Advances in Applied Microbiology, Volume 47

Seeing Red: The Story of Prodigiosin J. w. BENNETT

Department of Cell and Molecular Biology Tulane University New Orleans, Louisiana 70118

RONALD BENTLEY

Department of Biological Sciences University of Pittsburgh Pittsburgh, Pennsylvania 15260

I. Bread, Blood, and Bacteria II. Early Instances of "Blood" on Bread III. Red Bacteria and the History of Bacteriology A. Pre-Pasteurian Research B. Pigments and Paintings C. The Genus Serratia IV. Prodigiosin and Related Compounds A. Structures B. Biosynthesis V. From Saprophyte to Pathogen VI. Biological Activity of Prodigiosin and Related Compounds A. Possible Ecological Functions B. Pharmacological Activity VII. Final Comments References

I. Bread, Blood, and Bacteria Bread, b o t h l e a v e n e d a n d u n l e a v e n e d , p l a y s a crucial nutritional, religious, a n d e m o t i o n a l role in h u m a n lives. In the Old Testament, b r e a d is said to " s t r e n g t h e n e t h m a n ' s heart" (Psalms 104:151), a n d in the Lord's Prayer the request is "Give us this d a y o u r daily b r e a d " ( M a t t h e w 6:11). In J u d a i s m , u n l e a v e n e d b r e a d is the c e n t e r p i e c e of the P a s s o v e r meal. In Christianity, the Eucharist or s a c r a m e n t of the Lord's S u p p e r is c e l e b r a t e d t h r o u g h the c o n s e c r a t i o n a n d c o n s u m p t i o n of b r e a d a n d

1Biblical quotations are from the Authorized King James Version.

ADVANCES IN APPLIED MICROBIOLOGY, VOLUME 47 Copyright © 2000 by Academic Press All rights of reproduction in any form reserved. 0065-2164/00 $25.00

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J.w. BENNETT and RONALD BENTLEY

wine: "And as they did eat, Jesus took bread, and blessed, and brake it, and gave to them, and said, Take, eat: this is my body" (Mark 14:22). Bread is also, especially when not dried out, an excellent culture medium for the growth of many microorganisms, so much so that many present-day commercial breads contain calcium propionate "added to retard spoilage." In the pre-antibiotic era, microbial contamination of bread was used to good effect: the healing of wounds was facilitated by application of preparations made from moldy bread. A specific and early example is documented in an English herbal of 1760. Such preparations may well have contained penicillin, patulin, or other antibiotic materials formed by the fungi (Wainwright, 1990). However, in most cases when microbes use bread as a substrate for their growth, the result is spoilage. Contaminated breads can be detected by repellent flavors and distinctive coloration. Most spoilage of bread is caused by fungi: Aspergil]us niger forms black colonies, many members of the genus Penicillium are blue or green, while certain yeasts and bread molds such as Neurospora crassa form pink to red pigments. Bacteria are less commonly associated with deterioration of bread; however, under warm and humid conditions some strains of Serratia marcescens form distinctive red colonies on this substrate. The red color derives from the presence of the pigment prodigiosin and/or related materials (see later). As the bacterial colonies reach maturity, they dissolve into a fluid and viscous state with a mucilaginous appearance and an uncanny resemblance to blood. Indeed, from early times, there are many records of the appearance of "blood" on bread, beans, and other starchy foods such as polenta and potatoes. Like bread, blood is a substance with profound cultural implications beyond its physiological role. Human and animal sacrifice were practiced in many societies with the intent to propitiate the wrath of an all-powerful deity. The victim's blood was often associated with a mystical power. For the Aztecs, the sun god (Huitzilopochti) drove back the moon and stars each day. To carry out this tremendous task, he had to be nourished with human blood. In some cultures, prisoners of war were sacrificed and their blood consumed by the executioners, while in other cultures the drinking of blood was taboo (e.g., the ritual slaughter of animals by exsanguination as practiced by the Jews). The Old Testament is filled with blood imagery and stories of ritual sacrifice. During the momentous Passover devastation of all the firstborn (both men and beasts) in the land of Egypt, the Israelites were protected by the blood of an unblemished, 1-year old, male lamb spread on the side and upper door posts of their houses (Exodus 12). The paschal (Passover) lamb was a term later applied symbolically to Christ.

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To this day, many Christians proclaim that for believers redemption from sins is only possible by way of "the blood of the Lamb". At the Last Supper, Christ used wine to symbolize his blood: "This cup is the new testament in m y blood, which is shed for you" (Luke 22:20). The Roman Catholic faith has embraced the belief in transubstantiation, whereby the bread and wine of the Eucharist actually turn into the body and blood of Christ. This formal doctrine was specifically defined at the Fourth Lateran Council (1215) and reaffirmed at the Council of Trent (1551) (Cross and Livingstone, 1974). As noted above, mature colonies of pigmented Serratia are eerily bloodlike in appearance. More than a few microbiologists have hypothesized that the growth of these bacteria could be interpreted, in certain religious or symbolic contexts, as the miraculous appearance of blood. This paper discusses the possible role of Serratia m a r c e s c e n s in forming bloodlike material on starchy foods, and reviews many of the unusual properties of this fascinating bacterium and the red pigment(s) that it and other microorganisms form. In reviewing the historical record, we have made extensive use of previous publications (Harrison, 1924; Reid, 1936; Gaughran, 1969; Yu; 1979; Cullen, 1994). 2 II. Early Instances of "Blood" on Bread

It is impossible to know who first observed foodstuffs apparently carrying drops of blood. Red-spotted bread was probably observed in many parts of the world; however, only in European countries is there an extensive written record and only there did it come to play a role in religious controversy. The first known recorded report dates from Alexander the Great's siege of Tyre in 322 BCE. The disgruntled Macedonian troops were tired of the siege, w h e n a soldier noticed a trickle of blood inside a piece of broken bread. A soothsayer named Aristander interpreted the event as a good omen, opining that, had the droplets of blood been on the outside, the Macedonians would have been endangered. Since the flow was from the inside, it was an omen that Tyre would fall. Almost certainly, Aristander was well compensated for his ability to both calm the troops and prophesy the future. Christian Gottfried Ehrenberg (1795-1876) collected almost 100 European reports of the occurrence of blood, starting in the eleventh century; an English summary was provided by Gaughran (1969). Significantly, 2Unless otherwise apparent, the simple word "blood" will carry the meaning of "blood" or "bloodlike materials" to avoid much repetition. The genus abbreviation S. will refer to Serratia, and Streptomyces will not be abbreviated.

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blood was frequently observed on, or flowing from, the bread or wafers used as the Host in the Eucharistic liturgy. The prototype of many examples was recorded from Alsen, Denmark, in 1169. A village priest saw blood on a Host. Upon reporting this event to his superiors, the Chief Priest predicted the imminent shedding of Christian blood. A few days later, a pagan army overthrew churches, drove people into slavery, and killed those who resisted. Within several decades after the Alsen event, a strange myth grew up around reports of blood on Communion hosts. In 1247, near Berlin, a w o m a n removed a consecrated wafer from her mouth and sold it to Jews, who "stabbed it," resulting in the appearance of blood. The wafer was returned to the church, bringing it much fame, and the Jews were apparently unharmed. However, in other similar stories, Jews were persecuted and killed. Thus, in 1296 near Frankfurt, a purportedly stolen wafer was sold to Jews, stabbed, and yielded blood. A mob subsequently marched with banners, attacking Jews in Nuremberg, Rothenburg, Wfirzburg, and elsewhere, with a reported death toll of 10,000. Similar stereotypic reports of bloody Communion wafers led to repeated tormentations and executions of Jews for at least 200 years. Ominously, the geography of the persecutions overlapped that of the atrocities of the Holocaust several centuries later, with most of the incidents reported from cities in Germany and Poland. The number of those who perished will never be known. Scientists who believe that the explanation for bleeding hosts is the growth of Serratia are fond of quoting Scheurlen (1896), an early German observer: "dieser Saprophyt mehr menschen umgebracht hat als mancher pathogene Bacillus," or, in Isenberg's translation (1995), "This saprophyte has killed more humans than some pathogenic bacilli." In retrospect, whether or not the stories of red spots on Communion wafers were real or fabricated, the interpretation of the events and the reprisals against the Jews seem to make little sense. The Middle Ages was, of course, a time of many superstitions and profound religious belief. One wonders, however, w h y the Jews were not hailed as heroes for showing that the Host w o u l d bleed, thus providing a vivid demonstration of the truth of transubstantiation. It was nonsense to think that Jews wished to drink any sort of blood, since such an action was specifically forbidden to them. Deeper irrationalities were involved, with the incidents serving as a pretext to express a violent antisemitic prejudice. Eventually, the legend of the "blood libel" developed, which held that Jews needed the blood of children to make bread for Passover or for sorcery-related medicinal purposes; more recently, this libel was

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part of the many grotesqueries incorporated into Nazi propaganda (Trachtenberg, 1943). On the other hand, in a Christian context, the presence of blood on sacramental bread was interpreted in support of the doctrine of transubstantiation. In fact, one such incident became perhaps the most celebrated miracle of the thirteenth century (Cullen, 1994). According to Church history, a German priest with doubts about the doctrine of transubstantiation once celebrated mass at the Church of Santa Cristina in Bolsena, Italy in 1263. When blood dripped from the Host onto the altar linen and his vestments, his doubts were resolved and he sought absolution for his lack of faith. This event became celebrated as "The Miracle of Bolsena" and was later depicted in a Vatican fresco by Raphael. To commemorate the miracle, Pope Urban IV issued a bull that instituted the Feast of Corpus Christi and later decreed the construction of a n e w cathedral in Orvieto in which the host and vestment linens are preserved to this day. It has been suggested that the relics provide a tantalizing experimental opportunity. If enough DNA could be isolated from them, the polymerase chain reaction could be used to test the hypothesis that Serratia marcescens was involved in this medieval miracle. III. Red Bacteria and the History of Bacteriology A. PRE-PASTEURIAN RESEARCH

A giant leap forward in the understanding of microbiology in general and the formation of red-pigmented materials on foodstuffs in particular began with yet another event in Italy, this time in Legnaro (province of Padua) in 1819. The affected foodstuff was a bloody polenta (corn mush, corn porridge) found in the squalid home of a superstitious farmer named Antonio Pittarello. Eventually, more than 100 families in the region reported bloodlike materials on polenta or rice soup. A cooked chicken was described as "dripping with blood." Maleficent spirits were blamed for the event, and families who found bloodlike spots on their food were accused of evil activities. The event caused so much publicity that an official investigation was established under the direction of Dr. Vincenzo Sette, the medical officer at Pione di Sacco. Sette concluded that a fungus was responsible, while a botanist-priest, Pietro Melo, "claimed that the phenomenon was due to a spontaneous fermentation of the polenta which caused the corn meal to be transformed into a colored mucilage" (Breed and Breed, 1924). Sette eventually published his report in 1824, calling the organism Zaogalactina

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imetropha (from the Greek, "living slime situated on food"). On one occasion, he reddened polenta in a priest's house, thereby, according to the story, disposing of the theory that the phenomenon could only occur in the house of a sinner (it is a tribute to local piety that no one suggested the possibility that the priest was less than perfect!). Meanwhile, a pharmacist, Bartolomeo Bizio (then a student and later professor at the University of Padua), independently examined the red potenta, giving preliminary and detailed accounts of his work in 1819 and 1823 (Merlino, 1924). Bizio also classified the organism as a fungus and coined the further name Serratia marcescens. He used Serratia to honor a physicist, Serafino Serrati, who had run a steamboat on the Arno in 1787. Bizio believed that Serrati had a prior claim over "a foreigner" (presumably James Rumsey) as inventor of the steamboat and wished to honor his countryman. The second part of the binomial, marcescens, came from the appearance of the mature colonies that dissolved into "a fluid and viscous matter which has a mucilaginous appearance." Marcescens is the present participle of the Latin verb meaning "to decay or wither." Bizio performed experiments in which he used paper soaked with the red substance, or bits of red polenta, to transmit "seeds" of his fungus. As did Sette, Bizio made an honest mistake in identifying the causative bacterium as a fungus, but we are indebted to them for laying sure foundations for further investigations. Both Sette and Bizio were the first to provide evidence suggesting that the bloody material on food was due to a living organism, similar to the alga that caused pink snow on mountains, and transmissible from substrate to substrate by inoculation. Many years later, the spoilage of corn by fungal growth was investigated in connection with pellagra. With whole corn, "sometimes the embryo is colored reddish by Micrococcus prodigiosus"--one of the many binomials applied to S. marcescens (Black and Alsberg, 1910). These authors, however, did not refer to the spoilage of polenta. They are known for their discovery of penicillic acid in Penicillium puberulure and the rediscovery of mycophenolic acid in Penicillium brevicompactum (Alsberg and Black, 1913). Another chapter of the Serratia story picks up in 1848, when bloodlike spots were found on a boiled potato in a Berlin home. Ehrenberg investigated the phenomenon and became fascinated. A distinguished physician and protozoologist (he described more than 300 n e w species), Ehrenberg regarded the organism as a "tiny, oval animalcule," and renamed it Monas prodigiosa in 1849. Although aware of Sette's prior nomenclature, his historical efforts apparently did not lead him to Bizio. He came to believe that most historical accounts of bloody food could be attributed to the growth of M. prodigiosa. His colleague

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Scheurlen supported this belief, speaking of the "deceptive red color" of the microorganism and the fact that the hosts were a "particularly agreeable . . . m e d i u m " (Scheurlen, 1896: Gaughran, 1969). Cullen (1994) has analyzed retrospectively weather conditions that correlate humidity and warm air temperatures with historical reports of bloody foods. Another historical aspect may be noted. The laboratory culture of microorganisms on solid or semisolid media (e.g., cooked potato slices, gelatine, agar) developed towards the end of the nineteenth century; Koch's pioneering work with gelatine was reported in 1881. Although Sette and Bizio did not use pure cultures, their work with polenta is probably the first documented use of a solid m e d i u m for culturing microorganisms (Bulloch, 1938). A splendid color plate of S. marcescens growing on a potato is found in an early text (Crookshank, 1890). The organism is named as Bacterium prodigiosum with three other binomials but not including S. marcescens; it is also described picturesquely as "Blood Rain." B. PIGMENTS AND PAINTINGS At the time of Sette's work, a chemist, Pietro de Col, extracted the red pigment and used it to dye silk. He also created yet one more name, Mucot sanguineus, for the organism (Harrison, 1924). Similarly, both Sette and Bizio made ethanol extracts of the pigment and used them to dye silk and wool, sometimes with the aid of mordants. Alert to commercial possibilities, they were thwarted by the unfortunate sensitivity of the dye to light. It has to be remembered that in 1819 the major available red pigments were naturally occurring secondary metabolites derived with difficulty from insects (cochineal, kermes, lac) or from plants (madder); not until 1856 did Perkin produce the first synthetic dye, mauve. More than a century after Bizio and Sette's work, Alexander Fleming found a curious application for the red-pigmented bacterium. He made microbial "paintings" by outlining a drawing on blotting paper, placing it on a nutrient agar plate, and then inoculating with bacterial culture broths. On incubation a colored "germ painting" developed. Six of these "paintings" were reproduced on the endpapers of his biography by Andr6 Maurois. Clearly unfamiliar with the vagaries of bacterial nomenclature, Maurois used the superseded name in describing the possible colors--"the staphylococcus is yellow, the bacillus prodigiosus (sic) red, the bacillus violaceus (sic) blue" (Maurois, 1959). Even more recently, Serratia has also been used as "red ink"; at the 1956 Presidential Banquet of the American Society for Microbiologists in Houston, substitute "place cards" were fashioned from Petri plates containing

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appropriate media on which the names of the officers had been "written" with cultures of red-pigmented S. marcescens (Anonymous, 1999). C. THE GENUS SERRATIA The organism with this long and fascinating history is a member of the Enterobacteriaceae (aero-anaerobic, Gram-negative bacteria) and is motile (Blazevic, 1980; Grimont and Grimont, 1984, 1991). Some species and biotypes of Serratia produce reddish pigment(s) and, depending on colony age, the color ranges from dark red to pale pink. Pigment production is dependent on specific growth conditions, including m e d i u m composition, presence of certain ions and detergents, and temperature. It requires air, and the pigmentation is better developed w h e n Serratia cultures are incubated below 35°C or w h e n a low-phosphate agar without glucose (e.g., peptone-glycerol) is used. There is a strong tendency for clinical isolates to be nonpigmented and difficult to distinguish from other coliform organisms (Hejazi and Falkiner, 1997). Nonpigmented S. marcescens biotypes seem restricted to hospitalized patients, whereas pigmented biotypes are ubiquitous. In a 1978 review of the genus Serratia, three other species (in addition to S. marcescens) were recognized (Serratia liquefaciens, S. plymuthica, S. marinorubra), and a fourth, tentatively discussed as "strain 38" (Grimont and Grimont, 1978), was later named S. odorifera (Grimont et al., 1978). In the 1984 edition of Bergey's Manual of Determinative Bacteriology, S. marinorubra became S. rubidaea, and a sixth species, S. ficaria, was recognized (Holt and Krieg, 1984). In the second edition of The Prokaryotes, 10 species were mentioned and are presently known to belong in the genus Serratia (Grimont and Grimont, 1991). In addition to those already listed, the four other species are as follows: S. entomophila, S. fonticola, S. grimesii, and S. proteomaculans. Of these 10 species, only three--S, marcescens, S. plymuthica, S. rubidaea--produce prodigiosin (Grimont and Grimont, 1991). Bizio's patriotism in naming the first member of this genus as S. marcescens is admirable, and there is a pleasant euphony in the name; however, the steamboating physicist had nothing to do with the contaminated polenta. Bizio might have made a more relevant choice, for instance, the use of the peasant's name in whose house the "fungus" was found: Pittarella marcescens also has a fine ring to it! In fact, naming the red-pigmented bacterium became something of a cottage industry among early bacteriologists, with more than 20 names applied to this organism during the 100 years after Bizio's description. There was a tendency to retain the prodigious characteristic invented by

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Ehrenberg, probably in view of the supposed association with mira c l e s - f o r example, Bacillus prodigiosus and Bacterium prodigiosum. In 1920, the "final" report of The Committee on Classsification of the Society of American Bacteriologists recognized a possible priority for Erythrobacillus pyosepticus, which had been preserved as ATCC 275, and suggested the name Erythrobacillus prodigiosus (Grimont and Grimont, 1991). However, at that time, bacteriological nomenclature was governed by the International Botanical Code, which contained a priority principle requiring the oldest validly published name to be used. Erythrobacillus prodigiosus contradicted rules of priority and never gained acceptance outside the United States (Breed and Breed, 1924). It took the American bacteriologist Buchanan to apply the principle of priority and revive Serratia as the valid name. The first edition of Bergey's Manual of Determinative Bacteriology legitimized Bizio's priority more than a century after he had wished to honor Serrati (Bergey et al., 1923). It has been retained in subsequent editions of the Manual. Ironically, there is no proof that what is n o w called Serratia corresponds to Bizio's organism. The genus Serratia has the distinction in bacteriology of being outranked in age only by the genera Vibrio (1773) and Polyangium (1809). Even so, the acceptance of Serratia as a valid name has attracted considerable dissent. Specimens viewed by early microbiologists tended to be mixed cultures. The small size and morphological monotony of most bacteria provided few clues to the diversity of species. There was an unfortunate tendency to call all red-pigmented microorganisms Serratia simply because of their color. Red bacteria appearing on salted fish are a case in point. Such halophilic species do not ferment carbohydrates and are probably species of Halobacterium (Ayres et al., 1980). Several species of the yeast genus Rhodotorula form shiny, pink to red colonies on bread and other starchy foods and may also have been responsible for some of the incidents of bloody bread. In his delightful essay "Heretical Taxonomy for Bacteriologists," Cowan (1970) devoted a section to "The heresy of Serratia marcescens" referring to the uncertainty of applying names from the "pre-bacteriological era." Cowan felt that it was better to change the rules of nomenclature than to use pre-Pasteurian descriptions that were "in the modern sense, nothing short of the farcical" (Cowan, 1956). Later, in his posthumously published A Dictionary of Microbial Taxonomy, Cowan (1978) stated his opinion authoritatively: In my view it is a waste of time to try to find useful bacteriological information from observations made before bacteria were clearly distin-

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guished from algae, filamentous fungi and yeasts, and I believe we shall not lose anything by ignoring all work before the pioneer work of Pasteur. Nevertheless, many scientists, especially microbiologists, will have confidence that S. marcescens was responsible for most of the incidents involving blood on foodstuffs. There are other microorganisms forming pink or red colonies such as the yeasts Rhodotorula, Sporobolomyces salmonicolor, and Candida pulcherrima; the latter at least is ruled out since its pigment, pulcherrimin, is an iron complex and is insoluble in the usual organic solvents. Moreover, the characteristic dripping or flowing of S. marcescens cultures is not associated with the yeasts or fungi having red pigmentation. On the other hand, since other bacteria produce prodigiosin or prodigiosin-tike materials, some of the observed cases of blood-spotted food may have been due to organisms other than S. marcescens. The phenomenon was readily reproduced on "host bread" using S. marcescens cultures by Ehrenberg in the nineteenth century and more recently on polenta, unconsecrated Communion wafers (both Catholic and Protestant), and not-for-Passover matzos (Karp, 1988; Cullen, 1994); Protestant wafers gave the best results (Bennett, 1994). One of us has pointed out the lack of an appropriate control in these experiments; simple crackers or preservative-flee bread without religious significance should have been included! (Bentley, 1997). It may be noted that there have been reports of red spots on the cream layer of milk (Grimont and Grimont, 1978). Moreover, the range of materials subject to the development of red-spotted areas has been extended by the discovery of a "red spot disease" on culture beds of the kelp Laminaria japonica cultivated in the ocean around Hokkaido and used in the production of "makonbu" (Sawabe et al., 1998). The dried kelp, more colloquially known as "konbu," is usually used to flavor broth and soups, being then discarded. An aerobic, polarly flagellated, marine bacterium was identified as the causative agent of the red spot disease and the name Pseudoalteromonas bacteriolytica sp. nov. was proposed for it. It produces a prodigiosin-like pigment.

IV. Prodigiosin and Related Compounds A. STRUCTURES The major pigment of S. marcescens, originally named prodigiosine, was isolated in 1902, but a choice between alternate structural possibilities was not possible until its chemical synthesis was achieved in 1960, nearly a century and a half after the entrepreneurial hopes of Bizio and Sette had been dashed. Now termed prodigiosin, it is a very

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typical secondary metabolite. Prodigiosin, C20H25N3O, has an unusual structure with three pyrrole rings and is a pyrryldipyrrylmethene (Fig. 1A); two of the rings are directly linked to each other, and the third is attached by way of a methene bridge (Gerber, 1975; Williams and Qadri, 1980). It forms lustrous, square pyramidal crystals that are dark red with a green reflex; the hydrochloride C20H26C1N3 O forms crystals with a magenta color. The highly conjugated system of seven double bonds presumably accounts for the intense pigmentation. Secondary metabolites related to prodigiosin have been isolated from several bacterial genera. These related materials are frequently difficult to purify. Moreover, there has been considerable confusion with respect to naming them; to some extent, "prodigiosin" is used in the literature in a generic sense to include a family of similar materials. In devising trivial names for a group of related compounds it is useful to define a basic nucleus. Two such possibilities have been used for the prodigiosin-like materials (Gerber, 1975). The completely stripped down nucleus, devoid of all substituents, is termed "prodigiosene," while the portion common to most of the natural products, and containing a 6-methoxy substituent, is termed "prodiginine" (for structure and numbering, see Fig. 1A). The apparently bizarre numbering in which three carbons of the bipyrrole and one carbon of the monopyrrole are not numbered was devised "because substitutions on them would destroy the basic linear tripyrrole structure of prodigiosene" (Williams, 1973). Hence, prodigiosin could also be referred to as either 2-methyl-3-pentylprodiginine or 2-methyl-3-pentyl-6-methoxyprodigiosene (in some early papers, amyl is used for the preferred pentyl). Unfortunately, there has been no general agreement concerning the use of either prodigiosene or prodiginine, and, as will be seen, confusion has inevitably arisen with attempts to base nomenclature on prodigiosin itself. While there are advantages to the use of prodigiosene in connection with chemical syntheses, we believe that the use of prodiginine, as suggested by Gerber (1975) is the best solution. The few natural materials containing OH instead of O C H 3 at position 6 can conveniently be termed norprodiginines (Fig. 1A). Four structural types based on the prodiginine nucleus can be recognized: 1. Only straight chain alkyl substituents present: 1A. Alkyl substituents at both positions 2 and 3 (Fig. 1A). The prototype is prodigiosin itself with a methyl group at position 2 and a pentyl group at position 3. Higher homologues with methyl at position 2 and either hexyl or heptyl at position 3

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OCH 3

H

H A

/~4

H

IN R

B

H

R2 FIG. 1A,B FIG. 1. Prodigiosin and related compounds.

Structure 1A Prodigiosene Prodiginine Norprodigiosin Prodigiosin Undecylnorprodiginine Undecylprodigininea

R1 H CH30 HO CH30 HO CH30

R2 H H CH3 CH 3 CH3(CH2)lO CH3(CH2)lo

R3 H H CH3(CH2) 4 CH3(CH2) 4 H H

Structure 1B Metacycloprodigiosin

(ethyl-meta-cyclononylprodiginine) Butyl-meta-cycloheptylprodiginine has a similar meta structure

b u t w i t h only seven - - C H 2 - in the ring a n d a b u t y l substituent. Structure 1C Cycloprodigiosin hydrochloride Structure 1D C y c l o n o n y l p r o d i g i n i n e ; R -- H, n = 8 C y c l o m e t h y l d e c y l p r o d i g i n i n e ; R = CH3--, n :- 9 Tautomeric arrangements of the double b o n d systems are possible. One e x a m p l e is s h o w n in structure lB. aThis material is also referred to as u n d e c y l p r o d i g i o s i n a n d prodigiosin-25C (see text).

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OCH 3

OCH 3

C CH3

O~'.- (CH2)n -~ FIG. 1C,D

lB.

(along with prodigiosin) have been isolated from Pseudomonas magnesiorubra, the marine psychrophilic bacterium Vibrio psychroerythreus, and a sewage bacterium (Gerber, 1975). A river bacterium, Rugamonas rubra gen. nov., sp. nov., produces prodigiosin and (probably) the heptyl homologue (Austin and Moss, 1986). Norprodigiosin (2-methyl-3-pentylnorprodiginine) is formed by the S. marcescens mutant OF. Frequently, prodigiosin occurs as an adduct with macromolecules, typically protein. Alkyl substituents at position 2 only. Prodigiosin-like materials with an undecyl chain at position 2 were first fully characterized from certain actinomycetes (e.g., Streptomyces longisporus tuber) in 1966 (Wasserman et al., 1966; Harashima et al., 1967) and with a nonyl sidechain from Actinomadura madurae (formerly Nocardia madurae) (Gerber, 1975). The first is well known and has often been termed "undecylprodigiosin"--which would imply prodigiosin itself with an extra undecyl group. Another name, "prodigiosin-25 C" (or prodigiosin 25-C), is based on the total number of carbon atoms present with a designation of the chronological order of discovery (Harashima eta]., 1967). Thus, this name is meant to indicate that it is the third C25 material discove r e d - b u t is again not helpful in structural terms. Both of

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J.w. BENNETT and RONALDBENTLEY these terms are used in the current literature, as is the abbreviation Red, by molecular geneticists. Confusion can be avoided by naming this material as undecylprodiginine, as suggested by Gerber (1975), and this name will be used here. As with prodigiosin, a material with - - O H at position 6 is known, and is best named as undecylnorprodiginine.

2.

3.

Ring formation between positions 2 and 4 (i.e., a meta arrangement). A structure with a cyclononyl ring linked to positions 2 and 4 and carrying an additional ethyl substituent (Fig. 1B) was isolated from Streptomyces longisporus tuber in 1969. Generally known as metacycloprodigiosin, a more appropriate name w o u l d be ethylmeta-cyclononylprodiginine. In biosynthetic terms, it appears to result from a cyclization of undecylprodiginine. Metacycloprodigiosin is probably identical with streptorubin A from Streptomyces rubrireticuli var. pimprina (Gerber, 1975). Some organisms, for example, Streptomyces hiroshimensis, produce both metacycloprodigiosin and undecylprodiginine; this is also true for an actinomycete isolated from leek roots and belonging to the Streptoverticillium baldaccii cluster (Brambilla et al., 1995). A material identified as butyl-meta-cycloheptylprodiginine has also been isolated from an actinomycete, strain B 4358 (Laatsch et al., 1991); it resembles ethyl-meta-cyclononylprodiginine, but the ring between positions 2 and 4 contains only seven - - C H 2 - - groups and carries a butyl substituent instead of an ethyl. Butyl-meta-cycloheptylprodiginine is probably identical to the material previously identified as butylcycloheptylprodiginine (butyl-ortho-cycloheptylprodiginine, with ring formation at positions 2 and 3) (Gerber, 1975), and is also identical to the antibiotic streptorubin B isolated from Streptomyces roseoverticulatus var. albosporus (Gerber, 1975; F~rstner et al., 1998). It is likely to be another cyclized form of undecylprodiginine. Ring formation between positions 3 and 4 (i.e., an ortho arrangement) and with CH3 at position 2. The only example of this structural type is a cyclized form of prodigiosin itself, usually known as cycloprodigiosin; a more informative alternative to cycloprodigiosin w o u l d be to use the name 2-methyl (methyl-ortho-cyclobutyl)prodiginine. Originally isolated from a marine bacterium, Alteromonas ruber (Gerber and Gauthier, 1979), it was assigned an incorrect structure. A revision in 1983 (Gerber, 1983; Laatsch and Thomson, 1983) indicated that the ring contains three - - C H 2 - - and one ---CH3-CH-- group. Cycloprodigiosin was also found, together

SEEING RED: THE STORY OF PRODIGIOSIN

15

with prodigiosin itself, in the anaerobic marine bacterium Vibrio gazogenes (formerly Beneckea gazogenes) (Harwood, 1978; Gerber, 1983; Laatsch and Thomson, 1983) and as its hydrochloride (Fig. 1C) in Pseudoalteromonas denitrificans, a novel marine bacterium, isolated "from the sea near Japan" (Kawauchi et al., 1997). 4. Ring formation between position 2 of the monopyrryl unit and position 10 of the dipyrryl unit. Compounds described generically as "macrocyclic prodiginines" have been isolated from Actinomadura pelletieri (formerly Nocardia pelletieri) and Actinomadura madurae (formerly N. madurae) (Gerber, 1975). These structures contain a bridge with several --CH2-- groups between the first and third pyrrole rings (Fig. 1D). Again, the nomenclature is a problem since the structure of Figure 1D with R = H and n = 8 is named as "cyclononylprodiginine"; there could easily be confusion with the previously described ethyl-meta-cyclononylprodiginine. These "macrocyclic prodiginines" are apparently unique to the two organisms named. Finally, there has been an unfortunate nomenclature confusion between the red pigment prodigiosin and a material referred to in Russian literature as "prodigiosan." The latter is a polysaccharide or lipopolysaccharide also isolated from S. marcescens. In searching Biological Abstracts Online for "prodigiosin," some abstracts containing "prodigiosan" were obtained. Moreover, in some cases, the Russian word for prodigiosan was translated as prodigiosin--to take only one example, "activation of mononuclear phagocytes by a lipopolysaccharide (prodigiosin)" (Panin et aI., 1996). B. BIOSYNTHESIS

How is the strange pyrryldipyrrylmethene structure present in prodigiosin and related compounds constructed by bacteria? Isotope tracer studies with both stable and radioactive isotopes were undertaken almost half a century ago at a time when only prodigiosin itself was known and w h e n it was believed (incorrectly) to be a tripyrrylmethene; one of us still remembers carrying out 15N assays with a mass spectrometer for this kind of work (Hubbard and Rimington, 1950). This and other studies indicated important roles for acetic acid and glycine in prodigiosin biosynthesis, and later work additionally implicated proline, serine, and alanine. Owing to difficulties in carrying out chemical degradations of labeled samples to determine location of isotopes, progress was delayed until NMR methods became available; for a summary, see Gerber et al. (1978).

16

J.w. BENNETT and RONALD BENTLEY

It was eventually learned that the A and B rings (i.e., the dipyrryl unit; see Fig. 1A) of prodigiosin, undecylprodiginine, and metacycloprodigiosin were constructed similarly, but the C ring of prodigiosin was formed differently from that of undecylprodiginine and metacycloprodigiosin. In all cases, the methyl of the --OCH3 group at position 6 derived from S-adenosylmethionine. These results were complemented by extensive studies with mutants not producing prodigiosin (Mody et al., 1990). Thus, one such mutant, mutant 933, produced methoxybipyrrolecarboxaldehyde, MBC (Fig. 2A), and mutants Wl and WF produced methylpentylpyrrole (Fig. 2B), abbreviated MAP (from the early designation as methylamylpyrrole). If mutant 933 was supplied with MAP, prodigiosin was formed; similarly, Wl and WF produced prodigiosin if supplied with MBC. This process is referred to as "syntropic pigmentation." Initially, three mutant classes were described from nonpigmented clinical strains of S. marcescens (Ding and Williams, 1983); subsequently, five more have been recognized (Mody et al., 1990). Moreover, "nonnatural" monopyrroles could be added to 933, thus leading to novel prodiginine pigments (e.g., the use of 2,4-dimethylpyrrole gave a pigment with methyls at both positions 2 and 4). When pigmented strains are grown at 37-40°C, pigment is no longer formed. These elevated temperatures apparently interfere only with production of MBC since syntropic pigmentation occurred in most cases when it was added (Katz and Sobieski, 1980). The bipyrryl unit is invariable for the entire range of prodiginine pigments. The A ring is formed from the four noncarboxyl carbons and the nitrogen atom of proline. The B ring contains the proline carboxyl carbon, one acetate unit, and two carbons and one nitrogen from serine (the serine carboxyl is lost). For prodigiosin itself the monopyrryl unit (ring C) is constructed from two carbons and the nitrogen atom of alanine (the carboxyl of this amino acid also being lost) and a tetraketide unit from four acetate units (Fig. 2B). The monopyrryl unit for formation of undecylprodiginine derives from two carbons and the nitrogen of glycine (again a decarboxylation takes place) and a heptaketide unit from seven acetates (Fig. 2C}. A condensation of glycine with other polyketides can account for formation of the other actinomycete prodiginines. An important observation, made in 1985, was that undecylprodiginine, along with other prodigiosin-like materials, was produced by Streptomyces coelicolor (Tsao et al., 1985). Prodigiosin itself had been recognized earlier in an actinomycete (Perry, 1961). However, M. Bibb (personal communication) has suggested that this identification (based largely on a u.v. spectrum) was incorrect and that Perry's material was

SEEING RED: THE STORY OF PRODIGIOSIN

17

OCH3

A

CHO

+ PRO

H

~

//~--CH20H H2N COOH SER COOH •

H2 N

NH2

HOOC'~ / CH3 ALA

R'

HN

GLY

•

CH3

R"

B Ri:

_

ri

LC. -

C •

G

•

RII

--

LOH2-OH2JuOH3

r I

|

"1

•

FIc. 2. Biosynthesis of the two components required to form prodigiosin and undecylprodiginine. Standard three-letter abbreviations are used for amino acids. In all cases, filled-in squares denote carbon derived from the methyl of acetate and filled in circles denote carbon derived from the carboxyt of acetate. In actuality, the polyketide units are most likely formed from acetate plus polymalonate condensations. The arrows at the top of the monopyrrole units indicate the molecular position involved in reaction with the aldehyde group of MBC (structure A). A. The methoxybipyrrolecarboxaldehyde, MBC, is used for prodigiosin itself as well as undecylprodiginine and related compounds. It is constructed from all five carbons and the nitrogen of proline (pro), a single acetate unit, and two carbons and the nitrogen of serine (ser; the ser COOH is lost as CO2). B. The methylpentylpyrrole, MAP, required for prodigiosin formation is constructed from two carbons and the nitrogen of alanine (ala; the ala COOH is lost as CO2) and a tetraketide unit (i.e., eight carbons) formed from acetate. C. The undecylpyrrole required for undecylprodiginine is constructed from one carbon and the nitrogen of glycine (gly; the gly COOH is lost as CO2) and a heptaketide (i.e., 14 carbons) formed from acetate.

18

J.w. BENNETTand RONALDBENTLEY

actually undecylprodiginine. Streptomyces coelicolor also produces the aromatic polyketide actinorhodin and has been much used in molecular genetics studies (Cane, 1997). Undecylprodiginine played an important role in the first cloning of a gene, playing a defined role in the biosynthesis of an antibiotic; an O-methyltransferase gene was isolated by complementation and the color of undecylprodiginine was used as the selectable phenotype (Feitelson and Hopwood, 1983). The enzymatic product converted undecylnorprodiginine to undecylprodiginine; furthermore, two forms of the necessary enzyme, undecylnorprodiginine--S-adenosylmethionine O-methyltransferase--were detected, one with a very high molecular mass peak (Feitelson et al., 1985). The genes involved in actinorhodin and prodigiosin biosynthesis in Streptomyces coelicolor A3(2) have received very extensive investigation (see reviews by Hopwood et al., 1995; Bibb, 1996) and continue to be of interest (Chakraburtty and Bibb, 1997; White and Bibb, 1997; Guthrie et al., 1998). The regulation of prodigiosin biosynthesis is complex, being influenced by increased glucose levels and decreased by increased phosphate levels (Gyun-Kang et al., 1998). The S. marcescens genes encoding prodigiosin biosynthesis from the necessary mono- and bipyrryl units have been cloned and expressed in E. coli (Dauenhauer et al., 1984). No E. coli recombinants encoded the entire prodigiosin biosynthetic pathway. However, strain SAD400 produced prodigiosin when supplied with MBC by S. marcescens 933, while strain SAD757 required both MBC (from mutant 933) and MAP (from mutant WF). Clearly, SAD400 could form MAP, and both of the E. coli recombinants could condense the two portions of the molecule together. One strain of S. marcescens, ATCC 39006, has the unusual property of producing both prodigiosin and the [3-1actam antibiotic carbapenem (Thomson et al., 1997). Mutants defective in the production of these secondary metabolites had a mutation in a gene termed rap (for regulation of antibiotic and l~igment). It appears that this and related genes (e.g., in Erwinia and Yersinia) form a subfamily of proteins regulating diverse aspects of bacterial physiology. V. From Saprophyte to Pathogen For at least a century, S. marcescens was regarded as a harmless saprophyte. In fact, this pigmented bacterium was used extensively as a marker organism; generations of bacteriology students remember demonstrations of how a simple handshake can transmit microorganisms

SEEING RED: THE STORYOF PRODIGIOSIN

19

from one individual to another. More recently, S. m a r c e s c e n s was used as a test organism with pigskin as a substrate to evaluate topical antimicrobial action. Simulated handwashing protocols were evaluated in parallel with the i n - v i t r o model (McDonnell et al., 1999). In one dramatic public experiment to demonstrate the spread of microorganisms, the intrepid Dr. M. H. Gordon in 1906 gargled with an S. m a r c e s c e n s culture before reciting Shakespeare to the House of Commons. No MPs were present, but Petri dishes with an appropriate culture medium were placed at various distances (Yu, 1979). Recovery of red bacterial colonies demonstrated the role of speaking and coughing in spreading bacteria. Present-day microbiologists w o u l d not consider a repetition of the experiment since S. m a r c e s c e n s is now recognized as an opportunistic pathogen (see later). Another interesting prodigiosin story is the fascinating "red diaper syndrome" in Wisconsin of 1958 (Waisman and Stone, 1958). The child of a genetics professor, born uneventfully in the University of Wisconsin hospital, was brought home and a diaper delivery service was hired. The first pickup showed apparent blood stains on the diapers. Upon further examination, the child's urine and stool were normal; the bloody color developed only after the diapers were stored in a bin. The father, familiar with an abnormal tryptophan metabolism causing a "blue diaper syndrome," suspected a genetic abnormality. Eventually, of course, the guilty culprit turned out to be a strain of pigmented S. m a r c e s c e n s being used in the medical school to study aerosol techniques and genetics. The infant's intestines were heavily infected with the organism; use of sulfasuxidine and a controlled diet eventually restored a "normal" intestinal flora, but almost a year was required. Prodigiosin-producing S. m a r c e s c e n s has also been used as a marker organism in germ warfare research (Yu, 1979). In n o w notorious experiments conducted between 1950 and 1966, S. m a r c e s c e n s cultures were released by the U.S. Army on an unsuspecting population of involuntary and unwitting subjects in the New York City subways, and in locations in Calhoun County (Alabama), Key West (Florida), and San Francisco. In the Pacific Coast experiments, ships released cultures into the ocean, whereupon an aerosol was formed by wave action. Red-pigmented bacteria were recovered in air samples some 80 meters inland. When these secret experiments were finally acknowledged, the Army maintained that there had been no infections attributable to them. However, a documented outbreak of S. m a r c e s c e n s infections did occur in a San Francisco hospital in 1950-51 at the time of the aerosol experiments. One patient died from the first known case of serratial endocarditis, and his family sued the Department of Defense. While it

20

J.w. BENNETT and RONALD BENTLEY

was tempting to link the two events, later work by the Centers for Disease Control in Atlanta indicated that in 100 cases of S. m a r c e s c e n s infections in the United States, none had been caused by an organism with the same serotype and biotype as that used by the Army. The legal judgment was that the San Francisco case was probably the first in a series of new nosocomial infections rather than a consequence of the Army's program. Prior to about 1970, cases of serratial bacteremia were very rare, but in a single hospital (Stanford University Hospital) from 1968 to 1977, some 76 cases were reported (Yu et aL, 1979). S. m a r c e s c e n s is now implicated in many serious conditions; the list includes empyema, lung abscess, meningitis, osteomyelitis, peritonitis, pneumonia, sinusitis, urinary tract infection, and wound infection (Hejazi and Falkiner, 1997; von Graevenitz, 1980; Daschiner, 1980). A dramatic instance of a nosocomial outbreak occurred in Nashville, Tennessee involving an antibiotic-resistant strain infecting patients at four separate hospitals. The drug-resistant strain was isolated from the urine of a catheterized patient in April of 1973, and by late 1974 the same strain (characterized by serotype, phage type, and antibiotic-resistance pattern) had been isolated in the other hospitals. All four institutions were teaching hospitals and had regular rotations of physicians and nursing staff. A total of 210 patients were infected, 21 of them becoming bacteremic, with 8 fatalities (Williams and Qadri, 1980). Later, the organism was isolated from pooled hand rinsings of personnel. Dr. Gordon would not have been surprised. In another case, reported in 1996, an outbreak of S. m a r c e s c e n s occurred in a neurosurgery intensive care unit (Bosi et al., 1996). The responsible strain was located in a diluted hexetidine solution used as a mouthwash; the bottle of diluted antiseptic was the single source of this nosocomial outbreak. Other disinfectants (hexachlorophene, benzalkonium chloride) can also become contaminated (Yu, 1979). Medical equipment has also been implicated in the spread of S. m a r c e s c e n s in hospitals; a bizarre case involved shaving brush bristles used for personal grooming in an intensive care unit. Yet another problem arises with the use of solutions of chlorhexidine for disinfecting contact lenses. In one study, 11 of 12 strains of S. m a r c e s c e n s became adapted to the agent (Gandhi et al., 1993). Contaminated lens solutions have been associated with ocular infections (Mayo et al., 1987). The antibiotic resistance of many strains of S. m a r c e s c e n s is a serious problem (Yu, 1979; Yu et al., 1979; Farrar, 1980), with rapid horizontal transfer of drug resistance by plasmids. Although transfer to E. coli K12

SEEING RED: THE STORY OF PRODIGIOSIN

21

was inefficient, it was more effective to Klebsiella p n e u m o n i a e (Hedges, 1980). Clearly, new methods for controlling S. m a r c e s c e n s infections would be welcome. It has been suggested that a possible chemotherapeutic target might be the bacterial regulatory proteins formed by the rap gene, which were described earlier (Thomson et al., 1997). Cases of serratial endocarditis have been reported among heroin addicts in San Francisco; they were associated with a high frequency of embolic complications and a refractoriness to medical therapy. Perhaps the increased frequency with which serratial infections have been observed relates to the heavy use of antibiotics in the 1950s and the consequent development of drug-resistant strains. The absence of cases before the antibiotic era is striking. Some veterinary problems have been associated with Serratia species, and S. m a r c e s c e n s has been frequently recovered from healthy, diseased, and dead insects; both S. m a r c e s c e n s and S. liquefaciens are classified as potential insect pathogens (Grimont and Grimont, 1978). A possible role for S. m a r c e s c e n s in septic abortions in cows and buffaloes has been described (Das et al., 1988); most of the isolated strains produced prodigiosin.

VI. Biological Activity of Prodigiosin and Related Compounds A. POSSIBLE ECOLOGICAL FUNCTIONS

As typical secondary metabolites, prodigiosin and related materials have no clearly defined physiological functions in the producing organisms. However, it is possible that pigmented S. m a r c e s c e n s may have an advantage in ecological dispersion (Burger and Bennett, 1985). In studies of the drops produced by bursting air bubbles rising through bacterial suspensions, pigmented strains were enriched in the drops (relative to that of the bulk suspension) (Burger and Bennett, 1985; Syzdek, 1985). The pigmented cells appeared to have increased hydrophobicity, possibly due to the presence of prodigiosin. It was acknowledged, however, that the cell enrichment was a complex chain of events and was influenced by cultural conditions (Syzdek. 1985). Other workers indicated that clinical S. m a r c e s c e n s strains had hydrophobic properties in the absence of prodigiosin and that hydrophobicity was only shown by growth at 30°C but not at 37°C (Rosenberg et al., 1986). Pigment is not synthesized at the higher temperature. It seems clear that the cell-surface hydrophobicity of S. m a r c e s c e n s is not totally due to

22

J.W. BENNETT and RONALD BENTLEY

surface pigment. Nonpigmented cells contained an additional protein (Mr = 40,000) that may be responsible for the higher surface hydrophobicity of some nonpigmented mutants (Mallick, 1996). Color variation in Serratia has been correlated with amount of flagellar antigens, and there is an apparent coregulation of pigment and flagellin synthesis. Variation of these surface antigens may allow pathogenic stains to evade host immune systems (Paruchuri and Harshey, 1987). In a very detailed study by Van der Mei et al. (1992), S. marcescens strains were characterized by contact angle and zeta potential measurements, X-ray photoelectron spectroscopy, and infrared spectroscopy. Again, it appeared that the presence of prodigiosin did not influence the cell-surface hydrophobicity. It was suggested that the pigment was confined in deeper layers than those probed by contact angles (about 0.3-0.5 nm). Other results indicated that both pigmented and nonpigmented strains produced extracellular vesicles and had wetting activity when grown at 30°C, but not at 37°C. The wetting activity was probably important for spreading cells on the surfaces of porous or fibrous materials, especially those with hydrophobic properties (Matsuyama et al., 1986). Finally, the presence of the O-antigen may be important in adhesion of S. marcescens to plastic and glass and to human uroepithelial cells (Palomar et al., 1995).

B. PHARMACOLOGICAL ACTIVITY

In a review of pre-penicillin antibiosis, Abraham and Florey (1949) cited a report that "complete inhibition" of cholera Vibrio was achieved by an old culture of Micrococcus prodigiosus, an early name for S. marcescens, and gave instances of the antagonistic properties of "Chromobacterium prodigiosum" (i.e., S. marcescens) against other bacteria, trypanosomes (the "nagana trypanosome [Prowazek]"), protozoa, and fungi. Prodigiosin itself has been identified as having extremely broad antibiotic properties, being active against Gram-positive bacteria, protozoa, and pathogenic fungi. It has been used experimentally as a fungistatic/fungicidal agent against Coccidioides immitis, but, unfortunately, water-soluble solutions of the glutamic acid form caused venous sclerosis on injection (Williams and Hearn, 1967). Strains of Serratia plymuthica producing prodigiosin and other materials with antifungal properties were beneficial rhizobacteria for oilseed rape (Kalbe et aL, 1996).

SEEING RED: THE STORY OF PRODIGIOSIN

23

To some extent, prodigiosin prolongs the lives of mice infected with the malaria parasite Plasmodium berghei on subcutaneous administration in peanut oil (Castro, 1967). The macrocyclic structures cyclononylprodiginine (Fig. 1D, R -- H, n = 8) and cyclomethyldecylprodiginine (Fig. 1D, R = CH 3, n = 9) had activities comparable to that of prodigiosin itself, but undecylprodiginine was inactive (Gerber, 1975). At a time w h e n many malarial parasites are resistant to conventional treatment, there is an increased incentive to find new drugs. The antimalarial activity of prodigiosin makes it an attractive target for modern genetic and chemical manipulations. Prodigiosin showed significant cytotoxic activity against some cell cultures; it was particularly potent against P388 mouse leukemia (9 PS) with an IC50 (inhibitory concentration for 50% cell growth relative to untreated controls) of 3.7 x 10 -4 btg m1-1 (Boger and Patel, 1988). With L-1210 mouse lymphocytic leukemia, B16 mouse melanoma, and 9KB human epidermoid nasopharynx carcinoma, the ICso values were in the range 2.0-4.0 × 10 -2 btg m1-1. Within the last few years, a number of other interesting physiological activities have been associated with prodigiosin. In T-cell lymphoma YAC-1, undecylprodiginine strongly suppressed incorporation of [3H]acetate into the lipid fraction (Kataoka et al., 1995a). Using preparations from rat liver, undecylprodiginine had little or no inhibitory effect on fatty acid synthase, acetyl-CoA synthetase, and acetyl-CoA carboxylase. It appeared that undecylprodiginine perturbed permeation of acetate through the plasma membrane of YAC-1. Beginning in 1995, it was reported that undecylprodiginine uncoupled the vacuolar type H+-ATPase and inhibited vacuolar acidification in baby hamster kidney cells (Kataoka et al., 1995b; Ohkuma et al., 1998). The proton pump activity, but not the ATP hydrolytic activity, was inhibited in rat liver lysosomes, and glycoprotein processing was also suppressed. In addition, prodigiosin, metacycloprodigiosin, and undecylprodiginine uncouple the acidification mediated by F-type H +ATPases of both submitochondrial (rat liver) and E. coli inverted membrane vesicles (Konno eta]., 1998). Prodigiosins have an ionophoric nature and act as H+/C1- symporters (or OH-/C1- antiporters) in liposomes. Their uncoupling effect is likely due to their H+/C1- symport activity across biological membranes (Ohkuma et a]., 1998; Konno et al., 1998; Sato et al., 1998). The proton pumping activity of V-ATPase in osteoclastic cells is essential for bone resorption; undecylprodiginine and metacycloprodigiosin inhibition of acidification of vacuolar organ-

24

J.w. BENNETT and RONALD BENTLEY

elles suppresses parathyroid hormone-stimulated bone resorption (Woo et al., 1997). Undecylprodiginine (and other V-ATPase inhibitors) blocked the perforin-dependent cytotoxicity mediated by the CD8 ÷ cytotoxic T-cell clone; an acidic pH is needed to maintain the quantity and also the quality of perforin in the lytic granules (Togashi et al., 1997). The inhibitory effect of undecylprodiginine on acidification of intracellular organelles may account for some of its immunosuppressive activity (see later). In summary, these prodigiosins constitute a new group of useful probes for an analysis of vacuolar functions since their effects are selective to vacuolar pH in vivo with little or no effect on the cellular ATP level. The prodigiosin structure is small enough so that chemical modifications might lead to more active and less toxic compounds (Sato et al., 1998); a simple and elegant new synthesis of prodigiosins (D'Alessio and Rossi, 1996) and of a thienyl analogue of undecylprodiginine (D'Auria et al., 1999) have potential value in this connection. Other syntheses of prodigiosins have been summarized by F/_irstner et al. (1998). An even more remarkable property of prodigiosins is their immunosuppressive activity. Undecylprodiginine and metacycloprodigiosin, obtained from S t r e p t o m y c e s h i r o s h i m e n s i s , were found to be potent inhibitors of T-lymphocyte proliferation induced by concanavalin A and phytohemagglutinin, but were less suppressive against B-lymphocyte proliferation induced by lipopolysaccharide. There was little toxicity in mice (Nakamura et al., 1986, 1989). Cycloprodigiosin hydrochloride also had immunosuppressive properties (Kawauchi et al., 1997), and a recent paper described T-cell-specific immunosuppression by prodigiosin itself (Han et al., 1998). Metacycloprodigiosin was more effective in reducing splenic cytotoxic T-cell activity than in prolonging murine skin or cardiac allografts (Magae et al., 1996). It has also been stated that undecylprodiginine has a low therapeutic index in rats and is probably not selective for T-cell activation (Metcalfe et al., 1993). Since the action of prodigiosins is different from that of cyclosporin and rapamycin (Songia et al., 1997; Tsuji et al., 1990, 1992), it has been suggested that prodigiosin and related compounds may be useful as supplementary immunosuppressants in combined therapy (Kawauchi et al., 1997; Songia et al., 1997; Tsuji et al., 1992; Lee et al., 1998). If prodigiosin or a prodigiosin analogue emerged as a useful antimalarial drug or as an immunosuppressant in human therapy, it would provide a happy ending (or a bright new beginning) to the singular story of this already wondrous secondary metabolite.

SEEING RED: THE STORY OF PRODIGIOSIN

25

VII. Final Comments S. marcescens has played an important role in the history of bacterial taxonomy, in research on the transmission of bacterial aerosols, in the study of emerging nosocomial infections, and in the understanding of secondary metabolite biosynthesis. The prodigiosin pigments have intrigued organic chemists and pharmacologists, and may yet play roles in the treatment of infectious diseases such as malaria, and perhaps as immunosuppressant agents. However, a major reason for much of the continuing curiosity in the Serratia/prodigiosin story is the theory that these viscous, crimson bacterial colonies provide a naturalistic explanation for certain long-ago miracles involving the Eucharist, and that their appearance gives some credible explanation for the persistence of the antisemitic outrages associated with the "blood libel." The word miracle comes from the Latin miraculum, "a wonder, a marvel," and is related to the verb mirari, "to wonder, to be astonished, or amazed." There have been many extensive definitions, from the facetious to the profound. In ordinary language, the word miracle often conveys the idea of something wonderful, as in "the miracle of birth," or "penicillin is a miracle drug." To the religious community, a miracle is something more profound: an event that seems to contradict natural laws and that can be attributed to God or some other higher cause (Perschel and Perschel, 1988). Most scientists live by a creed that involves a belief that the world behaves according to the predictable laws of nature. Miracles are violations of natural laws; therefore, they do not occur. As George Santayana has written, "Miracles are propitious accidents, the causes of which are too complicated to be readily understood" (Santayana, 1930). Was the Miracle at Bolsena a divine event or merely a growth of S. marcescens? The answer to this question may be irrelevant; the crucial event (perhaps itself the miracle) was that the priest regained his faith (Vaclav, 1994). In like manner, standard accounts of the desecration of the Host, as told by microbiologists, grossly overemphasize the possible role of bacteriology, as if the invocation of a natural explanation can make medieval antisemitism more comprehensible. It is of course conceivable or even likely that prodigiosin or other microbial pigment may have played a role in some of the reports of blood on consecrated bread. If not, the dire consequences of the miracles must be kept in mind by those choosing to regard the incidents in religious terms (Isenberg, 1995).

26

J.W. BENNETT and RONALD BENTLEY

In general, too many scientists look back at these records with a smug, self-designated attitude of epistemological privilege. Ethicists, historians, philosophers, priests, rabbis, and other scholars and writers are better able to comprehend the mystery of human prejudice than are individuals using scientific facts and hypotheses alone. It is well to remember, in a paraphrase of Anatole France, that a person taking pride in being without prejudice has asserted a claim that is itself a very great prejudice--in his words, "He flattered himself on being a man without any prejudices; and this pretension itself is a very great prejudice" (France, 1938).

Acknowledgments We are grateful to Drs. F. and P. A. D. Grimont and Dr. Mervyn Bibb for reading the manuscript and for their very helpful suggestions. We thank Johanna Cullen and Ora Karp for sharing unpublished manuscripts with us, Drynda Johnston and Ann Rogers, Langley Library, University of Pittsburgh, for help in locating many references, and Mr. and Mrs. H. Katsuhisha for information on, and a sample of, "makonbu." Special thanks go to Scott Burger for stimulating our interest in prodigiosin, and to the Reverend Roger Boraas for guidance on the religious literature.

REFERENCES

Abraham, P., and Florey, H. W. (1949). Antibiotics from chromogenic bacteria. In "Antibiotics" (H. W. Florey, E. Chain, N. G. Heatley, M. A. Jennings, A. G. Sanders, E. P. Abraham, and M. E. Florey, eds.), Vol. 1, pp. 537-565. Oxford University Press, London. Alsberg, C. L., and Black, O. F. (1913). "Contribution to the Stndy of Maize Deterioration." U.S.D.A. Bureau of Plant Industry, Bulletin No. 270, Government Printing Office, Washington, DC. Anonymous (1999). A retrospective look at how far we have come. In "99th General Meeting Program Update," p. 1, American Society for Microbiology, Washington, DC. Austin, D. A., and Moss, M. D. (1986). Numerical taxonomy of red-pigmented bacteria isolated from a lowland river, with the description of a new taxon, Rugamonas rubra, new genus, and new species. J. Gen. Microbiol. 132, 1899-1910. Ayres, J. C., Mundt, J. O., and Sandine, W. E. (1980). "Microbiology of Foods." Freeman,

San Francisco. Bennett, J. W. (1994). More on the Miracle of Bolsena. Am. Soc. Microbiol. News 60, 403. Bentley, R. (1997). Secondary metabolites play primary roles in human affairs. Perspec. Biol. Med. 40, 197-221.

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Dauenhauer, S. A., Hull, R. A., and Williams, R. P. (1984). Cloning and expression in Escherichia coli of Serratia marcescens genes encoding prodigiosin biosynthesis. J. Bacteriol. 158, 1128-1132. D'Auria, M., DeLuca, E., Mauriello, G., and Racioppi, R. (1999). A short synthesis of a thienyl analogue of undecylprodigiosin. Syn. Commun. 29, 35-42 Ding, M.-J., and Williams, R. P. (1983). Biosynthesis of prodigiosin by white strains of Serratia marcescens isolated from patients. J. Clin. Microbiol. 17, 476-480. Farrar, W. E. (1980}. Antimicrobial susceptibility of clinical isolates, synergistic effects, and [3-1actamases of Serrotia. In "The Genus Serratia" (A. von Graevenitz and S. J. Rubin, eds.), pp. 121-138. CRC, Boca Raton, FL. Feitelson, J. S., and Hopwood, D. A. (1983). Cloning of a Streptomyces gene for O-methyltransferase involved in antibiotic biosynthesis. Mol. Gen. Genet. !90, 394-398. Feitelson, J. S., Malpartida, F., and Hopwood, D. A. (1985). Genetic and biochemical characterization of the red gene cluster of Streptomyces coelicolor A3(2). J. Gen. Microbiol. 131, 2431-2441. France, A. (1938). "The Crime of Sylvestre Bonnard," p. 128. Trans. L. Hearn. The Bodley Head, London. (Original publication date, 1908.) Fiirstner, H., Szillat, H., Gabor, B., and Mynott, R. (1998). Platinum- and acid-catalyzed enyne metathesis reactions: Mechanistic studies and applications to the syntheses of streptorubin B and metacycloprodigiosin. J. Am. Chem. Soc. 120, 8305-8314. Gandhi, P. A., Sawant, A. D., Wilson, L. A., and Ahearn, D. G. (1993). Adaptation and growth of Serratia marcescens in contact lens disinfectant solutions containing chlorhexidine gluconate. Appl. Environ. Microbiol. 59, 183-188. Gaughran, E. R. L. (1969). From superstition to science: The history of a bacterium. Ann. N. Y Acad. Sci. 31, 3-24. Gerber, N. N. (1975). Prodigiosin-like pigments. Crit. Rev. Microbiol. 3,469-485. Gerber, N. N. (1983). Cycloprodigiosin from Beneckea gazogenes. Tetrahedron Lett. 24, 2797-2798. Gerber, N. N., and Gauthier, M. J. (1979). New prodigiosin-like pigments from Alteromonas rubra. Appl. Environ. Microbiol. 37, 1176-1179. Gerber, N. N., McInnes, G. A., Smith, D. G., Waiters, J. A., Wright, J. L. C., and Vining, L. C. (1978). Biosynthesis of prodiginines: 13C resonance assignments and enrichment patterns in nonyl-, cyclononyl-, methylcyclodecyl- and butylcycloheptylprodigiosin produced by actinomycete cultures supplemented with laC-labeled acetate and 15Nlabeled nitrate. Can. J. Chem. 56, 1155-1163. Grimont, F., and Grimont, P. A. D. (1991). The genus Serratia. In "The Prokaryotes" (A. Balows, H. G. Triiper, M. Dworkin, W. Harder, and K.-H. Schleifer, eds.), Vol. 3, pp. 2822-2848. Springer-Verlag, New York. Grimont, P. A. D., and Grimont, F. (1978). The genus Serratia. Annu. Rev. Microbiol. 32, 221-248. Grimont, P. A. D., and Grimont, E (1984). In "Bergey's Manual of Systematic Bacteriology" (N. R. Krieg and J. G. Holt, eds.), pp. 477-484. Williams & Wilkins, Baltimore. Grimont, P. A. D., Grimont, F., Richard, C., Davis, B. R., Steigerwalt, A. G., and Brenner, D. J. (1978). Deoxyribonucleic acid relatedness between Serratia plymuthica and other Serratia species, with a description of Serratia odorifera sp. nov. (type strain: 1CPB 3995). Int. J. Syst. Bacteriol. 28, 453-463. Guthrie, E. P., Flaxman, C. S., White, J., Hodgson, D. A., Bibb, M. J., and Chater, K. F. (1998). A response-regulator-like activator of antibiotic synthesis from Streptomyces coelicolor A3(2) with an amino-terminal domain that lacks a phosphorylation pocket. Microbiology 144, 727-738. [for erratum see Microbiology 144, 2007.]

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Rosenberg, M., Blumberger, Y., Judes, H., Bar-Ness, R., Rubinstein, E., and Mazor, Y. (1986). Cell-surface hydrophobicity of pigmented and nonpigmented clinical Serratia m a r c e s c e n s strains. Infect. I m m u n . 51,932-935. Santayana, G. (1930). Introduction to Benedictus Spinoza, Ethics a n d "De lntellectus E m e n d a t i o n e . " Trans. A. Boyle. Dent and Sons, London. (Original publication date, 1910.) Sato, T., Konno, H., Tanaka, Y., Kataoka, T., Nagai, K., Wasserman, H. H., and Ohkuma, S. (1998). Prodigiosin as a new group of H÷/C1- symporters that uncouple proton translocators. J. Biol. Chem. 273, 21455-21462. Sawabe, T., Makino, H., Tatsumi, M., Nakano, K., Tajima, K., Iqbal, M. M., Yumoto, I., Ezura, Y., and Christen, R. (1998). P s e u d o a l t e r o m o n a s bacteriolytica sp. nov., a marine bacterium that is the causative agent of red spot disease of Laminaria japonica. Int. f. Syst. Bacteriol. 48, 769-774. Scheurlen, E. (1896). Geschichtliche und experimentelle Studien fiber der Prodigiosus. Arch. Hyg. 26, 1-31. Songia, S., Mortellaro, A., Taverna, S., Fornasiero, C., Scheiber, E. A., Erba, E., Colotta, E, Mantovani, A., Isetta, A.-M., and Golay, J. (1997). Characterization of the new immunosuppressive drug undecylprodigiosin in human lymphocytes: Retinoblastoma protein, cyclin-dependent kinase-2, and cyclin-dependent kinase-4 as molecular targets. J. I m m u n o l . 158, 3987-3995. Syzdek, L. D. (1985). Influence of Serratia marcescens pigmentation on cell concentrations in aerosols produced by bursting bubbles. App]. Environ. Microbiol. 49, 173178. Thomson, N. R., Cox, A., Bycroft, B. W., Stewart, G. S. A. B., Williams, P., and Salmond, G. P. C. (1997). The Rap and Hor proteins of Erwinia, Serratia and Yersinia: A novel subgroup in a growing superfamily of proteins regulating diverse physiological processes in bacterial pathogens. Mol. Microbiol. 26, 531-544. Togashi, K.-i., Kataoka, T., and Nagai, K. (1997). Characterization of a series of vacuolar type H÷-ATPase inhibitors on CTL-mediated cytotoxicity. I m m u n o l . Lett. 55,139-144. Trachtenberg, J. (1943). "The Devil and the Jews: The Medieval Conception of the Jew and Its Relation to Modern Antisemitism." Yale University Press, New Haven. Tsao, S.-W., Rudd, B. A. M., He, X.-G., Chang, C.-J., and Floss, H. G. (1985). Identification of a red pigment from Streptomyces coelicolor A3(2) as a mixture of prodigiosin derivatives. J. Antibiot. 38, 128-130. Tsuji, R. E, Yamamoto, M., Nakamura, A., Kataoka, T., Magae, J., Nagai, K., and Yamasaki, M. (1990). Selective immunosuppression of prodigiosin 25-C and FK506 in the murine immune system. J. Antibiot. 43, 1293-1301. Tsuji, R. F., Magae, J., Yamashita, M., Nagai, K., and Yamasaki, M. (1992). Immunomodulating properties of prodigiosin 25-C an antibiotic which preferentially suppresses induction of cytotoxic T cells. J. Antibiot. 45, 1295-1302. Vaclav, J. (1994). Any less a miracle? A m . Soc. Microbiol. N e w s 60, 579. Van der Mei, H. C., Cowan, M. M., Genet, M. J., Rouxhet, P. G., and Busscher, H. J. (1992). Structural and physicochemical surface properties of Serratia marcescens strains. Can. J. MicrobioI. 38, 1033-1041. yon Graevenitz, A. (1980). Infection and colonization with Serratia. In "The Genus Serratia" (A. von Graevenitz and S. J. Rubin, eds.), pp. 167-186. CRC, Boca Raton, FL. Wainwright, M. (1990). "Miracle Cure: The Story of Penicillin and the Golden Age of Antibiotics." Basil Blackwell, Oxford.

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Microbial/Enzymatic Synthesis of Chiral Drug Intermediates RAMESH N. PATEL Bristol-Myers Squibb Pharmaceutical Research Institute New Brunswick, New Jersey 08903

I. Introduction II. AntihypertensiveDrug: Vasopeptidase Inhibitor A. EnzymaticSynthesis of BMS-199541-01 B. EnzymaticSynthesis of L-6-Hydroxynorleucine C. EnzymaticSynthesisof AllysineEthyleneAcetal III. [~3-ReceptorAgonist A. MicrobialReduction of 4-Benzyloxy-3-Methanesulfonylamino2'-Bromoacetophenone B. EnzymaticResolution of Racemic c~-MethylPhenylalanineAmides C. AsymmetricHydrolysisof Racemic Methyl-(4-Methoxyphenyl)Propanedioic Acid, Ethyl Diester IV. AnticholesterolDrugs V. Deoxyspergualin VI. AntiviralAgents VII. StereoselectiveHydrolysisof Racemic Epoxide VIII. BiocatalyticDynamicResolution: Stereoinversion of Racemic Diol IX. Resolutionof Racemic SecondaryAlcohols X. Summary References

I. Introduction Currently much attention is being focused on the interaction of small molecules with biological macromolecules. The search for selective enzyme inhibitors and receptor agonists/antagonists is key for targetoriented research in the pharmaceutical and agrochemical industries. Increasing understanding of the mechanism of drug interactions on a molecular level has led to a strong awareness of the importance of chirality as the key to the efficacy of many drug products and agrochemicals. The production of optically active chiral intermediates is a subject of increasing importance in pharmaceutical industries. Increasing regulatory pressure to market homochiral drugs by the Food and Drug Administration has driven chemoenzymatic synthesis of chiral compounds. Organic synthesis has been one of the most successful 33 ADVANCES IN APPLIED MICROBIOLOGY, VOLUME 47 Copyright © 2000 by Academic Press All rights of reproduction in any form reserved, 0065-2164/00 $25.00

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RAMESH N. PATEL

scientific disciplines and has enormous practical utility. There have been many advances in organic synthesis, which have led to the synthesis of natural products, drugs, agricultural chemicals, polymers, and many classes of functional molecules. This raises the question of why biocatalysis? What does biocatalysis have to offer to synthetic organic chemists? Biocatalysis offers an added dimension and an enormous opportunity to prepare industrially useful chiral compounds. One major advantage of biocatalysis over chemical catalysis is that enzyme-catalyzed reactions are stereoselective and regioselective and can be carried out at ambient temperature and atmospheric pressure, which minimizes problems of isomerization, racemization, epimerization, and rearrangements that may occur during chemical processes. Biocatalytic processes catalyzed by microbial cells and the enzymes derived therefrom can be immobilized and reused for many cycles. In addition, enzymes can be overexpressed so as to make biocatalytic processes economically efficient. The ability to design biocatalysts that would act specifically in any desired reaction will change the face of synthesis. Tailor-made enzymes with modified activity and the preparation of thermostable and pH-stable enzymes produced by random and site-directed mutagenesis will lead to the production of novel stereoselective biocatalysts. The use of enzymes in organic solvents has led to hundreds of publications on enzyme-catalyzed asymmetric synthesis and resolution processes. Molecular recognition and selective catalysis are key chemical processes in life that are embodied in enzymes. A number of review articles have been published on the use of biocatalysis in organic synthesis (Sih and Chen, 1984; Jones, 1986; Crout et al., 1994; Davies et al., 1990; Csuz and Glanzer, 1991; Crosby, 1991; Kamphuis et al., 1990a; Sih et al., 1992; Santaneillo et al., 1992; Margolin, 1993; Cole, 1994; Patel, 1997, 1998, 1999; Mori, 1995; Wong and Whitesides, 1994). This chapter provides some specific examples of the use of microbial enzymes for the synthesis of chiral drug intermediates.

II. Antihypertensive Drug: Vasopeptidase Inhibitor A. ENZYMATIC SYNTHESIS OF

BMS-199541-01

[4S-(4a,7a,10ab)] 1-octahydro-5-oxo-4-[[(phenylmethoxy)carbonyl]amino]7H-pyrido-[2,1-b] [1,3]thiazepine-7-carboxylic acid methyl ester (BMS199541-01) is a key chiral intermediate for the synthesis of Omapatrilat (BMS-186716), a new vasopeptidase inhibitor presently under development (Robl et al., 1997). Our goal was to prepare the compound by a

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

35

simpler, more convenient route using an intermediate derived from L-lysine as a readily available starting material. An enzymatic process was developed for the oxidation of the e-amino group of lysine in the dipeptide dimer N2-[N[[(phenylmethoxy)carbonyl] L-homocysteinyl] Llysine)l,l-disulfide (BMS-201391-01) to produce BMS-199541-01 (Fig. 1) using L-lysine e-aminotransferase from Sphingomonas paucimobilis SC16113 (Patel et at., 1999a). This enzyme was overexpressed in Escherichia coli and a biotransformation process was developed using the recombinant enzyme. The aminotransferase reaction required a-ketoo glutarate as the amine acceptor. The glutamate formed during this reaction was recycled back to a-ketoglutarate by glutamate oxidase from Streptomyces noursei SC6007. A selective culture technique was used to isolate microorganisms able to utilize N-a-CBZ-L-lysine as the sole source of nitrogen. Using this technique, eight different types of colonies were isolated. Cultures were grown in shake flasks, and cell extracts prepared from cell suspensions were evaluated for oxidation of the e-amino group of L-lysine in the substrate dipeptide dimer BMS-201391o01. Product (BMS199541-01) formation (0.05-0.35 mg/ml) was observed with four cultures. One of the cultures, Z-2, later identified as Sphingomonas pau-

PH

(~ CO2H

0 CO2H ,;

Dlthlothmitol or

LS

Tributylphosphine

/ I .NH2

../.5

PHN"'~U'~N"~ l,. H~ ~ H2N)

H2N

DiDeotide Monomer

O CO2H

L-lysine eaminotransferase

Dipe~)tide Dimer BMS-201391-01

SphlngomonaSor rec E.peucimobiliScoli

If (~-ketoglut,,rata~'~ k~ ) Glutamate Oxidase ~',~ Glutamate _j/ Stmptomycesnoursel

P=CBZ

oooo. BMS-199541-01

O

L

o co,.-1

x

F

o co,.-7

j

Other protecting group P= phenoxyacetyl otphenylacetyl

FIG. 1. Enzymatic conversion of dipeptide dimer BMS-201391-01 to BMS-199541-01 by L-lysine e-aminotransferase.

36

RAMESH N. PATEL

cimobi]is SC16113, exhibited higher activity (0.35 mg/ml of product formed) and was used for further studies. The low mass balance and reaction yield were due to hydrolysis of the substrate dipeptide by cell extracts. S. paucimobilis SC16113 was grown in a 700-liter fermentor containing 500 liters of medium. During fermentation, cells were harvested from 200 ml of broth by centrifugation. Cells were suspended in buffer, and cell extracts were prepared. Cell extracts were evaluated for conversion of BMS-201391-01 to BMS-199541-01. Cultures grown for 48 to 60 hours had higher specific activity compared to cells harvested at 24 or 72 hours. A specific activity (milligrams of BMS-199541-01 formed per hour per gram of protein in cell extract) of 220 was obtained for cultures grown for 60 hours. A preparative batch for biotransformation of BMS-201391-01 to BMS199541-01 using 2 liters of cell extract of S. paucimobHis SC16113 was prepared. Substrate was used at a concentration of 1.5 g/liter. A reaction yield of only 10% (0.3 g of BMS-199541-01) was obtained after 1.75 hours. The product was isolated and identified by 1H-NMR, 13C-NMR, and mass analysis. Due to the low activity of L-lysine e-aminotransferase in S. paucimobilis SC16113, we decided to purify the enzyme, determine its sequence, and overexpress the protein in a suitable host. The enzyme was purified 254-fold to a specific activity (mg product formed per hour per gram of protein) of 36,600. After Sephacryl S-200 column chromatography, the purified enzyme showed a single protein band on SDS/PAGE using a silver stain. The molecular weight of the enzyme as determined by gel-filtration techniques was 81,000 daltons, and the subunit size as determined by SDS/PAGE was 40,000 daltons, indicating that the L-lysine ~-aminotransferase is a dimeric protein. The N-terminal and internal peptide sequence (generated by Lys-peptidase treatment) of purified L-lysine aminotransferase were determined. The purified L-lysine ~-aminotransferase was evaluated for cofactor requirements. The enzyme required ¢t-ketoglutarate as an amine acceptor. NAD or NADP were not required as cofactors, indicating that the enzyme was an aminotransferase and not a dehydrogenase. A reaction yield of 70 mol% was obtained for BMS-199541-01 with the complete system. Glutamate oxidase was required to recycle glutamate back to ~-ketoglutarate. In the absence of glutamate oxidase, a 35 tool% reaction yield of BMS-199541-01 was obtained (Table I). The L-lysine ~-aminotransferase was overexpressed in E. coli strain GI724(pAL781-LAT). The enzyme was produced in a 25-liter fermentor.

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

37

TABLE I COFACTORREQUIREMENTSOF L-LYSINEE-AMINOTRANSFERASE:CONVERSION Or BMS-201391-01 TO BMS-199541-01

Reaction system

BMS-201391-01 (mg/ml)

BMS-199541-01 (mg/ml)

Yield (%)

0.57 3 0.9 3 0.58

1.94 0 0.98 0 1.92

70 0 35 0 69

Complete system Minus ct-ketoglutarate Minus glutamate oxidase Minus aminotransferase Minus NAD or NADP

The complete reaction mixture in 10 ml contained 6 ml of purified L-lysine aminotransferase (Sephacryl S-200 fraction), 1 ml of i M potassium phosphate buffer, pH 7.8, containing 5 mM dithiothreitol, 1 m M EDTA, 20 mg c~-ketoglutarate and 30 mg of BMS-201391.3 ml of glutamate oxidase (7 U/ml) was added during the 5-hr reaction time. The concentrations of BMS-199541-01 and BMS-201391-01 were determined by HPLC.

The enzyme activity ranged from 1700 to 2425 units/liter of broth. The kinetics of enzyme production are shown in Figure 2. Screening of microbial cultures led to identification of Streptomyces noursei SC6007 as a source of extracellular glutamate oxidase. S. noursei SC6007 was grown in 380-liter fermentors. During fermenta-

"20

3000 "

•

(units/Lofbroth)

o

2000-

~6

== ==

_J "10

1000

v

"

Z

}-

==

O J! 0

•

,

•

10 Fermentation

,

20 Time

0

30

(hours)

F~c. 2. Fermentation of recombinant Escherichia coli: Production of L-lysine E-aminotransferase.

38

RAMESH N. PATEL

tion, cells were periodically harvested by centrifugation from 200 ml of culture broth. The supernatant solution was used for determination of extracellular glutamate oxidase activity. Glutamate oxidase activity correlated with growth of the culture in a fermentor and reached 0.75 units/ml at harvest (Fig. 3). At the end of fermentation, the fermentation broth was cooled to 8°C and cells were removed by centrifugation. Starting from the extracellular filtrate recovered after removal of cells from the fermentation broth, the glutamate oxidase was purified 260fold with a specific activity (units per milligram of protein) of 54. The purified enzyme showed a single protein band on SDS/PAGE using a silver stain. The molecular weight of the enzyme as determined by gel-filtration techniques was 125,000 daltons and the subunit size as determined by SDS/PAGE 60,000 daltons, indicating that the glutamate oxidase is a dimeric protein. The amino-terminal and internal peptide sequence of the purified enzyme were determined to allow for the synthesis of oligonucleotide probes for cloning and overexpression of the enzyme. Attempts to express the S. noursei SC6007 glutamate oxio dase using standard E. coli vectors and strains were unsuccessful. As an alternative, the SC6007 glutamate oxidase was expressed in Strep-

0.8

~'

.IL

P m t l s l v o l u m e of solids

"30 i

0.6

'20 c

o ~

0,4

o o >

10

,

~

0.2

.~ Q.

utamate oxldase 0 . 0 -0

10

20

30

40

Fermentation Time (Hours)

FIG. 3. F e r m e n t a t i o n of Streptomyces noursei SC6007: Production of glutamate oxidase.

39

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

tomyces lividans. The S. noursei SC6007 glutamate oxidase, including its native promoter sequence, was inserted into an S. lividans expression vector. Untransformed S. lividans does not have a native glutamate oxidase activity, while S. lividans transformed with the GOX expression plasmid demonstrated glutamate oxidase activity. SDS/PAGE analysis of the transformed S. lividans revealed a protein band not seen in an untransformed strain. This band was of the same molecular weight as the GOX protein purified from S. noursei SC6007, indicating that the glutamate oxidase activity present in the transformed strain arose from expression of the heterologous gene. About 0.4 units/ml of activity was detected from the S. lividans culture, indicating that the enzyme was expressed at a low level. Further research was required to overexpress this protein. Biotransformation of BMS-201391-01 to BMS-199541-01 was carried out using L-lysine s-aminotransferase from Escherichia coli GI724[pa1781-LAT] in the presence of ct-ketoglutarate and dithiothreitol (required to reduce the dipeptide dimer to a monomer). Glutamate produced during the reaction was recycled to o~-ketoglutarate by partially purified glutamate oxidase (7 units/ml) from S. noursei SC6007. Four different batches were carried out. Reaction yields of 65-70 mol% were obtained as shown in Table II. The kinetics of reaction are shown in Figure 4. Two n e w dipeptides, No[N[(phenylmethoxy)carbonyl]-L-methionyl]L-lysine (BMS-203528-01) and N,2-[S-acetyl-N-[(phenylmethoxy)carbonyl]-L-homocysteinyl]-L-lysine (BMS-204556), were evaluated as

TABLE II BIOTRANSFORMATIONOF BMS-201391-01 TO BMS-199541-01 BY L-LYSINE £-AMINOTRANSFERASEFROM ESCHERICHIACOLIGI724[pa1781-LAT] Experiment batch number

BMS-201391-01 input (g)

BMS-201391-01 remaining (g)

BMS-199541-01 01 {g)

BMS-199541-01 (mol% yield)

40455 40456 40457 40458

3 5 12 22

0.83 1.35 4.3 4.7

1.9 2.92 7.5 14.4

66.5 65 70 67

R e a c t i o n s were carried o u t as d e s c r i b e d in t h e text u s i n g cell extracts of Escherichia coli GI724[pa1781-LAT] in t h e p r e s e n c e of dithiothreitol a n d partially purified g l u t a m a t e o x i d a s e from Streptomyces noursei SC6007.

40

RAMESH N. PATEL

A .J ~,

~-

BMS-199541-01

r,

o

i

/ o 0

& i 100

BMS-201391-01

200

300

Reaction time (min)

FIG. 4. Kinetics of oxidation of dipeptide dimer BMS-201391-01 to BMS-199541-01 by L-lysine E-aminotransferase. Regeneration of c¢-ketoglutarate was carried out by glutamate oxidase.

substrates for L-lysine aminotransferase by cell-free extracts of E. coli GI724[pa1781-LAT] in the presence of a-ketoglutarate. The formation of new compounds from the enzymic reaction was investigated by liquid chromatography-mass spectrometry (LC-MS). The data indicate the of a n e w c o m p o u n d with a molecular weight of 392, which was assigned tentative structure 1. The e-NH 2 group of BMS-203528 was oxidized, and in the presence of trichloroacetic acid (TCA) the aldehyde was cyclized to the enamide with a loss of water (Fig. 5). When BMS-204556 was treated with cell-free extracts of E. coli GI724[pa1781-LAT] and o~-ketoglutarate, several new components were observed by LC-MS. The component with a molecular weight of 420.5 was assigned structure 2, formed by oxidation of the ~-NH2 group of BMS-204556 and subsequent dehydration to produce the cyclic enamide; the component with a molecular weight of 397 was proposed as desacetyl BMS-204556 3. The desacetyl BMS-204556 was then oxidized by the enzyme to BMS-199541-01 (MW = 378), as shown in Figure 6.

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

0

41

COOH L-lysine e - a n g n m r a n s f e r a s e f r o m S. pm~cimobilis o r rec E. coli

\S --NH2 CH3 I

BMS-203528-01 M o l wt. 411.62

~NCbz 0 |I"COOHH+ t ....

L | L_

CH3 M o l wt. 4 1 0

NCbz""~ CH3 Molwe.392 !

not observed in M S

FIG. 5. Enzymatic oxidation of BMS-203528-01 by L-lysine e-aminotransferase. Regeneration of a-ketoglutarate was carried out by glutamate oxidase.

To reduce the cost of producing two enzymes, the transamination reactions were carried out in the absence of glutamate oxidase and with higher levels of ~-ketoglutarate. The reaction yield in the absence of glutamate oxidase averaged about 33-35 mol%. With 40 mg/ml of t~-ketoglutarate (10-fold increase in concentration) and at 40°C, the reaction yield increased to 70 mol%, equivalent to that in the presence of glutamate oxidase. Phenylacetyl- or phenoxyacetyl-protected analogues of BMS-201391-01 (Fig. 1) also served as substrates for L-lysine e-aminotransferase, giving a reaction yield of 70 mol% for the corresponding BMS-199541-01 analogues. N-a-t-bntoxycarbonyl-L-lysine and N-a-carbobenzoxy-L-lysine were also oxidized by L-lysine aminotransferase from E. coli GI724[pa1781LAT]. The chiral compounds (S)-3,4-dihydro-l,2(2H)-pyridinedicarboxylic acid, 1-(phenylmethyl)ester (BMS-202665), (S)-3,4-dihydro-l,2(2H)-pyridinedicarboxylic acid, and 1,1-dimethylethyl ester (BMS264406) were obtained as products of oxidation of N-a-CBZ-L-lysine and N-ct-BOC-L-lysine, respectively (Fig. 7). A reaction yield of 80-85 mol% was obtained for each product. In the enzymatic reaction to convert BMS-201391-01 to BMS-19954101, we used dithiothreitol (DTT) to cleave the disulfide bond of the dipeptide dimer BMS-201391 to produce the dipeptide monomer, which was the substrate for the L-lysine aminotransferase. It was observed that tributylphosphine (an inexpensive compound) was as effective as DTT for the dipeptide dimer to monomer conversion. In the presence of 10-mM tributylphosphine, 3.5 mg/ml of BMS-201391-01, 40 mg/ml ~-ketoglutarate, and 0.1 units of transaminase, a 69 mol% yield of BMS199541-01 was obtained.

42

RAMESH N. PATEL

©

z

o

o~~

.~

z

u

[

~ g

tt~ e~

¢x]

'-..

0

Z

z

0

k Z

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

NH 2 NHP Nc~-protected-L-lysine P = BOC or CBZ

L-lysine e-AminotransferDase

H20

43

COOH BMS-202665, P=CBZ BMS-264406, P= BOC

FIG. 7. Enzymatic oxidation of N-a-protected L-lysine by L-lysine e-aminotransferase. Regeneration of c¢-ketoglutarate was carried out by glutamate oxidase.

To terminate the L-lysine aminotransferase reaction during conversion of BMS-201391-01 to BMS-199541-01, 10% vol/vol trichloroacetic acid (TCA) was used. It was also observed that a much cheaper compound, methanesulfonic acid, is equally effective as TCA, giving a 70 tool% yield of BMS-199541-01. Aminotransferases have been used extensively in the synthesis of L-amino acids from the corresponding ~-ketoacids (Stirling, 1992; Kamphuis et al., 1990b). L-lysine a-ketoglutarate aminotransferase from Flavobacterium fuscum was reported by Soda et al. (1968), and they demonstrated that the product of L-lysine oxidation is 1-piperidine-6carboxylic acid. In this aminotransferase reaction, the E-amino group of L-lysine is transferred to a-ketoglutarate to yield glutamate and a-aminoadipate-8-semialdehyde, which is immediately converted into the intramolecular dehydrated compound 1-piperidine-6-carboxylic acid. The oxidation of N-o~-carbobenzoxy and Noa-t-butoxycarbonyl L-lysine by Rhodotorula graminis to produce novel chiral compounds (S)-3,4dihydro-l,2(2H)-pyridinecarboxylic acid, 1-(phenylmethyl)ester, (S)3,4-dihydro-l,2(2H)-pyridinecarboxylic acid, and 1-dimethylethyl ester has been demonstrated by Patel et al. (1999b). Soda and Misono (1968) reported that L-lysine cx-ketoglutarate aminotransferase (MW = 116,000) contained two molecules of pyridoxal phosphate as a bound prosthetic group. Hammer and Bode (1992) purified L-lysine a-ketoglutarate aminotransferase from Candida utilis and reported that it is a dimeric 83,000-dalton protein. L-lysine aminotransferase from S. paucimobi]is SC16113 was a dimeric 80,000-dalton protein. B. ENZYMATICSYNTHESISOF L-6-HYDROXYNORLEUCINE L-6-hydroxynorleucine (4, Fig. 8) is a chiral intermediate useful for the synthesis of a vasopeptidase inhibitor now in clinical trials, and for the

44

RAMESH N. PATEL

glucose

",,,.._../

gluconicacid

glucosedehydrogenase NADH

~O~COzNa OH

O HoIV~

NAD

glutamate dehydrogenase ONa

NH3~

2-keto-6-hydroxyhexanoic acid, sodiumsalt5

H

~

NH2 OH

L-6-hydroxynorleucine 4

FIG. 8. Reductive amination of sodium 2-keto-6-hydroxyhexanoic acid 5 to L-6-hydroxynorleucine 4_by glutamate dehydrogenase.

synthesis of C-7 substituted azepinones as potential intermediates for other antihypertensive metalloprotease inhibitors (Robl and Cimarusti, 1994; Robl et al., 1997). It has also been used for the synthesis of siderophores, indospicines, and peptide hormone analogues (Maurer and Miller, 1981, 1982, 1983; Bodanszky et al., 1978; Dreyfuss, 1974). Previous synthetically useful methods for obtaining this intermediate have involved synthesis of the racemic compound followed by enzymatic resolution. D-amino acid oxidase has been used to convert Damino acid to the ketoacid, leaving the L-enantiomer that was isolated by ion exchange chromatography (Kern and Reitz, 1978). In a second approach, racemic N-acetylhydroxy norleucine has been treated with L-amino acid acylase to give the L-enantiomer (Robl et al., 1997). Both of these resolution methods give a maximum 50% yield and require separation of the desired product from the unreacted enantiomer at the end of the reaction. Reductive amination of ketoacids using amino acid dehydrogenases has been shown to be a useful method for synthesis of natural and unnatural amino acids (Bommarius, 1995; Galkin et al., 1997). We have developed the synthesis and conversion of 2-keto-6-hydroxyhexanoic acid 5 to L-6-hydroxy norleucine 4_ (Fig. 8) by reductive amination using beef liver glutamate dehydrogenase and glucose dehydrogenase from a Bacillus sp. for regeneration of NADH (Hanson et al., 1999). To avoid the lengthy chemical synthesis of the ketoacid, a second route was developed to prepare the ketoacid by treatment of racemic 6-hydroxy norleucine, readily available from hydrolysis of 5-(4-hydroxybutyl) hydantoin 6, with D-amino acid oxidase from porcine kidney or Trigonopsis variabilis and catalase followed by reductive amination to convert the mixture to L-6-hydroxynorleucine.

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

45

2-keto-6-hydroxyhexanoic acid 5 is converted completely to L-6hydroxy norleucine 4 by phenylalanine dehydrogenase from a Sporosarcina sp. and beef liver glutamate dehydrogenase, with formate dehydrogenase for regeneration of NADH (Hanson et al., 1999). Beef liver glutamate dehydrogenase was used for preparative reactions at a substrate concentration of 100 g/liter. As depicted in Figure 8, 2-keto6-hydroxyhexanoic acid 5, sodium salt, in equilibrium with 2-hydroxytetrahydropyran-2-carboxylic acid, sodium salt, was converted to L-6o hydroxynorleucine. The reaction requires ammonia and reduced NADH. NAD produced during the reaction was recycled to NADH by oxidation of glucose to gluconic acid using glucose dehydrogenase from Bacillus megaterium. The reaction was completed within about 3 hr with reaction yields of 89-92% and an enantiomeric excess of >98% for L-6-hydroxynorleucine. Chemical synthesis and isolation of 2-keto-6-hydroxyhexanoic acid required several steps. In a second more convenient process (Fig. 9),

o

~/

H2N~/~H C02H

H Ca{OH)p NaOH

="

H

Racemic 6-hydroxynorleucine

5-(4-hydroxybutyl) hydantoin _6 glucose

gluconic acid

glucose dehydrogenase NADH NH2

D-amino acid oxidase

0

5_

NAD

glutamate dehydrogenase O

NH3

H

~

NH~ OH

0 +

Racemic 6-hydroxynorleucine

NH2

H

~

OH © catalase + H202

H20 +

02

+ NH3

FIG. 9. Enzymatic conversion of racemic 6-hydroxynorleucine to L-6-hydroxynorleucine 4 by D-amino acid oxidase and glutamate dehydrogenase.

RAMESH N. PATEL

46

the ketoacid was prepared by treatment of racemic 6-hydroxynorleucine (produced by hydrolysis of 5-(4-hydroxy butyl)hydantoin 6) with D-amino acid oxidase and catalase. After the enantiomeric excess of the remaining L-6-hydroxynorleucine had risen to >99%, the reductive amination procedure was used to convert the mixture containing 2keto-6-hydroxy hexanoic acid and L-6-hydroxynorleucine entirely to L-6-hydroxynorleucine with yields of 91-97% and enantiomeric excess of >98%. Sigma porcine kidney D-amino acid oxidase and beef liver catalase or T. variabilis whole cells (source of oxidase and catalase) were used successfully for this transformation. C. ENZYMATIC SYNTHESIS OF ALLYSINE ETHYLENE ACETAL

(S)-2-amino-5-(1,3-dioxolan-2-yl)-pentanoic acid (allysine ethylene acetal _7) is one of three building blocks used for an alternative synthesis of omapatrilat, a vasopeptidase inhibitor (Robl et al., 1997). It was previously prepared in an eight-step synthesis from 3,4-dihydro-2Hpyran for conversion into 1-piperidine-6-carboxylic acid, an intermediate for biosynthesis of [3-1actam antibiotics (Rumbero et aL 1995). The reductive amination of ketoacid acetal 8 to acetal amino acid 7 was demonstrated using phenylalanine dehydrogenase from Thermoactinomyces intermedius (Fig. 10). The reaction requires ammonia and NADH. NAD produced during the reaction was recycled to NADH by oxidation of formate to CO2 using formate dehydrogenase (Hanson et

/--1

/--1O

o m o .nase Ammonium f o r m a ~

C02

NAD~--IcNAD OH

Phenylalanine dehydrogenase

H2N

COOH

8

FIG. 10. Reductive amination of ketoacid acetal 8 to amino acid acetal _7by phenylalanine dehydrogenase. Regeneration of NADH was carried out using formate dehydrogenase.

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

47

al., 2000). An initial process was developed using heat-dried cells of T. intermedius ATCC 33205 as a source of phenylalanine dehydrogenase and heat-dried cells of methanol-grown Candida boidinii as a source of formate dehydrogenase. An improved process using phenylalanine dehydrogenase from T. intermedius expressed in E. coli BL21(DE3) (pPDH155K) (SC16144) in combination with C. boidinii as a source of formate dehydrogenase and a third-generation process using methanol-grown Pichia pastoris containing endogenous formate dehydrogenase and expressing T. intermedius phenylalanine dehydrogenase were also developed (Hanson et al., 2000). Glutamate, alanine, leucine, and phenylalanine dehydrogenases (listed in order of increasing effectiveness) converted _8 to the desired amino acid (7) (Table III). The product was identical by HPLC and MS analysis to a chemically synthesized standard. Some alternative sources of phenylalanine dehydrogenase were tested. Sporosarcina ureae strains SC16048 and SC16049 had respective specific activities of 0.996 and 0.862 g/rag for reductive amination of phenylpyruvate, but amination of 8 was m u c h slower than with the enzyme from Thermoactinomyces. Using an extract of T. intermedius ATCC 33205 as a source of phenylalanine dehydrogenase and Boehringer Mannheim formate dehydrogenase for NADH regeneration increased the estimated yield to 80%, and the process was developed using this enzyme combination. Heat-dried cells of T. intermedius and C. boidinii SC13822 grown on methanol were used for the reaction. Phenylalanine dehydrogenase activities in cells recovered from fermentation and fermentor productivities are shown in Table IV. T. intermedius gave a useful activity on a small scale (15 liters), but lysed soon after the end of the growth period, making recovery of activity difficult

TABLE III REDUCTIVE AMINATIONOF KETO ACID 8 BY AMINO ACID DEHYDROGENASE

Dehydrogenase

Source

Glutamate

Beef liver

Alanine

Bacillus s u bti]is

Leucine

Bacillus s p h a e r i c u s

Phenylalanine

Sporosarcina spp.

Amount (units)

A m i n o acid _7 produced (mM)

76

1.03

35.7

11.77

22

14.01

12.6

51.7

48

RAMESH N. PATEL TABLE IV ACTIVITIESAND PRODUCTIVITIESOF PHENYLALANINEDEHYDROGENASEAND FORMATE DEHYDROGENASEFOR VARIOUSSTRAINSGROWNIN A FERMENTOR

Enzyme Phenylalanine dehydrogenase

Formate dehydrogenase

Strain

Thermoactinomyces intermedius Escherichia coil Pichia pastoris Candida boidinii Pichia pastoris

Specific activity (U/g wet ceils)

Volumetric activity (U/liter of broth)

510

185

900

10,000 ND

24,000 14,500

94,000 25,000

9 26

120 1950

350 3200

Producivity (U/liter/week)

or impossible on a large scale (4000 liters). The problem was solved by cloning and expressing the T. intermedius phenylalanine dehydrogenase in Escherichia coli, inducible by isopropylthiogalactoside. Fermentation of T. intermedius yielded 184 units of phenylalanine dehydrogenase activity per liter of whole broth in 6 hours. At harvest the fermentor needed to be cooled rapidly, because the activity was unstable. In contrast, E. coli produced more than 19,000 units per liter of whole broth in about 14 hours and was stable at harvest. C. boidinii grown on methanol was a useful source of formate dehydrogenase, as has been shown previously (Schfitte et al., 1976). In order to recover the cells on a large scale, it was helpful to add 0.5% methanol to stabilize the cells. P. pastoris grown on methanol was also a useful source of formate dehydrogenase (Hou et al., 1982). Expression of T. intermedius phenylalanine dehydrogenase in P. pastoris, inducible by methanol, allowed both enzymes to be obtained from a single fermentation. The expression of the two activities during a P. pastoris fermentation is shown in Figure 11. Formate dehydrogenase activity per gram wet cells was 2.7-fold greater than for C. boidinii, and fermentor productivity was increased by 8.7-fold compared to C. boidinii. Fermentor productivity for phenylalanine dehydrogenase in P. pastoris was about 28% of E. coli productivity. Formate dehydrogenase has been reported to have a pH optimum of 7.5 to 8.5 (Schfitte et al., 1976). The pH optimum for reductive amination of 8 by an extract of T. intermedius was found to be about 8.7.

49

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

3000

20000

--

PDH

¸2000

FDH 10000 ¸

1000

f.

t

--

0

i

• 20

40

80

60

Fermentation Time (Hours)

FrG. 11. Fermentation of Pichia pastoris for production of recombinant phenylalanine dehydrogenase and endogenous formate dehydrogenase.

Reductive amination reactions were carried out at pH 8.0. A summary of lab scale l-liter batches is shown in Table V. The time course for a representative batch showing conversion of ketoacid 8 to amino acid 7 is presented in Figure 12 using E. coli/C, boidinii heat-dried cells. The procedure using heat-dried cells of E. coli containing cloned phenylalanine dehydrogenase and heat-dried C. boidinii was scaled up

TABLE V LABORATORYSCALE(1 LITER)BATCHESFOR REDUCTIVEAMINAT1ONREACTIONS Phenylalanine dehydrogenase source

Formate dehydrogenase source

Reaction yield of _7 (%)

EE of product _7 (%)

T. intermedius

Candida boidinii

85

>99

Escherichia coil

Candida boidinii

90

>99

Pichia pastoris

Pichia pastoris

94

>99

50

RAMESH N. PATEL 60

50 I

40-

30-

20

10

0

10

20

30

Reaction T i m e (Hours)

FIG. 12. Kinetics of p r o d u c t i o n of amino acid acetal 7 from ketoacid acetal _8 by p h e n y l a l a n i n e dehydrogenase. Regeneration of NADH was carried out using formate dehydrogenase.

(Table VI), A total of 197 kg of compound 7 was produced in three 1600-liter batches using a 5% concentration of substrate _8 with an average yield of 91.1 mol% and enantiomeric excess greater than 98%.

TABLE VI PREPARATIVESCALEBATCHESFOR REDUCTIVEAMINATIONOF KETO ACID ~l

Phenylalanine dehydrogenase source

Formate dehydrogenase source

Keto acid 8 input (ks)

Amino acid 7 output (ks)

Reaction yield of _7 (tool%)

Escherichia coli Escherichia coli Escherichia coli Pichia pastoris

Candida boidinii Candida boidinii Candida boidinfi Pichia pastoris

80.17 79.96 89.6 18.05

62.40 66.75 67.61 15.51

92 96 86 97.5

EE of amino acid _7 (%) >99 >99 >99 >99

SYNTHESIS OF CHIRALDRUGINTERMEDIATES

51

A third-generation procedure using dried recombinant P. pastor& containing T. intermedius phenylalanine dehydrogenase inducible with methanol and endogenous formate dehydrogenase induced when P. pastoris was grown in medium containing methanol allowed both enzymes to be produced during a single fermentation, and they were conveniently produced in about the right ratio for the reaction. The Pichia reaction procedure had the following modifications of the E. coli/C, boidinii procedure: concentration of substrate was increased to 100 g/liter, a quarter of the amount of NAD was used, and DTT was omitted. The procedure with P. pastoris was also scaled up to produce 15.5 kg of 7 with a 97 mol% yield and enantiomeric excess greater than 98% (Table VI) in a 180-liter batch using 10% ketoacid 8 concentration. For reusability, formate dehydrogenase could be immobilized on Eupergit C and phenylalanine dehydrogenase on Eupergit C250L. The immobilized enzymes were tested for reusability in a jacketed reactor maintained at 40°C and were used five times for conversion of 8 to 7 without much loss of activity or productivity. At the end of each reaction, the solution was drained from the reactor through a 80/400 mesh stainless steel sieve, which retained the immobilized enzymes; then the reactor was recharged with flesh substrate solution. After five reuses, the reaction rate was decreased; however, the original reaction rate was restored in the seventh test study by addition of formate dehydrogenase. T, intermedius IFO14230 (ATCC 33205) was first identified as a source of phenylalanine dehydrogenase by Ohshima et al. (1991). The enzyme was purified and characterized, then cloned and expressed in E. coli by Takada et al. (1991). The enzyme was reported to be rather specific for deamination of phenylalanine (Ohshima et al., 1988), and to carry out amination of some ketoacids at a much lower rate than amination of phenylpyruvate. In our screening, the enzyme was the most effective amino acid dehydrogenase identified for bioconversion of 8 to 7. Formate dehydrogenase from C. boidinii was introduced by Shaked and Whitesides (1980), and by Kula and Wandrey (1987) for regeneration of NADH. The advantages of this enzyme reaction are that the product CO2 is easy to remove and that the negative reduction potential (E'° -- -0.42 V) for the formate dehydrogenase reaction drives reductive amination to completion. Previously, we prepared L-[3-hydroxyvaline from ~-keto-13-hydroxyisovalerate by the enzymatic reductive amination reaction using leucine dehydrogenase from Bacillus sphaericus ATCC 4525. L-[3-hydroxyvaline is a key chiral intermediate for the synthesis of tigemonam, an antiinfective drug (Hanson et al., 1990).

52

RAMESH N. PATEL

III. 133-Receptor Agonist [3-adrenoceptors have been classified as 131 and 132 (Land et al., 1967). Increased heart rate is the primary consequence of (31-receptor stimulation, while bronchodilation and smooth muscle relaxation are mediated from [32 receptor stimulation. Rat adipocyte lipolysis was initially thought to be a (51-mediated process (Land et al., 1967). However, recent results indicate that the receptor-mediated lipolysis involves neither I~1 nor [32, but "atypical" receptors, later called 133-adrenergic receptors (Arch, 1997). 133-adrenergic receptors are found on the cell surface of both white and brown adipocytes and are responsible for lipolysis, thermogenesis, and relaxation of intestinal smooth muscle (Arch et al., 1984). Consequently, several research groups are engaged in developing selective [~3 agonists for the treatment of gastrointestinal disorders, type II diabetes, and obesity (Wilson et al., 1984; Bloom et a]., 1989; Fisher et al., 1994; Sher, 1994). Efficient biocatalytic synthesis of chiral intermediates required for total chemical synthesis of [33 receptor agonist have been reported (Patel et al., 1998). These include: (a) microbial reduction of 4-benzyloxy-3-methanesulfonylamino-2'-bromoacetophenone 9 to corresponding (R)-alcohol 10 by Sphingomonas paucimobilis SC16113, (b) enzymatic resolution of racemic a-methyl phenylalanine amide 11 and c~-methyl-4-hydroxyphenylalanine amide 13 by amidase from Mycobacterium neoaurum ATCC 25795 to prepare the corresponding (S)-amino acids 12 and 14, and (c) asymmetric hydrolysis of methyl-(4-methoxyphenyl)propanedioic acid ethyl diester 15 to the corresponding (S)-monoester 16 by pig liver esterase. A. MICROBIAL REDUCTION OF 4-BENZYLOXY-3-METHANESULFONYLAMINO-2'-BROMOACETOPHENONE

Microbial reduction of 4-benzyloxy-3-methanesulfonylamino-2'-bromoacetophenone 9- to the corresponding (R)-alcohol 10 was demonstrated using S. paucimobilis SC16113 (Fig. 13). Among cultures evaluated, Hansenula anamola SC13833, Hansenula anamola SC16142, Rhodococcus rhodochrous ATCC 14347, and S. paucimobilis SC16113 gave the desired alcohol 10 in >96% enantiomeric excess and >15% reaction yield. S. paucimobilis SC16113 in the initial screening catalyzed the efficient conversion of ketone 9_to the desired chiral alcohol 10 with 58% reaction yield and >99.5% enantiomeric excess. Since substrate 9 is insoluble in water, the effect of solvents to dissolve substrate 9 and supply it in the biotransformation reaction mixture was evaluated. Dimethylformamide at 2-5% concentrations

SYNTHESIS OF CHIRALDRUG INTERMEDIATES

53

O / . . ~ ~ O

~HSO2CH3

OH Bspingomonaspaucimobilis

SC16113

OJ ~ NH B r

~"

SO2CH3 Product10 (R)-Alcohol

Substrate9 Ketone

OH

OH H o

H

(

/CH2R

CH3 ~I'~OCH2CO2H CI

BRL37344

NHSO2CH3~ J BMS-210620 OCH3

FIc. 13. Stereoselective reduction of 4-benzyloxy-3-methanesulfonylamino-2'-bromoacetophenone 9 to corresponding [R)-alcohol 10 by Sphingomonas paucimobilis SC16113. Structure of antiviral compounds BRL-37344 and BMS-210620,

was found to be the best cosolvent to supply the substrate in the biotransformation process. The fermentation of S. paucimobilis SC16113 was carried out in a 750-liter fermentor. From each fermentation batch, about 60 kg of wet cell paste was collected. Cells harvested from the fermentor were used to conduct the biotransformation in 1-, 10-, and 210-liter preparative batches under aerobic or anaerobic conditions. The cells were suspended in 80-mM potassium phosphate buffer (pH 6.0) to 20% (wt/vol, wet cells) concentration. Compound 9 (1-2 g/liter) and glucose (25 g/liter) were added to the fermentor, and the reduction reaction was carried out at 37°C. In some batches, the microfiltered and diafiltered ceils were used directly in the bioreduction process. In all biotransformation batches, a reaction yield of >85% and an enantiomeric excess of >98% were obtained. The isolation of chiral alcohol 10 from the 200-liter preparative batch was carried out to obtain 100 g of product 10. The isolated 10 gave a homogeneity index (HI) of 83% and an enantiomeric excess of 99.5% as analyzed by chiral HPLC. The MS and NMR data of isolated compound 10 and standard compound 10 were virtually identical. In an alternate process, frozen cells of S. paucimobilis SC16113 were used with resin-adsorbed (XAD-16 resin) substrate at 5 and 10 g/liter substrate concentrations. In this process, an average reaction yield of 85% and an enantiomeric excess of >99% were obtained for chiral

54

RAMESH N. PATEL

alcohol 10. At the end of the biotransformation, the reaction mixture was filtered on a 100-mesh (150 m) stainless steel screen, and the resin retained by the screen was washed with 2 liters of water. The product was then desorbed from the resin and crystallized in an overall 75 mol% yield with 91% homogeneity and 99.8% enantiomeric excess. The reduction of compound 9 to compound 10 was also carried out using cell extracts of S. paucimobilis SC16113. Glucose dehydrogenase was used to regenerate the cofactor NADPH required for the reduction. After a 90-min reaction time, 80% conversion of ketone 9 to chiral alcohol 10 was obtained. B. ENZYMATICRESOLUTION OF RACEMIC m-METHYL PHENYLALANINE AMIDES

The enzymatic resolution of racemic m-methyl phenylalanine amide 11 and ct-methyl-4-hydroxyphenylalanine amide 13 to the corresponding (S)-amino acids 12 and 14 (Fig. 14), respectively, by an amidase from Mycobacterium neoaurum ATCC 25795 was demonstrated by Patel et

0

0

M. neoaurumATCC2795

o

~

+

Product 12

Substrate 11

O H3C, ~ t

O NH2

H3C

M. neoaurumATCC25795

¢~ -

~ "OH

~.~

OCH 3

OCH 3

Substrate 13

Product 14

..,,,.~ R NH2

4-

H3

OCH 3

FIG. 14. Enantioselective enzymatic hydrolysis of ct-methyl phenylalanine amide 11 and c~-methyl-4-hydroxyphenylalanineamide 13 to corresponding (S)-amino acids by amidase from Mycobacterium neoaurum ATCC 25795.

SYNTHESIS OF CHIRALDRUG INTERMEDIATES

55

al. (1998). The chiral amino acids are intermediates for synthesis of a [33-receptor agonist (Bloom et al., 1989; Baroni eta]., 1994). The cells (10% wt/vol, wet cells) of M. n e o a u r u m ATCC 25795 were

evaluated for biotransformation of compound 11 to compound 12. The reaction was completed in 75 min with a reaction yield of 48 mol% (theoretical maximum = 50%) and an enantiomeric excess of 95% for the desired product 12. Freeze-dried cells of M. n e o a u r u m ATCC 25795 were suspended in 100-mMpotassium phosphate buffer (pH 7.0) at 1% concentration, and cell suspensions were used for biotransformation of compound 11. The reaction was completed in 60 min with a reaction yield of 49.5 mol% (theoretical maximum = 50%) and an enantiomeric excess of 99% for the desired product 12 (Fig. 14). Biotransformation of compound 11 was also carried out using a purified amidase. A reaction yield of 49 mol% and an enantiomeric excess of 99.8% were obtained for desired product 12 after a 60-min reaction time. Freeze-dried cells ofM. n e o a u r u m ATCC 25795 and partially purified amidase were used for biotransformation of compound 13. A reaction yield of 49 mol% and an enantiomeric excess of 78% were obtained for the desired product 14 using freeze-dried cells. The reaction was completed within 50 hours. Using partially purified amidase, a reaction yield of 49 tool% and a higher enantiomeric excess of 94% were obtained for desired product 14 after a 70-hr reaction time.

C. ASYMMETRICHYDROLYSISOF RACEMICMETHYL-(4-METHOXYPHENYL)-PROPANEDIOICACID, ETHYL DIESTER

The enzymatic asymmetric hydrolysis of methyl-(4-methoxyphenyl)propanedioic acid ethyl diester 15 to the corresponding (S)-monoester 16 by pig liver esterase has been demonstrated (Fig. 15). Chiral (S)monoester is a key intermediate for the synthesis of [33-receptor agonists. Various organic solvents were tested for the PLE-catalyzed asymmetric hydrolysis of diester 15 in a biphasic system. The results (Table VII) indicate that the reaction yields and enantiomeric excess of monoester 16 were dependent on the solvent used in asymmetric hydrolysis. Tetrahydrofuran, methyl ethyl ketone (MEK), methylisobutyl ketone (MIBK), hexane, and dichloromethane inhibited PLE. Lower reaction yields (28-56 mol%) and lower enantiomeric excess (59-72%) were obtained using t-butylmethyl ether, dimethylformamide (DMF), and

56

R A M E S H N. PATEL

/~

CO2C2H5 Pig LiverEsterase

H3CO-

~3CO-

...(.j

Diester 15

S-(-)-Monoester 16

FIG. 15. A s y m m e t r i c h y d r o l y s i s of r a c e m i c m e t h y l - ( 4 - m e t h o x y p h e n y l ) - p r o p a n e d i o i c acid e t h y l diester 15 to t h e c o r r e s p o n d i n g (S)-monoester b y pig liver esterase.

TABLE VII EFFECT OF SOLVENTON ASYMMETRIC HYDROLYSISOF METHYL-(4-METHOXYPHENYL)

PROPANEDIOIC ACID EHYL DIESTER 15 Enantiomeric Reaction time (hours)

Diester 15 (mg/ml)

Monoester 16 (mg/ml)

Yield (mol%)

of m o n o ester 16(%)

Methanol

22

0

0.65

37

92

Ethanol

22

0

1.7

96.7

96

Acetonitrile

22

0

0.5

28.2

59.3

Dimethylformamide

22

0

0.85

48.3

68.5

Dimethylsulfoxide

22

0.61

1

56.9

72

Acetone

22

0

1.44

81.9

65.1 82.1

Solvent

excess

Methylethylketone

48

0

1.36

77.3

Methylisobutylketone

64

2.01

0

0

-

t-butylmethylether

22

O. 76

0.8

46

64.4

Tetrahydrofuran

48

2

0

0

-

Toluene

22

0.18

0.59

33.6

91

Hexane

64

2.05

0

0

-

SYNTHESIS OF CHIRALDRUGINTERMEDIATES

57

dimethylsulfoxide (DMSO) as cosolvents. Higher enantiomeric excesses (>91%) were obtained using methanol, ethanol, and toluene as cosolvents. Ethanol gave the highest reaction yield (96.7%) and enantiomeric excess (96%) for monoester 16. The effect of temperature and pH were evaluated for the PLE-catalyzed hydrolysis of diester 15 in a biphasic system using ethanol as a cosolvent. It was observed that the enantiomeric excess of desired monoester 16 was increased by lowering the temperature from 25 to 10°C. The optimum pH for asymmetric hydrolysis of diester 15 in a biphasic system using ethanol as a cosolvent was 7.2 at 10°C. A semipreparative-scale asymmetric hydrolysis of diester 15 was carried out in a biphasic system using 10% ethanol as a cosolvent. Substrate (3 g) was used in a 300-ml reaction mixture. The reaction was carried out at 10°C, with 125-rpm agitation, and at a pH of 7.2 for 11 hours. A reaction yield of 96 mol% and an enantiomeric excess of 96.9% were obtained. From the reaction mixture, 2.6 g of monoester 16 were isolated in an 86.3 mol% overall yield. The enantiomeric excess of isolated S-(-)-monoester 16 was 96.9%. 1H NMR and MS of isolated product were consistent with monoester 13, and the specific rotation of monoester [~]D was --14.4 (c = 1.1 in methanol).

IV. Anticholesterol Drugs Pravastatin 17 and mevastatin 18 are anticholesterol drugs that act by competitively inhibiting HMG-CoA reductase (Endo et al., 1976a). Pravastatin sodium is produced in two fermentation steps. The first step is production of compound ML-236B by Penicillium citrinum (Endo et al., 1976b; Hosobuchi eta]., 1993a, 1993b). Purified compound is converted to its sodium salt 19 with sodium hydroxide and in the second step is hydroxylated to pravastatin sodium 17 (Fig. 16) by Streptomyces carbophilus (Serizawa et al., 1983). A cytochrome P450-containing enzyme system has been demonstrated from the S. carbophilus that catalyzed the hydroxylation reaction (Matsuoka et al., 1989). Squalene synthase is the first pathway-specific enzyme in the biosynthesis of cholesterol and catalyzes head-to-head condensation of two molecules of farnesyl pyrophosphate (FPP) to form squalene 20. It has been implicated in the transformation of FPP into presqualene pyro-

58

RAMESHN. PATEL A

.,,,OH

NaOOC" "~.... HOy

NaOOC/~.,.,,~OH

1 if,,. H 11.. ~ C H 3 CH3:" \V "<'Sodium

0

Streptomyces carbo_nhilus

H3C" ""( "O

HO~/] I

Ho:SLJ

ML-236B

Pravastatin Sodium 17 °

H3C~ Mevastatin 18

O .~H "~H CH3(/~,~ CH3 ~/~/

F~G.16. Hydroxylationof sodium ML-236Bto pravastatinsodium 17 by Streptomyces

carbophilus. Structure of mevastatin 18.

phosphate (Valentijn et al., 1995). FPP analogues are a major class of inhibitors of squalene synthase (Biller et al., 1991a, 1991b). However, the members of this class of compounds lack specificity and are potential inhibitors of other FPP-consuming transferases such as geranylgeranyl pyrophosphate synthase. To increase enzyme specificity, analogues of FPP and other mechanism-based enzyme inhibitors have been synthesized (Steiger et al., 1992). BMS-188494 is a potent squalene synthase inhibitor that is effective as an anticholesterol drug (Biller and Majnin, 1995; Lawrence et al., 1991). (S)[1-(acetoxyl)-4-(3-phenyl)butyl]phosphonic acid diethyl ester 21 is a key chiral intermediate required for total chemical synthesis of BMS-188494. The stereoselective acetylation of racemic [1-(hydroxy)-4-(3-phenyl)butyl]phosphonic acid diethyl ester 22 (Fig. 17) was carried out using G e o t r i c h u m c a n d i d u m lipase in toluene as solvent and isopropenyl acetate as acyl donor (Patel

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

59

N

o

-i

-

E

O

~

~

l° o~

O

N

Oo

O

O

~8

60

RAMESH N. PATEL

et al., 1997a). A reaction yield of 38% (theoretical maximum = 50%) and an enantiomeric excess of 95% were obtained for chiral 21.

V. Deoxyspergualin Spergualin, an antitumor antibiotic, was discovered in the culture filtrate of bacterial strain BMG162-aF2, which is related to Bacillus laterosporus, and its structure was determined to be (-)-(15S)-1-amino10-guanidino-11,15-dihydroxy-4,9,12-triazanonadecane-10,13-dione (Takeuchi et al., 1981; Umezawa et al., 1981). Total synthesis was accomplished by acid-catalyzed condensation of 11-amino-1,1-dihydroxy-3,8diazaundecane-2-one with (S)-7-guanidino-3-hydroxy-heptanamide followed by separation of the 11-epimeric mixture (Kondo et al., 1981). The antibacterial and antitumor activity of the enantiomeric mixture of spergualin was about half that of natural spergualin (Iwasawa et al., 1982), indicating the importance of the configuration at C-11 for antitumor activity. Umeda et al. (1987) demonstrated the optical resolution of the key intermediate of 15-deoxy-spergualin 23, racemic N-(7guanidino-heptanoyl)-a-alkoxyglycine, by using an exopeptidase (serine carboxy peptidase) and racemic N-(7-guanidino-heptanoyl)-a-alkoxyglycyl-L-amino acid as the substrate. Carboxypeptidase from Penicillum janthinellum catalyzed hydrolysis of the peptide bond of racemic N-(7-guanidinoheptanoyl)-a-methoxyglycyl-L-phenylalanine to yield (-)-N-(7-guanidinoheptanoyl)-a-methoxyglycine. The authors deduced that the absolute configuration of the carbon at 11 (C-11) of the bioactive (-)-enantiomer, and thus that of the natural spergualin, is (S). The (-)-enantiomer of 15-deoxy-spergualin was active against mouse leukemic L1210, while the (+)-enantiomer was almost inactive (Umeda et al., 1987). We have demonstrated an alternate and more direct route, the lipasecatalyzed stereoselective acetylation of racemic 7-[N,1V-bis(benzyloxycarbonyl) N-(guanidinoheptanoyl)]-ct-hydroxy-glycine 24 to the corresponding S-(-)-acetate 25 and unreacted alcohol (+)-24 (Patel et al., 1997b). S-(-)-acetate 25 (Fig. 18) is a key intermediate for total chemical synthesis of (-)-15-deoxyspergualin 23, a related immunosuppressive agent and antitumor antibiotic (Umeda et al., 1987; Maeda et a]., 1993). The reaction was carried out in methyl ethyl ketone (MEK) using lipase from Pseudomonas spp. (lipase AK). Vinyl acetate was used as an acylating agent. A reaction yield of 48% (theoretical maximum = 50%) and an enantiomeric excess of 98% were obtained for S-(-)-acetate 25.

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

ZHN H

H A A A H LipaseAK ZHN~-N NA

H (~ o.

z.

Racemlc alcohol 24

(+)-:Alcohol-- __24

H 2 N C H N ~ H N.

61

v

v

v

S- (--) Acetate __25

* ~ , , , ~ H ~ N H 2

II

0

I

OH

n

15-Deoxyspergualin 23

FIG. 18. Enantioselective enzymatic acetylation of racemic alcohol 24 to corresponding S-(-)-acetate 25 and unreacted R-(+)-alcohol 24 by lipase from a Pseudomonas sp. (lipase AK). Structure of 15-deoxyspergualin (antitumor antibiotic and immunosuppressive agent).

Unreacted alcohol (+)-24 was obtained in 41% yield and 98.5% enantiomeric excess.

Vl. Antiviral

Agents

An essential step in the life cycle of the human immunodeficiency virus (HIV-1) is proteolytic processing of its precursor proteins. This processing is accomplished by HIV-1 protease, a virally encoded enzyme. Inhibition of HIV-1 protease arrests replication of HIV in vitro. Thus, HIVol protease is an attractive target for chemotherapeutic intervention. Barrish et al. (1994) reported the discovery of a new class of selective HIV protease inhibitors that incorporate a C2 symmetric aminodiol core as one of its key structural features. Members of this class, and particularly BMS-186318, display potent anti-HIV activity in cell culture. The stereoselective microbial reduction of (1S)[3-chloro-2-oxo-l-(phenylmethyl)propyl] carbamic acid 1,1-dimethyl-ethyl ester 26 to 27a has been demonstrated (Fig. 19). Chiral alcohol 27a is a key intermediate for total chemical synthesis of BMS-186318 (Patel et al., 1997c). About 100 microorganisms were screened for stereoselective reduction of 26 to 27a. The reaction yields, diastereomeric purity, and enantiomeric excess of 27a obtained with the best six cultures are shown in Table VIII. The reaction yield and diastereomeric selectivity were dependent on the microorganism used during reduction of 26. S t r e p t o m y ces n o d o s u s SC13149, Candida boidinii SC13821, Mortierella r a m a n -

62

RAMESH N. PATEL

BOC

I 26

BOCI~/

O

I 27a OH

_

27_.._b.b

OH

27..__~c

OH

OH 27_.dd O

OH

OH

BMS-186318 (Antiviral agent) FIG. 19. Stereoselective enzymatic reduction of (1S)[3-chloro-2-oxo-l(phenylmethyl)propyl]carbamic acid 1,1-dimethyl-ethyl ester 26 to c o r r e s p o n d i n g chiral alcohol 27a by Streptomyces nodosus SC13149. Structure of antiviral agent BMS-186318.

TABLE VIII STEREOSELECTIVEMICROBIALREDUCTIONOF KETONE 26

Microorganisms

Streptomyces nodosus SC13149 Pullularia pullulans SC13849 Candida boidinii SC13821 Nocardioides albus SC13910 Mortierella ramanniana SC13850 Caldariomyces fumigo SC13901

Reaction yield of 27a (%) 56 47 39 5 54 52

Diastereomeric purity of 27a (%) >99 81 >99 85 91 93

EE of 27a (%) 99.9 99.9 99.9 99.9 99.9 99.9

SYNTHESIS OF CHIRALDRUG INTERMEDIATES

63

niana SC13850, and Caldariomyces fumago SC13901 gave >39% reaction yields, >91% diastereomeric purities, and 99.9% enantiomeric excess of product 27a. Further research was conducted using S. nodosus SC13149 and M. ramanniana SC13850 to convert ketone 26 to corresponding chiral alcohol 27a. Cells of S. nodosus SC13149 and M. ramanniana SC13850 were grown in a 25-liter fermentor for 48 hours. Cells were collected and suspended in 100-mM potassium phosphate buffer, pH 6.8, and the resulting cell suspensions were used to carry out the two-stage process for biotransformation of 26. After 24 hours a 67% reaction yield, 99% enantiomeric excess, and >99% diastereomeric purity were obtained for chiral alcohol 27a using cells of S. nodosus SC13149. M. ramanniana SC13850 gave a reaction yield of 54%, enantiomeric excess of 99.9%, and diastereomeric purity of 90% for chiral alcohol 27a. A single-stage fermentation-biotransformation process was developed for conversion of ketone 26 to chiral alcohol 27a with cells of S. nodosus SC13149, with a reaction yield of 80%, diasteromeric purity of >99%, and enantiomeric excess of 99.8%. From a 12-liter reaction mixture, 6.5 g of chiral alcohol 27a was isolated as white needle crystals (overall yield = 62%). The diastereomeric purity and enantiomeric excess of the isolated chiral alcohol were >99 and >99.8%, respectively.

VII. Stereoselective Hydrolysis of Racemic Epoxide Epoxide hydrolase catalyzes stereoselective hydrolysis of racemic epoxide to the corresponding chiral diol and nnreacted chiral epoxide. Furstoss and his coworkers used Aspergillus niger and Beauveria su]furescens for enantiospecific hydrolysis of epoxides, including many substituted styrene epoxides (Pedragosa-Moreau et al., 1993, 1994, 1996, 1997; Nellaiah, 1996). Faber and colleagues utilized epoxide hydrolases from Rhodococcus, Nocardia, and other species for enantiospecific hydrolysis (Hechtberger et al., 1993; Wandel eta]., 1995; Faber et a]., 1996; Kroutil et al., 1996, 1997a, 1997b). Enantioselective epoxide hydrolases from various fungal and other sources have been reported (Zhang et al., 1995; Grogan et al., 1997). Weijers (1997) found the yeast Rhodotorula glutinis to be effective for enantioselective hydrolysis of various epoxides. S-epoxide 28 is a key intermediate for a new prospective circadian modulator drug (Catt et al., 1998, 1999). Stereospecific hydrolysis of racemic epoxide RS-1-{2' (3'-dihydro benzo[b]furan-4'-yll-l,2-oxirane,

64

RAMESH N. PATEL

29) to corresponding R-diol 30 and unreacted chiral S-epoxide 28 (Fig. 20) was demonstrated by Goswami et al. (1999a). Epoxides are unstable under acidic and basic conditions. It is important to select an appropriate condition so as to minimize chemical hydrolysis of the epoxide during the biotransformation process. The stability of racemic epoxide 28 was determined under various conditions. About 100 and 91% of epoxide were hydrolyzed in 24 hours at pH 5 and 6, respectively. Even under neutral conditions (pH 7), the epoxide was not very stable, and 51% was hydrolyzed within 24 hours. Alkaline conditions (pH > 7) are better, and there was less hydrolysis (e.g., 38 and 30% in 24 hours at pH 8 and 9, respectively). Therefore, pH 8.0 was selected for conducting enzymatic hydrolysis. Even at pH 8, 19% of racemic epoxide 28 was hydrolyzed in 4 hours. Therefore, it was necessary to find a microorganism that hydrolyzes the racemic epoxide with high stereospecificity at a faster rate to prevent (or at least minimize) loss of unreacted desired S-epoxide 28 by chemical hydrolysis. Several fungi, yeast, and bacterial cultures were screened for stereospecific hydrolysis of the racemic epoxide. Two A. niger strains (SC16310, SC16311) and Rhodotorula glutinis SC16293 selectively hydrolyzed the R-epoxide, leaving behind S-epoxide 28. The enantiomer ratio (E) values (Chen et aL, 1982) for these microorganisms were -25. Unreacted S-epoxide 28 was obtained in >95% enantiomeric excess and at a 45% yield (theoretical maximum = 50%). Rhodococcus equi SC15835 did not hydrolyze the epoxide. Nocardia salmonicolor SC6310 hydrolyzed the racemic epoxide at a slow rate, and the enantiomeric excess of the S-epoxide was only 30%.

~spRhOdotorula glutinis?--~ ergillus~ger ~~V + 29

2~

OH-"OH ao

FIG. 20. Enantioselective hydrolysis of racemic epoxide 29 to corresponding (R)-diol 30 and unreacted (S)-epoxide 28.

65

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

From the initial screening studies, R. glutinis SC16293 and two A. niger strains, SC16310 and SC16311, were selected for further research. Hydrolysis of racemic epoxide by R. glutinis SC16293 was carried out. The desired S-epoxide 28 was obtained in 40% yield and >95% enantiomeric excess when the substrate was used at 2 g/liter and cells were used at 100 g/liter concentrations. Several solvents at 10% (vol/vol) were evaluated in an aqueous reaction mixture to improve the enantiomeric excess and yield (Table IX). Two solvents--cyclohexane and 1,1,2-trichloro-trifluoroethane (where the epoxide was not very soluble)--were used in higher amounts. Solvents had significant effects on both the extent of hydrolysis and the enantiomeric excess of unreacted S-epoxide 28. Most solvents, except for methyl tert-butyl ether (MTBE), gave lower enantiomeric excess than that of reactions catalyzed in buffer without any solvent supplement. The extent of hydrolysis in the presence of solvents was always lower than that in buffer. MTBE gave excellent results. A reaction yield of 45% (theoretical maximum = 50%) and an enantiomeric excess of 98.9% were obtained for unreacted S-epoxide 28. The hydrolysis reaction in the presence of MTBE gave an E value of 68. Two A. niger strains--SC16310 and SC16311--were evaluated in terms of their potential for stereospecific hydrolysis of the racemic epoxide. Both strains gave an enantiomeric excess of 97% and a yield of 45% of the remaining S-epoxide 28 when substrate was used at a 2

TABLE IX ENANTIOSELECTWEHYDROLYSISOF RACEMICEPOXIDE 2~ BY RHODOTORULA GLUTINIS SC16293 IN A BIPHASIC SYSTEM

Solvent

Reaction time (hours)

Remaining epoxide (%)

EE of (S)-epoxide 28 (%)

E value

Buffer

7

37

96.6

Cyclohexane

5

53

45.5

14 5

Toluene

5

66

45.9

29

1,1,2-trichlorotrifluoroethane

5

76

31.5

511

M e t h y l tert-butyl e t h e r (MTBE)

5

45

98.9

68

Methyl isobutyl ketone

5

68

22.7

4

n-Butanol

5

81

3.1

1

Dimethylsulfoxide

5

46

83.5

14

Dimethyl formamide

5

43

80

10

66

RAMESHN. PATEL

g/liter concentration. At a higher substrate concentration (5 g/liter) using a 100 g/liter cell concentration, a reaction yield of 51% and enantiomeric excess of 84% were obtained with SC16311.

VIII. Biocatalytic Dynamic Resolution: Stereoinversion of Racemic Diol One of the most often-used techniques for development of chiral compounds involves biocatalytic resolution. Though these kinetic resolution processes often provide compounds with high enantiomeric excess, the maximum theoretical yield of product or substrate is only 50%. In many cases, since the reaction mixture contains a 50:50 mixture of reactant and product with only slight difference in properties (e.g., hydrophobic alcohol and its acetate), separation becomes very difficult and impractical. These problems of kinetic resolution can be solved by employing a "dynamic resolution" process. The dynamic resolution process for alcohol is essentially a stereoinversion process. Only one enantiomer of the alcohol is enantiospecifically oxidized to the ketone, while the other enantiomer of the alcohol remains unchanged. The ketone is not isolated but is reduced to the opposite enantiomer of the alcohol during the process. The net result is conversion of the racemic alcohol to one enantiomer of the alcohol in high (theoretical maximum = 100%) yield. Dynamic resolution thus overcomes the limitation on maximum theoretical yield (50%) encountered during kinetic resolution of alcohol with enzymes. Only a handful of reports have appeared in the more recent literature on dynamic resolution of alcohols (Buisson et al., 1992; Nakamura et al., 1995; Fantin et al., 1995; Takahashi et al., 1995; Shimizu et al., 1987a, 1987b; Hasegawa et al., 1990; Stecher and Faber, 1997). Geotrichum c a n d i d u m , Candida parapsilosis, and a few other species are reported to be effective in such processes. Dynamic resolution involving a biocatalyst and metal-catalyzed in-situ racemizations has also been reported with limited success (Allen and Williams, 1996; Dinh et al., 1996). Chiral S-diol 31 (S-l-{2',3'-dihydrobenzo[b]furan-4'-yl)-ethane-l,2diol) is a key intermediate for a new prospective circadian modulator drug candidate (Catt et al., 1998, 1999). Dynamic resolution of racemic diol RS-l-{2',3"-dihydrobenzo[b]furan-4'-yll-ethane-l,2-diol32 to S-diol S-l-{2',3"-dihydrobenzo[b]furan-4'-yll-ethane-l,2-diol 31 (Fig. 21) was demonstrated by Goswami et al. (1999b).

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

~

OH RS-Diol OH 32

O

67

H OH S-Diol 31

T Reduction _ Oxidation OH

OH 0

R-Diol 34

Ketone 33

FIG. 21. Biocatalytic dynamic resolution. Stereoinversion ofracemic diol 32 to (S)-diol 31 by Candida boidinii and Pichia methanolica.

Seven cultures were selected from the screening of 20 microorganisms as leading candidates for dynamic resolution. These were Candida boidinii SC13821, SC13822, SC16115, Pichia methanolica SC13825, SC13860, and Hansenula polymorpha SC13895, SC13896. The relative proportions of S-diol 31 increased with time in biotransformations with the above cultures. At the end of 1 week, the enantiomeric excess of the remaining S-diol 31 was found to be in the range 87-100% with these microorganisms. Only two microorganisms, Candida parapsilosis SC16346 and Arthrobacter simplex SC6379, gave a higher yield of R-diol. A new c o m p o u n d was formed during these biotransformations, as seen by the appearance of a new peak in the HPLC of reaction mixture. This c o m p o u n d was slightly less polar than the diol. The identity of this compound was established as hydroxy ketone 33 from an LC-MS peak at mass 178. The starting RS-diol showed a mass peak at 180 by LC-MS. The area of the HPLC peak for hydroxy ketone 33 at first increased with time, reached a maximum, and then decreased. This w o u l d be expected from the proposed pathway of dynamic resolution (Fig. 21). Hydroxy ketone 33 was first formed by oxidation of R-diol 34, and then subsequently reduced back to diol, but only to S-diol 31.

68

RAMESH N. PATEL

The quantity and enantiomeric excess of the diol at various times were followed very carefully for transformation of RS-diol by the seven microorganisms described above. The reactions were also conducted with and without glucose to investigate the effect of glucose on the course of biotransformation (Table X). C. boidinii SC13822, C. boidinii SC16115, and P. methanolica SC13860 transformed RS-diol 32 in 3-4 days, and S-diol 31 was obtained with a yield in the range 62-71% and enantiomeric excess in the range 90-100%.

IX. Resolution of Racemic Secondary Alcohols The current interest in enzymatic production of chiral compounds lies in preparation of intermediates for pharmaceutical synthesis. S-(+)-2-pentanol is a key chiral intermediate required for synthesis of anti-Alzheimer's drugs that inhibit [3-amyloid peptide release and/or synthesis (Audia et al., 1996; Hamilton et al., 1996). The enzymatic

TABLE X DYNAMICRESOLUTION: STEREOINVERSIONOF RACEMICDIOL 32 TO CHIRAL (S)-DIOL 31

Medium

Reaction time (days)

Remaining diol

(S)-diol

(%)

31 (%)

EE of

Microorganism

Strain

Candida boidiniii

13821

Buffer Buffer + glucose

4 4

70 74

87 54

Candida boidinii

13822

Buffer Buffer+ glucose

4 4

66 62

90 100

Candida boidinii

16115

Buffer Buffer + glucose

3 4 3

74 64 71

95 100 94

Pichia methanolica

13825

Buffer Buffer + glucose

4 4

83 72

63 87

Pichia methanolica

13860

Buffer

3 4 2 3

65 46 57 67

100 100 89 100

Buffer + glucose

Hansenula polymorpha

13895

Buffer Buffer + glucose

4 3

84 100

44 32

Hansenula polymorpha

13896

Buffer Buffer + glucose

4 3

73 74

60 52

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

69

resolution of racemic 2-pentanol and 2-heptanol by lipase B from Candida antarctica has been demonstrated by Patel et al. (1999b). Commercially available lipases were screened for stereoselective acetylation of racemic 2-pentanol in an organic solvent (hexane) in the presence of vinyl acetate as an acyl donor. C. antarctica lipase B efficiently catalyzed enantioselective acetylation of racemic 2-pentanol. Reaction yields of 49% (theoretical maximum = 50%) and an enantiomeric excess of 99% were obtained for S-(+)-2-pentanol. Preparativescale acetylation of racemic 2-pentanol was carried out in an organic solvent (heptane) in the presence of vinyl acetate as an acyl donor using lipase B (Table XI). At the end of the reaction, 44.5 g of S-(+)-2-pentanol were estimated by HPLC analysis, with an enantiomeric excess of 98%. Among acylating agents tested, succinic anhydride was found to be the best choice due to easy recovery of (S)-2-pentanol at reaction end. Reactions were carried out using racemic 2-pentanol as solvent as well as substrate. Using 0.68 mole equivalent of succinic anhydride (Fig. 22) and 13 g of lipase B per kilogram of racemic 2-pentanol, a reaction yield of 43 mol% (theoretical maximum = 50%) and an enantiomeric excess of >98% were obtained for (S)°2-pentanol. Product was isolated in overall 36% yield (theoretical maximum = 50%). The results from three preparative batches are shown in Table XII. As described earlier, resolution of 2-heptanol was also carried out using lipase B. Reactions were carried out using racemic 2-heptanol as solvent as well as substrate. Using 0.68 mole equivalent of succinic anhydride and 13 g of lipase B

TABLE XI PREPARATIVESCALE ENZYMATICACETYLATIONOF RACEMIC 2-PENTANOL USING LIPASE B FROM CANDIDAANTARCTICA Reaction time (hours)

(S)-2-pentanol (g/liter)

(R)-2-pentanol (g/liter)

0 2 4 6

50 50 46 44.5

50 20 10 0.02

EE of (S)-2-pentanol (%) 0 32 60 98

Reaction m i x t u r e in 1 liter of h e p t a n e c o n t a i n i n g 100 g of r a c e m i c 2-pentanol, 1.02 m o l e e q u i v a l e n t of v i n y l acetate, a n d 1 g of lipase B from C. antarctica. T h e reaction w a s carried o u t at 35°C a n d 150 r p m .

70

RAMESH N. PATEL

OH

Racemic 2-pentanol

OH

Lipase B from C. antarctica

S-(+)-2-pentanol

o

°

R-(-)-2-pentylhemisuccinate

FIc. 22. Enzymatic resolution of racemic 2-pentanol to S-(+)-2-pentanol by Candida antarctica lipase.

TABLE XII ENZYMATICACYLATIONOF SECONDARYALCOHOLSUSINGSUCCINICANHYDRIDE ANDLIPASEB FROMCANDIDAANTARCTICA

Batch number 2-pentanoh 132 133 136

Batch number 2-heptanoh 140 141

2-pentanol input (kg)

(S)-2-pentanol (% yield)

EE of (S)-2pentanol (%)

0.5 0.5 0.9

42.0 43.5 43.7

>99 >98 99

2-heptanol input (kg)

0.1 0.5

(S)-2-heptanol (% yield)

43.0 44.5

EE of (S)-2heptanol (%)

>99 >99

Reaction mixture contained racemic 2-pentanol or 2-heptanol as solvent as well as substrate; 0.68 mole equivalent of succinic anhydride and 13 g of lipase B per kg of substrate input. The reaction was carried out at 38°C and 150 rpm.

per kilogram of racemic 2-heptanol, a reaction yield of 44 mol% (theoretical maximum = 50%) and an enantiomeric excess of >99% were o b t a i n e d f o r S - ( + ) - 2 - h e p t a n o l ( T a b l e XII). P r o d u c t w a s i s o l a t e d i n a n overall 40% yield (theoretical maximum = 50%).

SYNTHESIS OF CHIRAL DRUG INTERMEDIATES

71

X. Summary Biocatalytic processes were used to prepare chira] intermediates for pharmaceuticals. These include the following processes. Enzymatic synthesis of [4S-(4a,7a,10ab)]l-octahydro-5-oxo-4-[[(phenylmethoxy) carbonyl]amino]-7H-pyrido-[2,1-b] [1,3]thiazepine-7-carboxylic acid methyl ester (BMS-199541-01), a key chiral intermediate for synthesis of a new vasopeptidase inhibitor. Enzymatic oxidation of the e-amino group of lysine in dipeptide dimer N2-[N[[(phenylmethoxy)carbonyl] L-homocysteinyl] L-lysine)l,l-disulfide (BMS-201391-01) to produce BMS-199541-01 using a novel L-lysine ~-aminotransferase from S. paucimobilis SC16113 was demonstrated. This enzyme was overexpressed in E. coli, and a process was developed using recombinant enzyme. The aminotransferase reaction required ~-ketoglutarate as the amine acceptor. Glutamate formed during this reaction was recycled back to ~-ketoglutarate by glutamate oxidase from S. noursei SC6007. Synthesis and enzymatic conversion of 2-keto-6-hydroxyhexanoic acid 5 to L-6-hydroxy norleucine 4 was demonstrated by reductive amination using beef liver glutamate dehydrogenase. To avoid the lengthy chemical synthesis of ketoacid 5, a second route was developed to prepare the ketoacid by treatment of racemic 6-hydroxy norleucine (readily available from hydrolysis of 5-(4-hydroxybutyl) hydantoin, 6) with D-amino acid oxidase from porcine kidney or T. variabilis followed by reductive amination to convert the mixture to L-6-hydroxynorleucine in 98% yield and 99% enantiomeric excess. Enzymatic synthesis of (S)-2-amino-5-(1,3-dioxolan-2-yl)-pentanoic acid (allysine ethylene acetal, 7), one of three building blocks used for synthesis of a vasopeptidase inhibitor, was demonstrated using phenylalanine dehydrogenase from T. intermedius. The reaction requires ammonia and NADH. NAD produced during the reaction was recycled to NADH by oxidation of formate to CO2 using formate dehydrogenase. Efficient synthesis of chiral intermediates required for total chemical synthesis of a ~3 receptor agonist was demonstrated. These include: (a) microbial reduction of 4-benzyloxy-3-methanesulfonylamino-2"-bromoacetophenone 9 to corresponding (R)-alcohol 10 by S. paucimobilis SC16113, (b) enzymatic resolution of racemic m-methyl phenylalanine amide 11 and ~-methyl-4-hydroxyphenylalanine amide 13 by amidase from M. neoaurum ATCC 25795 to prepare corresponding (S)-amino acids 12 and 14, and (c) asymmetric hydrolysis of methyl-(4methoxyphenyl)-propanedioic acid ethyl diester 15 to corresponding (S)-monoester 16 by pig liver esterase.

72

RAMESH N. PATEL

(S)[1-(acetoxyl)-4-(3-phenyl)butyl]phosphonic acid diethyl ester 21, a key chiral intermediate required for total chemical synthesis of BMS188494 (an anticholesterol drug) was prepared by stereoselective acetylation of racemic [1-(hydroxy)-4-(3-phenyl)butyl]phosphonic acid diethyl ester 22 using G. candidum lipase. Lipase-catalyzed stereoselective acetylation of racemic 7-[N,N'-bis(benzyloxy-carbonyl)N-(guanidinoheptanoyl)]-~-hydroxy-glycine 24 to corresponding S-(-)-acetate 25 was demonstrated. S-(-)-acetate 25 is a key intermediate for total chemical synthesis of (-)-15-deoxyspergualin 23, an immunosuppressive agent and antitumor antibiotic. Stereoselective microbial reduction of (1S)[3-chloro-2-oxo-l-(phenylmethyl)propyl] carbamic acid, 1,1-dimethyl-ethyl ester 26 to corresponding chiral alcohol 27a (a key chiral intermediate for HIV protease inhibitors) was also demonstrated. Stereospecific enzymatic hydrolysis of racemic epoxide RS-I-{2',3'dihydro benzo[b]furan-4'-yl}-l,2-oxirane 29 the corresponding R-diol 30 and unreacted chiral S-epoxide 28 was demonstrated using R. glutinis and A. niger. Dynamic resolution of racemic diol RS-l-{2',3'-dihydrobenzo[b]furan-4'-yll-ethane-l,2-diol 32 to corresponding S-diol S-l-{2',3'-dihydrobenzo[b]furan-4'-yll-ethane-l,2-diol 31 was demonstrated using C. boidinii and P. methanolica. Chiral (S)-epoxide 28 and (S)-diol 31 are key intermediates for a new prospective circadian modulator drug. Enzymatic resolution of racemic 2-pentanol and 2-heptanol by lipase B from Candida antarctica was demonstrated. S-(+)-2-pentanol is a key chiral intermediate required for synthesis of anti-Alzheimer's drugs.

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Maurer, P. J., and Miller, M. J. (1981). Mycobactins: synthesis of (-)-cobaltin T from ~-hydroxynorleucine. J. Org. Chem. 46, 2835-2836. Maurer, P. J., and Miller, M. J. (1982). Microbial iron chelators: Total synthesis of aerobactin and its constituent amino acid, N6-acetyl-N6-hydroxylysine. J. Am. Chem. Soc. 104, 3096-3101. Maurer, P. J., and Miller, M. J. (1983). Total synthesis of a mycobactin: Mycobactin $2. J. Am. Chem. Soc. 105, 240-245. Mori, K. (1995). Biochemical methods in enantioselective synthesis of bioactive natural products. SYNLETT 11, 1097-1109. Nakamura, K., Inoue, Y., Matsuda, T., and Ohno, A. (1995). Microbial deracemization of 1-arylethanol. Tetrahedron Lett. 36, 6263-6266. Nellaiah, H., Morisseau, C., Archelas, A., Furstoss, R., and Baratti, J. (1996). Enantioselective hydrolysis of p-nitrostyrene oxide by an epoxide hydrolase preparation from Aspergillus niger. Biotechnol. Bioeng. 49, 70-77. Ohshima, T., Sugimoto, H., and Soda, K. (1988). Selective enzymic determination of L-phenylalanine and phenylpyruvate. Anal. Lett. 21, 2205-2215. Ohshima, T., Takada, H., Yoshimura, T., Esaki, N., and Soda, K. (1991). Distribution, purification, and characterization of thermostable phenylalanine dehydrogenase from thermophilic actinomycetes. J. Bacteriol. 173, 3943-3948. Patel, R. N. (1997). Stereoselective biotransformations in synthesis of some chiral pharmaceutical intermediates. Adv. Appl. Microbiol. 43, 91-140. Patel, R. N. (1998). Tour de paclitaxel: Biocatalysis for semisynthesis. Ann. Rev. Microbiol. 98, 361-395. Patel, R. N. (1999). Biocatalysis for synthesis of chiral drug intermediates. In "Stereoselective Biocatalysis" (R. Patel, ed.), pp 87-130. Marcel, New York. Patel, R. N., Banerjee, A., and Szarka, L. J. (1997a). Stereoselective acetylation of [1-(hydroxy)-4-(3-phenoxyphenyl)butyl]phosphonic acid diethyl ester. Tetrahedron: Asymmetry 8, 1055-1059. Patel, R. N., Banerjee, A., and Szarka, L. J. (1997b). Stereoselective acetylation of racemic 7-[N,N'-bis-benzyloxycarbonyl)-N-(guanidinoheptanoyl)]-a-hydroxyglycine.Tetrahedron Asymmetry 8, 1767-1771. Patel, R. N., Banerjee, A., McNamee C., Brzozowski, D., and Szarka, L. J. (1997c). Preparation of chiral synthon for HIV protease inhibitor: Stereoselective microbial reduction of N-protected a-aminochloroketone. Tetrahedron: Asymmetry 8, 2547-2552. Patel, R. N., Banerjee, A., Chu, L., Brzozowski, D., Nanduri, V., and Szarka, L. J. (1998). Microbial synthesis of chiral intermediates for [~3-receptor agonists. J. Am. Oil Chem. Soc. 75, 1473-1482. Patel, R. N., Banerjee, A., Nanduri, V., Goldberg, S., Johnston, R., Hanson, R., McNamee, C., Brzozowski, D., Tully, T., Ko, R., LaPorte, T., Cazzulino, D., Swaminathan, S., Parker, L., and Venit, J. (1999a). Biocatalytic preparation of a chiral synthon for a vasopeptidase inhibitor: Enzymatic conversion of N2-{N-[(phenylmethoxy)carbonyl]L-homocysteinyl]-L-lysine (11')-disulfide to [4S-(4a,7a,10ab)]l-octahydro-5•x••4•[(pheny•meth•xy)carb•ny•]amin•].7H.pyrid••[2•1•b][1•3]thiazepin•7•carb•xy• ic acid methyl ester by a novel L-lysine e-aminotransferase. Enzyme Microb. Technol. In press. Patel, R. N., Banerjee, A., Hanson, R. L, Brzozowski, D. B., Parker, L. W., and Szarka, L. J. (1999b). Oxidation of N-a-protected-L-lysine by Rhodotorula graminis to produce novel chiral compounds. Tetrahedron: Asymmetry 10, 31-36. Patel, R. N., Banerjee, A., Nanduri, V., and Comezoglu, F. T. (1999c). Enzymatic resolution of secondary alcohols. Biocatal. Biotransform. Manuscript submitted.

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Pedragosa-Moreau, S., Archelas, A. R., and Furstoss, R. (1993). Microbiological transformations, 28: Enantioselective epoxide hydrolyses as a preparative access to both enantiomers of styrene oxide. J. Org. Chem. 58, 5533-5536. Pedragosa-Moreau, S., Archelas, A., and Furstoss, R. (1994). Microbiological transformations, 29: Enantioselective hydrolysis of epoxides use microorganisms: A mechanistic study. Biorg. Med. Chem. 2,609-616. Pedragosa-Moreau, S., Morisseau, C., Zylber, J., Archelas, A , Baratti, J., and Furstoss, R. (1996). Microbiological transformations, 33: Fungal epoxide hydrolases applied to the synthesis of enantiopure para-substituted styrene oxides. A mechanistic approach. J. Org. Chem. 61, 7402-7407. Pedragosa-Moreau, S., Morisseau, C., Baratti, J., Zylber, J., Archelas, A., and Furstoss, R. (1997). Microbiological transformations, 37: An enantioconvergent synthesis of the [~-blocker Nifenalol® using a combined chemoenzymatic approach. Tetrahedron 53, 9707-9714. Robl, J. A., and Cimarusti, M. P. (1994). A synthetic route for the generation of C-7 substituted azepinones. Tetrahedron Lett. 35, 1393-1396. Robl, J. A., Sun, C., Stevenson, J., Ryono, D. E., Simpkins, L. M., Cimarusti, M. A., Dejneka, T., Slusarchyk, W. A., Chao, S., Stratton, L., Misra, R. N., Bednarz, M. S., Asaad, M. M., Cheung, H. S., Aboa-Offei, B. E., Smith, P. L., Mathers, P. D., Fox, M., Schaeffer, T. R., Seymour, A. A., and Trippodo, N. C. (1997). Dual metalloprotease inhibitors: Mercaptoacetyl-based fused heterocyclic dipeptide mimetics as inhibitors of angiotensin-converting enzyme and neutral endopeptidase. J. Med. Chem. 40, 1570-1577. Rumbero, A., Martin, J. C., Lumbreras, M. A., Liras, P., and Esmahan, C. (1995). Chemical synthesis of allysine ethylene acetal and conversion in situ into 1-piperidine-6-carboxylic acid: Key intermediate of the a-aminoadipic acid for [3-1actam antibiotic biosynthesis. Bioorg. Med. Chem. 3, 1237-1240. Santaneillo, E., Ferraboschi, P., Grisenti, P., and Manzocchi, A. (1992). The biocatalytic approach to the preparation of enantiomerically pure chiral building blocks. Chem. Rev. 92, 1071-1140. Schfitte, H. Flossdorf, J. Sahm, H., and Kula, M.-R. (1976). Purification and properties of formaldehyde dehydrogenase and formate dehydrogenase from Candida boidinii. Eur. J. Biochem., 62, 151-160. Serizawa, N., Serizawa, S., Nakagawa, K., Furuya, K., Okazaki, T., and Tarahara, A. (1983). Microbial hydroxylation of ML-236B (compactin): Studies on microorganisms capable of 3~-hydroxylation of M1-236B. J. Antibiot. 36, 887-891. Shaked, Z., and Whitesides, G. M. (1980). Enzyme-catalyzed organic synthesis: NADH regeneration by using formate dehydrogenase. J. Am. Chem. Soc. 102, 7104-7105. Sher, P. M. (1994). "Preparation of 5-Substituted Benzo-l,3-Dioxo]es as [33 Adrenergic Receptor Agonists." U.S. Patent Application 5321036, 6o14-94. Shimizu, S., Hattori, S., Hata, H., and Yamada, H. (1987a). Stereoselective enzymic oxidation and reduction system for the production of D-(-)-pantoyl lactone from a racemic mixture of pantoyl lactone. Enzyme Microb. TechnoI. 9, 411-416. Shimizu, S., Hattori, S., Hata, H., and Yamada, H. (1987b). One-steo microbial conversion of a racemic mixture of pantoyl lactone to optically active D-(-)-pantoyl lactone. AppI. Environ. Microbiol. 53,519-522. Sih, C. J., and Chen, C. S. (1984). Microbial asymmetric synthesis: Enantioselective reduction of ketones. Angew. Chem. 96, 556-566. Sih, C. J., Gu, Q.-M., Holdgrun, X., and Harris, K. (1992). Optically active compounds via biocatalytic methods. Chirality 4, 91-97.

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Soda, K., and Misono, H. (1968). L-lysine ~-ketoglutarate aminotransferase, II: Purification, crystallization, and properties. Biochemistry 7, 4110-4119. Soda, K., Misono, H., and Yamamoto, T. (1968). L-lysine c~-ketoglutarate aminotransferase, h Identification of product, 1-piperidine-6-carboxylic acid. Biochemistry 7, 41024109. Stecher, H., and Faber, K. (1997). Biocatalytic deracemization techniques: Dynamic resolutions and stereoinversion. Synthesis 1, 1-16. Steiger, A., Pyun, H. J., and Coates, R. M. (1992). Synthesis and characterization of aza analog inhibitors of squalene and geranylgeranyl diphosphate synthesis. J. Org. Chem. 57, 3444-3450. Stirling D. I. (1992). The use of aminotransferases for the production of chiral amino acids and amines. In "Chirality in Industry" (A. N. Collins, G. N. Sheldrake, and J. Crosby, eds.), pp 209-222. Wiley, New York. Takada, H., Yoshimura, T., Ohshima, T., Esaki, N., and Soda, K. (1991). Thermostable phenylalanine dehydrogenase of Thermoactinomyces intermedius: Cloning, expression, and sequencing of its gene. J. Biochem., 109, 371-376. Takahashi, E., Nakamichi, K., and Furui, M. (1995). R-(-)-mandelic acid production from racemic mandelic acids using Pseudomonas polycolor IFO 3918 and Micrococcus freudenreichi Ferm-P 13221. J. Ferment. Bioeng. 80, 247-250. Takeuchi, T., Iinuma, H., Kunimoto, S., Masuda, T., Ishizuka, M., Hamada, M., Naganawa, H., Kondo, S., and Umezawa, H. (1981). A new antitumor antibiotic, spergualin: Isolation and antitnmor activity. J. Antibiot. 34, 1619-1621. Umezawa, H., Kondo, S., Iinuma, H., Kunimoto, Y., Iwasawa, H., Ikeda, D., and Takeuchi, T. (1981). Structure of an antitumor antibiotic, spergualin. J. Antibiot. 34, 1622-1624. Umeda Y., Moriguchi, M., Katsushige, I., Kuroda, H., Nakamnra, T., Fujii, A., Takeuchi, T., and Humezawa, H. (1987). Synthesis and antitumor activity of spergualin analogues, III: Novel method for synthesis of optically active 15-deoxyspergualin and 15-deoxy-11-O-methylspergualin. J. Antibiot. 40, 1316-1324. Valentijn, A. R. P. M., de Harm, R., de Kant, E., van der Marel, G. A., Cohen, L. H., van Boom, J. H. (1995). Synthesis of a potential enzyme-specific inhibitor of squalene synthase. Red. Trans. Chim. Pays-Bas 114, 332-336. Wandel, U., Mischitz, M., Kroutil, W., and Faber, K. (1995). High selective asymmetric hydrolysis of 2,2-disubstituted epoxides using lyophilized cells of Rhodococcus sp. NCIMB 11216. J. Chem. Soc., Perkin Trans. 1, 735-736. Weijers, C. A. G. M. (1997). Enantioselective hydrolysis of aryl, alicyclic and apliphatic epoxides by Rhodotorula glutinis. Tetrahedron: Asymmetry 8, 639-647. Wilson, C., Wilson, S., Piercy, V., Sennitt, M. V., and Arch, J. R. S. (1984). The rat lipolytic beta-adrenoceptor: studies using novel [~-adrenoceptor agonist. Eur. J. Pharmacol. 100, 309-319. Wong, C.-H., and Whitesides, G. M. (1994). "Enzymes in Synthetic Organic Chemistry." Tetrahedron Organic Chemistry Series, VoL 12. Elsevier, New York. Zhang, J., Reddy, J., Roberge, C., Senanayake, C., Greasham, R., and Chartrain, M. (1995). Chiral bioresolution of racemic indene oxide by fungal epoxide hydrolases. J. Ferment. Bioeng. 80, 244-246.

Recent Developments in the Molecular Genetics of the Erythromycin-Producing Organism

Saccharopolyspora erythraea THOMAS

J.

VANDEN BOOM

Abbott Laboratories Fermentation Microbiology Research and Development North Chicago, Illinois 60064

I. II. III. IV.

V.

VI.

VII. VIII.

IX.

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Introduction Background Experimental Properties of S. erythraea Strains Characterization of the S. erythraea Genome A. Physical-Genetic Mapping of the Chromosome B. Genomic Polymorphisms in Industrially Improved S. erythraea Strains Introduction of DNA into S. erythraea A. Sonication-Dependent Electroporation B. Electroporation of Germinating Spores Transcriptional Organization and Regulation of the Erythromycin Biosynthetic Gene Cluster A. Previous Transcriptional Studies of the eryCI, ermE, and eryG Genes B. Construction and Analysis of Transcriptional Mutants in S. erythraea C. Erythromycin Biosynthetic Gene Cluster Promoters D. Transcriptional Overview of the ery Gene Cluster New Molecular Genetic Tools for Studying Gene Expression in S. erythraea Genetic-EngineeringApproaches to Industrial Strain Improvement A. Construction of High-Productivity Source Strains for Naturally Occurring Erythromycin Intermediates B. Two-Step Genetic-EngineeringApproaches for Optimization of Novel Macrolide Production in S. erythraea C. Introduction of the Vitreoscilla hemoglobin gene into S. erythraea Combinatorial Biosynthesis A. Manipulation of ery Biosynthetic Genes in Heterologous Streptomyces Hosts B. Manipulation of ery Biosynthetic Genes in S. erythraea Future Prospects References

I. Introduction N e a r l y 50 y e a r s s i n c e t h e m a c r o l i d e a n t i b i o t i c e r y t h r o m y c i n w a s first d e s c r i b e d ( M c G u i r e et al., 1952), t h e p r o d u c i n g m i c r o o r g a n i s m Saccharopolyspora erythraea r e m a i n s t h e s u b j e c t of k e e n i n d u s t r i a l i n t e r est. A n u m b e r of factors h a v e c o n t r i b u t e d to t h e o n g o i n g i n d u s t r i a l 79 ADVANCESINAPPLIEDMICROBIOLOGY.VOLUME47 Copyright©2000byAcademicPress Allrightsofreproductionin anyformreserved. 0065-2164/00$25.00

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THOMAS J. VANDENBOOM

research interest in both erythromycin and S. erythraea. Prominent among these is the fact that various dosage forms of erythromycin continue to enjoy widespread use globally for a variety of indications due to the excellent record of therapeutic efficacy and safety achieved by erythromycin-derived products. Moreover, the introduction of second-generation semisynthetic erythromycin derivatives in the early 1990s created additional demands for bulk erythromycin A as the starting raw material for these products. The two major commercial second-generation erythromycin species, clarithromycin and azithromycin, are shown in Figure 1. The emergence of clinical isolates resistant to the second-generation macrolide antibiotics (Weisblum, 1998) has fueled continuing research in a number of industrial laboratories to develop third-generation erythromycin derivatives (Ma et al., 1999; Agouridas et al., 1998; Phan et al., 1997). Perhaps the most promising class of third-generation candidates currently in clinical development is the 3-oxo-erythromycin derivatives, or "ketolides." Two leading clinical candidates in this class, ABT-773 and HMR-3647, are also shown in Figure 1. Finally, there appears to be growing interest in the genes involved in erythromycin biosynthesis in the emerging field of combinatorial biosynthesis. Both the type I polyketide synthase (PKS) from S. erythraea (for reviews, see Hutchinson, 1998, 1999; Cane et al., 1998) and the related desosamine deoxysugar biosynthesis genes from Streptomyces venezuelae (Zhao et aL, 1998) have been successfully manipulated to produce hybrid microbial metabolites. In this review, I discuss recent advances in the molecular genetics of S. erythraea, with particular emphasis on current topics of industrial interest. Our present knowledge of the S. erythraea genome, as well as recent advances in molecular genetic methods applicable to wild-type and industrially improved strains of this organism, are considered here. In addition, this review summarizes recent studies on the transcriptional organization and regulation of the erythromycin biosynthetic gene cluster. These studies have improved our understanding of erythromycin gene expression in this organism and provide a foundation for future genetic manipulations of this industrially significant metabolic pathway. Finally, I briefly consider genetic-engineering approaches to erythromycin strain improvement and the role of S. erythraea and erythromycin biosynthetic genes in the emerging field of combinatorial biosynthesis. S. erythraea has received considerable attention as a model system for the study of polyketide biosynthesis. This topic is beyond the scope of this review. A brief overview of the biosynthesis of the erythromycin polyketide backbone is included herein simply as background for this review. The interested reader is referred to several

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recent reviews that have appeared elsewhere for additional coverage of this topic (Staunton and Wilkinson, 1997; Katz, 1997; Khosla et al., 1999).

II. Background The Gram-positive actinomycete Saccharopolyspora erythraea produces the clinically significant macrolide antibiotic erythromycin A. The erythromycin biosynthetic gene cluster has been localized near one

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THOMAS J. VANDENBOOM

end of the linear S. erythraea chromosome (Reeves eta]., 1998). The gene cluster has been cloned and sequenced and contains at least 20 genes involved in the formation and modification of the 14-membered macrolide 6-deoxyerythronolide B (6-DEB) and in the synthesis, attachment, and modification of the two deoxysugars desosamine and mycarose (Salahbey et al., 1998; for reviews, see Staunton and Wilkinson, 1997; Katz, 1997). Functions for the majority of genes located in this cluster have been proposed based on an analysis of blocked mutants constructed through targeted gene inactivation. A schematic view of the erythromycin biosynthetic pathway through the first bioactive erythromycin intermediate erythromycin D is shown in Figure 2. During erythromycin biosynthesis, the aglycone backbone 6-DEB is produced by a type I modular PKS from one propionyl-CoA and six (2S)-methylmalonyl-CoA molecules in a process closely resembling fatty acid biosynthesis (Staunton and Wilkinson, 1997). The 6DEB synthase (DEBS) is encoded by three large genes, designated eryAI, eryAII, and eryAIII, located roughly in the center of the biosynthetic gene cluster. The three multifunctional enzymes encoded by these genes each contains two modules, or sets of enzymatic activities, responsible for a single round of polyketide chain extension. The catalytic activities present in these modules dictate the stereochemistry and extent of reduction during each round of chain extension. In addition, the specificity of the initial loading module dictates the preferred starter units used by the DEBS enzyme (Weissman et al., 1998b). Following synthesis of the 6-DEB polyketide backbone, a specific hydroxylation occurs at the C-6 position to produce erythronolide B (EB). The C-6 hydroxylase responsible for this reaction is encoded by the eryF gene (Weber et al., 1991). EB is then modified by sequential attachment of mycarose and desosamine at the C-3 and C-5 hydroxyl groups, respectively, to produce the first bioactive intermediate, erythromycin D. Mutations affecting the synthesis and attachment of mycarose define eryB genes and result in phenotypic accumulation of the aglycone EB, whereas mutations affecting the synthesis and attachment of desosamine define e ~ C genes and result in phenotypic accumulation of 3-a-mycarosyl erythronolide B. The terminal steps of the erythromycin biosynthetic pathway form a metabolic grid in which erythromycin D is converted to erythromycin A by two alternative pathways (Fig. 3). Two modification enzymes, a specific mycarosyl O-methyltransferase encoded by the eryG gene (Paulus et al., 1990; Haydock et al., 1991) and a C-12 hydroxylase encoded by the eryK gene (Stassi et al., 1993), compete for the erythromycin D

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pathway intermediate. Initial O-methylation of the mycarose moiety on erythromycin D leads to the bioactive intermediate erythromycin B. Subsequent C-12 hydroxylation of erythromycin B leads to erythromycin A. Alternatively, these reactions can be reversed. Initial C-12 hydroxylation of erythromycin D leads to the bioactive intermediate erythromycin C, which in turn is converted to erythromycin A by the specific mycarosyl O-methyltransferase. The latter route has been suggested as the preferred pathway based on kinetic studies of the C-12 hydroxylase enzyme (Lambalot eta]., 1995). II1. Experimental Properties of S.

erythraea Strains

Wild-type strains of S. erythraea (e.g., NRRL2338) are readily manipulated using minor variations of the molecular genetic methods devel-

MOLECULAR GENETICS OF

Saccharopolyspora erythraea

85

oped for the better-characterized Streptomyces species S. coelicolor and S. lividans (Hopwood et al., 1985). Strain NRRL2338, and closely related S. erythraea strains (e.g., ER720), have been widely used in molecular genetic studies to elucidate gene-function relationships in the erythromycin biosynthetic pathway. In addition, molecular genetic manipulations of the erythromycin polyketide synthase in these genetic backgrounds has led to isolation of a number of novel macrolide compounds (see below) and have provided some insights into the function of individual enzymatic domains in this complex enzyme system (Katz, 1997). Several variants of wild-type strain NRRL2338 are in use within the S. erythraea research community, including the NRRL2338 "red variant strain" used by Leadlay and coworkers (Hessler et al., 1997). These variants may differ slightly in their experimental handling properties (e.g., growth rate) and erythromycin productivities. In contrast to the wild-type variants noted above, a wide range of mutant strains of S. erythraea developed for the large-scale fermentative production of erythromycin have been less amenable to established streptomycete and wild-type S. erythraea molecular genetic methods (Fitzgerald eta]., 1998; Katz, 1997; Brunker et a]., 1998). Significant differences in protoplast formation and regeneration, recombination, genome structure, plasmid maintenance, drug resistance, and genetic stability have been observed between wild-type and certain industrially improved strains of this organism (unpublished observations). Industrially improved strains (e.g., CA340) represent the product of numerous cycles of mutagenesis and screening for improved erythromycin titers (or other desirable fermentation properties). Practical experimental differences between wild-type and industrially improved strains likely result from secondary mutations present in heavily mutagenized improved genetic backgrounds or reflect the pleiotropic nature of certain titer-enhancing mutations. These strain differences, although poorly understood, have provided an incentive for further development of molecular genetic tools applicable to the full range of wild-type and improved S. erythraea strains encountered in industrial applications. IV. Characterization of the S. erythraea Genome

A.

PHYSICAL-GENETIC MAPPING OF THE CHROMOSOME

A physical map of AseI and DraI restriction sites in the chromosome of S. er~hraea strain NRRL2338 was completed using high-resolution PFGE (Reeves et al., 1998). Summation of individual AseI, DraI, and AseI-DraI fragments revealed a chromosome size of roughly 8 Mb. This

86

THOMAS J. VANDEN BOOM

genome size is comparable to several previously characterized Streptomyces chromosomes (Kieser et al., 1992; LeBlond et al., 1993; Lezhava eta]., 1995; Pandza et al., 1997). The S. erythraea chromosome also shares several other features in common with previously characterized Streptomyces chromosomes. These features include a linear topology, the localization of genes involved in secondary metabolism near one end of the chromosome, and evidence for large genetically unstable regions of DNA (LeBlond et al., 1996; Aigle et al., 1996). In contrast to previously described Streptomyces chromosomes, no readily detectable terminal-inverted-repeat (TIR) sequences were observed in the S. erythraea chromosome when chromosomal end restriction fragments were hybridized to total AseI- and DraI-digested genomic DNA. It remains to be determined if short TIR sequences are present in this organism. Such TIR sequences would likely be undetectable using the large end restriction fragment probes employed in these experiments (Reeves et al., 1998). A total of 15 genetic loci have been mapped to specific AseI and DraI restriction fragments. The erythromycin biosynthetic gene cluster has been localized to an approximately 700-kb AseI-DraI restriction fragment located within 1.25 Mb from one end of the linear S. erythraea chromosome. Interestingly, several additional unlinked genes possibly involved in erythromycin biosynthesis or resistance, including ertX, a putative ABC-type transporter (O'Neill et al., 1995), gdh, a thymidine diphosphoglucose-4,6-dehydratase, and kde, a putative thymidine diphospho-4-keto-6-deoxyglucose 3,5-epimerase (Linton et al., 1995), have been localized to the same 7O0-kb region of the chromosome. In addition, this region also contains the attB site for the integrative S. erythraea plasmid pSE101 (Brown et al., 1988). B. GENOMIC POLYMORPHISMS IN INDUSTRIALLY IMPROVED S. ERYTHRAEA STRAINS

The AseI, DraI, and AseI-DraI chromosomal restriction digest profiles appear similar between wild-type and industrially improved strains of S. erythraea, except for two notable genomic polymorphisms. AseI chromosomal digests of strain CA340 reveal the loss of the 48-kb AseIN fragment and the appearance of an approximately 75-kb novel restriction fragment (Reeves et al., 1998). Strain CA340 produces roughly 10-fold more erythromycin than wild-type strain NRRL2338. It is at present not known if this AseI polymorphism is related to this productivity increase. In addition, more recent industrially improved erythro-

MOLECULARGENETICSOF Saccharopolyspora erythraea

87

mycin production strains, derived from strain CA340, harbor an additional roughly 150-kb chromosomal deletion (unpublished observation). Again, it is not known if this chromosomal polymorphism is directly related to the concomitant increase in erythromycin productivity observed in this genetic lineage. However, several genetic explanations seem plausible to explain the correlation between these genomic rearrangements and the phenotypic improvement in erythromycin biosynthesis: (1) competing secondary metabolic pathways might be deleted, (2) a negative trans-acting regulator might be deleted, or (3) a positive trans-acting regulator might be activated (or duplicated) as a result of the chromosomal rearrangement. Additional studies of these S. erythraea chromosomal polymorphisms should clarify the competing hypotheses outlined above, improve our understanding of the fluidity of the S. erythraea genome, and potentially facilitate future rational strain development efforts with this organism. The identification of the dispensable gene set present in the large 150-kb deleted region of the S. erythraea chromosome might also contribute to molecular genetic strain development approaches in other polyketide producing actinomycetes. V. Introduction of DNA into S. erythraea

Protoplast transformation techniques have proven effective for introduction of plasmid DNA into wild-type strains of S. erythraea at efficiencies of 105 to 106 transformants per ~g of replicating plasmid DNA (Yamamoto et al., 1986). Difficulties in extending this technique to industrially improved fermentation strains of S. erythraea led to the development of two complementary electroporation methods for introduction of DNA into this organism (Fitzgerald et al., 1998; English et aL, 1998). Although electroporation has found widespread application for introduction of DNA (and other macromolecules) into a broad range of cell types, there remain few reports describing application of this technology to industrially important filamentous organisms (Pigac and Schrempf, 1995; Tyurin and Livshits, 1996). A. SONICATION-DEPENDENTELECTROPORATION The development of electroporation techniques for S. erythraea was facilitated by the availability of a virulent bacteriophage for this host, designated CABT1, from the Abbott Laboratories Culture Collection (Fitzgerald et al., 1998). Concentrated preparations of the dsDNA

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genome of this bacteriophage are readily obtained using standard phage propagation and purification techniques (Sambrook et al., 1989). This reagent permitted development of a sensitive electrotransfection assay and avoided potential problems with S. erythraea host restriction of shuttle vectors propagated in heterologous genetic backgrounds. During attempts to prepare well-dispersed homogeneous suspensions of vegetative S. erythraea cultures in our laboratory, we made the fortuitous observation that sonication treatment rendered this organism electrocompetent. Subsequent experimentation demonstrated that the observed electrocompetence was strictly sonication dependent for the vegetative cultures being examined. Culture preparation, sonication, and electroporation conditions were optimized using the electrotransfection assay to achieve electroporation efficiencies of 1.2 x 103 plaque forming units per microgram of CABT1 DNA. A plasmid-based electrotransformation assay was also developed to optimize sonicationdependent electroporation conditions for plasmid DNA uptake. This system utilized an Escherichia coli-Streptomyces shuttle vector, designated pCD1, derived from the pJV1 replicon of Streptomyces phaeochromogenes (Bailey et al., 1986; Fitzgerald et al., 1998). This vector is poorly maintained by S. erythraea but permits recovery and scoring of primary thiostrepton-resistant transformants prior to eventual loss of the plasmid. The electrotransformation efficiency obtained with this plasmid was 1.0 × 104 thiostrepton-resistant transformants per microgram of pCD1 DNA. Interestingly, the positive effect of sonication on electrocompetence was eliminated when the sonicated hyphal fragments were returned to culture tubes and incubated with shaking for 60 min prior to electroporation, suggesting that the physical alteration responsible for electrocompetence was eliminated or repaired during this period. It is tempting to speculate that the mechanical disruption of long vegetative hyphal fragments by ultrasound treatment in this procedure perhaps exposes interior hyphal membranes more susceptible to electroporation mediated DNA transfer. Alternatively, the sonication treatment might alter the normal outer cell wall to facilitate DNA entry into the cell. B. ELECTROPORATION OF GERMINATING SPORES

The amount of nonviable cellular material in the sonicated S. erythraea culture preparations prompted us to look for alternative (sonication-independent) conditions to achieve electrocompetence. Toward this end,

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cellular preparations from different stages of the developmental life cycle of S. erythraea were examined (English et al., 1998). The life cycle of S. erythraea is typical of other filamentous actinomycetes and has been reviewed elsewhere (Chater, 1998). This approach led to the discovery that germinating spores (germlings) of this organism move through a remarkably brief window of electrocompetence during outgrowth to vegetative hyphae. Under the culture conditions reported, the optimum outgrowth period for the harvest of electrocompetent germlings was between roughly 16 and 18 hours. The cellular physiological and/or morphological changes occurring during this period that result in electrocompetence are at present not understood. The utility of this method was demonstrated by constructing a targeted gene disruption of the pccA locus in the industrially improved S. erythraea strain CA340 (English et a]., 1998). The pccA gene encodes the biotinylated c~-subunit of the propionyl-CoA carboxylase from this organism. Electroporation efficiencies of 4-8 x 103 transformants per microgram of DNA were obtained with the suicide vector pJAY4 used in these experiments. Taken together with the protoplast transformation technique of Yamamoto et al. (1986), the two electroporation procedures reviewed here permit introduction of DNA into the full range of S. erj/thraea strains we have examined.

Vl. Transcriptional Organization and Regulation of the Erythromycin Biosynthetic Gene Cluster A. PREVIOUS TRANSCRIPTIONAL STUDIES OF THE ERYCI, ERME, AND ERYG GENES

Although DNA sequence analysis of the erythromycin biosynthetic gene cluster has provided some insights into the potential transcriptional organization of this gene cluster (Dhillon et al., 1989; Cortes et al., 1990; Donadio et al., 1991; Haydock et al., 1991; Donadio and Katz, 1992; Stassi et a]., 1993; Gaisser et al., 1997, 1998; Summers et al., 1997), detailed transcriptional studies have been lacking. Previous detailed transcriptional studies of ery genes have been limited to analysis of the eryCI-ermE region of the ery gene cluster reported by Bibb et al. (1994). A preliminary transcriptional study of the eryG gene has also been reported (Weber eta]., 1989). The regulatory region involved in divergent transcription of the eryCI and ermE genes contains two ermE and two eryCI promoters (Bibb et al., 1994). The ermE gene encodes an N-methyltransferase, which confers resistance to erythromycin in the producing host through methyla-

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tion of the S. erythraea 23S rRNA (Thompson et al., 1982). $1 nuclease and exonuclease VII transcript mapping experiments identified two transcriptional start sites for ermE, designated ermEpl and ermEp2. These experiments were performed using RNA isolated from Streptomyces lividans strain TK24 containing the eryCI-ermE promoter region cloned on high-copy-number plasmids. The divergently transcribed erythromycin biosynthetic gene eryCI is also transcribed from two promoters, designated eryCIpl and eryCIp2. The eryCI gene is thought to encode an aminotransferase involved in the synthesis of desosamine. Interestingly, transcription from ermEpl is initiated at the same position as eryCIp2, but on the opposing DNA strand. The existence of tandemly arranged, divergently transcribed overlapping promoters in this region led to the suggestion by these workers that a high degree of coordinate regulation occurs with these genes. The transcriptional start site for ermEpl is located immediately adjacent to the ermE translational start codon. This results in transcription of a leaderless message for the ermE gene. The ermEpl, ermEp2, and eryCIpl promoters contain recognizable -10 and -35 regions. In contrast, no recognizable -10 or -35 sequences were evident in the eryCIp2 promoter region, suggesting the involvement of alternative transcription factors in initiation of transcription from this promoter. Transcription of the eryG gene, which encodes the mycarosyl Omethyltransferase enzyme, was previously examined by Weber et al. (1989). Using Northern hybridization experiments, these workers identified an approximately 1.3-kb eryG transcript. A possible promoter region immediately upstream was reported based on an analysis of a cloned S. erythraea DNA fragment in a luxAB reporter system in S. lividans. More recent evidence from our laboratory has identified, in addition to the approximately 1.3-kb transcript described earlier, an additional 2.6-kb eryG transcript identifiable in Northern hybridization experiments (Reeves et al., 1999). This larger transcript contains the eryBII gene located immediately upstream of eryG. From our analysis of transcriptional terminator mutations in this region (see below), the stable eryG and eryBII-eryG messages detected in Northern hybridization experiments appear to be derived primarily from one of two very large overlapping polycistronic transcripts in this region. There is no evidence for a functional promoter in S. erythraea immediately upstream of either the erzG or eryBII gene. The previous results from Weber and coworkers (1989) might be attributable to the use of the high-copy-number luxAB reporter group plasmid and heterologous genetic background used in the earlier experiments.

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B. CONSTRUCTIONAND ANALYSISOF TRANSCRIPTIONAL MUTANTSIN S. ERYTHRAEA The development of a novel transcriptional terminator cartridge for the targeted construction of polar transcriptional mutants in S. erythraea has facilitated transcriptional studies of the erythromycin gene cluster (Reeves et al., 1999). These studies have improved our understanding of the transcriptional organization of this industrially significant biosynthetic gene locus. The transcriptional terminator cartridge, designated trm, contains a 227-bp transcriptional terminator sequence obtained from the cloned S. erythraea ribosomal RNA rrnD operon. This terminator sequence is flanked by convenient multiple cloning site sequences to permit in-vitro insertion of the cartridge into targeted genes of interest. A native S. erythraea terminator sequence was selected to ensure functionality in this host. Two regions of predicted secondary structure are present in the trm sequence. The first is a stem-loop structure consisting of a 17-bp stem with a short 4-base loop. This stem-loop is immediately followed by a thymidine-rich region, characteristic of rho-independent terminators (Deng et al., 1987). The calculated AG of the stem-loop is -30 kcal/mol. The second predicted region of secondary structure in this sequence is a stem-loop consisting of an 18-bp stem with a 4-base loop, located 30 bp downstream from the first stem-loop. This structure has a predicted AG o f - 2 4 kcal/mol. The t~m cartridge was cloned into targeted ery genes and introduced into the corresponding S. erythraea chromosomal loci via homologous recombination using either a pWHM3-derived (Vara et al., 1989) or pJVI-derived (Bailey et al., 1986) integration vector. Several additional features make S. erythraea an attractive experimental system for molecular genetic studies of the erythromycin biosynthetic pathway. These include: (1) the availability of cloned ery genes and DNA sequences, (2) the availability of DNA transformation and gene replacement methods for this organism, (3) the availability of purified erythromycin pathway intermediates, and (4) the ability of this organism to utilize exogenously supplied pathway intermediates in erythromycin biosynthesis. In our genetic studies of the ery gene cluster, biotransformation experiments with erythromycin pathway intermediates provided a simple, but powerful method to analyze the effects of specific trn~ transcriptional terminator insertion mutations (designated ery::t~n~) on expression of downstream ery genes. A variety of molecular genetic and biochemical methods, including Northern hybridizations, Western blotting, and $1 nuclease protection assays, were

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also employed in the characterization of the ery::trnn mutants. Two examples of the recently described S. erythraea tm~ insertion mutants (Reeves et a]., 1999) are reviewed here to illustrate the use of this methodology in the study of ery gene expression in wild-type and industrially improved genetic backgrounds. Based on these studies, the tm~ genetic element should also be of general use for the study of other genetic loci in this host. The tmn transcriptional terminator element was introduced in vitro into a cloned segment of the eryAI gene. This disrupted gene sequence was then introduced into the chromosome of wild-type strain NRRL2338 via homologous recombination to generate mutant NRRL2338 eryAI::tm~. The eryAI::tmn insertion was also introduced into the industrially improved genetic background of strain CA340 to generate mutant CA340 eryAI::tm~. As predicted, these eryA blocked mutants do not produce erythromycin A or any erythromycin pathway intermediates, consistent with previously characterized blocked mutants in the polyketide synthase genes. The central region of the erythromycin biosynthetic gene cluster contains, in addition to the three large eryA genes, four additional downstream genes. These seven genes, which include (in order) eryAL eryAII, eryAIII, eryCII, eryCIII, eryBII, and eryG, span approximately 35 kb of the erythromycin gene cluster. Several lines of evidence obtained from our analyses of the eryAI::tm~ mutants suggest that this set of genes is transcribed as a very large 35-kb polycistronic message. In biotransformation experiments, the eryAI::tmn mutation in both NRRL2338 and CA340 shows a polar effect on downstream ery genes, including the eryG Oomethyltransferase gene. Cultures of the eryAI::t~ mutant, when supplemented with erythromycin C, show greatly reduced levels of bioconversion of the erythromycin C to erythromycin A. Similar results were obtained in feeding experiments with other pathway intermediates, indicating an additional polar effect on both downstream eryB and eryC genes, as predicted. $1 nuclease protection assays using an eryG probe were also performed using total RNA extracted from the NRRL2338 eryAI::trnn and CA340 eryAI::i~nn mutant strains. A significant reduction of the eryG signal was observed in the insertion mutant strains relative to the parental controls, hterestingly, the polar effect of eryAI::tr~ insertion on eryG transcript levels appeared to be more pronounced in the CA340 industrially improved genetic background. This suggests that the promoter region immediately upstream of eryAI plays a more significant role in expression of the downstream eryG gene in the industrially improved strain CA340 than in the wild-type strain NRRL2338. Western blot experiments were also performed on wild-type strain NRRL2338 and the

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NRRL2338 eryAI::tm~ mutant to determine if the eryG O-methyltransferase protein was present in cell free extracts of these strains. A crossreacting band was readily detected in extracts of the parental control strain, but not in similarly prepared extracts of the eryAI::trnn mutant. In order to examine whether the eryAI::tm, insertion was having an effect other than on termination of transcription, an independent mutant strain was constructed using oligonucleotide-directed mutagenesis. In this mutant, designated ARR50, the predicted eryAI promoter ATrich -10 hexamer sequence (TATTGT) was replaced with the Sinai restriction enzyme recognition sequence CCCGGG. The polar phenotype of this strain was identical to that of the eryAI::trnn insertion mutant. Taken together, these experiments provide strong evidence that the eryAI, eryAII, eryAIII, eryBII, eryCII, eryCIII, and eryG genes are primarily cotranscribed from a promoter upstream of eryAL In addition, these experiments demonstrate the general utility of this new genetic element in the study of complex transcriptional units in this organism. C. ERYTtIROMYCINBIOSYNTHETICGENECLUSTERPROMOTERS The transcription start sites for seven additional ery gene cluster promoters, located in four ery gene cluster regulatory regions, have been reported (Reeves et al., 1999). These promoter regions include the 224-bp eryAI-eryBIV intergenic region, the 188-bp eryBI-eryBIII intergenic region, the 83-bp eryCVI-eryBVIintergenic region, and the region immediately upstream of the eryK gene. The 224-bp eryAI-eryBIV intergenic region contains an eryAI promoter divergently transcribed from two eryBIV promoters, designated eryBIVP1 and eryBIVP2. The eryAI transcription start site was located 27 bp upstream of the translation start codon for this gene. Based on an analysis of the eryAI::trnnmutant described above, the eryAI-containing transcript extends from eryAI to eryG. The transcription start sites for the eryBIVP1 and eryBIVP2 promoters are located 84-88 and 132 bp upstream of the predicted translational start codon for eryBIV, respectively. The eryBIVP1 promoter appears to be the major rightward promoter (as shown in Fig. 4) for this region based on the relative abundance of the respective Sl-protected fragments. The eryBIV-containing transcript is thought to extend from eryBIVto eryBVII, based on an analysis of ery::tmn insertion mutations in this region. This segment of the ery gene cluster also contains a smaller overlapping polycistronic transcript that extends from eryBVI through eryCVI. The promoter for this transcript is located in the 83-bp eryBVI-eryCVIintergenic region. The -35 region of the minor eryBIV promoter, eryBIVP2, is predicted

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~0

~3

! .I:i. oa°t "@~O

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to overlap with the -35 region of the divergent eryAI promoter. On the basis of our transcriptional analysis of the cry gene cluster, the three promoters identified in this 224-bp regulatory region account for the transcription of roughly 80% of the gene cluster, or 14 of the 20 identified cry genes. The sequence of this regulatory region was determined for an industrially improved strain of S. erythraea and compared to the wild-type strain NRRL2338 sequence. Surprisingly, no mutational changes were identified in this regulatory region in the industrially improved genetic background (unpublished observation) despite the significant increase in cry transcript levels and erythromycin titers observed in this genetic strain lineage (Reeves eta]., 1999). The transcriptional activity of DNA fragments containing the 224-bp eryAI-eryBIV regulatory region was examined using a kanamycin/neomycin phosphotransferase (APH) reporter group in both S. erythraea strain NRRL2338 and S. lividans strain TK24 (Atkins and Baumberg, 1998). In S. lividans, appreciable APH activity was detected in APH fusion strains regardless of the orientation of the S. erythraea eryAI-eryBIV DNA fragment. This finding is consistent with the presence of divergently transcribed promoters in this region. The function of these promoters in S. lividans further supports the notion that a pathway-specific activator is not involved in regulation of the cry gene cluster and is consistent with the previous observation that the ermE and eryCI promoters also function in this heterologous host (Bibb et al., 1994). The 188-bp eryBI-eryBIII intergenic region contains two divergently transcribed promoters. Two potential transcriptional start sites, located 1-2 bp upstream of the predicted start codon, were identified upstream of the eryBIII gene. An eryBIII::tr,,n insertion mutant displayed a polar effect on the downstream eryF gene, indicating that the eryF gene is cotranscribed with eryBIII. The start site for the divergently transcribed monocistronic eryBI message is located 17-18 bp upstream of the predicted translational start for the eryBI gene. The transcription start site for the eryK gene, which encodes the P450-dependent C-12 hydroxylase, was localized to a region 45-50 bp upstream of the predicted TTG start codon for this gene and 9-14 bp from the predicted termination codon of the adjacent orf21 coding region (Pereda et al., 1997). This gene, which is located on one end of the cry gene cluster, is transcribed as a monocistronic message. D. TRANSCRIPTIONAL OVERVIEW OF THE ERY GENE CLUSTER

A current transcriptional map of the erythromycin biosynthetic gene cluster is shown in Figure 4. The monocistronic messages for eryCI and

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ermE, described earlier, along with the more recently characterized eryBI and eryK transcripts (Reeves et al., 1999), represent the exception in this gene cluster, as the majority of ery genes appear to be transcribed as large polycistronic messages. This includes the previously reported eryG gene described above. Four polycistronic messages, including a large transcript of approximately 35 kb, account for the expression of roughly 85% of the gene cluster (Reeves et al., 1999). The two largest transcripts in the gene cluster extend divergently from the eryA-eryBIV intergenic region. The large 35-kb transcript includes the eryAL eryAII, eryAIII, eryCII, eryCIII, eryBII, and eryG genes. The divergently expressed transcript includes the eryBIV, eryBV, eryCVI, eryBVI, eryCIV, eryCV, and eryBVII genes. A second promoter upstream of the eryBVI gene produces an overlapping transcript that includes the eryBVI, eryCIV, eryCV, and eryBVII genes. Finally, a bicistronic message appears to be involved in expression of the eryBIII and eryF genes, and is divergently expressed from the monocistronic eryBI message.

VII. New Molecular Genetic Tools for Studying Gene Expression in S. erythraea

The gene encoding the native S. erythraea c¢-galactosidase enzyme, designated melA, has been cloned and sequenced (manuscript in preparation). The MelA enzyme was purified to near homogeneity and the N-terminal amino-acid sequence determined. Oligonucleotide primers based on the N-terminal amino-acid sequence and a conserved downstream region within the a-galactosidase gene were used to generate a 640-bp PCR product probe. Using this probe, the complete ct-galactosidase gene was identified in a lambda phage library of S. erythraea chromosomal DNA. The S. erythraea melA gene has been used to construct a reporter system for the study of gene expression in this organism. This system includes two components: (1) a parental S. erythraea strain containing a chromosomal deletion of the melA gene, and (2) a promoterless melA gene engineered into a pJVl-based integration vector. The melA integration plasmid is designated pDPE185. The promoterless melA gene cartridge present in pDPE185 is preceded by stop codons engineered in all three frames and includes a convenient upstream multiple cloning site. A ribosomal RNA terminator is present both upstream and downstream of the melA gene. Expression of the melA gene is conveniently monitored in liquid and solid agar cultures using either p-nitrophenol-

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0~-D-galactopyranoside (pNP{xG) or 5-bromo-4-chloro-3-indoyl-~-D-galactoside (X-~-gal), respectively. The function of this reporter group system was examined using the ermE* promoter region. The ermE* promoter fragment was inserted into the multiple cloning site of pDPE185. The resulting plasmid was introduced into the S. erythraea melA deletion host strain. Strains containing the ermE* promoter upstream of the melA gene had 10- to 15-fold more o~-galactosidase activity, as measured using the pNPRGbased assay, than the parental control strain. The melA reporter group system should be useful for constructing new S. erythraea vectors (e.g., insertional inactivation applications), for identifying new promoters, and in the study of gene expression in this organism (Satter, 1998).

VIII. Genetic-Engineering Approaches to Industrial Strain Improvement A. CONSTRUCTION OF HIGH-PRODUCTIVITY SOURCE STRAINS FOR NATURALLY OCCURRING ERYTHROMYCIN INTERMEDIATES

Genetic-engineering approaches have been used to introduce null mutations in the eryG and eryK genes, both individually and in combination, into various wild-type and industrially improved strains of S. erythraea. The analysis of mutants harboring disruptions of these genetic loci have permitted assignment of biochemical functions to these gene products (Paulus et al., 1990; Stassi eta]., 1993). In addition, genetically engineered eryG, eryK, and eryG/eryK null mutants represent important fermentation source strains for erythromycins C, B, and D, respectively (English et al., 1998). These erythromycin intermediates represent potential starting materials for development of new semisynthetic fermentation-based products (Faghih eta]., 1998). Naturally occurring erythromycin intermediates also find routine use among erythromycin manufacturers as analytical reference standards. Introduction of these mutant alleles into industrially improved erythromycin A-producing strains had little or no impact on overall macrolide titers {unpublished observation). This approach illustrates the utility of exploiting macrolide productivity levels of existing erythromycin A-producing industrial strains for production of other naturally occurring erythromycin intermediates. Moreover, this approach should also be applicable to the wide range of novel polyketide derivatives being generated through combinatorial biosynthesis methods with this organism (see below).

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B. TWO-STEP GENETIC ENGINEERING APPROACHES FOR OPTIMIZATION OF NOVEL MACROLIDE PRODUCTION IN S. ERYTHRAEA

Two reports by Stassi and coworkers (1998a, 1998b) illustrate the potential of rational strain development approaches for optimization of novel polyketide yields in genetically engineered strains of S. erythraea. The first report involves construction of a double mutant with disruptions in both the eryF gene, encoding the erythromycin C-6 hydroxylase, and the eryK gene, encoding the erythromycin C-12 hydroxylase. The predicted product of this double mutant is 6,12-dideoxyerythromycin A. Fermentations of the double eryF/eryK S. erythraea mutant produced a mixture of the desired product--6,12-dideoxyerythromycin A--and the immediate biosynthetic precursor--6,12-dideoxyerythromycin D--with the precursor representing the dominant product in the fermentation. In order to facilitate conversion of the 6,12-dideoxyerythromycin D precursor to the desired end-product, an additional copy of the eryG gene, encoding the 3"-O-methyltransferase enzyme responsible for this final biosynthetic step, was engineered into the eryF/eryK double mutant. The additional copy of the eryG gene under the control of the ermE* promoter was integrated into a chromosomal locus unlinked to the erythromycin gene cluster. The eryG diploid strain had significantly higher specific activities of EryG throughout the course of a 5-day fermentation. This increase in EryG activity resulted in quantitative conversion of the 6,12-dideoxyerythromycin D precursor to the desired 6,12-dideoxyerythromycin A product in fermentations of the eryF/eryK eryC--diploid strain. The second report involves addition of a heterologous primary metabolic enzyme to S. erythraea to improve precursor availability for a genetically engineered hybrid PKS. The ethylmalonate-specific acyltransferase domain from module five of the niddamycin PKS of Streptomyces caelestis (Kakavas et al., 1997) was substituted for the methylmalonate-specific acyltransferase domain present in module four of the erythromycin PKS (Stassi et al., 1998b). The predicted product of this mutant is 6-desmethyl-6-ethylerythromycin A. Interestingly, S. erythraea strains harboring this hybrid PKS still produced erythromycin A instead of the predicted hybrid polyketide product. The authors attributed this surprising result to the relaxed substrate specificity of the niddamycin acyltransferase domain used in this mutant construction and the limited intracellular availability of the ethylmalonyl-CoA precursor (relative to methylmalonyl-CoA) in this organism. In order to increase the available ethylmalonyl-CoA pools in this hybrid PKS mutant, the ccr gene from Streptomyces collinus expressed from the ermE*

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promoter was integrated at the same unlinked genetic locus used in the eryF/eryK mutant described above (Stassi et al., 1998a). The ccr gene encodes the crotonyl-CoA reductase enzyme responsible for the last step in the reductive biosynthesis of butyryl-CoA from two molecules of acetyl-CoA (Wallace et al., 1996) in S. collinus. Butyryl-CoA can then be carboxylated to form the desired ethyl malonyl-CoA precursor (Wallace et al., 1997). S. erythraea strains containing the S. collinus ccr gene had roughly 20-fold higher Ccr activity than the parental control strain. The addition of this activity to the PKS mutant led to production of the desired 6-desmethyl-6-ethylerythromycin A compound as the predominant product in fermentations of this strain. Taken together, these examples clearly demonstrate the utility of two-step genetic-engineering approaches to optimize both primary and secondary metabolic activities required for production of novel polyketides in S. erythraea. C. INTRODUCTION OF THE VITREOSCILLA HEMOGLOBIN GENE INTO S. ERYTHRAEA

Introduction of the Vitreoscilla hemoglobin gene (vhb) into an industrial strain of S. erythraea has been reported (Minas et al., 1998; Brunker et al., 1998). The vhb gene has been introduced into a variety of microorganisms, including Acremonium chrysogenum (DeModena et al., 1993), Escherichia coli (Khosla and Bailey, 1988), Corynebacterium glutamicum (Sander et al., 1994), Bacillus subtilis (Kallio and Bailey, 1996), and S. coelicolor (Magnolo et al., 1991) in attempts to improve oxygen metabolism and product yields in these organisms. However, the role of this gene in enhancing specific product yields in these systems remains speculative. The dramatic 70% improvement in erythromycin titers reported by these authors is difficult to assess due to the lack of information provided on the industrially improved S. erythraea strain used in these experiments. Nonetheless, this work represents the first reported attempt to improve erythromycin titers in S. erythraea through introduction of a heterologous gene into this organism. Functional expression of the vhb gene in the genetically engineered S. erythraea strain was confirmed by CO-difference spectrum assays. In the absence of selective pressure, the vhb integrant appeared to be genetically stable through at least a 9-day fermentation cycle. The molecular role of the Vhb protein in improving erythromycin titers in this strain is at present poorly understood. However, the improvement in erythromycin production observed in the genetically modified industrial strain did not appear to be due to simply an increase in biomass yield or to the pattern of mycelial fragmentation. The commercial

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utility of the S. erythraea::vhb strain described by these workers remains to be determined. Due to differences in industrial S. erythraea strain lineages, it will be of interest to determine whether the observed improvement in erythromycin yields is of general utility or limited to the model strain used by these workers.

IX. Combinatorial Biosynthesis In the past decade, intensive work in a number of laboratories (for reviews, see Staunton and Wilkinson, 1997; Katz, 1997) has led to significant advances in our understanding of the genetics and biochemistry of the erythromycin type I polyketide synthase in S. erythraea. Early molecular genetic studies of this system revealed the modular structure of this complex enzyme system and provided significant insights into the functional roles of individual enzymatic domains within the eryA-encoded PKS (Cortes eta]., 1990; Donadio et al., 1991). These studies also led to a series of successful genetic-engineering efforts involving targeted gene replacements of specific PKS modules to produce a number of novel erythromycin derivatives (reviewed by Katz, 1997). Although providing an important validation of this genetic-engineering approach to produce new molecular structures, this methodology suffered from several significant limitations. The construction of individual mutants was labor intensive and involved single mutational changes. In addition, mutants were produced in wild-type S. erythraea genetic backgrounds, leading to relatively low titers of the novel erythromycin derivatives produced in these strains. The development of combinatorial genetic-engineering approaches that permit introduction of multiple genetic alterations into the ery PKS represents a significant technical advance and illustrates the potential of this gene cluster for production of a wide range of polyketide compounds (McDaniel et al., 1999). Parallel advances in precursor-directed biosynthesis of novel polyketide derivatives using genetically engineered variants of erythromycin PKS further demonstrate the promise of S. erythraea and the role of erythromycin biosynthetic genes in the emerging field of combinatorial biosynthesis (Marsden et a]., 1998; Pacey et al., 1998; Weissman et al., 1998a). These technical developments have generated considerable interest of late, as reflected in the number of reviews that have appeared on this topic (Hutchinson, 1998, 1999; Cane et al., 1998; Hallis and Liu, 1999). In the context of this review, I consider briefly here the complementary combinatorial biosynthesis experimental platforms that have emerged for manipulation of erythromycin biosynthetic genes in both the native host S. erythraea

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and heterologous bacterial hosts. Continued development of appropriate experimental platforms to fully exploit the potential of S. erythraea and the ery gene cluster in combinatorial biosynthesis applications will play a significant role in successfully translating this exciting new technology into therapeutically useful and commercially viable products. Ultimately, the success of these approaches in drug discovery programs will be measured not by the number of new chemical entities produced in submilligram quantities, but rather by the number of viable therapeutic leads that move into preclinical and clinical testing. A. MANIPULATIONOF ERY BIOSYNTHETICGENESIN HETEROLOGOUSSTREPTOMYCESHOSTS The milestone report by McDaniel et al. (1999) illustrates the successful use of a heterologous system to manipulate the erythromycin PKS system. These authors report production of >100 6-DEB derivatives through construction of erythromycin PKS mutants harboring multiple combinations of individual eryA mutations in either S. coelicolor or S. lividans. The mutations include AT substitution, KR deletion, KR gainof-function, and KR stereochemical alterations. The use of S. coelicolor and S. lividans offers numerous experimental advantages because of the well-developed molecular genetic systems in these organisms (Hopwood et al., 1985; Gusek and Kinsela, 1992; Hopwood, 1997). This includes a wide range of plasmid vectors and gene expression tools, well-established protoplast transformation methods, and available genomic information (Kieser et al., 1992). These genetic tools facilitate construction of the numerous polyketide synthase mutants required to produce a library of 6-DEB polyketide derivatives. The host strains used by McDaniel and coworkers, S. coelicolor strain CH999 and S. lividans strain K4-114, contain a chromosomal deletion of the entire actinorhodin polyketide gene cluster (Ziermann and Betlach, 1999). Deletion of the actinorhodin biosynthetic genes in these hosts simplifies quantitative analysis of the novel polyketides produced and reduces competing metabolic demands for production of extrachromosomally encoded PKS proteins. The erythromycin PKS genes are expressed in these strains from the pCK7 expression plasmid, which has been reviewed elsewhere (Katz, 1997). Several limitations of this heterologous experimental platform should also be considered. First, the S. coelicolor system described by McDaniel et al. (1999) involved only the polyketide synthase genes from S. erythraea to produce 6-DEB derivatives. The erythromycin biosynthetic genes required for biosynthesis and attachment of the two

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deoxysugars mycarose and desosamine, and for the hydroxylation and O-methylation modification reactions were not provided in the heterologous host. This required separate introduction of the PKS mutations into S. erythraea to examine the impact of the polyketide modifications on the downstream biosynthetic reactions. The importance of the glycosylations and further macrolide ring and sugar modifications on the bioactivity of erythromycin derivatives underscores the importance of this step in evaluating this combinatorial library. Presumably, the genetic versatility of S. coelicolor and S. ]ividans would permit further genetic engineering of these hosts to include these ancillary reactions if desired. However, further development of the native S. erythraea host might represent a more attractive option since this organism already contains the required secondary metabolic pathways necessary to supply biosynthetic precursors for these pathways. Second, the yield of 6-DEB in the S. coelicolor control strain harboring the wild-type erythromycin PKS genes was approximately 20 mg/liter. All mutational changes in the PKS genes resulted in lower yields from this parental baseline, presumably reflecting differences in substrate specificity and processivity in the downstream polyketide biosynthetic reactions. The authors also noted that yield losses correlated with particular single mutations appeared to be additive when engineered in multiple combinations into the hybrid PKS. The resulting yields of the 6-DEB derivatives produced in the S. coelicolor library ranged from <0.5 to 70% of the baseline 20 mg/liter yield. Further productivity losses are also likely at the post-PKS glycosylation and modification steps. Clearly, improving the relatively low yields of the novel polyketide compounds reported in this system represents an important technical challenge if this technology is to become a viable drug discovery tool. B.

MANIPULATION OF ERY BIOSYNTHETIC GENES IN S. ERYTHRAEA

Several laboratories have successfully engineered wild-type strains of S. erythraea to produce novel erythromycin derivatives using gene replacement methods to introduce mutations in the PKS genes in this organism (for review, see Katz, 1997). Although providing an important conceptual validation of genetic engineering strategies to produce new polyketides using this organism, these studies proved to be labor intensive and produced a relatively small number of new compounds. Several PKS mutant constructions failed to produce detectable levels of the desired polyketide compounds, although the genetic information was confirmed present in the chromosomal gene locus (Ruan et al., 1997;

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Stassi et al., 1998b). In addition, the dependence on wild-type S. erythraea strains to perform molecular genetic manipulations in these experiments significantly limited the yields of novel erythromycin derivatives even in successful genetic constructions (Katz, 1997). Taken together, these factors, including the relatively low throughput of compounds, the poor predictive ability in regard to permissible genetic alterations in the erythromycin PKS, and the low product yields of new erythromycin derivatives, were inconsistent with the demands of modern drug discovery programs and led to a reduction or elimination of drug discovery efforts utilizing these directed genetic approaches at several pharmaceutical houses in the late 1990s. The development of improved S. erythraea host strains, DNA transformation methods, and expression plasmids should stimulate renewed interest in these genetic-engineering approaches. The S. erythraea strain described by Rowe et al. (1998) should prove useful in combinatorial biosynthesis applications in this organism. This strain, designated JC2, harbors a deletion of almost the entire eryA region. The region of eryAIII encoding the chain-terminating TE domain has been retained at the chromosomal ery gene locus. This deletion mutant permits expression of genetically engineered PKS derivatives in the absence of a wild-type PKS. A homologous deletion construct introduced into an industrially improved genetic background would represent a particularly useful improvement in this technology. These authors also describe the construction and use of an expression vector, designated pCJR24, that permits expression of genetically engineered PKS genes under the control of the actI promoter and cognate activator protein ActII-ORF4 in strain JC2. The extension of precursor-directed polyketide biosynthesis in S. erflhraea by Leadlay and coworkers (Marsden et al., 1998; Pacey et al., 1998; Parsons et al., 1999) offers a promising complement to other combinatorial biosynthesis strategies in this organism. These workers substituted the S. avermitilis PKS loading domain involved in avermectin biosynthesis for the eryAI-encoded loading domain involved in erythromycin biosynthesis. Previous studies (Dutton eta]., 1991) demonstrated the broad substrate specificity of the avermectin PKS loading domain. This broad substrate specificity was exploited to incorporate a variety of substrates into the avermectin polyketide backbone, resulting in production of a series of novel C-25 substituted avermectins. The genetically engineered strain of S. erythraea containing this modified PKS, when supplemented with various exogenous fatty acids, produced a variety of novel C-13 erythromycin species as predicted.

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X. Future Prospects

The continued interest in erythromycin and semisynthetic second- and third-generation erythromycin derivatives as antiinfectives will continue to justify ongoing research in this molecule and in the producing organism, S. erythraea. Since large-scale production of semisynthetic compounds necessarily involves losses of the parent natural product in the synthetic reaction steps, there will be continued demand for improved fermentation manufacturing technologies to economically produce high-quality erythromycin A or advanced intermediates for these products. This is particularly evident from the structural complexity (and related complexity of the synthetic chemistries involved) of the lead third-generation ketolide drug candidates in clinical development (Fig. 1). The recent advances in the development of molecular genetic tools for manipulation of S. erythraea offer significant promise for further metabolic engineering improvements in the full range of fermentation strains supporting the manufacture of these products. Transcriptional studies of erythromycin biosynthesis have provided new insights to define rational strain improvement strategies for this pathway. These efforts should continue to provide improvements in erythromycin production technology in the years ahead. The recent developments in combinatorial biosynthesis using S. erythraea and erythromycin biosynthetic genes have significantly improved the prospects for discovery of new therapeutic candidates derived from this system. Two key issues will need to be addressed if we are to realize the potential of this technology. In order to compete with combinatorial chemistry and other alternative drug discovery approaches, combinatorial biosynthesis systems capable of producing significantly larger compound libraries will be necessary The progress in this area, reviewed by Hutchinson (1999), is encouraging in this regard. In addition, the productivity of genetically engineered S. erythraea mutants, used in either genetic or precursor-directed combinatorial biosynthesis applications, must be improved. This is essential both in order to support the drug discovery process and to facilitate timely preclinical and clinical development of lead drug candidates. Although further development of heterologous hosts like S. coelicolor is likely to improve the current baseline productivity of these systems, this is unlikely to be competitive with approaches utilizing the native host S. erythraea. Existing high-productivity mutants have not been fully exploited in this regard, probably owing to the separation of process

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development and drug discovery groups within large pharmaceutical houses and the general unavailability of these proprietary strains to academic laboratories and small biotechnology companies engaged in the field. This would perhaps be most effectively addressed through exploration of potential licensing and collaborative opportunities in this area. Existing technologies developed for large-scale fermentation of S. erythraea and the experience with this organism in industry represents a significant commercial, manufacturing, and regulatory advantage for development of new drug candidates using this platform. It seems likely that S. erythraea mutants producing several orders of magnitude higher polyketide levels would open a larger window of observation for new compounds during the drug discovery process and facilitate the isolation of gram to kilogram quantities for further preclinical and clinical testing. As an alternative, the combinatorial drug discovery phase could perhaps be supported by moderately improved yields in the heterologous host systems noted above. Appropriate vectors for transfer of the genetically engineered biosynthetic genes into higher-productivity S. erythraea genetic backgrounds could then be developed to support the demands of the later stages of the drug development process. The continuing advances in our ability to genetically manipulate S. erythraea, along with the highly developed fermentation technologies available for this organism, will ensure a continuing role for S. erythraea in the development and production of new drug candidates in the years ahead.

Acknowledgments I am grateful to the many researchers who have contributed their time and talents to the study of erythromycin and Saccharopolyspora erythraea over the past four decades. I am especially indebted to my Abbott colleagues, past and present, who have enriched my working life through their insights into this organism. In particular, I would like to acknowledge David Post, Diane Stassi, Leonard Katz, Martin Babcock, Jay Lampel, Mark Satter, Andrew Reeves, Samuel English, James Petzel, Bill Ellefson, Nancy Fitzgerald, Sandra Splinter, Donna Santucci, Tom Paulus, Janet DeWitt, and Vicki Luebke for their contributions toward improving our understanding of the genetics and physiology of this industrially important microorganism.

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Bioactive Products from

Streptomyces

VLADISLAV BI~HAL

Institute of Microbio]ogy Academy of Sciences of the Czech Republic Prague 14220, Czech Republic

I. Introduction II. Chemistry and Biosynthesis A. Peptide and Peptide-Derivative Antibiotics B. Polyketide Derivatives C. Other Bioactive Products III. Genetics and Molecular Genetics A. Preparation of High-Production Microorganisms B. Genetic Manipulation of Secondary Metabolite Producers IV. The Search for New Bioactive Secondary Metabolites A. Isolation from Natural Resources B. Producers of Bioactive Compounds C. Screening D. Senfisynthetic and Synthetic Bioactive Products E. Hybrid Bioactive Products and Combinatorial Biosynthesis V. Regulation of Secondary Metabolite Production A. Growth Phases of Streptomycetes B. Control of Fermentation by Basic Nutrients C. How Signals from the Medium Are Received D. Regulation by Low-Molecular-Weight Compounds E. Autoregulators F. Regulation by Phosphorylated Nucleotides G. Regulation by Metal Ions VI. Resistance to Bioactive Products A. Resistance of Secondary Metabolite Producers B. Resistance in Pathogenic Microorganisms References

I. Introduction

Medicine was transformed in the latter half of the twentieth century by the discovery of antibiotics and other bioactive secondary metabolites produced by microorganisms. Antibiotics are defined as microbial products that inhibit the growth of other microorganisms. After the antibacterial effect of penicillin had been observed by Fleming, a number of other antibiotics were discovered, mainly those produced by soil Streptomyces spp. and molds. Moreover, a broad spectrum of natural 113 ADVANCES IN APPLIED MICROBIOLOGY. VOLUME 47 Copyright © 2000 by Academic Press All rights of reproductiun in any form reserved. (1(165-2164/00 $25.00

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VLADISLAVBI~HAL

products having other effects on living organisms were found in microorganisms. In addition to antibiotics, the following compounds have been found: coccidiostatics used in poultry farming, antiparasitic compounds with a broad spectrum of activity against nematodes and arthropods, substances with antitumor activity, immunosuppressants, thrombolytics (staphylokinase), compounds affecting blood pressure, and so forth. Microbial metabolites also exist that exhibit good herbicide and pesticide activities and are biodegradable. However, microbial herbicides and pesticides are not widely used due to their high price. Another special group of natural products are the enzyme inhibitors synthesized by microorganisms (Umezawa et al., 1976). These compounds can inhibit antibiotic-degrading enzymes, as well as certain enzyme activities in human metabolism that cause illness. Many enzyme inhibitors are protease inhibitors, variously active against pepsin, papain, trypsin, chymotrypsin, cathepsin, elastase, renin, and so forth. Inhibitors of glucosidases, cyclic AMP, phosphodiesterase, different carbohydrases, esterases, kinases, phosphatases, and the like have been isolated from Streptomyces. The enzyme inhibitors that block synthesis of cholesterol are also important. Others exhibit immunosuppressive effects, the most well known being cyclosporin A (a cyclic undecapeptide) produced by filamentous fungi. Some macrolide antibiotics isolated from Streptomyces are also immunosuppressants. Several thousand biologically active compounds have been described, and each year new compounds are isolated from microorganisms. Microorganisms are a virtually unlimited source of novel chemical structures with many potential therapeutic applications. The term "secondary metabolite" was first used for some microbial products by Bu'Lock in 1961, and the suitability of this term was discussed by Bennett and Bentley (1989). Secondary metabolites are compounds that the microorganism can synthesize but are not essential for basic metabolic processes such as growth and reproduction. Nevertheless, many secondary compounds function as so-called signal molecules, used to control the producer's metabolism. Another function attributed to antibiotics is suppression of competing microorganisms in the environment, whereby the antibiotic-producing microorganisms have an advantage in competing for nutrients. The amount of secondary metabolites in microorganisms isolated from nature is rather low in most cases. To be usable for commercial production, high-yielding strains need to be selected through multiple mutations of the strain's genetic material, culture conditions must be optimized, and in some cases genetic engineering also increases yields.

BIOACTIVE PRODUCTS FROM Streptomyces

115

II. Chemistry and Biosynthesis Despite the complexity of some of their structures, bioactive secondary metabolites are synthesized from simple building blocks used in living organisms for the biosynthesis of cellular structures. These units include amino acids, acetate, propionate, sugars, and nucleotides. According to their structure and type of biosynthesis, secondary metabolites can be classified into several groups. A. PEPTIDE AND PEPTIDE-DERIVATIVEANTIBIOTICS

Microorganisms produce a number of peptides as secondary metabolites. These peptide antibiotics are not synthesized on ribosomes but on enzyme complexes called peptide synthetases (Lipmann, 1971; Laland and Zimmer, 1973). In peptide antibiotics, the peptide chain is often cyclic or branched. In addition to L-amino acids, other compounds can be present in the molecule, such as [3-amino acids, organic acids, pyrimidines, and sugar molecules. The best-known bioactive peptides, gramicidins and bacitracins, are produced by different strains of Bacillus licheniformis and Bacillus brevis; some peptide antibiotics are produced by Streptomyces spp. (Kleinkauf and von Doehren, 1986). The linear molecule of gramicidin A and the cyclic molecule of gramicidin S (Fig. 1) belong to the structurally simplest class of peptide antibiotics. Bacitracins are examples of cyclic peptides having a sidechain (Fig. 2). In the molecule of bleomycin, the sugars L-glucose and 3-O-carbamoyl-D-mannose are found. Peptide antibiotics containing an atom of iron or phosphorus in the molecule have also been isolated. If two molecules of cysteine are present in the peptide antibiotic, they are linked by a sulfide bridge. The - - C O - O - - bond in the antibiotic molecule is present in lactones. Such antibiotics are represented especially by the group of actinomycins that contain a phenoxazine dicarboxylic group bearing two peptide chains. The enniatine molecule consists of three residues of branched amino acids--L-valine, L-leucine, and L-isoleucine--and three residues of D-2hydroxyisovaleric acid (D-Hyiv) (Billich and Zocher, 1987). The amino acids and D-Hyiv are linked by alternating amide and ester bonds. The amide bonds are finally Nomethylated. Molecular conformation is important for the biological activity of peptide antibiotics, especially for the peptides capable of forming chelates with metals. Studies showed three-dimensional molecular structures with many hydrogen bonds (Iitaka, 1978). In the case of valinomycin (L--Val-D--Hyiv-D--Val-L--Lac)3, which transports K+ and Rb ÷ ions across natural and artificial membranes, the molecule is symmetrical in three dimensions if it forms a complex with the metal. If it is not in the form of the complex, it has only a pseudocentral symmetry.

116

VLADISLAV Bt~HAL

i

2

5

HCO - L - V a I - ~ L - G l y 15

12

L - T r p " = - D - Leu ~ i

14

D-Leu-"-

~

4

L-AIn -~

5

D-Leu --~L-AIn

11

I0

L-Trp ~

D-Leu ~

6

--l,'- D - V o l

9

S

/

17

L - T r p ' q - - D - V o l ~ - L.Vol

[5

L-Trp ~

NH CH 2 - - - - C H 2 0 H

/1I D -Phe L - L e u io i

i

GramicidinA

2L-Pr o

\

L- Orn 9

L-Val

a

L- ~ . ~

3 L- Phe

4 D-Phe s 5 L-Asn L-GIn J Gramicidin S

FIG. 1. Gramicidin A (top) and Gramicidin S (bottom).

! L-tie I

1 L-Cys

-"- L-Leu ---.--",- D-4Glu "----~='-L-Ile ~ L -

Lys

2 L- Asn

D - Orn

II

L-lie

D -Asp

\,o L- His

FIG. 2. Bacitracin.

.

,7 D - Phe

BIOACTIVE PRODUCTSFROM Streptomyces

117

The biosynthesis of peptide antibiotics takes place on a multienzyme complex (Kleinkauf and von Doehren, 1983, 1986). The individual amino acids are activated with the use of ATP to form aminoacyl adenylates. The aminoacyl groups are transferred to the enzyme thiol groups, where they are bound as thioesters. The structural arrangement of the thiol groups in the synthetases determines the order of amino acids in the peptide. Formation of peptide bonds is mediated by 4-phosphopantetheine, an integral part of the multifunctional multienzyme. The intermediate peptides are also bound to the synthetases by the thioester bond. The way in which the order of the amino acids in the molecule is regulated is not known. It is probably determined by the tertiary configuration of the enzyme. Our knowledge of the biosynthesis of peptide antibiotics comes mostly from studies of the gramicidin S and bacitracin synthetases. Gramicidin S synthetase consists of two complementary enzymes having molecular weights of 100 and 280 kD, while bacitracin synthetase consists of three subunits (Roland et al., 1977) (Fig. 3) having molecular weights of 200, 210, and 360 kD (lshihara et al., 1989). Each subunit contains phosphopantetheine. Enzyme A activates the first five amino acids of bacitracin, enzyme B activates L-Lys and L-Orn, and enzyme C activates the other five amino acids. D-amino acids are pro-

Asn

Orn

Ile ~

~ Lys 03

-1-

FIG. 3. Bacitracinsynthetase.

118

VLADISLAV BI~HAL

duced by racemization of their L-forms directly on the enzyme complex. Initiation and elongation start on subunit A up to the pentapeptide, independent of the presence of subunits B and C. The pentapeptide is transferred to subunit B, where two other amino acids are added. The heptapeptide is subsequently transferred to subunit C, where the biosynthesis of bacitracin is completed. Cyclization is achieved by binding the asparagine carboxy group to the e-amino group of lysine, whereas the isoleucine carboxyl group is bound to the a-amino group of the same lysine (Schlumbohm et al., 1985; Ishihara and Shimura, 1988; von Doehren, 1995). The antibiotic activity of bacitracin results in efficient inhibition of protein synthesis and cell wall synthesis, but other effects such as an interference with cytoplasmic membrane components and cation-dependent antifungal effects have been observed as well. In the case of gramicidin S, hemolytic effects, inhibition of protein phosphatases, and interaction with nucleotides have been observed in addition to antibacterial activity. Even though antibiotics normally have several mechanisms of action, the primary one is defined as the effect observed at the lowest active concentration. The peptide antibiotics are efficient mainly against Gram-positive bacteria. The [3-1actams are peptide-derived secondary metabolites. They are produced by many different microorganisms. Several reviews summarize research in this area (see, e.g., Martin and Liras, 1989; Jensen and Demain, 1995). The main producers are fungi (penicillins), but they are also produced by Streptomyces (clavulanic acid) and Cephalosporium (cephalosporins) spp. The main representatives of [~-lactams are penicillins and cephalosporins. Penicillins have a thiazoline ~-lactam ring in the molecule and differ, one from another, by sidechains linked via the amino group (Fig. 4). Cephalosporins have a basic structure similar to that of penicillins, and the derivatives are formed by variation of the sidechain (Fig. 5). The thiazolidine [3-1actam ring is synthesized using three amino acids: L-a-amino adipic acid, L-cysteine, and L-valine. By condensation of these three amino acids, a tripeptide is formed. It is transformed into the molecule of penicillin or cephalosporin through subsequent transformations. Clavulanic acid, produced by Streptomyces clavuligerus, also belongs to the [3-1actam family (Reading and Cole, 1977). This acid has a bicyclic ring structure resembling that of penicillin, except that oxygen replaces sulfur in the five-membered ring (Fig. 6). Clavulanic acid is an irreversible inhibitor of many [~-lactamases. The discovery of clavulanic

BIOACTIVE PRODUCTS FROM

R-CO -NH _ _ ~ S O/~.

Streptomyces

,..~ CH3 ~CH 3 --COOH

N

R

~ C H HO--~

2

Penicillin

CH2

Penicillin X

CH5- ( CH2)6

G

Penicillin K

0-CH2

Penicillin V

H2N - CH--lCH2)3 I COOH

Penicillin N

FIG, 4. Penicillins.

R,--CO--NH ~- ~ = L~

S~CH " 2R2 COOH

R~

R2

H2N --CH-- (CH2) 3 - - OCOCH3

Cephalosporin C

~'~CH

Cephalothin

2-

--

~ ' ~ s ~ CH2 ~S-CH Nw C -- CH2 --

2

OCOCH5

--N~

Cephaloridin

- OCOCH5

Cephacepirin

- OCOCH3

Cephacetril

Flc. 5. Cephalosporins.

119

120

VLADISLAV Bt~HAL ~O

F oJ N:) J

OH

COOH

FIG. 6. Clavulanic acid.

acid was a starting point for the development of penicillin analogues able to inactivate these enzymes. Penicillins are especially active against Gram-positive bacteria, but some semisynthetic penicillins, such as ampicillin, which is lipophilic as compared to, for example, benzyl penicillin, are also effective against Gram-negative bacteria. This effect is explained by their ability to readily enter the cells of Gram-negative bacteria, which have a high lipid content in the cell wall. ~-lactam antibiotics interfere with synthesis of bacterial cell walls and thus inhibit bacterial growth. Such a mechanism of action does little harm to the macroorganism to which ~-lactams are administered. Another example of amino acid bioactive substances are the glycopeptides, including semisynthetic derivatives (Zmijewski and Fayerman, 1995). The best known of these is vancomycin (Fig. 7) (Harris and Harris, 1982), effective against Gram-positive bacteria. This antibiotic is widely used in medicine, especially against [3-1actam-resistant strains. Vancomycin is not absorbed from the gastrointestinal tract and is used to treat enterocolitis caused mainly by Clostridium difficile. Vancomycin is produced by many species, of which Amycolotopsis orientalis is used for commercial production. Glycopeptides are composed of either seven modified or unusual aromatic amino acids or a mix of aromatic and aliphatic amino acids. By substitution of amino acids in the amino-acid core, derivatives of amino glycosides are formed. In vancomycin the aminosugar vancosamine is bound to the amino-acid core. Removal of aminosugar reduces the activity of vancomycin by two- to fivefold. The sugars seem to play an important role in imparting enhanced pharmacokinetic properties for vancomycintype glycopeptide antibiotics (Williams eta]., 1990). B. POLYKETIDEDERIVATIVES

Polyketides are a large group of secondary metabolites synthesized by decarboxylative condensation of malonyl units, often with subsequent

BIOACTIVE PRODUCTS FROM

H3C ....~/~

Streptomyces

(~-~'~ CH2OH CH 3 ~-O "O

CI

NO -

..-

"'H H " ~

L.'H

,o

".-4. ,

NH2

HO /

"
121

OH

NH

O

Hj

"~'CH 3

OH

FIG. 7. Vancomycin.

cyclization of the polyketide chain. The starter group may be an acetate, but it may also be pyruvate, butyrate, ethyl malonate, or paraaminobenzoic acid, among others. Formation of the initial polyketide chain is similar to that taking place during biosynthesis of fatty acids and is catalyzed by polyketide synthases. Simple carboxylic acids are activated as thioesters (acyl-SCoA), which are carboxylated to form malonyl-CoA, methylmalonyl-CoA, and ethylmalonyl-CoA, and are polymerized after decarboxylation (Lynen and Reichert, 1951; Lynen, 1959; Lynen and Tada, 1961). A principal role is played by the acyl carrier protein (ACP) (Goldman and Vagelos, 1962). ACP detected throughout the growth of Streptomyces glaucescens was purified to homogeneity and found to behave like many other ACPs from bacteria and plants. The ACP prosthetic group in many microorganisms is 4'-phosphopantothenic acid. Its terminal groups and acyls produced by polymerization are bound via the --SH group. The acyls are transferred to the other --SH group, which is part of the cysteine molecule. 6-methyl salicylic acid (6MS) is one of the simplest polyketides formed by condensation and subsequent aromatization of one acetylCoA molecule and three malonyl-CoA molecules. This compound was isolated from Penicillium patulum (Bu'Lock and Ryan, 1958). By other metabolic steps 6MS is transformed to produce a toxin called patulin

122

VLADISLAVBI~HAL

COOH OH

H}C~

HZ~-~

I~

~--"~1~ 142u I"-SpH .^ •

~'~-s --~S_H P

'

7

CO-CH~-CI~

\ CoA-SH ~ CoA" S'CO- CH~ COOH

CoA -S-CO-CH 5 CoA-SH

"

*~,1:" " s p ' c o ' c H 3 L %.ScH teA-I-CO.¢ -4~0t4

o.~-~T,

E_:f #'

-co~'~ c~

I~'SPH / r . S~H

CoA-SH

Vs~.co.c,z_coo,

CH3

co,

\tHai CO'CHz-COOte

~ /co-cH 2 cH\ I~ -Sp CH -ScH o,c /

\E::C

E "$pH l "Sc'CO'CH2"CO'CH3

c,. , co c.icoo.~E

\CH 3

CaA- 1114 .o,.

" Sp-CO'CH2" CO'CH3 - ScH

:

_,....

~,¢~,~.o..

oI,

c.z-.,.~, ! H20

co 2 I

CH2 CH-OH I

CH2 I C =0 I

C=O I CH3

C:O I CH3

Fzc. 8. 6-methylsalicylic acid synthetase.

(Sekiguchi, 1983; Sekiguchi et al., 1983). The synthesis of 6MS takes place on an enzymatic complex called 6MS synthetase (Fig. 8) (Dimroth et al., 1970, 1976). Tetracyclines are important antibiotics that are used widely in human and veterinary medicine. The chemical structure of some typical tetracyclines is shown in Figure 9, and their biosynthesis is depicted in Figures 10 and 11 (McCormick, 1965). Chlortetracycline (CTC) and tetracycline (TC) are produced by Streptomyces aureofaciens, whereas oxytetracycline (OTC) and tetracycline are produced by Streptomyces rimosus. For more extensive coverage of research, articles by B~hal et

BIOACTIVE PRODUCTS FROM Streptomyces

123

19 a Me~N~MO ~

~ N H 2 OH 0 OH 0

Tetracycline Chlortetracycline 6-Demethyltetracycline 6-Demethylchlortetracycline Oxytetracycline 6-Deocytetracycline Minocycline

0

5

6

6

7

H H H

CH3 CH3 H 14 CH3 CH3 H

OH OH OH OH OH H H

H CI 11 CI [l H N(CH3)3

H

OH OH H

FIG. 9. Tetracyclines.

O

H2NCO - CH?_ CO$CoA - -

8 HOOC- CH2 -- COSCoA

~

~

c

o

s

-- E

O

0

0

0

0 )

~

COS -- E

COOH

CONH z 0

0

Me

0

0

0

OH

0

OH

OH

Mo

OH

CONH2

CONH2 OH

0

OH

Oxanthranol

OH

OH

6 - Methylpretetramide

FIe. 10. Tetracycline biosynthesis.

124

V L A D I S L A V BI~HAL

T

~

O

~

£

z z o

0 ~

8

z 0

I 0

o-~ E8 0

~c

o

g

4

a.

o ci z

z

£ ~

z

~

O

o

£

z 0

T 0

"-

z

u

6~ =o

o

9= "r, o o

=o

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o

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og

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T: ~,o

o

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3"

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~4

d

Z

0

£

z

o

I

0

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o

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T

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o

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K

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-0

4 ~

z o

.li ,° _o o

BIOACTIVE PRODUCTS FROM

Streptomyces

125

al. (1983), B~hal (1987), and B~hal and Hunter (1995) should be consuited. Tetracyclines act as inhibitors of protein synthesis. They are considered to be broad-spectrum antibiotics that are efficient against both Gram-postive and Gram-negative bacteria, but with significant side effects in humans. They are preferably used only w h e n other less toxic antibiotics are not effective. Anthracyclines are synthesized in a similar way as tetracyclines; however, they often have one or several sugar residues in the molecule. Most often deoxysugars, synthesized from glucose, are present in the anthracycline molecule. Daunorubicin and doxorubicin (adriamycin) (Fig. 12) are excellent antitumor agents, and are widely used in the treatment of a number of solid tumors and leukemias in humans. Unfortunately, these drugs have dose-limiting toxicities such as cardiac damage and bone marrow inhibition. A variety of drug delivery systems for anthracyclines have been reported. In most cases, the drugs were linked to high-molecular-weight compounds such as dextran (LeviSchaff et al., 1982; Tanaka, 1994) and DNA (Campeneere et al., 1979). Anthracyclines are produced by many Streptomyces spp. and the genetics of their production is well elaborated (Hutchinson, 1995). Macrolides are usually classified so as to include proper macrolides having 12-, 14-, or 16-membered macrocyclic lactone rings to which at least one sugar is bound, and polyenes having 26- to 38-atom lactone rings containing two to seven unsaturated bonds. Aside from the sugars bound to the lactone ring, an additional aromatic part is normally present in the polyene molecule. Both macrolides and polyenes are biosynthesized in the same way using identical building blocks. Macrolides represent a broad group of compounds, and new substances

o [

~

CH30

HO ~

0

o 1

HO

~OHcHzR O

HO ~---' NH2 daunorubicin (2), R = H doxorubicin (3), R = OH

FIG. 12. Anthracyclines.

126

VLADISLAVBI~HAL

have been frequently added to the list. Macrolides usually possess antibacterial activity, whereas polyenes are mostly fungicides. Erythromycins produced by Saccharopolyspora erythrea (Fig. 13), together with oleandomycin and picromycin, belong to the best-known 14-membered lactone ring macrolides (Harris et al., 1965). Macrolides with a 16-membered ring are represented by tylosin (Fig. 14) (Omura

CH 3 CH3 ' ~

0

HO

N(CH3)2

OR2

0

O

I -"~OR I CH 3

R, = Ra = H, R2 = CH 3, R4 = OH

erythromycin A

Ri = Ra = R4 = H, R2 = CH a

erythromycin

R I = R2 = R3 = H, R4 = OH

erythromycin C

Rt = R2 = Ra = R4 = H

erythromycin D

B

FIG.13. Erythromycins.

CH_ CH2R mycamlnose - mykarose

i

OH 0 myclnos -R = CHO

Tylosln

R = CH20H Relomycin

FIG.14. Tylosinand relomycin.

Streptomyces

BIOACTIVE PRODUCTS FROM

127

et al., 1975), which is produced by Streptomyces fradiae, as well as by leucomycin, spiramycin, and so on. Nystatin is the best-known polyene secondary metabolite (Fig. 15). Candicidin is another well-known secondary metabolite belonging to the polyene group. Its molecule includes p-aminoacetophenone as the terminal group. 4-amino benzoic acid (PABA) was identified as a precursor of the aromatic part of the candicidin molecule (Liu eta]., 1972; Martin, 1977). Synthesis of the lactone ring is similar to that observed in the case of other polyketides. In contrast to aromatics, pyruvate and butyrate units are more often used in biosynthesis instead of acetate units. The greatest difference, however, is the fact that, instead of aromatic rings, a lactone ring is formed. Keto- and methyl groups of the polyketide chain, from which macrolides are formed, are normally transformed more frequently. The sugars found in macrolide and polyene molecules are not usually encountered in microbial cells. They include both basic and neutral sugar molecules, and L-forms are often found. So far, at least 15 different sugars have been described in macrolides and polyenes. All of them are 6-deoxysugars; some are N-methylated, while others have a methyl group on either the oxygen or carbon atom. It has been repeatedly proven (Corcoran and Chick, 1966; Salah-Bey et al., 1998) that glucose

OH

CH3 H2 OH"

R = H

Nystatin

A~

CH 3 R =

Nystatin

Aa

FIG. 15. The nystatins.

I

128

VLADISLAV Bt~HAL

is primarily incorporated into macrolide sugar residues. Also in Streptomyces griseus, glucose, mannose, and galactose were incorporated to a greater extent into the mycosamine candicidin, as compared to its aglycone (Martin and Gi], 1979). The transformation of glucose to a corresponding sugar takes place in the form of nucleoside diphosphate derivatives, which is similar to the situation with other secondary metabolites. Avermectins consist of a 16-membered macrocyclic lactone to which the disaccharide oleandrose is bound (Fig. 16) (Burg et al., 1979; Miller et a]., 1979). Avermectins are produced by Streptomyces avermectilis. The macrocyclic ring of avermectins is synthesized, as are other polyketides, by producing a chain from acetate, propionate, and butyrate building blocks. Oleandrose (2,6-dideoxy-3-O-methylated hexose) is synthesized from glucose. The oleandrose moiety is methyl° ated before attachment of the sugar to the macrolide ring (Shulman and Ruby, 1987). Avermectins are potent antiparasitic compounds active against a broad spectrum of nematode and arthropod parasites, but lack antifungal or antibacterial activity. They bind to a specific high-affinity site present in nematodes but not in vertebrates. The dosage for animals and humans is extremely low. Ivermectin (22,23-dihydroavermectin B1) is a semisynthetic compound that is used to control internal and external

OCH3

H0~,..1~ .,L J .

OCH3

H3C4 "0" " 0 , , . . ( . ~

I

,CH

H

H3C#" -0-0,,,,.~0.,,.[.,,. -0.~= ~ CH3 °

FIG. 16. An example of an avermectin.

BIOACTIVE PRODUCTS FROM Streptomyces

129

parasites in animals and is the most potent anthelminthic compound. Avermectins are also employed in human medicine and for plant protection. Detailed reviews on the uses and biosynthesis of avermectins can be found in several monographs (e.g., MacNeil, 1995; Ikeda and Omura, 1995). Polyethers form a large group of structurally related natural products mainly produced by Streptomyces spp. (Birch and Robinson, 1995). They are potent coccidiostats (monensin, salinomycin) and are used in the agricultural arena (Westley, 1977). Polyethers are compounds possessing the ability to form lipid-soluble complexes that provide a vehicle for a wide variety of cations to traverse lipid barriers. This ion-bearing property led to their being named ionophores (Moore and Pressman, 1994). The backbones of polyethers are synthesized from acetate, propionate, and butyrate (monensin A) units. Isobutyrate and n-butyrate are efficiently incorporated into polyether antibiotics (Pospf~il et al., 1983). Incorporation of isobutyrate was explained by formal conversion of isobutyryl-CoA into N-butyryl-CoA or methylmalonyl-CoA by isobutyryl-CoA mutase and methylmalonyl-CoA mutase, respectively. C. OTHER BIOACTIVE PRODUCTS

Chloramphenicol (Fig. 17) is produced by Streptomyces venezuelae (Vining and Westlake, 1984). At present, however, the antibiotic is produced commercially using a fully synthetic process. In contrast to polyketides, the aromatic ring of the chloramphenicol molecule is synthesized from glucose via chorismic acid and p-amino benzoic acid. Chloramphenicol itself exhibits a broad spectrum of antibacterial activity, and it inhibits protein synthesis by binding to the 50S subunit of

Cl2-CH - CO

I

HN

I

CH:tOH C-H

I

H --C-OH

NO 2

FIG. 17. Chloramphenicol.

130

VLADISLAV BI~HAL

prokaryotic ribosomes to block the peptidyl transferase reaction (Pongs, 1979). Streptomycin (Fig. 18) is a well-known aminoglycoside antibiotic originally discovered by Schatz et al. (1944). It is synthesized by many streptomycetes, which produce a number of derivatives. The molecule of streptomycin consists of three components: streptidine, L-streptose, and N-methyl-L-glucosamine. None of these components has been found in the primary metabolism of microorganisms. The steps of streptomycin biosynthesis were disclosed by Walker and colleagues (Walker and Walker, 1971), who also studied the rele-

~H2 ~:= N H ;

NH 2 I .JL C = NH~

NH

OH

I

OH

0

\

v

t

R3,./' i OH

I

Streptomycin Dihydrostreptomycin N-Demethylstreptomycin Hydroxystreptomycin Manosidostreptomycin Manosidohydroxystreptom

R1

R2

R3

R4

CHO CH20H CHO CHO CHO CHO

CH3 CH~ CH3 CH20 H CH3 CH20H

CH3 CH 3 H CH 3 CH 3 CH~

H H H H mannose mannose

FIG. 18. Streptomycins.

BIOACTIVE PRODUCTS FROM Streptomycos

131

vant enzymes (Walker, 1975). The importance of streptomycin consists mainly in its ability to suppress Mycobacterium tuberculosis, resulting in effective suppression of tuberculosis. Bialaphos is formed from two L-alanine residues and the amino acid phosphinothricine. The latter compound is synthesized by streptomycetes from acetyl-CoA and phosphoenolpyruvate, and subsequently methylated using methionine as the methyl donor (Bayer et al., 1972). The producing microorganisms are Streptomyces hygroscopicus and

Streptomyces viridochromogenes. III. Genetics and Molecular Genetics A. PREPARATION OF HIGH-PRODUCTION MICROORGANISMS

Microorganisms that are isolated from nature (wild-type strains) usually produce small amounts of secondary metabolites. Sometimes during selection and subsequent cultivation in the laboratory, changes occur, making the cultivated strain nonidentical with the original strain. In such cases it should be noted that the term wild-type strain only refers to the fact that the strain did not undergo an "artificial" genetic change. In order for the commercial production of secondary metabolites to be profitable, higher levels of secondary metabolite synthesis are reached via genetic changes of the producers. Mutants are isolated by exposure of spores to UV irradiation, X-rays, 7-rays, o~-particles, or chemical mutagens (nitrogen mustards, N-methyl-N'-nitro-N-nitroso guanidine). Combined mutagenesis using various mutagens is often used. The surviving spores give rise to individual colonies, whose level of secondary metabolite production is tested; strains with higher yields are selected and subjected to further rounds of mutagenesis. Mutants that exhibit poor growth and sporulation ability are not suitable candidates for further improvement, even if their secondary metabolite production may exceed that of the original strain. Today's high-production strains, which synthesize as much as 10,000-fold levels of secondary metabolites compared to the original strains, are the result of many years of costly strain improvement. Unfortunately, these high-production strains sometimes can revert to lose their overproduction through spontaneous mutagenesis. When high-production strains are prepared by mutagenesis, a type of mutant that loses some of the structural genes can also be obtained. Such a mutant can exhibit a higher level of a secondary metabolite intermediate whose transformation stopped due to the absence of the corresponding enzyme. By crossing these mutants, some biosynthetic

132

VLADISLAVBI~HAL

pathways used to synthesize secondary metabolites were elucidated, for example, tetracyclines (McCormick et al., 1960). In some cases, the loss of the capability for secondary metabolite production in strains where extrachromosomal DNA was removed (e.g., by using acriflavine or ethidium bromide) suggested that the regulatory genes were located on plasmids (Hotta et al., 1977; Akagava et al., 1979; Ikeda et al., 1982). On the contrary, the loss of antibiotic productivity in Micromonospora rosario and Micromonospora purpurea treated by acriflavine is due to chromosomal mutation but not to loss of a plasmid (Kim et al., 1990).

B. GENETICMANIPULATIONOF SECONDARYMETABOLITEPRODUCERS The structural genes encoding the enzymes that synthesize secondary metabolites are located mostly on chromosomes. They are often organized in gene clusters (Binnie et al., 1989; Malpartida and Hopwood, 1984; Martin, 1992). A primary genetic resource for any organism is a genomic library, which can lead to construction of a set of ordered clones that includes the whole genome in manageable fragments and, eventually, to a complete DNA sequence. Resistance genes of the producer to its own products are usually located either at the beginning or at the end of a cluster, often in both positions (Ohnuki et al., 1985). In addition to the resistance and structural genes, regulatory genes are important in secondary metabolite production; however, their function is poorly understood. Structural genes for a number of secondary metabolites have been cloned into host microorganisms; genes for secondary metabolite resistance and other regulatory genes have also been cloned. Streptomyces coelicolor and S. lividans were developed as genetic models. A delightful summary of this work has been presented by Hopwood (1999). The polyketides comprise a large group of bioactive metabolites that are biosynthesized by decarboxylative condensation of activated simple carboxylic acids. This reaction is catalyzed by enzymes known as polyketide synthases. The structure and function of polyketide synthases in antibiotic producers was reviewed by Robinson (1991) and Bentley and Bennett (1999). As opposed to fatty acid synthase reactions, the polyketoacyl intermediates are not reduced but are differently modified. Polyketide synthases are classified into two types: type I and type II. They consist of multifunctional or monofunctional proteins, respectively, which are believed to act on covalently bound substrates

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attached by thioesthers to an acyl carrier protein domain or to a separate acyl carrier protein (Hopwood and Sherman, 1990). The properties of polyketide synthases have been deduced mostly from an analysis of DNA sequences of their cloned genes. The genes for erythromycin biosynthesis in Saccharopolyspora erythrea are organized in a series of functional modules. The enzymatic functions of this type I polyketide synthase are carried as a linear array of domains (Donadio eta]., 1991). Combinatorial biosynthesis involves interchanging secondary metabolism genes between antibiotic-producing microorganisms to create unusual gene combinations or hybrid genes. Novel metabolites can be made by both approaches. The method has been particularly successful with polyketide synthase genes. Derivatives of medically important macrolide antibiotics and unusual polycyclic compounds have been produced by combination of the type I and II polyketide synthase genes (Hutchinson, 1998). When polyketide synthase genes of microorganisms producing various polyketides were hybridized, a great deal of similarity of polyketide synthases from various streptomycetes was evidenced and new polyketides were constructed (Malpartida et al., 1987; Hopwood and Sherman, 1990; Khosla et al., 1993). When the gene encoding the acyl carrier protein of actinorhodin polyketide synthase of Streptomyces coelicolor A3 was replaced with homologues from the granaticin, oxytetracycline, tetracenomycin, and putative frenolicin polysynthase gene clusters, all of the replacements led to expression of functional synthases. Recombinant aromatic polyketides similar to actinorhodin or to shunt products by mutants defective in the actinorhodin pathway were synthesized (Khosla et al., 1993). These results reaffirm the idea that construction of hybrid polyketide synthases will be a useful approach for dissecting the molecular basis of the specificity of polyketide synthase-catalyzed reactions. IV. The Search for New Bioactive Secondary Metabolites

A. ISOLATION FROM NATURAL RESOURCES In spite of the fact that several thousand compounds isolated from microorganisms having some biological activity are known, new substances are still sought by major pharmaceutical companies. The probability of finding a new compound that would be usable as a new antibiotic or other biologically active compound is low, so a great number of microorganisms have to be screened. A rough estimate is that about 100,000 microorganisms are being screened each year for the

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presence of biologically active compounds. Modern screens are highly automated. The selection methods used, the targets, and the methods for detection of biological activity are usually not published. Preparation of a new biologically active compound and its introduction into clinical practice require the cooperation of scientists from various scientific disciplines and years of clinical trials. This effort can be subdivided into three parts (Yarbrough et al., 1993): 1. microbiology collection of source samples (soil) isolation of diverse microbes fermentation under different conditions to enhance diversity reproduce fermentation enhance the production for isolation taxonomy of the organism 2. molecular biology/pharmacology target selection screen design/implementation high-throughput screening identification of active compounds efficacy studies mechanism of action 3. chemistry active compound identification characterization/replication isolation/purification structure elucidation

B. PRODUCERS OF BIOACTIVE COMPOUNDS About 70% of the known bioactive substances are produced by streptomycetes, and the rest mainly by molds and nonfilamentous bacteria. With the increasing spectrum of efficiency of microbial metabolites, new nontraditional sources of such compounds have been tapped. These include microorganisms living under extreme conditions (e.g., high and low temperatures), sea-living microorganisms, and multicellular plants and animals. Another important source of new compounds are mutants of the producers of known active substances, for example, blocked mutants.

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C. SCREENING

The enterprise of screening microbial metabolites for new leads, first exploited by antibiotic researchers and today expanded to virtually all fields of therapeutic interest, has proven successful and will continue as an important avenue in the discovery of new drugs. The original method for determination of antibiotic efficiency consisted of application of test extract to wells made in an agar medium layer in Petri dishes, into which the sensitive (target) microorganism was inoculated. Most often, Staphylococcus aureus, Sarcina lutea, Klebsiella pneumoniae, Salmonella gallinarium, Pseudomonas spp., Bacillus subtilis, and Candida albicans were used. When a compound with an antibiotic activity toward the testing microorganism was put into a well, it diffused through the agar medium and a halo was formed around the well, as a result of suppressed microorganism growth. This classic plate assay has been modified and improved in many ways. The tests of other biological activity require different and frequently sophisticated methods. This is especially true when enzyme inhibitors are the subject. Thus, Ogawara et al. (1986) chose a tyrosine protein kinase associated with malignant transformation of cells caused by retroviruses as the target in a biochemical screen. They found genistein, an isoflavone from a Pseudomonas sp., exhibiting a specific inhibitory activity. Production of target enzymes using recombinant DNA methodology has dramatically expanded the number of potential targets that can be feasibly screened. A screen for inhibitors of HIV reverse transcriptase is an example. The enzyme was produced in Escherichia coli, purified by affinity chromatography, and used to test natural products for the activity (Tak6 et al., 1989).

D. SEMISYNTHETICAND SYNTHETIC BIOACTIVE PRODUCTS Natural products can be modified in various ways. The nonspecificity of biosynthetic enzyme systems facilitates synthesis of certain secondary metabolites through addition of selected precursors to the growth medium. Thus, the reaction equilibrium can be shifted to promote production of the derivative required, for example, preparation of penicillins with different sidechains. The individual derivatives of penicillin and cephalosporin have slightly different antimicrobial spectra and are active against microorganisms resistant to other derivatives. The structure of polypeptide antibiotics can also be modified by addition of amino acids to the growing culture.

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Replacement of a part of the metabolite molecule can be accomplished chemically or enzymatically. In this way, semisynthetic penicillins, cephalosporins, tetracyclines, and other antibiotics can be prepared. Production of semisynthetic penicillins and cephalosporins is facilitated by the fact that 6-amino penicillanic and 7-amino cephalosporanic acids are easily prepared. The sidechain is removed by the action of an enzyme or by chemical hydrolysis (Fig. 19), then another acyl is bound chemically or enzymatically to the amino group in position 6 (penicillins) or 7 (cephalosporins). Glycylcyclines, semisynthetic tetracyclines, N,N-dimethylglycylamido derivatives of minocycline, and 6-demethyl-6-deoxytetracycline were significantly more active against strains resistant to minocycline and tetracycline. These compounds show potent activity against a broad spectrum of Gram-positive and Gram-negative bacteria, including strains that carry the major tetracycline determinants, efflux and ribosomal protection (Eliopoulos et al., 1994; Testa et al., 1993; Sum et al., 1998). Much research has been devoted to synthesizing derivatives or analogues of macrolides with improved chemical, biological, and pharmacological properties. New 14-membered macrolide derivatives of erythromycin A have shown improved pharmacokinetics due to modifications of the hydroxyl group at C-6, a proton at C-8, or a ketone at C-9. In addition, a new 15-membered macrolide, azithromycin, shows good activity against Gram-positive bacteria (Mazzei et al., 1993).

O~

N

~

CH 5 CH 3 COOH

6-aminopenicillanicacid HOOC-

NH

I

7-aminocephalosporanicacid

J~'rS

COOH

FIG. 19. 6-amino penicillanic acid and 7-amino cephalosporanic acid.

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As the majority of bioactive products have rather complex structures, their chemical synthesis is generally more expensive than their production by fermentation. An exception to the rule seems to be chloramphenicol, which is normally prepared using chemical synthesis. E. HYBRID BIOACTIVE PRODUCTS AND COMBINATORIALBIOSYNTHESIS

Genetic engineering methods have recently advanced so much that now we can suitably combine structural genes of two or even more bioactive secondary metabolite producers. If these genes are expressed, hybrid bioactive products are synthesized that cannot be found in nature (Hutchinson, 1987, 1988; Tomich, 1988; Hopwood, 1993). Hopwood et a]. (1985, 1986) used this method with the genes of actinorhodin synthesis and obtained related hybrid macrolides, mederrhodin A and B, dihydromederrhodin A, and dihydrogranatirhodin (see Omura et al., 1986). New anthracyclines were produced when a DNA segment was cloned from Streptomyces purpurascens ATCC 25489 close to a region that hybridized to a probe containing part of the actinorhodin polyketide synthase from Streptomyces galilaeus ATCC 31615 (Niemi et al., 1994). Exogenous addition of designed synthetic molecules to cultures of a mutant of Saccharopolyspora erythrea that was not able to synthesize 6-deoxyerythronolide B, the macrocyclic precursor of erythromycin, resulted in highly selective multimilligram production of unnatural polyketides (Jacobsen et al., 1997). This illustrates the catalytic versatility of modular polyketide synthases.

V. Regulation of Secondary Metabolite Production A. GROWTH PHASES OF STREPTOMYCETES

In cultures of streptomycetes capable of secondary metabolite production, several growth phases representing different physiological states can be distinguished: 1. Preparatoryphase (lag phase): The rate of biomass increase is low; the culture is adapting to its new environment. 2. Growth phase (the term "logarithmic phase" is not suitable for most Streptomyces spp. since their growth curves are not exponential functions): Intensive growth is taking place, accompanied by low secondary metabolite synthesis. This phase is roughly equivalent to the "trophophase."

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3. Transition phase: Characterized by a decreased growth rate; secondary metabolite production is begun. The enzymes of secondary metabolism are synthesized (B~hal, 1986a,b) and protein synthesis is slowed down. 4. Production phase: Characterized by a significant reduction in the growth rate (sometimes growth even ceases), a negligible change in biomass concentration, and intensive synthesis of secondary metabolite. This phase is sometimes called the "idiophase." Producers of secondary metabolites are mostly filamentous bacteria or fungi, which means that in their culture cells of various age and at different stages of development are present. The microorganisms grow in pellets, inside which growth conditions differ from those on the pellet surface (e.g., nutrient concentrations, oxygen concentration). An increase in dry weight does not always correlate with an increase in growth, since in streptomycetes a thickening of the cell wall or glycocalyx formation often occurs that increases the dry weight without increasing the number of cells (Vo~f~ek et al., 1983). Since individual cells in a fermentation can be at different stages of development (i.e., in different physiological states), the physiological state of the whole culture represents an average of the physiological states of individual cells. B. CONTROLOF FERMENTATIONBY BASIC NUTRIENTS

Sufficient biomass is required to reach a high yield of a secondary metabolite. Moreover, the danger of contamination is diminished and the economic parameters of the fermentation device are optimal if growth is rapid. For this reason, readily utilizable sources of carbon, nitrogen, and phosphorus are used. However, production of a secondary metabolite does not usually take place until one or more nutrients become limited. Therefore, the culture medium should be designed in such a way that, after the biomass has increased sufficiently, at least one nutrient source will become depleted. The carbon source, nitrogen source, and phosphate limitation all have been described as important triggers in different systems. Many secondary metabolites are produced in fed-batch systems, that is, a certain amount of the culture medium is inoculated with the producing microorganism, and, after a certain time interval, another dose of nutrients is added to the fermentor. Thus, prolonged cultivation can be accomplished that enables one to increase the yield of secondary metabolite. The inflow of nutrients makes it possible to maintain opti-

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mal levels. A depiction of production cultivation of Streptomyces aureofaciens is given in Figure 20 (B~hal, 1987). In cultivations whose course is well known, nutrient inflow is programmed in advance. The inhibition of penicillin synthesis by glucose was observed shortly after its discovery in media containing glucose and lactose (Demain, 1974). The antibiotic was found to be synthesized only after glucose was depleted from the medium and lactose began to be metabolized. Similarly, glycerin was observed to inhibit biosynthesis of cephalosporins (Demain, 1983). Using this information, fermentation protocols were worked out in which the level of glucose was kept low so as not to inhibit antibiotic production. The mechanism of inhibition by readily utilizable sugars of the synthesis of secondary metabolites is probably due to repression of the enzymes of secondary metabolism (Revilla eta]., 1986; Erban et al., 1983).

A

B

:~

pH

~" NH3. N

. i "_ I_ SUCROSE

()

:~0

i0

60

80

100

120

HOURS

FIG. 20. Parameters of industrial fermentation of S. aureofaciens: (1) chlortetracycline production (g/liter), (2) ATC oxygenase (pkat/mg proteins x 2), (3) NH3-nitrogen (g/liter x 0.1), (4) sucrose (g/liter x 10), (5) pH. Ammonium supplements were added at points A and B.

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Readily utilizable nitrogen sources can also negatively influence production of secondary metabolites. A m m o n i u m ions often decrease secondary metabolite synthesis, and, therefore, their concentration in production media is limited, while soy flour, peanut flour, and other substances are preferred nitrogen sources. These latter nitrogen sources are more similar to those used in nature by the microorganisms producing secondary metabolites. Readily utilizable nitrogen sources repress enzymes of secondary metabolism in Cephalosporium acremonium (Shen et al., 1984) during biosynthesis of cephalosporin and in Streptomyces clavuligerus producing cephamycin (Demain and Brafm, 1986). Similarly, inhibition of biosynthesis of leucomycin (Omura et al., 1980a), tylosin (Omura et al., 1980b), and erythromycin (Flores and S~inches, 1985) are explained by repression of the enzymes of secondary metabolism. A m m o n i u m salts also inhibit the activity of anhydrotetracycline oxygenase isolated from S. aureofaciens (B~hal et al., 1983). The overproduction of most secondary metabolites can only be achieved if phosphate is limited. Inorganic phosphate has to be added carefully in doses to the m e d i u m to achieve an optimal ratio between biomass production and secondary metabolite biosynthesis. When bound to organic compounds normally added to the medium (e.g., soya flour), phosphate does not affect secondary metabolite production. In general, secondary metabolite biosynthesis is started when the concentration of phosphate decreases below a certain level. At this point, the producer culture undergoes a shift from the physiological state characteristic for the growth phase to that of the overproduction phase. Inorganic phosphate also causes repression of the synthesis of enzymes of secondary metabolism (Behal et al., 1979b; Madry and Pape; 1981, Martin et al., 1981). After phosphate was depleted from the medium, a significant decrease in the rate of protein synthesis was observed during tetracycline biosynthesis (Behal, 1982). If phosphate was kept above the threshold concentration, the decrease in the rate of protein synthesis did not occur and enzymes of secondary metabolism were not synthesized. Addition of phosphate to the medium at the beginning of the production phase, after the phosphorus source was depleted and the enzymes of secondary metabolism synthesis initiated, resulted in a decrease in the levels of the enzymes of secondary metabolism in the culture and acceleration of protein synthesis. McDowall et al. (1999) proved that production of oxytetracycline by Streptomyces rimosus is controlled, at least in part, at the level of transcription from promoters overlapped by tandem repeats similar to those of the DNAbinding sites of the OmpR family.

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C. H o w SIGNALS FROM THE MEDIUM ARE RECEIVED

Reception of signals from the environment, which result in initiation of secondary metabolite synthesis, does not significantly differ from transduction of signals for other metabolic processes. Catabolite repression signals or those signaling the depletion of nitrogen or phosphate, or initiation of sporulation, are transduced via two-component signal proteins (Doull and Vining, 1995). With some structural variation, these proteins are characterized by common mechanistic features and conserved amino-acid sequences. The two-component system consists of a cytoplasmic membrane-linked sensor-transmitter protein and a response-regulator protein, located in the cytoplasm. The sensor-transmitter is composed of a sensor domain located near its N terminal; the N terminal is found outside the cytoplasm. A specific effector is capable of binding directly to this N terminal. The transmitter domain is located in the cytoplasm to be linked to the sensor domain via a hydrophobic amino-acid sequence stretching across the membrane. The sensortransmitter proteins are histidine-protein kinases, capable of autophosphorylation at their C termini on receiving the proper signal. The phosphorylated protein becomes a donor in reactions transferring phosphorus. The acceptor is the cytoplasmic response-regulator protein. Two-component signal proteins thus transfer information concerning the conditions that can affect cell action. D. REGULATION BY Low-MOLECULAR-WEIGHT COMPOUNDS The expression of structural genes is also regulated by some low-molecular-weight compounds. The mechanism of their action is not understood. For example, tryptophan exhibited a stimulatory effect on the production of mucidin in the basidiomycete Oudemansiella mucida (Nerud et al., 1984) and actinomycin in Streptomyces parvulus (Troast et al., 1980). Methionine was found to promote synthesis of cephalosporin C. Neither tryptophan nor methionine are used as building blocks for these metabolites. Benzyl thiocyanate is one of the low-molecular-weight compounds that affect chlortetracycline biosynthesis. It increases production of both chlortetracycline and tetracycline in S. aureofaciens, although it does not influence production of oxytetracycline in S. rimosus. The effect on the metabolism of S. aureofaciens is multiple (Novotnfi et al., 1995), including effects on a number of enzymes, including the enzymes of secondary metabolism (B~hal et al., 1982). Benzyl thiocyanate is able to raise the level of secondary metabolite production only if it

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is added in the lag phase, in the growth phase, or at the beginning of the production phase. Its effect is more pronounced in low-production strains, where the enzyme level and chlortetracycline production are increased 10- to 2O-fold, as compared to high-production strains where the increase is only twofold.

E. AUTOREGULATORS

Low-molecular-weight diffusible compounds from streptomycetes have been discovered that regulate the metabolism of the producing strain (Horinuchi and Beppu, 1990, 1992, 1995). The best known is factor A, 7-butyrolactone (Fig. 21), which was discovered in Streptomyces griseus (Khokhlov, 1982). A nonproducing strain started the synthesis of streptomycin after factor A was added to the culture simultaneously; the culture formed aerial mycelium. Factor A is synthesized by many streptomycetes, but the regulatory effect was observed only in Streptomyces griseus, Streptomyces bikiniensis, and Streptomyces actuosus. The addition of factor A to blocked mutants of Streptomyces griseus JA 5142 caused resumption of synthesis of anthracyclines and leukaemomycin (anthracycline-type antibiotic) (Graefe et al., 1982). Resistance to streptomycin linked with enzymatic phosphorylation of the antibiotic was also induced by factor A (Hara and Beppu, 1982).

NH2

R

0

OH

I

Factor A

o=P,-O Factor B FIG. 21. Factor A and factor B.

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Analogues of factor A have also been found, all of them being 7-butyrolactones. Virginiae butanolides were detected in Streptomyces virginiae (Yanagimoto et al., 1979). Factor I was isolated from Streptomyces sp. FR1-5, and its effective concentration was 0.6 ng/ml culture. Most of the factor A analogues, however, were not biologically active. Factor B was isolated from the yeast Saccharomyces cerevisiae. This substance was capable of eliciting production of rifamycin in a blocked mutant of a Nocardia species (Fig. 21) (Kawaguchi et al., 1984). Factor B was effective at a concentration of 10 -8 M, with one molecule eliciting a synthesis of about 1500 molecules of the rifamycin. The structure of factor B is similar to cAMP, but none of the derivatives of known nucleotides exhibited a comparable effect. Chemically prepared derivatives of factor B have also been tested. Activity was observed with those that contained a C2-C12 acyl moiety; octylester was the most effective (Kawaguchi et al., 1984). A substitution of guanosine for adenine did not result in a loss of the biological activity of factor B. Factor C was isolated from the fermentation medium of Streptomyces griseus. This compound causes cytodifferentiation of nondifferentiating mutants (Szabo et al., 1967). Factor C is a protein having a molecular weight of about 34,500 daltons, and it is rich in hydrophobic amino acids. The effect of autoregulators is easily observable if they elicit morphological changes such as the formation of aerial mycelia. Carbazomycinal and 6-methoxycarbazomycinal, isolated from Streptoverticillium spp., inhibit aerial mycelium formation at a concentration of 0.5 to 1 pg/ml. Autoregulators affecting sporulation were found in Streptomyces venezue]ae (Scribner et al., 1973), Streptomyces avermectilis (Nov~ik et al., 1992), and Streptomyces viridochromogenes NRRL B-1551 (Hirsch and Ensign, 1978). From the same strain of S. viridochromogenes, germicidin was isolated by Petersen and coworkers (1993). The compound had an inhibitory effect on the germination of arthrospores of S. viridochromogenes at a concentration as low as 40 pg/ml. Germicidin (6-(2butyl)-3-ethyl-4-hydroxy-2-pyrone) is the first known autoregulative inhibitor of spore germination in the genus Streptomyces and was isolated from the supernatant of germinated spores and also from the supernatant of a submerged culture. Mutants of Streptomyces cinnamonensis resistant to high concentrations of butyrate and isobutyrate produce an anti-isobutyrate (AIB) factor that is excreted into the culture medium (Pospfgil, 1991). On plates, AIB factor efficiently counteracted toxic concentrations of is-

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obutyrate, acetate, propionate, butyrate, 2-methylbutyrate, valerate, and isovalerate in S. cinnamonensis and other Streptomyces species.

E REGULATIONBY PHOSPHORYLATEDNUCLEOTIDES

Global control mechanisms for secondary metabolite biosynthesis have been investigated. The energetic state of the cell is thought to be such a general control mechanism. The intracellular ATP level reflects the content of free energy in the cell. In some cases, the start of secondary metabolite synthesis is linked with a decrease in intracellular ATP level. Such a relationship was observed in Streptomyces aureofaciens and Streptomyces fradiae during production of tetracycline (Janglovfi et al., 1969; Curdovfi et al., 1976) and tylosin (Vu-Trong et al., 1980). Even though the regulatory role of ATP cannot be strictly excluded, the results seem to support a hypothesis that a higher ATP level accompanies active primary metabolism. A slowdown in growth and primary metabolism is accompanied by a decrease in the ATP level. The role of cAMP in the metabolism of secondary metabolite producers was also studied, especially in connection with glucose regulation. To date, no indication has been obtained suggesting a specific role of cAMP in regulation of secondary metabolite production (Chatterjee and Vining, 1981).

G. REGULATIONBY METAL IONS

Metal ions act as a part of enzyme active centers. The optimal concentrations of metal ions for cultivation of secondary metabolite-producing strains have usually been determined empirically. In complex media it is generally not necessary to add specific metal ions; however, in defined media their presence is essential.

Vl. Resistance to Bioactive Products

Resistance against bioactive products has been studied mainly in antibiotic producers. Antibiotic resistance is usually looked at from two angles: first, emergence of drug-resistant strains, and, second, "self-resistance" of antibiotic-producing strains. The ways in which these two types of resistance are achieved is often similar.

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A. RESISTANCE OF SECONDARY METABOLITE PRODUCERS

The basic metabolic processes of wild-type secondary-metabolite producing microorganisms are not inhibited if the secondary metabolites are synthesized at low concentrations. After strain improvement, strains with 100- to lO00-fold increases in secondary metabolite yields have been isolated. Genome changes of the improved strains include a number of deletions and amplifications in chromosomal DNA, as well as changes in extrachromosomal DNA. Low-production strains, whose resistance to their own product is low (i.e., higher concentrations of the product inhibit their growth) regulate their secondary metabolite production by inhibiting the enzyme activities that participate in synthesis of the secondary metabolite. In highproduction strains, such controls are lost and the strains have to find a way to survive in the presence of a high concentration of the antibiotic (Vining, 1979). The genes for self-resistance are often located at the beginning of the cluster of structural genes. As a result, they are expressed simultaneously with the structural genes. The genes of newly gained resistances, however, are mostly located on plasmids. Some antibiotics function by attacking active centers of enzymes. However, if the active center is modified, the antibiotic cannot bind to it and then resistance comes into existence. It is not known whether a decreased ability to bind the secondary metabolites results from posttranslational modification of the active center or if resistant molecules of the enzyme are synthesized de novo. To date, there is no clear evidence to support the latter situation. Many antibiotics inhibit protein synthesis, the target site being at the ribosome level. Often, the functions of Tu and G elongation factors are also impaired, together with reduced synthesis of guanosine penta- and tetraphosphates (Weiser eta]., 1981). The antibiotic producers (mostly Streptomyces spp.), as well as the bacteria against which the antibiotic is used, protect themselves by posttranscriptional modification of rRNA. Adenine is methylated to obtain N6-dimethyladenine rRNA in the 23S subunit. Such modified ribosomes do not bind the antibiotic. In other cases, adenine is methylated to yield 2-O-methyladenosine (Cundliffe and Thompson, 1979; Mikul/k et al., 1983; Thompson et al., 1982). However, methylation-modified ribosomes can be sensitive to the effect of other antibiotics. The genes coding for methylases that catalyze methylation of adenine in some streptomycetes were cloned into Streptomyces lividans, and the ribosomes of the mutants prepared

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were resistant to the corresponding antibiotics (Calcutt and Cundliffe, 1990; Kojic eta]., 1992). The most important mechanism of resistance observed in secondary metabolite producers seems to be export from the cell into the environment. In Streptomyces rimosus, an oxytetracycline producer, genes for the enzymes increasing the antibiotic transport rate precede the structural genes on the chromosome. Genes for the resistance, consisting in protection of ribosomes via synthesis of an unidentified protein, are located at the end of the structural gene cluster (Ohnuki et al., 1985). Producers of bioactive secondary metabolites also have to solve the problem of reverse flow of products into the cell. Some secondary metabolites are bound to the cell wall; others are complexed in the medium (tetracyclines in the presence of Ca2+ ions). Cytoplasmic membranes of resistant strains are often less sensitive to the effect of secondary metabolites. This kind of resistance is thought to be connected to the content of phospholipids in the cell. Secondary metabolite producers can use several types of resistance simultaneously. Tetracyclines, which strongly inhibit protein synthesis, interfere with binding of the ternary complex amino acyl-tRNA-EFTuGTP to ribosomes (Gavrilova et al., 1976). The genes for resistance were cloned from Streptomyces rimosus into Streptomyces griseus, sensitive to tetracyclines, using pOA15 as the vector plasmid (Ohnuki et al., 1985). After mapping the plasmids in resistant strains using restriction nucleases, two types of plasmids capable of transfer of different types of resistance were found. One type consisted of an increased ability of tetracycline transport to the medium, and the other in an increased resistance of ribosomes to the effect of tetracyclines. These ribosomes bore a compound(s), bound to their surface, that could be removed by washing with 1 M NH4C1 solution. The ribosomes lost their resistance after the washing, which was demonstrated with both the ribosomes of S. griseus and those of the original strain of S. rimosus. The two types of resistance were both constitutive and inducible. The inhibiting concentrations of chlortetracycline in Streptomyces aureofaciens are higher in the production phase as compared to the growth phase (Behal et a]., 1979a). Thus, the resistance can be increased even during the fermentation process. Resistance to oxytetracycline of S. rimosus increases in proportion to production (McDowall et al., 1999). Another way secondary metabolite producers can avoid the effect of their products is to situate the distal enzymes of the secondary metabolite biosynthetic pathway (synthases) outside the cell, most often in the periplasm. In Streptomyces aureofaciens, a higher proportion of the

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terminal enzyme of tetracycline synthase was found under high-production conditions in the periplasm, as compared to low-production conditions (Erban et al., 1985). B. RESISTANCE IN PATHOGENIC MICROORGANISMS

Shortly after antibiotics were introduced into clinical practice on a massive scale, strains of hitherto-sensitive microorganisms started to appear. These resistant strains required the use of m u c h higher antibiotic concentrations or proved completely resistant to these antibiotics. The resistant strains originated from clones that survived the antibiotic treatment, especially if the treatment was terminated before all pathogenic microorganisms were killed or the antibiotic was applied at sublethal doses. There are several ways in which microorganisms can gain resistance (Ogawara, 1981). These include: 1. creation of an alternative metabolic pathway producing a compound whose biosynthesis is blocked by the bioactive metabolite 2. production of a metabolite that can antagonize the inhibitory effects of the bioactive metabolite 3. an increase in the amount of the enzyme inhibited by the secondary metabolite 4. a decrease in the cell's metabolic requirement for the reaction inhibited by the secondary metabolite 5. detoxification or inactivation of the bioactive metabolite 6. a change in the target site 7. blocking of the transport of the bioactive metabolite into the cell In most resistant microorganisms, the mechanisms of resistance in items 5, 6, and 7 are encountered. Penicillins and cephalosporins are degraded in three ways: (1) by the enzyme penicillin amidase, which cleaves the amidic bond by which the sidechain is bound to the [3-1actam ring, (2) by the enzyme acetyl esterase that hydrolyzes the acetyl group at C-3 on the dihydrazine ring of cephalosporins, and (3) by the enzyme ~-lactamase, which catalyzes hydrolysis of the ~-lactam ring of penicillins and cephalosporins. Penicillin amidases are rarely used by microorganisms to build up resistance to [~-lactam antibiotics; however, these enzymes are often employed for the synthesis of semisynthetic antibiotics. Acetyl esterase also is not important from the point of view of antibiotic resistance. In

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most cases, ~-lactams are inactivated by 15-1actamase, which destroys one of the important sites for their antibiotic activity; the damage is irreversible. However, [3-1actamases are synthesized not only by microorganisms that came into contact with penicillins; constitutive synthesis of these enzymes have been found in three quarters of all Streptomyces strains. One can suppose that the genes for synthesis of ~-lactamases were transferred horizontally. Recent studies indicate frequent and promiscuous gene transfer even between distantly related bacterial species. A possibility of direct transfer from a streptomycete to a pseudomonad, for example, may seem unlikely. However, it is not necessary to invoke direct exchanges. It is more reasonable to imagine that exchanges between distantly related organisms result from a cascade of transfer between related species (Davis, 1992). Another way of inactivating a bioactive metabolite molecule is N-acetylation of the amino group or O-phosphorylation of the hydroxyl. Bialaphos was found to be inactivated by acetylation. The substance itself is not toxic, but phosphinothricine is liberated in the cell, which inhibits the glutamine synthetases, key enzymes of the inorganic nitrogen assimilation pathway (Donn et al., 1984).

Acknowledgments I thank Joan W. Bennett (Tulane University) for her critical reading of an early version of the manuscript. This work was supported by the Grant Agency of the Czech Republic (grant no. 203/98/0421)

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Salah-Bey, K., Doumith, M., Michel, J. M., Haydock, S., Cortes, J., Leadlay, P. F., and Raynal, M. C. (1998). Targeted gene inactivation for the elucidation of deoxysugar biosynthesis in the erythromycin producer Saccharopolyspora erythraea. Mol. Gen. Genet. 257, 542-553. Schatz, A., Bugle, E., and Waksman, S. A. (1944). Streptomycin, a substance exhibiting antibiotic activity against Gram-positive and Gram-negative bacteria. Proc. Soc. E~Yp. Biol. Med. 55, 66-69. Schlumbohm, W., Vater, J., and Kleinkauf, H. (1985). Reactive sulfhydryl groups involved in the aminoacyl adenylate activation reactions of the gramicidin S synthetase. Biol. Chem. Hoppe Sey]er 366, 925-930. Scribner, H. E., Tang, T., and Bradley, S. G. (1973). Production of a sporulation pigment by Streptomyces venezuelae. App]. Microbiol. 25,873-879. Sekiguchi, R. (1983). The biosynthesis of mycotoxin patulin. Hakkokogaku 61,129-137. Sekiguchi, J., Shimato, T., Yamada, Y., and Gaucher, G. M. (1983). Patulin biosynthesis: Enzymatic and nonenzymatic transformations of the mycotoxin (E)-Ascladiol. Appl. Environ. Microbiol. 45, 1939-1942. Shen, Y. C., Hein, J., Solomon, N. A., Wolfe, S., and Demain, A. L. (1984). Repression of [5-1actam production in Cephalosporium acremonium by nitrogen sources. J. Antibiot. 37, 503-512. Shulman, M. D., and Ruby, C. (1987). Methylation of demethylavermectins. Antimicrob. Agents Chemother. 31,964-965. Sum, F. J., Sum, F. W., and Projan, S, J. (1998). Recent development in tetracycline antibiotics. Curt. Pharm. Res. 4, 119-132. Szab6, G., Bekeshi, I., and Vitalis, S. (1967). Mode of action of factor C, a substance of regulatory function in cytodifferentiation. Biochem. Biophys. Acta 145, 159-165. Tak6, Y., Inouye, Y., Nakamura, S., Allaudeen, H. S., and Kubo, A. (1989). Comparative studies of the inhibitory properties of antibiotics on human immunodeficiency virus reverse transcriptase and cellular DNA polymerases. J. Antibiot. 42,107-115. Tanaka, H., Kominato, K., Yamamoto, R., Yoshika, T., Nishida, H., Tone, H., and Okamoto, R. (1994). Synthesis of doxorubicin-cyclodextrin conjugates. J. Antibiot. 47, 10251029. Testa, R. T., Petersen, P. J., Jacobus, N. V., Sum, P. E. Lee, V. J., and Tally, F. P. (1993). In vitro and in vivo antibacterial activities of the glycylcyclines, a new class of semisynthetic tetracyclines. Antimicrob. Agents Chemothez: 37, 2270-2277. Thompson, J., Cundliffe E., and Stark, M. J. R. (1982). The mode of action of berninamycin and mechanism of resistance in producing organism Streptomyces bernesis. J. Gen. Microbiol. 128, 875-884. Tomich, P. K. (1988). Streptomyces cloning: Possible construction af novel compounds and regulation of antibiotic biosynthesis genes. Antimicrob. Agents Chemother. 32, 1472-1476. Tomoda, H., and Omura, S. (1990). New strategy for discovery of enzymes inhibitors: Screening with intact mammalian cell or intact microorganisms having special functions. J. Antibiot. 43, 1207-1222. Troast, T., Hitchcock, M. J. M., and Katz, E. (1980). Distinct kinureninase and hydroxykinurinase enzymes in an actinomycin-producing strain of Streptomyces paravulus. Biochem. Biophys. Acta 612, 97-106. Umezawa, K., Aoyagi, T., Suda, D., Hamada, M., and Takeuchi, T. (1976). Bestatin, an inhibitor of aminopeptidase B, produced by actinomycetes. J. Antibiot. 30, 170-173. Vining, L. C. (1979). Antibiotic tolerance in producer organisms. Adv. Appl. Microbiol. 25, 147-168.

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Vining, L. C., and Westlake, D. W. S. (1984). Chloramphenicol. In "Biotechnology of Industrial Antibiotics" (E. J. Vandamme, ed.). pp. 387-411. Dekker, New York. von Doehren, H. (1995). Peptides, In "Genetics and Biochemistry of Antibiotics Production" (L. C. Vining and C. Stuttard, eds.), pp. 129-171. Butterworth-Heinemann, Boston. Vo[f]ek, J., Curdov~, E., Jechov~, V., Lenc, B., and Hogt~lek, Z. (1983). Electron-cytochemical demonstration of polyphosphates and the appropriate phosphates in the glycocalyx of Streptomyces aureofaciens. Curt. Microbio]. 8, 31-36. Vu-Trong, K., Bhuwapathanapun, S., and Gray, P. P. (1980). Metabolic regulation in tylosin-producing Streptomycesfradiae: Regulatory role of adenylate nucleotide pool and enzyme involved in biosynthesis of tylonolide precursors. Antimicrob. Agents Chemother. 17, 519-525. Walker, J. B. (1975). ATP: Streptomycin 6-phosphotransferase. Methods Enzymol. 43, 428-470. Walker, M. S., and Walker, J. B. (1971). Streptomycin biosynthesis. J. Biol. Chem. 246, 7034-7040. Weiser, J., Mikulfk, K., and Bosh, L. (1981). Studies on the elongation factor Tu from Streptomyces aureofaciens. Biochem. Biophys. Res. Commun. 99, 16-20. Westley, J. (1977). Polyether antibiotics: Versatile carboxylic acid ionophores by Streptomyces. Adv. App]. Microbio]. 22, 177-223. Williams, D. H., Stone, M. J., Mortishire-Smith, R. J., and Hauck, P. R. (1990). Molecular recognition by secondary metabolites. Biochem. Pharmacol. 40, 27-34. Yanagimoto, M. Yamada, Y., and Terui, G. (1979). Physiological study on the production of staphylomycin, 3: Extraction and purification of inducing material produced in staphylomycin fermentation. Hakko Kogeku Zassi 57, 6-19. Yarbrough, G. G, Taylor, D. P., Rowlands, R. T., Crawford, M. S., and Lasure, L. L. (1993). Screening microbial metabolites for new drugs-theoretical and practical issues. J. Antibiot. 46, 535-544. Zmijewski Jr., M. J., and Fayerman, J. T. (1995). Glycopeptides. In "Genetics and Biochemistry of Antibiotics Production" (L. C. Vining and C. Stuttard, eds.), pp. 269281. Butterworth-Heinemann, Boston.

Advances in Phytase Research EDWARD J. MULLANEY, CATHERINE B. DALY,

AND ABUL H. J. ULLAH Southern Regional Research Center Agricultural Research Service United States Department of Agriculture New Orleans, Louisiana 70124

I. Introduction A. Phytase Today B. Phytic Acid in Plants C. Increased Use of Plant Meals in Animal Feed D. Early Research Identifies a Fungal Phytase II. Phytases That Are Histidine Acid Phosphatases (HAPs) A. PhyA B. PhyB C. E. coli HAP Phytase D. Yeast HAP Phytase E. Plants III. Phytases with an Undefined Active Site A. Bacillus Phytase (Phytase C) B. Klebsiella Phytase C. Yeast D. Plants and Microbes IV. Increased Phosphorus Levels in Our Environment Creates Need for Phytase V. Engineering Phytase A. Heat Tolerance B. Temperature and pH Optima C. Substrate Specificity D. Enzyme Stability E. Synergistic Effect VI. Enzyme Production A. Filamentous Fungi B. Expression in Yeast C. Expression in Plants D. Transgenic Animals VII. Expanding Uses of Phytase A. Potential in Aquaculture B. Phytase as a Soil Amendment C. Production of myo-Inositol Phosphates D. Semisynthesis of Peroxidase VIII. Occupational Health Concerns IX. Future Prospects A. Phosphorus for Future Generations B. Phytase's Role in Our Future X. Summary References 157 A D V A N C E S IN APPLIED MICROBIOLOGY, V O L U M E 47 006S-2164/00 $25.00

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I. Introduction

A. PHYTASE TODAY At the close of the twentieth century, annual sales of phytase as an animal feed additive were estimated to be $500 million and growing (Abelson, 1999). Evolution of the market for this feed additive can be attributed to a chain of events during the late twentieth century that both created the need for this enzyme and provided the means for its commercial development. Earlier reviews on phytase (Wodzinski and Ullah, 1996; Dvorakova, 1998) have chronicled the events since 1907, when Suzuki et al. (1907) first discovered phytase, up to its commercialization in 1994. Since then, both the use of, and the research on, phytase have expanded considerably. A search of the scientific literature for the period 1992-98 for studies involving phytase research (Fig. 1) demonstrates the recent increased interest in this enzyme. An examination of current phytase research and its relationship to a contemporary environmental concern provides insight into how a convergence of technologies fosters additional research and development on this key biocatalyst.

140

120

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0 1992

1993

1994

1995

1996

1997

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FIG. 1. Number of published studies pertaining to phytase each year for the period 1992-98. The total for each year was obtained by a search of PubMed, Food and Science Technology Abstracts (FSTA), Agricola, CAB abstracts, and Biological Abstracts for research involving phytase.

ADVANCES IN PHYTASE RESEARCH

159

B. PHYTIC ACID IN PLANTS

Phytic acid (myo-inositol 1,2,3,4,5,6-hexakis dihydrogen phosphate) is the primary storage form of phosphorus and the source of inositol in plant seeds (Reddy et al., 1982). Studies on the structure of phytic acid (phytate) and the amounts of phytin phosphorus in various feedstuffs obtained from various plant meals have been published (Nelson et al., 1968a; Reddy et al., 1982; Wodzinski and Ullah, 1996). The storage function of phytase may be a part of a larger regulatory function that enables plants to control their phosphate and mineral concentrations (Raboy and Gerbasi, 1996). In addition to its role in phosphate storage, phytase may function as an antioxidant in the seeds (Graf et al., 1987; Graf and Eaton, 1990). In plant seeds, most of the iron is complexed with phytate (Morris and Ellis, 1976), thereby alleviating the potentially lethal combination of free iron and unsaturated fatty acids in close proximity. Very low phytic acid maize mutants produce either stunted plants or seeds that are unable to germinate (Maugenest et al., 1999). C. INCREASED USE OF PLANT MEALS IN ANIMAL FEED

The soybean industry has successfully expanded its markets in the latter half of the twentieth century. Feeds for monogastric animals increasingly use soybeans, grains, and other plant seeds that contain high levels of phytic acid. Considerable research efforts have resulted in a transition in one segment of agriculture into an "animal agriculture" that requires large quantities of cereals and meals (Berlan et al., 1977). Between 1972 and 1992, the poultry industry was able to switch from fish meal as its primary protein source in its feed ration to lowercost plant protein sources such as soybean meal (Rumsey, 1993). This was achieved by the efforts of poultry nutritionists who successfully removed inhibitors and antinutritional factors from plant meals, and then developed amino-acid and mineral supplements that enhanced the nutritional profile of these plant meals until they were essentially equal to fish meal. The phytin phosphorus in plant meals was unavailable to monogastric animals because they lack phytase. The lower cost achieved through substituting soybean and other plant meals in these animal feeds was a major consideration for this research. Economic pressures continue to create a trend toward larger animal production units coupled with lower production costs (Mallin, 2000). Cost-effectiveness also dictated the means to deal with high levels of phytic acid in these meals. All the phytin phosphorus in these meals was unavailable to monogastric animals because they lack phytase.

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This lack of adequate phosphorus was corrected by supplementing soybean and other meals with relatively inexpensive rock phosphate, which provided the animal with this necessary nutrient. The excess phytin phosphorus was disposed of in the animals' manure. However, this practice added even more phosphate to the animals' feed ration and resulted ultimately in phosphorus levels in the manure that far exceeded the land's capability. D. EARLY RESEARCH IDENTIFIES A FUNGAL PHYTASE The growth of the market for phosphate to supplement animal feed fostered a critical step in the commercial development of phytase. Wodzinski and Ullah (1996) detailed the role that International Mineral and Chemical (IMC), a supplier of rock phosphate to the feed industry, had in initiation of the research that first identified a phytase from Aspergillus niger (ficuum) NRRL 3135 (ATCC 66876) in 1968 (Nelson et al., 1968b) with high enough activity to be considered a candidate for production as an animal feed supplement. In a remarkable display of foresight, IMC management sponsored research that surveyed hundreds of microorganisms for phytase activity. They apparently realized that a microbial phytase might one day be marketed as an effective means to hydrolyze phytic acid in plant meals and that they could supply this enzyme to animal feed producers. However, the necessary techniques to achieve overexpression of fungal phytase had not yet been developed and the project was terminated in 1968 before commercialization could be achieved. Dr. Rudy Wodzinski, a member of that IMC research effort, remained convinced of the merits of developing phytase as an animal feed additive. His continued interest in this project served to attract other scientists to this research, and he remained until his death in 1997 a strong advocate for the future potential of this enzyme. One of the early problems for researchers was the lack of a commercial supply of phytase. When Dr. Wodzinski was contacted by the Linus Pauling Institute in 1986 about a request for a supply of phytase, he supplied Sigma Chemical Company with both the necessary growth requirements and the A. niger isolate to produce enough phytase to supply the research efforts of several scientists. Dr. Wodzinski also served as a consultant to the Agricultural Research Service of the U.S.D.A. in 1984 when it started its own phytase research program. His support of this ARS study resulted in the initial characterization of phytase (Ullah and Gibson, 1987) and partial cloning of the phytase gene (Mullaney et a]., 1991). In time, when advances in biotechnology made it possible, the phytase gene from A. niger NRRL 3135 was overexpressed and its product commercialized (Van Hartingsveldt et al., 1993).

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161

II. Phytases that are Histidine Acid Phosphatases (HAPs) A. PHYA Several types of acid phosphatases have been reported in biological systems. These include purple acid phosphatases with a dinuclear Fe-Fe or Fe-Zn center in their active site (Klabunde et al., 1996), the low-molecular-weight acid phosphatases, and the high-molecularweight acid phosphatases (Vincent et al., 1992). A. niger NRRL 3135 phytase A (phyA) belongs to this last group and features a conserved active site motif, RHGXRXP (Fig. 2), unique to this class of enzyme, that hydrolyzes phosphomonoesters in a two-step mechanism (Ullah et al., 1991; Van Etten et al., 1991). Several other fungal (Table I), bacterial, and plant phytases are now known to belong to the histidine acid phosphatase (HAP) class of enzymes (Wodzinski and Ullah, 1996). All these phytases share this common active site structure. As one of the best characterized HAPs, the A. niger NRRL 3135 phytase molecule is being employed as a model to better understand this class of enzymes.

TABLE I FUNGALHAP PHYTASES

PhyA

A. niger (ficuum) NRRL 3135 A. niger var. awamori ATCC 38854 A. niger T213 (A. niger CB) A. niger SK-57 A. fumigatus ATCC 130703 a Emericella nidulans (A. nidulans)

Accession numbers JN0656 P34753

Number of Cys

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-

(Roche Nr. R1288)

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10

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aErroneously designated ATCC 34625 in Wyss et al., 1998.

bThielavia heterothallica.

8

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FIG. 2. The A. niger NRRL 3135 phyA amino-acid sequence (NCBI Accession No. JN0656). The conserved residues having sidechains protruding into the reaction cavity have an asterisk (*) above them (Pasamontes et al., 1997b) and the # above the 10 Ash residues denotes glycosylation. The N-terminal (N) (RHGXRXP) and C-terminal (C) (HD) motifs found in histidine acid phosphates are highlighted. The number above each of the 10 cysteine residues refers to the individual disulfide bridge to which it belongs (Kostrewa et al., 1997). The two acidic and four basic amino acids, respectively, that compose the substrate specificity site (Kostrewa et al., 1999)--Glu228, Asp262, Lys91, Lys94, Lys3OO, and Lys3Ol--are underlined.

A . n i g e r NRRL 3135 p h y A is a m o n o m e r i c p r o t e i n w i t h a m o l e c u l a r w e i g h t of 48.5 kDa for the u n g l y c o s y l a t e d e n z y m e (Ullah a n d Dischinger, 1993). H o w e v e r , the n a t i v e e n z y m e , w h i c h is h e a v i l y glycosylated, s h o w s a m o l e c u l a r w e i g h t of 85 kDa (Ullah, 1988). Fungal p h y t a s e , w h e n p u r i f i e d f r o m s h a k e - f l a s k - p r o d u c e d cells g r o w n in the p r e s e n c e of starch a n d l o w p h o s p h a t e c o n d i t i o n s , s h o w s m i c r o h e t e r o g e n e i t y in m o l e c u l a r w e i g h t d i s t r i b u t i o n as e v i d e n c e d b y the diffuse b a n d in

ADVANCES IN PHYTASE RESEARCH

163

SDS-PAGE gel electrophoresis giving a molecular weight of 85-100 kDa (Ullah and Gibson, 1987). It is noteworthy that A. niger NRRL 3135 phyA shows inherent thermostability since its optimum temperature was determined to be 58°C (Ullah and Gibson, 1987). One of the reasons for higher thermostability in this enzyme could be the presence of 10 cysteine (Cys) residues, which allow the protein to have 5 disulfide bridges (Fig. 2). Both biochemical (Ullah and Mullaney, 1996) and X-ray crystallographic data (Kostrewa et al., 1997) indicate that all 10 Cys residues are involved in forming 5 disulfide bridges. When the putative phyA gene from Arabidopsis thaliana was checked for Cys residues, it also revealed the presence of 10 Cys residues (Mullaney and Ullah, 1998b). The presence of five disulfide bridges is becoming a hallmark of phyA proteins, especially in microbes (Table I). The active site residues in phyA from various sources are well described. The active site motif containing the conserved septapeptide had been discovered independently by two labs in 1991 (Ullah et al., 1991; Van Etten et al., 1991). The vestiges of phyA's active site, however, were first described in 1989 by Bob Fletterick's research team while searching for sequence similarities among diverse phosphate-metabolizing enzymes such as fructose 2,6-biphosphatase, phosphate glycerate mutase, and acid phosphatases (Bazan et al., 1989). The other hallmark of phyA protein is the C-terminal HD motif (His361 and Asp362, Fig. 2), which had been recognized first by the Van Etten group as early as 1991 (Van Etten et al., 1991). The HD motif is well conserved in a wide variety of phytase sequences (Ehrlich et al., 1993; Mitchell et al., 1997; Pasamontes et al., 1997b). The catalytic His (His82) and Asp (Asp362) residues come from very different parts of the protein and are located in close proximity to each other in the active site area of the phytase molecule, as shown in the X-ray-deduced three-dimensional structure of phyA (Kostrewa et aL, 1997). The equivalent His residue in E. coli acid phosphatase is thought to perform a nucleophilic attack on the scissile phosphoester bond (Ostanin et al., 1992), and the equivalent Asp residue is thought to protonate the leaving group (Ostanin and Van Etten, 1993). A. niger phytase is extensively glycosylated, as pointed out earlier by the banding pattern in SDS-PAGE (Ullah and Gibson, 1987). The secreted phytase was found to be stable for months in the crude culture filtrate. In addition, during chemical sequencing of the protein it was necessary to add urea to a concentration of 4.0 M so that trypsin and chymotrypsin could cleave the peptide bonds (Ullah and Dischinger, 1993). Thus, it was thought that glyco-conjugates that are present in phytase must be preventing the protease from degrading the peptide

164

EDWARD J. MULLANEY et al.

bonds. In phytase, there are 10 (originally reported as 9) asparagine (Asn) residues that could be glycosylated, and all of them were found to be glycosylated (Ullah and Dischinger, 1993). The role of glycosylation in the functional expression of A. niger phyA in Pichia pastoris has been investigated by Han and Lei (1999). They found that glycosylation was vital to the enzyme's thermostability. Glycosylation patterns for several fungal phyAs expressed in different expression systems have been determined (Wyss et al., 1999b). The extent of glycosylation varied, but no significant effect on specific activity, thermostability, or the refolding properties of individual phytases was reported. The role of glycosylation in fungal phyA protein was studied by expressing the gene in E. coli where glycosylation would not be possible. When the gene was expressed, the protein did not fold properly to produce activity (Phil]ippy and Mullaney, 1997). Thus, glycosylation may be assisting in folding the enzyme. In fungi grown in phosphatelimiting conditions, the p h y A gene product has to travel across the membrane to reach the phosphate source, that is, starch in the medium. It is plausible that glycosylation of Ash residues not only facilitates the enzyme's transport across the membrane, but it may also help the linear polypeptide to fold appropriately as it exits the cell. To the contrary, in E. coil where A. niger p h y A gene was overexpressed, the enzyme was found in inclusion bodies where it did not fold properly. This may explain why fungal phytase has not been successfully cloned and expressed in bacteria. In 1996, a process for the deglycosylation of proteins for crystallization using a recombinant glycosidase fusion protein was developed (Grueninger-Leitch et al., 1996) that enabled Kostrewa et al. (1997) to first crystallize A. niger NRRL 3135 phyA and then determine its structure by X-ray crystallography. The X-ray crystal structure of phyA established how the 5 disulfide bridges are formed from the 10 Cys residues: Cys31-Cys40, Cys71-Cys414, Cys215-Cys465, Cys264Cys282, and Cys436-Cys444 (Fig. 2). The structure can be subdivided into a large ~/~ and a smaller s-domain. A deep indentation that contains the catalytically essential amino acids Arg81 and His82 is formed at the interface of these two domains. By comparing the crystal structure of A. niger NRRL 3135 phyA with the previously determined structure of another HAP, rat acid phosphatase (Schneider et al., 1993), Kostrewa concluded that several conserved residues--Arg81, His82, Arg85, Arg165, His361, and Asp362-from the ~/[3 domain are essential for the catalytic process of phosphoester hydrolysis. In a model of substrate binding for this study, all these conserved amino-acid residues are involved with the scissile 3-phosphate group of phytate when hydrolysis is initiated. Several

165

ADVANCES IN PHYTASE RESEARCH

applications of this X-ray crystallization study to facilitate enzyme engineering of phytase are provided in Section V. A phytase from Peniophora lycii, a basidiomycete, has recently been reported and is awaiting approval for marketing as Bio-Feed ® Phytase by Novo Nordisk (Lassen et al., 1997). Based on a comparison of its amino-acid sequence with A. niger NRRL 3135 phyA, it appears to be an HAP with 10 cysteine residues; 4 of the 5 disulfide bridges show strong conservation (Fig. 3). Only the two N-terminal cysteine residues, forming the first disulfide bridges in A. niger phyA, are different in P. lycii phytase. One unique feature of P. lycii phytase is that, unlike A. niger NRRL 3135 phyA, it is a 6-phytase. All the known Aspergillus phytases start the hydrolysis of phytic acid at the third phosphorus group, that is, 3-phytase. Preliminary studies suggest that this enzyme

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166

EDWARDJ. MULLANEYet al.

has a high initial rate of liberating phosphate from phytic acid coupled with high specific activity. B. PHYB Another extracellular enzyme from A. niger NRRL 3135 with phytase activity, which is also an HAP, is phytase B (phyB). This enzyme had been referred to in the literature initially as a pH-2.5-optimum acid phosphatase and was thought not to have phytase activity (Ullah and Cummins, 1987). However, additional studies have now shown that it does (Ullah and Phillippy, 1994), and one study has recently reported on its overexpression in a fungal expression system and the high phytase activity from the recombinant enzyme (Meittinen-Oinonen et al., 1997). The deduced amino-acid sequence is currently available for three phyB genes: phyB from A. niger NRRL 3135 (Ehrlich et al., 1993), aph from A. niger var. Awamori ATCC 38854 (Piddington et al., 1993), and pH-2.5 acid phosphatase A. niger T213 (Kostrewa et al., 1999). Irving and Cosgrove (1972) were first to identify an A. niger NRRL 3135 phosphomonoesterase with phytase activity and a pH optimum for phytic acid of 2.0, which later was shown to be phyB. The enzyme was not purified, and, as such, molecular details and kinetic characterization were not available at the time. In 1987 the purification and characterization of a phosphomonoesterase with a pH optimum of 2.5 was reported (Ullah and Cummins, 1987). When this enzyme was tested for phytate breakdown at pH 5.0, the results were negative. Later, when the same enzyme from A. niger was assayed at pH 2.5 for phytate hydrolysis, it turned out that this enzyme is also an efficient phytase with a turnover number of 628 per second as opposed to 348 per second for phyA when phytate was used as a substrate (Ullah and Phillippy, 1994). Like phyA, pH-2.5 acid phosphatase or phyB is a 3-phytase (Irving and Cosgrove, 1972). The phyB gene, which has been reported in other Aspergillus isolates, was also cloned from Aspergillus awamori by ALKO, a Finnish biotechnology company (Piddington et al., 1993). Wyss and colleagues showed a fungal phyB to have low phytase activity (Wyss et al., 1999a). A close examination indicates that the activity of the enzyme was tested at a substrate concentration of 5 mM phytate (Fig. 2, Wyss et al., 1999a). We examined the effect of substrate concentration on enzyme activity with A. niger NRRL 3135 phyB phytase at pH 5.0 and 2.5. Maximum activity was observed at 1.5 mM phytate concentration. A concentration above this value was found to be inhibitory. This inhibition by excess substrate followed a concentration de-

ADVANCES IN PHYTASE RESEARCH

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pendency (Ullah, unpublished data). For A. niger NRRL 3135 phyA, at pH 2.5, a substrate concentration above 4 mM completely inactivated phytase activity (Fig. 3; Ullah, 1988). Wyss and his coworkers have interpreted their results to favor the view that phyB had low phytatedegrading ability. If the enzyme assays had been carried out at 1 mM phytate concentration, these authors would have drawn a very different conclusion. Thus, it appears that substrate concentration should be maintained at or about the 1 mM level to assess the enzymatic activity of extracellular phytase (Ullah, 1988). The X-ray crystal structure of A. niger T213 phyB is now known (Kostrewa et al., 1999). Unlike A. niger phyA, it is a tetramer formed by two dimers. Wyss et al. (1998) had reported A. niger T213 phyB (pH-2.5 acid phosphatase) to be an oligomer that is most likely composed of four identical protomers. The crystal structure reveals that the main contacts for each unit come from the N terminus, each interfacing with its neighboring molecule in that region. In this structure two dimers form a tetramer that allows each active site ready access to the substrates. As in phyA, phyB also shows five disulfide bridges in its X-ray crystal structure. The three bridges analogous to Cys71-Cys414, Cys215Cys465, and Cys436-Cys444 (Fig. 2) in phyA are conserved in phyB. The N terminus of phyB stretches out to allow for interfacing with its neighbor, while in phyA the N terminus is a disulfide bridge formed by Cys31-Cys40, which results in a compact loop configuration in this region. Despite having almost identical catalytic centers, phyA and phyB have different pH profiles for hydrolysis of phytate. PhyA hydrolyzes phytate at both pH 2.5 and 5.0, but phyB displays optimum phytase activity at pH 2.5 and lacks activity at pH 5.0. Kostrewa et al. (1999) attribute this variation to differences in the charge distribution at the substrate specificity sites. In the A. niger T213 phyB site, there are only two acidic amino acids, Asp75 and Glu272 (Fig. 4). At the A. niger NRRL 3135 phyA substrate specificity site, there are two acidic and four basic amino acids: Glu228, Asp262, Lys91, Lys94, Lys300, and Lys301 (Fig. 2) [the last two Lys residues were erroneously given as 250 (227) 1 and 251 (228) 1 in the report of Kostrewa et al., 1999]. The active site of phyB is thus more acidic than the active site of phyA. Therefore, at pH 2.5 the acidic amino acids (Asp75 and Glu272, Fig. 4) in the phyB substrate specificity site are uncharged and can accommodate the negatively charged phytate as a substrate. At this pH in phyA the four basic 1The amino-acid residue location in the abbreviated sequence in Fig. 4 (Kostrewa et al., 1999).

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EDWARD J. MULLANEY et aL 1

T213 phyB

FSYGAAIPQSTQEKQFSQEFRDGYSILKHYGGNGPYSERVSYGIARDPPTSCEVDQVIMV

T213 phyB

~ S P S A G ~ I E ~ d ~ V Y S I N T r E Y K G D I d ~ I . ~ W T Y Y V P N E ~ r Y N / ~ " I ~ S G P Y 120

60

2 N

T213 phyB

AGLLDAYNHGNDYKARYGHLWNGETVVPFFSSGYGRVIETNEGFFGYNYSTNAALN

180

T213 phyB

3 4 I IS ESEVMGADS LTPTCDTDNDQTTCDNLTYQLPQFKVAAARLNSQNPGMNLTASDVYNL

240

T213 phyB

•HVMASFELNARPFSNWINAPTQDEWVSFGYVHDLNYYYCAGPGDKNMAAVGAVYANASLT

4 300

T213 phyB

LLNQGPKEAGSLFFNFA~{{~I~ITPI LAALGVL I PNEDL PLDRVAFGN PYS IGN IVPMGGH 360 C 1 5 5 LT I ERLSCQATALSDEGTYVRLVLNEAVLPFNDCTSG PGYSCPLANYTS I I/¢KNLPDYTT 420

T213 phyB

TCNVSASYPQYLSFWWNY~'fT~ELNYRSS PIACQEGDAMD

T213 phyB

3

2 460

FIG. 4. The amino-acid sequence from the crystal structure study of the A. niger T213 phyB phytase gene (Kostrewa et al., 1999). The N-terminal (N) (RHGXRXP1 and C-terminal (C) (HD) motifs found in histidine acid phosphatase are underlined. The acidic amino acids of its substrate specificity site (Kostrewa et al., 1999) are highlighted. The number above each of the 10 cysteine residues refers to the individual disulfide bridge to which it belongs (Kostrewa et al., 1999).

amino acids (Lys91, Lys94, Lys300, and Lys301; Fig. 2) of the substrate specificity site are all positively charged and would attract the negatively charged phosphate groups of the phytate molecule. When the pH is raised to 5.0, the acidic amino acids become negatively charged, while the basic amino acids remain positively charged. The substrate binding site of phyB would thus repulse the negatively charged phytate molecule while the site in phyA would still attract the phosphate groups of phytate. Applying this model for the substrate specificity site to A. niger NRRL 3135 phyB, it is interesting to note that only one of the two amino acids is acidic. In A. nigerNRRL 3135 phyB, while Glu272 is conserved, Asp75 is serine, an aliphatic neutral amino acid with a hydroxyl sidechain (Fig. 5). This may indicate that only a single acidic residue at position 272 is necessary. PhyB has been reported to have a broader substrate specificity than A. nigerphyA (Wyss et al., 1999a; Ullah and Cummins, 1988). The more

169

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3135 phyB T213 phyB

MPRTS LLTLACALATGASAFSYGA~ PQSTQEKQFSQEFRDGYS I LKHYGGNGPYSERVS FSYG~AI PQSTQEKQFSQEFRDGYS I LKHYGGNGPYS ERVS

3135 phyB T213 phyB

1 YGI~RDP P ~ E V D Q V I M ~ PSAGKS I E E A I ~ S INTTEYKGDLAPLNDW 120 YG IARDPPTflCEVDQVIMV~GRRYPSPSAGK~I EEALAKVYS INTTEYKGDLAFLNDW 101

3135 phyB T213 phyB

2 TYYVPNECYYNAETTSGPYAGLLDAYNHGNDYKARYGHLWNGETVVP FFS SGYGRV I ETA 180 TYYVPNECY~SGPYAGLLDAYNHGNDYKARYGHLWNGET~FFSSGYGRVI ETA 161

3135 phyB T213 phyB

3 4 P3CFGEGFFGYNYSTNAALN I IS E S EVMGADS LT PTCDTDNDQTTCDNLTYQLPQF KVAAA 240 RKFGEGFFGYNYSTN~J~LN I I SESEVMGADSLTPTCDTDNDQTTCDNLTYQLPQFKVAAA 221

3135 phyB T213 phyB

RLNS Q N P G M N L T A S D ~ I~4AS F E I ~ P F S N W I N ~ D E W V S FGYVEDL~CAG 300 R/~S Q N P C ~ T A S D b ~ Y I ~ q ~ FELNARP FS NW INAFTQDEWVS F G Y V E D I ~ CAG 281

3135 phyB T213 phyB 3135 phyB T213 phyB

PGDK~AAVGAVY~N~.S L ~ Y = L N Q G P ~ P L F F N F ~ ITP I L ~ I PNEDLPLD P G D K N M A A V G A V Y A N A S L T L L N O G P K E A G ~ L F F N F A H ~ I T P I L A A L ~ I PNEDLPLD C 1 5 RVAFGNPYS I GN I V P M ~ T IERLS CQATALS D R G ~ ~ V L P F N I ~ S G PGYS RV~'GNPY S I GN IVPMGGHLT I ERLS CQATALSDEGTI'VRL~I~EA~rLPFNDCTSGPGYS

3135 phyB T213 phyB

C PLUS CPI~S

S

60 41

*

4

5

3

I LNRIqLPD~'~-~-I'C~VS~YPQYLS F ~ £ ' F r E I ~ ILNKNLPD~'FFI~VSASYPQYLS F W ~ ' T T ~ L ~

360 341

420 401

2

SP IACQ~D/t~ S P IACQEGDAMD

479 460

Fie. 5. A comparison of the amino-acid sequences of phyB of A. niger NRRL 3135 (NCBI Accession No. P34754) and the partial phyB sequence of A. niger T213 (Kostrewa et al., 1999). The N-terminal (N) (RHGXRXP) and C-terminal (C) (HD) motifs found in histidine acid phosphatase are underlined. The acidic amino acids of its substrate specificity site (Kostrewa et al., 1999) are denoted by an *. The number above each of the 10 cysteine residues refers to the individual disulfide bridge to which it belongs (Kostrewa et al., 1999). All of the nonconserved amino-acid sequence in T213 is highlighted.

neutral electrostatic field of the phyB substrate specificity site (Kostrewa et al., 1999) provides a reason for this. A wider variety of phosphomonoesters can be utilized effectively by phyB at its optimum pH as a substrate. The highly positive electrostatic field of phyA's substratespecific site is optimized for binding negatively charged phytate. Consequently, other less charged substrates bind less effectively at that site. PhyB (pH-2,5 acid phosphatase) from A. niger T213 is more thermostable than recombinant phyA from either A. niger T213 or A. fumigatus ATCC 13070 (erroneously designated ATCC 34625) (Wyss et al.,

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EDWARD J. MULLANEY et al.

1998). However, exposure of A. niger T213 phyB to 90°C results in an irreversible conformational change and complete inactivation of the enzyme. PhyA from A. niger and A. fumigatus were denatured at temperatures between 50 and 70°C. A. fumigatus phyA, after cooling to 30°C, refolds into a native-like conformation and regains most of its activity. But the less heat-tolerant A. niger T213 phyA is capable of properly refolding only after heating to 50°C. The tetrameric structure of A. niger T213 phyB provides this higher thermostability. But it also explains why its individual protomers are unable to properly reassociate into an active tetramer once the molecule is denatured.

C. E. c o u HAP PHYTASE Members of the HAP group also occur in prokaryotes. For example, E. cob HAP phytase is also known. Just as A. niger phyB was first reported as a pH-2.5 acid phosphatase, this periplasmic enzyme encoded by the E. cob appA gene was first identified in the literature as an acid phosphatase with a pH optimum of 2.5 (Dassa and Boquet, 1985). Subsequently, it was denoted a phytase P2, because of its high phytase activity (Greiner et al., 1903). This study also elucidated the hydrolytic pathway this enzyme employs for phytin and accordingly established it as a 6-phytase. Recently, a recombinant form of E. cob phytase was purified and crystallized to provide a three-dimensional structure of a 6-phytase (Jia et al., 1098). This X-ray-deduced model can be used with the crystal structure of A. niger phyA, a 3-phytase (Kostrewa et al., 1997) to define the structural basis for their different catalytic pathways. The complete nucleotide sequence of the E. cob appA gene has been determined (Dassa et al., 1990). Rodriguez et al. (1999) utilized this sequence to design primers in order to clone the phytase gene from an E. coli isolate selected for its high phytase activity. The nucleotide sequence for this second E. cob phytase gene, appA2, was 95% homologous to appA. This translated into seven different amino acids in its deduced sequence. The significance of these sequence changes was established when both the E. cob appA and appA2 genes were expressed in Pichia pastoris and their recombinant proteins, r-appA and roappA2, were markedly different in their pH profile and other catalytic characteristics (Rodriquez et al., 1999). Both r-appA and r-appA2 have identical sequence in the regions of the N-terminal motif (RHGVRAP, positions 38-44) and the C-terminal motif (HD, positions 325-326) that is characteristic of HAP. It was by a

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site-directed mutagenesis study with this gene, E. coli appA, that Ostanin and Van Etten (1993) demonstrated the critical nature of the C-terminal motif, HD, to the catalytic function of the HAP class of enzymes. D. YEASTHAP PHYTASE Phytases have been reported previously in yeast (Nayini and Markakis, 1984). Several yeast acid phosphatase genes have now been cloned and characterized. For a review, see Wodzinski and Ullah (1996). In this review, three yeast acid phosphates were surveyed as HAP representatives and shown to have both the RHGXRXP and HD motifs in their amino-acid sequence. These three enzymes are encoded by the following genes: Schizosaccharomyces pombe phol, Saccharomyces cerevisiae pho3, and Schizosaccharomyces pombo pho4. Subsequently the S. cerevisiao pho3 gene was cloned and transformed into an Asporgillus oryzae expression system (Moore et al., 1995). An assay of the resulting recombinant S. cerevisiae HAP indicated high phytase activity. In addition, two other S. cerevisiae HAP genes were included in that screening, and they were shown to encode recombinant S. cerevisiae HAP with phytase activity. This suggests that similar yeast HAPs also can effectively hydrolyze phytic acid. Additional details of this transformation study are given in Section V.E. E. PLANTS Genes containing the HAP active site motif have recently been reported in diverse species of plants. In maize, a phytase cDNA, phy $11, was cloned and sequenced (Maugenest et al., 1997). This cDNA was then utilized to screen a maize genomic library. Two different genes, PHYT I and PHYT II, were identified (Maugenest et al., 1999). The study indicated that both PHYT I and PHYT II are expressed in germination of this monocotyledon, but only PHYT I is expressed in adult roots. A high level of homology is evident in the transcribed sequences of these two genes. However, other than partial homology to the region of amino acids around the HAP consensus motif, RHGXRXP, little sequence homology is found with A. niger phytase. A putative Arabidopsis thaliana phytase, recently discovered by searching the Genbank database, shares more features with A. niger phyA phytase (Mullaney and Ullah, 1998b). The enzyme encoded by the A. thaliana gene has not only the septapeptide active sequence

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EDWARDI. MULLANEYet al.

RHGXRXP, but also the dipeptidic HD region necessary for phosphatase activity in fungal phytase. In addition, A. thaliana and A. niger HAP both have 10 Cys residues, necessary for proper structure folding by forming 5 crucial disulfide bridges in A. niger phyA. Moreover, the primary structure of both the plant and funga] phyA are comprised of 464 to 448 amino-acid residues, respectively. III. Phytases with an Undefined Active Site

A. BACILLUSPHYTASE(PHYTASEC) A Bacillus subtilis phytase that hydrolyzed only phytate was reported by Powar and Jagannathan (1982). It required calcium for activity and had optimum activity at pH 7.5. A thermostable phytase from Bacillus sp. DS11, later identified as B. amylo]iquefaciens (Ha eta]., 1999), was then purified and characterized (Kim et aI., 1998a) and its gene (phy) cloned and overexpressed in E. coil (Kim et al., 1998b). In that same year a differentresearch group, Kerovuoand colleagues(1998), reported on the characterization, gene cloning, and sequencing of a phytase (phyC) from Bacillus subtilis VTT E-68013. Sequence from the two cloned phytases indicates that neither of these contain the active site motif found in HAPs. Analysis of their putative amino-acid sequence shows they are highly homologous (93% sequence identity). However, while they both require calcium for activity and have a similar pH optimum, B. subtilis VTT E-68013 pH 7-7.5 and B. amyloliquefaciens pH 7-8.0, have different optimum temperatures: 55°C for B. subti]is and 70°C for B. amyloliquefaciens. Preliminary X-ray crystallographic analysis of the B. amy]o]iquefaciens thermostable phytase (TS-Phy) has been initiated (Ha et al., 1999). Additional research on the contributions that their divergent sequence contributes to different temperature optima will advance our understanding of this novel class of phytase. B. KLEBSIELLAPHYTASE

Phytases have been isolated from both Klebsiella terrigena and K. aerogenes. A cytoplasmic phytase has been isolated from K. terrigena (Greiner et al., 1997). It is reported to be a monomeric 3-phytase, with a molecular weight of 40 kDa. Like A. niger NRRL 3135 phyA, the K. terrigena phytase has an optimum temperature of 58°C and is not a metalloenzyme. This study also indicated that it is rather specific for phytate and has an optimum pH of 5.0. Tambe et aL (1994) found two inducible molecular forms of phytase from K. aerogenes (Aerobacter

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aerogenes). The molecular weight of the large form is 700 kDa, and of the other form 10-13 kDa, making it the smallest known fraction to exhibit phytase activity. This indicates that, despite its size, an intact active site is present in the 13-kDa form. The possibility that the 700kDa fraction was formed by noncovalent bonding of the 13-kDa enzyme with other larger proteins was discussed. The two also differed in their optimum pH value: 5.2 for the low-molecular-weight form and 4.5 for the larger. C. YEAST Two yeast species, Schwanniomyces castellii (Schwanniomyces occidentalis) (Segueilha et al., 1992) and Arxula adeninivorans (Sano et al., 1999), have secreted phytase that has been characterized. S. castelli phytase is tetrameric in structure with a molecular weight of 490 kDa. When deglycosylated, this tetramer is composed of one large unit (125 kDa) and three identical subunits (70 kDa). Its phytase has been reported to be thermostable up to 74°C. Conditions to optimize S. castelli CBS 2863 phytase yield have been determined (Lambrechts et a]., 1993). Both S. castelli CBS 2863 and A. adeninivorans secreted phytases have high optimum temperatures: 77 and 75°C, respectively.

D. PLANTSAND MICROBES A phytase has been purified from soybeans (Morgan et al., 1998). Based on limited amino-acid sequence, it does not show homology with any known HAP phytase. Amino-acid sequence analysis reveals its only match is with a region in the N terminus of a putative purple acid phosphatase in Arabidopsis (NCBI Accession #AAC04486). Phytase from scallion (Allium fistulosum L.) leaves has also been purified and determined to have a maximum activity at pH 5.5 and an optimum temperature of 51°C (Phillippy, 1998). The phytase from another monocotyledon, oats (Arena sativa), has also been isolated. It is a monomeric enzyme with a molecular weight of about 67 kDa and has pH and temperature optima of 5.0 and 35°C, respectively (Greiner and Alminger, 1999). McElhinney and Mitchell (1993) demonstrated phytase activity in ectomycorrhizal fungi. Isolates of Paxillus involutus, Suillus grevillei, and two unidentified basidiomycetes from Sitka spruce were shown to have phytase activity. The enzyme activity appears to be membrane-

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EDWARD J. MULLANEY eta].

bound and not extracellular. It was postulated that this might be an adaptation for growth in soils where available phosphorus is low and there is intensive competition for limiting nutrients. Paramecium phytase was found to degrade phytate to Ins(1,2)P2 via the dephosphorylation sequence 6/5/4/1 (Van Der Kaay and Van Haastert, 1995). In investigations of phytate synthesis, the Paramecium phytase allows the kinetics of incorporation and release of radiolabeled phosphate to be precisely followed. IV. Increased Phosphorus Levels in Our Environment Creates Need for Phytase After the Second World War, industrial nitrogen fixation technology that was originally developed for munitions production was adapted to supply agriculture with nitrogen for fertilizer. This ushered in the age of fertilizer and transformed agriculture. With this development, nitrogen could be combined with phosphorus and potassium to precisely match the nutrient requirement of crops. This has contributed to the specialization found in agriculture, and in the 1990s the upper Midwestern states became the major users of phosphorus fertilizer, primarily for feed-grain production; this grain was then exported (Lanyon, 1999). This abundance of feed grain has fostered the emergence of industrial animal production units in various regions of this country (Mallin, 2000). During the last decade, numerous large poultry and swine production units have been constructed in coastal regions of the Southeastern United States. Traditional reliance on crops to utilize and crop lands to bind the high level of phosphorus in the manure from these operations has been inadequate. When the capacity of the soil to bind phosphate is exceeded, it enters groundwater or adjacent surface water bodies. This is especially problematic in the sandy soils that are typical in coastal plains. An increased concern has recently emerged about the environmental consequences of this nutrient runoff from agricultural operations. This concern has been generated in part by an increasing number of blooms of toxin-producing microbes around the world. A recent example of the phenomenon was large-scale fish kills in the Chesapeake Bay attributed to a toxin-producing dinoflagellate, Pfiesteria piscicida, which in 1997 attracted national news coverage (Mlot, 1998). Very limited research has been carried out on the effects of phytic acid or inositol phosphates on aquatic microbes. Chu (1946) studied

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the ability of marine diatoms to utilize various phosphorus compounds. Phytin was shown to support the growth of Phaeocystis pouchetii, Skeletonema costatum, and Nitzschia closterium. The oceanic dinoflagellate Pyrocystis noctiluca is also known to utilize phytic acid (Rivkin and Swift, 1980). The involvement of inositol phosphates has been cited in cyst formation in another dinoflagellate, Crypthecodium cohnii (Tsim et al., 1998). Research on Pfiesteria piscicida has established that organic phosphorus does stimulate both its toxic form and the nontoxic zoospores. Phosphorus is also believed to play a role in the transformation of various stages of P. piscicida into a toxic zoospore stage (Burkholder and Glasgow, 1997). The widespread concern caused by the 1997 Pfiesteria fish kill in the Chesapeake Bay resulted in a Blue Ribbon Citizens' Pfiesteria Action Commission. This commission investigated the phenomenon and issued a report to the Governor of Maryland. Among its findings was that the large poultry-producing region of the state was applying more phosphorus in manure than the crops could utilize (Mallin, 2000). A complex of factors resulting from developments in the fertilizer and feed industries have resulted in the phosphorus supplies on many farms now exceeding their crops' nutrient requirements (Lanyon, 1999). The commission considered several means for reducing the amount of phosphorus manure applied to the land. They concluded that the most encouraging proposal in this area was the use of phytase in animal feed. The efficacy of phytase in reducing phosphorus levels in livestock manure has been established (Yano et al., 1999; Ward, 1993). Acting on this part of the commission's report, Governor Glendening proposed that as of 1 January 2000 phytase was to be added to all poultry feed produced in Maryland. To help defray the added cost of preparing the feed mills to use this enzyme, it was also proposed that the state support a cost-sharing program with feed producers.

V. Engineering Phytase A. HEAT TOLERANCE

The cost of using phytase as an animal feed additive can be reduced if the heat tolerance of the enzyme is increased. This is because, during processing the plant meal, most of the meal mash is heated briefly to a high temperature (65-95°C). The A. niger phytase that is currently marketed would be denatured at these temperatures. This means an additional processing step is required after pelletization to add the phytase, thus adding to the cost. A phytase that combines the desirable

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EDWARDJ. MULLANEYet al.

traits in the A. niger NRRL 3135 phytase that is now commercially available with high heat tolerance would therefore be a superior enzyme for most current animal feed applications. The search for higher thermostability has led to the cloning of phytase genes from thermophilic fungi. The phytase genes from Myceliophthora thermophila (Mitchell et al., 1997) and Talaromyces thermophilis (Pasamontes et al., 1997b) have been cloned. Analysis of their predicted amino-acid sequence revealed that both have a high identity to known phytases and are HAPs. Their optimum temperatures were not reported, but the optimum pH for enzyme activity for phytic acid for M. thermophila is 5.5 (Wyss et al., 1999b). Another thermophilic fungus, Thermomyces lanuginosus, has been cloned and the expressed enzyme characterized (Berka et al., 1998). T. lanuginosus phyA encodes a mature protein of 442 amino acids that has 47% homology with the A. niger phytase (phyA). The T. lanuginosus phytase gene was transformed and expressed in Fusarium venenatum. The optimum temperature for phytase activity for the recombinant phytase was 65°C. While the pH profiles for A. niger and T. lanuginosus phytase (phyA) are similar, T. lanuginosus has a slightly higher optimum pH, pH 6, and its enzyme is active at neutral pH, where A. niger phytase lacks activity. Several other phytases have been investigated as possible enzymes with increased heat tolerance. They include phytase from Aspergillus terreus No. 9A-1, optimum temperature 70°C (Yamada et al., 1968); Schwanniomyces castelii (Segueilha et al., 1992); Arxula adeninivorans (Sano et al., 1999); Bacillus sp. DS11 (Kim et al., 1998a); and A. fumigatus ATCC 13073 (Pasamontes et al., 1997a). The A. fumigatus ATCC 13073 phytase gene has been cloned and overexpressed in A. niger NW205 (Pasamontes et al., 1997a). The recombinant phytase was reported to withstand temperatures up to 100°C for more than 20 minutes with only minimal loss (10%) of its enzymatic activity. In a subsequent study (Wyss et al., 1998), this recombinant phytase was compared with the phytase encoded by the A. niger T213 phytase gene, which was cloned and also overexpressed in A. niger NW205. A comparison of the two recombinant phytases revealed that, while A. niger T213 phytase does not have the capacity to refold properly after heat denaturation, the recombinant A. fumigatus phytase refolds into a native-like and fully active configuration. Studies to support the potential use of A. fumigatus ATCC 13073 phytase as a feed additive that can withstand the brief period of high temperatures required in pelletization of animal feed have been performed. When the recombinant A. fumigatus phytase was added to feed

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mash that was then pelleted at 85°C, approximately 50% of the enzymatic activity was recovered (Wyss et al., 1998). Nunes and Guggenbuhl (1998) evaluated the efficacy of recombinant A. fumigatus phytase in a feeding trial with pigs. Recombinant A. fumigatus phytase was shown to significantly decrease the phosphorus content in feces, and it also increased the growth rate for the pigs over pigs fed with a control diet. Analysis of the A. fumigatus ATCC 13073 phytase sequence reveals no obvious reason for its higher heat tolerance. The only unique feature of its sequence is a higher pI (7.28) than the other known fungal phytases (Wyss et al., 1999b). This pI value is also higher than most of the extracellular proteins produced by the expression strains used (A. niger NW205). This would make it especially well suited for rapid purification if it is produced on an industrial scale. Using a new biotechnology technique, the first "engineered" phytase has already been assembled. A phytase with an increased optimum temperature has been reported by using a consensus technique. This technique compares the amino-acid sequence of several known phyAs and then selects the most conserved choice for each residue. This sequence is then back-translated into a DNA sequence, and then this DNA is transformed into an expression system (Lehmann, 1998). A high degree of conservation of amino-acid sequence and features such as preservation of cysteines for disulfide bridges are observed in a comparison of the consensus and the A. niger NRRL 3135 phyA (Fig. 6). The use of compounds to enhance the thermostability of phytase has also been investigated (Phillippy, 2000). Phytate is reported to enhance the activity of A. adeninivorans phytase (Sano et al., 1999), and calcium contributes to the heat tolerance of Bacillus sp. DS11 (Kim et al., 1998a). The effects of different buffers on the heat tolerance of A. fumigatus phytase expressed in Pichia pastoris indicates they can facilitate refolding of the enzyme into the native-like, active configuration after heat denaturation (Rodriguez et al., 2000). Immobilization of E. coli phytase is reported to enhance its thermostability (Greiner and Konietzny, 1996). Glycosylation of recombinant A. niger phyA expressed in S. cerevisiae (Hanet al., 1999) and in Pichia pastoris (Han and Lei, 1999) contributes to the thermostability of the enzyme produced. B. TEMPERATURE AND PH OPTIMA

The optimum temperature for A. niger NRRL 3135 phyA activity is 58°C (Ullah and Gibson, 1987). This is approximately 20 ° higher than the

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EDWARD J. MULLANEY et al. 1

1

3135 phyA Consensus

phyA

60 60

313S phyA Consensus

phyA

120 120

3135 p h y A Consensus phyA

180 180

3135 phyA

240 240

Consensus phyA

3135 phyA

~

~

i

s

T

4

Consensus phyA

4

s ~ ~ z ~ ~ v ~ s ~ ~

3135 phyA

360 360

Consensus phyA

3135 phyA

420 420

Consensus phTA S

3135 phyA Consensus phyA

300 300

5

3 467 467

FIG. 6. A comparison of the amino-acid sequences of phyA of A. niger NRRL 3135 (NCBI Accession No. JN0656) and the consensus phytase (Lehmann, 1998). Regions of conserved amino-acid sequence are highlighted, and the number above each of the 10 cysteine residues refers to the individual disulfide bridge to which it belongs (Kostrewa et al., 1999).

body temperatures of poultry and swine, at which maximum activity is desirable. When used in aquaculture, with lower body temperatures of the animals, there is even a more pronounced activity reduction. Having a pH optimum similar to the pH level found in the digestive tract of the animal is essential for maximum effectiveness of the enzyme. Having a broad pH optimum for phytase activity as reported for A. fumigatus, pH 2.5-7.5 (Pasamontes et al., 1997a), would expand the enzyme's potential usefulness as a feed additive. Crystal structure studies of differences in charge distribution of the substrate specificity site (Kostrewa et al., 1999) offers the first insight into the molecular basis for the pH optima of both phyA and phyB (see §II.B). When this pH optimum model is applied to P. lycii phytase, the local electrostatic

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charge of the respective amino acids in the substrate specificity site (Fig. 3) at pH 5.0 would be negative. This would explain the lower pH optimum, pH 4.0, reported for P. lycii phytase (Lassen et al., 1997). Further research into these mechanisms offers the potential to eventually engineer a feed additive superior to the native fungal phytase that is in commercial use today. C. SUBSTRATE SPECIFICITY

To be effective in an animal diet high in phytic acid, high phosphohydrolytic activity associated with phytase is essential. The phytases that have been characterized to date do not all have the same affinity for phytate as a substrate. Wyss et al. (1999a) have categorized E. coli and several fungal phytases into two groups based on activity levels for phytic acid and the ability of the enzyme to hydrolyze other substrates. The first group has high activity (102 to 811 U/mg) but a narrow substrate specificity and includes A. niger, A. terreus, and E. coli phytase. The second, with a low activity (23 to 41 U/mg), but a broad substrate range, includes phytases from A. fumigatus, A. nidulans, and M. thormophila. The activity of the second group is similar to the activity level of A. niger NRRL 3135 pH-6.0-optimum acid phosphatase (Ullah and Cummins, 1988; Ullah and Dischinger, 1993), which is a metalloenzyme (Mullaney and Ullah, 1998a). A molecular basis for this division is not known at this time. Kostrewa et al. (1999) do offer some insight into the narrow substrate specificity of phyA for phytate. It is the positive charge of its substrate specificity site that, while optimized for binding phytate, is less attractive to other substrates not as negatively charged. D. ENZYME STABILITY

The stability of plant and microbial phytases have been reviewed by Phillippy (2000). A. niger phytase is known to be significantly more resistant to proteolytic digestion than either wheat (Phillippy, 1999) or A. fumigatus phytase (Wyss et al., 1999b). Recombinant phytases from A. fumigatus, A. nidulans, A. terrus 9A1, and M. thermophila when expressed in A. niger undergo proteolytic degradation. To address this problem, Wyss et al. (1999b) have successfully engineered the higher level of proteinase resistance found in A. niger phytase into A. fumigatus phytase. They compared the N-terminal amino-acid sequence of A. fumigatus phytase to the three-dimensional structure model of A.

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niger phytase, and deduced that the proteolytic cleavage site was within one of the exposed loop structures. A site between amino acids 152 and 153 of A. fumigatus phytase (Wyss et al., 1999b) was then identified as the probable cleavage site. Site-directed mutagenesis at that site with $152N and R151L/S152N yielded mutant proteins with reduced susceptibility to proteolysis. The utilization of this information from the available three-dimensional structure model marks a significant achievement toward the goal of actually being able to engineer a phytase molecule with improved enzymatic characteristics for use as an animal feed additive.

E. SYNERGISTIC EFFECT

When A. niger (ficuum) NRRL 3135 is grown under limiting phosphate conditions, it produces four different extracellular acid phosphatases. These four enzymes are phyA (Ullah and Gibson, 1987), phyB (Ullah and Cummins, 1987; Ullah and Phillippy, 1994), Apase6 (Ullah and Cummins, 1988; Mullaney et al., 1995), and phoA (Ehrlich et al., 1994). Whereas phyA and phyB, both histidine acid phosphatases, can effectively hydrolyze phytic acid, Apase6, which is a purple acid phosphatase, cannot. The active site motif and hydrolytic mechanism of phoA is not known, but this enzyme cannot effectively utilize phytic acid as a substrate. The genes for these enzymes are not clustered, but rather are dispersed throughout its genome (Table II). Their simultaneous expression by A. niger suggests that in nature, when this fungus experiences certain nutrient conditions, it needs all these enzymes to scavenge enough phosphorus from the available sources. A single acid phosphatase/phytase did not evolve with superior hydrolytic activity for all the phosphate sources this mold encounters in its surroundings. It also may be advantageous to combine different phytases or acid phosphatases in animal feed rations to achieve a more efficient utilization of the phytin phosphorus by the animal. A study by Moore et al. (1995) suggested that an increase in phytin phosphorus availability may be achieved by combining two acid phosphatases. Three Saccharomyces cerevisiae histidine acid phosphatase genes (pho3, pho5, and pho11) were cloned by polymerase chain reaction and overexpressed separately in an Aspergillus oryzae isolate. Phytase activity was then measured in the control A. oryzae and in individual isolates from the three transformations. All three transformations yielded isolates that displayed up to a four- to sixfold increase

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TABLE II CHROMOSOMELOCATIONOF A. NIGER ACID PHOSPHATASEGENES Gene

Chromosome

phyA phyB aphA phoA

III VIII IV

I

Chromosome location of the four A. niger phosphate-repressible acid phosphatase genes determined by CHEF electrophoresis and Southern blot analysis (Mullaney, unpublished data).

in phytase activity. Phytase activity of the native yeast acid phosphatases were not, however, reported in that study. In the same study, the A. niger aphA gene (MacRae et al., 1988; Mullaney et al., 1995), which encodes Apase6, a purple acid phosphatase that has low phytase activity, was also cloned and overexpressed in A. oryzae. One isolate from this transformation, with an estimated number of 20 copies of the aphA gene, was reported to have a fivefold higher level of phytase activity than the control A. oryzae. This suggests that the increase in phytase activity was due to a synergistic effect between the recombinant purple acid phosphatase and the host A. oryzae phytase. The efficacy of the pho5 and the aphA recombinant acid phosphatases was established in a feeding trial. In a chicken feed trial, the basal diet contained 0.25% unavailable P (phytic acid). Both enzymes were effective and raised the plasma phosphorus levels in the test animals to a level equivalent to that obtained in chickens fed a diet of supplemental inorganic phosphorus. Compared to a commercially available phytase, the recombinant acid phosphatases increased P utilization by 40% compared to 48% for phytase. Park et al. (1999) showed that more enzymatic activity might be induced by combining A. niger NRRL 3135 phyA and B. amyloliquefaeiens phyC. Their study indicated that different combination ratios of phyA and phyC, because of their different pH profiles, would be more effective at hydrolyzing phytate over the entire gastrointestinal tract than either single phytase alone.

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Another synergistic effect from combining two acid phosphatases was recently reported by Wyss et al. (1999a). Their study showed that A. fumigatus and several other fungal phyAs only release five of the six phosphate groups in phytic acid. However, when a combination of A. fmnigatus phytase (phyA) and A. niger pH-2.5 acid phosphatase (phyB) was utilized, all six of the phosphate groups were released. This suggests that this combination of enzymes would be beneficial in increasing phosphorus utilization in animal feed rations with high phytic acid levels. Evidence supporting this synergistic effect has also been described by Vanderbeke et al. (1994). By blending phyA and phyB, a higher synergetic phytate hydrolyzing efficiency after thermal treatment was observed. The higher thermal stability of phyB (pH-2.5 acid phoso phatase) and its importance in achieving this effect was noted. The molecular basis for phyB's greater heat stability is discussed in Section II.B. However, in a recent hog feeding trial and an in vitro study, no evidence of any synergistic interaction between phyA and phyB was reported (Nasi et al., 1999). In this study, genes encoding the A. niger var. Awamori phytase (phyA) and pH-optimum-2.5 acid phosphatase (phyB) (Piddington et al., 1993) were transformed separately into Trichoderma reesei. The transgenic phytases were then utilized in both in vitro and in vivo tests that showed no significant synergistic effect in hydrolyzing phytin phosphorus from barley, maize, and soybean meal, when the diet was fed without thermal processing. Research was also conducted on the benefits of producing phytase in a fungal expression system that simultaneously produces another hydrolytic enzyme. The gene for A. niger pH-2.5 acid phosphatase (phyB) (Piddington et al., 1993) was transformed and expressed in a high-celo lulase-production strain of Trichoderma reesei (Meittinen-Oinonen et al., 1997). The recombinant phytase was secreted into the culture medium broth and retained its activity. This enzyme mixture would increase the nutritional value of animal feeds containing significant amounts of both phytic acid and cellulose. An enzymic "cocktail" composed of phytase, an acid phosphatase, an Aspergillus saitoi acid protease, citric acid, and an A. niger pectinase was studied under simulated intestinal conditions to measure enzymatic dephosphorylation of corn (Zyla et al., 1995). The results indicated that the "cocktail" improved digestibility of phytate phosphorus, protein, and carbohydrates. The use of citric acid alone to lower gastric pH levels and thus enhance phytase efficacy has been tested in hog

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feeding experiments (Goihl, 1998). However, no significant effects were reported. VI. Enzyme Production

A. FILAMENTOUSFUNGI Today, large-scale fermentation operations employ overexpression and other biotechnological techniques to produce nearly all the recombinant phytase used by the animal feed industry. One company, Gist-Brocades, has cloned multiple copies of the A. nigerNRRL 3135 phyA gene into their PluGBug® system that yields high levels of phytase in their A. niger host. This product is now being marketed as Natuphos TM (van Dijck, 1999). Another enzyme producer, Novo Nordisk, is replacing their current phytase product, Phytase Novo TM, with another phytase cloned from Peniophora lycii, a basidiomycete (Novo Nordisk A/S, 1999). This P. lycii phytase will be overexpressed in an A. niger expression system and sold under the product name Bio-Feed Phytase (Ronozyme TM P). Alltech Inc. produces another phytase, Allzyme Phytase 115. This is a nonrecombinant phytase from a proprietary isolate.

B. EXPRESSIONIN YEAST The potential to employ a yeast expression system for commercial phytase production has been examined in several studies. Han et al. (1999) described the use of a relatively low-cost medium containing yeast extract-peptone-dextrose (YEPD) to produce A. niger NRRL 3135 phyA in a Saccharomyces cerevisiae system. This research utilized the pYES2 expression vectors (Invitrogen) to construct a plasmid, pYPP1, that when transformed and expressed in S. cerevisiae yielded up to 2797 units per liter of extracellular phytase activity in the medium supernatant within 15 hours. Hansenula polymorpha and Pichia pastor&, facultative methylotrophic yeasts, have been investigated as potential high-yield production systems for phytase. A. niger NRRL 3135 phyA has been expressed in P. pastoris and with the production of high levels of active phytase (25-65 U/ml of medium) (Han and Lei, 1999). This recombinant phyA shared the same characteristics with phyA overexpressed by A. niger with slightly improved thermostability profile. Phytase titers in the fermentation supernatant up to 13.5 g/liter for a "consensus" phytase (Lehmann, 1998) have been achieved by an expression system using a

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recombinant strain of H. polymorpha (Mayer et al., 1999). This system featured the use of an economical carbon source, glucose or glucose syrups, as a means to make this a low-cost process. However, lower yields (6.1 g/liter) were obtained when the A. fumigatus phyA gene was expressed in the system.

C. EXPRESSIONIN PLANTS

The gene for fungal phytase has been successfully overexpressed in several transgenic plants (Day, 1996; Verwoerd et al., 1995; Li et al., 1997). The ability to express recombinant phytase in plants offers the possibility for the development of plant varieties that would contain sufficient amounts of phytase in their grain or seed so that phytase supplements would not be required. In addition, the potential use of crop plants to serve as bioreactors to produce phytase commercially is being investigated. In 1995, Verwoerd and coworkers in Holland did express a functional phytase in Nicotiana tabacum through a constitutive expression of phytase cDNA and showed that the enzyme was secreted out of the cells. They achieved secretion to the extracellular fluid by the use of a signal sequence from the tobacco pathogen-related protein S (Verwoerd et al., 1995}. The expressed phytase was found to be biologically active and accumulated in leaves up to 14.4% of total soluble protein during plant maturation. Researchers at the University of Wisconsin Biotechnology Center have independently expressed the A. niger NRRL 3135 phyA gene in tobacco leaves. The full 441-aa protein was made in leaf tissue, which was purified to homogeneity and extensively characterized (Ullah et al., 1999). Except for a decrease in molecular mass due to reduced glycosylation, the expressed recombinant phytase was virtually the same as native fungal phytase. The catalytic properties of the cloned phytase were encouraging enough to open the possibility of overexpressing the fungal phyA gene in other crop plants. This could pave the way for producing phytase commercially in field crops. University of Wisconsin researchers have also developed alfalfa plants to commercially produce phytase. They performed cloning and expression of the fungal phytase gene, so that most of its product was contained in the juice collected after the alfalfa was processed (Gutknecht, 1997). The equipment investment for this biofarming process is minimal and potentially turns a byproduct into a source of additional income for the farmer. Other enzymes have also been expressed in these

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plant "bioreactors." But the results achieved with phytase enhances the feasibility of future development of this technology to produce this enzyme commercially. Another application of biotechnology in plants is to reduce the need for phytase by lowering phytate levels in the plant's cereal or meal. Maize cultivars with reduced levels of phytic acid have already been produced (Raboy and Gerbasi, 1996). Transgenic soybean isolates that overexpress fungal phytases and thus eliminate or reduce the need to supplement meal with phytase are also being pursued. Li et al. (1997) have expressed the A. niger NRRL 3135 phytase gene in soybean (Glycine max). The recombinant phytase had a lower molecular weight than the native fungal enzyme, but its temperature and pH optimum were almost identical to that of the native enzyme. This strategy of having transgenic plant overexpressed phytase still requires a heat-tolerant phytase to survive the elevated temperatures often required in feed production. Recent work at the Swiss Federal Institute of Technology in Zurich details the transfer of a heat-tolerant A. fumigatus phytase gene by transformation and its expression in rice (Oryza sativa L.). This research is targeted at improving the nutritional profile of rice by reducing the amount of phytate. Phytate binds up to 95% of the iron in rice and keeps it from being absorbed. Therefore, individuals in parts of the world with a high-rice diet are prone to iron deficiency (Gura, 1999). D. TRANSGENIC ANIMALS In the future, transgenic poultry, hogs, and so on may produce phytase in their own digestive tract. Several attempts have already been made to transform and express a fungal phytase in an animal (privileged information, personal communication). To date none of these attempts have been successful. Similar results were obtained when the phyA gene was expressed in E. coli (Phillippy and Mullaney, 1997). The problem could very well be associated with glycosylation or its lack in animal and bacterial cells, respectively. Native fungal phyA protein contains 10 asparagine (Ash) residues with glycosylation signals that are all N-glycosylated. In E. coli these Ash residues are not glycosylated, and perhaps the recombinant protein does not fold appropriately to produce active site geometry, which is essential for activity. Similarly, in animal systems where O-glycosylation is preferred, the 10 N-glycosylation sites will be left unglycosylated. This may explain the lack of activity of phyA protein expressed in animals. Perhaps engi-

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neering O-glycosylation sites at these locations may allow the mammalian cells to glycosylate either the threonine or serine residues and thereby allow for appropriate folding of recombinant phytase. This, however, needs to be tested. As more is understood about the structure-function relationships of the microbial phytases, another possible avenue is to modify an animal's HAP, that is, rat acid phosphatase (Kostrewa et al., 1997; Schneider et al., 1993), or multiple inositol polyphosphate phosphatase (Craxton et al., 1997) to enhance its ability to hydrolyze phytin. If successful, this would reduce the difficulty of obtaining expression in animal tissue. VII. Expanding Uses of Phytase A. POTENTIAL IN AQUACULTURE

Numerous studies have been conducted on the use of soybean meal or other plant meals in aquaculture, including feeding studies on rainbow trout (Watanabe and Pongmaneerat, 1993; Mwachireya et al., 1999), the greenback flounder (Bransden and Carter, 1999), and the African catfish (van Weerd et al., 1999). By substituting lower-cost plant protein for a more expensive protein source, such as menhaden fish meal, a significant cost reduction could be achieved. Feed costs constitute up to 70% of total fish production costs (Rumsey, 1993). The consumer price index over the period of 1982-92 showed that, while the price index for seafood increased by more than 50%, the cost of alternative proteins increased only an average of 30% (Chamberlain, 1993). As in poultry and hogs, fish lack an adequate digestive enzyme to effectively utilize the phytin phosphorus in this feed. Moreover, as aquatic animals, the problems associated with high phosphorus levels in the water from their waste is an immediate problem. Therefore, phytase has been evaluated as a means to both increase the use of low-cost plant meals in the aquaculture industry, and also to maintain acceptable phosphorus levels in the water. Several fish feeding studies have documented the potential value of phytase in diets containing high levels of plant feedstuffs (Robinson et al., 1996; Oliva-Teles et al., 1998: Mwachireya et al., 1999). Chamberlain (1993) has projected that global seafood consumption will increase 35% by 2025. The aquaculture industry will supply an increasing amount of the world's need for seafood. Whereas the major market for phytase today is as a food additive in poultry and hog feed, there is great potential to expand the market into feedstuffs used in

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aquaculture. The higher temperature required for pelletization of feed in aquaculture and the lower body temperature of fish may require the development of commercial phytases tailored for aquaculture feeds. B. PHYTASE AS A SOIL AMENDMENT

In certain locations, phytic acid and its derivatives may represent up to 50% of the total organic phosphorus in the soil (Dalai, 1978). This abundance of phytic acid in the soil and the possibility that the addition of phytase might stimulate plant growth in these soils has been investigated. Findenegg and Nelemans (1993) studied the effect of phytase (phyA) on the availability of phosphorus from phytic acid in the soil for maize plants. Growth stimulation was reported as the result of an increased rate of phytin hydrolysis when phytase was added to the soil. However, the amount of phytase necessary for a significant effect meant that this was not a practical technique at this time. This study also suggested that the expression of phytase in the roots of transgenic plants might increase the availability of phosphorus to plant roots (Day, 1996). To better understand the role root phytase plays in the phosphorus nutrition of plants, the phytase and acid-phosphatase activity of extracts from several temperate pasture grass and legumes were isolated and studied (Hayes et al., 1999). C. PRODUCTION OF MYO-INOSITOL PHOSPHATES

Greiner and Konietzny (1996) investigated the use of E. cob phytase to generate specific breakdown products from phytic acid. A packed-bed bioreactor containing covalently attached E. coli phytase was constructed in this study to economically produce special isomers of the lower myo-inositol phosphate esters. The bioreactor chiefly yielded I(1,2,3,4,5)P5, I(2,3,4,5)P4, I(2,4,5)P3, and I(2,5)P2 isomer forms. Because only one major isomer of each myo-inositol phosphate species was formed, further purification could be easily achieved by ion-exchange chromatography. The Km for phytate increased from 130 pM for free enzyme to 240 pM when E. coli phytase was immobilized. However, the catalytic turnover number was lowered from 6209 per second for free phytase to 1182 per second. Thus, on crosslinking, E. coli phytase's catalytic activity was slowed down drastically. However, the immobilized bacterial enzyme performed much better than the immobilized A. niger NRRL 3135 phyA (Ullah and Cummins, 1988; Dischinger and Ullah, 1992). One reason

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that the fungal phytase performed so poorly on immobilization could be due to extensive crosslinking of the enzyme through its glyco-conjugates. The active site geometry of Aspergillus niger phytase was altered significantly to cause such loss of activity. To the contrary, crosslinking of protein due to glycosylation would not be a problem when E. coli phytase is used in a bioreactor.

D. SEMISYNTHESIS OF PEROXIDASE A semisynthetic peroxidase was designed by taking advantage of the structural similarity of the active site of vanadium-dependent haloperoxidases and fungal phytases and acid phosphatases (van de Velde et al., 2000). The Delft group incorporated vanadate ion into the active site of A. niger (ficuum) NRRL 3135 phytase. This resulted in transformation of native phosphohydrolase activity of phytase into semisynthetic peroxidase. The "new" enzyme was able to catalyze enantioselective oxidation of prochiral sulfides, with /-/202 affording the S-sulfoxide. Under the reaction conditions, this semisynthetic vanadium peroxidase was found to be stable for over 3 days with only a slight loss in turnover number. The other exceptional feature of this "new" enzyme, being polar water-miscible, was that cosolvents, such as methanol, dioxane, and dimethoxyethane, could be used up to a concentration of 30% (v/v) with only a slight loss in activity. It is remarkable that of all acid phosphatases and phytases tested by the Delft group, only those enzymes belonging to the "Histidine Acid Phosphatase" class with the active site sequence RHGXRXP could function as a peroxidase when vanadate ion was incorporated into the active site. The idea of transforming certain acid phosphatases into enzymes acting as vanadium chloroperoxidase stems from the observation that vanadium chloroperoxidase shares structural similarities with some membrane-bound acid phosphates. Furthermore, the apoenzyme of vanadium chloroperoxidase could exhibit phosphatase-like activity (Hemrika et al., 1997; Neuwald, 1997). VIII. Occupational Health Concerns There have been several occasions when workers exposed to microbial enzymes have suffered allergic responses (Slavin and Lewis, 1971). Being aware of this, commercial producers of phytase now routinely

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include a safety warning on their product about the need for workers to exercise caution w h e n handling the enzyme. Extended exposure and breathing of dust are the main concerns. Bio-Feed ® Phytase from Novo Nordisk is available in a coated granulated form that is advertised as a nondusty product that offers several advantages over the powdered enzyme. Doekes et al. (1999) present evidence that phytase is an occupational allergen that can cause specific IgE immune responses among exposed workers. Because the commercially available phytase in this study was not purified to homogeneity, this allergic response could not be positively linked to phytase. However, it was concluded that measures to prevent airborne occupational exposure should be implemented at sites where phytase is handled. Recombinant phytase expressed in A. niger still contains a glyco-conjugate with Noacteyl-glucosamine and a high mannose chain. Prior research has demonstrated that these oligosaccharides are immunodominant in Aspergillus, and antibodies to these epitopes are readily detected in the serum of individuals with aspergillosis (Hearn and Shimizu, 1996). Noglycosylation of asparagine residues may play a role in the antigenicity of phytase. However, this needs to be studied in detail before one can conclusively ascertain what role, if any, this glyco-conjugate is playing in stimulating the IgE immune response.

IX. Future Prospects A. PHOSPHORUS FOR FUTURE GENERATIONS Today, agricultural operations are supplying rock phosphate at minimal cost from mining operations throughout the world. The current major use of this phosphate is in fertilizers. Phosphorus, like nitrogen, is essential for plants, just as it is for all other forms of life (Abelson, 1999). It is a basic component in nucleic acids, ATP, and numerous other biological compounds. Nitrogen, however, has a cycle that constantly replenishes the earth's supply. To the contrary, phosphorus has no analogous cycle. The geologic phosphorus deposits that are being mined today are millions of years old. They contain the phosphorus that was removed from our biosphere w h e n it precipitated from prehistoric oceans (Lanyon, 1999). This fact is beginning to generate concern that future generations may face a shortage as the demands on the world's phosphate reserves accelerates. Phytase is now being recognized for its beneficial environmental role in reducing the phosphorus levels in manure. But as the need to conserve the world's phosphate reserves increases in significance, the role of phytase may broaden.

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Some insight in gauging the future role of phytase may be obtained by examining the development of another group of industrial enzymes, subtilisin proteases. It is estimated that, of the total amount of enzymes produced in the world, 60% is destined for the laundry detergent market (Horikoshi, 1999). This huge market for subtilisin proteases and other enzymes developed in the last quarter of the twentieth century because of the phosphorus loading of the Great Lakes and other waterways (Alexander, 1977). Biotechnology responded to the need to replace detergent phosphate with alkaliphilic enzymes that would perform effectively as laundry proteases. Considerable research effort was channeled into the development of the superior laundry proteases that are available today (Wolff et al., 1996). The parallel between phytase and subtilisins is that they both were developed in response to a need to reduce excess phosphate in our environment. Phytase seems destined to become increasingly important, along with other innovations, as a measure to maximize efficient utilization of the earth's phosphorus supply. While it is not possible to predict how greatly the market for this enzyme will expand, trends in agriculture today suggest continued growth in demand. The development of a consensus phytase (Lehmann, 1998) points to still more research to develop a second-generation phytase with superior attributes as an animal feed additive. Continued research on lowering the production cost and expanding its utilization to other applications also suggests an increased importance of phytase in the immediate future.

X. Summary Since its discovery in 1907, a complex of technological developments has created a potential $500 million market for phytase as an animal feed additive. During the last 30 years, research has led to increased use of soybean meal and other plant material as protein sources in animal feed. One problem that had to be overcome was the presence of antinutritional factors, including phytate, in plant meal. Phytate phosphorus is not digested by monogastric animals (e.g., hogs and poultry), and in order to supply enough of this nutrient, additional phosphate was required in the feed ration. Rock phosphate soon proved to be a cost-effective means of supplying this additional phosphorus, and the excess phytin phosphorus could be disposed of easily with the animals' manure. However, this additional phosphorus creates a massive environmental problem when the land's ability to bind it is exceeded. Over

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the last decade, numerous feed studies have established the efficacy of a fungal phytase, A. niger NRRL 3135, to hydrolyze phytin phosphorus in an animal's digestive tract, which benefits the animal while reducing total phosphorus levels in manure. The gene for phytase has now been cloned and overexpressed to provide a commercial source of phytase. This monomeric enzyme, a type of histidine acid phophatase (HAP), has been characterized and extensively studied. HAPs are also found in other fungi, plants, and animals. Several microbial and plant HAPs are known to have significant phytase activity. A second A. niger phytase (phyB), a tetramer, is known and, like phyA, has had its X-ray crystal structure determined. The model provided by this crystal structure research has provided an enhanced understanding of how these molecules function. In addition to the HAP phytase, several other phytases that lack the unique HAP active site motif RHGXRXP have been studied. The best known group of the non-HAPs is phytase C (phyC) from the genus Bacillus. While a preliminary X-ray crystallographic analysis has been initiated, no enzymatic mechanism has been proposed. Perhaps the pivotal event in the last century that created the need for phytase was the development of modern fertilizers after the Second World War. This fostered a transformation in agriculture and a tremendous increase in feed-grain production. These large quantities of cereals and meal in turn led to the transition of one segment of agriculture into "animal agriculture," with their its animal production capability. The huge volumes of manure spawned by these production units in time exceeded both the capacity of their crops and crop lands to utilize or bind the increased amount of phosphorus. Nutrient runoff from this land has now been linked to a number of blooms of toxin-producing microbes. Fish kills associated with these blooms have attracted public and governmental concern, as well as greater interest in phytase as a means to reduce this phosphorus pollution. Phytase research efforts now are focused on the engineering of an improved enzyme. Improved heat tolerance to allow the enzyme to survive the brief period of elevated temperature during the pelletization process is seen as an essential step to lower its cost in animal feed. Information from the X-ray crystal structure of phytase is also relevant to improving the pH optimum, substrate specificity, and enzyme stability. Several studies on new strategies that involve synergistic interactions between phytase and other hydrolytic enzymes have shown positive results.

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Further reduction in the production cost of phytase is also being pursued. Several studies have already investigated the use of various yeast expression systems as an alternative to the current production method for phytase using overexpression in filamentous fungi. Expression in plants is underway as a means to commercially produce phytase, as in biofarming in which plants such as alfalfa are used as "bioreactors," and also by developing plant cultivars that would produce enough transgenic phytase so that additional supplementation of their grain or meals is not necessary. Ultimately, transgenic poultry and hogs may produce their own digestive phytase. Another active area of current phytase research is expanding its usage. One area that offers tremendous opportunity is increasing the use of phytase in aquaculture. Research is currently centered on utilizing phytase to allow producers in this industry to switch to lower-cost plant protein in their feed formulations. Development of a phytase for this application could significantly lower production costs. Other areas for expanded use range from the use of phytase as a soil amendment, to its use in a bioreactor to generate specific myo-inositol phosphate species. The transformation of phytase into a peroxidase may lead to another novel use for this enzyme. As attempts are made to widen the use of phytase, it is also important that extended exposure and breathing its dust be avoided as prudent safety measures to avoid possible allergic responses. In expanding the use of phytase, another important consideration has been achieved. Conservation of the world's deposits of rock phosphate is recognized as important for future generations. Phosphorus is a basic component of life like nitrogen, but, unlike nitrogen, phosphorus does not have a cycle to constantly replenish its supply. It is very likely that the use of phytase will expand as the need to conserve the world's phosphate reserves increases. REFERENCES Abelson, P. H. (1999). A potential phosphate crisis. Science 283, 2015. Alexander Jr., G. R. (1977). The rationale for a ban on detergent phosphate in the Great Lakes Basin. Ciba Found. Sxmp. 57, 269-284. Bazan, J. F., Fletterick, R. J., and Pilkis, S. J. (1989). Evolution of a bifunctional enzyme: 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase. Proc. Natl. Acad. Sci. U.S.A. B6, 9642-9646. Berka, R. M., Rey, M. W., Brown, K. M., Byun, T., and Klotz, A. V. (1998). Molecular characterization and expression of a phytase gene from the thermophilic fungus Thermomyces lanuginosus. AppI. Environ. Microbiol. 64, 4423-4427. Berlan, J.-P., Bertrand, J.-P., and Lebas, L. (1977). The growth of the American "soybean complex." Eur. R. Agric. Eco. 4, 395-416.

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Biotransformation of Unsaturated Fatty Acids to Industrial Products CHING T. HOU

Oil Chemical Research United States Department of Agriculture Peoria, Illinois 61604

I. Introduction II. Monohydroxy Fatty Acids A. Production of lO-Hydroxystearic Acid B. Positional Specificity of Strain DS5 Hydratase C. Hydration of Other Fatty Acids III. Dihydroxy Unsaturated Fatty Acids IV. Trihydroxy Unsaturated Fatty Acids V. Other Reaction Products from the Strain ALA2 System References

I. Introduction

Hydroxy fatty acids a r e important industrial materials. The hydroxy group gives a fatty acid special properties, such as higher viscosity and reactivity compared to other fatty acids. Because of their special chemical attributes, hydroxy fatty acids are used in a wide range of products, including resins, waxes, nylons, plastics, corrosion inhibitors, cosmetics, and coatings. Furthermore, they are used in grease formulations for high-performance military and industrial equipment. Plant systems produce hydroxy fatty acids; presently, imported castor oil and its derivatives are the only commercial source of these industrial hydroxy fatty acids. Ricinoleic and sebacic acids, two castor oil derivatives, are classified by the Department of Defense as strategic and critical materials. 12-hydroxystearates (esters with C10-12 alcohols) are used in leather coatings requiring oil resistance and waterproofing, and in roll leaf foils because of their alcohol solubility and excellent wetting and adhesion to metallic particle (Naughton, 1974). Although a Japanese patent application by Soda and Kido (1987) claimed the production of ricinoleic acid from oleic acid by Bacillus pumilus, other investigators could not repeat their claim, including Soda's own group. Like ricinoleic acid, lesquerella's hydroxy fatty acids also have double bonds and a carboxyl group that provide sites where chemical reactions can occur (Fig. 1). Several newly discovered hydroxy unsaturated fatty ac201 A D V A N C E S IN APPLIED MICROBIOLOGY, V O L U M E 47 0 0 6 5 - 2 1 6 4 / 0 0 $25,00

202

CHING T. HOU

Ricinoleic Acid (18carbon chain, one double b o n d )

OH I CH3(CH2)s-CH--CH2-CH'-CH-(CH2)7C02 H

Lesquerolic Acid (20 carbon chain, one double bond)

OH I CH3(CH2)s-CH- CH2-CH'- CH-(CH2)sCO2H

Densipolic Acid (18 carbonchain, two doublebonds)

OH

Auricolic Acid (20 carbonchain, two doublebonds)

I

CH3CH2-CH=CH-(CH2)2-cH-CH2-CH=CH-(CH2)7c02H OH

I

CH3CH2-CH=CH--(CH2)2-CH--CH2--CH=CH-(CH2)gCO2H

FIG. 1. Hydroxy fatty acids from castor oil and lesquerella oil.

ids were reported to have interesting physiological activity (Hou, 1998a; Kato et al., 1983; Masui et al., 1989). Because of fluctuating supplies and prices for castor oil, some companies have sought alternative raw materials, primarily petroleum-based feedstock. Microbial enzyme systems biotransform unsaturated fatty acids to three types of hydroxy fatty acid products, namely, monohydroxy, dihydroxy, and trihydroxy fatty acids. These new products have potential industrial applications. Microbial oxidation of unsaturated fatty acids was reviewed (Hou, 1995a). II. Monohydroxy Fatty Acid A. PRODUCTION OF 10-HYDROXYSTEARIC ACID

Microbial hydration of unsaturated fatty acid was first reported by Wallen and colleagues in 1962. They found that a pseudomonad isolated from fatty material hydrated oleic acid at the cis 9 double bond to produce 10-hydroxystearic acid (10-HAS) with a 14% yield. The 10-HSA is optically active (Schroepfer and Block, 1963, 1965) and has the D-configuration. Incubation of this organism with oleic acid in a medium enriched with deuterium oxide yielded 10-HSA containing one stable-bound deuterium atom (Schroepfer and Block, 1965). This deuterium was shown to be on carbon atom 9 in the L-configuration. By using soluble cell-free extracts of a pseudomonad, Niehaus and

BIOTRANSFORMATION OF UNSATURATED FATTY ACIDS

203

Schroepfer (1965) were able to demonstrate conversion of oleate to 10-HSA under anaerobic conditions and the reversibility of the reaction. These findings, coupled with the observed stereospecific uptake of one atom of solvent hydrogen into 10-HSA and the lack of conversion of either the cis- or trans-9,10-epoxystearic acid to 10-HAS, are compatible with a mechanism involving hydration of the double bond of oleic acid and rule out epoxide intermediate (Schroepfer, 1966). The same enzyme preparation was later found to catalyze both the hydration of cis- and trans-9,1O-epoxystearic acids to yield threo- and erythro-9,10-dihydroxystearicacid, respectively (Niehaus and Schroepfer, 1967). Niehaus et al. (1970) demonstrated the interconversion of oleic acid and 10-HSA by a soluble (105,000g supernatant) enzyme preparation from a pseudomonad. This further ruled out the possible intermediate role of an epoxide in the overall conversion. The enzymatic conversion of oleate to 10-HSA was observed to proceed under anaerobic conditions, a feature not characteristic of enzymatic epoxidations of olefins. Moreover, neither the DL-cis-9,10-epoxystearate nor the DL-trans-9,10-epoxystearate served as precursors of either oleate or 10-hydroxystearate under the conditions studied. By using a squalene screening method (Yamada et al., 1975), Seo et al. (1981) isolated a culture, Corynebacterium sp. S-401, from soil that hydrates the squalene molecule to form tertiary alcohols. Resting cells of strain S-401 also stereospecifically hydrated oleic acid to 10-ketostearic (10-KSA) and (-)-10R-hydroxystearic acids with 22.4 and 9.1% yield, respectively. Strain S-401 failed to catalyze hydration of oleoamide, oleonitrile, oleyl alcohol, oleyl aldehyde, or cis-9-octadecene. Accordingly, the carboxy group of oleic acid seems to be essential in this reaction. Cells of Rhodococcus rhodochrous also hydrated oleic acid to 10HSA and 10-KSA at 55 and 12% yields, respectively (Litchfield and Pierce, 1986). Hydration of oleic acid to 1O-HSA was also demonstrated in resting cell suspensions of seven Nocardia species under anaerobic conditions (Koritala et al., 1989). Nocardia cholestero]icum NRRL 5769 gave a yield exceeding 90% with optimum conditions at pH 6.5 and 40°C. A minor amount of 10-KSA was detected. The reaction proceeds via hydration of the double bond, as shown by labeling experiments with deuterium oxide and 180-labeled water. The system was specific for fatty acids with cis unsaturation at the 9-position. Anaerobiosis favored bioconversion to 10-HAS (Davis et al., 1969), and higher pH favored bioconversion to 10-KSA (Koritala et al., 1989).

204

CHING T. HOU

So far, microbial hydration of oleic acid was found in Pseudomonas (Wallen et al., 1962), Nocardia (Rhodococcus) (Litchfield and Pierce 1986; Koritala et al., 1989), Corynebacterium (Seo et al., 1981), Sphingobacterium (Kaneshiro et al., 1994) and Micrococcus (Blank et al., 1991) species. The work of E1-Sharkway et al. (1992) considerably extended the genera of microorganisms known to hydrate oleic acid to include a range of eukaryotic organisms. Strains from several other genera, including Absida, Aspergillus, Candida, Mycobacterium, and Schizosaccharomyces, were also found capable of catalyzing the hydration of oleic acid. Resting cells of Saccharomyces cerevisae (baker's yeast, type II:sigma) converted oleic acid to 10-HSA with a 45% yield (E1-Sharkway eta]., 1992). Three other cultures--Nocardia aurantia ATCC 12674, Nocardia sp. NRRL 5646, and Mycobacterium fortuitum UI 53378--converted oleic acid to 10-KSA with 65, 55, and 80% yields, respectively. Small amounts of 10-HSA were also produced by these cultures, except for strain NRRL 5646. The stereospecificity of microbial hydration of oleic acid to 10-HSA was investigated by Yang et al. (1993) based on a 1H-nuclear magnetic resonance spectral analysis of diastereomeric S-(+)-O-acetylmandelate esters of hydroxystearates (E1-Sharkway et al., 1992). While R. rhodochrous ATCC 12674 mediated hydration of oleic acid gave mixtures of enantiomers 10(R)-hydroxystearic acid and 10(S)-hydroxystearic acid, Pseudomonas sp. NRRL B-3266 produced optically pure 10(R)°hydroxystearic acid. The remaining microorganisms investigated stereoselectively hydrated oleic acid to 1O(R)-hydroxystearic acid containing 2 and 18% of the contaminating 10(S)-hydroxystearic acid (Kaneshiro et al., 1994). Although hydration of oleic acid to 10-HSA was investigated using a cell-flee system (Niehaus and Schroepfer, 1965, 1967; Niehaus et al., 1970; Schroepfer 1966), attempts to purify hydratase were not successfill. Very little was known about the physical and chemical properties of oleate hydratase. Purification and characterization of oleate hydratase from Nocardia cholesterolicum NRRL 5767 were investigated by Huang et al. (1991a). The cell-flee extracts obtained after French Press disintegration of the cells and centrifugation were fractionated by amm o n i u m sulfate. The enzyme activity was found in the fraction of 60-75% ammonium sulfate saturation. The enzyme fraction was further purified through Mono-Q ion exchange and Supherose gel filtration column chromatography. The purified enzyme fraction showed a single protein band on acrylamide gel electrophoresis. The hydration proceeded linearly for 6 hours. The optimum pH for the enzyme reac-

BIOTRANSFORMATIONOF UNSATURATEDFATTYACIDS

205

tion was between 6.5 and 7. The Krn value for the hydratase reaction at 30°C was 2.82 x 10 -4 M. The molecular weight estimated from a Superose HR 10/30 gel filtration column was about 120,000 daltons and from SDS-PAGE about 32,000 daltons (Huang et al., 1991b), suggesting that oleate hydratase is a tetramer, and is composed of four identical subunits. Lanser (1993) reported the conversion of oleic acid to 10-ketostearic acid by microorganisms from other genera (e.g., Staphylococcus spp.). The yield was greater than 90% with less than 5% of byproduct--10hydroxystearic acid. In addition, Kuo et al. (1999) reported the conversion of oleic acid to 10-ketostearic acid by Sphingobacterium sp. strain 022. Flavobacterium sp. DS5 converted oleic acid to 10-KSA in 85% yield Hou (1994a). Optimum time, pH, and temperature for the production of 10-KSA are: 36 hr, 7.5, and 30°C, respectively. A small amount of 10-HSA (about 10% of the main product, 10-KSA) is also produced during bioconversion. 10-KSA is not further metabolized by strain DS5 and accumulates in the medium. In contrast to growing cells, resting cell suspensions of strain DS5 produced 10-HSA and 10-KSA in a ratio of 1:3. The cell-free crude extract obtained from ultrasonic disruption of the cells converts oleic acid to mainly 10-HSA (10-HSA/10-KSA = 97/3). This result strongly suggested that oleic acid was converted to 10-KSA via 10-HSA. The stereochemistry of product 10-HSA from strain DS5, determined by 1H-NMR of the mandelate esters of methyl10-hydroxystearate obtained from DS5, showed 66% enantiomeric excess in 10(R) form. The Flavobacterium DS5 enzyme system also catalyzed conversion of linoleic acid. In contrast to oleic acid substrate that yielded mainly the keto product, linoleic acid substrate yielded mainly 10-hydroxy12(Z)-octadecenoic acid (10-HOA) with a 55% yield (Hou, 1994b). The optimum conditions for the production of 10-HOA were: pH 7.5, temperature 20 to 35°C, and 36 hours of incubation. Two minor products produced were 10-methoxy-12-octadecenoic acid and 10-keto-12-octadecenoic acid (10-KOA). Strain DS5 oxidized unsaturated but not saturated fatty acids. The relative activities were in the following order: oleic > palmitoleic > arachidonic > linoleic > linolenic > 7-1inolenic > myristoleic acids. With the resting cell suspension, the 10-HOA/10-KOA ratio was 97/3. Less 10-KOA was produced compared to that of growing cells. The cells were disrupted with ultrasonic oscillation and centrifuged to obtain cell-free crude extract. The linoleic acid conversion enzyme(s) resided

206

CHING T. HOU

in the cell-flee crude extract, and only 10-HOA was produced from linoleic acid. B. POSITIONAL SPEGIFICITYOF STRAIN DS5 HYDRATASE

From substrate specificity studies (Hou, 1994a,b), it seems that DS5 hydratase hydrates a specific carbon position of the unsaturated fatty acid substrates. In order to clear up this point and the effect of substrate carbon chain length on the strain DS5 hydratase activity, we studied the hydration of mono-, di, and tri-unsaturated C18 fatty acids as well as other carbon chain-length monounsaturated fatty acids. Strain DS5 converted o~-linolenic acid to 10-hydroxy-12,15-octadecadienoic acid and a minor product--10-keto-12,15-octadecadienoic acid (Hou, 1994b). Strain DS5 also converted 7-1inolenic acid to 10-hydroxy-6(Z),12(Z)-octadecadienoic acid. The enzyme hydrated 9-unsaturation but did not alter the original 6,12-unsaturations. Strain DS5 converted myristoleic acid to two products: 10-keto myristic and 10-hydroxymyristic acids. Palmitoleic acid also gave two bioconversion products: 10-ketopalmitic and 10-hydroxypalmitic acid. Previously, the strain DS5 bioconversion products from oleic and linoleic acids were identified as 10-ketostearic (Hou, 1994a) and 10-hydroxy-12(Z)-octadecenoic (Hou, 1994b) acid, respectively. It is interesting to note that all unsaturated fatty acids tested are hydrated at the 9and 10-positions with the oxygen functionality at C-10, despite their varying degree of unsaturation. DS5 hydratase was not active on saturated fatty acids, and other non-9(Z)-unsaturated fatty acids such as elaidic [9(E)-octadecenoic], arachidonic [5(E),8(E),11(E),14(E)-eicosatetraenoic], and erucic [13(E)-docosanoic] acids (Hou, 1995b,c). It was concluded that DS5 hydratase is indeed a C-10-positional-specific enzyme. The fact that elaidic acid was not hydrated indicates that the unsaturation must be in the cis configuration for DS5 hydratase activity. The strain DS5 system produced more keto product from palmitoleic and oleic acids and more hydroxy product from myristoleic, linoleic, and a- and 7-1inolenic acids. The reason for this product preference is not clear. Among the 18-carbon unsaturated fatty acids, additional double bonds on either side of position C-10 lower enzyme hydration activity. A literature search revealed that all known microbial hydratases hydrate oleic and linoleic acids at the C-10 position (Fig. 2). Therefore, the positional specificity of microbial hydratases might be universal. C. HYDRATIONOF OTHER FATTY ACIDS

Hydration of unsaturated fatty acids other than oleic acid was also reported. Wallen et al. (1971) prepared three new unsaturated 10-hydroxy fatty acids, all optically active, by the anaerobic microbial hydra-

BIOTRANSFORMATIONOF UNSATURATEDFATTYACIDS

1. Oleic acid ~

207

OH I H3C-(CH2)7-CH-CH 2- (CH2)7COOH

0

II

H3C-(CH2)7 - C- CH2-(cH 2)7c00H OH i 2. Linoleicacid ----~ H3C-(CH2)4 -CH~- CH-CH2-CH-CH2--(CH2)7COOH OH I 3. o~-Linolenicacid--~ H3C-CH 2 -CH--CH--CH2-CH=CH - C H 2 - C H - CH2--(CH2)7COOH

OH I

4. y-Linolenicacid---~ H3C-(CH2) 4-CH-CH-CH2-CH-CH2-CH2-CH'-CH-(CH2)4COOH

Fic. 2. Bioconversionproducts fromunsaturatedfatty acids by strain DS5 hydratase.

tion of a cis-9-double bond. Substrates that formed these new hydroxy fatty acids were linoleic, linolenic, and ricinoleic acids. The yields were linoleic acid to 10-hydroxy-12(Z)-octadecenoic acid, 20 mol%; linolenic acid to 10-hydroxy-12(Z),15(Z)-octadecadienoic acid, 21 mol%; and ricinoleic acid to 10,12-dihydroxystearic acid, 41 mol%. GieselBuhler et al (1987) reported production of 10-hydroxy-12-octadecenoic acid from linoleic acid by resting cells of Acetobacterium woodii through hydration. Litchfield and Pierce (1986) claimed that cells of Rhodococcus rhodochrous catalyzed hydration of linoleic acid to 10-hydroxy-12-octadecenoic acid at 22% yield with 10-keto-12-octadecenoic acid as a coproduct. In the early stages of cell growth, the hydration enzyme is inducible by the presence of oleic acid. Using washed resting cells suspension of Nocardia cholesterolicum under anaerobic conditions, Koritala and Bagby (1992) more recently reported the hydration of linoleic and linolenic acids to 10-hydroxy12(Z)-octadecenoic (yield 71%) and 10-hydroxy-12(Z),15(Z)-octadecadienoic acids (yield 77%), respectively. The production of 1O-hydroxy fatty acids by hydratase from various microbes is summarized in Table I. The hydroxylation of oleic acid has also been reported. Lanser et al. (1992) found that two strains of Bacillus pumilus (NRRL BD-174 and BD-226) produced 15-, 16-, and 17-hydroxy-9-cis-octadecenoic acids.

208

CHING T. HOU

III. Dihydroxy Unsaturated Fatty Acids As part of a screening program to find new industrial chemicals from vegetable oils and their component fatty acids, a new bacterial strain (PR3) was isolated. This strain converted oleic acid to a new compound: 7,10-dihydroxy-8(E)-octadecenoic acid (DOD) (Hou and Bagby, 1991; Hou et al., 1991). Isolated from a water sample at a pig farm in Morton, Illinois, strain PR3 formed a smooth, round, white colony on agar plate. The bacteria were motile, short, and rod-shaped with multiple polar flagella, and could not grow anaerobically The cells were oxidase-positive. Strain PR3 was classified in the genus Pseudomonas (Hou and Bagby, 1991). The strain produced fluorescein on King's medium B as well as pyocyanin on King's medium A, suggesting that the organism was a strain ofP. aeruginosa. Further identification was conducted with DNA reassociation measurements (Hou et al., 1993). The chemical structure of DOD was determined by GC/MS, FTIR, and NMR (Hou et al., 1991). Its production from oleic acid reached a maximum after 48 hr of incubation with a yield of 63% (Hou, 1999). The yield was later improved to greater than 80% by modifying the culture medium and reaction parameters (Kuo et al., 1998). The production of

TABLE I HYDRATASESFROMVARIOUSMICROBESTHAT PRODUCE10-HYDROXYPRODUCT Microbes Pseudomonas Corynebacterium Rhodococcus Bacillus

Nocardia Micrococcus S a rcin a

Aspergillus Candida Mycobacterium Schizosaccharomyces Staphylococcus Flavobacterium Sphingobacterium

References Wallen et al. (1962) Seo et al. (1981) Litchfield and Pierce (1986) Soda and Kido (1987) Koritala et al. (1989) Blank eta]. (1991) Blank et al. (1991) EI-Sharkaway et a]. (1992) E1-Sharkaway et al. (1992) E1-Sharkaway et al. (1992) E1-Sharkaway et a]. (1992) E1-Sharkaway et al. (1992) Lanser (1993) Hou (1994a,b) Kaneshiro et a]. (1994)

BIOTRANSFORMATION OF UNSATURATED FATTY ACIDS

209

DOD with a cell-free enzyme preparation was also demonstrated at a greater than 90% yield. The absolute configuration of DOD was originally determined with the aid of circular dichroism to be R,R (Knothe et at,, 1992). Recently, an alternative method to CD was used to determine the absolute configuration of DOD that involved formation of the (-)-methoxycarbonyl (MCO) derivative of the two hydroxyls, oxidative cleavage of the double bond (Fig. 3) (Hamberg, 1971; Hamberg ot al., 1986), and gas chromatographic analysis of the two methyl esterified diastereomeric fragments: methyl-2-MCO-decanoate and dimethyl-2-MCO-octandioate. As described by previous workers, the 2(S)-MCO derivatives elute at earlier

•••••V• ~

OCH3

. ) (-)-menthoxycarbonylchloride

O

~

~

~

~

V

O

~

~

OCH3

0

HMnO:-Aceticacid

0

Y

"0~ ~-0 o

o

o==~

~

o

.~oo.. 2 . . . . . . ~ . ~. -i1- v

v

v

"OCH3

o

FIG. 3. Method for producing (-)-methoxycarbonyl (MCO) derivatives for chiral analysis by GC.

210

CHING T. HOU

times by GC than the 2(R)-MCO derivatives. Comparing the GC analysis of the two MCO derivatives obtained from DOD with that obtained from a partially racemized sample, DOD was determined to be 7(S),10(S)-dihydroxy-8(E)-octadecenoic acid (Gardner and Hou, 1999). Production of DOD from oleic acid is unique in that it involves addition of two hydroxy groups at two positions and rearrangement of the double bond of the substrate molecule. Subsequent investigation of reactions catalyzed by PR3 led to isolation of another new compound: 10-hydroxy-8-octadecenoic acid (HOD) (Hou and Bagby, 1992). From the structure similarity between HOD and DOD, it is likely that HOD is an intermediate in the formation of DOD from oleic acid by strain PR3. Kinetic studies (Hou and Bagby, 1992) showed that conversion from HOD to DOD is not a rate-limiting step. The bioconversion pathway for production of DOD from oleic acid is postulated with HOD as the intermediate, and unsaturation at the 8-position carbon is possibly in cis configuration. We more recently determined that the rearranged double bond of HOD was in trans form by NMR and FTIR analyses (Kim et al., 2000a). The absolute configuration of the hydroxy group at carbon 10 of HOD was also determined to be in the S configuration by methoxycarbonyl (MCO) derivation of the hydroxy group followed by oxidative cleavage of the double bond and methyl esterification (Hamberg et al., 1986). This result coincided with our other findings that the main final product DOD represented 7(S),lO(S)-dihydroxy configuration (Gardner and Hou, 1999). In addition, a minor isomer of HOD (about 3%) with 10(R) configuration was also detected. A postulated pathway for bioconversion of oleic acid to DOD by strain PR3 is depicted in Figure 4a. Oleic acid is first converted to HOD. During this step, one hydroxyl group is introduced at C-10(S) and a double bond is shifted from C-9 cis to C-8 trans, suggesting that there may be at least two or more enzymes involved in this first step for cis-trans-shifted isomerization of the double bond and further hydroxylation introducing a hydroxyl group at C-7(S). In Flavobacterium sp. DS5, a C-10-position-specific and cis-specific hydratase was involved in hydration of unsaturated fatty acids in which the C-10 hydroxyl group was introduced with removal of the C-9 cis double bond, typical of the hydration reaction of fatty acids Hou (1995b,c). It is unlikely that a hydratase is involved in the PR3 reaction in that the double bond at C-9 of substrate was retained as a shifted trans-configured form during hydroxylation until the formation of DOD. A similar type of compound, dihydroxyoctadecenoic acid, is produced by Pseudomonas 42A2. However, the positions for the double bond and hydroxy groups in their report were determined later

BIOTRANSFORMATIONOF UNSATURATEDFATTY ACIDS a.

Oleicacid

211

91i* ~ v / ~ / ~ j ~ C O O H I

Hydroxylationwithdoublebondmigration

i HOD ~

C

O

O

HO Hydroxylation~ DOD ~

C

O

H OH

O

H

HO b.

91~ i ' ~ ~ / ~ C ° ° H lo %..d'.,./%~-~./ OH I

Ricinoleicacid

Hydroxylationwithdoublebondmigration

i DHOD~

C

O

Hydroxylation~

O

H

OH

TOD ~ ~ ~ ~ ~ % j C O O H

H" FIG.4. Biosynthesis of (a) 7,10-dihydroxy-8(E)-octadecenoicacid produced from oleic acid and (b) TOD from ricinoleic acid by Pseudomonas aeruginosa PR3.

(Mercade et al., 1988; de Andres et al., 1994; Guerrero et al., 1997). This group also reported the oxidation of oleic acid to (E)-10-hydroperoxy8-octadecenoic acid and (E)-lO-hydroxy-8-octadecenoic acid by 42A2 (Guerrero et al., 1997). Strain PR3 converts ricinoleic acid to a more polar compound. The structure of this n e w product was determined by GC/MS, FTIR, and NMR to be 7,10,12-trihydroxy-8(E)-octadecenoic acid (TOD) at 35% yield (Kuo et al., 1998; Kuo and Hou 1999). The reaction mechanism is the same as that for the conversion of oleic acid to DOD. Another new c o m p o u n d was isolated from the ricinoleic acid-PR3 system; its

212

CHING T. HOU

structure was determined by MS, FTIR, and NMR to be 10,12-dihydroxy-8(E)-octadecenoic acid (DHOD). Evidence obtained strongly suggested that DHOD is an intermediate in the bioconversion of ricinoleic acid to TOD (Fig. 4b). The optimum conditions for production of DHOD were pH 6.5, temperature 25°C, and 30 hr (Kim et al., 2000b). Physiologic activity tests revealed that DOD has some antibiotic activity against Bacillus subtilis and a common yeast pathogen, Candida albicans.

IV. Trihydroxy Unsaturated Fatty Acids A new compound, 12,13,17-trihydroxy-9(Z)-octadecenoic acid, was produced from linoleic acid by a new microbial isolate. The microorganism that performs this unique reaction was isolated from a dry soil sample collected from McCalla, Alabama. Strain ALA2 is a Gram-positive nonmotile rod (0.5 x 2 jam) classified as C]avibacter sp. ALA2 (Hou eta]., 1997). The chemical structure of the new compound was determined by MS, FTIR, and NMR. The chemical ionization mass spectrum of the methyl ester prepared with diazomethane gave a molecular ion of m/z 345. Fragments of 327 (M-18) and 309 (M-2 x 18) were also seen. The electron impact spectrum of the methylated product provided more fragments for structural analysis. Large fragments corresponding to o~-cleavage with ions m/z 227 (25%) and 129 (100%) place the two hydroxy groups at positions C-12 and C-13, and the third hydroxy group at a position higher than carbon 13. Proton and 13C NMR analyses further confirmed the structure. Resonance signals (ppm) and corresponding molecular assignments given in Table II located three hydroxy groups at C-12, C-13, and C-17, and further confirmed the identity of the bioconversion product as 12,13,17-trihydroxy-9(Z)-octadecenoic acid. The coupling constant of 10.7 Hz at C-9,10 confirmed our infrared data that the unsaturation is in cis configuration (Hou, 1996, 1998a) (Fig. 5). It is interesting to note that all three types of hydroxy fatty acids-mono-, di-, and trihydroxy--were discovered by scientists at the U.S.D.A.'s National Center for Agricultural Utilization Research (Wallen eta]., 1962; Hou and Bagby, 1991; Hou eta]., 1991; Hou, 1996). Production of trihydroxy unsaturated fatty acids in nature is rare. Trace amounts have been found in plants. 8,9,13-trihydroxy docosanoic acid has been produced by yeast as an extracellular lipid (Stodola et al., 1965). 9,10,13-trihydroxy-11(E)- and 9,12,13-trihydroxy-10(E)-octade-

BIOTRANSFORMATION

OF UNSATURATED FATTY ACIDS

213

T A B L E II PROTON AND 13C NUCLEAR MAGNETIC RESONANCE SIGNALS AND MOLECULAR ASSIGNMENTS FOR B1OCONVERSION PRODUCT Resonance C h e m i c a l shifts ( p p m ) / c o u p l i n g (Hz) Carbon number

1'3C

1

174.4

2

34.1

2.29

t

3

24.9

1.60

m

4

29.0

1.30

bs

5

29.0

1.30

bs

6

29.0

1.30

bs

7

29.5

1.30

bs

8

27.3

2.04

m

J8,9 = 7.0

9

133.8

5.55

m

J9,10 = 10.7

10

J10,11 = 7.2

Proton

-

124.6

5.40

m

11 12 b

31.7 73.7

2.29 3.48

m m

136

73.8

3.48

m

14

33.5

1.48

m

15

21.7

1.30

bs

16

39.1

1.45

m

17

68.0

3.82

m

18

23.5

1.18

d

OCH3

51.5

3.65

s

J2,3 = 7.4 a

J17,18 = 6.1

a C o u p l i n g c o n s t a n t J i n Hz. bShift m a y b e i n t e r c h a n g e d .

cenoic acids were detected in beer (Graveland, 1970). It has been suggested that these trihydroxy fatty acids are formed from linoleic acid during malting and mashing of barley (Baur and Grosch, 1977). Gardner et al. (1984) reported the production of diastereomeric (Z)-11,12,13-trihydroxy-9-octadecenoic acids and four isomers of (E)-9,12,13(9,10,13)trihydroxy-10(11)-octadecenoic acids by acid-catalyzed transformation of 13(S)-hydroperoxylinoleic acid. Hydroxy and epoxy unsaturated fatty acids present in some rice cultivars acted as antifungal substances and were active against rice blast fungus (Kato et al., 1983, 1984). It was postulated that these fatty acids were derivatives of linoleic and

214

CHING T. HOU

OH

//

o

oH

-(oo

12,13,17-Trihydroxy-9(Z)-octadecenoic acid

FIG. 5. 12,13,17-trihydroxy-9(Z)-octadecenoic acid produced from linoleic acid by Clavibacter sp. ALA2.

linolenic acid hydroperoxides. Mixed hydroxy fatty acids have also been isolated from Sasanishiki rice plants that suffered from rice blast disease, and they were shown to be active against the fungus (Kato et al., 1985). Their structures were identified as 9S,12S,13S-trihydroxy10-octadecenoic acid and 9S,12S,13S-trihydroxy-10,15-octadecadienoic acid (Kato eta]., 1986; Suemune et al., 1988). 9,12,13-trihydroxy10(E)-octadecenoic acid isolated from Colocasia antiquorum inoculated with Ceratocystis fimbriata showed anti-black rot fungal activity (Masui et al., 1989). Other than extraction from plant materials, our discovery is the first report on production of trihydroxy unsaturated fatty acids by microbial transformation, as well as the first evidence that the structure of THOA resembles that of plant self-defense substances. Therefore, the biological activity of THOA at 200 p p m was tested against many plant pathogenic fungi (Hou, 1998b). The results, expressed as percentage growth inhibition, are shown in Table III. THOA inhibited the growth of Erisyphe graminis f. sp. tritici (common called "wheat powdery mildew"), Puccinia recondita (wheat leaf rust), Phytophthora infestans (potato late blight), and Botrytis cinerea (cucumber botrytis). It seems that the position of the hydroxy groups on the fatty acid molecule plays an

215

BIOTRANSFORMATION OF UNSATURATED FATTY ACIDS TABLE III ANTIFUNGALACTIVITYOF THOA (200 ppm) Fungus

Disease

Erisyphe graminis Puccinia recondita Pseudocercosporella herpotrichoides Septoria nodorum Pyricularia grisea Rhizoctonia solani Phflophthora infestans Botrytis cinerea

% inhibition

Wheat powdery mildew Wheat leaf rust Wheat foot rot

77 86 0

Wheat glume blotch Rice blast Rice sheath blight Potato late blight Cucumber botrytis

0 0 0 56 63

important role in determining activity against certain plant pathogenic fungi. V. Other Reaction Products from the Strain ALA2 System A GC analysis of typical reaction products produced from linoleic acid by strain ALA2 is shown in Figure 6. In addition to the main reaction product at GC retention time (Rt) 24 min, there were small amounts of

100 123

8

i 0

0

10

20 Time (rnin)

30

40

FIG. 6. A typical gas chromatogram of strain ALA2 reaction products: (1) internal standard, palmitic acid; (2) substrate, linoleic acid; (3-7) unknown; (8) product 12,13,17trihydroxy-9(Z)-octadecenoic acid.

216

CHING T. HOU

(a)

(b)

~ ~

OH COOH

OH

OH COOH

(c)

(d)

FIG. 7. Chemical structures of minor reaction products obtained from linoleic acidstrain ALA2 system: (a) 12-[5-ethyl-2-tetrahydrofuranyl]-12-hydroxy-9(ZJ-dodecenoic acid; (b) 12-[5-ethyl-2-tetrahydrofuranyl]-7,12-dihydroxy-9(Z)-dodecenoic; (c) 12,17; 13,17-diepoxy-9(Z)-octadecenoic acid; (d) 12,17;13,17-diepoxy-7-hydroxy-9(Z)-octadecenoic acid.

products at 7, 10, 13, and 17 min. Mass spectrum analysis of fragments indicated that they are 12-[5-ethyl-2-tetrahydrofuranyl]-12-hydroxy9(Z)-dodecanoic acid for Rtl0 and 12-[5-ethyl-2-tetrahydrofuranyl]7,12-dihydroxy-9(Z)-dodecanoic for Rt17 (Fig. 7a,b). The yield of the main product (THOA) was 35%, and the relative amounts of products produced were 9 (THOA), 1 (Rtl0), and 1.3 (Rt17) (Hou et al., 1998). The other two minor products--GC Rt7 and 13--from the ALA2-1inoleic acid system were isolated, and their chemical structures determined by GC/MS and NMR. Products Rt7 and Rt13 were found to be the novel bicyclic fatty acids 12,17;13,17-diepoxy-9(Z)-octadecenoic acid and 12,17;13,17-diepoxy-7-hydroxy-9(Z)-octadecenoic acid, respectively (Hou et al., 2000). These structures are shown in Figure 7c,d. The relationship among these products in the metabolic pathway of the strain ALA2 system is currently under investigation. The optimum conditions for bioconversion of linoleic acid to THOA are a pH of 7.0 and a production temperature of 30°C (Hou et al., 1997). Maximum THOA production is found after 5-6 days of reaction.

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REFERENCES Baur, C., and Grosch, W. (1977). Study on the taste of di-, tri- and tetrahydroxy fatty acids. Z. Lebensm. Unters. Forseh. 165, 82-84. Blank, W., Takayanagi, H., Kido, T., Meussdoerffer, F., Esaki, N., and Soda, K. (1991). Transformation of oleic acid and its esters by Sarcina lutea. Agric. Biol. Chem. 55, 2651-2652. Davis, E. N., Wallen, L. L., Goodin, J. C., Rohwedder, W. K,, and Rhodes, R. A. (1969). Microbial hydration of eis-9-alkenoic acids. Lipids 4, 356-362. de Andres, C., Mercade, E., Guinea, J., and Manresa, A. (1994). 7,10-dihydroxy-8E-octadecenoic acid produced by Pseudomonas sp. 42A2: Evaluation of different cultural parameters of the fermentation. World ]. Microbiol. Biotechnol. 10, 106-109. E1-Sharkawy, S. H., Yang, W., Dostal, L., and Rosazza, J. P. N. (1992). Microbial oxidation of oleic acid. App]. Environ. Microbiol. 58, 2116-2122. Gardner, H. W., and Hou, C. T. (1999). All (S) stereo configuration of 7,10-dihydroxy-8(E)~ octadecenoic acid from bioconversion of oleic acid by Pseudomonas aeruginosa. J. Am. Oi] Chem. Soc. 76, 1151-1156. Gardner, H. W., Nelson, E. C., Tjarks, L. W., and England, R. E. (1984). Acid-catalyzed transformation of 3(S)-hydroperoxy-linoleic acid into epoxyhydroxyoctadecenoic acid and tri-hydroxyoctadecenoic acids. Chem. Phys. Lipids 35, 87-101. Giesel-Buhler, H., Bartsch, O., Hueifel, H., Sahm, H., and Schmid, R. (1987). In "Proceedings of the International Symposium on 'Biocatalysis in Organic Media,' Wageningen" (Laane, Tramper, and Lilly, eds.), p. 241. Elsevier, Amsterdam. Graveland, A. (1970). Enzymatic oxidation of linoleic acid and glycerol-l-monolinoleate in doughs and flour-water suspensions. J. Am. Oil Chem. Soc. 47, 352-361. Guerrero, A., Casals, I., Busquets, M., Leon, Y., and Manresa, A. (1997). Oxidation of oleic acid to (E)-10-hydroperoxy-8-octadecenoic and (E)-10-hydroxy-8-octadecenoic acids by Pseudomonas sp. 42A2. Biochim. Biophys. Acta 1347, 75-81. Hamberg, M. (1971). Steric analysis of hydroperoxides formed by lipoxygenase oxygenation of linoleic acid. Anal. Biochem. 43, 515-526. Hamberg, M., Herman, R. P., and Jacobson, U. (1986). Stereochemistry of two epoxy alcohols from Saprolegnia parasitica. Biochem. Biophys. Acta 879,410-418. Hou, C. T. (1994a). Production of 10-ketostearic acid from oleic acid by a new microbial isolate, Flavobacterium sp. NRRL B-14859. Appl. Environ. Microbiol. 60, 3760-3763. Hou, C. T. (1994b). Conversion of linoleic acid to 10-hydroxy-12(Z)-octadecenoic acid by Flavobacterium sp. DS5. J. Am. Oil Chem. Soc. 71, 975-978. Hou, C. T. (1995a). Microbial oxidation of unsaturated fatty acids. Adv. AppI. Microbial. 41, 1-23. Hou, C. T. (1995b). Production of hydroxy fatty acids from unsaturated fatty acids by Flavobacterium sp. DS5 hydratase, a C-10 positional- and cis-unsaturation-specific enzyme. J. Am. Oil Chem. Soc. 72, 1265-1270. Hou, C. T. (1995c). Is strain DS5 hydratase a C-10 positional specific enzyme? Identification of bioconversion products from c~- and y-linolenic acids by Flavobacterium sp. DS5. ]. Ind. Microbiol. 14, 31-34. Hou, C. T. (1996). A novel compound, 12,13,17-trihydroxy-9(Z)-octadecenoic acid, from linoleic acid by a new microbial isolate Clavibacter sp. ALA2. L Am. Oil Chem. Soc. 73, 1359-1362. Hou, C. T. (1998a). "12,13,17-trihydroxy-9(Z)-Octadecenoic Acid and Derivatives and Microbial Isolate for Production of the Acid." U.S. Patent 5,852,196. Hou, C. T. (1998b). Antimicrobial activity of hydroxy fatty acids. Paper presented at SIMB Annual Meeting, Denver.

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Hou, C. T. (1999). "Microbial Production of a Novel Compound, 7,10-Dihydroxy-8-Octadecenoic Acid from Oleic Acid." U.S. Patent 5,900,496. Hou, C. T., and Bagby, M. O. (1991). Production of a new compound, 7,10-dihydroxy8(E)-octadecenoic acid, from oleic acid by Pseudomonas sp. PR3. J. Ind. Microbiol. 7, 123-130. Hou, C. T., and Bagby, M. O. (1992). 1O-hydroxy-8(Z)-octadecenoic acid, an intermediate in the formation of 7,1O-dihydroxy-8(E)-octadecenoic acid from oleic acid by Pseudomonas sp. PR3. J. Ind. Microbiol. 9, 103-107. Hou, C. T., Bagby, M. O., Platner, R. D., and Koritala, S. (1991). A novel compound, 7,10-dihydroxy-8(E)-octadecenoic acid, from oleic acid by bioconversion, f. Am. Oil Chem. Soe. 68, 99-101. Hou, C. T., Nakamura, L. K., Weisleder, D., Peterson, R. E., and Bagby, M. O. (1993). Identification of NRRL strain B-18602 (PR3) as Pseudomonas aeruginosa and effect of phenazine-l-carboxylic acid formation on 7,10-dihydroxy-8(E)-octadecenoic acid accumulation. World J. Microbiol. BiotechnoL 9, 570-573. Hou, C. T., Brown, W., Labeda, D. P., Abbott, T. P., and Weisleder, D. (1997). Microbial production of a novel trihydroxy unsaturated fatty acid from linoleic acid. J. Ind. Microbiol. BiotechnoL 19, 34-38. Hou, C. T., Gardner, H., and Brawn, W. K. (1998). Production of polyhydroxy fatty acids from linoleic acid by Clavibacter sp. ALA2. J. Am. Oil Chem. Soc. 75, 1483-1487. Hou, C. T., Gardner, H., Weisleder, D., and Brown, W. (2000). Biotransformation of linoleic acid by C]avibacter ALA2: Isolation and characterization of bicyclic fatty acids. Abstr. Ann. Mtg. Am. Oil Chem. Soc., San Diego. Abstr. #B-2. Huang, J.-K., Hou, C. T., and Bagby, M. O. (1991a). Isolation and characterization of oleate hydratase from Noeardia cholesterolicum NRRL 5767. Abstr. Ann. Mtg. Soc. Ind. Microbiol., Philadelphia. Abstr. #29. Huang, J.-K., Hou, C. T., and Bagby, M. O. (1991b). Purification and characterization of oleate hydratase from Nocardia cholesterolicum NRRL 5767: Physical and chemical properties. Abstr. 34th West Central States Biochem. Conf., Ames, Iowa. Abstr. #51. Kaneshiro, T., Huang, J.-K., Weisleder, D., and Bagby, M. O. (1994). 10R-hydroxystearic acid production by a novel microbe, NRRL B-14797, isolated from compost. J. Ind. Microbial. 13, 351-355. Kato, T., Yamaguchi, Y., Uyehara, T., Yokoyama, T., Namai, T., and Yamanaka, S. (1983). Self-defensive substances in rice plant against rice blast disease. Tetrahedron Lett. 24, 4715-4718. Kato, T., Yamaguchi, Y, Abe, N., Uyeharaa, T., Nakai, T., Yamanaka, S., and Harada, N. (1984). Unsaturated hydroxy fatty acids, the self-defensive substances in rice plant against rice blast disease. Chem. Lett. 25,409-412. Kato, T., Yamaguchi, Y., Abe, N., Uyehara, T., Namai, T., Kodama, M., and Shiobara, Y. (1985). Structure and synthesis of unsaturated trihydroxy C-18 fatty acids in rice plant suffering from rice blast disease. Tetrahedron Lett. 26, 2357-2360. Kato, T., Yamaguchi, Y., Ohnuma, S., Uyehara, T., Namai, T., Kodama, M., and Shiobara, Y. (1986). Structure and synthesis of 11,12,13-trihydroxy-9(Z),15(Z)-octadecadienoic acids from rice plant suffering from rice blast disease. Chemistry Lett., pp. 577-580. Kim, H., Gardner, H. W., and Hou, C. T. (2000a). 10(S)-hydroxy-8(E)-octadeeenoic acid, an intermediate in the conversion of oleic acid to 7,10-dihydroxy-8(E)-octadecenoic acid. J. Am. Oil Chem. Soc. 77, 95-99.

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Kim, H., Kuo, T. M., and Hou, C. T. (2000b). Production of 10,12-dihydroxy-8(E)-octadecenoic acid, an intermediate in the conversion of ricinoleic acid to 7,10,12-trihydroxy-8(E)-octadecenoic acid by Pseudomonas aeruginosa PR3. Ind. Microbiol. Biotechnol. 24, 167-172. Knothe, G., Bagby, M. O., Peterson, R. E., and Hou, C. T. (1992). 7,1O-hydroxy-8(EJ-octadecenoic acid: Stereochemistry and a novel derivative, 7,10-dihydroxyoctadecanoic acid. J. Am. Oil Chem. Soc. 69, 367-371. Koritala, S., and Bagby, M. O. (1992). Microbial conversion of linoleic and linolenic acids to unsaturated hydroxy fatty acids. J. Am. Oil. Chem. Soe. 69, 575-578. Koritala, S., Hosie, L., Hou, C. T., Hesseltine, C. W., and Bagby, M. O. (1989). Microbial conversion of oleic acid to 10-hydroxystearic acid. Appl. MicrobioI. Biotechnol. 32, 299-304, Kuo, T. M., and Hou, C. T. (1999). Bioconversion of unsaturated fatty acid by Pseudomonas aeruginosa PR3. Recent Res. Dev. Oil Chem. 3, 1-10. Kuo, T. M., Manthey, L. K., and Hou, C. T. (1998). Fatty acid bioconversion by Pseudomonas aeruginosa PR3. J. Am. Oil Chem. Soc. 75,875-879. Kuo, T. M., Lanser, A. C., Kaneshiro, T., and Hou, C. T. (1999). Conversion of oleic acid to 10-ketostearic acid by Sphingobacterium sp. strain 022. J. Am. Oil Chem. Soc. 76, 709-712. Lanser, A. C. (1993). Conversion of oleic acid to 10-ketostearic acid by Staphylococcus sp. J. Am. Oil Chem. Soc. 70, 543-545. Lanser, A. C., Plattner, R. D., and Bagby, M. O. (1992). Production of 15-, 16-, and 17-hydroxy-9-octadecenoic acids by bioconversion of oleic acid with Bacillus pumilus. J. Am. Oil Chem. Soc. 69, 363-366. Litchfield, J. H., and Pierce, G. E. (1986). "Microbiological Synthesis of Hydroxy-Fatty Acids and Keto-Fatty Acids." U.S. Patent 4,582,804. Masui, H., Kondo, T., and Kojima, M. (1989). An antifungal compound, 9,12,13-trihydroxy-(E)-10-octadecenoic acid, from Colocasia antiquorum inoculated with Ceratocystis fimbriata. Phytochemistry 28, 2613-2615. Mercade, E., Robert, M., Espuny, M. J., Bosch, M. P., Manreesa, M. A., Parra, J. L., and Guinea, J. (1988). New surfactant isolated from Pseudomonas sp. 42A2. J. Am. Oil Chem. Soe. 65, 1915-1916. Naughton, F. C. (1974). Production, chemistry and commercial applications of various chemicals from castor oil. J'. Am. Oil Chem. Soc. 51, 65-71. Niehaus, W. G., and Schroepfer Jr., G. J. (1965). The reversible hydration of oleic acid to 10-D-hydroxystearic acid. Biochem. Biophys. Res. Commun. 21, 271-275. Niehaus, W. G., and Schroepfer Jr., G. J. (1967). Enzymatic stereospecificity in the hydration of epoxy fatty acids. J. Am. Chem. Soc. 89, 4227-4228. Niehaus, W. G., Kisic, A., Torkelson, A., Bednarczyk, D.J., and Schroepfer Jr., G. J. (1970). Stereospecific hydration of the Ag-double bond of oleic acid. J. Biol. Chem. 245, 3790-3797. Schroepfer Jr., G. J. (1966). Conversion of oleic acid to 10-hydroxystearic acid. J. Biol. Chem. 241, 5441-5447. Schroepfer Jr., G. J., and Block, K. J. (1963). Enzymatic stereospecificity in the dehydrogenation of stearic acid to oleic acid. J. Am. Chem. Soc. 85, 3310-3315. Schroepfer Jr., G. J., and Block, K. J. (1965). Enzymatic stereospecificity in the conversion of oleic acid to 10-hydroxystearic acid. J. Biol. Chem. 240, 54-65. Seo, C. W., Yamada, Y., Takada, N., and Okada, H. (1981). Hydration of squalene and oleic acid by Corynebaeterium sp. S-401. Agric. Biol. Chem. 45, 2025-2030.

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Soda, K., and Kido, T. (1987). "Manufacture of Hydroxy Unsaturated Fatty Acids with Bacilluspumilus." Japanese Kokai Tokyo Koho JP 01051092 A2 890227 Heisei. Patent Application JP87-207994. Stodola, F. H., Vesonder, R. E, and Wickerham, L. J. (1965). 8,9,13-trihydroxydocosanoic acid, an extracellular lipid produced by a yeast. Biochem&try 4, 1390-1394. Suemune, H., Harabe, T., and Sakai, K. (1988). Synthesis of unsaturated trihydroxy C-18 fatty acids isolated from rice plants suffering from rice blast disease. Chem. Pharmacol. Bull. 36, 3632-3637. Wallen, L. L., Benedict, R. G., and Jackson, R. W. (1962). The microbial production of 1O-hydroxystearic acid. Arch. Biochem. Biophys. 99, 249-253. Wallen, L. L., Davis, E. N., Wu, Y. V., and Rohwedder, W. K. (1971). Stereospecific hydration of unsaturated fatty acids by bacteria. Lipids. 6, 745-750. Yamada, Y., Motoi, H., Kinoshita, S., Takada, N., and Okada, H. (1975). Oxidative degradation of squalene by Arthrobacter species. AppI. Microbiol. 29,400-404. Yang, W., Dostal, L., and Rosazza, J. P. N. (1993). Stereospecificity of microbial hydration of oleic acid to 10-hydroxystearic acid. Appl. Environ. Microbiol. 59, 281-284.

Ethanol and Thermotolerance in the Bioconversion of Xylose by Yeasts THOMAS W. JEFFRIES

Institute for Microbial and Biochemical Technology Forest Service, Forest Products Laboratory United States Department of Agriculture Madison, Wisconsin 53705-2366 and Department of Bacteriology, University of Wisconsin, Madison Madison, Wisconsin 53706-1527

YONG-SU JIN

Department of Food Science University of Wisconsin, Madison Madison, Wisconsin 53706-1527

I. Introduction II. Lignocellulose A. Pretreatment B. Simultaneous Saccharification and Fermentation III. Xylose-Fermenting Microbes A. Bacteria B. Xylose-Fermenting Yeasts and Fungi C. Genetic Studies with P. stipitis and P. tannophilus D. Expression of Pichia Genes in Saccharomyces IV. Critical Parameters for Yeast Xylose Fermentation A. Carbon Source B. Temperature C. pH D. Aeration E. Nutrient Uptake V. Factors Affecting Thermo- and Ethanol Tolerance A. Membrane Lipids B. Plasma Membrane H+-ATPase C. Mitochondrial Stability D. Trehalose E. Heat-Shock Proteins VI. Summary References

221 ADVANCESIN APPLIEDMICROBIOLOGY,VOLUME47 CopyrightO 2000by AcademicPress All rightsof reproductionin any formreserved. 0065-2164/00 $25.00

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I. Introduction

Most ethanol produced in the world today is derived from starch or sucrose (Gong et al., 1999). These storage carbohydrates are readily hydrolyzed by enzymes, and Saccharomyces cerevisiae easily ferments the resulting sugars--glucose and fructose--to high concentrations of ethanol (e.g., van Hoek et al., 1998). Ethanol fermentations are traditionally carried out for wine or beer production, but ethanol for transportation is a large and growing use. In 1997, about 13,000 tons of ethanol were produced for fuel worldwide (Wilke, 1999). Ethanol is clean burning. Its oxygen content decreases emissions when combusted with gasoline, and because ethanol is derived ultimately from plant matter, its use as a fuel does not contribute to the net accumulation of carbon dioxide in the atmosphere (Costello and Chum, 1998). Starches and sugars are abundant in many crops, but expansion of ethanol production as an automotive fuel in the new millennium will require feedstocks that do not compete for food or fiber (Wheals eta]., 1999). Such feedstocks include lignocellulosic byproduct residues from agriculture and silviculture (Saddler, 1993). Their utilization will require new technologies for efficient, inexpensive bioprocessing. Lignocellulose is a generic term for plant matter derived from wood and agricultural residues. It is composed mainly of lignin and cellulose, but the lignocellulosic phytomass also contains significant amounts of hemicellulose. Xylose is the principal component of hemicellulose found in angiosperm agricultural and silvicultural residues. It is obtained by acid or enzymatic hydrolysis of xylan. The cellulosic fiber component of wood and many agricultural residues is in demand for fiber production, but the hemicellulose and lignin components are available for recovery through biorefining and bioconversion. Biorefining is analogous to petrochemical refining in which a crude feedstock is separated into its higher-value components. While petrochemical refining is carried out at high temperatures on a complex mixture of relatively low-molecular-weight monomers, biorefining is carried out at moderate temperatures on mixtures of polymeric and monomeric materials. Corn wet milling best represents biorefining as it is practiced today. In this large-scale industrial process, corn is separated into its starch, oil, gluten, fiber, and nitrogenous liquid components. Subsequent bioconversion involves enzymatic hydrolysis of the starch and fermentation of the resulting glucose to ethanol, citrate, lactate, and various other products such as antibiotics. Biorefining of lignocellulose to useful products is more difficult, because wood and agricultural residues are composed of structural polymers rather than

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the storage polymers found in grains. Moreover, the various extractives released and degradation products formed during pretreatment often make lignocellulosic hydrolysates harder to use. Lignocellulose contains five major sugars, the abundance of which varies with the feedstock (Pettersen, 1984). They are the hexoses D-glucose, D-mannose, and D-galactose, and the pentoses D-xylose and L-arabinose. Fructose is not normally found in lignocellulose. Commercial bioconversion of lignocellulose to ethanol requires efficient fermentation of sugar mixtures--including xylose (Hinman et al., 1989). Otherwise, product yields are low and waste disposal costs excessive. The fermentation of glucose and fructose has been established through thousands of years of practice. The prevailing yeast strains used for producing wine, beer, and bread have been isolated from many different sources. They belong to S. cerevisiae and a few other taxonomic groups (Vaughn-Martini and Martini, 1995). In contrast, the objective of producing ethanol from pentose sugars has arisen relatively recently, and, despite much effort in several laboratories around the world, it remains problematic. Even though anhydrides of xylose are abundant, xylose itself does not usually occur as a free sugar. Moreover, the five-carbon structure of xylose does not lend itself readily to fermentations in which ethanol is the sole product. Therefore, little natural selection for xylose-fermenting species has taken place. This review focuses on achieving high ethanol concentrations at elevated temperatures--conditions that would be appropriate for enzymatic saccharification and cofermentation of lignocellulosic feedstocks.

II. Lignocellulose Lignocellulosic materials are complex matrices of lignin, cellulose, hemicellulose, various extractives, and inorganic components. The compositions vary widely with plant species, age, time of harvest, and condition or stage of growth (Higuchi 1997). Analysis is challenging (Puls, 1993). About 45% of the total dry weight of wood is cellulose, the hydrolysis of which yields glucose. In agricultural residues, cellulose comprises 8 to 35% of the total dry weight. Glucose is also present in hemicellulosic sugars. Overall, glucose averages 30% of the total dry weight in these materials (Pettersen and Schwandt, 1991; Pettersen, 1984; Krull and Inglett, 1980). The prevalence of glucose in starch and other storage carbohydrates such as sucrose makes D-glucose the most abundant carbohydrate in terrestrial plants. Xylose is the second most abundant sugar. It is especially prevalent in angiosperms (flowering plants). In woody angiosperms (hardwoods), D-xylose averages about

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17% of the total dry weight, but in herbaceous angiosperms, such as residues from agricultural crops, it can comprise up to 31%. Other sugars--such as mannose, galactose, and arabinose--are found in the glucomannan, arabinoxylan, and glucuronoxylan hemicellulosic components. Glucomannan is the main hemicellulose of gymnosperms (softwoods). In these plants, mannose comprises about 10% of the total dry weight. The lignin content is slightly higher in softwoods than in hardwoods, and because the lignin is more crosslinked in softwoods it is harder to remove. Although the glucose and mannose present in softwoods can be fermented readily, timber and pulp manufacture places a high value on straight trunks and long fibers. Therefore, agricultural residues and fast-growing hardwood species are most commonly considered for fuel ethanol production. The high content of xylose in these materials requires that it be used efficiently. A. PRETREATMENT

Lignocellulosic materials must be treated with physical, chemical, or thermal processes in order to release fermentable sugars or increase their susceptibility to enzymatic hydrolysis. Several pretreatments are presently under investigation. They include lime (Chang et al., 1997), ammonia (Dale et al., 1999), high-temperature dilute acid (Lee et al., 1999), and concentrated acid (Goldstein et al., 1989). These and other pretreatments have been reviewed more recently (Szczodrak and Fiedurek, 1996). Acid hydrolysis is one of the oldest and most established technologies for converting lignocellulose into fermentable sugars. There are two principal approaches: dilute sulfuric acid and concentrated-acid hydrolysis. Dilute-acid hydrolysis is often carried out in two stages. In the first stage, a relatively mild hydrolysis is used to recover the hemicellulosic sugars. Depending on the substrate and the conditions used, between 80 and 95 % of the hemicellulosic sugars can be recovered from the lignocellulosic feedstock (Torget and Hsu, 1994; Torget et al., 1996; Katzen and Fowler, 1994). In the second stage, a higher concentration of acid and a higher temperature hydrolyze the cellulose to glucose. The first- and second-stage hydrolysates can be recovered and fermented separately by different organisms or combined and fermented. Other variations include a mild acid pretreatment stage combined with subsequent enzymatic saccharification and fermentation (see §I.B). One of the difficulties with dilute-acid hydrolysis is that it degrades the lignin into a nonreactive form, and it generates large amounts of toxic byproducts that inhibit the growth of fermentative microbes

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(Larsson et al., 1999a; Mandenius et al., 1999). A variation on diluteacid hydrolysis uses continuous countercurrent high-temperature dilute acid to reduce formation of degradation products in the second stage (Lee et al., 1999). While dilute-acid hydrolysis itself is inexpensive and can be used with a wide variety of feedstocks, the detoxification steps add complexity and cost (Larsson et al., 1999b). The hightemperature countercurrent continuous approach results in a higher product yield, but generates a much more dilute sugar stream. With batch-wise dilute-acid hydrolysis, only about 50 to 55% of the cellulose in wood can be converted to sugar. The balance of the material is either left as residual cellulose or is degraded. Therefore, while the technology is inexpensive, it is not sufficiently effective for commercial development unless the feedstock is very cheap. Advanced forms of high-temperature countercurrent hydrolysis or dilute-acid pretreatment combined with enzymatic saccharification and cofermentation may prove to be cost-effective. Concentrated-acid hydrolysis is carried out at lower temperatures and generates fewer degradation byproducts, so fermentation of the resulting sugars is much less problematic. However, concentrated sup furic or hydrochloric acid is difficult to work with, and essentially all of the acid must be recovered and reconcentrated in order for the process to be economical. Electrodialysis (Goldstein, 1989) and ion-exchange chromatography have been investigated as technological approaches to acid recovery, and are presently being pursued for commercialization.

B. SIMULTANEOUS SACCHARIFICATION AND FERMENTATION Simultaneous saccharification and fermentation (SSF) is the most efficient way to convert pretreated lignocellulose to ethanol (McMillan et a]., 1999; Banat et al., 1998; Wyman, 1994; Philippidis et al., 1993). It is often effective when combined with dilute-acid or high-temperature hot-water pretreatment (Sreenath et al., 1999). In SSF, cellulases and xylanases convert the carbohydrate polymers to fermentable sugars. These enzymes are notoriously susceptible to feedback inhibition by the products--glucose, xylose, cellobiose, and other oligosaccharides. But the efficiency of enzymatic saccharification increases if the resulting sugars are converted to ethanol. Because cellulases function well at relatively high temperatures (50 to 70°C), the limiting factor is fermentation. The efficiency of product formation increases with increasing ethanol concentration up to about 5% on a w/w basis, so fermentation

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at high temperatures (>40°C) and at or above 5% ethanol are priorities for commercialization of this technology. A few yeasts will ferment glucose at temperatures up to 45°C. The best studied of these is Kluyveromyces marxianus IMB3 (Fleming et al., 1993; Singh et al., 1998). Strains of this yeast have been adapted by continuous culture in the presence of ethanol to ferment glucose to 44 g/liter ethanol and at temperatures as high as 45°C (Hack and Marchant, 1998). However, growth is slight and the trait is not stable. Researchers have sought thermotolerant yeasts to use in SSF processes for almost 20 years (McCracken and Gong, 1983), but this effort has been largely limited to empirical screening studies. Accumulated knowledge about the basis for ethanol and thermotolerance will provide new opportunities for developing better strains through molecular genetics.

III. Xylose-Fermenting Microbes A. BACTERIA Bacteria have been known to ferment pentoses since the studies of lactobacilli by Fred et al. (1920). Today, xyloseofermenting bacteria include both native and genetically engineered organisms, and many have characteristics useful for simultaneous saccharification and fermentation (Table I). Bacterial fermentations of xylose for ethanol production are being commercialized, but yeasts have several perceived advantages. With a few exceptions, such as lactic acid fermentation (Anuradha et al., 1999), bacteria produce a wide mixture of metabolic products and exhibit much lower ethanol tolerance. This makes product recovery more difficult. Yeasts have larger cells and thicker cell walls than bacteria, which makes cell harvest and recycle easier. Perhaps most importantly, yeast fermentations are not as susceptible to contamination by bacteria or viruses. For these reasons many industrial ethanol processors retain an interest in xylose-fermenting yeasts. Aspects of xylose fermentation by bacteria have been reviewed previously (Jeffries, 1983). B. XYLOSE-FERMENTINGYEASTSAND FUNGI Karczewska (1959) first reported direct conversion of xylose to ethanol by yeast. Even though many yeasts were known to assimilate xylose, this discovery did not enter the review literature (e.g., Lodder, 1971), and was not cited for over 20 years. Various laboratories began to reexamine yeast xylose fermentation after Wang et al. (1980) reported that S. cerevisiae and Schizosaccharomyces p o m b e could ferment xylulose, a keto pentulose, to ethanol. Soon afterward, Schneider et al.

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TABLE I NATIVE AND ENGINEERED BACTERIALSPECIES CAPABLEOF FERMENTING XYLOSETO ETHANOL

Species

Characteristics

References

Clostridium acetobutilicum

Useful in fermentation of xylose to acetone and butanol; ethanol produced in low yield.

E1 Kanouni et al. (1998)

Clostridium thermocel]um

Capable of converting cellulose directly to ethanol and acetic acid; ethanol concentrations are generally less than 5 g/liter.

Herrero and Gomez (1980)

Escherichia coli

Native strains ferment xylose to a mixture of ethanol, succininc, and acetic acids but lack ethanol tolerance; genetically engineered strains predominantly produce ethanol.

Lindsay et al. (1995), Yamano et al. (1998)

Klebsiella oxytoca

Native strains rapidly ferment xylose and cellobiose; engineered to ferment cellulose and produce ethanol predominantly.

Ingram et al. (1999)

Lactobacillus pentoaceticus

Consumes xylose and arabinose. Slowly uses glucose and cellobiose. Acetic acid is produced along with lactic in 1:1 ratio.

Chaillou et al. (1998), Sreenath et al. (1999)

Lactobacillus casei

Ferments lactose very well; particularly useful for bioconversion of whey.

Chaillou et al. (1999), Roukas and Kotzekidou (1998)

Lactobacillus

Uses cellobiose if nutrients are supplied; uses D-glucose, D-xylose, and L-arabinose.

Sreenath, personal communication (1999)

Lactobacil]us pentosus

Homolactic fermentation. Some strains produce lactic acid from sulfite waste liquors.

Chaillou et al. (1999)

Lactobacillus plantarum

Consumes cellobiose more rapidly than glucose, xylose, or arabinose. Appears to depolymerize pectins; produces lactic acid from agricultural residues.

Chaillou et al. (1999), Sreenath et al. (1999)

Zymomonas mobil&

Normally ferments glucose and fructose; engineered to ferment xylose.

Zhang et al. (1995)

xyIosus

228

THOMAS W. JEFFRIES and YONG-SU JIN

(1981) and Slininger et al. (1982) reported that Pachysolen tannophilus could ferment xylose to ethanol. Gong et al. (1981) reported a mutant Candida sp. that would produce ethanol from xylose, and about that same time, Jeffries (1981) reported that Candida tropicalis required aeration to convert xylose to ethanol. The early progress in this field has been reviewed by several authors (e.g., Jeffries and Kurtzman, 1994; Hahn-H~gerdal et al., 1994). Other information on metabolic engineering and regulation of yeast xylose fermentation has been reviewed (Jeffries and Shi, 1999). The current review focuses on attaining elevated levels of ethanol at temperatures that are normally considered extreme for yeast growth. At least 22 yeast strains have been shown to produce some ethanol from D-xylose (Toivola et al., 1984; du Preez and van der Walt, 1983; Schneider et al., 1981) (Table II). However, only six of these

TABLE II NATIVEOR ENGINEEREDYEASTANDFUNGALSPECIESCAPABLE OF FERMENTINGXYLOSETO ETHANOL Species

Characteristics

References

Has both active and passive transport systems for xylose uptake; produces moderate amounts of xylitol; does not grow anaerobically; requires biotin and thiamine.

du Preez and van der Walt (1983)

Candida boidinii

Produces large amounts of xylitol; oxidizes methanol.

Vandeska et al. (1996)

Pichia stipitis

Ferments all sugars found in wood; some strains ferment xylan.

Lee et al. (1986)

Fusarium oxysporum; Fusarium oxysporum var. lini

Ferments 20 different carbon sources, including xylitol; does not use xylan or cellulose; converts xylose to ethanol, CO2, and acetic acid.

White and Williams (1928), Gibbs et al. (1954), Suihko et al. (1991), Suihko (1983)

Mucor sp.

Ferments pentoses and alditols to ethanol.

Ueng and Gong, 1982

Pachysolen tannophi]us

Ferments xylose, glucose and glycerol; metabolizes xylose anaerobically; produces large amounts of xylitol.

Schneider et al. (1981), Slininger et al. (1982)

Candida shehatae

THERMO- AND ETHANOL TOLERANCE

229

(Brettanomyces naardenensis, Candida shehatae, Candida tenuis, P. tannophilus, Pichia segobiensis, and Pichia stipitis) produce significant amounts of ethanol, and of these only three (C. shehatae, R tannophilus, and R stipitis) have been studied extensively. Several research groups have developed genetic transformation systems for P. tannophi]us (Wedlock and Thornton, 1989; Reiser et al., 1990; Hayman and Bolen, 1993) and for P. stipitis (Ho et al., 1991; Morosoli et al., 1993; Yang eta]., 1994; Lu et al., 1998a; Piontek eta]., 1998). It is possible to transform auxotrophic strains of C. shehatae with selectable markers and vectors designed for P. stipitis, but the absence of a sexual mating system in C. shehatae makes it less useful for fundamental studies. C. GENETIC STUDIES WITH e. STIPITIS AND P. TANNOPHILUS

Cloning and disruption of genes for the critical enzymes involved in xylose metabolism or their overexpression in S. cerevisiae have led to greatly improved understanding of the rate-limiting steps in yeast xylose metabolism. At the end of 1999, GenBank listed approximately 26 entries for genes cloned from P. stipitis or P. tannophilus--in addition to ribosomal genes used for taxonomic classification studies (Kurtzman, 1994). Stevis and Ho (1987) did some of the earliest research on the genes for xylose metabolism by creating a xylulokinase mutant of Escherichia coli (Stevis et al., 1987), then using it to clone the xylulokinase gene from P. tannophilus. Subsequently, Ho and Chang (1989) used as similar approach to clone a gene for xylulokinase from S. cerevisiae. K6tter et al. (1990) first reported cloning the gene for xylitol dehydrogenase, Xyl2, from P. stipitis, and, more recently, S h i e t al. (2000) sequenced a more complete clone from another strain. Takuma et al. (1991) and Hallborn et al. (1991) independently cloned the gene for aldose reductase (Xyll) from P. stipitis. Billard et al. (1995) cloned a gene with 62% identity to P stipitis Xyll, and many other related genes are on deposit in GenBank. Hallborn et al. (1995) isolated a short-chain dehydrogenase gene from P. stipitis that has D-arabinitol dehydrogenase activity. The biochemical differences among these and other dehydrogenases have been reviewed previously (Jeffries and Shi, 1999). Walfridsson et al. (1995) isolated the R stipitis Tkll and Ta]l genes for transketolase and transaldolase, and Weierstall et al. (1999) isolated the genes for the glucose transporters of P. stipitis. Other genes cloned from P stipitis include the X y n A gene for xylanase (Lee et al., 1986; Basaran et al., 1999), the Cycl gene for cytochrome c (Shiet a]., 1999), the selectable markers LEU2 (Lu et al., 1998a) and Ura3 (Yang et al., 1994), an A R S 2

230

THOMAS W. JEFFRIESand YONG-SUJIN

sequence that confers autonomous replication (Yang et al., 1994), the P d h e l a component for pyruvate dehydrogenase (Davis and Jeffries, 1997), two genes for pyruvate decarboxylase (Pdcl, Pdc2) (Lu et al., 1998b), and two genes for alcohol dehydrogenase (Adhl, Adh2) (Cho and Jeffries, 1998; Passoth et al., 1998). Fewer genes have been cloned from P. tannophilus. They include genes for aldose reductase (Bolen et al., 1996), ornithine carbamoyltransferase (Skrzypek et a]., 1990), UDP galactose-4-epimerase (Skrzypek and Maleszka, 1994), cytochrome c (Clark-Walker, 1999a), cytochrome c oxidase subunit 2 (Clark-Walker, 1999b), and OMP decarboxylase (Ura3) (Clark-Walker, 1998). Functions for most of these genes can be surmised through comparative genomics, but when two or more isomers are present or when the gene products are active in a poorly defined pathway, their functions cannot be determined until physiological effects have been demonstrated. For example, Cho and Jeffries (1998) showed that disruption of the P s A d h l gene resulted in accumulation of xylitol in P. stipitis, whereas disruption of PsAdh2 had no significant effect on cells growing on xylose under oxygen limited conditions. Xylitol dehydrogenase is a member of the alcohol dehydrogenase family, but the principal substrate that it acts on is 15 biochemical steps away from ethanol. Xylitol dehydrogenase and alcohol dehydrogenase both use NAD(H) as a cofactor, so deletion of PsAdh 1 is thought to cause accumulation of xylitol by increasing the internal concentration of this cofactor. These findings are in accord with the observed expression of PsAdhl under fermentative conditions (Passoth et al., 1998; Cho and Jeffries, 1999). The PsAdhl gene therefore appears to keep intracellular NADH levels low and to be responsible for ethanol formation. Pyruvate decarboxylase (PsPdcl} has unusual kinetic (Passoth et al., 1996) and structural (Lu et al., 1998b) features that could be important in supplying acetaldehyde as an electron sink. Some evidence suggests that the PsAdh2 gene might be involved in ethanol oxidation. Disruption of the Cycl gene in P. stipitis has the interesting effect of greatly reducing both respiration and cell growth, thereby diverting reductant into ethanol production (Shi and Jeffries, 1999). The cells are able to survive apparently because they possess an alternative oxidase (Jeppsson et al., 1995). P. stipitis has not been used as extensively as S. cerevisiae for heterologous expression, but its unique physiology has allowed for a few interesting experiments. The heterologous expression of a Cryptococcus xylanase enables P. stipitis to ferment xylan (Morosoli et al., 1993), and

THERMO- AND ETHANOLTOLERANCE

231

the heterologous expression of S. cerevisiae Ural, which codes for a dihydroorotate dehydrogenase that uses fumarate as an alternative electron acceptor, enables R stipitis to grow anaerobically on glucose (Shi and Jeffries, 1998). D. EXPRESSIONOF PICHIA GENESIN SACCHAROMYCES Even though S. cerevisiae does not ferment D-xylose, it does convert D-xylulose to ethanol under microaerobic conditions (Maleszka and Schneider, 1984). It possesses a gene for aldose reductase (Garay-Arroyo and Covar-Rubias, 1999), and it will produce xylitol from xylose. It has a gene that is very similar to the xylitol dehydrogenase of P. stipitis (Richard et al., 1999), and it also possesses and expresses a gene for D-xylulokinase activity (Deng and Ho, 1990). Therefore, the question is "Why does S. cerevisiae NOT assimilate and produce ethanol on xylose?" Researchers have addressed this by examining the effects of heterologous gene expression of various genes from P. stipitis. The metabolic engineering of xylose fermentation in S. cerevisiae has been progressively more successful. Genes for B stipitis xylose reductase (Xyll) (Amore et al., 1991; Hallborn et al., 1991), Xyll plus xylitol dehydrogenase (Xyl2) (K6tter et al., 1990; Tantirungkij et al., 1993, 1994a), transketolase (Tkt) plus Xyll and Xyl2 (Metzger et al., 1994), or Tkt plus transaldolase ( Tal) and Xyll and Xyl2 (Walfridsson et al., 1995) have been expressed in S. cerevisiae in order to impart xylose fermentation. The introduction of Xyll from R stipitis did not enable S. cerevisiae to grow on or produce ethanol from xylose. However, heterologous expression of Xyll does enable it to make xylitol from xylose, as long as a supplemental carbon source is provided. Galactose is particularly useful because it does not compete with xylose for transport (K6tter and Ciriacy, 1993). The presence of both Xyll and Xyl2 enables S. cerevisiae to grow on xylose (Tantirungkij et al., 1994a,b; Meinander et al., 1996), but it is necessary to also overexpress the gene for xylulokinase (Xksl) in order to obtain significant growth or ethanol production on xylose. An S. cerevisiae fusion strain containing PsXyll, PsXyl2, and the Saccharomyces gene for xylulokinase (Xksl) shows higher fermentative capacity on glucose and xylose (Chang and Ho, 1988; Ho et al., 1998). Presumably, overexpression of the native Saccharomyces Xksl improves xylose metabolism in this strain. Rodriguez-Pefia et al. (1998) examined the function of Xksl in well-defined laboratory strains of S. cerevisiae, and they reported that deletion of Xksl blocked growth on xylulose, but overexpression also had a negative effect on growth. This

232

THOMAS W. JEFFRIES and YONG-SU JIN

suggests that the role for xylulokinase might be rather complex, and that its expression needs to be closely regulated in S. cerevisiae. Expression ofP. stipitis Tkt and Tk! genes does not show significant effects in recombinant S. cerevisiae strains, which indicates that the nonoxidative portion of the pentose phosphate pathway is not limiting in this yeast. It is particularly difficult to engineer S. cerevisiae for the fermentation of mixtures of glucose and xylose because glucose interferes with xylose transport in S. cerevisiae, and in the absence of glucose the enzymes responsible for fermentation are not induced. To better understand the problem of xylose uptake, van Zyl et al. (1999) expressed P. stipitis Xyll and Xyl2 on a multicopy vector in a uracil phosphoribosyltransferase-deficient (furl) strain of S. cerevisiae. This enabled cultivation of the transformants in a rich medium with autoselection for growth on xylose. The researchers used either glucose or raffinose, a slowly metabolized carbon source, as co-metabolizable carbon sources. By using a complex medium rather than a minimal defined medium, they increased xylose utilization twofold. Addition of glucose or raffinose as a cosubstrate increased xylose utilization another threefold. In rich medium with raffinose as the cosubstrate, the transformants consumed 50 g/liter of xylose and produced about 5 g/liter each of xylitol and ethanol after 80 hr. Overexpression of X k s l in this genetic background and with these nutritional conditions would probably increase ethanol production further. It is unclear from the current literature whether the conditions for ethanol production from xylose have actually been optimized with recombinant S. cerevisiae strains--particularly with respect to aeration. In most instances, the amounts of ethanol produced are well below what would be considered toxic levels for this organism, so the limiting factors are still not well understood. The fermentation characteristics of various microbes are shown on Table IIL IV. Critical Parameters for Yeast Xylose Fermentation

Carbon and nitrogen sources, aeration, pH, and temperature are important for cell growth and product formation. Aeration plays a critical role. Oxygen limitation induces fermentation in P. stipitis and C. shehatae (Alexander eta]., 1988; Alexander and Jeffries, 1990; Cho and Jeffries, 1998; Passoth et al., 1998). At the same time, these yeasts require oxygen for growth and maximal ethanol production (Neirinck et el., 1984; Rizzi et al., 1989). Aeration is also important for pentose

233

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234

THOMAS W. JEFFRIESand YONG-SUJIN

metabolism in S. cerevisiae (Maleszka and Schneider, 1984). Attaining and maintaining optimum aeration rates present special challenges. A. CARBONSOURCE Du Preez et al. (1989) found that the maximum ethanol concentration attained by P. stipitis and C. shehatae was not affected by using D-glucose rather than D-xylose as a substrate. However, Meyrial eta]. (1995a) found that the ethanol concentration resulting in growth inhibition depended on the sugar consumed. In the case of xylose, growth inhibition occurred at 30 g/liter, but with glucose, cells continued to grow up to 34 g ethanol liter-1. The higher ethanol tolerance observed with glucose as a carbon source correlated with higher plasma membrane H+-ATPase activity (see below, §V.B). Moreover, the carbon source also determined whether or not the ATPase activity was stimulated by addition of 10 g/liter ethanol (Meyrial et al., 1995b). P. stipitis produced more ethanol in the studies reported by du Preez et al. (1989) than it did in the studies by Meyrial et al. (1995a). Probably, du Preez attained the higher concentrations by using fed-batch fermentation because sugar concentrations above 50 g/liter adversely affect both P. stipitis and C. shehatae. The fed-batch fermentation might also account for the different effects observed of sugars on ethanol tolerance. The ethanol tolerance of P. tannophilus changes with the carbon source used for growth (Jeffries eta]., 1985). When cultivated on xylose as a sole carbon source, this yeast produces only about 20 g/liter of ethanol, but it produces up to 55 g/liter when cultivated on glucose. P. tannophilus also produces ethanol much more rapidly on glucose than on xylose. Addition of glucose to xylose fermentations by P. tannophiIns increases ethanol production from the former sugar. Jeffries et al. (1985) hypothesized that this might be attributable to repression of ethanol respiration, but glucose could have affected ethanol tolerance as well. B. TEMPERATURE The effects of temperature on growth and ethanol concentration have been well studied in the xylose-fermenting yeasts. Prior eta]. (1989), Hahn-H~igerdal et al. (1994), du Preez (1994), and McMillan (1994) have reviewed the literature on optimum conditions for yeast xylose fermentation in detail. Temperatures that provide for optimum biomass and ethanol productivities do not necessarily enable maximum ethanol accumulation

THERMO-AND ETHANOLTOLERANCE

235

(Slininger et al., 1990). This implies that ethanol toxicity affects production. In P. stipitis, xylitol and residual xylose concentrations increase with temperature. Maximum biomass and ethanol productivity by P. stipitis occurred at 26 to 35°C. Maximum ethanol selectivity was achieved at 25 to 26°C. The optimum pH range for growth and fermentation on xylose was 4-7 at 25°C. In contrast to the low-temperature optimum for xylose, ethanol productivity and accumulation for glucose were optimal at 34°C. Slininger et al. (1985, 1990) obtained a maximum of 57 g/liter ethanol with P. stipitis cultivated at 25°C. Du Preez et al. (1986a,b) observed that the limit for ethanol production with P. stipitis increased from 33 g/liter at 30°C to 43 g/liter at 25°C. Du Preez et al. (1989) reported that ethanol inhibits growth of P. stipitis and C. shehatae at lower levels than it inhibits ethanol production. This difference in inhibitory concentrations has also been observed with S. cerevisiae (Brown et al., 1981; Lee et al., 1980), Kluyveromyces fragilis (Rosa et al., 1986), and the bacterium Z y m o m o n a s mobilis (JSbses and Roels, 1986). These observations are probably related. Microbes rely on proton gradients for active transport, and if plasma membrane ATPase activities are disrupted by ethanol, cell growth will stop as nutrient uptake ceases. Facilitated diffusion could continue to support sugar uptake and fermentation, but growth will be impaired in the absence of mechanisms for nitrogen uptake. Some strains of xylose-fermenting yeasts produce up to 47 g ethanol • liter-1 at 30°C (Table IV). Du Preez et al. (1989) found that in R stipitis and C. shehatae ethanol production was mainly growth associated and that the volumetric rate of ethanol production increased linearly with the volumetric growth rate over a three- to fivefold range. They could not increase the maximum ethanol concentration on D-xylose by increasing the initial cell density, and they concluded that the low ethanol tolerance of these xylose-fermenting yeast strains is not a consequence of the metabolic pathway used during pentose fermentation. These results suggest to this reviewer that the inhibitory effect of ethanol could result from impaired active uptake systems. Because ethanol production is growth related, any loss of viability or decrease in growth rate will reduce ethanol formation. Lucas and van Uden (1985) found that ethanol enhances thermal death of C. shehatae. The specific growth rate did not vary significantly from its maximum (-31°C) down to 20°C. Maximum ethanol tolerance (6% v/v) occurred over a temperature plateau (10 to 17.5°C). Ethanol depressed the maximum temperature for growth from 31 to 17.5°C and increased the minimum temperature for growth from 2.5 to 10°C.

236

THOMAS W. JEFFRIES and YONG-SU JIN TABLE IV

CONCENTRATIONSOF ETHANOL INHIBITORYTO GROWTH AND ETHANOL PRODUCTIONBY PICHIA

STIPITIS AND CANDIDA SHEHATAE IN FED-BATCH FERMENTATIONSON XYLOSE AND GLUCOSEa

Yeast strain

Carbon source

Growth (g/liter)

C. shehatae Y492 C. shehatae Y798 C. shehatae Y981 C. shehatae Y981 P. stipitis Y663 P. stipitis Y663

Xylose Xylose Xylose Glucose Xylose Glucose

32.3 30.5 31.2 34.9 35.1 34.9

Fermentation (g/liter) 44 38.9 45.4 44.8 47.1 43.8

aData from du Preez et al. (1989).

Acetic acid also shifts growth and thermal death profiles to lower temperatures (Rodrigues-Alves et al., 1992}. Forced cycling of pH 0.5 units above and below the optimum of 4.5 decreased the fermentation rate but did not affect ethanol yield (Ryding et al., 1993). Ethanol tolerance of C. shehatae decreases when inhibitory compounds are present. This is particularly conspicuous with acid hydrolysates of wood (Hahn-H~igerdal et al., 1991). C. shehatae was able to tolerate up to 0.4% (v/v) acetic acid at pH 4.5, but its presence in the medium reduced the temperature growth range from 5 to 34°C to between 21 and 27°C. Acetic acid decreased the cell yield by 64% and tolerance to added ethanol from 5% (v/v) to 2% (Rodrigues-Alves et al., 1992). It is hypothesized that un-ionized acetic acid disrupts proton gradients by diffusing from the acidified external medium into the cells and dissociating in the more neutral environment of the cytoplasm. With loss of the proton gradient to drive nutrient uptake, ethanol tolerance declines. C. P H

Sanchez et al. (1997) found that the best initial pH for ethanol production from D-xylose by C. shehatae in batch fermentation was 4.5. Under these conditions, the maximum specific growth rate (Pmax) was 0.329 hr -1 and the specific ethanol production rate (qE) was 0.72 kg • k g - 1 hr -1. The average xylitol yield was 0.078 kg - kg-1, and the overall ethanol yield was 0.41 kg • kg-1. Both the specific substrate uptake rate (qS) and qE diminished once the exponential growth phase was over. A maxi-

237

THERMO- AND ETHANOL TOLERANCE

mum qE of 0.72 kg • kg-1 hr-1 equates to about 15.6 mmol ethanol g-1 hr-1. By comparison, van Hoek et al. (1998) showed that the fermentative capacity of S. cerevisiae increased with the specific growth rate and ranges between 10 and 22 mmol of ethanol g-1 hr-1. Thus, under optimal conditions the fermentative capacities of P. stipitis on xylose and S. cerevisiae on glucose do not differ greatly. The performance characteristics of several xylose-fermenting yeasts are listed in Table V. The pH and temperature optima for biomass accumulation by P. tannophilus on xylose were 3.7 and 31.5, respectively, at an initial xylose concentration of 50 g/liter (Roebuck et al., 1995). As in the case for P. stipitis and C. shehatae, P. tannophilus attains maximum ethanol productivity under microaerobic conditions (Kruse and Schugerl,

TABLE V PERFORMANCEOF XYLOSE-FERMENTINGYEASTSON GLUCOSEAND XYLOSEa

Carbon

Strains

Pichia stipitis

source (g/liter) Xylose (40)

Aeration Aerobic Oxygen limited

CBS7126

Anaerobic Glucose (40)

Aerobic Oxygen limited

Anaerobic

Candida shehatae

Xylose (40)

Aerobic Oxygen limited

Anaerobic

CBS 2779

Glucose (40)

Aerobic Oxygen limited

Anaerobic

PachysoIen

Xylose (40)

Aerobic

tannophilus

Oxygen limited

NRRL Y-2460

Anaerobic Glucose (40)

Aerobic Oxygen limited

Anaerobic aData from Ligthelm et al. (1986).

Eth-

Produc-

Xy-

Bio-

anol yield

tivity (g/g.

litol yield

mass yield

(g/g)

1. hr)

(g/g)

(g/g)

0.18 0.47 0.40

0.17 0.20 0.02

0 0.06 0

0.39 0.05 0.03

0.26 0.38 0.33

0.17 0.28 0.13

0.22 0.37 0.41

0.21 0.32 0.15

0.33 0.42 0.44

0.35 0.51 0.29

0.10 0.28 0.26

0.04 0.10 0.07

0.31 0.43 0.42

0.38 0.49 0.18

0.23 0.14 0.10 0.04 0.13 0.18

0.33 0.01 0.01 0.21 0.03 0.02

0.17 0.30 0.30

0.25 0.01 0.01 0.14 0.06 0.04

238

THOMAS W. JEFFRIESand YONG-SUJIN

1996). However, R tannophilus xylitol yields tend to be much greater. With a detoxified hemicellulose hydrolysate at pH between 6.0 and 7.5, R tannophilus converted 90% of the available xylose into xylitoh At pH values outside this range, cells respired up to 30% of the xylose (Converti et al., 1999). Kavinaugh and Whittaker (1994) reported that recycling cells of R tannophilus NCYC 614 from batch fermentations over a period of 31 days increased ethanol tolerance in this strain. This effect did not apply to all instances, because they were not able to see a significant increase in ethanol tolerance with R tannophilus CBS 4045. D. AERATION The dissolved oxygen tension (DOT) is particularly critical in attaining maximal ethanol production with xylose-fermenting yeasts. R stipitis and C. shehatae require aeration for maximal ethanol production (du Preez et al., 1986a,b). Under anoxic conditions, the specific ethanol productivity of R stipitis and C. shehatae decreased, and especially in the case of C. shehatae, xylitol production increased (du Preez et al., 1989). This requirement is not unique to P. stipitis or C. shehatae, because native S. cerevisiae also requires oxygen to metabolize xylulose (Maleszka and Schneider, 1984), and recombinant S. cerevisiae requires oxygen to produce ethanol from xylose (K6tter et al., 1990). Even Fusarium oxysporum requires oxygen for the fermentation of D-xylose and D-glucose (Singh et al., 1992). The oxygen requirement for ethanol production was considered novel when first reported (Jeffries, 1981), but it is apparent that oxygen plays various roles in the metabolism of xylose by eukaryotes. It is important for a xylose-fermenting yeast to possess an aldose reductase that is active with both NADH and NADPH in order to maintain redox balances during xylose assimilation (Verduyn et al., 1985), but oxygen enters into xylose metabolism in other ways as well. One of the factors limiting ethanol production is its simultaneous assimilation. With R stipitis at ethanol concentrations in excess of 28 g/liter, ethanol assimilation exceeds production, even when the dissolved oxygen tension is kept to 0.2% of saturation. In the absence of aeration, ethanol accumulation continues, but at a much lower rate, and xylitol production increases (du Preez et al., 1989). Reduced respiration capacity could be the reason that R stipitis cycA strains (Shi et al., 1999) exhibit higher specific ethanol production rates, du Preez et al. (1989] attained 47 g • liter-1 ethanol at 30°C with the DOT controlled at 0.2%

THERMO- AND ETHANOL TOLERANCE

239

of air saturation. Jeffries and Alexander (1990) attained 56 g/liter ethanol within 38 hr (1.53 g. liter -1. h r -1) with C. shehatae in fed-batch fermentation after the cell inoculum had been cultivated at a high dilution rate and shifted to oxygen limited conditions. Candida shehatae requires oxygen to maintain viability. Kastner et al. (1999) showed that oxygen starvation induces cell death in C. shehatae w h e n it is grown on D-xylose, but not w h e n it is cultivated on D-glucose. Growth of C. shehatae was limited to one division or less w h e n cells cultivated aerobically on either glucose or xylose are shifted from aerobic to anaerobic conditions. Cell viability rapidly declined with cells cultivated on xylose, but cells cultivated on glucose remained viable nine times longer. Shi and Jeffries (1998) also observed differences between glucose and xylose during anaerobic cultivation of P. stipitis. The basis for this difference is not fully understood, but it could reflect differences in the cells to produce metabolic energy on glucose and xylose under anaerobic conditions or the ability of glucose-grown cells to maintain nutrient transport systems. For example, Meyrial et al. (1995a) reported that cultivation of P. stipitis on glucose increases the activity of plasma membrane ATPase threefold in comparison to the activity obtained w h e n cells are grown on xylose. The pH and temperature optima did not shift, and the enzymatic activities showed similar affinity for ATP. However, the glucose-activated enzyme was less sensitive to ethanol. These results show that plasma membrane ATPase activity, which is critical for transport, correlates with ethanol tolerance and the inhibitory effect of ethanol on growth. Plasma membrane ATPase is essential for maintaining the proton gradient that is responsible for uptake of nutrients. Biosynthesis of ergosterol, cardiolipin, and unsaturated fatty acids requires oxygen (Hossack and Rose, 1976; Mandal et al., 1978), and exogenous supplies are necessary for the anaerobic growth of S. cerevisiae (Hossack et al., 1977). However, these lipids are not sufficient for the anaerobic growth of R tannophilus (Neirinck et al., 1984) or B stipitis (Shi and Jeffries, 1998). These yeasts, like most other eukaryotes, require active electron transport for the synthesis of uracil, and hence cannot make mRNA under anaerobic conditions. The critical enzyme step that imposes this limitation is dihydroorotate dehydrogenase (DHODase). In most eukaryotes, it is located in the mitochondria, and regeneration of its cofactor requires active electron transport coupled with respiration. S. cerevisiae possesses an unusual DHODase (ScUral) that resides in the cytoplasm, which couples the reduction of fumarate to succinate with the regeneration of its cofactor. Expression of ScUral

240

THOMAS W. JEFFRIES and YONG-SU JIN

in R stipitis enables this yeast to grow anaerobically on glucose, but not on xylose, so some other factor such as sugar transport or energetics probably limits anaerobic xylose metabolism in this organism. E. NUTRIENT UPTAKE

Membrane transport is mediated by two different systems in yeasts: facilitated diffusion and active proton symport. Facilitated diffusion is energy independent and functions well at elevated sugar concentrations. Proton symport requires generation of a proton gradient but is useful during growth at low extracellular sugar concentrations. In S. cerevisiae, the transport of glucose into the cells plays a direct role in sensing glucose and in signal transduction. S. cerevisiae uses facilitated diffusion systems to take up hexoses but uses proton symport systems to take up disaccharides. S. cerevisiae can handle wide ranges of sugar concentrations up to 1.5 M by developing a group of hexose transport (Hxt) proteins. The presence and the concentration of appropriate substrates tightly regulate expression of these enzymes. In S. cerevisiae, hexose uptake is mediated by a large number of related transporter proteins. Six out of 20 genes for hexose transport mediate the uptake of glucose, fructose, and mannose at metabolically relevant rates (Boles and Hollenberg, 1997). Two others catalyze the transport of only small amounts of these sugars. One protein is a galactose transporter but is also able to transport glucose. Hexose-transport-deficient mutants (hxt) have no clearly detectable phenotypes. Expression of Hxtl, 2, 3, 4, 6, or 7 is sufficient to allow various degrees of glucose utilization (Reifenberger et al., 1997) In yeasts that utilize both xylose and glucose, these sugars share the same transporter systems (Boles and Hollenberg, 1997). Glucose can inhibit xylose uptake by competing with the xylose transporters. Even 0.05 mM glucose can compete with xylose uptake, which significantly reduces xylose transport. Xylose transport in P. stipitis is mediated by low- and high-affinity proton symporters (Weierstall et al., 1999). Both transporters are constitutively expressed with low Vmax values. The low-affinity system takes up glucose in the range of 0.3-1 mM. Moreover, inhibitor studies indicate that uptake of xylose requires aerobic respiration, which suggests that both systems involve proton symport (Loureiro-Dias and Santos, 1990). A putative xylose transporter gene from R stipitis, PsStul, has been cloned recently (Weierstall et al., 1999). This gene can confer high-affinity uptake of glucose and growth to a S. cerevisiae h x t l - 7 strain (Boles and Hollenberg, 1997). The deduced amino-acid sequence shows

THERMO- AND ETHANOL TOLERANCE

241

54% identity to the Hxt glucose transporters of S. cerevisiae. When P s S t u l is introduced into S. cerevisiae, it can transport xylose but with a considerably lower affinity than what is achieved in P. stipitis. This suggests that the transport system in P. stipitis is coupled to other elements. Two other Stu-related genes have been identified from P. stipitis by crosshybridization with S. cerevisiae transporter genes as probes. Uptake of xylose in S. cerevisiae is mediated nonspecifically and with low affinity by the hexose transporters, so other factors would be required to increase xylose metabolism in this yeast. In S. cerevisiae genetically engineered for xylose uptake, glucose, mannose, and fructose inhibited xylose conversion by 99, 77, and 78%, respectively. These sugars are transported with by the same high-affinity transport system as xylose, and the results are thought to reflect competitive inhibition of xylose transport (Meinander and Hahn-H/igerdal, 1997). Galactose is less inhibitory to xylose transport than is glucose and was therefore a better co-metabolizable carbon source for xylitol production. Membrane transport plays an important role in the utilization of xylose and other sugars in lignocellulose hydrolysates, and it can limit utilization of sugars (Spencer-Martins, 1994). V. Factors Affecting Thermo- and Ethanol Tolerance

Ethanol tolerance is very important in brewing, wine making, and especially in the biosynthesis of industrial ethanol. Because it is so critical, it has been studied extensively (e.g., Mishra and Singh, 1993). The mechanisms underlying ethanol resistance are very complex. They differ from one yeast to another (Alexandre et al., 1994) and with the conditions for cultivation. Many genes appear to be involved (D'Amore et al., 1990), and the exact basis for ethanol tolerance is not fully understood (Chi et al., 1999). Factors that affect ethanol tolerance include the proportion of ergosterol in the cellular membranes, phospholipid biosynthesis, the degree of unsaturation of membrane fatty acids (Alexandre et al., 1994), temperature, the activity of plasma membrane ATPase, superoxide dismutase, and the capacity of a strain to produce trehalose. Sublethal heat and ethanol exposure induce essentially identical stress responses in yeast (Piper, 1995). Factors affecting the capacity of yeast to survive at high temperatures include the presence of stress-response pathways to signal induction of appropriate heat-shock proteins. One induced protein, Hspl04, contributes to both thermotolerance and ethanol tolerance. Heat and ethanol stress cause similar changes to plasma membrane protein composition, reducing the levels

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of plasma membrane H*-ATPase protein and inducing the plasma membrane-associated Hsp30. A. MEMBRANELIPIDS Temperature is a critical variable that affects both growth and ethanol tolerance. Decreasing temperature decreases membrane fluidity; increasing temperature increases membrane fluidity. According to the principle of homeoviscous or homeophasic adaptation, the cell must compensate for environmental changes by altering its composition to maintain fluidity at the new temperature (Vigh et al., 1998). The manner in which this adaptation occurs, however, can vary with cell type and the conditions. Organisms commonly adapt to low temperatures by increasing the proportion of cis-unsaturated fatty-acyl groups in their membrane lipids. Physical principles suggest that fluidity would decrease as the ratio of saturated to unsaturated fatty acids increases because desaturation introduces a bend in the fatty acid chain. However, the bulk of fatty acids in the membranes of S. cerevisiae are unsaturated (Fig. 1), so other factors may be more important. For example, when S. cerevisiae cells are grown for an extended period at 37°C, they adapt to the higher temperature by increasing their content of unsaturated fatty acids. The cells adapted to 37°C require a higher temperature in order to induce heat-shock proteins (Chatterjee et al., 1997). Guerzoni et al. (1997) found that the unsaturation level of S. eerevisiae cellular fatty acids increases at both sublethal or supraoptimal temperatures. The adaptation can be reversed by cultivation at 25°C. They hypothesized that a high content of unsaturated fatty acids is not in itself a prerequisite for withstanding temperature stress, but rather results from activation of oxygen-consuming desaturase activity. Membrane fluidity is affected by the ratios of cell lipids and proteins. These vary with the yeast strain and the conditions under which it is cultivated. Alexandre et al. (1994) found that the average protein, lipid, phospholipid, and sterol contents of two S. cerevisiae strains were about 30, 15, 3.5, and 1.5%, respectively. When cultivated with 4 or 10% added ethanol, the lipid, phospholipid, and sterol concentrations all decreased. S. cerevisiae FY 169--the strain used in the European Union Yeast Genome Sequencing Program--is atypical in that it has significantly higher palmitic (C16:0) and lower amounts of oleic (C18:1) fatty acids as compared to other wild-type yeasts. Therefore, the data shown in Figure 1 would be even more biased toward unsaturated fatty acids in a typical strain of S. cerevisiae (Daum et al., 1999).

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Glucose grown cells

[ ] C_<14 [ ] C16:1 PM

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Ethanolgrowncells

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C16:0

[ ] C18:1

1171

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:

Mito Mic V'ac Subcellular fraction

FIG. 1. Fatty acid composition of subcel]u]ar membranes from ]ate logarithmic phase

cells of Saccharomyces cerevisiae FY 169 cultivated on 2% glucose or 2% ethanol at 30°C under aerobic conditions (Tuller et al., 1999). PM = plasma membrane, Mito = mitochondria, Mic = microsomes (endoplasmic reticulum), Vac = vacuoles; C numbers indicate chain length and degree of unsaturation.

Decreases in the sterol:protein and s t e r o l : p h o s p h o l i p i d ratios, and an increase in the u n s a t u r a t i o n i n d e x all increase m e m b r a n e fluidity. However, fluidity i n d i r e c t l y d e d u c e d from the sterol:protein, sterol:phosp h o l i p i d , or u n s a t u r a t e d : s a t u r a t e d fatty acid ratios does not always reflect the real state of the plasma m e m b r a n e (Alexandre et al., 1994). It is n e c e s s a r y to m e a s u r e m e m b r a n e fluidity directly. Swan and Watson (1999) substituted various u n s a t u r a t e d fatty acids in a Ag-desaturase (oleil) m u t a n t and f o u n d that the most heat- and ethanol-tolerant cells had m e m b r a n e s e n r i c h e d in oleic acid (C18:1). S. cerevisiae does not p r o d u c e the p o l y u n s a t u r a t e d linoleic (18:2) and linolenic (18:3) fatty acids, w h i c h are present in other plants and fungi. To investigate the

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possible roles of such lipids in ethanol resistance, Kajiwara et al. (1996) examined the heterologous expression of an Arabidopsis thaliana A12 desaturase in S. cerevisiae. They showed that the transformed yeast is capable of accumulating hexadecadienoylw (16:2) and lineloeoyl- (18:2) substituted membrane lipids and that the cells had greater resistance to ethanol than the controls. Ethanol also affects membrane fluidity, but through different mechanisms. Alcohols lower the temperature required for maximal activation of heat-shock genes, and the concentration of alcohol needed decreases with alcohol chain length (Vigh et al., 1998). Ethanol is thought to alter membrane organization and permeability by entering the hydrophobic interior and increasing the polarity of this region (Alexandre et al., 1993). This would weaken hydrophobic interactions and increase the diffusion of polar molecules through the membrane. Ethanol in the cell membrane is said to weaken the water lattice structure, decrease the strength of lipophilic interactions, promote membrane leakage, and decrease the integrity of the semipermeable barrier (Sajbidor and Grego, 1992; Sajbidor, 1997). Such changes would put extraordinary demands on active transport systems for nutrient uptake. Jones and Greenfeld (1987) found that, beyond a critical threshold concentration, the fluidity of the yeast plasma membrane increases exponentially. Cells grown under anaerobiosis have a lower level of fatty acids than cells grown aerobically, but the anaerobically grown cells exhibit higher membrane fluidity. Sensitivity to both heat and oxidative stress depends on membrane lipid composition. In the case of anaerobically grown cells, the most stress resistant have membranes enriched in saturated fatty acids (Steels et al., 1994). An increase in fatty acid unsaturation in cellular membranes increases ethanol tolerance (Alexandre et al., 1994). Sterols, especially ergosterol, promote growth and ethanol tolerance by providing rigidity to the cell membrane (Zinser et al., 1991). The plasma membrane is particularly rich in ergosterol, and the fraction increases with growth on ethanol (Fig. 2). Total sterol content of cells increases with cultivation under anaerobic conditions and decreases under aerobic conditions (Sajbidor et al., 1995). Ergosterol plays an important role in yeast stress tolerance. It stabilizes cell membranes independently of heatshock proteins or trehalose. Swan and Watson (1999) concluded that membrane lipid composition plays a more consistent role in stress tolerance than trehalose, heat-shock proteins, or ergosterol. The S. cerevisiae FY 169 genome type strain also has a relatively low total sterol content and tends to accumulate lanosterol, the sterol precursor (Danm et al., 1999).

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Glucose grown cells 10g

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Subcellular fraction FIG. 2. Sterol composition of subcellular membranes from late logarithmic phase cells of Saccharomyces cerevisiae FY 169 cultivated on 2% glucose or 2% ethanol at 30°C under aerobic conditions (Tuller et al., 1999). PM = plasma membrane, Mito = mitochondria, Mic = microsomes (endoplasmic reticulum).

Phospholipids have not been examined as extensively as fatty acids and ergosterol for their effects on ethanol tolerance, but recent studies by Chi et al. (1999) suggest that phosphatidylinositol (PI) may be critical in determining ethanol tolerance in yeast strains that are capable of high ethanol production. PI is among the most abundant phospholipids of yeast membranes (Fig. 3), and addition of inositol increases phospholipid synthesis. Inositol can become the factor determining ethanol tolerance in cells that have sufficient fatty acids and ergosterol. Synthesis of phosphatidylcholine (PC) from phosphatidylethanolamine (PE) is induced by inositol (Greenberg and Lopes, 1996). Early work by Mishra and Prasad (1988) showed that cells enriched with phosphatidylserine had greater tolerance to ethanol, and they concluded that

246

THOMAS W. JEFFRIES and YONG-SU JIN Glucose grown cells

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[ ] PtdDM-Etn [] CL

[ ] PA [ ] Lyso-PL [ ] Lyso-Pt:lEtn [ ] Others

Subcellular fraction F]c. 3. P h o s p h o l i p i d c o m p o s i t i o n of subcellular m e m b r a n e s from late logarithmic p h a s e cells of Saccharomyces cerevisiae FY 169 cultivated on 2% glucose or 2% ethanol at 3O°C u n d e r aerobic c o n d i t i o n s (Tuller et al., 1999). PM -- p l a s m a m e m b r a n e = Mito = mitochondria, Mic = m i c r o s o m e s (endoplasmic reticulum), Vac = vacuoles, PtdCho = p h o s p h a t i d y l c h o l i n e , PtdEtn = p h o s p h a t i d y l e t h a n o l a m i n e , PtdIns = p h o s p h a t i d y l i n o s i tol, PtdSer = p h o s p h a t i d y l s e r i n e , PtdDMEtn -- p h o s p h a t i d y l m e t h y l e t h a n o l a m i n e , CL = cardiolipin, PA = p h o s p h a t i d i c acid, Lyso-PL = l y s o p h o s p h o l i p i d s , Lyso-PtdEtn = lysophosphatidylethanolamine.

this resulted from an alteration in the charge of the membrane phospholipids rather than changes in membrane fluidity. Even so, ergosterol appears to be more important than phospholipids. For example, Chi et al. (1999) found that a more ethanol-tolerant strain of S. cerevisiae had a higher ergosterohphospholipid ratio, a higher incorporation of longchain fatty acids in total phospholipids, and a slightly higher proportion of unsaturated fatty acids in total phospholipids than a less ethanol-tolerant strain. Likewise, el Dein (1997) found that in Candida

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lambica, a yeast with only moderate ethanol tolerance, growth in up to 8% (v/v) ethanol was accompanied by an increase in total fatty acids but a decrease in phospholipids. Murakami et al. (1996) did not observe a significant correlation between phospholipid content and tolerance to freezing, but it is possible that these and other researchers have neglected inositol as a limiting factor in yeast phospholipid biosynthesis. Neirinck et al. (1984) and Shi and Jeffries (1998) have examined the effects of lipids on anaerobic growth of pentose-fermenting yeasts. Neirinck et al. (1984) were not able to obtain anaerobic growth of P. tannophilus even after incorporating essential lipids into the medium. Shi and Jeffries (1998) did obtain anaerobic growth of P. stipitis transformed with ScUral by adding ergosterol, Tween 80, and linoleic acid to the medium (Jessens et al., 1983) along with fumarate as a terminal electron acceptor for the DHODase activity (Nagy et al., 1992). However, the transformants grew on glucose but not on xylose. Ethanol yields were comparable to those attained by S. cerevisiae control cultures. To our knowledge, the effect of ergosterol, unsaturated fatty acids, inositol, or phospholipids on ethanol or thermotolerance has not been investigated with xylose-fermenting yeasts. Eubacteriales do not produce ergosterol, and, for the most part, they do not tolerate high ethanol concentrations either. Z y m o m o n a s mobilis produces large amounts of ethanol rapidly. It has cardiolipin, phosphatidylethanolamine, phosphatidylglycerol, and phosphatidylcholine as its major phospholipids (Carey and Ingram, 1983). In yeasts, cardiolipin is found almost exclusively in the mitochondria (Fig. 3), and its concentration increases when cells are cultivated in the presence of ethanol. Z. mobilis also produces large amounts of pentacycline triterpanoid lipids known as hopanoids (Moreau et al., 1997). The structures of these compounds superficially resemble sterols. In the absence of ethanol, hopanoid content can be as high as 30% of total cellular lipids. In the presence of ethanol, complex changes occur in the levels of hopanoids and other membrane constituents. Various researchers have hypothesized that either hopanoid or cis-vaccenic acid in the phospholipids of the bacterial membrane could account for the ethanol tolerance of Z. mobilis, but such relationships have not been established unambiguously. The heterologous expression of genes for hopanoid biosynthesis (Perzl et al., 1998) could help establish whether these lipids play a major role in bacterial ethanol tolerance. Hobley and Pamment (1994) showed that one of the reasons Z. mobilis is able to tolerate high concentrations of ethanol is that it adapts rapidly to changes in extracellular ethanol without significant viability loss.

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THOMAS W. JEFFRIESand YONG-SUJIN

The biochemical, molecular, genetic (Daum et al., 1998), and environmental (Sajbidor, 1997) factors affecting lipid synthesis in S. cerevisiae and other organisms have been reviewed. Lipid composition by itself probably does not affect ethanol tolerance. Rather, the physiological function that the various lipids play in maintaining membrane integrity is the critical factor, and the most important function of the membrane is nutrient transport and accumulation. B. PLASMAMEMBRANEH+-ATPASE The plasma membrane (PM) proton pump (H÷-ATPase) of yeast couples ATP hydrolysis to proton extrusion, thereby providing the means for solute uptake by secondary transporters and for regulating cytoplasmic pH. By pumping protons out of the cytoplasm, the H÷-ATPase acidifies the external medium, and makes the cytoplasm relatively alkaline. The PM H÷-ATPase is a highly abundant, essential enzyme in S. cerevisiae. It belongs to the family of P-type ATPases, a class of enzymes that includes the Na÷,K÷-ATPase and the gastric H÷,K÷-ATPase (Monk et aL, 1995). S. cerevisiae possesses two plasma membrane H÷-ATPase isoforms: Pmal and Pma2. They are 89% identical at the protein level (Supply eta]., 1995), but they exhibit different activation, kinetic, and regulatory properties, which suggests that they have different functions. The major yeast plasma membrane H÷-ATPase is encoded by the essential P m a I gene. The P m a 2 gene encodes an H÷-ATPase that is functionally interchangeable with the one encoded by P m a I , but it is expressed at a much lower level than P m a I and it is not essential (Fernandes and S~-Correia, 1999). P m a l is primarily responsible for proton translocation in S. cerevisiae. The expression of P m a l is regulated by glucose and by the Tuf/Rapl/Grfl transcription factor in S. cerevisiae (Rao et al., 1993). Deletion of A p a I causes defective regulation of P m a 1 expression by glucose but has no noticeable effect on expression of other Tuf-regulated genes (Garcia-Arranz et al., 1994). Pmal is activated by glucose (Serrano, 1983) through a mechanism that involves the Cdc25 gene product (Portillo and Mazon, 1986). Cdc25 and Ras are two proteins required for cAMP signaling in S. cerevisiae. Cdc25 is the prototype guanine nucleotide exchange protein that activates Ras. Ras, in turn, activates adenylyl cyclase (Mintzer and Field, 1999). The N-terminal portion of Cdc25 is important in signal processing (Gross et aL, 1999). The specific activity of Pmal increases with growth temperature. However, this activation does not result from increased Pmal synthesis.

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Expression levels actually decrease along with the amount of enzyme present in the membrane (Viegas et al., 1995). Ethanol also activates ATPase in S. cerevisiae (Monteiro and S~i-Correia, 1998). Ethanol-induced stimulation of plasma membrane H+-ATPase increases the Vma~. The affinity for Mg+ATP decreases. Sublethal concentrations of ethanol enhance expression of Pma2 while reducing expression of P m a l . The inhibition of P m a l expression by ethanol correlates with a decline in the content of plasma membrane ATPase as measured by immunoassays (Monteiro et al., 1994). Therefore, the observed increase in activity following stress is not attributable to synthesis of new protein, but rather to activation of the existing enzyme. Piper et al. (1997) reported that in S. cerevisiae, Hsp30 is a stress-inducible regulator of ATPase activity. Hsp30 is induced by heat shock, ethanol exposure, severe osmostress, weak organic acid exposure, and glucose limitation. Hsp30 induction downregulates stimulation of H ÷ATPase caused by stress. Plasma membrane H+-ATPase consumes a substantial fraction of the ATP generated by the cell. Hsp30 might therefore play an energy conservation role during prolonged stress exposure or glucose limitation (Piper et al., 1997). Barbosa and Lee (1991) were the first to study the effects of alcohols on PM ATPase activity in a xylose-fermenting yeast. In-vitro studies showed that the amount of alcohol required to inhibit ATPase activity 50% decreased with increasing chain length. Moreover, the concentration required to inhibit ATPase activity corresponded approximately with the concentration required to inhibit growth. Meyria] et al. (1995a,b, 1997) carried out extensive studies of PM ATPase activity in R stipitis. The enzyme from this yeast attained its highest activity between pH 7.3 and 7.5 at 35°C. This value is very far from the ATPase activities in S. cerevisiae or K. marxianus (Rosa and SS-Correia, 1992), and Meyrial et aL (1995a) concluded from a comparison of the pH optima of ATPases reported from different yeasts that the most ethanol-tolerant have ATPases with low pH optima. They also noted that the B stipitis and K. marxianus K m values for ATP are about 2- to 10-fold higher than the values reported for S. cerevisiae. A higher affinity for ATP should enable the ATPase to maintain proton gradients even under conditions where energy generation is low, such as during anaerobic growth. They found that glucose stimulates the activity of P. stipitis PM ATPase threefold in comparison to the activity measured when xylose is used as the carbon source (Meyrial eta]., 1995a). The stimulation occurs mainly through doubling of Vm~,,. The Km value for ATP remains similar in both glucose- and xylose-grown cells. The stimulation of ATPase activity correlates with an increase in the maxi-

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THOMAS W. JEFFRIES and YONG-SU JIN

mum amount of ethanol produced by glucose-grown cells. It is possible that glucose activation of ATPase is related to the observation by Shi and Jeffries (1998) that B stipitis transformed with ScUral will grow anaerobically on glucose but not on xylose. The concentration of ethanol that is required to cause a 50% inhibition of ATPase activity is 34.5 and 41.4 g/liter in xylose- and glucose-grown cells, respectively. For both glucose- and xylose-grown cells, the concentration of ethanol required to inhibit growth reduced ATPase activity of control cells (grown without ethanol) by 60%. Ethanol also stimulates ATPase activity in P. stipitis (Meyrial eta]., 1995b); unlike S. cerevisiae, however, the extent of stimulation depends on the carbon source used for growth. With glucose, addition of 1% ethanol doubles ATPase activity. No significant increase in ATPase activity is observed with xylose-grown cells. Moreover, the stimulatory effects of glucose and ethanol are additive. Similar activation of ATPase activity has been reported for S. cerevisiae and K. marxianus (Rosa and Sfi-Correia, 1991, 1992). It is unclear whether ATPase activity in P. stipitis involves one protein or two, as in the case of S. cerevisiae. Physiological concentrations of ethanol do not affect passive entry of protons into either glucose- or xylose-grown P. stipitis (Meyrial et al., 1997). In contrast, active proton extrusion is affected differently by ethanol, depending on the carbon source used for cell growth. Low, physiological concentrations of ethanol decrease the proton extrusion rate in glucose-grown cells, but only nonphysiological ethanol concentrations reduce proton extrusion in xylose-grown cells. By comparing the rates of proton efflux with ATPase activity, Meyrial et al. (1997) concluded that glucose activates both ATP hydrolysis and the protonpumping activities of H÷-ATPase, whereas ethanol causes an uncoupling between ATP hydrolysis and proton extrusion. Heat shock and ethanol stress cause similar changes in the protein composition of the PM. Heat shock and ethanol stress each reduce the levels of plasma membrane H+-ATPase while simultaneously increasing the specific activity of the remaining enzyme. Proton efflux places an energy demand but may help to counteract the adverse effects on membrane permeability that result from stress (Piper, 1995). Plasma membrane ATPase activity is essential for basal heat resistance. Moreover, thermotolerance is enhanced by prior exposure to stress. Prestressed cells are able to protect the proton gradient longer than cells that have not adapted to heat or ethanol (Coote et a]., 1994). Stress resistance is largely conferred by a mechanism independent of the ATPase. A mutant strain with reduced expression of plasma membrane ATPase showed significantly less resistance to lethal heat than

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the wild-type parent. However, prior exposure to sublethal temperatures induced levels of thermotolerance similar to the parent strain, suggesting that the mechanism of sublethal heat-induced thermotolerance is independent of ATPase activity (Coote et al., 1994). C. MITOCHONDRIAL STABILITY

Mitochondria are essential for ethanol tolerance in S. cerevisiae (Aguilera and Benffez, 1985). This presents a difficulty because ethanol also tends to induce the formation of petite mutants (Jim~nez et al., 1988). In fact, the mitochondria in populations of flor yeasts isolated from sherry wine show an unusual level of mitochondrial DNA polymorphism (Ibeas and Jim~nez, 1997). Treatment with ethanol did not cause an increase in the spontaneous mutation rate of nuclear genes, but petite mutants increased from a low spontaneous level (about 1%) to nearly half the survivors at 24% ethanol (Bandas and Zakharov, 1980). Chi and Arneborg (1999) demonstrated that ethanol tolerance correlated with a lower frequency of ethanol-induced respiratory-deficient mutants, a higher ergosterohphospholipid ratio, a higher proportion of phosphatidylcholine, a lower proportion of phosphatidylethanolamine, a higher incorporation of long-chain fatty acids in total phospholipids, and a slightly higher proportion of unsaturated fatty acids in total phospholipids (Chi and Arneborg, 1999), Manganese superoxide dismutase (MnSOD), which is found in mitochondria, is essential for ethanol tolerance in S. cerevisiae (Costa et al., 1993). In E. coli, MnSOD is associated with DNA (Steinman et al., 1994), and in yeasts it might serve to prevent damage to mitochondrial DNA. Ethanol toxicity correlates with the production of reactive oxygen species in the mitochondria, as evidenced from results obtained with respiration-deficient mutants (Costa et al., 1997). The freezing-andthawing process also generates free radicals and oxidative stress, which cause major injury to cells during aerobic freezing and thawing. The superoxide radical is mainly generated on the matrix side of the inner mitochondrial membrane of yeast cells (Balzan et al., 1999). Superoxide radicals from the mitochondrial electron transport chain initiate damage in the cytoplasm (Park et al., 1998). Normally such radicals are dispersed by cytoplasmic Cu and Zn superoxide dismutase. It is also possible to protect the cells against oxidative stress by heterologous expression of Escherichia coli iron superoxide dismutase (FeSOD) when it is targeted to the mitochondrial matrix (Balzan et al., 1995). Nedeva eta]. (1993) used thermotolerance as a screening mechanism to identify yeasts that produce heat-stable superoxide dismutase. After

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THOMAS W. JEFFRIES and YONG-SU JIN

screening 10 thermotolerant yeasts from the genera Saccharomyces, Kluyveromyces, and Pichia, they found that higher cell yields at 45°C were obtained with two thermotolerant strains of K. lactis, which also produced thermostable superoxide dismutase. Because the free radicals generated during respiration are so destructive, they might be the root cause for the frequency of petites formed at high temperatures and elevated ethanol level. D. TREHALOSE Tolerance to heat, freezing, oxidation, and desiccation overlap to some extent with ethanol tolerance, and all but ethanol tolerance correlate with trehalose accumulation (Piper, 1995; Eleutherio eta]., 1995; Lewis et al., 1997). Roles for trehalose in preventing protein denaturation during desiccation or at elevated temperature are fairly well established, but its role in ethanol tolerance is less clear. Trehalose is barely detectable in log-phase cells grown on glucose, but it accumulates up to 20% of the total dry weight of the cell in stationary phase cells as well as in exponential-phase cells growing at elevated temperature. Cells growing on nonfermentable carbon sources have high levels of trehalose in both log-phase and stationary-phase cells (Nwaka and Holzer, 1998). Trehalose is a nonreducing disaccharide of glucose that is present in yeasts and higher organisms. It enables cells to survive desiccation and osmotic and thermal stress (Singer and Lindquist, 1998). Trehalose accumulates in the cytosol of yeast during heat shock, where it provides protection to proteins. Trehalose is synthesized from UDP-glucose and glucose-6-phosphate in a two-step reaction catalyzed by the trehalose6-phosphate synthase (Tpsl) and trehalose-6-phosphate phosphatase complex (Piper, 1993). Ribeiro et al. (1997) used a tpsI mutant of the fission yeast Schizosaccharomyces pombe in a genetic approach to determine the specific role of trehalose in a heat-induced cell. They showed that the tpsl mutant has a serious defect in heat-shock-induced acquisition of thermotolerance at 40 to 42.5°C. Earlier studies that used cycloheximide to block protein synthesis found that heat shock did not induce S. pombe to produce trehalose (DeVirgilio et al., 1990). The discrepancy between these two studies might be attributable to the toxicity of cycloheximide. When cells are heat shocked, trehalose immediately starts to accumulate, even before the induction of enzymes for trehalose metabolism (Neves and Francois, 1992). This shows that rapid increase in trehalose is related not to induction of enzymes for trehalose synthesis but to activation of preexisting enzymes. Trehalose

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stabilizes enzymes and also greatly reduces their activities. Trehalose degradation is therefore necessary in order for the cell to resume activity once the stress condition has passed. This is carried out by trehalase. In vitro, the activity of trehalase decreases as the activity of the trehalose-6-phosphate synthase-phosphate complex increases (Neves and Francois, 1992). Hexokinase also appears to act synergistically with trehalose synthesis because it provides glucose-6-phosphate for trehalose synthesis. After incubation in D-xylose, an inhibitor of hexokinase, trehalose levels in these cells dropped almost in 90%, confirming the involvement of both hexokinases in the accumulation of trehalose (Peixoto and Panek, 1999). Probably trehalose will be found to play a role in thermotolerance, osmotolerance, and resistance to ethanol because the mechanism is so widely distributed among yeasts, fungi, and plants. For example, trehalose synthesis is an important factor in thermotolerance of the fission yeast S. pombe. On induction of trehalose-6P synthase, the elevated levels of intracellular trehalose correlated not only with increased tolerance to heat shock but also with resistance to freezing and thawing, dehydration, osmostress, and toxic levels of ethanol (Soto et al., 1999). The broad nature of this stress response suggests that trehalose may be the metabolite underlying the overlap in induced tolerance. An exhaustive search of the current literature did not uncover any reports about trehalose synthesis in xylose-fermenting yeasts. E. HEAT-SHOCK PROTEINS

When yeasts confront extreme temperatures (50°C), synthesis of heatshock proteins (Hsps) is induced (Parsell and Lindquist, 1993). These proteins play important roles in protecting cells from extreme temperature. In S. cerevisiae, Hsp90 and Hsp70 are expressed even at normal temperature and are only modestly induced by heat (Craig and Jacobsen, 1984). Hspl04 is expressed at a very low level at normal temperatures and is very strongly induced by heat. Hspl04 expression is critical for survival following heat shock in yeast (Lindquist and Kim, 1996). Recent research indicates that Hspl04 promotes survival by resolubilizing heat-damaged aggregated proteins (Parsell et al., 1994; Schirmer et al., 1998). The function of Hspl04 is to repair damage after stress rather than to prevent damage from stress, as in the case of trehalose. Hsp30, the integral plasma membrane heat-shock protein of S. cerevisiae, is a stress-inducible regulator of plasma membrane H+-ATPase (Piper et al., 1997). Cells lacking Hsp30 showed greater plasma membrane H+-ATPase activity than cells with normal levels of Hsp30. This

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THOMAS W. JEFFRIES and YONG-SU JIN

indicates that Hsp30 reduces stimulation of H÷-ATPase. Hsp30 reduces the Vmaxof H÷-ATPase in heat-shocked cells. The action of Hsp30 is lost with deletion of the C-terminal 11 amino acids of H÷-ATPase, a deletion that does not abolish stress stimulation of this enzyme (Braley and Piper, 1997). This suggests that Hsp3O modulates ATPase activity by interacting with this portion of the enzyme.

Vl. Summary The mechanisms underlying ethanol and heat tolerance are complex. Many different genes are involved, and the exact basis is not fully understood. The integrity of cytoplasmic and mitochondrial membranes is critical to maintain proton gradients for metabolic energy and nutrient uptake. Heat and ethanol stress adversely affect membrane integrity. These factors are particularly detrimental to xylose-fermenting yeasts because they require oxygen for biosynthesis of essential cell membrane and nucleic acid constituents, and they depend on respiration for the generation of ATP. Physiological responses to ethanol and heat shock have been studied most extensively in S. cerevisiae. However, comparative biochemical studies with other organisms suggest that similar mechanisms will be important in xylose-fermenting yeasts. The composition of a cell's membrane lipids shifts with temperature, ethanol concentration, and stage of cultivation. Levels of unsaturated fatty acids and ergosterol increase in response to temperature and ethanol stress. Inositol is involved in phospholipid biosynthesis, and it can increase ethanol tolerance when provided as a supplement. Membrane integrity determines the cell's ability to maintain proton gradients for nutrient uptake. Plasma membrane ATPase generates the proton gradient, and the biochemical characteristics of this enzyme contribute to ethanol tolerance. Organisms with higher ethanol tolerance have ATPase activities with low pH optima and high affinity for ATE Likewise, organisms with ATPase activities that resist ethanol inhibition also function better at high ethanol concentrations. ATPase consumes a significant fraction of the total cellular ATP, and under stress conditions when membrane gradients are compromised the activity of ATPase is regulated. In xylose-fermenting yeasts, the carbon source used for growth affects both ATPase activity and ethanol tolerance. Cells can adapt to heat and ethanol stress by synthesizing trehalose and heat-shock proteins, which stabilize and repair denatured proteins. The capacity of cells to produce trehalose and induce HSPs correlate with their thermotolerance. Both heat and ethanol increase the frequency of petite mutations and kill cells. This might be attribut-

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able to membrane effects, but it could also arise from oxidative damage. Cytoplasmic and mitochondrial superoxide dismutases can destroy oxidative radicals and thereby maintain cell viability. Improved knowledge of the mechanisms underlying ethanol and thermotolerance in S. cerevisiae should enable the genetic engineering of these traits in xylose-fermenting yeasts.

Acknowledgments The authors acknowledge H. K. Sreenath for the use of unpublished data and N. Q. Shi for a critical reading of the manuscript. Research for this review was supported in part by NREL contract No. XXL-9-2903402 to T. W. J.

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Microbial Degradation of the Pesticide Lindane (7-Hexachlorocyclohexane) BRAJESH KUMAR SINGH AND RAMESH CHANDERKUHAD

Department of Microbiology University of Delhi New Delhi 110021, India

AJAY SINGH

Department of Biology University of Waterloo Waterloo, Ontario, N2L 3G1, Canada

K. K. TRIPATHI AND P. K. GHOSH

Department of Biotechnology Ministry of Science and Technology New Delhi 110003, India

I. II. III. IV. V. VI. VII. VIII. IX.

Introduction Environmental Fate of Hexachlorocyclohe×ane Toxicological Effects of Hexachlorocyclohexane Biochemica] Mechanisms of Hexachlorocyclohexane Degradation Bacterial Degradation of Hexachlorocyclohexane Fungal Degradation of Hexachlorocyclohexane Algal and Cynobacterial Degradation of Hexachlorocyclohexane Future Prospects Nomenclature References

I. Introduction Increased and indiscriminate use of chemical pesticides causes considerable environmental pollution and human health problems due to their toxicity, persistence, and transformation into hazardous metabolites. On the other hand, the use of chemica] pesticides cannot be discontinued in the near future. Halogenated aromatic pesticides are an important group, as halogens are often implicated as a reason for persistence and toxicity of such compounds (Neilson et al., /985). Halogenated aromatic compounds, particularly chlorinated aromatics, have been utilized in agriculture and industry for many years as insecticides, 269 ADVANCES1NAPPLIEDMICROBIOLOGY,VOLUME47 Copyright© 2000by AcademicPress All rightsof reproductionin any formreserved. 0065-2164/00$25.00

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herbicides, solvents, lubricants, plasticizers, and insulators (Singh et a]., 1987; Fetzner, 1998). Hexachlorocyclohexane (HCH), commonly known as BHC (benzene hexachloride), is one of the most extensively used organochlorine insecticides. HCH is a collective term that encompasses the eight isomers of 1,2,3,4,5,6-hexachlorocyclohexane (Willett et a]., 1998). Figure 1 shows structures of different HCH isomers. HCH is available in two technical grades: technical HCH (a mixture of different isomers, including {x-, [~-, 7-, and ~-HCH) and 7-HCH, also known as lindane (CAS CI

CI

CI

CI

Alpha enantiomers

t;I CI

CI CI

Beta

Gamma CI

CI

~1

CI

C!

Delta

Epsilon CI

CI

CI

CI

CI

Eta

Theta

FIG. 1. Structures of eight isomers of hexachlorocyclohexane. Two ~-enantiomers of c~-hexachlorocyclohexane exist.

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MICROBIAL DEGRADATION OF THE PESTICIDE LINDANE

Registry No. 608-73-1). As a broad-spectrum insecticide, technical HCH has been used for both agricultural and nonagricultural purposes. The various formulations of lindane and technical HCH have many trade names, including Agrocide, Ben-Hex, Gammexane, Kwell, Lindatox, Quellada, and Tri-6. They have been used in seed and soil treatment on a variety of crops, in widespread insecticidal applications on crops, ornamental trees, lawns, and wood products, and for vector control (Li et al., 1998a). Medicinally, 7-HCH has been applied topically to people for the treatment of lice and scabies. The isomers of HCH differ significantly from each other with respect to their persistence and toxicity to insects, birds, mammals, and other nontarget organisms. Lindane (7HCH) is the only isomer with insecticidal activity, and the use of all other isomers has been banned internationally. Lindane has been extensively used worldwide despite its persistent nature, tendency toward bioaccumulation, and toxicity to higher animals, as well as the possible toxicological and environmental problems of its residues (Marks et al., 1989; Waliszewski, 1993; Graham, 1993). Earlier, the United States, China, India, and Japan were the major users of technical HCH (Li et al., 1998a). China produced more than four million tons of HCH from 1952 to 1983. Up until the 1970s, the United States and Japan used about 350,000 and 400,000 tons of HCH, respectively (Table I). It has been estimated that HCH use in India was about one million tons until 1995. In the 1970s, most developed countries either imposed a total ban or restricted the use of HCH, while many developing nations restricted/banned the production and usage of this insecticide in the 1980s (Voldner and Li, 1995). Nevertheless, organochlorine insecticides such as lindane are now widespread contaminants in the global ecosystem (Li et al., 1998b). Ideally, pesticides should be detoxified after their initial benefits to agriculture or industry TABLE I MalOR PRODUCERSAND USERSOF TECHNICALHEXACHLOROCYCLOHEXANE(FAO, 1993)

Country China India Japan USA

Years of usage/ production 1952-83 1948-95 1948-70 1947-76

Total amount (mt) 4.46 1.00 0.40 0.35

Total usage density (t/kha) 44.4 5.9 83.9 1.9

272

B.K. SINGH et al.

have been achieved. Most studies on pesticide degradation have aimed at gaining a basic understanding of biodegradation activities of microorganisms and to devise suitable bioremediation methods for removing or detoxifying concentrated pesticide residues (Engst et al., 1977, 1979; Macholz and Kujawa, 1985; Deo et al., 1994). The isolation and genetic manipulation of microorganisms for use in bioremediation has received considerable attention (Pemberton, 1983; Pemberton and Wynne, 1984; Reineke, 1986; Singh et al., 1989; Imai et al., 1991; Lal et al., 1995). Factors that affect the degradation of organochlorine compounds include the number of attached halogens, the stability of the genes encoding the detoxification pathway, the mass transfer properties of the contaminant, and the contaminant-degrading organisms and properties of the soil or water (Cork and Khalil, 1995). Various species of bacteria belonging to the genera Pseudomonas, Bacillus, and Clostridium can degrade chloroaromatic compounds by utilizing them as a sole carbon and energy source (Lal and Saxena, 1982; Bouwer and Zehnder, 1992; Choudhry and Chapalamadugu, 1992). White-rot fungi also degrade chlorinated hydrocarbons. The role of several basidiomycetes (e.g., Phanerochaete chrysosporium, Pleurotus sajor caju, Phanerochaete sordida, Cyathus bulled and Coriolus versicolor) and their enzyme systems have been reviewed (Bumpus et al., 1985; Kennedy et al., 1990; Higson, 1991; Shah et al., 1992; Barr and Anst, 1994; Mougin et al., 1996; Arisoy and Kotankaya, 1997; Singh and Kuhad, 1999). The capabilities of microalgae for degradation of aromatic compounds have been explored only relatively recently (Semple et al., 1999). General reviews that deal with the environmental fate of different pesticides have been published (Chakraborty, 1982; Barik, 1984; Lal et al., 1986; Commandeur and Parsons, 1990; H/iggblom, 1992; Heitzer and Sayler, 1992; Liu and Suflita, 1992; Deo et al., 1994; Kumar et al., 1996; Fetzner, 1998). A comprehensive account of environmental persistence and toxicology, and the biochemical and molecular aspects of HCH biodegradation is presented in this chapter.

II. Environmental Fate of Hexachlorocyclohexane

Organochlorine compounds are generally recalcitrant to degradation. HCH isomers persist in soils in a wide variety of climates. Lichtenstein and Polivka (1959) reported that under certain conditions lindane could persist in the environment for up to 11 years, with about 10-14% of the applied lindane extracted from silt loam soil 3 years after application (Lichtenstein and Schulz, 1959). According to Kearney et al.

MICROBIALDEGRADATIONOF THE PESTICIDELINDANE

273

(1964), 75-100% of lindane had disappeared from soil within about 30 years. Wheatley (1965) calculated the half-life of lindane as 4-6 weeks for surface treatment and 15-20 weeks for soil incorporation. In contrast, Edwards (1966) estimated the half-life of lindane in soil to be 1.2 years. One research group found residual levels of 3-8% of applied lindane in soil 15 years after application (Voerman and Besemer, 1970), whereas another group (Stewart and Fox, 1971) failed to detect any lindane residue 12 years after a surface spray. In tropical regions, the environmental persistence of organochlorine compounds including HCH is slightly less than in temperate regions. Lindane, apparently the least persistent of HCH isomers in the environment, undergoes rapid decomposition in submerged soils in tropical regions (Raghu and MacRae, 1966; MacRae et al., 1967; Tsukano and Kobayashi, 1972). Kathpal eta]. (1976) reported that 75-100% of BHC had disappeared from soils 2-3 months after application. Kushwaha et al. (1977) observed 70-80% loss of BHC after 3 months in a sandy loam soil. In clay loam soil, 73-90% of applied BHC was not detected after 3 months (Srivastava and Yadav, 1977). In flooded paddy soil, between 45 and 100% of applied lindane disappeared within 1 month as compared to a 2% loss in unflooded soil during this period (Yoshida and Castro, 1970). P s e u d o m o n a s putida isolated from soil was found to convert 7-HCH to o~-HCH, which explains the high level of ~-HCH in the environment (Benzet and Matsumura, 1973). However, ~-HCH and 7-HCH both disappeared rapidly from flooded soil (Siddaramppa and Sethunathan, 1975). The contamination of water by organochlorine pesticides including HCH has been widely reported. Edwards (1973) found 0.011 to 28 rag/liter of HCH in river water in the United States, with 0.16 mg/liter of lindane in the northern part of the Mississippi (Rihan et al., 1978). Miliadis (1994) reported 0.005 to 0.014 mg/liter of lindane in the water of the Lake Lliki (Athens, Greece). Contamination of Ganges water with HCH or lindane at different places in India has been reported (Agnihotri et al., 1994; Ahmad et al., 1996). Kumari eta]. (1996) observed 2.2 ± 0.9 pg/liter of HCH residues in pond water in Haryana, India. Dua et al. (1996) found concentrations above the maximum permissible limits of HCH in drinking water in 13 villages of Uttar Pradesh, India. They also observed that HCH residues were 3.6 times higher in pond water without fish as compared to their respective concentration in ponds with fish, perhaps implying that HCH is accumulated in fishes. [~-HCH had the lowest value, indicating that accumulation of this isomer in the environment was due possibly to its stability and resistance to micro-

274

B.K. SINGH et al.

bial degradation (Agnihotri et al., 1994). HCH residues at concentrations higher than the acceptable daily intake (ADI) were reported in water from five lakes of Nainital, India, although HCH had not been used nearby. Atmospheric transportation of HCH was suggested as the mechanism of contamination (Dua et al., 1998a). HCH residues in air have been reported (Edwards, 1973). Tanabe and Tatsukawa (1980) measured HCH in air over the Bay of Bengal. HCH residues in the air over Delhi varied from O to 21797 ng -3 (Kaushik et al., 1987). HCH residues have also been detected in food grains such as chickpea, pigeon pea, rice, and wheat (Mukherjee et al., 1989; Kaphalia eta]., 1990; Ahuja and Awasthi, 1993).

III. Toxicological Effects of Hexachlorocyclohexane The large-scale use of HCH coupled with its extreme persistence and slow degradation have led to potential health hazards (Banerjee et al., 1996a). Residues of HCH find their way into various food commodities and ultimately carry over through food chains into humans (Ramachandran et al., 1984). HCH residues in human milk have been reported from different parts of the world (WHO, 1991, 1992; Ejobi, 1996a,b; Barkatina, 1998), including India (Krishnamurti, 1984; Jani et al., 1988; Nair et a]., 1996). Banerjee et al. (1996b) reported considerably higher concentrations of ~-, ~-, and 7-HCH in human milk in the Delhi region of India than those reported from other countries. Such higher concentrations may be due to several reasons, such as the magnitude and frequency of applications, the efficiency of absorption and excretion, the age of the individuals, and their nutritional and socioeconomic status. The presence of HCH in mothers' milk may result in exposure of breast-fed infants to levels higher than accepted daily limits. In addition, Mussalo eta]. (1990) have warned about the possible role of organochlorine residues in breast cancer. Krieger et a]. (1994) detected increased amounts of organochlorine in certain cancerous tissues. In contrast, Newcomb et al. (1994) reported a reduction of breast cancer risk with lactation-linked elimination of organochlorine as a possible mechanism. The deleterious effects of these pesticides on the immune system have also been reported (Saha and Banerjee, 1993; Banerjee et al., 1996a,b). Ejobi eta]. (1996a) found high concentrations of lindane and DDE in Ugandan cow's milk, where both children and adults are exposed to contamination.

MICROBIAL DEGRADATION OF THE PESTICIDE LINDANE

275

The organochlorine pesticides are widely recognized as neurotoxic substances affecting the peripheral and central nervous systems, and causing a hyperexcitability of nerves and muscles (Hassal, 1983). Residues of organochlorine have been detected all over the world in the blood of exposed workers (Griffith and Duncan, 1985; Bouman et al., 1991; Chandra et al., 1992) as well as in people without occupational exposure (Leoni et al., 1989; Sasaki eta]., 1991; Chandra et al., 1992). Chandra et al. (1992) observed that the mean total HCH serum level was 15.9 pg/liter among 52 mango growers in Malihabad, India. High concentrations of HCH in blood serum from the general population in Ahmadabad (India) were also reported (Bhatnagar et al., 1992; Kashyap et al., 1993). Recently, Dua et al. (1998b) observed high concentrations of HCH from the skin lipids of exposed people in India and found a direct relationship between the concentration of HCH in lipids and in blood. Other harmful effects of HCH deposition in human fat are disturbances of lipid metabolism (Carlson and Kalmodin°Hidman, 1972; Barros et al., 1991) and alterations in some membrane-bound proteins and membrane permeability (Magour et al., 1984; Carrero et al., 1989). Shukla et al. (1996) found high absorption (80-95%) of HCH in rat intestinal cells. Siddiqui et al. (1996) studied bioaccumulation of HCH isomers in different tissues of young and old rats. They observed that 7-HCH concentrations were highest in kidney followed by lung, liver, brain, spleen, heart, and blood in younger-aged groups, whereas in older-aged groups the order was spleen, lung, kidney, brain, liver, heart, and blood, suggesting that age influences the distribution profile of 7-HCH. Residues of HCH were also found in various tissues of dolphins (Willet et al., 1998). Harmful effects of organochlorines on nonmammalian animals have also been reported. These chemicals enter into aquatic systems by drifting, runoff, or water application. They accumulate through the food chain. For example, HCH has been reported from the eggs of birds that prey on fishes (Ohlendorf et al., 1985; Hothem and Zador, 1995). Goutner et al. (1997) found higher concentrations of lindane in eggs of the little tern Sterna albifrons. The population of these birds is on the decline in Europe, and lindane contamination of their eggs and its potential effect on egg hatchability are implicated. Becker (1989) and Becker et al. (1993) reported contamination by lindane and its metabolites in larid eggs. Lindane contamination in water birds (Fasola, 1987) and in herons and egrets (Findholt and Trost, 1985; Eitzner et al., 1988) has also been observed. Hazarika and Das (1998) studied histological

276

B.K. SINGH et al.

pictures of fish exposed to different concentrations of HCH and found significant changes as compared to controls. Important changes included disruption of ovarian follicles, vacuolation of cytoplasm of germinal cells, and reproduction in the number of matured ova and secondary oocytes. Bioaccumulation of this organochlorine pesticide also was observed in mussels. Hickey et al. (1997) reported high concentrations of lindane in the freshwater mussel Hyridella m e n z i e s i from the Waikatu River of New Zealand. Livestock meat and dairy products are the prime sources of human dietary exposure to organochlorines, contributing about 60-85% of mean daily intake. These percentages are in accordance with the welldocumented fact that organochlorines predominantly accumulate in the lipid fraction of foods. Recently, Osibanjo and Adeyeye (1997) reported that lindane had the highest mean daily intake among all organochlorine pesticides in Nigeria. Many workers have also studied the toxicological effects of HCH on microorganisms. In flooded soil, HCH inhibited both oxidation and reduction reactions (Ray et al., 1980; Pal et al., 1980). Addition of HCH retarded the drop in redox potential and maintained the soil in an oxidized state (Pal et al., 1980). Kumaraswamy et al. (1997) reported that HCH was inhibitory to the population of total bacteria, methanotrophs, and ammonium oxidizers. Despite oxidized conditions in the flooded soil amended with HCH, methane oxidation (monitored aerobically) was inhibited, which suggested that HCH is also toxic to methane oxidizing bacteria. Since autotrophic ammonium oxidizers have been implicated in oxidation of methane (Bedard and Knowles, 1989), inhibition of autotrophic ammonium oxidizers and other methane oxidizers by HCH would explain the low methane oxidizing activity in soil samples amended with HCH. HCH application at recommended levels affected nitrogen fixation by stimulating nitrogenase activity (Patnaik et al., 1994). However, HCH has been reported to inhibit nitrification in a rice field (Ray et al., 1980). Chemical, physical, or biological agents may degrade pesticides reaching the soil sediments or water ecosystem. Biological and chemical decomposition of xenobiotic compounds in such environments is always influenced by changes in many physicochemical parameters, such as temperature, pH, ion concentration, and redox potential. However, the microorganisms present in soil and water are a major factor in the degradation of these pesticides. Transformation of pesticides does not occur only by one type of organism, or by a few selected organisms, but is affected by consortia of microorganisms such as actinomycetes,

MICROBIAL DEGRADATION OF THE PESTICIDE LINDANE

277

fungi, and bacteria, and complete mineralization is brought about by the whole microflora inhabiting a contaminated site (Kumar et al., 1991). Table II presents a list of microorganisms capable of degrading HCH. IV. Biochemical Mechanisms of Hexachlorocyclohexane Degradation

HCH isomers are biodegraded via dehalogenation, dehydrohalogenation, isomerization, and oxidation. Chemical reactions involved in degradation of HCH are shown in Figure 2.

TABLE II HEXACHLOROCYCLOHEXANE-DEGRADING MICROORGANISMS Organism

Reference

Bacteria Aerobacter aerogenes Bacillus cereus Bacillus megaterium Citrobacter freundii Clostridium rectum Escherichia coli Pseudomonas flourescens Pseudomonas putida Pseudomonas paucimobilis Pseudomonas sp. Sphingomonas paucimobilis

Mecksongsee and Guthrie (1965) Mecksongsee and Guthrie (1965) Mecksongsee and Guthrie (1965) Jagnow et al. (1977) Jagnow et al. (1977) Francis et al. (1975) Mecksongess and Guthrie (1965) Benzet and Matsumura (1973) Bachmann et al. (1988a,b) Sahu et al. (1990) Johri et al. (1998)

Cyanobacteria Anabaena spp. Nostoc ellipsossorum

Kurtiz and Wolk (1995) Kurtiz and Wolk (1995)

Microalgae Chlorella vulgaris Chlamydomonas reinhardtii

Kobayashi and Rittmann (1982) Kobayashi and Rittmann (1982)

Fungi Coriolus ( Trametes) hirsutus Cyathus bulleri Phanerochaete chrysosposium Phanerochaete sordida Pleurotus sajor-caju

Singh and Kuhad (1999) Singh (1999) Bumpus et al. (1985) Singh (1999) Arisoy and Kolankaya (1997)

B.K. SINGHet

278

~

CI

I~C1

al.

Ch~CI

CI CI T-TCCH ~

CI~ ~ ~CI CI / "/-PCCH /

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CI

CI" T ~C1 CI T-HCH Isomeriz/

CI

CI

OxidativeIntermediates?

CP T "C! CI a, [3 or 8 - HCH

FIG.2. Chemical reactions involved in hexachlorocyclohexane degradation.

Reductive dehalogenation, where a chlorine atom is replaced with a hydrogen atom, is one of the most common pathways of microbial degradation of HCH isomers. 7-HCH (lindane) is converted into 3,4,5,6tetrachlorocyclohexane (7-TCCH). Other metabolites such as tri- and tetrachlorinated benzene and phenols have also been observed. A glutathione-dependent reductive dehalogenase (LinD) is involved in the degradation of 7-HCH by Sphingomonas paucimobilis UT26 (Miyauchi et al., 1998). This enzyme catalyzes the reductive dechlorination of 2,5-dichlorohydroquinone to chlorohydroquinone and then to hydro° quinone. Anaerobic degradation of lindane by Clostridium rectum, Pseudomonas putida, and white-rot fungi occurs via this mechanism (Jagnow et al., 1977; Ohisa et al., 1980). Hydrolytic dehalogenases, involved in hydrolytic dehalogenation of heterocyclic, aromatic, and aliphatic compounds, are the largest group

MICROBIALDEGRADATIONOF THE PESTICIDELINDANE

279

of dehalogenases described to date (Slater et al., 1995, 1997). The degradation of 7-HCH also involves hydrolytic steps, catalyzed by tetrachlorocyclohexadiene dehalogenase (LinB) (Nagata et al., 1993a). Tetrachlorocyclohexadiene dehalogenase (LinB), haloalkane dehalogenase (DhlA), 1-chloroalkane helihydrolase (DlmA), and haloacetate dehalogenase (DehH1) from Moraxella sp. strain B (Kawasaki et al., 1992) share significant sequence homology, suggesting a mechanistic similarity and indicating that they belong to the structural group of "a/J3 hydrolase fold" hydrolytic enzymes (Krooshof et al., 1997) Dehydrohalogenation reactions take place between the saturated chlorinated carbon and the adjacent hydrogen on the neighboring carbon and eliminate HC1 from the haloorganic substrate. Dehydrohalogenation occurs during mineralization of 7-HCH by S. paucimobilis UT26 (Nagata et al., 1993b). The LinA gene encoding the dehydrochlorinase shows no homology to any known dehalogenase gene (Imai et a]., 1991). 7-HCH is converted into 7-pentachlorocyclohexane (7-PCCH) by most of the bacteria studied to date (Yule et a]., 1967; Imai eta]., 1989; Sahu et a]., 1990). Many workers reported the interconversion of four different isomers of HCH (~, [3, 7, and 8) by heating, temperature change, and change in UV light (Wheatstone et al., 1953; Roemer eta]., 1972; Steinwandter, 1976). Some workers have also observed biological isomerization. Benzet and Matsumura (1973) isolated a strain of Pseudomonas putida from soil that converted 7-HCH into a-HCH under laboratory conditions. Other workers also reported such isomerizations in plants, animals, and microbes (Macholz and Kujawa, 1985; Srimathi et al., 1985; Deo et al., 1994). Due to the lack of mixed function oxidase systems in microorganisms, oxidation reactions are more common in higher organisms. V. Bacterial Degradation of Hexachlorocyclohexane Anaerobic degradation of 7-HCH occurs rapidly in paddy soil (MacRae et a]., 1967; Tsukano and Kobayashi, 1972). The pathway of ¥-HCH degradation of Clostridium spp. has been established (MacRae et a]., 1969). Seven out of 13 Clostridium strains tested were able to degrade 7-HCH, but none of the strains were able to transform [3- and 8-HCH. According to the proposed pathway, 7-HCH is converted into 7-TCCH (Heritage and MacRae, 1977), which is further dechlorinated into chlorobenzene (Ohisa et al., 1980). All other known bacteria that degrade lindane anaerobically follow a similar pathway (Jagnow et al., 1977). Complete degradation of lindane occurred in 4-6 days with Clostridium

280

B.K. SINGHet al.

butyricum and C. p a s t e u r i a n u m , while other facultative anaerobes were less active (Jagnow et al., 1977). Lindane degradation was associated with a membrane fraction in C. spheroides, and this activity required reduced glutathione (Heritage and MacRae, 1977). Anaerobic bacteria contain large amounts of corrinoids and other cofactors such as F430. Corrinoids and other transition metal-containing porphyrins have been found to transform lindane to TCCH, chlorobenzene, and benzene in vitro (Marks et al., 1989). Van Eekert et al. (1998) studied anaerobic transformation of [~-HCFI by methanogenic granular sludge from upflow anaerobic sludge blanket (UASB) reactors fed with methanol, volatile fatty acids, or sucrose. The sludge, which had not previously been exposed to HCH, transformed [3-HCH to benzene and chlorobenzene. Both 13- and c~-HCH present in contaminated soil were converted to benzene and chlorobenzene on incubation of the soil under anaerobic conditions. These results indicate that ~-HCH transforming bacteria are present in different anaerobic environments. Biotransformation of four isomers--~-, ~-, 7-, and 8-HCH--under methanogenic conditions was investigated (Middeldorp et al., 1996). In a flow-through column packed with polluted sediment, [3-HCH was completely removed with chlorobenzene detected as the end-product in the effluent. A ~-HCH transforming anaerobic enrichment culture from the column was able to dechlorinate o~-HCH at a comparable rate and 7- and 8-HCH at lower rates. 8-2,3,4,5-TCCH was proposed as an intermediate, and benzene and chlorobenzene were formed as stable end-products. The above results provide a good perspective for the application of a sequential anaerobic/aerobic biological treatment of soils and aquifers polluted with HCH isomers. Middeldorp et al. (1996) proposed a pathway for transformation of ~-HCH under methanogenic conditions (Fig. 3). There are few reports on aerobic biodegradation of 7-HCH. Matsumura et al. (1976) found that P s e u d o m o n a s putida (ATCC 17484) degraded 7-HCH through dechlorination. Tu (1976) reported degradation of 7-PCCH, 7-TCCH, and tetrachlorobenzene (TCB) by Pseudom o n a s NO62. Subsequently, a strain of P s e u d o m o n a s paucimobilis SS86 was isolated from an upland experimental field where 7-HCH had been used once a year for 12 years (Senoo and Wada, 1989; Wada et al., 1989). This organism also degraded 7-HCH through dehydrochlorination (Imai et al., 1989; Yabuchi et al., 1990). Sahu et al. (1990) isolated a fourth aerobe, a P s e u d o m o n a s sp., from the rhizosphere of HCH-

MICROBIALDEGRADATIONOF THE PESTICIDELINDANE CI

CI

CI~ CI

281

C! C[

13-Hexachlorocyciohexane ~l Dihalo-climination CI

C1

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8-Tetrachlorocyclohexane ~l Dilmlo-elilnilLation

Dchydr°halV°gcna I CI

Chlorobenzene

~ CI

__

alo-climination

CI

Dichlorocyclohexadiene

© Benzene

FIG.3. Biotransformationof [3-hexachlorocyclohexaneunder methanogeniccondi-

tions.

treated sugarcane plants. This strain degraded not only 7-HCH but also utilized the thermodynamically stable [~-isomer of HCH as a sole source of carbon under aerobic conditions. This was the first report of the isolation of an aerobe that can metabolize ~-HCH. Subsequently, Nagata et al. (1993b) identified 7-PCCH, 1,2,4-TCB, 2,5-DCP, 2,5-DDOL, and 2,5-DCHQ as metabolites of 7-HCH in Sphingomonas paucimobilis UT26 (Fig. 4).

B.K. SINGH et

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2,5-DCHQ

CO2 FIG. 4. Biodegradation of 7-hexachlorocyclohexane. Bold arrows show aerobic degradation.

('I

MICROBIAL DEGRADATION OF THE PESTICIDE LINDANE

283

Biodegradation of o~- and ~-HCH was undertaken in soil slurries under different redox conditions: aerobic, methanogenic, denitrifying, and sulfate reducing. The aerobic conditions proved to be best for microbial transformation of o~-HCH (Bachmann et al., 1998a). The ~HCH was mineralized at an initial rate of 23 mg/kg of soil per day by a soil microbial community under aerobic conditions; however, under methanogenic conditions the mineralization rate was 13 mg/kg of soil per day. No significant degradation was observed under denitrifying and sulfate-reducing conditions in the case of [LHCH. Physical factors such as temperature, auxiliary carbon source, and substrate concentration were found to influence microbial degradation of o~-HCH. The most favorable temperature range for o~-HCH degradation was 20-30°C. The addition of auxiliary organic carbon compounds showed repressive effects on R-HCH mineralization. High concentrations of 1,4-DCB, 1,2,4TCB, and TCB were found to have adverse effects on the transformation of o~-HCH. The proposed pathway of o~-HCH degradation is shown in Figure 5. Johri et al. (1998) reported degradation of o~-, [3-, 7-, and 8-HCH by Sphingomonas paucimobilis. The bacterium degraded o~-HCH after 3 days; with [3- and 7-HCH, and with 8-HCH, respectively, 98 and 56% degradation occurred after 12 and 8 days. PCCH was the primary metabolite during the degradation of all the HCH isomers. The isolation of microbes for aerobic degradation of 7-HCH made it possible to clone catabolic genes responsible for degradation of this compound (Johri et al., 1996). Imai et al. (1991) initially characterized the degradative pathway of lindane via the chemically unstable intermediate 2,4,5-DNOL in R paucimobilis UT26. A genomic library of P. paucimobilis UT26 was constructed in P. putida by using pKS13, a broad host range cosmid vector. One of the clones was further characterized after subcloning the lindane degradative gene. This clone contained a recombinant plasmid (pKSRI), which was actually pKS13 with a 25-kb insert. A 5-kb HindlII fragment from pKSR1 was subcloned into pUC118 (pIMAI). Recombinant E. coli cells containing pIMAI retained the ability to transform lindane to 1,2,4-TCB thereby showing that the 5-kb HindlII fragment was responsible for the degradation of lindane. Deletion analysis of the 5-kb insert of pIMAI revealed that a 5OO-bp fragment contained the region for the activity that converted lindane to 1,2,4-TCB. By nucleotide sequencing, an open reading frame of 465 base pairs was found within the 500-bp region that encodes for 7-HCH

284

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i C02 + H20 FIG.5. Aerobicbiodegradationof ct-hexachlorocyclohexane.

MICROBIAL DEGRADATION OF THE PESTICIDE LINDANE

285

dehydrochlorinase, which is responsible for dehydrochlorination, yielding 1,2,4-TCB from lindane (Imai et al., 1991). In P. paucimobilis UT26, lindane was first transformed by two steps of dehydrochlorination to an intermediate, 1,4-TCDN, and then to 2,5-DDOL by two dehalogenation steps via an intermediate, 2,4,5-DNOL (Imai et al., 1989). A genomic library of P. paucimobilis UT26 was constructed in P. putida PpY101LA in order to clone a gene encoding the enzyme responsible for conversion of 1,4-TCDN and 2,4,5-DNOL (Nagata et al., 1993a). When an 8-kb BgllI fragment from one of the cosmid clones that converted lindane to 2,5-DDOL was subcloned, a 1.1-kb region was found responsible for the activity. An open reading flame of 885 base pairs (LinB) within the region was revealed by nucleotide analysis. The protein product of the LinB gene was shown to be a 32-kDa protein by SDS-PAGE analysis. LinB could degrade 1-chlorodecane, 1-chlorobutane, and 2-chlorobutane, which are otherwise poor substrates for LinA, suggesting that LinB may be a member of broad substrate-specificity haloalkane dehalogenases (Nagata et al., 1993a). Further, Nagata et al. (1994) cloned the/3"nC gene encoding a 2,5-DDOL dehydrogenase that converts 2,5-ODOL to 2,5-DCHQ in P. paucimobilis.

Vl. Fungal Degradation of Hexachiorocyclohexane Until recently, research into pesticide degradation by microorganisms has focused primarily on bacteria, and fewer studies have been performed with fungi. The main reasons are: • bacteria are easy to culture and grow more quickly than fungi, • bacteria are more amenable to genetic manipulation techniques, • bacteria are less likely to form mutant revertants, and • bacteria use pesticides in laboratory conditions as their sole source of carbon. Fungi as future bioremedial agents have several advantages over bacteria such as: • ability to tolerate low pH values, • fewer complex nutritional requirements, • capability to degrade and utilize a wide range of complex substrates such as cellulose, hemicellulose, lignin and pectin,

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• white-rot fungi with a wide range of extracellular enzymes degrade organopollutants nonspecifically in soil or water environments and make the resultant metabolites more vulnerable for further microbial attack, • white-rot fungi degrade various pesticides including lindane by co-metabolism, and • white-rot fungi tolerate higher concentrations of toxic pollutants than bacteria. Various white-rot basidiomycetes have been investigated for their lindane-degrading capabilities: Phanerochaete chrysosporium (Kohler et al., 1988; Mougin et al., 1996), Pleurotus sajor-caju (Arisoy and Kolankaya, 1997), Trametes ( Coriolus) hirsutus, Phanerochaete sordida, and Cyathus bulleri (Singh and Kuhad, 1997, 1998; Singh, 1999). Phanerochaete chrysosporium, a lignin-degrading white-rot fungus, has received considerable attention for its reported ability to degrade and mineralize a wide range of environmental pollutants such as polychlorinated biphenyls (PCBs), polyaromatic hydrocarbons (PAHs), dioxin, 2,4,5-T, 2,4-D, lindane, and DDT (Higson, 1991; Shah et al., 1992; Joshi and Gold, 1993). The extensive biodegradative properties of the white-rot fungi toward numerous environmentally persistent chemicals have been credited to its lignin-degrading system (LDS), which includes mainly extracellular lignin peroxidases (LiPs), manganese-dependent peroxidases (MnPs), and laccase (Higson, 1991; Shah et al., 1992; Barr and Aust, 1994). These enzymes are produced under substrate-limiting growth conditions, and are not induced by the presence of pollutants. However, Kohler eta]. (1988) and Mougin et al. (1996) studied mineralization of DDT and lindane by P. chrysosporium and found that mineralization of these compounds was independent of LDS. Similarly, other workers have postulated that enzyme systems other than LDS might be involved in catalyzing the pollutant degradation reactions as an alternate or complement to LDS (Dhawale et al., 1992; Joshi and Gold, 1993). Some reports also suggest that both intra- and extracellular enzymes may be sequentially involved in the degradation process of organic compounds (Armenante et al., 1994). More recent experiments carried out in our laboratory support a previous study of Mougin et al. (1996), where only lindane was extracted from homogenized mycelium. These observations indicate that at least the initial steps in lindane degradation do not involve an intracellular enzyme system. Based on these studies and earlier reports, we are of the opinion that LDS might not be playing a role in lindane metabolism. The white-rot fungi have been shown to produce different combinations of

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these extracellular enzymes. P. chrysosporium is known to produce several isozymes of LiP and MnP and produce laccase in a liquid culture m e d i u m containing cellulose (Srinivasan et al., 1995). Phlebia brevispora and Ceriporiopsis subvermispora possess MnP and laccase but not LiP activity (Ruttiman et al., 1992), while P. sordida is reported to produce only MnP (Ruttiman et al., 1994). Lindane-degrading strains of Phlebia radiata, Coriolus versicolor, and C. bulleri are unique in their ability to produce all three enzymes (i.e., LiP, MnP, and laccase) (Kuhad et al., 1997). All white-rot fungi studied have been found to degrade lindane, although they have different sets of lignin-degrading systems, thereby suggesting that a complete LDS may not be involved in lindane degradation and some other extracellular enzyme system might be catalyzing degradation of the pesticide. Mougin et al. (1996) studied lindane transformation by P. chrysosporium. It was observed that lindane was partially mineralized (34.5%) by this fungus after 14 days of incubation, and that TCCH, TCCE, and TCCOL were the major intermediates of lindane biodegradation. They suggested that LiPs and MnPs may not be involved in the first step of lindane degradation, and that P450 oxidases could be the active system for pesticide detoxification in R chrysosporium. TCCH and TCCOL were also shown as intermediates in lindane metabolism by P. sordida and T. hirsutus (Singh and Kuhad, 1999). C. bulleri was the most efficient lindane degrader among different white-rot fungi tested (Singh, 1999).

VII. Algal and Cynobacterial Degradation of Hexachlorocyclohexane A range of naturally occurring microorganisms degrades organic pollutants in aquatic environments. Although studies have concentrated mainly on bacteria and fungi, Semple eta]. (1999) reviewed reports of biodegradation of aromatic compounds by microalgae. In addition, Kobayashi and Rittmann (1982) compiled information on algae capable not only of bioaccumulation of pesticides but also of transforming some of these pollutants: Chlorella vulgaris and Chlamydomonas reinhardtii both transform lindane to PCCH under aerobic conditions. Since algae are obligate aerobes, they require oxygen to degrade organic compounds. However, the biochemical mechanisms by which microalgae catabolize these pollutants are still unclear. There is some evidence that algae use dioxygenase systems to oxidize aromatic compounds (Semple eta]., 1999). Catabolism of phenol to catechol was

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confirmed by isolation of product from incubations in which catechol 2,3-dioxygenase was inhibited with 3-chlorocatechol (Semple and Cain, 1996). 3-chlorocatechol had also been shown to suicide-inhibit the enzyme (Bartels et al., 1984). Microbial degradation of 7-HCH has been reported in the cyanobacteria Anabaena and Nostoc (Kurtiz and Wolk, 1995). When the strains of Anabaena sp. PCC 7120 and Nostoc ellipsossorum were supplied with the fcb ABC operon from Arthrobacter globiformis, they acquired the capacity to dechlorinate 4-CBA (Tsoi eta]., 1991). Further genetic manipulation of these nitrogen-fixing strains by addition of the LinA gene enhanced degradation of lindane (Kurtiz and Wolk, 1995). Kurtiz et al. (1997) reported that dechlorination of lindane by Anabaena sp. PCC 7120 was dependent on the function of the nir operon, encoding enzymes for nitrate utilization. VIII. Future Prospects Long-term persistence and excessive use of lindane has heavily contaminated the terrestrial and aquatic environments. Bioremediation has promise for cleaning up polluted habitats. However, the use of microorganisms for bioremediation requires a thorough understanding of the physiological and biochemical parameters involved in pollutant transformation. Both microorganisms and cell-free enzymes have been studied for degradation of these xenobiotics by various researchers (Lal et a]., 1986; Crecchio eta]., 1995). So far, only a few microorganisms are known to degrade lindane under laboratory conditions. The biochemical mechanisms of lindane degradation, the enzymes catalyzing such reactions, and the genes encoding these enzymes are still poorly defined. However, manipulation of catabolic genes has provided some basic understanding of lindane metabolism by bacteria. Recently, white-rot fungi have attracted considerable attention as potential bioremedial agents for HCH contamination. Lindane degradation by other fungi has also been reported. While the enzymes catalyzing lindane transformation by white-rot fungi have not yet been characterized, further research may lead to isolation of new strains with better abilities to degrade or detoxify HCH as well as to identify enzymes catalyzing different steps in HCH metabolism. The "real-world" application of microorganisms and their enzymes has been limited for several reasons. Genetically engineered microorganisms (GEMs) can be used to bring about rapid degradation of pesticide residues by adding large amounts of prepared inoculum. However, various arguments have

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been put forward as to w h y GEMs may not produce desirable effects (Johri et al., 1996). The major arguments state that: (1) most GEMs w o u l d be unable to compete with indigenous microorganisms and so w o u l d not survive long enough to remove the toxic chemicals, and (2) GEMs w o u l d have no incentive to attack the toxic chemicals because of the presence of large amounts of other and more easily available nutrient sources. Environmentalists are concerned about safety when assessing the effects of releasing GEMs into the environment. The introduction of foreign genes into wild-type microbes not only results in genotype alterations but can also produce unanticipated changes in phenotypic expression of the recipient organism, which, in turn, can result in unanticipated environmental impacts (Doyle et al., 1995). The effects that GEMs may have on the ecosystem could also be affected by their mode of introduction, their spatial and temporal distribution, and the physicochemical and biological characteristics of the environment to which they are released (Yin and Stotzky, 1997). As our knowledge of the pathways of 7-HCH degradation becomes more precise, and as our capabilities for constructing effective recombinant organisms capable of persistence in adverse conditions increases, it would become possible to more effectively remediate environmental pollution by 7HCH more effectively. IX. Nomenclature

2,4,5-DNOL 2,5-DCHQ 2,5-DDOL 4-CBA 5-CHQ BHC DBP DCB DCP HCH LDS PCCH PCPA PNP TCB TCCH TCDN TCP

2,4,5-dichloro-2,5-cyclohexadiene-l,4-diol 2,5-dichlorohydroquinone 2,5-dichloro-2,5-cyclohexadiene-l,4-diol 4-chlorobenzoic acid 5-chloro-2-hydroxy-l,4-benzoquinone Benzene hexachloride Dichlorobenzophenone Dichlorobenzene Dichlorophenol Hexachlorocyclohexane Lignin-degrading system Pentachlorocyclohexane p-chlorophenoxyacetic acid p-nitrophenol Trichlorobenzene Tetrachlorocyclohexane Tetrachlorocyclohexadiene Trichlorophenot

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Microbial Production of Oligosaccharides: A Review S. G. PRAPULLA, ~

SUBHAPRADA, AND

N. G.

KARANTH

Fermentation Technology and Bioengineering Department Central Food Technological Research Institute Mysore 570013, India

I. Introduction A. Classification of Oligosaccharides B. Physicochemical and Functional Properties C. Health Benefits II. Enzymatic Mechanisms of Oligosaccharide Synthesis A. Synthesis of Oligosaccharides by Glycosidases in Reverse B. Synthesis of Oligosaccharides Using Glycosyltransferases III. Microbial Production of Oligosaccharides A. Fructooligosaccharides B. Galactooligosaccharides C. Isomaltooligosaccharides D. Novel Oligosaccharides E. Purification of Oligosaccharides IV. New Approaches to Microbial Oligosaccharide Synthesis A. Use of Organic Solvents B. Use of Aqueous Two-Phase Systems C. Use of Immobilized Systems D. Use of Lipid-Coated Glycosidases E. Use of Recombinants V. Assays and Structural Determination of Oligosaccharides A. High-Performance Liquid Chromatography B. Paper Ghromatography C. Gas-Liquid Chromatography D. Thin-Layer Chromatography E. 13C-NMR Analysis F. FAB Mass Spectrometry VI. Applications of Oligosaccharides A. Nonfood Applications B. Food Applications VII. Conclusions References

I. Introduction

Oligosaccharides are relatively new functional food components that are rapidly becoming popular. They have great potential for improving the quality of the flavor and physicochemical properties of foods. In 299 ADVANCESINAPPLIEDMICROBIOLOGY,VOLUME47 Copyright© 2000by AcademicPress All rights of reproductionin any formreserved. 0065-2164/00 $25.00

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addition, they have the advantages of being low in calories, noncariogenic, and bifidogenic. The increasing consumer demand for healthy and low-calorie foods manufactured using natural compounds is spurring interest in oligosaccharides. World production (excluding lactulose) was estimated at 35,000 tons in 1990 and 65,000 tons in 1995 (Crittenden and Playne, 1996). The oligosaccharides being studied for use in the food industry include the fructooligosaccharides, the galactooligosaccharides, and the isomaltooligosaccharides. The microbial fructooligosaccharides have attracted special attention because their sweet taste is very similar to that of sucrose. Oligosaccharides are classically defined as glycosides that contain between 3 and 10 sugar moieties. Food-grade oligosaccharides also include functionally similar disaccharides like lactulose. Oligosaccharides prepared for use in the food industry are not pure products, but are mixtures containing oligosaccharides with different degrees of oligomerization: the parent disaccharide and the monomer sugar. Various types of oligosaccharides are found as natural components in many common foods, including fruits, vegetables, milk, and honey. Food-grade oligosaccharides are subdivided into 12 groups (Crittenden and Playne, 1996). A. CLASSIFICATIONOF OLIGOSACCHARIDES 1. Galactooligosaccharides

Galactooligosaccharides are a-D-Glu(1-->4)-[13-D-Gal(1-~6)]n, where n = 2-5. They are produced commercially from lactose using the galactosyltransferase activity of 13-galactosidase, which dominates lactose hydrolysis at high lactose concentrations. They are used as prebiotic food ingredients since they enhance growth of bifidobacteria in the intestines. 2. Lactulose

Lactulose is ]]-D-Gal(1--->4)-[3-D-Fru. It is produced by an alkaline isomerization process that converts the glucose moiety in lactose to a fructose residue. Lactulose is not digested by humans and promotes preferential growth of bifidobacteria. It is therefore used as a prebiotic food ingredient and a low-calorie sweetener. In addition, it is widely used as a pharmaceutical for control of constipation and portosystemic encephalopathy.

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3. Lactosucrose Lactosucrose is [3-D-Gal(1--~4)-C~-D-Glu-(1-~2)-~-D-Fru. This trisaccharide is manufactured from a mixture of lactose and sucrose, using the transfructosylation activity of the enzyme ~-fructofuranosidase. This trisaccharide is incorporated in foods as a bifidogenic ingredient.

4. Fructooligosaccharides Fructooligosaccharides (FOSs) are ~-D-Glu(1-~2)-[[3-D-Fru(1--~2] n, where n = 2--4, ~-o-Fru(1--~2)-[[~-D-Fru(1--)2]n, where n = 1-9, or ~-D-Glu(1--+2)-[[~-D-Fru(1--)2],,, where n = 2-9. They are manufactured by two different processes, which result in slightly different end-products: (1) from the disaccharide sucrose using the transfructosylation activity of the enzyme [~-fructofuranosidase, and (2) by controlled enzymatic hydrolysis of inulin with inulinase. The fructooligosaccharides manufactured by each of these methods have their own applications: while the transfructosylation fructooligosaccharides find use as prebiotic ingredients, the longer-chain oligosaccharides derived from inulin are used as fat replacers.

5. Palatinose or Isomaltulose Oligosaccharides Palatinose or isomaltulose oligosaccharides are (~-o-Glu(1--,6)-D-Fru),, where n = 2-4. They are produced from sucrose using an immobilized isomaltulose synthase. The palatinose oligosaccharides so made are digested in the small intestine and thus cannot act as prebiotic agents. They are important as a low-cariogenic sweetener. Palatinose oligosaccharides are also formed by intermolecular dehydration of palatinose. These are not digestible and function as bifidogenic factors.

6. Glycosyl Sucrose (Coupling Sugar) Glycosyl sucrose is (C~-D-GIu(1--~4)-Ot-D-Glu(1-~2)-~-D-Fru. It is manufactured from the disaccharides maltose and sucrose using the enzyme cyclomaltodextrin glucanotransferase. Glycosyl sucrose is used as a substitute sweetener (it is approximately half as sweet as sucrose) and considerably reduces dental caries. Glycosyl sucrose is not bifidogenic. It is largely utilized in the food industry to suppress crystal formation, the browning reaction, and retrogradation of starch.

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7. Ma]too]igosaccharides Maltooligosaccharides are [a-D-Glu(1-o4] n, where n -- 2-7. Commercially, they are produced from starch by the action of debranching enzymes such as pullulanase and isoamylase, combined with hydrolysis by various a-amylases. These a-amylases have differing reaction specificities and produce maltooligosaccharides of different chain lengths. Consumption of maltooligosaccharides is found to improve colonic conditions by reducing the levels of intestinal putrefactive bacteria such as C. perfringens and members of the family Enterobacteriaceae. However, they are not known to be bifidogenic.

8. Isomaltooligosaccharides Isomaltooligosaccharides are [a-D-Glu(1-~6]n, where n -- 2-5. They are produced using a combination of enzymes in two stages. In the first stage, the starch is liquefied by a-amylase; in the second stage, the liquefied starch is hydrolyzed to maltose, which is then converted to isomaltooligosaccharides by the transglucosidase activity of a-glycosidase. Isomaltooligosacharides are bifidogenic and can thus be included in food as prebiotic ingredients.

9. Cyclodextrins Cyclodextrins are cyclic: [a-D-Glu(1-~4]n, where n -- 6-12. They are formed from starch digestion by the action of cyclomaltodextrin glucanotransferase. These oligosaccharides are capable of forming inclusion complexes with various organic compounds and lead to desirable changes in the physical and chemical properties of the incorporated compound. They are, however, not bifidogenic. 10. Gentiooligosaccharides Gentiooligosaccharides are [[3-D-Glu(1-~6]n, where n -- 2-5. They are produced from glucose syrup by enzymatic transglucosylation. Gentiooligosaccharides are found to promote the growth of bifidobacteria and lactobacilli (Nakakuki, 1993).

11. Soybean Oligosaccharides Soybean oligosaccharides are [a-D-Gal(1-~6)]n-a-D-Glu(1-~2)-[~-D-Fru, where n -- 1-2. They are extracted directly from the raw material, without enzymatic processes, by removal of protein and salts. They

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include raffinose and stachyose and are not digestible but reach the colon intact, thereby acting as prebiotics. 12.

Xylooligosaccharides

Xylooligosaccharides are [[3-Xy1(1-~4]n, where n = 2-9. They are manufactured by controlled hydrolysis of xylan, with the enzyme endo-l,4[3-xylanase. These oligoasaccharides promote the growth of bifidobacteria in the colon and are used predominantly in prebiotic drinks. B. PHYSICOCHEMICALAND FUNCTIONALPROPERTIES The physicochemical and physiological properties of oligosaccharides vary with the type of mixture. Chemically, oligosaccharides consist of monosaccharide residues, linked by oxygen bridges between the hemiacetal hydroxyl of the anomeric carbon of one residue and an alcoholic hydroxyl of another residue. Although many studies are being carried out on food-grade oligosaccharides, less information is available with regard to their physicochemical properties. Gross (1962) reported the properties of such kestosides as 1-kestose, 6-kestose, and neokestose. 1-kestose can easily be crystallized as white needles. The crystals of 1-kestose exhibit a specific rotation of [ct]~° + 28.5 °, and the melting temperature of 1-kestose ranges between 199 and 200°C. It is nonreducing to Fehling solution. The sweetness values of the ffuctooligosaccharides as compared to 10% sucrose are 31, 22, and 16% for 1-kestose, nystose, and l~'-fructofuranosylnystose, respectively. They are highly hygroscopic. The viscosity of a fructooligosaccharide solution is relatively higher than that of sucrose at the same concentration. Fructooligosaccharides have higher thermal stability than sucrose. These are highly stable at the normal pH range of foods (pH 4.0-7.0) and are stable at refrigerated conditions for over a year. They are reported to resemble sucrose in many properties, including solubility, freezing and boiling point, and crystal data (Yun and Song, 1996). Incorporation of oligosaccharides enhances the physiological and rheological properties of foods. This is primarily due to their specific properties: • They improve "mouth feel" since they have a higher molecular weight than the mono- and disaccharides. • They are used as bulking agents when it is desirable to have relatively low sweetness, to enhance other food flavors, and to alter the freezing temperature of frozen foods.

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• They are used to control the amount of browning due to Maillard reactions in heat-processed foods. • They help control microbial contamination, since they have a high moisture-retaining capacity with low water activity and also prevent excessive drying. • They act as strong inhibitors of starch retrogradation. • They are widely used in beverages, infant milk powders, confections, bakery products, and yoghurt and dairy desserts. C. HEALTH BENEFITS

Among the greatest advantages of oligosaccharides are the health benefits they impart. Many of them are not digestible in the intestine and therefore are noncalorific. Another major advantage is that, being nonfermentable they are noncariogenic, that is, they prevent dental caries caused by oral flora, mainly Streptococcus mutans. The primary health benefit bestowed by oligosaccharides is stimulation of beneficial bacteria in the colon, especially Bifi'dobacterium spp. The adult human gastrointestinal tract is a complex balanced ecosystem consisting of many species of bifidobacteria and lactobacilli, most being advantageous or benign to a healthy individual. Disturbances in the ecological balance caused by changes in diet, stress, or antibiotic treatment can lead to overgrowth of deleterious bacteria, leading to gastrointestinal disorders ranging from mild discomfort and flatulence to such more serious conditions as colitis, diarrhea, and irritable bowel syndrome. The increase in incidence of disease with age is postulated to be due to a decrease in intestinal bifidobacteria in old age, resulting from diminished secretion of gastrointestinal juices. The toxic metabolites formed during fermentation of food in the colon--for example, nitrosamines, indoles, and skatoles (carcinogens), and phenols and cresols (cancer-promoting agents)--have an adverse health effect. Putrefactive intestinal bacteria are known to promote cancer, aside from being mutagenic and hepatotoxic. A plausible way to reduce this toxicity is to suppress the growth of putrefactive microflora, allowing enhanced growth of indigenous bifidobacteria. Bifidobacteria are believed to be beneficial to the host, as they assist in digestion and absorption, produce vitamins, prevent growth of putrefactive pathogenic bacteria, and stimulate the immune response. The effective bifidogenic dose of oligosaccharides is generally less than 15 g/day (Tomomatsu, 1994).

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Commercial manufacture of food-grade oligosaccharides involves the use of enzymatic processes, with the exception of soybean oligosaccharides (produced by direct extraction) and lactulose (produced by an alkali-catalyzed reaction). They are manufactured by hydrolysis of such polysaccharides as starch, inulin, and xylan; however, the buildup of oligosaccharides from simple sugars like sucrose or lactose by enzymatic processes allows synthesis of controlled, "tailor-made" oligosaccharides. Depending on the enzyme source, oligosaccharides with different linkages are formed. For example, fructosyltransferase derived from Aspergi]lus niger produces only 1-kestose (1F-~-fructosyl sucrose), while enzymes of Claviceps purpurea and asparagus produce both 1-kestose and neokestose (6c-[~-fructosyl sucrose). Though, in general, mass production of oligosaccharides is not complicated, the use of microbial enzymes is preferred over plant enzymes because the yields of oligosaccharides using enzymes from plant sources is low and mass production of enzymes is limited by seasonal conditions. The use of microbial enzymes is therefore an effective alternative to chemical synthesis. The industrial production of microbial oligosaccharides is a field that offers tremendous scope and opportunity. It is continuing to assume greater importance, and there have been some successful ventures: for example, microbial FOS production with fructosyltransferase from Aspergiflus or Aeureobasidium pullulans (Barthomeuf and Pourrat, 1995). Meiji Seika (Japan) was the first company to meet success with commercial FOS production (brand name Neosugar®). More recently, the M/s Cheils Foods and Chemical Company (Korea) has been successful with FOS production using immobilized cells of A. pullulans (Yun et al., 1992).

II. Enzymatic Mechanisms of Oligosaccharide Synthesis Oligosaccharides possess great structural diversity and have a wide range of roles within the cell. Commercial and chemical synthesis and synthetic strategies have been well developed (Garegg, 1990). However, the protocols are complicated and give very poor yields. For these reasons, there is a great deal of interest in developing methodologies for enzymatic synthesis of these oligosaccharides (Toone et al., 1989; Bucke and Rastal, 1990; Ichikawa eta]., 1992). The suggested approaches for oligosaccharide synthesis using glycosidases are indicated in Figure 1. Two main classes of enzymes

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s . G . PRAPULLA et al. (a) EQUILIBRIUM APPROACH

Mannose

b~

~

Mannose,2

a-mannosidase (60-80%, w/w sugar 60°C)

(b) KINETIC APPROACH

p-Nitrophenyl-ct-mannoside ct-mannosidase ct-marmosyl-Enzyme+ p-Nitrophenol complex

L

~l~

p-Nitrophenyl-13-glucoside

Man - tx (l-x) - 13GIc - P NPH + a-Mannosidase

FIG. 1. Synthesis of oligosaccharides using glycosidases. Reprinted with permission from Rastall and Bucke (1992).

currently being used for in-vitro synthesis of oligosaccharides are glycosidases and glycosyltransferases. Glycosidases have many advantages over the sucrose-dependent glycosyltransferases and are therefore more widely used. A.

SYNTHESIS OF OLIGOSACCHARIDES BY GLYCOSIDASES IN REVERSE

Glycosidases have been used sporadically as synthetic reagents since the late nineteenth century (Croft-Hill, 1898). There has been a remarkable growth in the understanding of the mechanisms involved. Bucke and Rastall (1990) described two that allow glycosidase reversal: the equilibrium approach and the kinetic approach (Fig. 1).

MICROBIAL PRODUCTION OF OLIGOSACCHARIDES

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The equilibriumapproach is based on the fact that all enzyme reactions are reversible. The reversal of the normal glycoside hydrolysis reaction is achieved by increasing the concentration of the products of the forward reaction and reducing the concentration of reactants. This is done by incubating the enzyme in a highly concentrated solution of monosaccharides. Johannson et al. (1989) and Rastall et al. (1992) reported that sugar concentrations of the order of 70-80% (w/w) are needed to reduce aw to the point where the reverse reaction becomes dominant, but at such concentrations the reaction rates are slow. The temperature of the reaction is therefore increased to 50-60°C to allow equilibrium to be reached within a reasonable time period. The enzyme is protected from denaturation by the stabilizing effects of high sugar and low water concentrations (Rastall and Bucke, 1992). The kinetic approach takes advantage of the fact that the hydrolysis of glycosidic bonds proceeds through a two-stage process involving a covalently linked sugar-enzyme intermediate (Viratelle et al., 1969; Stokes and Wilson, 1972). This covalent bond is broken by a nucleophilic displacement reaction involving water as an acceptor molecule; however, other compounds such as alcohols (Viratelle et al., 1969; Shiffin and Hunn, 1969) and other carbohydrates (Nilsson, 1987) can act as acceptors. For the synthesis of an oligosaccharide, a reaction mixture is prepared containing a reactive donor substrate such as a p-nitrophenyl (p-PNP) glycoside, which promotes rapid glycosylation of the enzyme, and a glycoside acceptor. Manipulation of the nature of the glycosidic bond formed is possible to a certain extent by careful selection of the aglycone (Nilsson, 1986). Glycosidases are generally nonspecific for acceptor sugars, so heterooligosaccharides can be synthesized by co-condensation of two monosaccharides or a monosaccharide and an oligosaccharide. The production of heterooligosaccharide can be promoted over the production of homooligosaccharides by increasing the percentage of acceptor sugar in the mixture, but this leads to a decrease in the total yield of oligosaccharide products (Rastall and Bucke, 1992). The absolute concentration of the sugar mixture is critical and depends on the nature of the acceptor. B. SYNTHESIS OF OLIGOSACCHARIDES USING GLYCOSYLTRANSFERASES

Many glycosyltransferases have been purified and applied to in-vitro synthesis of oligosaccharides (Toone et al., 1989; Ichikawa et al., 1992). Galactosyltransferase is the best characterized of all the glycosyltransferases. Other glycosyltranferases used are fucosyltransferase (Rosevear

308

s.G. PRAPULLA et al.

eta]., 1980; Palcic et al., 1987) and glycosyltransferase (Wiezemann et al., 1989). The first stage in the synthesis, as described by Rastall and Bucke (1992), involves the formation of sugarol-phosphates by a kinase enzyme followed by transfer of the sugar residue to a nucleoside triphosphate by a nucleoside transferase. The second stage involves transfer of sugar units from the sugar nucleotides to sugar acceptors, catalyzed by glycosyltransferases (Fig. 2). The primary advantage of glycosyltransferases is their great specificity, allowing for precise and sequential construction of oligosaccharides. However, this requires the presence of a very wide range of enzymes; thus, the availability of purified enzymes is a major limiting factor (Rastall and Bucke, 1992). Another major problem with the use of glycosyltransferases as synthetic reagents is that these enzymes are generally unstable in solution (Wong et al., 1982). Currently, glycosidases are preferred for commercial use as they are easily purified, stable, and not required in completely pure form (Rastall and Bucke, 1992). The disadvantage of these enzymes is that regulation of the proportions of the various oligosaccharides in the final product is difficult.

III. Microbial Production of Oligosaccharides Conventionally, chemical methods have been used for the synthesis of oligosaccharides. These methods have many disadvantages, though. They are laborious, require continuous and rigid monitoring, and require the use of pure and often hazardous chemicals. Moreover, they are expensive and give low yields.

Sugar

Kinase

Sugar-l-p

+ NTP nucleoside transferase

NDP - Sugar + PPi

glycosyl transferase Sugar acceptor molecule OLIGOSACCHARIDE

FIG, 2. Synthesis of oligosaccharides using glycosyltransferase. mission from Rastall and Bucke (1992).

Reprinted with per-

MICROBIALPRODUCTIONOF OLIGOSACCHARIDES

309

These disadvantages become evident, even with the synthesis of an extremely simple disaccharide such as sucrose. Sucrose has an ~-l-glucose-[3-2-frnctose linkage. To ensure a linkage between only the - - O H 1 of the glucose and the - - O H 2 of the fructose, it is essential to block all the other hydroxyl groups on the two monosaccharides by acetylation, and so on. After formation of the disaccharide, deblocking of these groups has to be done. Chromatographic separation of the ~ and [3 forms of the newly formed product is also often required. Microbial methods for production of oligosaccharides provide costeffective and convenient alternatives to chemical synthesis. A. FRUCTOOLIGOSACCHARIDES Industrially important fructooligosaccharides consist mainly of 1-kestose, nystose, and fructofuranosylnystose, in which one to three fructosyl units are bound to the [~-2,1 position of the sucrose. FOSs are rapidly gaining importance and are being widely studied because of their specific physiological effects on humans, especially growth, stimulation of beneficial bifidobacteria in the digestive tract, decrease of total cholesterol and lipids, constipation relief, and generally improved health. They are also used as stabilizers and/or bulking agents and for production of sweeteners (Barthomeuf and Pourrat, 1995). Chemical FOS synthesis has not been practical because of the laborious nature of conventional methods, the use of hazardous/expensive chemicals, the multiple reaction steps, and the resultant low yields (Rastall and Bucke, 1992). Microbial production is therefore becoming increasingly attractive, fructooligosaccharides are produced by transfer of fructose residues to sucrose molecules by the action of [3-fructofuranosidase (E.C. 3.2.1.26) or [~-fructosyltransferase (FTF) (E.C. 2.4.1.9). The enzymes involved could either be intracellular or extracellular.

1. Production of 1-Kestose by Aspergillus phoenicis 1-kestose is a noncaloric, noncariogenic fructooligosaccharide and is used as a bulking agent in combination with intense sweeteners. Balken et al. (1991) carried out studies on the enzymatic production of 1kestose by sucrose-lF-fructosyltransferase (SFT) (E.C. 2.4.1.99) from Aspergillus phoenicis CBS 294.80. A. phoenicis CBS 294.80 has been used for production of fructose from inulin. This organism, which is known to produce a thermostable inulinase, is also reported to produce SFT (E.C. 2.4.199) (Balken et al., 1991). The organism can be grown in a m e d i u m (pH 6.8) composed of FeS04 • 7H20 (0.1 g/liter), MnS04 • 4H20 (0.1 g/liter), K2HPO4 (5.0 g/liter), NaC1 (1.0 g/liter), yeast extract (3 g/liter), MgSO4.7H20 (0.2 g/liter), sucrose (100 g/liter), and agar (20

310

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g/liter) for 8 days at 28°C. This is solely used for the production of spores. The fermentation medium, which was the same as the precultivation m e d i u m (without the agar), was inoculated with 108 spores per liter and incubated with 200-rpm agitation at 30°C. At the end of fermentation, mycelia were separated and whole cells were used, after a single wash with 0.15 MNaC1, as the SFT source. By using 2191 m M sucrose in a 0.1 m M potassium phosphate buffer at pH 8.0, using 0.18 g dry mycelium per 100 ml reaction volume at 55°C, they could obtain 644 m M (-325 g/liter) 1-kestose on incubation for 12 hours, corresponding to a yield of 60% (w/w) FOS from sucrose (Balken et al., 1991).

2. Production by Aspergillus japonicus A. japonicus TIT-90076 was reported by Chen and Liu (1996) to produce a high-transfructosylating ~-fructofuranosidase. The organism was grown on potato dextrose agar at 30°C, and suspensions of about 4 x 107 spores/ml were prepared. A 1-ml aliquot of this suspension was inoculated into 100 ml of culture m e d i u m consisting of NaNO3 (20 g/liter), MgSO4 • 7H20 (0.5 g/liter), K2HPO4 (0.5 g/liter) with carbon and nitrogen sources. Fermentation was carried out for 96 hours at 30°C with aeration. Homogenized cell suspension was used as the source of enzyme. The transfructosylating and hydrolytic activities of the homogenized cell suspension were estimated by measuring the released glucose (G) with a glucose analyzer and reducing sugars (R) by the DNSA method. Chen and Liu (1996) determined the optimum pH for maximum transfructosylating activity and m i n i m u m hydrolytic activity to be between pH 5.0 and 6.0. The enzyme had almost no transfructosylating activity below pH 3.0 and above pH 10.0. The temperature optimum was reported to be between 55 and 65°C. Sucrose was the best substrate, as it possesses a ~-1,2 linkage that can be cleaved by the [3-fructofuranosidase and acts as an inducer for enzyme production by A. japonicus TIT-90076. Optimal enzyme production was reported at 25 % sucrose concentration. The study by Chen and Liu (1996) details the optimization of cultural conditions for m a x i m u m enzyme production. They have carried out a detailed investigation on the effect of different carbon sources, nitrogen sources, inorganic salts, etc., on enzyme production. Their study also includes the effect of surfactants and polymer additives on the morphology of mold and enzyme production. 3. Production by Aspergillus niger A. niger ATCC 20611 was reported by Hidaka et al. (1988) to be the most suitable strain for FOS production, since the strain showed very

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high enzyme productivity. Its transfructosylating activity was found to be stronger compared to its hydrolyzing activity. The culture was inoculated into a fermentation broth containing 5% sucrose, 3.5% yeast extract, and 0.5% carboxymethyl cellulose at pH 6.0 and incubated at 28°C. FOSs were produced by incubating a mixture of 50% (w/v) sucrose and cells of A. niger ATCC 20611 (to get 2.5 units of transfructosylating activity per gram of sucrose) in a 0.04 M buffer, pH 5.0, at 40°C for 72 hours with agitation. At the end of 8 hours, there was production primarily of 1-kestose, while at the end of 72 hours a significant amount of nystose was formed.

4. Production by Fusarium oxysporum F. oxysporum 172464, 141117, and 141110 (International Mycological Institute, U.K.) were studied for their fructosyltransferase production by Patel et al. (1994). The selected fungus was cultivated in modified Czapek's m e d i u m at an initial pH of 5.5. FOS production in 8 hr with a concomitant increase in fructosyltransferase activity was reported. After 48 hr, invertase activity increased and the FOS concentration in the m e d i u m decreased as it was utilized to meet the energy demands for growth. 5. Production of 1-Kestose by Scopulariopsis brevicaulis S. brevicaulis N-01, a fungus, was isolated from soil by Takeda et al. (1994). They detailed the procedure for obtaining a theoretical yield of 85% of 1-kestose. The amount of 1-kestose produced was 95.6 g/liter with a yield of 64.0%, which corresponded to a theoretical yield of 85.0% (Fig. 3). In contrast to S. brevicaulis, A. niger was reported to produce only 24% 1-kestose, with significant amounts of nystose (23%) and fructofuranosyl nystose (15%) (Hidaka et al., 1988). 6. Production by Penicillium frequentans Usami et al. (1991) reported the production of 13-fructofuranosidase showing fructose-transferring activity by P. frequentans WU-1S. An inoculum of conidia was used at a concentration of 106 per milliliter, and fermentation was carried out at 30°C with a maximum transfructosylation activity of 5.40/ml. 7. Production by Penicillium rugulosum Barthomeuf and Pourrat (1995) reported production of high-content FOSs from sucrose by a crude fructosyltransferase of P. rugulosum. The enzyme was obtained by growing the fungus P. rugulosum for 3 days at 29-30°C in Czapek's m e d i u m containing sucrose (3)%, NH4C1 (1%),

312

s . G . PRAPULLA et al.

- - e - Sucrose

- - e - 1 - Kestose

- - e - Growth

160 30

140

120

LU CO 100

20

0

I-C/) LU v

"v

8O

O o"

uJ

~ 60 10

CO

40

20

~ 0

20

40

60

0 80

T I M E (h)

FIG. 3. Production of 1-kestose by S. brevicaulis under optimal conditions. Reprinted with permission from Takeda et al. (1994).

and soya peptone (0.75%). They used an inoculum of about 108 spores per liter and maintained pH at 5.5. Barthomeuf and Pourrat demonstrated that the crude enzyme (culture filtrate) acts as a mixed-enzyme source of fructosyltransferase (FTF) (Class 2 by Enzyme Nomenclature) and glycosidase (Class 3 by Enzyme Nomenclature). They obtained yields of about 80% in the presence of 750 g/liter sucrose with enzyme concentrations of 5 FTF units per gram of sucrose at 55°C and pH 5.5. Accumulation of high concentrations of nystose (412 g/liter) and fructofuranosyl nystose (76 g/liter) were reported. The major advantage of the crude FTF was reported to be formation of a high FOS concentration within a relatively short time: 650 g/liter in 10 hr as opposed to 363 g/liter in 25 hr as obtained by the mixed-enzyme system of FTF and glucose oxidase, where sucrose concentration has to be limited to 400 g/liter. The reaction is carried out at 40°C to prevent inactivation of

MICROBIALPRODUCTIONOF OLIGOSACCHARIDES

313

glucose oxidase, which oxidizes glucose. The glucose oxidase mixedenzyme system is detailed in a subsequent section.

8. Production by Aureobasidium pullulans Jung et al. (1987) investigated the conditions for production of fructosyltransferase from A. pullulans KFCC 10245. The fermentation medium consisted of sucrose (20%), yeast extract (1%), K2HPO 4 (0.5%), MgSO4 • 7H20 (0.05%), and NaNO 3 (1%), adjusted to pH 6.5. The cells were cultivated for 4 days at 28°C and 180 rpm. They reported production of both intra- and extracellular enzyme; the production of both increased with sucrose concentration (Fig. 4). They demonstrated that both 0.5% phosphate and 2% sodium nitrate had a positive effect on enzyme production. They were also able to increase intracellular enzyme production to 140% by increasing the concentration of magne-

120

100

90 100

80 70

o~ 8O >b-

60

f--

tO ,<

60

50

32

rn

LU

>

t.--

w n,

40

t.U

tO

>rY

40

30

~

20

20

10

0

0 0

5

10

15

20

25

30

35

40

45

SUCROSE (%)

FIG. 4, Effect of sucrose concentration on e n z y m e production. • = intracellular enzyme activity; [3 extracellular e n z y m e activity; • dry cell weight. Reprinted w i t h permission from Jung e t al. (1987).

314

s.G. PRAPULLAet al.

sium sulfate in the culture medium from 0.05 to 0.2%. FOS production using A. pullulans entrapped in calcium alginate gel was reported by Yun et al. (1992). The fungus was cultivated at 30°C for 6 hr in a medium similar to that described by Jung et al. (1987). Immobilization of cells increased stability to 30 months at 50°C. Higher enzyme activity was reported if the culture was cultivated in a medium containing 55% (w/w) of Mg2÷. Hayashi et al. (1991) reported immobilization of a free fructosyl-transferring enzyme from Aureobasidium ATCC 20524 on Shirasu (volcanic ash) porous glass. 9. Production by Arthrobacter spp.

Fujita et al. (1990) described FOS production through the transfructosylating activity of ~-fructofuranosidase ([3-FFase) (E.C. 3.2.1.26) of Arthrobacter sp. K1. They cultivated the strain for 2 days at 30°C and 110 rpm in a medium composed of yeast extract (1.2%), polypeptone (0.8%), lactose (4.0%), (NH4)2HPO4 (0.4%), and MgSO 4 • 7H20 (0.1%), adjusted to pH 7.0, primarily for inoculum buildup. The medium for FOS production consisted of CSL (5.0%), sucrose (3.0%), (NH4)2HPO4 (0.4%), and MgSO4 • 7H20 (0.1%) at pH 7.0. Fermentation was carried out at 37°C for 25 hr under aeration at 6 liter/min. Fujita et al. (1994) also reported production of novel fructooligosaccharides by using different monosaccharides and disaccharides as acceptors of the fructosyl residue. They carried out all reactions at 40°C and pH 6.5 for 5-20 hours using a reaction mixture of sucrose, acceptor, and [3-FFase in 50 mM phosphate buffer. The main transition product to most reducing mono- and disaccharides was a nonreducing oligosaccharide with a fructosyl residue linked to the hemiacetal hydroxyl group, for example, 2-O-[3-D-fructofaronsyl-~-L-sorbo pyzanoside with L-sorbose as the acceptor. Formation of reducing oligosaccharides was reported in addition to nonreducing oligosaccharides when D-galactose or L-arabinose was used as an acceptor, for example, 3-O-[~-D-fructofuranosyl-D-galactopyranose and 4-O-~-D-fructofuranosyl-L-arabinopyranose, respectively. 10. Production Using Mixed-Enzyme Systems There have been reports on the use of mixed-enzyme systems for FOS production apart from many reports with isolated single-enzyme systems. Cultures of A. niger AN10a were reported by Novak et al. (1996) to produce FOSs simultaneously with gluconate. The culture produces

MICROBIALPRODUCTIONOF OLIGOSACCHARIDES

315

an active invertase that cleaves sucrose to form glucose and fructose; the glucose is converted to gluconate while fructose is transferred through a transfructosylation mechanism to a molecule of sucrose, yielding FOSs. They were able to achieve a maximum yield of 40% of consumed sucrose for 1-kestose and 8% for nystose. These relatively high FOS yields elucidated by simultaneous oxidation of glucose are reported to be due to a decrease in the competitive inhibition of transfructosylation (Jung et al., 1987). Production was carried out by Novak et a]. (1996) in a m e d i u m containing sucrose (250 g/liter), Ca(NO3) 2 • 2H20 (2.0 g/liter), KH2PO 4 (0.25 g/liter), MgSO 4 - 7H20 (0.25 g/liter), KC1 (0.25 g/liter), and FeC13 (0.01 g/liter). Studies on production of high-content fructooligosaccharides from sucrose by the mixed-enzyme system of fructosyltransferase (E.C. 3.2.1.26, A. pullulans KFCC 10524) and glucose oxidase (E.C. 1.1.3.4., A. niger) have also been carried out by Yun and Song (1993). They carried out the mixed-enzyme reaction in a stirred tank reactor containing 40% (w/v) sucrose with 10 units of fructosyltransferase and 10 units of glucose oxidase per gram of sucrose for 25 hours at 40°C and pH 5.5. Highly concentrated fructooligosaccharides (up to 90%) were reported with this system. In another study by the same authors (Yun et al., 1994a), a high FOS content (up to 98%) was reported using the same system with agitation at 550 rpm and an oxygen flow rate of 0.71 liter per minute with 10 units of [3-fructofuranosidase and 15 units of glucose oxidase. A higher content of nystose and only a trace amount of fructofuranosyl nystose were formed with the mixed-enzyme system. The use of glucose isomerase instead of glucose oxidase to eliminate glucose inhibition was studied by Yun and Song (1993), but was found to be impractical due to its equilibrium characteristics. Glucose oxidase is very effective in allowing continuous conversion of fructooligosaccharides by complete removal of glucose. The mixed-enzyme systems could thus be applied to large-scale production of high-content fructooligosaccharides.

B. GALACTOOLIGOSACCHARIDES Galactosides possess great structural diversity and have a wide range of roles. For example: 1. In a living cell, glycolipids and glycoproteins that contain lipid/ protein conjugates with unique galactooligosaccharides are impor-

316

S.G. PRAPULLA et al.

taut as recognition molecules and have been implicated in cell-virus, cell-bacterial, and cell-cell interactions (Rastall and Bucke, 1992). They are also important as bifidus factors (Pivarnik et al., 1995). 2. In the food industry they are used as additives for a variety of functional purposes, including solubilizers, stabilizers, sweeteners, and humectants (Rastall and Bucke, 1992; Pivarnik et al., 1995). Most of these molecules occur in very small amounts in complex biological matrices, making their extraction cumbersome and expensive; as a result, these compounds are not available on a scale large enough to permit extensive research. Chemical strategies for synthesis of galactooligosaccharides do exist but are very complicated, labor-intensive, and costly. Products so manufactured are necessarily expensive, since the yields are low and scale-up is difficult (Rastall and Bucke, 1992). Microbial production of galactooligosaccharides is fast developing into a convenient and economic alternative to chemical syntheses. Galactooligosaccharides are produced by transfer of galactose from glucose (in the lactose molecule) to hydroxyl-containing acceptors (other than water) catalyzed by the enzyme ~-galactosidase (E.C. 3.2.1.32). This enzyme exhibits preference for the formation of a [3(1-46) linkage when sugars are the acceptor molecules. Thus, for example, in the presence of only lactose, allolactose (6-O-[~-D-galactopyranosyl-D-galactose) and galactobiose (~)-D-galactose(1-*6)-D-galactose) are the commonly formed disaccharides (Kinsella and Taylor, 1995). 1. Production from Fungal Sources

A wide range of fungi produce ~-galactosidases that can be used for lactose hydrolysis. Many of them can also be used for production of oligosaccharides by transgalactosylation. a. Production by Aspergillus niger. Wierzbicki and Kosikowski (1973) reported the formation of oligosaccharides by the action of [3-galactosidase of A. niger on 4% lactose, with oligosaccharide formation of only 1-2% of total lactose. They found that oligosaccharide production was influenced by substrate concentration and reaction time. Adachi (1977) identified these oligosaccharides as 2-O-~-D-galactopyranosyl-D-glucose, 3-O-[3-D-galactopyranosyl-D-glucose, 6-O-[~-D-ga-

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lactopyranosyl-D-glucose, 3-O-[~-D-galactopyranosyl-D-galactose, and 6O-[3-D-galactopyranosyl-D-galactose. b. Production by A. niger (Exogalactanase). Bonnin and Thibault (1996) reported that the exo-~-l,4-galactanase of A. niger, which hydrolyzes galactans in exomanner mode, was also able to catalyze transfer of galactobiosyl residues to the such acceptor molecules as alcohols, saccharides, hydroxybenzenes, and glycerol, forming oligosaccharides. The yield of transgalactosylation was shown to be most efficient when the acceptor was used at a 10 mM final concentration, and galactan (1.5% w/v) was the donor with 5 nkat/ml exogalactanase, at pH 7 and 40°C. They further detailed that each incubation parameter influenced the transglycosylation in the same way irrespective of the degree of substrate polymerization, and they obtained a maximum yield of 30.9% when a dimer was used as the substrate. c. Production by Aspergillus oryzae. Lopez-Leiva and Guzman (1995) reported conversion of lactose in whey ultrafiltrate permeates to oligosaccharides when passed through an immobilized enzyme reactor of galactosidase from A. oryzae. They found the optimum reaction time for maximum production of oligosaccharides to be in the range of 5 to 15 seconds, wherein a conversion of 25-45% of initial lactose concentration was reported. Their studies indicate that the highest possible concentration of initial lactose should be chosen for high production of oligosaccharides. d. Production by Trichoderma harzianum. Trichoderma is known for the production of highly cellulolytic enzymes, and it is commonly used for production of many glycosidic enzymes (e.g., glucosidase, xylanase, CMCase). Prakash et al. (1989) reported that the culture also produces [3-galactosidase with high transgalactosylation activity. They successfully produced more than 30% oligosaccharides in a high-lactose growth medium composed of lactose (150 g/liter), cottage cheese whey (spray-dried) (5 g/liter), KH2PO4 (1 g/liter), NH4NO3 (2.5 g/liter), MgSO4 • 7H20 (0.2 g/liter), FeC13.6H20 (0.02 g/liter), CaC12 (0.01 g/liter), and NaC1 (0.01 g/liter) at pH 7.0 and 30°C, with agitation at 100 rpm for 2 weeks. After separation and structure elucidation, they reported that in the total oligosaccharides [3(1~6) linkages were about 90%, and the order of linkage formation was ~(1--~6) > ~(1-~3) > ~(1--~4), although oligosaccharides with the latter two types of linkages were produced in small quantity. High production of O-~-D-Gal(I~6)-[3-DGal(1-~4)-D-Glc is important as it occurs in human milk and is a bifidus factor.

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s . G . P R A P U L L A et al.

2. Production from Bacterial Sources a. Lactic Acid Bacteria. Toba et al. (1982) reported formation of 3-11 types of galactooligosaccharides during lactose hydrolysis by cell-free extracts of 12 strains of Lactobacillus and from Streptococcus thermophilus. Further investigation led to classification of these bacteria into four groups based on their chromatographic patterns (Table I). Toba et al. (1982) carried out investigations on oligosaccharide synthesis in a lactose medium containing lactose (10 g/liter), Difco trypton (20 g/liter), yeast extract (5 g/liter), gelatin (2.5 g/liter), NaC1 (4 g/liter), and L-ascorbic acid (0.5 g/liter). They optimized conditions for maximum transgalactosylation. The optimum conditions were pH 6.5-7.0 and temperature 50-60°C. b. Production by Caldocellum saccharolyticum. The thermophilic bacterium C. saccharolflicum was reported by Stevenson et al. (1996) to produce a 13-galactosidase with significantly greater resistance to inactivation by heat and organic solvent and with better transgalactosylation efficiency than the enzyme from A. oryzae. The A. oryzae enzyme

TABLE I CLASSIFICATIONOF LACTICACID BACTERIABY THE CHROMATOGRAPHICPATTERNOF OLIGOSACCHARIDESFORMED FROM LACTOSEBY TRANSGALACTOSYLATIONREACTION

Group 1 2

3

Organism

Oligosaccharides formed

L. helveticus B-1 L. helveticus LH-17

6-O-[~-D-galactopyranosyl-D-glucose

L. acidophilus L. casei C-9 L. plantarum 118 L. plantarum 128 L. plantarum LP-1

6-O-v-D-galactopyranosyl-D-glucose

L. helveticus Bf-9 L. lactis L-3

6-O-I~-D-galactopyranosyl-D-glucose 3 -O-[3-D-galactopyranosyl-D-glucose

trisaccharide 6-O-[~-D-galactopyranosyl-D-galactose trisaccharide

trisaccharide L. L. L. S.

bulgaricus B-6 bulgaricus B-56 bulgaricus yB-62 thermophi]us 510

6-O-[~-D-galactopyranosyl-D-glucose 6-O-13-D-galactopyranosyl-D-galactose 3 - O-[~-D-galactopyranosyl-D-glucose 2 - O-[~-D-galactopyranosyl-D-glucose

tri- and higher oligosaccharides Reprinted with permission from Toba et al. (1982).

MICROBIAL PRODUCTION OF OLIGOSACCHARIDES

319

gave better oligosaccharide yields at lower lactose concentrations, while at higher concentrations (above 50% w/w) the C. saccharolyticum enzyme was significantly better, yielding a sugar mixture containing 42% by weight of tri- and tetrasaccharides, from a 70% w/w lactose solution, compared with 31% with the A. oryzae enzyme. The main advantage of the C. saccharolyticum enzyme is its thermostability (80°C), allowing dissolution of high concentrations of lactose, thereby maximizing the efficiency of oligosaccharide synthesis (Stevenson et al., 1996). c. Production by Bifidobacterium bifidum. Dumortier et al. (1994) reported that the ~-galactosidase from B. bifidum exhibits specific galactosyltransferase activity, in addition to its hydrolytic activity. While lactose hydrolysis occurred optimally at pH 6.5 and 37°C, transgalactosylation was optimum at pH 4.8 and 45°C. The amount of galactooligosaccharides was 29% of total sugars, with maximum production achieved when 60% of the initial lactose was transformed by t3. bifidum cells after incubation for 80 hours with 500 mM lactose at pH 4.25. Among all the effectors tested, only EDTA (5 mM) increased transgalactosylation activity. 3. Production from Actinomycetes Production by Saccharopolyspora rectivirgula. Nakao et al. (1993) isolated a thermophilic actinomycete (S. rectivirgula) from hay that secreted a thermostable ~-galactosidase with high transgalactosylation activity. An oligosaccharide yield of 41% (w/w) was achieved when the enzyme was reacted with 1.75 M lactose at 70°C and pH 7.0 for 22 hr. This enzyme is suitable for immobilization to allow efficient production of oligosaccharides from lactose at high temperatures (>60°C). 4. Production from Yeasts There have been reports on transgalactosylation activity of [~-galactosidase from some yeasts such as Saccharomyces lactis (Burvall et al., 1979) and Kluyveromyces lactis (Dickson et al., 1978; Pivarnik and Rand, 1991). Yeast lactases, however, produce a lesser number of total transgalactosylase products and a lower portion of trisaccharides when compared to mold-derived lactases (Pivarnik et al., 1995). ~-galactosidase from S. lactis has been used for preparing low-lactose milk under the trade name "Maxilact" (Burvall eta]., 1979).

320

S.G. PRAPULLA et al. C. ISOMALTOOLIGOSACCHARIDES

Together with fructooligosaccharides and several other oligosaccharides, isomaltooligosaccharides have received much attention and have been produced commercially in Japan since the early 1980s. Panose, composed of three glucose units with a(1~4) and a(1~6) glucosidic linkages, is known to prevent dental caries. Incorporation of panose into foods, especially confections, is being widely studied. It has conventionally been produced from starch by cooperative reactions of a starch-degrading enzyme such as a-amylase and a-glucosidase, and debranching enzymes such as pullulanase (Takaku, 1988). Yun et al. (1994b) reported that panose was the major isomaltooligosaccharide selectively produced from maltose using a transglucosylation reaction catalyzed by intact cells of A. pullulans. They cultivated A. pullulans KFCC 10245 in a medium composed of 0.5% (w/v) sucrose, 5% maltose, 0.5% yeast extract, 0.2% K2HPO4, 0.1% KC1, and 0.15% MgSO4 at 25°C for 96 hr. When maltose was used at 50% (w/v), about 50% (w/w) of panose was reported to accumulate after 120 hours of reaction at 55°C. The organism produced a fructosyltransferase when cultivated in a sucrose m e d i u m at a higher temperatures. D. NOVELOLIGOSACCHARIDES Glycosidases have the natural function of hydrolyzing terminal glycosyl residues from di- or oligosaccharides, in which process water acts as the glycosyl acceptor. The glycosyl residue enzyme complex can be intercepted by nucleophiles other than water. Often the substrate itself may act as the acceptor, as in the case of FOS synthesis from sucrose. Novel galactosides and oligosaccharides can be synthesized if different sugars and nonsugar alcohols are used as glycoside acceptors (Toone et al., 1989). These galactosides are of potential interest as fine-grade chemicals and as substrates for lipase-catalyzed construction of surfactants and natural emulsifiers such as galactosyl glycerides (Stevenson et al., 1993). The practicality of using lactose, a low-cost galactosyl donor, for large-scale synthesis of potentially important galactosides was studied by Stevenson et al. (1996). Their study augmented observations from various earlier studies that glucosidases generally have little acceptor specificity and that the only limitation was the water solubility of the acceptor molecule. Although water-insoluble acceptors (e.g., octanol) could be galactosylated in good yield, phenyl galactoside, an expensive amphiphilic substrate, had to be used as a galactosyl donor. Co-solvents

MICROBIAL PRODUCTION OF OLIGOSACCHARIDES

321

can improve yields with moderately hydrophobic acceptors (e.g., benzyl alcohol). Even the best co-solvents caused significant enzyme inactivation; hence, relatively large quantities of enzyme are needed. Optimization of galactoside yield by an excess of either lactose or acceptor is impractical due to solubility limitations. The main limitation to the acceptor concentration is its solubility and its effect on donor solubility and enzyme stability. Trincone et al. (1991) carried out synthesis of alkyl glucosides by using a crude homogenate of the thermophilic archaen Sulfiblobus solfataricus, which possesses [~-galactosidase activity. Synthesis of different alkyl-O-D-glucosides by transglycosylation using phenyl-[3-D-glucoside, phenyl-[~-D-galactoside, and lactose as carbohydrate donors was performed using a 14 molar excess of alcohols at 75°C and in the presence of 25% acetonitrile as co-solvent. They obtained a 25% yield within 20 minutes using a crude homogenate containing 7.2 mg total protein and a 14 molar excess of alcohol. Under the same conditions, they obtained a 19% yield of octyl [~-D-glucoside, higher than that reported by Mitsuo et al. (1984) with a 10 molar excess of alcohol and 100 mg of lactose from Kluyveromyces lactis, where 13% yield was obtained within 30 min. Trincone and colleagues (1991) obtained a 97% yield in a two-phase system with 5% water. Various sugars can act as acceptor molecules, resulting in formation of a wide range of oligosaccharides (Rastall and Bucke, 1992). Table II shows the different oligosaccharides that can be produced by using different sugar acceptors.

TABLE II NOVELOLIGOSACCHARIDESSYNTHESISEDUSING DIFFERENTACCEPTORS(o~-GLucOSIDASE) Acceptor sugar

Product

D-xylose

D-GIc-(c~-I~4D-Xyl D-Glc-c~-I--~IL-Xyl

D-ribose D-galactose D-mannose D-arabinose

D-Glc-c~-1---~4-D-Rib D-Glc-R-1---~6-D-Gal D-Glc-c~-l---~6-D-Man D-Glc-c~-I~2-D-Ara

Reprinted with permission from Rastall and Bucke (1992).

322

s.G. PRAPULLA et al. E. PURIFICATION OF OLIGOSACCHARIDES

Efficient extraction and isolation of oligosaccharides from the fermentation broth, where they are generally present along with unutilized substrate and hydrolysis products such as monosaccharides, is essential. Pure products are required for use in the food industry, where they are incorporated into health foods and energy foods, or used as texture modifiers and so forth. Use of high-grade pure oligosaccharides is also essential in the medical usage, in such applications as cell-pathogen interaction studies and blood-group determination. Relatively few reports are available on isolation and purification of oligosaccharides.

1. Purification of Fructooligosaccharides Takeda et al. (1994) studied purification of fructooligosaccharides with reference to 1-kestose applicable for large-scale purification. They detailed a procedure that involves three primary steps: (1) preparation of seed crystals, (2) crystallization, and (3) recrystallization.

2. Purification of Galactoo]igosaccharides Lopez-Leiva and Guzman (1995) carried out trials on the concentration of galactooligosaccharides present in whey hydrolysates by means of membrane filtration (nanofiltration). They used whey ultrafiltrate (UF) permeates that had been hydrolyzed to about 20%. A whey UF permeate previously hydrolyzed to about 20% was subjected to nanofiltration for fractionation of the oligosaccharides from smaller carbohydrates: lactose, glucose, and galactose. They obtained a retention factor of 37.0% for lactose, 15.4% for glucose and galactose, and 74.7% for oligosaccharides, with the magnitude of the retention factor corresponding to the size of the corresponding molecules. Oligosaccharides were retained most when the monosaccharides were only slightly retained.

3. Purification of Chitooligosaccharides Samain et al. (1997) described utilization of oligosaccharide adsorption on activated carbon for purification. In this method, chitooligosaccharides were recovered exclusively in the pellet obtained on centrifugation of the culture broth. The cells were disrupted by boiling, and cell debris was removed by centrifugation. Chitooligosaccharides were purified by adsorption on activated charcoal followed by elution with

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323

30% ethanol. The yields ranged from 50 to 60%. Yields ranged from 30 to 35% when the chitooligosaccharides purified on Dowex 50WX4 eluted with 2% aqueous ammonia.

IV. New Approaches to Microbial Oligosaccharide Synthesis The growing market for oligosaccharides necessitates continued efforts to develop better, high-yield, and efficient methods of their synthesis. Many researchers have studied different approaches to obtain both high yields and allow synthesis of novel oligosaccharides. The disadvantages of the conventional methods include the high cost of the enzyme, the low yields due to poor reactants solubility, the narrow products range, and the instability of enzymes. The new approaches discussed below allow much a larger scope for addressing and overcoming these problems. A. USE OF ORGANIC SOLVENTS

Synthesis of oligosaccharides using glycosidases in a reverse reaction requires that the conditions be modified to allow the equilibrium to shift in the direction of synthesis and not hydrolysis. Conventionally, this has been carried out by using high concentrations of mono- and disaccharides. The most significant reactant, however, is water, and many workers have experimented with the use of organic solvents to reduce the water activity of their reaction systems. Shiffin and Hunn (1969) found that [3-galactosidase is activated by the presence of lower levels of alcohols in concentrations up to 20-30% (v/v). Addition of alcohols increased the speed of the rate-limiting deglycosylation step, resulting in formation of the appropriate alkyl glycoside. Alcohols have also been shown to be good acceptors of the sugar residue. Shinoyama et al. (1988) reported the synthesis of many alkyl xylosides (e.g., methyl, ethyl, 2-phenyl ethyl, cyclohexyl) by using the ~-xylosidase from A. niger on xylobiose in the presence of alcohols, with yields ranging from 16 to 114%. Nilsson (1986) found that organic co-solvents such as N,N-dimethylformamide reduced the yield of oligosaccharides formed by ~-galactosidase, possibly due to a reduced interaction between CH and C H 2 groups on the substrate and amino acids in the active site of the enzyme. While working with different systems, Ogawa et al. (1990) and Usui and Murata (1988) described the use of alcohols as aids to oligosaccharide synthesis, with methanol being more effective than ethanol. However,

324

s.G. PRAPULLA et al.

high concentrations of alcohol are inhibitory to enzymes and can bring about denaturation. B. USE OF AQUEOUSTwO-PHASE SYSTEMS Aqueous two-phase systems are formed when certain water-soluble polymers are mixed in aqueous solution. Their incompatibility results in the formation of two aqueous phases. The application of aqueous two-phase systems in biotechnology and biochemistry has largely been in extractive bioconversion and protein purification (Rastall and Bucke, 1992). Bartlett et al. (1992) used a novel kinetic approach in a polyethylene glycol (PEG)/dextran two-phase system for synthesis of oligosaccharities. They used jack bean ~-mannosidase and found that the enzyme partitioned very strongly into the dextran phase, while the p-nitrophenyl c~-mannoside donor and p-nitrophenyl 13-galactoside acceptor partitioned evenly. They manipulated the phase volume ratios to obtain an excess of the PEG phase and reported a 10-fold increase in oligosaccharide formation per unit of enzyme compared to a normal aqueous system. Bartlett and colleagues (1992) recommended the twophase system for large-scale synthetic reactions (Fig. 5). C. USE OF IMMOBILIZED SYSTEMS

The most expensive component of an enzymically catalyzed oligosaccharide synthesis reaction is the enzyme itself. The mono- and disaccharide reactants can be recycled after separation from products. Immobilization allows efficient use of an enzyme for practical applications. Immobilization also stabilizes the enzyme when used at high temperatures or in the presence of organic solvents (Larsson et al., 1987). Careful choice of immobilization conditions could also make it possible to influence the product spectrum (Rastall and Bucke, 1992). Ajisaka et al. (1987) used the equilibrium approach for immobilization of ~-galactosidase and synthesized lactulose and allolactosamine from mixtures of galactose, fructose, and n-acetyl glucosamine. They immobilized the E. coli enzyme on either Leupergit C or Sepharose 4-B and carried out synthesis by circulating solutions containing 10% (w/v) galactose and either 30% (w/v) N-acetyl glucosamine or 50% (w/v) fructose through columns of the immobilized enzyme. They connected the enzyme columns with a column of activated carbon to remove the products and thereby shift the equilibrium in the direction of condensation and increased yield. Oligosaccharides were eluted by

MICROBIAL PRODUCTION OF OLIGOSACCHARIDES

325

iiii ii

•'. PEG "."

•i )i

!i!Dextran ii 1) Initial two-phase system

~ 1 1 )

iiiiiiiiiii! :::PEal:::

Remove PEG phase, [ isolate products I Replenish system with fresh PEG phase containingdonor and acceptor

~Se~a~~ V

Mix and incubate

PEG phase contains:syntheticproducts, acceptor, p-nitrophenol,monosaccharide

Dextran phase contains:Enzyme,products, acceptor,p-nitrophenol,monosaccharides

3) Allowto settle

F~c. 5. Synthesis of oligosaccharides in aqueous two-phase systems. Reprinted with permission from Rastall and Bucke (1992).

washing the column with 10-50% (v/v) ethanol. They reported yields of 0.7-0.9% N-acetyllactosamine and 7.0-9.1% N-acetylallo]actosamine per day. In a study carried out by Ajisaka and Fujimoto (1989) using the same immobilized E. coli ~-galactosidase system with 10% (w/v) galactose and 50% (w/v) sucrose, formation of only isoraffinose was reported in comparison to a mixture of isoraffinose and D-Glcp-~(1-->2)-[D-Galp~ [~(1-->6)]-[3-D-fructose obtained from the batch reaction. This regioselectivity was probably due to immediate binding of trisaccharide products to the activated carbon column, thereby preventing attainment of equilibrium. The product spectrum was thus a consequence of differences in the activation energies of formation. Immobilized enzymes have also been used to synthesize oligosaccharides by the kinetic approach. Larsson et al. (1987) used immobilized

326

s.G. PRAPULLAet al.

~-galactosidase (on Sepharose CL-4B) to hydrolyze lactose in the presence of N-acetylgalactosamine, resulting in synthesis of D-Gal-[3(1-~6)D-Gal-NAC. Mozaffar et al. (1988) immobilized Bacillus circulans ~galactosidase on porous silica gel by crosslinkage with glutaraldehyde, and synthesized a range of oligosaccharides during lactose hydrolysis. Kery et al. (1991) reported the successful use of cellulose beads as a support matrix for immobilized [3-galactosidase. They synthesized a mixture of 6- and 3-1inked disaccharides and trisaccharides from p-nitrophenyl-~-galactoside as a glycosyl donor and methyl-c~-galactoside as an acceptor. Hayashi et al. (1991) immobilized a fructosyl-transferring enzyme from Aureobasidium sp. ATCC 20524, which produces 1-kestose from sucrose, onto a Shirasu (volcanic ash) porous glass (1142 U/g support). The packed column was stable for more than 30 days during continuous operation. Yun et al. (1992) investigated continuous production of fructooligosaccharides employing a packed-bed reactor charged with cells of A. pullulans KFCC 10245 immobilized on calcium alginate. They optimized conditions for reactor operation at a feed concentration of 860 g/liter, a feed rate (as superficial space velocity) of 0.2 per hour, and a temperature of 50°C. Under these optimal conditions they obtained a productivity of 180 g/liter/hr, with initial activity maintained for more than 100 days. They successfully scaled up the reactor to a production scale of 1000 liters. Yun and Song (1996) carried out continuous production of fructooligosaccharides from sucrose using fructosyltransferase immobilized on a highly porous resin, Diaion HPA 25. The optimal operation conditions of the immobilized enzyme column were a 600 g/liter of sucrose feed concentration and a feed rate as superficial space velocity of 2.7 per hour at pH 5.5 and 55°C. An 8% loss of initial activity was reported after 30 days of continuous operation when the column was rn at 50°C; a productivity of 1174 g/liter/hr was reported during this period. D. USE OF LIPID-COATEDGLYCOSIDASES The addition of water-miscible organic solvents increases the yield of transglycosylation catalyzed by glycosidases in a reverse reaction (Mori et al., 1997). However, the transglycosylation yields in these examples are usually low because the hydrolysis reactions proceed fast relative to transglycosylation in homogeneous aqueous organic media. Mori et al. (1997) prepared a lipid-coated [~-D-galactosidase in which the enzyme surface is covered with a lipid monolayer, and two long lipophilic alkyl tails serve to solubilize the enzyme in organic solvents. In a two-phase aqueous organic system, the lipid-coated enzyme exists

MICROBIAL PRODUCTION OF OLIGOSACCHARIDES

327

in the organic (2-propyl ether) phase and acts as an efficient transglycosylation catalyst for various hydrophobic alcohols with lactose in the aqueous buffer solution. Native [3-D-galactosidase, however, gave poor yields of both transglycosylation and hydrolysis reactions in a twophase system due to denaturation of the enzyme at the interface. E. USE OF RECOMBINANTS

Large-scale synthesis of specific tailor-made oligosaccharides is increasingly becoming essential. This has increased the use of glycosyltransferases over the glycosyl hydrolases, as the former display high regiospecificity for the acceptor and the donor substrate. The major drawback in utilization of glycosyltransferases is a lack of availability and the prohibitive cost of the sugar nucleotides used as activated sugar donors. To tackle this drawback, Samain et al. (1997) used growing bacterial cells as natural minireactors for continuous regeneration of sugar nucleotides and used the intracellular pool of sugar nucleosides as the substrate for in-vivo synthesis of "recombinant" oligosaccharides by recombinant glycosyltransferases. The method reported by them described cultivation of E. coli harboring genes for oligosaccharide synthesis. They produced pento-Noacetyl chitopentose and its deacetylated derivative tetra-N-acetyl-chitopentose in high yields (up to 2.5 g/liter) by cultivating at high density cells of E. coli expressing nodC or nodBC genes (nodC encodes for chitooligosaccharide synthesis and nodBC for chitooligosaccharide N-deacetylase). V. Assays and Structural Determination of Oligosaccharides Measurement of enzyme activity and determination of oligosaccharide concentration and yield require a knowledge of the methods for estimation of the compound itself. Assays for oligosaccharides have largely been reported using chromatographic methods, primarily by high-performance liquid chromatography (HPLC). Other techniques reported by various authors include paper chromatography, thin-layer chromatography (TLC), mass spectrometry (MS), and nuclear magnetic resonance (NMR).

A. HIGH-PERFORMANCE LIQUID CHROMATOGRAPHY Advances in high-performance liquid chromatography (HPLC) with particular reference to column and instrument technology have led to speedy analysis of oligosaccharides (Table I/I). Jeon and Mantha (1985) reported that both polar-bonded-phase and resin-based HPLC columns are commonly used for carbohydrate separation. The polar-bonded phases are efficient, and carbohydrates elute in order of increasing

328

S. G. P R A P U L L A e t a l . T A B L E III ANALYSIS OF OLIGOSACCHARIDES USING H P L C

Column Aminospheri-5 (4.6 × 250 mm) Spherogel carbohydrate (7.5 x 30 cm) Shimadzu PNH2 (4.6 × 150 mm)

Column temperature

Mobile phases

Flow rate ml/min

Detection system

Source

Ambient

Acet onitrile:distilled water 75:25 (v/v)

2.0

Refractive index (RI)

Jeon & Mantha (1985)

80°C

Distilled water

0.6

RI

Jeon & Mantha (1985) (1985)

Ambient

Acetonitrile:distilled water 75:25 (v/v)

1.0

RI

Hidaka etal. (1988)

Acetonitrile:distilled water 75:25 (v/v)

1.5

RI

Jung etal. (1987) Duan eta]. (1994)

Bondapak carbohydrate (10.4 × 300 mm) Wakopack WB-T-130E (7.8 × 300 mm)

60°C

Distilled water

0.2

RI

Hayashi et el. (1991)

Shodex Ionopak ks-801 (8 × 300 nm)

7O°C

Distilled water

1.0

RI

Nakao et el. (1993)

Asahipak NH2P-50 (5 × 250 mm)

25°C

Acetonitrile:distilled water 75:25 (v/v)

1.0

RI

Nakao eta]. (1993)

A m i n e x HP X 42C (7.8 × 300 mm)

85°C

Water

1.0

RI

Yun etal. (1995, 1997), Yun & Song (1993, 1996)

A m i n e x HP X 42C (7.8 x 300 mm)

85°C

Acetonitrile:distilled water 75:25 (v/v)

0.75

RI

Dumortier etal. (1994)

Waters p Bondasphere 5 p NH2 - IoOA (3.9 × 150 mm)

30°C

Acetonitrile:distilled water 75:25 (v/v)

1.0

RI

Takeda etal. (1994)

Sperisorb NH2 (4.6 × 250 ram)

Acetonitrile:distilled water 75:25 (v/v)

1.8

RI

Barthomeuf & Pourrat (1995)

Nucleosil capcell 5 pm

A m m o n i u m acetate 50 m M

0.6

RI

Samain et al. (1997)

Capcell Pak C18 (3.9 x 250 mm)

Distilled water

-

*

Barthomeuf etal. (1997)

*168-diode array spectrophotometer.

MICROBIALPRODUCTIONOF OLIGOSACCHARIDES

329

monosaccharide chain length. On the other hand, components elute in order of decreasing molecular size from resin-based columns. Resinbased columns are increasingly being studied for use in analysis of small oligosaccharides (with resolution of polymers limited to a degree of polymerization equal to 4). Concentrations of oligosaccharides can be determined by weighing the individual peaks and comparing them to standard curves. This method has been widely used. The accuracy, sensitivity, reproducibility, and ease of operation with which the samples can be analyzed has made this a very popular technique. B. PAPERCHROMATOGRAPHY Toba et al. (1980) detailed a method for estimation of oligosaccharides by paper chromatography using sheets (40 x 40 cm) of Toyo No. 514 paper. The papers were developed in a butanol:pyridine:water (6:4:3) medium. Sugars were detected by an aniline hydrogen phthalate (AHP) reagent. Toba and Adachi (1978) described two-dimensional analysis of oligosaccharides by paper chromatoelectrophoresis. They carried out detection and characterization of oligosaccharides with the use of diphenyl amine aniline phosphoric acid (DAAP), triphenyl tetrazolium chloride (TTC), and AHP. This method is simple and is now widely used for separation of individual oligosaccharides, but it lacks sophistication and accuracy and is more suited to qualitative analysis rather than quantification of reaction products. Paper chromatoelectrophoresis is an improved version of this method, it is not routinely used because of its high cost. C. GAS-LIQUIDCHROMATOGRAPHY Toba and Adachi (1978) carried out gas-liquid chromatography (GLC) of oligosaccharide trimethylsilyl (TMS) esters with a Hitachi model 063 gas chromatograph fitted with a hydrogen flame ionization detector and a stainless steel column (2 m x 3 mm). Nitrogen was used as a carrier gas at a flow rate of 30 ml/min, with temperatures rising from 150 to 300°C at 3 °C/rain. They obtained trimethyl derivatives of the transfer reaction products by shaking the reaction products with pyridine:hexamethyldisilazane:trifluoroacetic acid (10:9:1, v/v), and separating on a column of Chromosorb W coated with 1.5% SE-52. They also made a detailed GLC analysis of the methyl glycosides using a stainless steel

330

s.G. PRAPULLAet al.

column (2 m X 3 mm) packed with 8% diethylene glycol succinate on Diasolid L operated at 200°C, with nitrogen as a carrier gas at 20 ml/min. They achieved methylation of oligosaccharides by shaking 0.5-2.0 mg of sample with 0.2 ml methyl iodide, 0.2 ml N,N-dimethylformamide, and 0.2 g silver oxide at room temperature in the dark for 18 hours. The mixture was then filtered, the residue washed with chloroform, and the filtrate evaporated to dryness. The products were boiled with 5% methanolic hydrogen chloride for 8 hours. The resulting methylglycosides were identified by comparing their retention times with that of standards. Burvall e t al. (1979) reported the use of a 2-m glass column packed with 3% ECNSS-M on Gas Chrom Q (100-200 mesh) at a specified column temperature for sugar alditol acetates and at 160°C for partially methylated alditol acetates. For permethylated alditol derivatives of dito tetrasaccharides, they recommended the use of a glass capillary column (25 m x 0.25 mm) wall coated with SE-30 at a specified column temperature and at 160°C for partially methylated alditol acetates. Stevenson e t al. (1993) prepared samples for GLC analysis by adding 10 gl of quenched reaction sample to acetonitrile (100 gl) and evaporating to dryness at 35°C under a stream of air in a block heater. The solid residue was trifluoroacetylated by addition of dry pyridine (5 gl), dichloromethane (65 ~1), and trifluoroacetic anhydride (30 ~1). They injected 1 ~1 of sample after incubation at 35°C for 15 min onto a gas chromatograph fitted with an S.G.EBP1 (0.22 mm x 25 m) capillary column and a flame ionization detector. They maintained injector and detector temperatures at 300°C and oven temperature at 120°C for 2.5 min after injection, then increased by 10°C/min to 270°C. This is a powerful technique for both qualitative and quantitative analysis with a high degree of accuracy. However, reaction products need to be converted to trimethyl derivatives, which is a laborious process. Much care needs to be exercised during methylation of sugars; otherwise, incomplete methylation leads to erroneous results and affects column performance. D. THIN-LAYERCHROMATOGRAPHY Wierzbicki and Kosikowski (1973) recommended using 0.25 mm thick plates of silica gel for analysis of oligosaccharides. A mixture of n-butanol:acetic acid:diethylether:water (9:6:3:1) was used as the developing solvent and the color-developing reagent was prepared by dissolv-

MICROBIALPRODUCTIONOF OLIGOSACCHARIDES

331

ing 5 g of benzidine in acidified ethanol. After development, the plates were dried in an oven at 100°C for 10-15 minutes, sprayed with color reagent, and heated to 100°C for 25 minutes. Mono- and oligosaccharides gave yellow brown spots under daylight and green spots under short-wavelength UV. Balken et al. (1991) qualitatively determined oligosaccharides by TLC on silica gel plates using of chloroform:ammonium hydroxide:methanol solvent system (60:20:54). Spots were visualized by spraying with 1-naphthol or phosphomolybdate. Prakash et al. (1989) suggested a similar process with use of ethyl acetate:acetic acid:water (2:1:1, v/v/v) as the solvent system. Quantitative estimation was done by extracting individual spots developed on TLC using a mixture of 40% trichloroacetic acid in water, acetic acid, and ethanol (1:1:8) and measuring the optical density at 400 mn. Concentrations were determined from a standard curve prepared with lactose (Wierzbicki and Kosikowski, 1973). Stevenson et al. (1993) performed TLC using silica gel 60 F254 on aluminum foil plates with ethyl acetate:pyridine:water (33:13:4). Compounds were visualized by dipping the plates in a mixture of 5% (v/v) concentrated sulfuric acid and 2% (w/v) anisaldehyde in 95% ethanol and heating with hot air until spots appeared. Trincone et al. (1991) monitored formation of phenyl iS-D-glycosides by carrying out TLC using chloroform:water (8:2, v/v) as the eluant. TLC is one of the simplest methods for analysis. However, the accuracy with which the eluted spots can be extracted is poor, so that it does not find wide application. F,. 13C-NMR ANALYSIS This technique has largely been used for elucidation of the structure elucidation of the oligosaccharides. Prakash et al. (1989) obtained proton-decoupled 13C-NMR spectra of oligosaccharides at 25 MHz with a JEOL FX100 operated in pulsed Fourier transform mode at 30°C using D20 as a solvent. Chemical shifts were referenced to an external standard 2,2,3,3-tetra-deuterio-4,4-dimethyl-4-sila pentanoate (TSP-d4). Barthomeuf et al. (1997) carried out NMR analysis of oligosaccharides at 100 MHz using a Bruker AC 400 spectrophotometer and a sample containing approximately 5 mg/ml sugar in deuterium oxide:water (1:2, v/v). They expressed the results obtained in ppm from the signal of trimethyl silane (TMS) and referenced chemical shifts using acetone as an internal standard. Samples used for NMR analysis were obtained by semipreparative HPLC. NMR is one of the most powerful techniques

332

s . G . PRAPULLA et al.

available for structural elucidation. The cost of the equipment is its major constraint. Moreover, the protocol for sample preparation is much more elaborate, and it is necessary to carry out semipreparative HPLC before samples are subjected to NMR analysis. F. FAB MASS SPECTROMETRY Prakash eta]. (1989) used FAB mass sPectrometry for elucidation of the major galactooligosaccharides formed by Trichoderma harzianum. They recorded positive-ion fast atom bombardment mass spectra in a JMS DX300 mass spectrometer. The sample and stainless steel probe tip were introduced into the FAB source and bombarded by a 3-keV argon beam. They dissolved the oligosaccharide sample (-5 mg) in about 5% sodium acetate and loaded it on a glycerol-coated probe tip. Similar to NMR, this method is more applicable for structural elucidation rather than routine analysis. Elaborate sample preparation steps and the initial cost of the equipment are limitations on its wide application.

Vl. Applications of Oligosaccharides Oligosaccharides have application in both nonfood and food-related fields. A. NONFOOD APPLICATIONS

The most important nonfood applications of oligosaccharides are in the medical and biotechnological fields. These compounds play a wide range of roles within living cells, often acting as recognition molecules. Microbial production allows not only easy availability of oligosaccharides but also facilitates synthesis of "tailor-made" compounds, thus giving additional thrust to studies focused on elucidating the molecular mechanism of a variety of biological functions. Oligosaccharides, generally as glycoconjugates, have been implicated in cell-cell, cell-virus, and cell-bacteria interactions (Rademacher et al., 1988; Schnaar, 1991). The lipopolysaccharide O antigen found in Gram-negative bacteria, the blood group determinants, and several tumor-associated antigens have potential as diagnostic tools. Oligosaccharides regulate cellular differentiation and act as inducers of disease-resistance responses following invasion by fllngal pathogens (McNeil et al., 1984). Bacterial cell-surface oligosaccharides are reported to be

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responsible for host recognition and binding following infections of root hair with nitrogen-fixing bacteria (Lerouge et al., 1990). Hedbys et al. (1984) suggested the potential use of 6-O-~-D-galactopyranosyl-2acetamido-2-deoxy-D-galactose, formed by the transgalactosylase activity of E. coli [5-galactosidase, as a cell surface receptor. Ooi et al. (1985) used the [~-galactosidase of A. oryzae to synthesize glycosides of digoxigenin. A [3-galactosyl-serine-glycoprotein component was synthesized separately by Cantacuzene et al. (1991) and Sauerbrei and Thiem (1992) using the ~-galactosidase of E. coli and A. niger. Hedbys et al. (1989) were able to synthesize the blood group determinant Gal-[31-3Glc-NAC[~-SEt using [~-galactosidase from E. coil A Salmonella-inhibiting therapeutic composition containing fructooligosaccharide (largely Neosugar) as the effective compound was patented by Coors Biotechnology (Speights, 1990; Speights et al., 1991). Other nonfood applications of oligosaccharides include their use in drug delivery, cosmetics, and mouthwash. A Japanese company has patented the use of an enzymatically synthesized, nonreducing, neotrehalose-like oligosaccharide for use in cosmetics and pharmaceuticals. This compound is stable, not susceptible to crystallization, and has appropriate viscosity Aga eta]., 1995). Nonfermentable oligosaccharides have been used to manufacture a high-density liquid preparation of yeast (>800 g/liter) with improved activity retention, easy dissolution, uniform suspendability, and greater tolerance to repeated freeze-thaw operations (Suoranta, 1991). Chitooligosaccharides are also emerging as an important class of oligosaccharides with a diversity of applications, including use as antimicrobials, plant growth inhibitors, and cosmetic components (Crittenden and Playne, 1996). B. FOODAPPLICATIONS Oligosaccharides have grown immensely popular as food ingredients, particularly in Japan and Europe, largely due to the health benefits associated with their consumption. Several foods containing oligosaccharides as the functional ingredient have obtained FOSHU (Foods of Specific Health Use) status under Japanese federal legislation. In fact, foods incorporating oligosaccharides comprise 34 of the 58 approved foods in the 1996 FOSHU list. Oligosaccharides have excellent functional properties and improve the general colonic environment. They are bifidogenic factors and improve the growth of indigenous bifidobacteria (Crittenden and Playne,

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1996). The growth of bifidobacteria has many beneficial effects, such as improved digestion and absorption, increased vitamin availability, and prevention of gut colonization by pathogens and putrefactive bacteria. The current interest in application of bifidobacteria to improve colonic health has made oligosaccharides quite popular. Oligosaccharides are being included in probiotic yogurt and yogurt drinks to produce synbiotic products. "Bifiel," produced by Yakult Honsha, Japan (Kan et al., 1989) contains galactooligosaccharides, while fructooligosaccharides have been incorporated into "Symbalance" (Toni Milch, Switzerland), "Fyos" (Nutricia, Belgium), and "Fysiq" (Mona, Netherlands). Barthomeuf et al. (1997) reported that bifidogenic activity is optimum with short oligofructosaccharides in which the fructosyl units (n -- 2-8), are bound by a 132-1 linkage. New physiological effects of oligosaccharides consumption continue to be elucidated, including possible protection against the development of colon cancer. Oligosaccharides help prevent constipation because they are indigestible and so have an effect similar to dietary fiber. Apart from general enhancement of the colonic environment, oligosaccharides have been shown to exhibit protective action against diarrhea caused by the heat-stable enterotoxin of E. coli (Sta). Sta c a u s e s diarrhea by stimulating intestinally bound guanylate cyclase. Using the T84 human colon carcinoma cell line, these studies showed that fucosylated oligosaccharides could inhibit Sta-stimulated guanylate cyclase by 6080%, whereas other oligosaccharides exhibited a 17-18% inhibitory effect on the enzyme (Daniel et al., 1996). Oligosaccharides are incorporated into several health drinks and athletic beverages. A general-purpose sports beverage made up of oligosaccharides (20 g/liter), sucrose (55 g/liter), citric acid (1.8 g/liter), citric aroma (1.0 g/liter), and NaC1 (1.0 g/liter) has been patented (Korduner et al., 1982). Another product especially adapted for rapid administration of water and carbohydrates during heavy muscle work has been patented by Pripps Bryggerier (Gyllang et al., 1986). It is a monosaccharide-free solution containing 3-25% (by weight) of a mixture of soluble oligosaccharides. The same company holds an international patent (Newsholme et al., 1988) for an instant beverage mix for athletes with 0-750 g/liter oligosaccharides with 0-55 g/liter monosaccharides and 2-40 g/liter amino acids. A Japanese company holds an international patent (Takaichi et al., 1991) for a liquid nutrient product containing three to six oligosaccharides.

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1. Oligosaccharides in Processed Foods

Oligosaccharides are used in the confectionery, beverage, and bakery industries not as pure products, but as mixtures containing oligosaccharides with various degrees of polymerization. The choice of an appropriate mixture for a particular food application depends on the specific physicochemical and physiological properties of the oligosaccharides, which in turn depends on their structure. Oligosaccharides are typically 30-60% as sweet as sucrose, which makes them desirable in foods when a low sweetness level is used to enhance other food flavors. In conjunction with intense artificial sweeteners, they help to mask certain aftertastes. The primary advantage of using oligosaccharides is that they provide bulking properties almost identical to that of sucrose and, being indigestible, they are safe for consumption by individuals suffering from insulin-dependent diabetes. Oligosaccharides are therefore being widely used in confections, jams, marmalades, and desserts (Crittenden and Playne, 1996) as low-calorie, noncariogenic sugar substitutes. The use of 15-40% oligosaccharides in confectionery syrup was patented by the Corn Products Company (Walon, 1971). Hayashibira (1974) patented the use of fructooligosaccharides as sweetening agents in the manufacture of frozen dairy desserts, ice creams, sweetened condensed milk, sweetened dry milk, and other dairy products. The use of oligosaccharides with one to four fructose units linked to sucrose as low cariogenic sweeteners was patented by Meiji Seika Kaisha (Adachi and Hidaka, 1981) in 1981. The same company holds an American patent (Adachi and Hidaka, 1987) for the use of microbially produced fructooligosaccharides as low-calorie noncariogenic sweeteners. Tate and Lyle, a British company, patented (Beyts, 1989) the use of glucooligosaccharides in a synergistic mixture of sweeteners to be used in dietetic foods, beverages, bakery products, and confections. The use of fructooligosaccharides as normal and low-calorie sweeteners is covered by a French patent (Biton et al., 1989) held by Ronssel-Uclaf of South Africa. Meiji Seika patented a process for production of a candy that has pressurized gas entrapped within it along with incorporation of oligosaccharides. On consumption, the candy releases the gas, generating a pleasant taste in the mouth (Sumi et a]., 1991). Roquette Fr~res patented (Mentink and Serpelloni, 1992) a low-calorie chocolate in which the sweetener is an oligosaccharide.

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Raffinerie Tirlemontoise, a Belgium company, obtained an international patent (de Soete and Freund, 1993) for a low-calorie chocolate confectionery product in which sugar is wholly or partially replaced by linear or branched fructooligosaccharides with or without the addition of intense sweeteners. Unilever holds a European patent (Plug, 1995) describing the use of oligosaccharides in foods. Due to their noncariogenicity, low calorific value, and viscosity, oligosaccharides are being increasingly incorporated into a variety of chewing gums. The Wrigley Company (United States) has developed a petroleum wax-free chewing gum incorporating oligosaccharides that act as binders. The oligosaccharides described were fructooligosaccharide, isomaltose, and oligofructose (Yatka et al., 1994). The same company holds a patent (Yatka et al., 1995) for the manufacture of other chewing gums containing oligosaccharides. Embodiments of the product include use of oligosaccharides as a rolling compound applied to the product, use as a coating such as a hard shell, for pellet gum, and as center fill in chewing gum. The patent also covers co-drying of oligofructose with other sweeteners and evaporation to make syrups for use as encapsulating agents for high-intensity sweeteners or flavors used in gums. The various functional and physicochemical properties of oligosaccharides have popularized their use in the bakery industry, especially since they have also been shown to act as strong inhibitors of starch retrogradation (Nakakuki, 1993). A 1988 patent (Kono et al., 1988) describes oligosaccharides originating from agar and/or carrageenan as being effective in preventing retrogradation of gelatinized starch. Yakult Honsa (Japan) described a method for producing bread with a galactooligosaccharide incorporated at 2.6% by weight. They also hold a patent for the process of making bread containing Gal-(Gal)n (Sonoike et al., 1992). Partially esterified oligosaccharides are suitable as fat substitutes since they have good sensory properties and have no calorific value, as they are not substantially hydrolyzed in the digestive tract. They have excellent mouth feel and many other properties similar to vegetable oils and fats (White, 1990, 1991). Carboxylic acid esters of oligosaccharides have been used in the preparation of edible water-in-oil and oil-in-water emulsions with reduced energy and fat content, as in the preparation of margarine, cream, and mayonnaise. A novel branched oligosaccharide synthesized by Aspergillus s y d o w i fructosyltransferase has also been reported to be useful as a food additive (Mieth et al., 1985).

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VII. Conclusions

Oligosaccharides have emerged as functional foods with varied applications in response to an increasing consumer demand for healthier foods. The functional properties of oligosaccharides make them unique in food applications, and their popularity and demand is continually increasing. In order to meet the rising demand, various research groups have successfully produced them using microbes. Microbial production of oligosaccharides is an effective alternative to chemical synthesis. A whole range of novel oligosaccharides are being developed. Research efforts directed at production of novel oligosaccharides with varied functional properties will be a future trend. Application of novel production techniques for better yields is also envisaged. There is tremendous potential for broader elucidation of the physiological effects of oligosaccharides. Unique physicochemical properties, combined with health benefits, make the oligosaccharides interesting from the point of view of research and development, along with the large scope of potential commercialization.

Acknowledgments The authors thank Mr. M. N. Ramesh, a scientist in the Food Engineering Department, who helped us compile and prepare this chapter.

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Index

A ABT-773, 80-81

Acetobacterium woodii, fatty acid hydration by, 207 Acetyl esterase, antibiotic resistance and, 147 Acriflavine, 132 Actinomadura madurae, prodigiosins, 15 Actinomadura pel]etieri, prodigiosins, 15 Actinomycetes, oligosaccharide biosynthesis by, 319 Actinomycin, biosynthesis of, 141 Actinorhodin, 18 Acyl carrier protein (ACP), 121 Adriamycin, 125 AIB factor. See Anti-isobntyrate factor Air pollution, by organochlorine pesticides, 274 Alcohols, biocatalytic dynamic resolution of, 66-70 Algae, degradation of lindane by, 272, 287-288 Allysine ethylene acetal, enzymatic synthesis of, 46-51, 71 Allzyme phytase 115,183 Alteromonas ruber, prodigiosins, 14 Amidase, enzymatic synthesis of chiral drug intermediates, 54-55 Amycolotopsis orientalis, bioactive products from, 120 Anabaena, degradation of lindane by, 288 Animal feed, phytase and, 159-160, 175-177 Anthracyclines biosynthesis of, 125 hybrid bioactive products of, 137 Anti-Alzheimer's drug, enzymatic synthesis of chiral drug intermediates, 66, 72 Antibiotic properties, of prodigiosin, 22-23 Antibiotic resistance, 144-148

Anticholesterol drugs, enzymatic synthesis of chiral drug intermediates, 57-60, 72 Antihypertensive drugs enzymatic synthesis of allysine ethylene acetal, 46-51, 71 BMS-199541-01, 34-43 L-6-hydroxynorleucine, 43-46 Anti-isobutyrate (AIB) factor, 143-144 Antimalarial activity, of prodigiosin, 23 Antiviral agents, enzymatic synthesis of chiral drug intermediates, 61-63 Apase6, 181 aphA gene, 181 appA gene, 170 Aquaculture, phytase use in, 186-187 Arabidopsis thaliana phytase, 171-172 Arthrobacter globiformis, degradation of lindane by, 288 Arthrobacter simplex, enzymatic synthesis of chiral drug intermediates, 67 Arthrobacter spp., oligosaccharide biosynthesis by, 314 Arxula adeninivorans phytase, 173,176 Aspergillus fumigatus phytase, 176, 180, 182

Aspergillus japonicus, fructooligosaccharide biosynthesis by, 310

Aspergillus niger enzymatic synthesis of chiral drug intermediates, 63-65 oligosaccharide biosynthesis by, 310-311,314, 316-317 Aspergillus niger phytase, 160-170, 175-182, 184 Aspergillus oryzae, oligosaccharide biosynthesis by, 317-319 Aspergillus phoenicis, 1-kestose biosynthesis by, 309-310 Aspergillus sydowi, oligosaccharide biosynthesis by, 336 Aspergillus terreus phytase, 176

345

346

Aureobasidium pullulans,

INDEX

~-lactams antibiotic resistance, 148 biosynthesis of, 118-120 Bialaphos, 131 Bifidobacterium bifidum, oligosaccharide biosynthesis by, 319 Bifidobacteriam spp., in human gastrointestinal tract, 304,333-334 Bioactive products from Streptomyces, 113-114 biosynthesis of, 115-131 B derivatives of, 133-137 molecular genetics of, 131-133 [~3-adrenergic receptors, 52 regulation of production, 137-144 [~3-receptor agonists, enzymatic synthesis resistance to, 144-147 of, 52-57, 71 Biocatalysis. See Microbial/enzymatic Bacillus amyloliquefaciens phytase, 172 synthesis Bacillus brevis, bioactive products Biocatalytic dynamic resolution, of from, 115 racemic diol, 66-68, 72 Bacillus licheniformis, bioactive products Biodegradation from, 115 of lindane Bacillus megaterium, enzymatic by algae and cyanobacteria, 272,277, synthesis of chiral drug 287-288 intermediates, 45 by bacteria, 272, 277,279-285 Bacillus pumilus, fatty acid hydration by, by fungi, 285-287 207 Bio-Feed phytase, 183 Bacillus subtilis phytase, 172, 176 Biorefining, 222 Bacitracins, biosynthesis of, 115-116 Bizio, Bartolomeo, 6, 8 Bacitracin synthetase, 117 Bleomycin, 115 Bacteria. See also Bioactive products; Blood, cultural history of, 2 Microbial/enzymatic synthesis; "Blue diaper syndrome," 19 Secondary metabolites BMS-186318, biotransformation of, 61 degradation of lindane by, 272,279-285 BMS-186716, microbial/enzymatic genetically engineered microorganisms synthesis of, 34 (GEMs), 288-289 BMS-188494, biotransformation of, 58, 72 oligosaccharide biosynthesis by, BMS-199541-01, biotransformation of, 309-315,318-319 34-43, 71 prodigiosins, 2-26 BMS-201391-01, biotransformation of, xylose-fermenting, 226-227, 233 35-43, 71 Bacterium prodigiosum, prodigiosins, 7 BMS-202665, biotransformation of, 41, 43 Bacteroides polypragmatus, xylose BMS-203528-01, biotransformation of, fermentation with, 233 39-41 Barley, trihydroxy fatty acids from, 213 BMS-204556, biotransformation of, Basidiomycetes, biodegradation of 39-40, 42 organochlorine compounds by, BMS-264406, biotransformation of, 41, 43 272,286 Bread, red spoilage of, 2-5 4-Benzyloxy-3-methanesulfonylamino-2'-br Butyl-meta-cycloheptylprodiginine, 14 omoacetophenone, microbial 7-Bntyrolactone, 142 reduction of, 52-54 oligosaccharide biosynthesis by, 305,313-314,320 Autoregulators, bioactive products from Streptomyces, 143 Avermectins, biosynthesis of, 128 Azithromycin, 80-81

INDEX C

Caldariomyces fumigo, enzymatic synthesis of chiral drug intermediates, 61-63 Caldocellus saccharolyticum, oligosaccharide biosynthesis by, 318-319 Candicidin, biosynthesis of, 127 Candida antarctica, enzymatic synthesis of chiral drug intermediates, 69-72 Candida boidinii enzymatic synthesis of chiral drug intermediates, 46-51, 61-63, 66-68, 72 xylose fermentation by, 228 Candida parapsilosis, enzymatic synthesis of chiral drug intermediates, 66-67 Candida pulcherrima, red pigmentation of, 10 Candida shehatae, xylose fermentation by, 228-229, 233-239 Candida tropicalis, xylose fermentation by, 228 Candida utilis, enzymatic synthesis of chiral drug intermediates, 43 Carbazomycinal, biosynthesis of, 143 ccr gene, 98-99 Cephalosporins biosynthesis of, 136, 139, 141 degradation of, 147 structure of, 118-119 Cephalosporium acremonium, bioactive products from, 140 Ceriporiopsis subvermispora, 287 Chiral drug intermediates microbial/enzymatic synthesis of anticholesterol drugs, 57-60, 71-72 antihypertensives, 34-51 antiviral agents, 61-63 [~3-receptor agonists, 52-57, 71 biocatalytic dynamic resolution, 66-68 deoxyspergualin, 60-61, 72 racemic diol stereoinversion, 66-68, 72 racemic epoxide hydrolysis, 63-66 racemic secondary alcohol resolution, 68-72

347

Chitooligosaccharides, purification of, 322-323 Chlamydomonas reinhardtii, lindane degradation by, 287 Chloramphenicel, biosynthesis of, 129-130 Chlorella vulgaris, lindane degradation by, 287 1-Chloroalkane helihydrolase (DhaA), 279 Chlortetracycline biosynthesis of, 122,124, 141 structure of, 123 Chromatography, oligosaccharide assay by, 327-331 Chromobacterium prodigiosum, 22 Clarithromycin, 80-81 Clavibacter spp., ALA2, 212,215-216 Clavulanic acid, biosynthesis of, 118, 120 Clostridium acetobutilicum, xylose fermentation by, 227 Clostridium butyricum, lindane degradation by, 279-280 Clostridium pasteurianum, lindane degradation by, 280 Clostridium rectum, lindane degradation by, 278 Clostridium saccharolyticum, xylose fermentation with, 233 Clostridium spp., lindane degradation by, 279-280 Clostridium thermocellum, xylose fermentation by, 227 Combinatorial biosynthesis, 80 of bioactive products, 137 Saccharopolyspora erythraea, 100-103 Coriolus bulleri, 287 Coriolus hirsutus, 286 Coriolus versicolor, 287 Corynebacterium spp., oleic acid hydration by, 203 Coupling sugar, 301 Cyanobacteria, degradation of lindane by, 287-288 Cyath us bulleri, 286 Cycl gene, 230 Cyclodextrins, 302 Cyclononylprodiginine, 15 Cycloprodigiosin, 14

348

INDEX

Daunorubicin, 125 6-DEB synthase (6-DEBS), 82 de Col, Pietro, 7 DehH1, 279 6-Deoxyerythronolide B (6-DEB), 82 Deoxyspergualin, enzymatic synthesis of chiral drug intermediates, 60-61, 72 Desosamine deoxysugar biosynthesis genes, from Streptomyces venezuelae, 80 DhaA, 279 DhIA, 279 DHOD. See 10,12-Dihydroxy-8(E)-octadecenoic acid Dihydrogranatirhodin, 137 Dihydromederrhodin, 137 7,10-Dihydroxy-8(E)-octadecenoic acid (DOD), biosynthesis, 208-210, 212 lO,12-Dihydroxy-8(E/-octadecenoic acid (DHOD), biosynthesis of, 211-212 Dihydroxy unsaturated fatty acids, biotransformation of, 208-212 Dinoflagellates, phytic acid in, 174-175 Diols, racemic, biocatalytic dynamic resolution of, 66-68, 72 DNA, introduction into Saccharopolyspora erythraea, 87-89 DOD. See 7,10-Dihydroxy-8(E)-octadecenoic acid Doxorubicin, 125 Dyes, red bacteria, 7

Electroporation, DNA introduction into Saccharopolyspora erythraea, 87-89 Environment, phosphorus levels in, 160, 174-175 Enzymatic/microbial synthesis. See Microbial/enzymatic synthesis Epoxide, racemic, biocatalyzed stereoselective hydrolysis of, 63-66 ermE gene, 89-90, 97 ertX gene, 75 eryCI gene, 89-90

eryG gene, 90, 98 Erythromycin, 79 biosynthesis of, 82-84 from Saccharopolyspora erythreae, 85-100, 126, 133 chemical structure of, 81 Erythromycin biosynthetic gene cluster, 80-82, 86, 89-96 Escherichia cob HAP phytase from, 170-171 xylose fermentation by, 227,233 Esterase, enzymatic synthesis of chiral drug intermediates, 55-57 Ethanol biosynthesis of bacteria, 226-227, 233 critical parameters, 232,234-241 ethanol tolerance, 241-255 heat shock proteins, 253-254 Pichia spp., 229-232 thermal tolerance, 241-255 trehalose, 252-253 yeasts and fungi, 226,228-229, 233 uses of, 222 Ethyl-meta-cyclononylprodiginine, 14

FAB mass spectrometry, of oligosaccharides, 332 Factor A, 142 Factor B, 142-143 Factor C, 143 Factor I, 143 Farnesyl pyrophosphate (FPP), 57-58 Fatty acids, unsaturated, biotransformation of, 201-216 Flavobacterium fuscum, enzymatic synthesis of chiral drug intermediates, 43 Ftavobacterium spp. oleic acid hydration by, 205 sp. DS5,210 Foods oligosaccharides in, 303-304,333-337 red spoilage of, 2-6 Formate dehydrogenase, enzymatic synthesis of chiral drug intermediates, 47, 48-49

INDEX FPP. See Farnesyl pyrophosphate Frnctooligosaccharides, 301,309 biosynthesis of, 309-315 properties of, 303 purification of, 322 uses of, 335-336 Fructosyltransferase, 305 Fucosyltransferase, 307-308 Fungi degradation of lindane by, 285-287 galactooligosaccharide biosynthesis by, 316-317 phytase from, 160-170, 183 xylose-fermenting, 226,228-229, 233 Fusarium oxysporum fructooligosaccharide biosynthesis by, 311 xylose fermentation by, 228, 233

G Galactooligosaccharides, 300 biosynthesis of, 315-319 purification of, 322 Galactosides, 315-316, 320 Galactosyltransferase, 307 Gas-liquid chromatography (GLC), oligosaccharide assay by, 329-330 gdh gene, 75 Genetically engineered microorganisms (GEMs), 288-289 Genistein, 135 Gentiooligosaccharides, 302 Geotrichum candidum, enzymatic synthesis of chiral drug intermediates, 58-59, 66, 72 Germicidin, biosynthesis of, 143 GLC. See Gas-liquid chromatography Glucomannan, 224 Glucooligosaccharides, uses of, 335 Glucose dehydrogenase, enzymatic synthesis of chiral drug intermediates, 54 Glutamate dehydrogenase, enzymatic synthesis of chiral drug intermediates, 44-46 Glutathione-dependent reductive dehalogenase (LinD), 278 Glycopeptides, biosynthesis of, 120

349

Glycosidases lipid-coated, 326-327 oligosaccharide biosynthesis and, 306-307,320 Glycosyl sucrose, 301 Glycosyltransferases, 307-308 Gramicidins, biosynthesis of, 115-116 Gramicidin S synthetase, 117

H Haloacetate dehalogenase (DehH1), 279 Haloalkane dehalogenase (DhlA), 279 Hansen ula polymorpha enzymatic synthesis of chiral drug intermediates, 66, 67 phytase production from, 183 HAPs. See Histidine acid phosphatases HCH. See Hexachlorocyclohexane Heat-shock proteins, 253-254 Hexachlorocyclohexane (HCH) biodegradation of by algae and cyanobacteria, 272,277, 287-288 by bacteria, 272,277, 279-285 by fungi, 285-287 environmental contamination by, 272-274 producers of, 271 structure of, 270-271 toxicological effects of, 274-277 High-performance liquid chromatography (HPLC), oligosaccharide assay by, 327-329 Histidine acid phosphatases (HAPs), 161-172 HIV protease inhibitors, enzymatic synthesis of chiral drug intermediates, 61, 72 HMR-3647, 80-81 HOD. See lO-Hydroxy-8-octadecenoic acid HPLC. See High-performance liquid chromatography Hxt gene, 240 Hydrolytic dehalogenases, 278-279 10-Hydroxy fatty acids, biosynthesis of, 207-208

350 Hydroxy fatty acids dihydroxy, 208-212 monohydroxy, 202-207 strain ALA2 system, 212,215-216 trihydroxy, 212-215 uses of, 201 L-6-Hydroxynorleucine, enzymatic synthesis of, 43-46, 71 10-Hydroxy-8-octadecenoic acid (HOD), biosynthesis of, 210 10-Hydroxystearic acid, biosynthesis of, 202-206

Immunosuppressive activity, of prodigiosin, 24 Isomaltooligosaccharides, 302,320 Isomaltulose oligosaccharides, 301 Ivermectin, 128-129

&de gene, 75 Kelp, "red spot disease," 10 1-Kestose, 303,309-310 Ketolides, 80-81 Klehsiella aerogenes phytase, 172-173 Klebsiella oxytoca, xylose fermentation by, 227, 233 Klebsiella terrigena phytase, 172-173 Kluyveromyces fragilis, oligosaccharide biosynthesis by, 319 Kluyveromyces lactis, oligosaccharide biosynthesis by, 319 Konbu, "red spot disease," 10

[~-Lactams antibiotic resistance, 148 biosynthesis of, 118-120 Lactobacillus casei, xylose fermentation by, 227

INDEX

Lactobacillus pentoaceticus, xylose fermentation by, 227 Lactobacillus pentosus, xylose fermentation by, 227 Lactohacillus plantarum, xylose fermentation by, 227 Lactobacillus spp., oligosaccharide biosynthesis by, 318 Lactobacillus xylosus, xylose fermentation by, 227 Lactose, galactoside synthesis using, 320-321 Lactosucrose, 301 Lactulose, 300, 305 Lesquerella hydroxy fatty acids, 201-202 Lignocellulose, 222-224 biorefining of, 222-223 aeration, 238-240 ethanol tolerance, 241-255 heat-shock proteins, 253-254 nutrient uptake, 240-241 p/I, 236-238 pretreatment, 224 simultaneous saccharification and fermentation (SSF), 225-226 temperature, 234-236 thermal tolerance, 241-255 trehalose, 252-253 xylose-fermenting microbes, 226-233 LinB, 279,285 /rinD, 278 Lindane, 270-271 biodegradation of by algae and cyanobacteria, 272,277, 287-288 by bacteria, 272,277, 279-285 by fungi, 285-287 environmental contamination by, 272-274 toxicological effects of, 274-277 Linoleic acid, biocatalyzed hydration of, 207 Linolenic acid, biocatalyzed hydration of, 207 Lipase, enzymatic synthesis of chiral drug intermediates, 60-61, 69, 72 L-Lysine-e-aminotransferase, biotransformation of, 35-37, 39-43

INDEX M

Macrocyclic prodiginines, 15 Macrolides, 80-81, 125-126 biosynthesis of, 125-127 derivatives of, 136 hybrid macrolides, 137 structure of, 126 Maltooligosaccharides, 302 Maxilact, 319 Mederrhodin, 137 MelA enzyme, 96 melA gene, 96-97 Metacycloprodigiosin, 12, 14, 23-24 6-Methoxycarbazomycinal, biosynthesis of, 143 Methyl-(4-methoxyphenyl)-propanedioic acid ethyl diester, racemic, asymmetric hydrolysis of, 55-57 c~-Methyl phenylalanine amide, racemic, enzymatic resolution of, 54-55 6-Methyl salicylic acid (6MS), biosynthesis of, 121-122 6-Methyl salicylic acid synthase, 122 Microalgae, biodegradation of organochlorine compounds by, 272 Microbial/enzymatic synthesis chiral drug intermediates, 33-34, 71-72 anticholesterol drugs, 57-60, 72 antihypertensives, 34-51 antiviral agents, 61-64 [~3-receptor agonists, 52-57, 71 biocatalytic dynamic resolution, 66-68 deoxyspergualin, 60-61, 72 racemic diol stereoinversion, 66-68, 72 racemic epoxide hydrolysis, 63-66 racemic secondary alcohol resolution, 68-72 of ethanol bacteria, 226, 227, 233 critical parameters, 232, 234-241 ethanol tolerance, 241-255 heat shock proteins, 253-254 Pichia spp., 229-232 thermal tolerance, 241-255 trehalose, 252-253 yeasts and fungi, 226, 228-229, 233

351

of oligosaccharides, 299-300, 308-309, 337 enzymatic mechanism, 305-308 fructooligosaccharides, 309-315,322 galactooligosaccharides, 315-319,322 immobilized systems, 324-326 isomaltooligosaccharides, 320 lipid-coated glycosidases, 326-327 novel oligosaccharides, 320-321 organic solvents for, 323-324 purification of, 322-323 recombinants for, 327 two-phase systems, 324 of unsaturated fatty acids, 201-202 dihydroxy, 208-212 monohydroxy, 202-207 strain ALA2 system, 212,215-216 trihydroxy, 212-215 Microbial products, 113-114 Micrococcus prodigiosus, 6, 22 Micromonospora purpurea, secondary metabolite production by, 132 Micromonospora rosaria, secondary metabolite production by, 132 Minocycline, derivatives of, 136 Monas prodigiosa, 6 Monensin, 129 Monohydroxy fatty acids, biotransformation of, 202-207 Mortierella ramanniana, enzymatic synthesis of chiral drug intermediates, 61-63 Mucor sanguineus, 7 Mucor spp., xylose fermentation by, 228 Myceliophthora thermophila, phytase genes, 176 Mycobacterium fortuitum, oleic acid hydration by, 204 Mycobacterium neoaurum, enzymatic synthesis of chiral drug intermediates, 54-55, 71 Myo-inositol phosphate, production of, 187-188

N

Natuphos, 183 Neokestose, 303,305 Neosugar, 305

352

INDEX

Neurospora crassa, xylose fermentation by, 233 NMR analysis, of oligosaccharides, 331-332 Nocardia aurantia, oleic acid hydration by, 204 Nocardia cholesterolicum, fatty acid hydration by, 203,204,207 Nocardia madurae, prodigiosins, 15 Nocardia pelletieri, prodigiosins, 15 Nocardia salmonicolor, enzymatic synthesis of chiral drug intermediates, 64 Norprodigiosin, 12-13 Nostoe ellipsossorum, degradation of lindane by, 288 Nystatin, structure of, 127

in chewing gums, 336 classification of, 300-303 foods, use in, 303-304,333-337 functions of, 332 health benefits of, 304-305 properties of, 303-304 Omapatrilat, microbial/enzymatic synthesis of chiral intermediates, 34, 46 Organochlorine compounds biodegradation of by algae and cyanobacteria, 272,277, 287-288 by bacteria, 272,277, 279-285 by fungi, 285-287 degradation of, 272 toxicological effects of, 274-277 Oxytetracyclines (OTC), biosynthesis of, 122,140, 141

O Occupational health concerns, phytase production, 188-189 Oleandomycin, 126 Oleandrose, 128 Oleic acid biotransformation to 10-HSA, 202-206 hydroxylation of, 207 Oligosaccharides, 299-300,337 applications of, 332-336 assay of, 327 FAB mass spectrometry, 332 GLC, 329-330 HPLC, 327-328 NMR, 331-332 paper chromatography, 329 TLC, 330-331 biosynthesis of, 308-309 enzymatic mechanisms, 305-308 fructooligosaccharides, 309-315,322 galactooligosaccharides, 315-319, 322 immobilized systems, 324-326 isomaltooligosaccharides, 320 lipid-coated glycosidases, 326-327 novel oligosaccharides, 320-321 organic solvents for, 323-324 purification of, 322-323 recombinants for, 327 two-phase systems, 324

Pachysolen tannophilus, xylose fermentation by, 228,229-231, 233-234, 237, 239 Palatinose oligosaccharides, 301 Paper chromatography, oligosaccharide assay by, 329 Paramecium phytase, 174 Penicillin amidases, antibiotic resistance and, 147 Penicillins biosynthesis of, 118-120, 136 degradation of, 147 Penici]lium citrinum, enzymatic synthesis of chiral drug intermediates, 57

Penici]lium frequentans, fructooligosaccharide biosynthesis by, 311 Penicillium patulum, bioactive products from, 121

Penicillius rugulosum, fruct ooligosaccharide biosynthesis by, 311-313 Peniophora lycii phytase, 165, 183 2-Pentanol, biocatalytic dynamic resolution of, 69, 72

INDEX Peptide antibiotics, biosynthesis of, 115-120 Peroxidase, semisynthesis of, 188 Pesticides biodegradation of by algae and cyanobacteria, 272, 277,287-288 by bacteria, 277, 279-285 by fungi, 285-287 Phanerochaete chrysosporium, environmental pollutant degradation by, 286-287 Phanerochaete sordida, environmental pollutant degradation by, 286 Phenylalanine dehydrogenase, enzymatic synthesis of chiral drug intermediates, 46-51 Phenyl galactoside, 320 Phlebia brevispora, 287 Phlebia radiata, 287 pho genes, 171 Phosphorus levels, environmental, 160, 174-175 PhyA, 161-166, 176 phyA gene, 164 PhyB, 166-170, 182 phyB gene, 166 PhyC, 172 Phytase, 158, 189-192 animal feed and, 159-160, 175-177, 183 Bacillus phytase, 172 bioengineering, 175-183,185 E. coli HAP phytase, 170-171 environmental phosphorus levels and, 160, 174-175 enzyme production, 183 enzyme specificity of, 179 functions of, 159 fungal phytase, 160, 183 heat tolerance of, 175-177 histidine acid phosphatases, 161-172 Klebsiella phytase, 172-173 occupational health issues, 188-189 pH optimum of, 177-179 PhyA, 161-166, 176 PhyB, 166-170 PhyC, 172 plants, 171-174, 184-185 substrate specificity of, 179

353

synergistic effect of, 180-183 temperature optimum of, 177-179 in transgenic animals, 185-186 uses of, 159-160, 186-188 yeast phytase, 171,173,183-184 Phytic acid, 159, 175 PHYT I gene, 171 PHYT II gene, 171 Pichia pastor& enzymatic synthesis of chiral drug intermediates, 47-51 phytase production from, 183 Pichia stipitis, xylose fermentation by, 228, 229-231,233-239,249-251 Picromycin, 126 Pithia methanolica, enzymatic synthesis of chiral drug intermediates, 66-68, 72 PKS. See Polyketide synthase Plants phytase from, 171-174, 184-185 phytic acid in, 159 Pleurotus sajor-caju, 286 Pmal gene, 248 Pma2 gene, 248 Polenta, red spoilage of, 5-6 Polyenes, 125-127 Polyethers, bioactivity of, 129 Polyketides, biosynthesis of, 120-129, 132 Polyketide synthase (PKS), 80, 98, 132-133 Polymorphisms, of Saccharopolyspora erythraea, 86-87 Pravastatin, enzymatic synthesis of chiral drug intermediates, 57-60 Prodiginine, 11-12 Prodigiosan, 15 Prodigiosene, 11-12 Prodigiosin, 8, 10-26 antibiotic properties, 22-23 antimalarial activity, 23 biosynthesis of, 15-18 ecological functions, 21-22 immunosuppressive activity, 24 pharmacological activity, 22-24 Serratia spp. producing, 8 structure, 10-15 Prodigiosin 25-C, 13 Protease inhibitors, enzymatic synthesis of chiral drug intermediates, 61, 72

354

INDEX

PsAdhl gene, 230 PsAdh2 gene, 230 Pseudoalteromonas bacteriolytica, "red spot disease," 10

Pseudoalteromonas denitrificans, prodigiosins, 15

Pseudomonas enzymatic synthesis of chiral drug intermediates, 60-61 strain 42A2, 210-211

Pseudomonas magnesiorubra, prodigiosins, 13

Pseudomonas paucimobilis, lindane

Rhodococcus rhodochrous, fatty acid hydration by, 203,207

Rhodotorula glutinis, enzymatic synthesis of chiral drug intermediates, 63-66 Rhodotorula graminis, enzymatic synthesis of chiral drug intermediates, 43 Rice, trihydroxy fatty acids from, 213-214 Ricinoleic acid, 201 Rifamycin, biosynthesis of, 143 Ronozyme P, 183 Rugomonas rubra, prodigiosins, 13

degradation by, 280, 285

Pseudomonas putida, lindane degradation by, 280,285

Pseudomonas spp. lindane degradation by, 280-281 strain PR3,208, 211 PsStul gene, 240 Pulcherrimin, 10 Pyruvate decarboxylase, 230

Saccharomyces cerevisiae bioactive products from, 143 expression of Pichia genes in, 231-232 fermentation by, 226,233,240, 242-248 yeast phytase, 171 Saccharomyces lactis, oligosaccharide biosynthesis by, 319

Saccharopolyspora erythraea R

Racemic diol, biocatalytic dynamic resolution of, 66-68, 72 Racemic epoxide, biocatalyzed stereoselective hydrolysis of, 63-66 Racemic methyl-(4-methoxyphenyl)-propanedioic acid ethyl diester, asymmetric hydrolysis of, 55-57 Racemic c~-methyl phenylalanine amide, enzymatic resolution of, 54-55 Racemic secondary alcohols, biocatalytic dynamic resolution of, 66-68 rap gene, 18 Recombinant oligosaccharides, in-vivo synthesis of, 327 Red bacteria history of, 2-9 naming, 6-10 pigments and paintings, 7-8 "Red diaper syndrome," 19 "Red spot disease," 10 Relomycin, structure of, 126 Response-regulator protein, 141

combinatorial biosynthesis with, 80, 100-103, 137 experimental properties of, 84-85 molecular genetics of, 85-105,133 erythromycin biosynthetic gene cluster, 80, 81-82, 86, 89-96 industrial strain improvement, 97-100 introduction of DNA into, 87-89 mapping, 85-86 new tools for, 96-97 polymorphisms, 86-87 optimization of macrolide production, 98-99 polyketide synthase from, 80

Saccharopolyspora rectivirgula, oligosaccharide biosynthesis by, 319 Salinomycin, 129

Schizosaccharomyces pombe fermentation by, 226,233,252 yeast phytase, 171 Schwanniomyces castellii phytase, 173,176 Schwanniomyces occidenta]is phytase, 173

INDEX

35 5

Scopulariopsis brevicaulis,

Sphingomonas paucimobilis

fructooligosaccharide biosynthesis by, 311 Sebacic acid, 201 Secondary alcohols, biocatalytic dynamic resolution of, 68-70 Secondary metabolites, 114 biosynthesis of bialaphos, 131 chloramphenicol, 129-130 peptide antibiotics, 115-120 polyketides, 120-129, 132 streptomycin, 130-131 derivatives of, 135-137 hybrid bioactive products, 137 isolation of, 133-134 microbial producers of, 131-134 regulation of production, 137-144 resistance to, 144-148 screening for, 135 Sensor-transmitter protein, 141 Serrati, Serafino, 6

enzymatic synthesis of chiral drug intermediates antihypertensives, 35-36, 43 [~3-receptor agonist, 52-56, 71 lindane degradation by, 278,281,283 Squalene synthase, enzymatic synthesis of chiral drug intermediates, 58-60 SSF. See Simultaneous saccharification and fermentation

Serratia ficaria, 8 Serratia liquefaciens, 8, 21 Serratia marcescens, 25 antibiotic resistance, 20-21 cell-surface hydrophobicity, 22 prodigiosin, 8, 10, 15-16, 18, 21-22 Serratia marcescens infection, 19-21

Serratia marinorubra, 8 Serratia odorifera, 8 Serratia plymuthica, 8, 22 Serratia rubidaea, 8 Serratia spp., 8-10 color variations in, 22 prodigiosin, 8, 10-26 Simultaneous saccharification and fermentation (SSF), 225-226 Soil amendments, phytase as, 187 Sonication-dependent electroporation, DNA introduction into Saccharopolyspora erythraea, 87-88 Soybean oligosaccharides, 302-303,305 Spergualin, enzymatic synthesis of chiral drug intermediates, 60-61, 72 Sphingobacterium spp., oleic acid hydration by, 205

Streptococcus thermophilus, oligosaccharide biosynthesis by, 318

Streptomyces aureofaciens, bioactive products from, 122,139, 141,144, 146-147 Streptomyces avermectilis, bioactive products from, 143 Streptomyces carbophilus, enzymatic synthesis of chiral drug intermediates, 57 Streptomyces cinnamonensis, bioactive products from, 143 Streptomyces clavuligerus, bioactive products from, 118, 140 Streptomyces coelicolor, prodigiosins, 16, 18 Streptomyces collinus, ccr gene, 98-99 Streptomyces fradiae, bioactive products from, 127,144 Streptomyces galilaeus, combinatorial biosynthesis with, 137 Streptomyces griseus, bioactive products from, 128,142-143

Streptomyces hiroshimensis, prodigiosins, 14

Streptomyces hydroscopicus, bioactive products from, 131

Streptomyces lividans bioengineering, 145-146 enzymatic synthesis of chiral drug intermediates, 39 Streptomyces longisporus ruber, prodigiosins, 13-14 Streptomyces nodosus, enzymatic synthesis of chiral drug intermediates, 61-63 Streptomyces noursei, enzymatic synthesis of chiral drug intermediates, 35, 37-39, 71

356

INDEX

Streptomyces parvulus, bioactive products from, 141

Streptomyces purpurascens, combinatorial biosynthesis with, 137

Streptomyces rimosus bioactive products from, 122,140-141 bioengineering, 146 Streptomyces rubriretuculi var. pimprina, prodigiosins, 14 Streptomyces spp. bioactive products biosynthesis of, 115-131 derivatives of, 133-137 molecular genetics of, 131-133 regulation of production, 137-144 resistance to, 144-147 bioactive products from, 125 growth phases of, 137-138

Streptomyees venezuelae bioactive products from, 143 chloramphenicol biosynthesis from, 129 desosamine deoxysugar biosynthesis genes, 80

Streptomyces viridochromogenes, bioactive products from, 131, 143 Streptomycin, biosynthesis of, 130-131 Streptorubin A, 14

StreptoverticiHium baldaccii, prodigiosins, 14

Streptoverticillium spp., bioactive products from, 143

Sulfiblobus solfataricus, 321 Syntropic pigmentation, 16

Talaromyces lanuginosus phytase, 176 Talaromyces thermophi]is, phytase genes, 176 Tetrachlorocyclohexadiene dehalogenase (LinB), 279 Tetracyclines, 125 biosynthesis of, 122-125, 140-141 structure of, 122-123

Thermoactinomyces intermedius, enzymatic synthesis of chiral drug intermediates, 46-48, 51, 71

Thin-layer chromatography (TLC), oligosaccharide assay by, 330-331 THOA. See 9,12,13-Trihydroxy10(E}-octadecenoic acid Tigemonam, chiral intermediate of, 51 Tk! genes, 232 Tkt genes, 232 TLC. See Thin-layer chromatography TOD. See 7,10,12-Trihydroxy-8(E}octadecenoic acid Trametes hirsutus, 286-287 Transcriptional terminator cartridge, in Saccharopolysporo erythraea, 91-92 Transgenic animals, phytase, 185-186 Transgenic plants, phytase, 185 Trehalose, 252-253 Trichoderma hardanum, oligosaccharide biosynthesis by, 317 Trigonopsis variabilis, enzymatic synthesis of chiral drug intermediates, 44, 46 8,9,13-Trihydroxy docosanoic acid, biosynthesis of, 212 7,10,12-Trihydroxy-8(E)-octadecenoic acid (TOD), biosynthesis of, 211 9,12,13-Trihydroxy-10(E)-octadecenoic acid (THOA), biosynthesis of, 212-215 12,13,17-Trihydroxy-(Z)-octadecenoic acid, biosynthesis of, 212 Trihydroxy unsaturated fatty acids, biotransformation of, 212-215 Tylosin biosynthesis of, 127 structure of, 126

U Undecylprodiginine, 12, 16, 18, 23-24 Undecylprodigiosin, 12-13 Unsaturated fatty acids, 201-202 dihydroxy, 208-212 monohydroxy, 202-207 strain ALA2 system, 212,215-216 trihydroxy, 212-215 uses of, 201

INDEX

357 X

V Vancomycin biosynthesis of, 120 structure of, 121 Vasopeptidase inhibitor enzymatic synthesis of, 71 allysine ethylene acetal, 46-51, 71 BMS-199541-01, 34-43 L-6-hydroxynorleucine, 43-46 vhb gene, introduction into Saccharopolyspora erythraea, 99-100 Vibrio gazogenes, prodigiosins, 15 Vibrio psychroerythreus, prodigiosins, 13 Vitreoscilla hemoglobin gene, introduction into Saccharopolyspora erythraea, 99-100

W

Water pollution, by organochlorine pesticides, 273-274 White-rot fungi, biodegradation of organochlorine compounds by, 272, 286-287

Xylitol dehydrogenase, 230 Xylooligosaccharides, 303 Xylose, 222-223 bioconversion of bacteria, 226-227,233 critical parameters, 232,234-241 ethanol tolerance, 241-255 heat shock proteins, 253-254 Pichia spp., 229-232 thermal tolerance, 241-255 trehalose, 252-253 yeasts and fungi, 226, 228-229, 233 lignocellulose, 223-226 Y Yeast phytase, 171,173 Yeasts oligosaccharide biosynthesis by, 319 xylose-fermenting, 226, 228-229,233 Z

Zymomonas mobilis, xylose fermentation by, 227,233

CONTENTS OF PREVIOUS VOLUMES

Volume 37

Microbial Degradation of the Nitroaromatic Compounds Frank K. Higson An Evaluation of Bacterial Standards and Disinfection Practices Used for the Assessment and Treatment of Stormwater Marie L. O'Shea and Richard Field Haloperoxidases: Their Properties and Their Use in Organic Synthesis M. C. R. Franssen and H. C. van der Plas Medicinal Benefits of the Mushroom Ganoderma S. C. Jong and J. M. Birmingham Microbial Degradation of Biphenyl and Its Derivatives Frank K. Higson The Sensitivities of Biocatalysts to Hydrodynamic Shear Stress Ale~ Prokop and Rakesh K. Bajpai Biopotentialities of the Basidiomacromycetes Somasundaram Rajarathnam, Mysore Nanjarajurs Shashirekha, and Zakia Bano INDEX 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, and 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 P. Lillehoj and Vedpal S. Malik Anoxygenic Phototrophic Bacteria: Physiology and Advances in Hydrogen Production Technology K. Sasikala, Ch. V. Ramana, R Rahuveer Rao, and K. L. Kovacs INDEX

Volume 39

Asepsis in Bioreactors M. C. Sharma and A. K. Gartu Lipids of n-Alkane-UtilizingMicroorganisms and Their Application Potential Samir S. Radwan and Naser A. Sorkhoh Microbial Pentose Utilization Prashant Mishra and Ajay 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 Hallaert, and Erick J. Vandamme 359

360

CONTENTS OF PREVIOUS VOLUMES

Physicochemical and Biological Treatments for Enzymatic/Microbial Conversion of Lignocellulosic Biomass Purnendu Ghosh and Ajay Singh

Volume 41

INDEX

Improving Productivity of Heterologous Proteins in Recombinant Saccharomyces cerevisiae Fermentations Amit Vasavada

Volume 40

Microbial Cellulases: Protein Architecture, Molecular Properties, and Biosynthesis Ajay Singh and K3"yoshi 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 Ale~ Prokop Challenges in Commercial Biotechnology. Part II. Product, Process, and Market Development Ale~ 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

Microbial Oxidation of Unsaturated Fatty Acids Ching T. Hou

Manipulations of Catabolic Genes for the Degradation and Detoxification of Xenobiotics Rup Lal, Sukanya Lal, P. S. Dhanaraj, and D. M. Saxena Aqueous Two-Phase Extraction for Downstream Processing of Enzymes/Proteins K. S. M. S. Raghava Rao, N. K. Rastogi, M. K. Gowthaman, and N. G. Karanth 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 II. Biopolyesters, Biopesticide, Biofuel, and Biofertilizer Ch. Sasikala and Ch. V. Ramana INDEX

Detection, Isolation, and Stability of Megaplasmic-Encoded Chloroaromatic Herbicide-Degrading Genes within Pseudomonas Species Douglas J. Cork and Amjad Khalil INDEX

Volume 42

The Insecticidal Proteins of Bacillus th uringiensis P. Ananda Kumar, R. P. Sharma, and V S. Malik

CONTENTS OF PREVIOUS VOLUMES Microbiological Production of Lactic Acid

361

Volume 44

John H. Litchfield

Biodegradable Polyesters

Biologically Active Fungal Metabolites Cedric Pearce

Ch. Sasikala

The Utility of Strains of Morphological Group II Bacillus Samuel Singer

Old and New Synthetic Capacities of Baker's Yeast P. D'Arrigo, G. Pedrocchi-Fantoni, and S. Servi

Phytase Rudy J. Wodzinski and A. H. J. Ullah INDEX

Investigation of the Carbon- and Sulfur-Oxidizing Capabilities of Microorganisms by Active-Site Modeling Herbert L. Holland

Volume 43

Production of Acetic Acid by Clostridium thermoaceticum Munir Cheryan, Sarad Parekh, Minish Shah, and Kusuma Witjitra

Contact Lenses, Disinfectants, and Acanthamoeba Keratitis Donald G. Ahearn and Manal M. Gabriel

Marine Microorganisms as a Source of New Natural Products V. S. Bernan, M. Greenstein, and W. M. Maiese

Stereoselective Biotransformations in Synthesis of Some Pharmaceutical Intermediates Ramesh N. Patel

Microbial Xylanolytic Enzyme System: Properties and Applications Pratima Bajpai

Oleaginous Microorganisms: An Assessment of the Potential Jacek Leman INDEX

Microbial Synthesis of D-Ribose: Metabolic Deregulation and Fermentation Process R de Wulf and E. J. Vandamme

Production and Application of Tannin Acyl Hydrolase: State of the Art B K. Lekha and B. K. Lonsane

Ethanol Production from Agricultural Biomass Substrates Rodney /. Botbast and Bada] C. Saha

Thermal Processing of Foods, A Retrospective, Part I: Uncertainties in Thermal Processing and Statistical Analysis M. N. Ramesh, S. G. Prapulla, M. A. Kumar, and M. Mahadevaiah

Thermal Processing of Foods, A Retrospective, Part II: On-Line Methods for Ensuring Commercial Sterility M. N. Ramesh, M. A. Kumar, S. G. Prapulla, and M. Mahadevaiah INDEX

362

CONTENTS OF PREVIOUS VOLUMES

Volume 45

One Gene to Whole Pathway: The Role of Norsolorinic Acid in Aflatoxin Research J. W. Bennett, P.-K. Chang, and D. Bhatnagar Formation of Flavor Compounds in Cheese P. F Fox and J. M. Wallace

Breathing Manganese and Iron: Solid-State Respiration Kenneth H. Nealson and Brenda Little Enzymatic Deinking Pratima Bajpai Microbial Production of Docosahexaenoic Acid (DHA, C22:6) Ajay Singh and Owen P. Ward

The Role of Microorganisms in Soy Sauce Production Desmond K. O'Toole

INDEX

Gene Transfer Among Bacteria in Natural Environments Xiaoming }Tin and G. Stotzky

Volume 46

Cumulative Subject Index