyces cerevisiae Fermentations Amit Vasavada Manipulations of Catabolic Genes for the Degradation and Detoxification of Xenobiotics R u p Lal, Sukanya Lal, I! S. Dhanaraj, and D. M. Saxena Aqueous Two-Phase Extraction for Downstream Processing of EnzymesIProteins K. S. M. S . Raghava Roo, N . K. Rastogi, M. K. Gowthaman, and N . G. Karanth

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CONTENTS OF PREVIOUS VOLUMES

Biotechnological Potentials of Anoxygenic Phototrophic Bacteria. Part I. Production of Single Cell Protein, Vitamins, Ubiquinones, Hormones, and Enzymes and Use in Waste Treatment Ch. Sasikala and Ch. V: Ramana Biotechnological Potentials of Anoxygenic Phototrophic Bacteria. Part 11. Biopolyesters, Biopesticide, Biofuel, and Biofertilizer Ch. Sasikala and Ch. V: Ramana INDEX

Volume 42 The Insecticidal Proteins of Bacillus thrtringiensis I! Ananda Kurnar, R. I! Sharma, and I/: S. Malik Microbiological Production of Lactic Acid John H. Litchfield Biodegradable Polyesters Ch. Sasikala The Utility of Strains of Morphological Group I1 Bacillus Samuel Singer Phytase Rudy J. Wodzinski and A . H. J. Ullah INDEX