Biology - Chemistry Interface A Tribute to Koji Nakanishi The [1 ed.] 9780824771164, 0-8247-7116-8

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The Bio Iogy-C he mistry Interface

ATribute to Koji Nakanishi

edited by

Raymond Cooper

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Pharmanex, Inc. San Francisco, California

John K. Snyder Boston University Boston, Massachusetts

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MARCEL

MARCELDEKKER, INC. D E K K E R

NEWYORK BASEL

ISBN: 0-8247-7116-8 This book is printed on acid-free paper. Headquarters Marcel Dekker, Inc. 270 Madison Avenue, New York, NY 10016 tel: 212-696-9000; fax: 212-685-4540 Eastern Hemisphere Distribution Marcel Dekker AG Hutgasse 4, Postfach 812, CH-4001 Basel, Switzerland tel: 41-61-261-8482; fax: 41-61-261-8896 World Wide Web http:/ /www.dekker.com The publisher offers discounts on this book when ordered in bulk quantities. For more information, write to Special Sales/Professional Marketing at the headquarters address above. Copyright  1999 by Marcel Dekker, Inc. All Rights Reserved. Neither this book nor any part may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, microfilming, and recording, or by any information storage and retrieval system, without permission in writing from the publisher. Current printing (last digit): 10 9 8 7 6 5 4 3 2 1 PRINTED IN THE UNITED STATES OF AMERICA

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Preface

Natural products science, a fascinating cornerstone of modern research, has long bridged the traditional frontier between chemistry and biology. Humankind has always been intrigued by the power and potential of plants and nature. Many old texts reveal how the ancient cultures drew on the beneficial properties of plants. They learned the wisdom of extracting the ingredients and using such potions as foods, medicines, and mood enhancers long before anyone understood how these worked. Slowly we have found the tools to explore the chemistry of these ingredients, and thus the systematic study of natural products began. Morphine was isolated in 1805 and strychnine in 1819, although their structures remained mysteries for more than 100 years, and pure camphor has been an article of commerce for centuries. Today the biosynthetic machinery of plants and other organisms is purposefully manipulated to produce new ‘‘natural products’’ of biological significance in medicine and agriculture. In the nineteenth century, early progress in natural products research centered on the study of pigments from flowers as colored dyes. Originally, extraction of drug compounds, particularly alkaloids, from plants was achieved by using simple isolation methods: a water steep or a solvent (generally alcohol) extraction. The impetus was set to explore this research area further and, not surprisingly, more and more intellectual pursuit of natural sciences and our environment encouraged universities and scientific centers throughout the world to study natural products, which then formed the nucleus of chemistry programs. As source material to begin any research investigation, plants were abundant and easily obtained. The first natural products to be studied in detail were generally the major constituents of plants, since these often precipitated from solution and could be purified through recrystallization. How well do we recall

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today that the purity of chemicals up to the latter half of the twentieth century was determined solely by melting point? As increasing numbers of chemical constituents with more complexity were found, structural analysis relied on chemical transformations and degradation studies. Total synthesis was confirmatory. The discovery of one or two compounds based on these research studies was usually acceptable to earn a doctorate. The second half of the twentieth century has witnessed incredible advances in natural products research. These have been achieved through the discovery of new chromatographic separation methods and remarkable advances in spectroscopy. As new technologies for isolation and structure identification have evolved, the isolation and detection of ever-diminishing amounts of natural products, coupled with the determination of structures on a microscale, have become almost routine. In addition to identifying important targets for total synthesis, and thereby spurring innumerable advances in fundamental organic chemistry, studies of natural products have led to significant research efforts in the related fields of bioorganic chemistry and biosynthesis, as chemists, biologists, and biochemists have striven to understand how these molecules are produced in nature and to establish the molecular basis of the biological activity of these compounds. The structural determination of natural products has impacted our basic understanding of nature. One very important aspect of structure determination is the use of spectroscopy, particularly nuclear magnetic resonance, mass spectrometry, circular dichroism, and x-ray diffraction methods. Circular dichroism is particularly important in establishing absolute stereochemistry, as chirality is correlated directly to biological activity of the biomolecule. Thus, as we approach the end of this century, we see the challenging questions in biology requiring answers at the molecular level being met by increasingly sophisticated techniques and comprehension of the chemistry of nature. Professor Koji Nakanishi has been a pioneer and a towering figure in natural products research. He has been a major contributor at the crossroads of bioorganic chemistry over the past 40 years. His extraordinary and broad vision of natural products chemistry and its close relationship to bioorganic studies is now universally accepted and was the inspiration for this book. He has constantly looked at challenges in bioorganic chemistry and pushed ever closer the boundaries at the interface between chemistry and biology. He has achieved this through his lifelong studies in natural products, his investigations into the chemistry of vision, his pursuit of new and ever more powerful analytical and spectroscopic microtechniques for solving complex structural problems, and his study of infrared and circular dichroism and their applications to bioorganic science. Koji’s curiosity and insights in applying the right solution to challenging problems are among his legacies, to which we as students of his are deeply grateful. Koji Nakanishi was born in 1925 in Hong Kong to parents of Japanese

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descent. As a result of his father’s business postings abroad, Koji’s early childhood was spent in various European capitals as well as in Alexandria, Egypt, thereby giving birth to and nurturing his unique world vision. He returned to Japan for his formal university training and received his B.Sc. degree at Nagoya University in 1947. He first came to the United States in 1950–52 to study with the legendary Professor Louis Fieser at Harvard University, and he returned to Japan as an assistant professor to embark on his remarkable career in natural products and bioorganic chemistry. He completed his Ph.D. in 1954 under the mentorship of Professors Egami and Hirata, then took positions at Nagoya University (1955–58), Tokyo Kyoika University (1958–63), and Tohoku University (1963–69). In 1969 he was invited to join the faculty at Columbia University, New York, where he currently holds the chair of ‘‘Centennial Professor of Chemistry.’’ Indeed, it was at Columbia University that former and current students, postdoctoral fellows, and esteemed colleagues of Professor Nakanishi gathered to celebrate his 70th birthday and to honor him for his years of mentorship and friendship, as well as for his considerable contributions to bioorganic science. Two days of stimulating presentations on various topics in bioorganic chemistry gave birth to the idea behind this book: to produce a volume with contributed chapters from his former students in his honor. This text reflects Koji’s own research interests in its scope and attempts to bridge the gap between biology and chemistry: a gap that is rapidly diminishing as investigators use the tools and vision that Koji has provided. Koji humbly reminds us that he is ‘‘only a technician’’; we respectfully differ. He is a visionary, and in essence the investigatory seeds planted by Koji are now in full bloom in research gardens headed by those he taught. We choose to highlight current research activities from former members of his research groups from Asia, the United States, Europe, and Australia, thereby illustrating Koji’s global scientific influence. As with Koji’s own research, one goal of this text is to further dissolve the boundary that has kept chemistry and biology apart; the contributions in this volume are by investigators for whom this boundary has long since disappeared. It is hoped that readers will come to understand the highly interactive nature of research in biological chemistry and chemical biology, and find the transition between chemistry and biology far less intimidating. Thus, this book reflects the ideals of Professor Nakanishi and his impact. Although the chapter titles may at first glance seem to suggest a relatively large breadth of subjects, in fact they all fit snugly within the focus of the chemical basis of biological activity. Subjects range from hydrolytic enzymes to combinatorial chemistry, yet all the chapters strive to elucidate biological responses at the molecular level. The contributing authors provide detailed accounts of their current research rather than presenting formal reviews of disparate subjects. The

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rationale for this approach is to emphasize the interactive nature of the research in bioorganic chemistry. The unifying theme throughout is the original skills that developed in natural products research and chemistry of vision. The microanalytical techiques, the spectroscopic challenges, have now evolved into the application of chemical minds to biological problems. Thus, the selection of authors reflects a blend of investigations in academic and industrial research. Koji’s pioneering contributions and world vision of science have inspired several generations of chemists from around the globe, and demonstrations of his mastery of the magical arts have left numerous audiences of the brightest minds completely and delightfully mystified. We can identify the defining moment of our education and scientific growth as the time we spent with Koji, and we offer our profoundest gratitude to him for his tireless leadership and support. Raymond Cooper John K. Snyder

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Contents

Preface Contributors Tribute Letters 1. Insect Antifeeding Limonoids from the Chinaberry Tree Melia azedarach Linn. and Related Compounds Munehiro Nakatani 2. Polygodial and Warburganal, Antifungal Sesquiterpene Dialdehydes and Their Synergists Isao Kubo 3. Marine Bromoperoxidases—Chemoenzymatic Applications Chris A. Moore and Roy K. Okuda 4. LC-Hyphenated Techniques in the Search for New Bioactive Plant Constituents Kurt Hostett aryse Hostettmann, Sylvain Rodriguez, and Jean-Luc Wolfender 5. Determination of the Absolute Configuration of Biologically Active Compounds by the Modified Mosher’s Method Takenori Kusumi and Ikuko I. Ohtani 6. Circular Dichroism Spectroscopy and the Absolute Stereochemistry of Biologically Active Compounds Nobuyuki Harada

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7. Recent Applications of Circular Dichroism to Carbohydrate Conformational Analysis and Direct Determination of Drug Levels Jesu´s Trujillo Va´zquez 8. Furan-Terminated Cationic π-Cyclizations in the Synthesis of Natural Products Steven P. Tanis 9. Chemistry and Biology of Semisynthetic Avermectins Timothy A. Blizzard

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10.

Chemical and Biological Approaches to Molecular Diversity: Applications to Drug Discovery Harold V. Meyers

11.

Imidazoline Receptors and Their Endogenous Ligands Colin J. Barrow and Ian F. Musgrave

12.

Oxidoredox Suppression of Fungal Infections by Novel Pharmacophores Valeria Balogh-Nair

13.

A Mechanistic Analysis of C—O Bond Cleavage Events with a Comparison to 3,6-Dideoxysugar Formation David A. Johnson and Hung-wen Liu

14.

The Molecular Mechanism of Amyloidosis in Alzheimer’s Disease Michael G. Zagorski

15.

Bacteriorhodopsin Structure/Function Studies: Use of the Demethyl Retinal Analogues for Probing of the Arg82Ala Mutant Rosalie K. Crouch, Donald R. Menick, Yan Feng, Rajni ee, and Thomas G. Ebrey

16.

Autonomous Genomes David G. Lynn

17.

Stereochemical Considerations of Immunoglobulin Heavy Chain Enhancer Activation Barbara S. Nikolajczyk and Ranjan Sen

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Contributors

Valeria Balogh-Nair Department of Chemistry, The City College of the City University, New York, New York Colin J. Barrow School of Chemistry, The University of Melbourne, Parkville, Victoria, Australia Timothy A. Blizzard Medicinal Chemistry, Merck Research Laboratories, Rahway, New Jersey Rosalie K. Crouch Department of Ophthalmology, Medical University of South Carolina, Charleston, South Carolina Thomas G. Ebrey School of Cellular and Molecular Biology, University of Illinois at Urbana-Champaign, Urbana, Illinois Yan Feng Department of Ophthalmology, Medical University of South Carolina, Charleston, South Carolina Rajni Govindjee Center for Biophysics and Computational Biology and Department of Cell and Structural Biology, University of Illinois at Urbana-Champaign, Urbana, Illinois

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Contributors

Nobuyuki Harada Institute for Chemical Reaction Science, Tohoku University, Sendai, Japan Kurt Hostettmann Institut de Pharmacognosie et Phytochimie, Universite´ de Lausanne, Lausanne, Switzerland Maryse Hostettmann Institut de Pharmacognosie et Phytochimie, Universite´ de Lausanne, Lausanne, Switzerland Department of Chemistry, University of Minnesota, Minne-

David A. Johnson apolis, Minnesota

Isao Kubo Department of Environmental Science, Policy, and Management, University of California, Berkeley, California Takenori Kusumi Faculty of Pharmaceutical Sciences, Tokushima University, Tokushima, Japan Hung-wen Liu Department of Chemistry, University of Minnesota, Minneapolis, Minnesota David G. Lynn cago, Illinois

Department of Chemistry, The University of Chicago, Chi-

Donald R. Menick Departments of Medicine, and Biochemistry and Molecular Biology, Medical University of South Carolina, Charleston, South Carolina Harold V. Meyers Chemistry and Drug Discovery Group, New Chemical Entities, Inc., Framingham, Massachusetts Chris A. Moore California

Department of Chemistry, San Jose´ State University, San Jose´,

Ian F. Musgrave toria, Australia

Prince Henry’s Institute for Medical Research, Clayton, Vic-

Munehiro Nakatani Department of Chemistry and Bioscience, Kagoshima University, Kagoshima, Japan

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Barbara S. Nikolajczyk* Brandeis University, Waltham, Massachusetts Ikuko I. Ohtani Department of Chemistry, Biology, and Marine Science, University of the Ryukyus, Okinawa, Japan Roy K. Okuda Department of Chemistry, San Jose´ State University, San Jose´, California Sylvain Rodriguez Institut de Pharmacognosie et Phytochimie, Universite´ de Lausanne, Lausanne, Switzerland Ranjan Sen Rosenstiel Research Center and Department of Biology, Brandeis University, Waltham, Massachusetts Steven P. Tanis Medicinal Chemistry I, Pharmacia & Upjohn, Inc., Kalamazoo, Michigan Jesu´s Trujillo Va´zquez Instituto Universitario de Bio-Orga´nica ‘‘Antonio Gonza´lez,’’ Universidad de La Laguna, Tenerife, Spain Jean-Luc Wolfender Institut de Pharmacognosie et Phytochimie, Universite´ de Lausanne, Lausanne, Switzerland Michael G. Zagorski Department of Chemistry, Case Western Reserve University, Cleveland, Ohio

* Current affiliation: Immunobiology Unit, Departments of Medicine and Microbiology, Boston University Medical School, Boston, Massachusetts.

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Tribute Letters

March 16, 1998 Dear Koji, On the occasion of your 70th birthday, many of your old friends and colleagues came to Columbia for a wonderful celebration in 1995. Now we are assembling a permanent record of our appreciation for your friendship and as a tribute to your many elegant and important contributions to the chemistry and biology of natural products. I can remember our first meeting, 34 years ago, in Tokyo, at the Presymposium to the IUPAC meeting in Kyoto. In the next two weeks you were our host almost every evening and introduced us to Japanese food and customs. It was a great awakening to the realization that Japanese chemistry was rapidly gaining tremendous momentum and a turning point in my career. Since that meeting I have had the pleasure and privilege of working with 26 Japanese colleagues (several of whom came from your lab). It was a pleasure to repay a little of your hospitality when you visited our homes in Sussex, New Haven and College Station, and you know that you and your wife are always welcome in Texas. You are a true pioneer in solving different problems at the chemistry– biology interface using every possible technique on vanishingly small amounts of material and your work continues to be an inspiration to all of us. Most importantly your personal qualities have ensured a permanent legacy in your many students who have done so well in our profession. It must make you feel very

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proud to have had so many loyal and dedicated coworkers. Above all (and almost uniquely in our field) you have remained a gentleman, with the highest ethical standards in dealing with your colleagues. I don’t know how you manage to work so hard yet still find time for your magic and your friends. I can guess the secret of your success in chemistry and life—that you are fortunate like myself, to have such a long and happy marriage. Betty joins with me in wishing you and your wife continued health, happiness and success for many more birthdays to come. As always! Yours very sincerely,

A. I. Scott, F.R.S. Davidson Professor of Science Director of Center for Biological NMR Texas A&M University, College Station, Texas

March 8, 1998 Dear Koji, ‘‘They’’ never stop celebrating you! ‘‘They,’’ of course, are those who have had the privilege to obtain from you, as post-docs or as Ph.D. students, part of their life baggage. They have been also kind enough to associate to them some of your long time friends, and it was indeed a great pleasure to have the opportunity to pay tribute to you in Columbia nearly half a century after we had first met in the basement of Converse Laboratory, at Harvard, in Louis Fieser’s group. When I was invited to contribute to this volume with a letter, I tried to call back the oldest memories of our meetings I could muster. For some odd reason, even though I am neither a gourmet nor a gourmand, they were nearly all memories of food. The experience of learning from you (and from Huang Wey Yuan) to use chopsticks (a very useful lesson), the dinners of frog1 or lamb2 legs in 1

My wife was working at Harvard Medical School in Pharmacology with Fieser’s friend Prof. O. Krayer, on the action of the Veratrum alkaloids on frog heart. A frog: one heart, two legs. We always had a few dozen frozen legs aside for our friends. These legs, and frozen guinea pigs (one heart, one guinea pig), helped us survive on our starvation scholarships. 2 On affluent months, for a change from frogs and guinea pigs, I was buying lamb by the half at the

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the stable-boy’s rooms of the mock-French castle I was living in with Paula in Brookline.3 Food apart, another very old memory I can retrieve is that of the innumerable small sealed tubes in which you were desperately heating pristimerin with something (was it zinc or selenium?), to find its structure. Food and pristimerin apart, I also revive with a little nostalgy our outing to White Mountains, to a mountain the name of which escapes me, when we had to climb very large oblique stone slabs and you lost grip at the top, to slide down slowly at first, then quickly, on your fingers and stomach, to arrive at the bottom half skinned. But, food and pristimerin and outing apart, it is also then that I first became one of your favorite stooges, always ready to serve on any available stage, many, many times later, as a faire-valoir to your other profession. A simpleton glad to oblige. Koji, I have only one regret: that I could never find a pretext to share some (serious) work with you, in Japan, in Nairobi or in New York, even though we have both always held the same conviction that chemistry and biology share more than one border and that to follow fashion is silly when there is so much else to explore. Merci, Koji, pour ton amitie´

Professor Guy Ourisson Vice President de l’Acade´mie des Sciences Strasbourg, France

Italian market. The subway passengers had to sit next to a very un-American young man carrying half a lamb protruding from his rucksack. 3 We were living in the small rooms above the stables built by a former U.S. Ambassador to Italy and to Japan, in the middle of the huge estate he had bequeathed to the township of Brookline, Lars Andersen Park. The stables were in the form of the Chaˆteau de Chambord, or nearly so, and were the seat of the Veteran Motor Cars Association of America, the ‘‘vie de chaˆteau,’’ which we shared with some 100 old cars.

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1 Insect Antifeeding Limonoids from the Chinaberry Tree Melia azedarach Linn. and Related Compounds Munehiro Nakatani Kagoshima University, Kagoshima, Japan

I. INTRODUCTION Limonoids are tetranortriterpenoids derived from euphane (H-20β) or tirucallane (H-20α) triterpenoids with a 4,4,8-trimethyl-17-furanylsteroidal skeleton [1]. Over 300 limonoids have been isolated to date, and they are the most distinctive secondary metabolites of the plants in the order Rutales. Particularly, they characterize members of the family Meliaceae, where they are abundant and varied [2– 4]. Almost every part of the trees of this family has been used in folkloric and traditional systems of medicine [5,6]. Recent work has established a wide range of biological activities for these compounds, including insect antifeedant and growth-regulating properties, a variety of medicinal effects in animals and humans, and antifungal, bacteriocidal, and antiviral activities. The biological activities of limonoids from Rutales have been reviewed [7]. In particular, the limonoids from the neem tree Melia azadirachta indica Juss and the Chinaberry tree Melia azedarach Linn. have attracted considerable interest because of their marked insect antifeedant properties and intriguing structural variety. The most potent insect antifeedants are azadirachtin and related highly oxidized C-seco limonoids from M. azadirachta. Their antifeedant activities and structure–activity relationships have been reviewed [8–10].

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Nakatani

Similarly, Melia azedarach is known to produce azadirachtin-type limonoids that are potent antifeedants such as the meliacarpinins, and C-19/C-29 bridged lactols and acyl acetals such as the azedarachins, and trichilins. We have investigated the limonoid constituents of M. azedarach and some related plants and evaluated the antifeedant properties of more than 30 limonoids in this work, which includes all of the types of limonoids isolated from M. azedarach excluding glycosides. The structures and antifeedant activities of these compounds are detailed here. II. LIMONOIDS FROM Melia azedarach Linn. Melia azedarach is a native of Persia, India, and China but is naturalized in a number of continents including Africa, Australia, and the Americas. Thus, the constituents of this tree from many regions have been studied. Limonoids may be found in all tissues of the plant, but different organs within an individual may produce different types of limonoids. Since the initial isolation of gedunin (29), nimbolin A (9), and nimbolin B (32) from the trunk wood, reported in 1969 [11], about 55 limonoids have been isolated from the fruits, stem bark, and root bark to date. A variety of oxidation and skeletal rearrangements of the basic limonoid skeleton (meliacane) are found (Figs. 1–4). A characteristic oxidation of the skeleton that is found in M. azedarach produces C-19/C-29 bridged acyl acetals with a 14,15-epoxide as in the azedarachins (10–16, Fig. 2) and trichilins (17–23, Fig. 2), or their D-ring keto compounds (24–28, Fig. 2). In addition, spirosendan (35, Fig. 3), recently isolated by us, is a novel spiro limonoid possessing a C-12/C30 bridged system, the first such limonoid substructure found in nature, to the best of our knowledge [12a]. In limonoids of other origins, the A- or D-rings commonly is oxidized to lactones (A- or D-seco limonoids), in the latter case presumably by introduction of a carbonyl function at C-16 followed by a BaeyerVilliger-type oxidation. But these derivatives are rarely found in M. azedarach, with the sole exceptions to date being gedunin (29) and the related 3-glucoside (30). Many limonoids possess a 14,15-epoxide. Furthermore, the C-ring is fre-

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Figure 1 Limonoids from M. azedarach, Group 1: apo-euphol limonoids.

quently oxidatively cleaved to a carboxylic acid or lactone (C-seco limonoids). The C-seco limonoids, which are major ring-seco limonoids of M. azedarach, M. azadirachta, and M. toosendan in the family Meliaceae and are found only in these three related species, may occur by one of several mechanisms, and there is controversy about the mechanism by which the C-12/C-13 bond of the C-ring is opened [1,13,14]. The series of sendanal (8), ohchinal (31), nimbolidins A (43), and B (44), and nimbolin B (32) illustrates one possible process involved, which includes introduction of an oxygen function at C-12, followed by oxidation to a ketone. Cleavage of the C-12/C-13 bond is accompanied by the simultaneous opening of a 14,15-epoxide to generate the CHO-12 and allylic 15-OH functions

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Nakatani

Figure 2 Limonoids from M. azedarach, Group 1: apo-euphol limonoids (continued ), and Group 2: D-seco limonoids.

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Figure 3 Limonoids from M. azedarach, Group 3: C-seco limonoids.

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Nakatani

Figure 4 Limonoids from M. azedarach, Group 3: C-seco limonoids, highly oxidized compounds.

Scheme 1

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(Scheme 1, Route 1) as found in (31). Rotation about the C-8/C-14 bond would allow the 15-OH to recyclize with CHO-12 to form a lactol C-ring as in (32). Alternatively, an electron transfer process may generate a radical cation that subsequently opens to an allylic radical with a remote C-12 acylium-type ion intermediate. Capture of the allyl radical by a neighboring C-7 oxygen substituent would lead to C-seco derivatives with the C-7/C-15 oxo bridge such as (31) (Scheme 1, Route 2). The C-12 keto limonoid nimbidinin found in the seeds of Azadirachta indica [15] may serve as a precursor for this latter route. Meliacarpinin A (46, Fig. 4), one of the most potent insect antifeedants, was isolated from the root bark of Okinawan M. azedarach in 1993 [16]. This limonoid and the related meliacarpinins (47–54) are more highly oxidized natural products of the C-seco class similar to the azedarachins. 1-Cinnamoylmelianolone (45), found in the fruits of the American variety, may be a precursor to the meliacarpinins. It is of interest that none of the azadirachtin-type limonoids isolated from M. azedarach possesses a 4β-carboxylate group like azadirachtin, but rather a 4β-methyl group [17].

III. STUDIES ON THE LIMONOID ANTIFEEDANTS OF THE CHINESE Melia azedarach A.

Isolation of Limonoids from the Root Bark

The ether and methanol extracts of the root bark contained a variety of limonoids, which were detectable by the characteristic color upon treatment with Ehrlich’s reagent on thin layer chromatography (TLC). Antifeedant limonoids are often very sensitive to traces of acid and gradually decompose on a silica gel column. It was, therefore, necessary to use droplet countercurrent chromatography (DCCC) [18,19], flash chromatography, and high-performance liquid chromatography (HPLC) separation techniques that eliminate or at least minimize contact with a potentially acidic stationary phase. The isolation of the various limonoid congeners was often a tedious process requiring careful use of HPLC. The isolations, which were monitored by antifeedant assays described in the next section, were accomplished as outlined in Fig. 5. B.

Bioassay of Antifeedants

The antifeedant assay results reported here were obtained with the Southern army worm Spodoptera eridania (Boisduval) [20] as the test species. Spodoptera species are distributed throughout the world and constitute a major agricultural threat. The feeding bioassay was carried out by the conventional leaf disk method [21], using 2-cm-diameter leaf disks cut from the Chinese cabbage Brassica campestris L. var. chinensis (Cruciferae) with a cork-borer. Each disk was dipped

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Nakatani

Figure 5 Isolation scheme for the limonoids from the root bark of M. azedarach.

for 2 s in an acetone solution of the sample; 5 treated disks were arranged alternately with another 5 control disks (immersed for 2 s in acetone alone), all concentrically placed near the periphery in a Petri dish, as illustrated in Fig. 6. Subsequently, 10 third-instar larvae were placed in the center of the dish, and the treated and untreated leaves eaten by the larvae in 2-hour to 24-hour periods were evaluated at appropriate intervals. The bioassay was terminated after the

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Figure 6 Strategy for the antifeedant bioassay.

larvae had eaten approximately 50% of the control disks, which usually took 6– 12 h. When the average eaten area of the treated disks was visually judged to be less than 50% of that of the control disks, the test compound was judged to be active. This bioassay was used to guide the isolations to the active limonoids, as well as to assess their relative antifeedant activities. To determine the minimum inhibitory concentration, this choice test was done at 50, 100, 150, 200, 300, 400, 500, and 1000 ppm, with 50 ppm corresponding to a concentration of ca. 1 µg/ leaf-cm2.

C.

Structures of the Trichilins

The series of limonoids called the trichilins was first isolated from the root bark of the East African medicinal plant Trichilia roka (Meliaceae) [22]. The structures of trichilin B (17) and its 12-epimer (trichilin A) were elucidated by extensive 1H and 13C nuclear magnetic resonance imaging (NMR) studies and through chemical correlation. Some pertinent points related to these structural studies are listed as follows: 1. Irradiation of the 13- and 8-Me peaks induced 30% and 12% Nuclear Overhauser Effect (NOE) enhancements of the 9-H and 19-H signals, respectively (see conformational drawing, Fig. 7). 2. The assignment of the 12-OH stereochemistry as β in trichilin A and α in trichilin B (17) was deduced from the finding that in their 12-pbromobenzoates, the aromatic protons of the benzoate and furan rings were at higher field in trichilin B. Thus, the shifts of the p-bromobenzoate protons were (for trichilins A/B) o-H δ 7.99/7.65, m-H δ 7.51/

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Figure 7 1H NMR data for trichilin B (17) in CDCl3, 400 MHz, in δ (multiplicity and J values), and NOE correlations (double-headed arrows). NMR, nuclear magnetic resonance; NOE, nuclear overhauser effect.

7.59, and those of the furan were 21-H δ 7.20/7.02, 22-H δ 6.36/5.98, and 23-H δ 7.37/7.10. The higher-field chemical shifts of these aromatic protons in trichilin B can be accounted for by the mutual shielding induced by the ring currents of the two aromatic rings, which are located on the same side of the molecule. More directly, however, the 12-OH configurations were independently derived from a new addi-

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tivity relation found in the Cotton effects of multiple coupled chromophores in the CD spectra [23]. For the trichilins A and B, the 7,12bis-p-bromobenzoate exciton couplings between the two benzoates as well as with the furan chromophores distinguished the C12 stereochemistry. 3. Oxidation of trichilins A and B with pyridinium chlorochromate (PCC) in CH2Cl2 afforded the same α-diketone I (Scheme 2) as the 7,11,12trione; thus A and B are 12-epimers. 4. When trichilins A and B were treated with a catalytic amount of pTsOH, they were converted quantitatively into the isomeric trichilin C (Scheme 2) and its 12-epimer, respectively. 5. Finally, treatment of trichilin B with Zn(BH4)2 led unexpectedly to a Lewis acid–catalyzed acyl migration in ring A to give a mixture of trichilin B and its 1,2-diacetyl analogue aphanastatin (23), the structure of which had been determined by x-ray analysis [24]. This unexpected conversion confirmed the structure of trichilin B.

Scheme 2

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D.

The Structure of Meliacarpinin A

The meliacarpinins (46–50, and 52) are highly oxidized C-seco limonoids isolated by us from the Chinaberry tree M. azedarach that are structurally very similar to the azadirachtinins. The structure of meliacarpinin A (46) [25] was mainly elucidated by comparing the NMR data with those of the azadirachtins isolated from M. azadirachta. The 1H NMR spectrum was nearly superimposable on that of 1-tigloyl-3-acetoxy-11-methoxyazadirachtinin [26] except for the change of the 4β-methyl carboxylate to a 4β-methyl group. The NMR data and selected NOE connectivities used to elucidate the structure and stereochemistry are shown in Fig. 8.

Figure 8 1H NMR data for meliacarpinin A (46, cin ⫽ cinnamoyl) in CDCl3, 400 MHz, in δ (multiplicity and J values), and NOE correlations (double-headed arrows). NMR, nuclear magnetic resonance; NOE, nuclear overhauser effect.

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IV. ANTIFEEDANT ACTIVITY OF THE LIMONOIDS FROM Melia azedarach The limonoids isolated from M. azedarach and their effects on insect feeding are summarized in Table 1. To develop a quantitative understanding of structure– activity relationships among limonoids, the data are summarized primarily for the activity against the third-instar larvae of a voracious pest insect, Spodoptera eridania (Boisduval). It is important to note that differences in the response of test insects when compared with different test species, or even different life cycle stages of the same species, can mask any meaningful observations of structure– activity relations. This is apparent from a comparison of the data reported against different insects for compounds (11, 22, 39, and 40) in Table 1. Even where the same bioassay species has been used, differences in the larval stage tested may make comparisons invalid. Some quantitative trends are apparent in the data in Table 1. Aside from the highly oxidized C-seco meliacarpinins and azadirachtins, the most active compounds appear to be intact apo-euphol limonoids with a 14,15-epoxide and a C-19/C-29 acyl acetal bridged system. Structure–activity relationships in this class will be discussed next. Within the C-seco class, the highly oxidized meliacarpinins are the most active of the limonoids from M. azedarach. Although their activities may be weaker than that of the azadirachtins from M. azadirachta (effective dose of 50 ppm for 46–52 against S. eridania in comparison to 14 ppm for azadirachtin against Epilachna varivestis), they are much more potent antifeedants than the less oxidized class of C-seco limonoids exemplified by salannin (39), nimbolin B (32), and ohchinolide B (37). The second most active class of limonoids from M. azedarach is the C-19/C-29 bridged acetal class with the intact apo-euphol skeleton (10–28) (effective dose 150–500 ppm). The two members of the intact apo-euphol limonoid class without this C-19/C29 acetal bridge that were tested against S. eridania, azadiron (1), and nimbolin A (9) showed little or no activity (effective dose ⱖ 1000 ppm).

V.

STRUCTURE–ACTIVITY RELATIONSHIPS IN C-19/C-29 BRIDGED ACETALS

During the course of our structural and chemical correlation studies, about 50 compounds belonging to the C-19/C-29 bridged acetal class of limonoids were made available. Antifeedant assays with S. eridania (leaf disk method) showed several interesting structure–activity correlations. First, activity is insensitive to substituent variation in the A-ring except for the nature of the C29 bridging position: A hemiacetal bridge (hydroxyl group at C29) increases the activity in comparison to that of an acylated hemiacetal (com-

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Table 1 Antifeeding Activity of Limonoids from Melia azedarach

Limonoid Group 1: Intact apo-euphol limonoids Azadirone (1) Meldenin (2) 6-Acetoxy-7α-hydroxy-3-oxo-14β,15β-epoxymeliac-1,5-diene (3) 6-Acetoxy-3β-hydroxy-7-oxo-14β,15β-epoxymeliac-1,5-diene 3-O-β-d-glucopyranoside (4) 6-Acetoxy-3β-hydroxy-7-oxo-14β,15β-epoxymeliac-1,5-diene 3-O-β-d-xylopyranoside (5) 6-Acetoxy-3β,11α-dihydroxy-7-oxo-14β,15β-epoxymeliac-1,5-diene 3-O-α-l-rhamnopyranoside (6) 6,11α-Diacetoxy-3β-hydroxy-7-oxo-14β,15β-epoxymeliac-1,5-diene 3-O-β-d-glucopyranoside (7) Sendanal (8) Nimbolin A (9) C-19/C-29 bridged acyl acetals Amoorastatin (10) Toosendanin (11) Azedarachin A (12) 12-O-Acetylazedarachin A (13) Azedarachin C (14) Azedarachin B (15) 12-O-Acetylazedarachin B (16) Trichilin B (17)

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Test insect

Effective concentration (ppm)

Spodoptera eridania

Inactive

S. eridania

1000

S. eridania O. furnacalis S. eridania S. eridania S. eridania S. eridania S. eridania S. eridania S. eridania

150 200 300 200 400 400 200 400 200

References Isolation

Activity a

27 28,29 29 29

12 NA NA

30

NA

31

NA

32

NA

33 11

NA 12

34,35 36,37

12 35 20 37 37 37 12 37 40

38 38 39 12 40 22,40

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12-O-Acetyltrichilin B (18) 1,12-Di-O-acetyltrichilin B (19) Trichilin D (20) Trichilin H (21) Meliatoxin A 2 (22) Aphanastatin (23) Amoorastatone (24) 12-Hydroxyamoorastatone (25) iso-Chuanliansu (26) Meliatoxin B 1 (27) Meliatoxin B 2 (28) Group 2: D-seco limonoids Gedunin (29) 7α-Acetoxy-3β-hydroxy-14β,15β-epoxygedunan-1-ene 3-O-β-d-glucopyranoside (30) Group 3: C-seco limonoids Ohchinal (31) Nimbolin B (32) Nimbolinin B (33) 1-Deacetylnimbolinin B (34) Spirosendan (35) Ohchinolide A (36) Ohchinolide B (37) Ohchinolal (38) Salannin (39) Deacetylsalannin (40) Ohchinin (41)

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S. S. S. S. S. S. S. S. S. S. S. S.

eridania eridania eridania eridania litura eridania eridania eridania eridania eridania eridania eridania

400 400 400 400 300 400 200 400 300 400 500 500

40 40 22,40 40 40,41 24 34,35 35 37 41 41

20 40 40 40 42 40 20 12 12 12 12 12

O. nubilalis

500

11 44

43 NA

S. frugiperda S. eridania S. eridania

Inactive 1000 1000

S. eridania S. eridania S. eridania S. littoralis S. eridania E. varivestis

700 1000 1000 100 1000 30

45 11 47 47 12 47,49 47,49 50 51

46 12 48 NA NA NA 12 12 48 52 12 53 NA

51 50

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Table 1 Continued

Limonoid Ohchinin acetate (42) Nimbolidin A (43) Nimbolidin B (44) Highly oxidized compounds 1-Cinnamoylmelianolone (45) 1-Cinnamoyl-3-acetyl-11-methoxymeliacarpinin (46, Meliacarpinin A) 3-Deacetylmeliacapinin A (47) Meliacarpinin B (48) Meliacarpinin C (49) Meliacarpinin E (50) 20-O-Acetylmeliacarpinin C (51) Meliacarpinin D (52) 20-O-Acetylmeliacarpinin D (53) 1-Deoxy-3-methacrylyl-11-methoxymeliacarpinin (54) Azadirachtin (55, Azadirachtin A) Other trichilins from different species and reaction products 1: Natural products Trichilin A (56) (C12-epimer of trichilin B) 7-O-Acetyltrichilin A (57) Trichilin C (58) (15-Keto isomer of trichilin A)

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Test insect

Effective concentration (ppm)

References Isolation

Activity a

S. eridania S. eridania S. eridania

1000 500 500

45 47 47

12 20 20

S. eridania

50

54 25

NA 25

S. eridania S. eridania S. eridania

50 50 50

S. eridania

50

E. varivestis

14

55 48 48 12 55 48 55 55 56,57

NA 48 48 12 NA 48 NA NA 53

S. eridania

300

22

22

S. eridania S. eridania

400 500

59 22

12 12

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12α-Epimer of trichilin C (59) Trichilin E (60) (C12-Epimer of aphanastatin) Trichilin F (61) (1-O-Acetyl-3-deacetyltrichilin A) Trichilin G (62) (1-O-Acetyl-2,3-deacetyltrichilin A) 2: Acetylated compounds 12-O-Acetyltrichilin A (63) 7,12-Di-O-acetyltrichilin A (64) 1,7,12-Tri-O-acetyl-2-deacetyltrichilin A (65) 1,7,12-Tri-O-acetyl-3-deacetyltrichilin A (66) 7,12-Di-O-acetyltrichilin B (67) 1-O-Acetyl-3-deacetyltrichilin B (68) 12-O-Acetyltrichilin C (69) 3: Oxidized compounds 7-Oxotrichilin A (70) 7-Oxotrichilin B (71) 12-Oxotrichilin B (72) 1,12-Dioxotrichilin B (73) 7,12-Dioxotrichilin B (74) a

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NA ⫽ not available.

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S. eridania S. eridania

400 300

12 22

12 12

S. eridania

300

60

60

S. eridania

300

60

60

S. S. S. S. S. S. S.

eridania eridania eridania eridania eridania eridania eridania

400 500 500 500 500 200 ⬎500

58 58 61 61 58 61 12

58 58 61 61 58 61 12

S. S. S. S. S.

eridania eridania eridania eridania eridania

Inactive Inactive 1000 Inactive Inactive

22 22 22 22 22

58 58 12 58 58

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pare the activities of 10 and 11 with 12–28, Table 1). All natural C-19/C-29 bridged acetals possess the 11-keto group, and the configuration of the C-12 hydroxyl group has a pronounced effect on the activity (compare the activities of (17) vs. 56, 23 vs. 60, and 58 vs. 59). The activity of compounds lacking the 12-OH (for example, 14 vs. 15, 20 vs. 17, and 24 vs. 25) or in which the C7 (57 vs. 56) and/or C12 positions (10 vs. 11, 12 vs. 13, 15 vs. 16, 17 vs. 18 vs. 67, and 56 vs. 63 vs. 64, and 58 and 59 vs. 69) are acetylated or oxidized to ketones (17 vs. 71, 72, and 74, and 56 vs. 70 and 72) are reduced. Replacement of the 14,15-epoxide with a C15 ketone also results in reduced activity (10 vs. 24, 11 vs. 25, 17 vs. 59, 20 vs. 27, and 56 vs. 58). These structure–activity relationships in the trichilins are summarized in Fig. 9.

Figure 9 Structure–activity relations in the trichilins (F ⫽ 3-furyl). (Source: Modified from Ref. 58.)

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Figure 10 Biogenesis and antifeedant activity of the limonoids from M. azedarach.

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VI.

CONCLUSIONS

Research on biologically active limonoids has been motivated by the quest to find useful compounds for specific agricultural or medicinal applications. Though bioassay designs and choice of bioassay species have varied tremendously in the literature, the common conclusion undoubtedly is that limonoids function primarily as ‘‘antifeedants’’ in the host species. This suggests that the primary selective advantage of the production of limonoids by the host plant is protection against insect herbivory. About 350 limonoids have been isolated to date. Although limonoids may be found in all tissues of the plant, limonoid biosynthesis is characterized by an evolutionary pattern of increasing oxidation and rearrangement of the original limonoid skeleton [62]. Relations of biogenesis of the limonoids in M. azedarach with their antifeedant activity are summarized in Fig. 10. From the scheme, it can be seen that the evolutionary trend of increasing oxidation correlates with increasing activity against insects.

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2 Polygodial and Warburganal, Antifungal Sesquiterpene Dialdehydes and Their Synergists Isao Kubo University of California, Berkeley, California

I. INTRODUCTION Many human pathogenic microorganisms can now be controlled with the antibiotics that are presently available. However, with the increase in drug resistance and prevalence of opportunistic infections, there is still a great need for new, more effective agents. For example, systemic infections caused by filamentous fungi, especially in patients with impaired immune defense mechanisms, have become an increasingly serious, worldwide problem. Although various new antifungal agents have been introduced, control of most fungal diseases has not yet been achieved [1]. Hence, in our continuous search for antimicrobial agents from tropical plants, a new emphasis has been placed on antifungal agents, particularly against Candida albicans, considered one of the most devastating fungi responsible for human opportunistic systemic infections [2]. Tropical plants are exposed throughout the entire year to predation by various parasites such as bacteria, fungi, and insects. In order to survive these demanding conditions, efficient built-in chemical defense mechanisms have evolved, and thus tropical plants offer a rich and intriguing source of secondary metabolites possessing attractive biological activities with potential medicinal applications. These plants remain a good source of new antifungal agents [3]. However, there is a need to investigate them from a point of view different from that of the traditional approach of mere isolation and structure determination of biologically active principles. Today, understanding the mechanism, as well as

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enhancing the biological activity, have become important goals of natural product chemists. This largely reflects the recent rapid developments in separation and spectroscopic techniques that render isolation and structure determination more routine. The plants chosen for study were identified primarily on the basis of information provided by medicine men in East Africa, mainly in Kenya and Tanzania [4]. Botanically identified plants were then collected and extracted with methanol at ambient temperatures. The extracts were first tested for antimicrobial activity against four representative microorganisms, Bacillus subtilis, Escherichia coli, Saccharomyces cerevisiae, and Penicillium chrysogenum, at a concentration of 100 µg/mL [5]. The active extracts were then tested against a larger number of microorganisms to determine the scope of activity. Interestingly, the information gathered from medicine men, known as Bwana Mganga in Swahili [4], proved to be very useful in identifying species with significant activities. The plants collected on the basis of their information had a much higher probability of producing active extracts than those collected randomly [6]. From 79 extracts, which included 72 species of plants distributed among 35 families, 40 extracts initially gave positive results indicative of antimicrobial activity at the 100 µg/mL level against one or more of the microorganisms. Noticeably few extracts showed activity against fungi, especially C. albicans. This review focuses on a particular group of compounds: the antifungal sesquiterpene dialdehydes isolated from trees of the Warburgia genus. These sesquiterpene dialdehydes are among the rare phytochemicals that exhibit potent antifungal activity against C. albicans.

II. INITIAL STUDIES The genus Warburgia, endemic to East Africa, belongs to a small family, Canellaceae, and consists of only two species: W. ugandensis and W. stuhlmannii. We first became interested in the tree W. ugandensis while I was on a tenured appointment at the International Centre of Insect Physiology and Ecology (ICIPE), located in Nairobi, Kenya, because of its strong, hot taste. The methanolic extract of the bark of W. ugendensis collected on the ICIPE grounds was originally found to exhibit potent insect antifeedant activity against the African armyworm, Spodoptera exempta [7]. Fractionation guided by the antifeedant assay led to three active principles isolated from the bark, leaves, and fruit of this tree [8,9]. These active compounds, isolated after repeated chromatography and characterized by their unique hot taste, were the sesquiterpene dialdehydes [10] warburganal (1) [11] and muzigadial (also known as canellal) (2) [12,13], in addition to a known congener, polygodial (3). Most of the initial chemical study was performed in Professor Koji Nakanishi’s labs together with his colleagues at Columbia University in New York [11,12,14]. Subsequently, these antifeedant sesquiterpene dialdehydes received considerable attention from synthetic organic

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chemists, and various syntheses have been reported [15–17]. Included among these efforts is the conversion of 1-abietic acid into (⫺)-warburganal, the first reported synthesis of (1) with the natural absolute stereochemistry [18].

Since the Warburgia plants are widely used in folk medicine in East Africa [19], their extracts were also submitted for testing in various available pharmacological assays together with other plant extracts. Among the extracts tested, the two from the Warburgia species were found by Professor Makoto Taniguchi of Osaka City University to exhibit a broad-spectrum antimicrobial activity, including activity against Candida utilis [5]. This finding initiated an investigation to determine the antifungal principles, which was achieved in collaboration with Professor Taniguchi and his coworkers [20–24].

III. ISOLATION AND IDENTIFICATION OF ANTIFUNGAL PRINCIPLES A bioassay-directed fractionation of the n-hexane extract of the bark of W. ugandensis using Bacillus subtilis and Saccharomyces cerevisiae as test organisms resulted in the isolation of three active sesquiterpene dialdehydes 1–3, previously identified as the insect antifeedants from the same source [8,11,12,14]. Their structures were identified by spectroscopic methods [11,12]. In the course of this work it was noted that methanol, originally used to extract the Warburgia bark, at least partly if not totally, inactivated the antifungal activity of these dialdehydes through acetal formation. Therefore, the use of alcohols was avoided throughout the isolation procedure. Polygodial (3) was first isolated as a hot tasting substance from the sprouts of Polygonum hydropiper (Polygonaceae) [25], which has been used in folk medicine and food spices in several Asian countries including Japan and Vietnam. For example, the sprout of P. hydropiper is a well-known relish for sashimi in Japan [26,27]. Subsequently, warburganal (1) was also isolated from the same source in minute amounts [28]. In addition to the antifungal sesquiterpene dialdehydes 1–3, a number of congeners such as mukaadial (4) [29], ugandensidial (also known as cinnamodial) (5) [30], epipolygodial (6) [14], as well as confertifolin (7), 9α-hydroxycinnamolide (8) [31], cinnamosnolide (9) [14], colorata-4 [13], 8-dienolide (10), and bemadienolide (11) [14] were also isolated from

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Scheme 1

ous parts of W. ugandensis, but none exhibited antifungal activity at concentrations of 100 µg/mL. Their structures were all identified by spectroscopic methods, particularly by nuclear magnetic resonance (NMR) spectroscopy. All the sesquiterpenoids isolated (1–11) are considered oxidation products of the drimane skeleton. The same sesquiterpenoids were also isolated from the bark of W. stuhlmannii [11,12,32,34], as well as from several other plants [31–33]. It was also discovered that each dialdehyde gave the corresponding lactone upon treatment with 2N HCl or 1N NaOH. For example, alkaline treatment of (1) at room temperature gave (8) through an intramolecular Cannizzaro reaction, and acid treatment yielded futronolide (12) (Scheme 1). Similarly, alkaline treatment of (5) yielded the lactone (13), which was converted to (9) by acetylation [29]. Nevertheless, these more stable lactones (in comparison to 3) were found to be no longer active.

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IV. ANTIFUNGAL ACTIVITY With the exception of (1)–(3), none of the other sesquiterpenoids (4)–(11) showed any antifungal activity at concentrations up to 100 µg/mL. Bioassay results for these sesquiterpenoids against additional microorganisms are listed in Table 1.

Table 1 Antimicrobial Activity of the Warburgia sesquiterpenoids a MIC (µg/mL) Microorganisms tested Staphylococcus aureus Bacillus subtilis Streptococcus mutans Micrococcus luteus M. lysodeikticus Escherichia coli Enterobacter aerogenes Proteus vulgaris Pseudomonas aeruginosa Helicobacter pylori Saccharomyces cerevisiae Schizosaccharomyces pombe Hansenula anomala Candida utilis C. albicans C. krusei Cryptococcus neoformans Pityrosporon ovale Sclerotinia libertiana Mucor mucedo Rhizopus chinensis Aspergillus niger A. flavus Trichophyton rubrum T. mentagrophytes Penicillium crustsum P. marneffei a

1

2

3

4

6

100 ⬎100 ⬎100 ⬎100 ⬎100 ⬎100 ⬎100 ⬎100 ⬎100 — 3.13 12.5 12.5 3.13 6.25 — — 25 3.13 25 100 50 — — 6.25 50 —

⬎100 ⬎100 ⬎100 ⬎100 ⬎100 ⬎100 — ⬎100 ⬎100 — 1.56 25 25 3.13 — — — — 3.13 25 100 50 — — — 50 —

100 ⬎100 ⬎100 ⬎100 ⬎100 ⬎100 ⬎100 ⬎100 ⬎100 ⬎100 0.78 6.25 1.56 1.56 3.13 6.25 3.13 50 1.56 6.25 12.5 25 50 0.78 3.13 25 3.13

⬎100 ⬎100 — ⬎100 ⬎100 ⬎100 — ⬎100 ⬎100 — ⬎100 ⬎100 ⬎100 ⬎100 — — — — 100 ⬎100 ⬎100 ⬎100 — — — ⬎100 —

⬎100 ⬎100 — ⬎100 ⬎100 ⬎100 — ⬎100 ⬎100 — ⬎100 ⬎100 ⬎100 ⬎100 — — — — ⬎100 ⬎100 ⬎100 ⬎100 — — — ⬎100 —

The rate of activity varied slightly with the seed culture media, the physiological age of the culture and the type of culture medium. The MIC was measured by twofold serial broth dilution. The MIC was the lowest concentration of sample at which no growth of the test microorganism was visible. —, not tested. Source: Refs. 7, 35, and 36.

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Warburganal (1), muzigadial (2), and polygodial (3) exhibited activity against all fungi tested [6]. In particular, high activity was shown against Candida utilis, Saccharomyces cerevisiae, Pityrosporon ovale, Trichophyton rubrum, Penicillium chrysogenum, Hansenula anomala, and Sclerotinia libertiana. The activities of (1) and (3) against the pathogenic fungus Candida albicans were also quite high: their minimum inhibitory concentrations (MICs) were 3.13 and 6.25 µg/mL, respectively [35]. Among the three antifungal sesquiterpene dialdehydes 1–3, the structurally simplest, (3), exhibited the most potent activity [20]. In addition, the activity of (3) against fungi was of broad scope. Polygodial (3) was two to eight times more active than (1) and (2) against most species of fungi tested, and its potency against these fungi was comparable to that of amphotericin B, one of the most potent antibiotics known against filamentous fungi [22], although its high toxicity limits extensive use. Therefore, (3) may be potent enough to be considered for practical application, and further studies with (3) are ongoing. The structural similarity of (2) with (3) suggests that similarly potent antifungal activity should be found for this dialdehyde as well. Although the results in Table 1 indicate that this is the case, (2) could not be extensively tested because of its limited availability. Interestingly, in contrast to (3), the C-9 epimer, epipolygodial (6), did not show antimicrobial activity at concentrations of 100 µg/mL. This difference in antimicrobial activity between (3) and (6) may reflect differences in abilities to cross the cell membranes of the target microorganisms since a marked difference in their water solubilities was noted: (6) is much less soluble in water than (3). The facile, reversible conversion of (3) to the cyclic dihemiacetal (14) in aqueous media (Scheme 2) may assist its water solubilization [20]. Such hydration to a cyclic dihemiacetal would be difficult in (6) with the pseudoaxial C-9α-aldehyde orientation. In support of this suggestion, reduction of the C-9 aldehyde group of (3) resulted in loss of activity. In addition to (6), mukaadial (4), possessing two additional hydroxyl groups at C-6 and C-9, did not exhibit any activity at concentrations up to 100 µg/mL. Numerous phytochemicals have been isolated as potential antifungal agents [3,36]. As natural products, these phytochemicals are typically biodegradable and, more importantly, renewable. The efficient utilization of such renewable

Scheme 2

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natural products is becoming increasingly important worldwide to conserve natural resources. Unfortunately, although these antifungal phytochemicals may play an important role in the defense of the host plants against fungal predation, their biological activities are usually not sufficiently potent for practical application in medicine. This is a common dilemma when the biological activities of phytochemicals are considered; hence, studies to enhance activities are needed for efficient utilization of renewable natural products. An attempt to enhance the antimicrobial activity of some purified antimicrobial agents through combination with other agents was made. In a preliminary experiment, 3 was first tested with several antibiotics such as actinomycin D and rifampicin. In these experiments, one agent is maintained at a concentration below the MIC (for example, 1/2MIC) while the concentration of the second agent is varied to determine its MIC in the presence of the first agent. The roles of the two agents are then reversed. Polygodial (3) significantly enhanced the antifungal activity of these antibiotics against C. utilis and S. cerevisiae, but not vice versa [20,22,23,37]. Polygodial (3) also synergized the antifungal activity of maesanin (15), isolated from the fruit of the East African medicinal plant Maesa lanceolata (Myrsinaceae) [38], against C. utilis [39].

The reason for these combination effects seems to be an increase in the permeability of the plasma membrane toward the antibiotics induced by 3 [40,41]. Thus, when cells of S. cerevisiae were treated with 3, ultrastructural changes in the cell membrane were observed [22], as shown in Fig. 1. These morphological alterations suggested that the primary site of action of 3 is the plasma membrane with simultaneous organelle disorganization followed by the fatal loss of cellular constituents such as proteins and polysaccharides, as illustrated in Fig. 2 [37]. The addition, of excess Ca 2⫹ was found to protect against the polygodial-induced cell membrane damage in S. cerevisiae, and this protection was abolished by the addition of EDTA [24]. It has been suggested that Ca 2⫹ stabilizes the structure of membranes by forming ion bridges between phosphate groups of phospholipids and the carboxyl groups of membrane proteins, similar to bacterial membranes [42]. Interestingly, the protection seems to be specific for Ca 2⫹ since Mg 2⫹, a similar divalent cation, did not give this protection, though both Ca 2⫹ and Mg 2⫹ suppressed miconazole-induced leakage and slightly suppressed amphotericin B– induced leakage. This result indicates that 3, miconazole, and amphotericin B differ in their modes of action on fungal cell membranes [24]. The Ca 2⫹ cation has many properties that make it better suited for chelation than the more abun-

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(a)

(b) Figure 1 (a) Section of untreated control cell of Saccharomyces cerevisiae. (b) Section of S. cerevisiae cell treated with 50 µg/mL of polygodial (3) for 10 min. CW, cell wall; PM, plasma membrane (cell membrane); N, nucleus; M, mitochondrion; V, vacuole.

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Figure 2 Leakage of (a) folin-reagent and (b) phenol-H2 SO4-positive substances from , None; , Saccharomyces cerevisiae cells during incubation with polygodial (3). , 10 µg/mL of 3. 1 µg/mL of 3;

dant Mg 2⫹, such as its larger radius, its lower energy of hydration, and the presence of d-orbitals, allowing it to chelate more readily. In addition, both 3 and 1 have been reported to exhibit the membrane leakage activity in human neuroblastoma cells [43]. The binding site of 3 on the cell membrane and the mechanism of the suppressive effects of Ca 2⫹ on the polygodial-induced leakage remain to be established.

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V.

COMBINATION EFFECTS

In our continuing search for antimicrobial agents from tropical plants, several phenylpropanoids, such as anethole (16), isolated from the seeds of Pimpinella anisum (Umbelliferae) [44], and eugenol (17), methyleugenol (18), and safrole (19), in addition to anethole from the seeds of Licaria puchuri-major (Lauraceae) [45], were isolated as antimicrobial principles in rather large quantities. All exhibited moderate but broad-spectrum activity. Their MICs, ranging from 100 to 800 µg/mL, were not potent enough to warrant further studies on the individual compounds. However, it was still considered worthwhile to investigate the possibility of their use as antifungal agents in combination with other antimicrobial natural products, since they were all isolated from various food spices that have long been consumed by people from many different cultures. Hence, they were first examined in combination with 3 in an attempt to enhance their antifungal activity against several fungi such as C. albicans and S. cerevisiae as well as the dermatomycotic fungus Pityrosporon ovale. Unexpectedly, 3 did not synergize the antifungal activity of any of these phenylpropanoids, though its antifungal activity was significantly increased when combined with one of the phenylpropanoid compounds. Notably, a dramatic increase in the antifungal activity of 3 occurred when it was combined with a sublethal amount (1/2MIC) of 16: the activity of 3 against C. albicans and S. cerevisiae was increased 32- and 64-fold, respectively. Thus, the MIC of 3 against C. albicans was lowered from 3.13 to 0.098 µg/mL, and against S. cerevisiae, from 1.56 to 0.024 µg/mL, when 3 was combined with 100 µg/mL of 16 (1/2MIC for both C. albicans and S. cerevisiae) [44].

VI.

FUNGICIDAL ACTIVITY

These combination effects, based on the MICs obtained after 48-h incubations, do not fully characterize the antifungal activity. For example, it was not clear whether the combination of 3 and 16 was fungicidal or fungistatic. Hence, the viable count method [46] to analyze the growth curve of C. albicans was used

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in order to obtain the minimum fungicidal concentration (MFC). The growth curves of C. albicans in the presence of 3 and 16 are illustrated in Fig. 3 [47]. Polygodial (3) and anethole (16) exhibited fungicidal activity against C. albicans at 3.13 and 200 µg/mL, respectively; however, they were not fungicidal at 1.56 and 100 µg/mL, respectively. Thus, C. albicans was not viable after 18 and 42 h at 3.13 µg/mL of 3 and 200 µg/mL of 16. The MFCs of both 3 and 16 against C. albicans were therefore the same as their MICs. The combination of 3 and 16, however, was found to possess fungicidal rather than fungistatic activity at concentrations far below their MICs. Fungicidal activity against C. albicans with the combination of 3 and 16 is shown in Fig. 3. When 100 µg/mL of 16 (1/ 2MIC for C. albicans) was combined with more than 0.098 µg/mL of 3, C. albicans was not viable after 6 or 12 h. Thus, the fungicidal activity of 3 against C. albicans was increased 32-fold by 16. In contrast, the fungicidal activity of 16 was enhanced only 4-fold by 3. After this discovery, warburganal (1) was also examined in combination with 16 to see whether 16 had the same enhancing activity [35]. As expected, 16 also significantly increased the activity of 1 against both C. albicans and S. cerevisiae. In this combination, the activity of 1 against C. albicans and S. cerevisiae was enhanced 32- and 256-fold, respectively, when combined with 100 µg/mL of 16. The MIC of 1 against C. albicans was lowered from 6.25 to 0.20 µg/mL, and against S. cerevisiae from 6.25 to 0.024 µg/mL. Anethole (16) also enhanced the activity of 3 and 1 against P. ovale but not as much as against C. albicans and S. cerevisiae. The MICs were reduced only from 50 to 6.25 µg/ mL for 3, and from 25 to 3.13 µg/mL for 1 in combination with 50 µg/mL 16

Figure 3 Growth curves of Candida albicans in the presence of anethole (16), polygodial (3), and their combination.

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(1/2MIC for P. ovale). Similarly, the antifungal activity of 3 was significantly increased when combined with subinhibitory concentrations of methyleugenol (18) and safrole (19), whereas eugenol (17) did not induce any meaningful enhancement of activity, as shown in Table 2. As shown by the results presented in Table 2, 18 induced a 128-fold enhancement of the activity of 3 against P. ovale; the MIC was lowered from 50 to 0.39 µg/mL. A preliminary analysis suggests that those phenylpropanoids that do not possess a free phenolic group, such as 16, 18, and 19, enhance the activity of 3 more than 17, which has a free phenolic group, though the number of samples tested to date is rather limited. Anethole (16) was very effective in enhancing the antifungal activity of 3 and 1 against C. albicans, C. utilis, and S. cerevisiae, and the combinations with 18 were the most effective against P. ovale [44]. In addition, 16 was also combined with other antifungal agents, such as amphotericin B. This combination was investigated since amphotericin B is also known to damage the plasma membrane by interacting with sterols [41] in fungal cells [40]. However, 16 did not enhance the antifungal activity of amphotericin B against C. albicans and S. cerevisiae; in fact, its antifungal activity was somewhat antagonized by 16, as shown in Table 3. Increasing amounts of 16 decreased the antifungal activity of amphotericin B significantly. Specifically, the activity of amphotericin B against C. albicans and S. cerevisiae decreased 16-fold when it was combined with 25 and 50 µg/ mL of 16, respectively. In contrast, 16 did increase the antifungal activity of amphotericin B against P. ovale 8-fold. Thus, the MIC was lowered from 3.13 to 0.39 µg/mL when amphotericin B was combined with 50 µg/mL of 16. These results indicate that a chemical reaction between 16 and amphotericin B did not occur prior to the assay. The effect of 16 on the activity of amphotericin B and

Table 2 Antifungal Activity of Polygodial (3) in Combination with 1/2MIC of Phenylpropanoids 16–19 MIC (µg/mL) Compounds combined

Candida albicans

Saccharomyces cerevisiae

Polygodial (3) alone ⫹ Anethole (16) ⫹ Eugenol (17) ⫹ Methyleugenol (18) ⫹ Safrole (19)

6.25 0.20 3.13 1.56 0.78

3.13 0.049 1.56 0.78 0.39

Source: Refs. 35, 44, 45, and 47.

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Pityrosporon ovale 50 6.25 25 0.39 6.25

Table 3 Antifungal Activity of Amphotericin B in Combination with Anethole (16) MIC of amphotericin B (µg/mL) Concentration of 16 (µg/mL) 0 6.25 12.5 25 50 100

Candida albicans

Saccharomyces cerevisiae

Pityrosporon ovale

0.78 1.56 3.13 12.5 6.25 0.20

0.78 1.56 3.13 6.25 12.5 3.13

3.13 3.13 0.78 0.78 0.39 —

Source: Ref. 44.

other compounds therefore depends on the species of fungi being tested and the antifungal agents being combined. Although accumulation of this kind of knowledge may provide new insight into the molecular basis of fungicidal activity, the mechanism of the combination treatment remains to be established.

VII. MODES OF FUNGICIDAL ACTION A deeper probe into the mechanism of the antifungal activity of polygodial 3 was also undertaken. It is known that the addition of glucose to an unbuffered suspension of S. cerevisiae cells results in the extrusion of acid. The change in external pH upon the addition of glucose is characteristic of yeast cells and is accepted to be due to the action of the plasma membrane H⫹ –adenosine triphosphatase (H⫹-ATPase) [48]. The activation of H⫹-ATPase by glucose at the molecular level is not yet fully understood, but the maintenance of internal pH homeostasis is essential for the cell to survive since intracellular pH is important for the activity of a number of enzymes with pH optima [49,50]. So presumably, the added glucose results in an increase in intracellular acidity that must be eliminated. This glucose-induced medium acidification was inhibited by 3, as illustrated in Fig. 4. The inhibition was presumably caused by inhibition of H⫹-ATPase. In support of this conclusion, 3 was also found to inhibit the isolated H⫹-ATPase of S. cerevisiae. Therefore, it is possible that the potent antifungal activity of 3 is, at least in part, due to its inhibition of the plasma membrane H⫹-ATPase. Interestingly, the inhibitory action of 3 on the glucose-induced acidification of the medium was strongly suppressed by Ca 2⫹, but only weakly by Mg2⫹.

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Figure 4 Inhibitory effect of polygodial (3) on medium acidification by the plasma membrane H ⫹-ATPase of Saccharomyces cerevisiae. The medium acidification was induced by adding the glucose solution (final concentration 2%) and was evaluated with the mole concentration of protons calculated with external medium pH. The ratio of inhibition (percentage) was calculated as follows: (1 ⫺ [H ⫹]inhibitor /[H ⫹]inhibitor free) ⫻ 100. H ⫹-ATPase, H ⫹ –adenosine triphosphatase.

It should also be noted that the activity of 3 is enhanced under acidic conditions [22]. It is known that yeast cells are able to maintain a normal internal pH when suspended in an acidic medium with relatively little change in the intracellular pH. The acidic conditions appear to stimulate the plasma membrane H⫹ATPase activity and ‘‘excess’’ protons are pumped out to the external medium, maintaining constant internal pH during growth [48]. As a result of the inhibition of the plasma membrane H⫹-ATPase by 3, the intracellular pH may drop into the range where phosphofructokinase is sensitive [51]. The subsequent inhibition of glycolysis caused by this inactivation of phosphofructokinase results in a drop in adenosine triphosphate (ATP) levels and thus restricts growth [49]. This rationale may explain why 3 is more potent in the acidic conditions. In contrast to the potent antifungal activity of 3, its congener 4 did not exhibit any activity up to 800 µg/mL; 1 was active, though to a lesser extent than 3 [33]. Thus, the activity decreased for each additional hydroxyl group ‘‘added’’ to the molecular framework of 3. The fungicidal activity of 3 was explained as the result of the structural disruption of the cell membrane [21,52,53], as illustrated in Fig. 1. Moreover, in previous work using human neuroblastoma cells [43], the increase in membrane permeability was demonstrated to depend

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on the accumulation of the unsaturated dialdehydes in the membranes with the reactive aldehyde groups oriented toward the membrane surface. Considering the results so far reported, the antifungal mechanism of 3 may result, at least in part, from its ability to function as a nonionic detergent, similar to long-chain alcohols [54]. The greater activity of the dialdehydes 1 and 3 could be due primarily to a balance between the hydrophilicity of the unsaturated aldehyde subunits and the hydrophobicity of the decalin portion of the molecules; 4 does not possess this balance because of its increased hydrophilicity and, hence, is inactive. As a nonionic detergent, 3 would likely approach the binding site with the electronegativity of the aldehyde oxygen atom. The aldehyde oxygens are potent hydrogen bond acceptors that will disrupt existing hydrogen bonds. For example, in the lipid bilayer the hydroxyl group of ergosterol, a major component of the plasma membrane of S. cerevisiae, resides near the membrane–water interface and is likely hydrogen bonded with the carbonyl group of phospholipids [55,56]. Since ergosterol owes its membrane-fixing properties to its rigid, longitudinal orientation in the membrane and has profound influence on membrane structure and function, if these hydrogen bonds are disrupted, the cell will die. Interestingly, the polygodial-induced membrane damage was prevented by Ca 2⫹, but this protection is eliminated by adding EDTA [24]. The role of Ca 2⫹ is still unclear and many mechanisms to explain it seem possible. For example, the possibility of Ca 2⫹ binding to the negatively charged phosphate oxygen atoms on the membrane, similar to bacterial membranes [42], cannot be entirely ruled out. If this is so, it would result in formation of a cross-linked membrane structure that may impede the approach of 3 to the binding site on the cell membrane. Needless to say, further study is needed to clarify the Ca 2⫹ protection mechanism. Given the detergentlike properties of 3, it is possible to suggest that 3 also acts at the lipid–protein interface of H ⫹-ATPase, denaturing its functioning conformation. In a system containing both lipids and proteins, it is difficult to determine whether a conformational change of a protein is the result of a direct H ⫹ATPase interaction or of motional or conformational modification of the lipids themselves that exist at the lipid–protein interface. Nevertheless, the binding of 3 as a nonionic detergent can only involve relatively weak head group interactions, such as hydrogen bonding. It is suggested that the intrinsic proteins of membranes are held in position by hydrogen bonding, as well as by hydrophobic and electrostatic forces, and that hydrogen bonding also mediates the penetration of membranes by proteins. As proposed, hydrogen bonds may be disrupted by 3 and redirected. Thereby the conformation of the protein may be changed, and consequently the H ⫹-ATPase in particular may lose its functioning conformation. Although H ⫹-ATPase is the most abundant plasma membrane protein, constituting over 20% of the total membrane protein in S. cerevisiae, other plasma membrane proteins may also be disrupted by 3. All of this is consistent with the previous report that the primary active site of 3 is at the membrane [22].

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All the data so far obtained can be explained by the detergent concept. It should be remembered that 3 showed potent fungicidal activity, particularly against yeasts, but the detergent mechanism mentioned previously should not be specific. The specificity of 3 against yeasts is likely based on its hydrophobic decalin moiety. The activity of polygodial may be increased by modifying this hydrophobic decalin moiety, but experimentally this may not be practical. In addition, α,β-unsaturated aldehydes are highly reactive substances, and they readily react with biologically important nucleophiles, such as sulfhydryl, amino, and hydroxyl groups. The main reaction appears to be 1,4-addition under physiological conditions, although the formation of Schiff bases is also possible [10,57]. An earlier report demonstrated a good correlation between the antifungal activity and the papain inhibitory activity of 3. Both activities appear to result from highly specific reactivity of sulfhydryl groups with the enal group [21]. Yeast plasma-membrane H ⫹-ATPase is reported to contain nine cysteines [58]. Polygodial (3) may bind directly to the plasma-membrane H ⫹-ATPase, possibly with sulfhydryl groups of the three cysteines in the presumed transmembrane segments (C148A, C312S, C867A). However, Petrov and Slayman [59] reported that no single cysteine is required for the enzyme activity, on the basis of their site-directed mutagenesis study. This does not exclude, however, the possibility that 3 breaks a hydrogen bond as a detergent and then reacts with the freed sulfhydryl group of the H ⫹-ATPase. This is supported by the previous report by Monk and his colleagues [60] that covalent modification of the conserved C148 in the transmembrane segment 2 may be important for inhibition of H ⫹-ATPase activity and cell growth. Nevertheless, the involvement of this kind of biochemical reaction is still unclear.

VIII.

CONCLUSIONS

The data so far obtained indicate that polygodial (3) initially acts as a nonionic detergent. More importantly, 3 inhibits the plasma-membrane H ⫹-ATPase by disrupting and disorganizing the hydrogen bonds at the lipid bilayer–protein interface. It seems that 3 targets the extracytoplasmic region and thus does not need to enter the cell, thereby avoiding most cellular pump–based resistance mechanisms. For the otherwise healthy person, fungal infections are more of a nuisance than health-threatening and are normally kept in check by a strong immune system and by otherwise innocuous bacteria of the throat and gut. However, when outside forces such as cancer chemotherapy or heavy doses of antibiotics disrupt the body’s natural defenses, fungal populations can sharply increase and cause serious health problems. Synergistic substances that enable physicians to use lower, safer antifungal dosages would be a useful addition to the therapeutic

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arsenal. Further studies of increased potency of currently used antifungal agents due to combination with 3, as well as 1 and 2, may provide a more rational and scientific approach for the development of new and extraordinarily powerful antifungal treatments, especially against the deadly C. albicans.

ACKNOWLEDGMENT This work has involved a number of scientists. I am greatly indebted to my colleagues cited in the references, especially Professor K. Nakanishi, Professor M. Taniguchi, Dr. M. Himejima, and Dr. S. H. Lee.

REFERENCES 1. S. Sternberg, Science, 266: 1632 (1994). 2. Y. Fukuzawa and K. Kagawa, in Filamentous Microorganisms (T. Arai, ed.) Japan Scientific Societies, Tokyo, pp. 247–253. (1985). 3. L. A. Mitscher, S. Drake, S. R. Gollapudi, and S. K. Okwute, J. Nat. Prod., 50: 1025 (1987). 4. I. Kubo, in Science Year 1982, World Book-Childcraft International, Chicago, pp. 126–137. (1981). 5. M. Taniguchi, A. Chapya, I. Kubo, and K. Nakanishi, Chem. Pharm. Bull., 26: 2910 (1978). 6. M. Taniguchi and I. Kubo, J. Nat. Prod., 56: 153, (1993). 7. I. Kubo, in Methods in Plant Biochemistry, Vol. 6 (K. Hosttetmann, ed.) Academic Press, London, pp. 179–193. (1991). 8. I. Kubo and K. Nakanishi, in The Chemical Basis for Host Plant Resistance to Pest, ACS Symposium Series 62, (P. A. Hedin, ed.), American Chemical Society, Washington, D.C., pp. 165–178. (1977). 9. I. Kubo, in Recent Advances in Phytochemistry: Phytochemical Potential of Tropical Plants, Vol. 27, (K. R. Downum, J. T. Romeo, and H. A. Stafford, eds.) Plenum, New York, pp. 133–151. (1993). 10. I. Kubo and I. Ganjian, Experientia, 37: 1063 (1981). 11. I. Kubo, Y. W. Lee, M. Pettei, F. Pilkiewicz, and K. Nakanishi, J. Chem. Soc. Chem. Commun., 1013 (1976). 12. I. Kubo, I. Miura, M. Pettei, Y. W. Lee, F. Pilkiewicz, and K. Nakanishi, Tetrahedron Lett., 4553 (1977). 13. F. S. El-Feraly, T. McPhail, and K. D. Onan, J. Chem. Soc. Chem. Commun., 75 (1978). 14. K. Nakanishi and I. Kubo, Isr. J. Chem., 16: 28 (1978). 15. S. P. Tanis and K. Nakanishi, J. Am. Chem. Soc., 101: 4399 (1979). 16. M. Jalani-Naini, D. Guillerm, and J. Y. Lallemand, Tetrahedron, 39: 749 (1983). 17. K. Mori and H. Watanabe, Tetrahedron, 42: 273 (1986).

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18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50. 51.

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H. Okawara, H. Nakai, and M. Ohno, Tetrahedron Lett., 23: 1087 (1982). J. O. Kokwaro, in Medicinal Plants of East Africa, East African Literature Bureau, Nairobi, p. 45 (1976). M. Taniguchi, T. Adachi, S. Oi, A. Kimura, S. Katsumura, S. Isoe, and I. Kubo, Agric. Biol. Chem., 48: 73 (1984). M. Taniguchi, T. Adachi, H. Haraguchi, S. Oi, and I. Kubo, J. Biochem., 94: 149 (1983). M. Taniguchi, Y. Yano, E. Tada, K. Ikenishi, S. Oi, H. Haraguchi, K. Hashimoto, and I. Kubo, Agric. Biol. Chem., 52: 1409 (1988). M. Taniguchi, Y. Yano, K. Motoba, S. Oi, H. Haraguchi, K. Hashimoto, and I. Kubo, Agric. Biol. Chem., 52: 1881 (1988). Y. Yano, M. Taniguchi, T. Tanaka, S. Oi, and I. Kubo, Agric. Biol. Chem., 55: 603 (1991). C. S. Barnes and J. W. Loder, Aust. J. Chem., 15: 322 (1962). A. Ohsuka, Nippon Kagaku Zasshi, 84: 748 (1963). I. Kubo, Drug News Persp., 2: 292 (1989). Y. Fukuyama, T. Sato, I. Miura, and Y. Asaka, Phytochemistry, 24: 1521 (1985). I. Kubo, T. Matsumoto, A. B. Kakooko, and N. K. Mubiru, Chem. Lett., 979 (1983). C. J. W. Brooks and G. H. Draffan, Tetrahedron, 25: 2887 (1969). D. Kioy, A. I. Gray, and P. G. Waterman, J. Nat. Prod., 52: 174 (1989). R. F. McCallion, A. L. J. Cole, J. R. L. Walker, J. W. Blunt, and M. H. G. Munro, Planta Med., 44: 134 (1982). M. S. Al-Said, S. M. El-Khawaja, F. S. El-Feraly, and C. H. Hufford, Phytochemistry, 29: 975 (1990). D. Kioy, A. I. Gray, and P. G. Waterman, Phytochemistry, 29: 3535 (1990). I. Kubo and M. Himejima, Experientia, 48: 1162 (1992). L. A. Mitscher, R. P. Leu, M. S. Bathala, W. N. Wu, J. L. Beal, and R. White, Lloydia, 35: 157 (1972). I. Kubo and M. Taniguchi, J. Nat. Prod., 51: 22 (1988). I. Kubo, T. Kamikawa, and I. Miura, Tetrahedron Lett., 24: 3825 (1983). M. Taniguchi, Y. Yano, E. Tada, T. Tanaka, S. Oi, H. Haraguchi, K. Hashimoto, and I. Kubo, Agric. Biol. Chem., 53: 1525 (1989). S. C. Kinsky, Annu. Rev. Pharmacol., 10: 119, (1970). J. M. T. Hamilton-Miller, Adv. Appl. Microbiol., 17: 109 (1974). M. A. Asbell and R. G. Eagon, J. Bacteriol., 92: 380 (1966). A. Forsby, E. Walum, and O. Sterner, Chem. Biol. Interact., 84: 85 (1992). I. Kubo and M. Himejima, J. Agric. Food Chem., 39: 2290 (1991). M. Himejima and I. Kubo, J. Nat. Prod., 55: 620 (1992). C. W. Norden, H. Wentzel, and E. Keleti, J. Infec. Dis., 140: 629 (1979). M. Himejima and I. Kubo, J. Agric. Food Chem., 41: 1776 (1993). P. Eraso and C. Gancedo, FEBS Lett., 224: 187 (1987). W. B. Busa and R. Nuccitelli, Am. J. Physiol., 246: 409 (1984). S. Ramos, M. Balbin, M. Raposo, E. Valle, and L. A. Pardo, J. Gen. Microbiol., 135: 2413 (1989). H. A. Krebs, D. Wiggins, and M. Stubbs, Biochem. J., 214: 657 (1983).

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52. M. N. Jones and D. Chapman, Micelles, Monolayers, and Biomembranes, WileyLiss, New York (1995). 53. A. Kockova´-Kratochvi´lova´, Yeasts and Yeast-like Organisms, VCH, Weinheim (1990). 54. I. Kubo, H. Muroi, and A. Kubo, Bioorg. Med. Chem. Lett., 3: 873 (1995). 55. H. Brockerhoff, Lipids, 9: 645 (1970). 56. V. P. S. Chauhan, L. S. Ramsammy, and H. Brockerhoff, Biochim. Biophys. Acta., 772: 239 (1984). 57. E. Schauenstein, H. Esterbauer, and H. Zollner, in Aldehydes in Biological Systems, Pion, London, pp. 172–200 (1977). 58. R. Serrano, M. C. Kielland-Brandt, and G. R. Fink, Nature, 319: 689 (1986). 59. V. V. Petrov and C. W. Slayman, J. Biol. Chem., 270: 28535 (1995). 60. B. C. Monk, A. B. Mason, G. Abramochkin, J. E. Haber, D. Seto-Young, and D. S. Perlin, Biochim. Biophys. Acta, 1239: 81 (1995).

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3 Marine Bromoperoxidases— Chemoenzymatic Applications Chris A. Moore and Roy K. Okuda San Jose´ State University, San Jose´, California

I. INTRODUCTION A distinctive feature of many marine natural products is the presence of halogen atoms, particularly chlorine, bromine, and iodine, as structural components (Fig. 1). There is little doubt that the organically bound halogens are derived from halide salts that occur naturally in seawater and are incorporated during the biosynthetic process. Approximately 2400 marine natural products that are known contain one or more halogen atoms, of which chlorine and bromine are the most abundant [1,2]. Considering that the concentration of bromide in seawater is 1/300 that of chloride (65 mg/L bromide vs. 19,000 mg/L chloride) [3], the frequent occurrence of bromine in marine natural products suggests that it may be selected for incorporation into secondary products over chlorine. In contrast to marine secondary products, a much smaller number of terrestrially derived natural compounds contains halogen. Indeed, only a handful of brominated natural products from terrestrial sources has been reported [2]. To date, the only enzymes that are known to incorporate halogen into organic substrates are known collectively as the haloperoxidases. Chloroperoxidases are described primarily from the fungi and some bacteria [4]. These enzymes in the presence of both hydrogen peroxide, a ‘‘suitable’’ organic substrate (defined later), and either chloride, bromide, or iodide catalyzes the halogenation of the substrate [5]. Bromoperoxidases are able to utilize only bromide and iodide but are much more widely distributed in nature, having been reported from numerous species of bacteria, marine algae, and some marine invertebrates (e.g.,

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worms and mollusks). Iodoperoxidases are able to utilize only iodide as the halide and are found in marine algae and in the thyroid gland in humans [6]. Although haloperoxidases are believed to be responsible for the incorporation of halogen during the biosynthetic process, this activity has only been definitively demonstrated in a small number of cases [7,8]. However, the existence of these enzymes in organisms that also produce significant quantities of halogenated secondary metabolites is unlikely to be coincidental. It should be noted that some haloperoxidases are also found in organisms that are not known to produce halogenated organics [9]. This may be due to either as yet undetected halogenated organic products (e.g., proteins or nucleic acids) or a possible alternative role for the enzyme in the source organism. This chapter will focus only on those marine-derived bromoperoxidases, which compose the largest group of haloperoxidases reported and on which much recent research has focused. The distribution of these enzymes among diverse taxonomic sources leads to an excellent potential for finding variants that have distinct chemoenzymatic applications.

II. BACKGROUND A.

The Enzymes

The first haloperoxidase to be characterized was named chloroperoxidase and was identified from the fungus Caldariomyces fumago [10]. This enzyme contains heme-bound iron as the cofactor and has been extensively investigated [11]. Several bacteria are reported to contain a nonheme chloroperoxidase, which is involved in the chlorination step of chlorinated metabolites [4]. Except for a small number of cases of enzymes from a marine sponge [12] and a marine worm [13], all chloroperoxidases reported to date have been isolated from nonmarine fungi [6]. Bromoperoxidases are so named because of their requirement for hydrogen peroxide and bromide (or iodide) for catalytic activity. If an organic substrate containing a functional group that is susceptible to electrophilic attack (e.g. alkene, alkyne, aromatic, β-diketone) is also present, an electrophilic substitution or addition occurs. In the mechanistic sense, the bromide is oxidized to a bromonium ion equivalent and reacts as an electrophilic species. In some cases, the bromide becomes incorporated into the substrate; in others, it may be part of a transient intermediate and may not be found in the product [6]. In addition to being classified according to the halides used by an enzyme, haloperoxidases are also categorized by the metal cofactor present. Two general classes are recognized—those that contain heme (the ‘‘H’’ haloperoxidases, such as chloroperoxidase), and the nonheme type (the ‘‘NH’’ enzymes, which include

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most marine bromoperoxidases). Both H and NH bromoperoxidases have been found in marine organisms. Interestingly, a significant number of marine algal bromoperoxidases has been found to have vanadium species as the metal cofactor, with or without nonheme iron present [14]. The bioinorganic chemical processes of this system have been of considerable interest. Indeed, chemical mimics of the vanadium bromoperoxidases that have been prepared and exhibit similar gross catalytic activity [15,16].

B.

Distribution in Marine Organisms

Among marine organisms, bromoperoxidases have been most widely reported from algae. In particular, these enzymes appear to be common in many species of red algae. These enzymes are also found in a number of species of brown algae but are reported only from several green and blue–green algae. Two comprehensive surveys of marine algae and bromoperoxidases have appeared [9,17]. Table 1 lists some of the marine algae from which a significant level of bromoperoxidase activity has been reported. 1). Bromoperoxidases have also been reported from a small number of marine invertebrates (Table 1). One report indicates their presence in a marine annelid and in a marine hemichordate, or ‘‘acorn worm’’ [22], which is known to produce copious quantities of halogenated aromatic metabolites [28]. That brominating enzymes have been found in marine algae and invertebrates is perhaps not surprising, since many are known to produce significant quantities of halogenated natural products (Fig. 1). Hewson and Hager [17] found a strong correlation between the presence of bromoperoxidase and the presence of halogenated lipids in marine algae collected from the Gulf of California. Presumably, the enzymes are involved in the halogenation step(s) of the biosynthesis of these compounds. In our own work, we have found bromoperoxidases in many species of algae that are not known to produce halogenated compounds. This suggests that these enzymes may serve another catalytic role in the algae, other than biological halogenation [9]. It is interesting to note that although chlorinated, as well as brominated and iodinated, metabolites appear in marine natural products, few chlorinating enzymes have been reported from marine sources. An ‘‘abnormal’’ bromoperoxidase with chlorinating ability was reported from the sponge Iotrochoa birotulata [12]. The haloperoxidase from the brown alga Ascophyllum nodosum can be induced to incorporate chloride into organic substrates, but at a rate 500 times less that if bromide were used [29]. Although nearly all marine-derived haloperoxidases have been bromoperoxidases, recently an unusual flavin-containing chloroperoxidase has been reported from a marine polychaete worm [13].

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Table 1 Representative Marine Organisms That Contain Bromoperoxidase Marine algae Reference Red algae (phylum Rhodophyta) Asparagopsis taxiformis Bonnemaissonia hamifera Ceramiun rubrum Corallina officinalis C. pilulifera C. vancouveriensis Cystoclonium purpureum Gracilaria sjoestedtii Laurencia nipponica Plocamium cartilageneum Portieria hornemanni Rhodomela larix Brown algae (phylum Phaeophyta) Ascophyllum nodosum Fucus distachia Laminaria digitata Macrocystis pyrifera Green algae (phylum Chlorophyta) Codium cylindricum Halimeda incrassata Penicillus capitatus P. lamourouxii Rhipocephalus phoenix Udotea flabellum

United States (Hawaii) Mexico (Gulf of California); United States (New Hampshire) Netherlands Great Britain; United States (Massachusetts) Japan United States (California) United Japan United United United

States (California) States (California) States (Hawaii) States (Washington)

9 17 9 18 14 19 20 9 21 9 8 9 9 22

Netherlands Netherlands France United States (California)

23 24 23 24

Japan (Okinawa) Mexico (Gulf of California) United States (Florida) United States (Florida) United States (Florida) Mexico (Gulf of California)

9 17 25 26 26 17

Invertebrates Sponge Iotrochoa birotulata Worms Ptychodera flava (acorn worm) Thelepus setosus (annelid) Gastropod (phylum Mollusca) Murex trunculus

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United States (Florida)

12

United States (Hawaii) United States (Hawaii)

22 22

Not given in ref.

27

Figure 1 Examples of halogenated marine natural products.

C.

Mechanism of Action

Several articles have reviewed the mechanism of action of marine bromoperoxidases [5,30], and thus only a brief discussion will be given here. Heme bromoperoxidases react via oxidation of the heme iron by hydrogen peroxide to yield an activated species, called Compound 1, which then oxidizes the halide to a halonium ion equivalent. It is unclear whether the halonium ion is enzyme bound or exists as a hypohalous acid (Fig. 2) [6]. Nonheme vanadium bromoperoxidases from marine algae are reported to yield an ‘‘enzyme-trapped’’ bromonium species; if an organic substrate is present, it is bound to the enzyme, whereupon reaction occurs [31]. Bromoperoxidases react primarily with substrates that are susceptible to electrophilic attack. In biosynthesis, the bromoperoxidases obviously exert their halogenating capabilities in a regio- and stereoselective fashion, which gives rise to many natural products containing chiral halogens. Since chemical routes to chiral halogen moieties starting from achiral substrates are still rather circuitous, if bromoperoxidases can be induced to perform enantioselective halogenation in a chemoenzymatic mode, these enzymes could be extremely useful for synthetic applications. To date, enantioselective halogenation has not been achieved by using bromoperoxidases. However, work in this area is still in its preliminary development, and the enzymes may require selection of proper experimental conditions before progress is made.

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Figure 2 Possible mechanism of iron-heme bromoperoxidase reactions, ‘‘EOX’’ vs. HOBr. (Source : Ref. 10.)

III. CHEMOENZYMATIC APPLICATIONS OF MARINE BROMOPEROXIDASES A.

Key Parameters for Enzymatic Activity

As with all chemoenzymatic applications, a number of reaction parameters must be carefully manipulated to give optimal yield of the product. With bromoperoxidases, the most important conditions are related to the pH of the buffer, the concentration and rate of addition of hydrogen peroxide, and, to a lesser extent, the concentration of bromide in solution. These reactions are typically conducted in aqueous buffer solutions, but if a substrate is insoluble in water, organic cosolvents may be used. In general, the bromoperoxidases are reported to be relatively ‘‘robust’’ and are fairly stable to storage, in contrast to other enzymes [14]. It is rather difficult to establish one common set of reaction conditions that

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will work best with all bromoperoxidases, since each investigator uses a unique set of parameters to perform reactions with these enzymes. Furthermore, enzymes from different biological sources are also likely to have different reaction characteristics, thus adding to the complexity. However, for most of the marine algal bromoperoxidases, which by far are the largest group under study, the optimal reaction parameters fall within a definable range. In nearly all cases, the optimal ranges of enzyme activity have been determined by using the standard monochlorodimedone (MCD) assay for haloperoxidases (Fig. 3) [10]. Although this may not necessarily reflect the reactivity of the bromoperoxidase with other substrates, the general use of this assay at least gives benchmark values across which all of these enzymes may be compared. The major reaction parameters determined by the MCD assay will be briefly discussed for representative bromoperoxidases. 1. pH Nearly all marine bromoperoxidases have a pH optimum for reactivity that falls in the range of 5.0 and 7.0. This is in marked contrast to fungal chloroperoxidase, which has a pH optimum near pH 3.3 [10]. The pH optimum for most reported marine bromoperoxidases is a fairly sharp peak, and generally activity drops to less than 50% of the maximum if the solution is ⫾2 pH units from the optimal value [9]. Fig. 4 shows examples of pH optima for some marine algal bromoperoxidases. 2. Hydrogen Peroxide (H 2 O 2) Concentration and Rate of Addition Hydrogen peroxide provides the oxidative impulse necessary for the bromoperoxidase to activate bromide to an electrophilic species. In too-high titers, however, H 2O 2 has been shown to inactivate bromoperoxidase, probably by irreversible

Figure 3 Monochlorodimedone (MCD) assay for haloperoxidase activity.

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Figure 4 pH Optima of some algal bromoperoxidases. (Source : Ref. 9.)

oxidation of amino acid residues or of the metal cofactor. Hydrogen peroxide is reported to be a noncompetitive inhibitor of A. nodosum bromoperoxidase, with greater inhibition at higher pH [32]. This sensitivity may reflect that in the natural system, the enzyme is subjected to only very low levels of hydrogen peroxide at all times. Product formation occurs at much higher yield if peroxide is added in small aliquots over an extended period [33]. Examples of enzyme activity vs. added hydrogen peroxide are shown in Fig. 5.

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Figure 5 Hydrogen peroxide optima of some algal bromoperoxidases. (Source: Ref. 9.)

3. Bromide Concentration Optimal bromide concentrations for bromoperoxidase activity vary rather widely among different sources of the enzyme. In some cases, the enzyme activity diminishes significantly upon addition of additional bromide. Interestingly, some bromoperoxidases are still active at very high levels of bromide (e.g., 1M NaBr) [9]. 4. Buffer and Organic Cosolvents One problem commonly associated with chemoenzymatic reactions is that these reactions are often carried out in aqueous buffered solutions. Hence, the organic substrates must be water soluble in order to react. We have found that many nonpolar organic compounds do eventually dissolve in water over extended periods and show a high level of reactivity with these enzymes. However, bromoperoxidases are known to be quite robust enzymes [14] and maintain a good level of catalytic activity if used in mixtures of aqueous buffer and an organic solvent (e.g. methanol, dimethyl sulfoxide) to solubilize the substrate. Bromoperoxidase immobilized on agarose also maintains good catalytic activity in some organic solvents [34].

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Figure 6 Bromide optima of some algal bromoperoxidases. (Source: Ref. 9.)

B.

Functional Group Modifications

The following section summarizes information on the reactions of organic substrates with marine bromoperoxidases. Examples are taken from published literature and from the authors’ laboratory. It is important to note that, in most cases, a single set of reaction conditions was used, so any yields shown are not optimized for the particular substrate. It is assumed that all appropriate controls have been performed with each substrate, to ensure that the product is not spontaneously formed. 1. Alkenes and Alkynes (Table 2) The primary products obtained from reaction of alkenes with bromoperoxidases are bromohydrins. The mechanism is probably via electrophilic addition of the bromonium ion (or equivalent) to the alkene, followed by the addition of a nucleophile. However, although the Markovnikov regiochemistry is typically observed, in several cases, anti-Markovnikov products are also found [35,36]. This suggests that the enzyme is able to orient the substrate to some extent before the addition occurs, and thus impart some degree of regioselectivity.

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Table 2 Alkene and Alkyne Substrates Substrate

Product(s)

Propene 9

:

Cinnamyl alcohol

[α] D ⫽ 0

Cinnamic acid

[α] D ⫽ 0

Styrene

cis-Propenyl phosphonic acid Geraniol

Reference

Laurencia pacifica Corallina offinalis

36

Corallina sp.

37

C. pilulifera

38

C. pilulifera

38

C. pilulifera

38

C. pilulifera

38

C. pilulifera

38

Corallina vancouveriensis

35

1

Allyl alcohol

Cyclohexene

Enzyme source

[α]D ⫽ 0

[α]D ⫽ 0

[α]D ⫽ 0

X ⫽ Br, Y ⫽ OH X ⫽ OH, Y ⫽ Br

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Table 2 Continued Substrate

Product(s)

Linalool X ⫽ Y ⫽ Br X ⫽ OH, Y ⫽ B X ⫽ BR, Y ⫽ OH

α-Pinene a

a

Enzyme source

Reference

Corallina vancouveriensis

35

Corallina vancouveriensis

35

See Fig. 7.

Dibromides and epoxides are also found in alkene product mixtures. By using a mixture of chloride and bromide in a ratio of 500: 1 (mimicking the concentration in seawater), Geigert et al. [36] were able to produce mixed dihalides (Table 2). Epoxides are also seen as products of alkenes, such as geraniol, which are more likely derived from intramolecular cyclization of the bromohydrins. α-Pinene yielded several ring-opened products when reacted with Corallina vancouveriensis bromoperoxidase (Fig. 7) [35]. Itoh et al. measured the optical activity of four halohydrin products that contained chiral centers and found that the [α] D was 0° in all cases [38]. When reacted with the bromoperoxidase from C. vancouveriensis, α-pinene and linalool both result in a surprising array of products. Some of the identified products are shown in Table 2; a scheme explaining the origin of the pinene products is also shown (Fig. 7). Enzymatic products of linalool also possess optical rotation measurements of 0 [35]. Alkynes appear to be poor substrates for bromoperoxidases. Only phenyl acetylene was found to react with C. officinalis enzymes, although the product was not structurally characterized and may involve reaction with the aromatic ring, and not the alkyne itself [38]. 2. β-Diketones and Enolizable Carbonyl Compounds (Table 3) The principal assay used by all researchers to quantify the activity of a haloperoxidase is based on the α-halogenation of monochlorodimedone (2-chloro-3,3dimethyl-1,5-cyclohexanedione, or MCD), shown in Figure 3. In aqueous buffer

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Figure 7 Proposed mechanism of formation of α-pinene products.

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Table 3 β-Diketones and Enolizable Ketones Substrate

Product(s) CHBr3

Enzyme source

Reference

Penicillus capitatus

39

3-oxo-octanoic acid R ⫽ CH 2 Br CHBr2 CBr3 3-oxo-octanoic acid

Monochlorodimedone (MCD) a

CHBr3 CH 2 Br2

Dihalodimedone

a

Bonnemassonia hamifera

Standard assay; see Fig. 3

7

10

The mono- and dibromoheptanones were also converted to the tribromoheptanone, but the formation of CHBr 3 is nonenzymatic.

solutions, the enol form, which has an ⑀ value of 20,000/M-cm at 290 nm, predominates. If a haloperoxidase is added to a solution of MCD, hydrogen peroxide, and bromide, the dihalo product is formed, which has an ⑀ of 200/M-cm. One unit (U) of bromoperoxidase activity is defined as the amount of enzyme necessary to convert one micromole of MCD to dihalo product at 25°C in 1 min [10]. The role of bromoperoxidases in the biosynthesis of brominated carbonyl compounds was investigated by Theiler et al. [7]. When reacted with 3-oxooctanoic acid, the enzyme from Bonnemaissonia hamifera yielded a mixture of dibromomethane, bromoform, and 1-bromopentane. In subsequent studies, the bromoperoxidase from Penicillus capitatus produced brominated heptanones and bromoform from the same substrate, which were similar or identical to compounds found naturally in B. hamifera. The authors suggest that the difference in product formation may be due to either distinctive catalytic abilities of each or two simultaneous reaction pathways [39]. The ready formation of bromoform by action of bromoperoxidases has led to the suggestion that these enzymes, through their natural catalytic activity with simple carbonyl compounds, are responsible for a significant percentage of atmospheric halomethanes, which are contributing factors to the degradation of the earth’s ozone layer. Indeed, bromoperoxidases from marine algae are suggested to be ultimately responsible for the formation of significant quantities of volatile

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organohalogen compounds that are found in the atmosphere [40,41], and it has been suggested that these compounds are responsible for some damage to the earth’s ozone layer [42,43]. 3. Aromatic Compounds (Table 4) Benzene rings containing substitutents that activate the ring toward electrophilic aromatic substitution (e.g., phenols) react readily with bromoperoxidase, yielding brominated products in good to high yields. Compounds with strongly deactivating substituents alone are not reactive, but if present with an activating substituent (e.g., OH), may yield a brominated product. In all cases, the regiochemistry appears to follow the expected patterns for electrophilic aromatic substitution. Multiple brominated products are possible, but the yields and ratio of single vs. multiple halogenated products may vary widely with reaction conditions. 4. Heterocycles (Table 5) A variety of nitrogen heterocycles and one sulfur heterocycle have yielded products with bromoperoxidases. Itoh et al. have compared the relative halogenating abilities of fungal chloroperoxidase and the bromoperoxidase from the alga Corallina pilulifera and report some significant differences between them. Using these enzymes, they were able to produce halogenated derivatives of nucleoside and nucleotide bases, which are known to have activity as potential anticancer agents. In this study, the reactions of the same substrates with fungal chloroperoxidase were compared. With the exception of the chlorinated products, most of the brominated and iodinated nucleic acid base products were similar. However, the report indicates several practical differences that favor the use of bromoperoxidase over the fungal enzyme [44]. Barbituric acid and derivatives were brominated by the enzyme from Ascophyllum nodosum to only mono- or dibromo products [45]. 5. Amino Acids and Proteins (Table 5) Using the bromoperoxidase from the brown alga Ascophyllum nodosum, a protein was successfully radiolabeled with 77 Br and the conditions were optimized [46]. This represents a novel technique for labeling of therapeutically useful proteins. The amino acid derivative methoxytyrosine yielded interesting decarboxylated nitrile and aldehyde products [47]. 6. Role of Bromoperoxidase in the Cyclization of a Complex Natural Product Elegant studies by Fukuzawa et al. [8] have demonstrated that the bromoperoxidase from the red alga Laurencia nipponica is involved in the final stages of the

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Table 4

Aromatic Compounds

Substrate

Product(s)

Phenol

2,4,6-Tribromophenol

Anisole

o-Bromoanisole p-Bromoanisole 2,4,6-Tribromophenol

o-Hydroxybenzyl alcohol m-Hydroxybenzyl alcohol p-Hydroxybenzyl alcohol

Enzyme source

3-Hydroxy-2,4,6-tribromobenzyl alcohol 4-Bromo-3-hydroxybenzyl alcohol 2,6-Dibromo-4-hydroxy benzyl alcohol

Methyl salicylate

Reference

Corallina pilulifera C. vancouveriensis C. pilulifera

20 33 38

Amphiroa ephedraea Corallina pilulifera C. vancouveriensis C. vancouveriensis

20 33 33

Rhodomela larix Ptychodera flava C. vancouveriensis

22

C. vancouveriensis

33

C. pilulifera

38

33

(91% yield)

Thymol

56 1-Methoxynaphthalene

Phenol red

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Bromphenol blue

:

31

Most bromoperoxidases (may be used as a general indicator)

9

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C

Table 5

Heterocyclic Compounds

Substrate

Product(s)

Enzyme source

Reference

Thiophene

C. pilulifera

38

Pyrazole

C. pilulifera

44

‘‘V-bromoperoxidase’’ (from marine alga)

31

A. nodosum

45

R ⫽ Br, I

Indoles R ⫽ Me, Phenyl

Barbituric acid and derivatives

R1 ⫽ R2 ⫽ R3 ⫽ H R1 ⫽ R2 ⫽ H Nucleic acid bases and nucleosides Cytosine 5-Bromocytosine C. pilulifera Uracil 5-Bromouracil, 5-iodouracil Cytidine 5-Bromocytidine Thymine (Decomposes) (No reaction with adenine, adenosine, guanine, and 2-deoxyuridine) Amino acid P. capitatus R ⫽ CH 2-CN CH2-CHO

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44

47

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C

biosynthesis of cyclic metabolites that occur naturally in the organic extract. Using laurediols (also found as trace metabolites in L. nipponica) as substrate for both lactoperoxidase and later the actual bromoperoxidase from the Laurencia, it was demonstrated that each laurediol isomer yielded a cyclized product that was identical to a natural product found in the alga (Fig. 8). This is the first example in which a direct bromonium-induced cyclization is clearly demonstrated to be catalyzed by a bromoperoxidase. Many halogenated marine-derived products may also be envisioned to be derived from halonium ion cyclization [48].

IV.

CONCLUSIONS

This review indicates bromoperoxidases from marine organisms are capable of halogenating a wide variety of organic substrates, often in high yields. The general mechanism appears to be via electrophilic substitution or addition routes, and this is supported by most of the regiochemistry that is observed in the products. However, in the reaction with certain alkenes, anti-Markovnikov products are observed along with Markovnikov products, suggesting some difference from the ‘‘normal’’ chemical route to this functionality. Although stereoselectivity has not yet been observed through the use of bromoperoxidases, it is also clear that the development of these enzymes as chemoenzymatic reagents still requires a great deal of effort. Most studies (including those from our laboratory) report the use of these reactions under only one set of reaction conditions. Relatively little work has been done in the optimization of the reaction conditions under which bromoperoxidases might show enhanced selectivity. As an example of the critical role of these parameters, Allain et al. [49] have shown that chloroperoxidase may be induced to produce chiral epoxides in high enantiomeric excess from disubstituted alkenes, but only, if the rate of hydrogen peroxide is precisely controlled. Subsequent work with chloroperoxidase has demonstrated that chiral epoxides with a high level of enantioselectivity may be formed with certain alkene substrates [50,51]. The catalytic activity of the bromoperoxidases involves numerous experimental parameters and will require a significant amount of effort to determine whether enantioselectivity can be shown in the laboratory. Bromoperoxidases are widespread in the marine environment, with the marine algae probably being the largest natural resource of these enzymes. The bromoperoxidase from Corallina officinalis is now offered commercially from Sigma Chemical Co., and recent reports on the immobilization of this enzyme on agarose have appeared [34]. Numerous other sources of bromoperoxidases are known but have not been investigated in any detail (or reported in the literature) [9,17]. No reports of bromoperoxidases from marine fungi or marine bacteria have appeared, although halogenated metabolites have been reported from both groups. This is probably due to lack of investigation, rather than absence

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Figure 8 Cyclization of laurediols by Laurencia nipponica bromoperoxidase. (Source : Ref. 8.)

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of the enzyme since bromoperoxidases are known from Pseudomonas sp., Streptomyces sp., and other bacteria [4,52], and fungi are the traditional source of chloroperoxidases. Genes encoding for bromoperoxidases have been cloned [53], and thus genetic manipulation to produce quantities of these enzymes is feasible. Bromoperoxidases have a significant advantage over chemical halogenation methods in organic synthesis. In contrast to classical halogenation methods (e.g., Br 2 in CCl 4), the bromoperoxidases are extraordinarily simple to use and require no unusual precautions. The most hazardous component of the reaction system is hydrogen peroxide (usually diluted). The workup of the reaction merely involves extraction of the product by an organic solvent. The resulting aqueous solution may then be relatively easily disposed. Because the bromoperoxidase reaction involves reagents that are relatively innocuous, it may be considered an ‘‘environmentally benign’’ process, especially when compared to typical chemical methods to achieve the same synthetic goals. Nearly all of the conventional chemical halogenation reactions require either a significant level of caution and/or special disposal arrangements to minimize environmental contamination. Especially for industrial applications, the use of enzymes may offer a beneficial alternative to chemical methods. One potential application of bromoperoxidase is the radiolabeling of proteins for diagnostic application using 77 Br. [54] One major benefit of the use of bromoperoxidases will be realized if stereoor regiochemistry can be controlled for specific substrates, reflecting the natural ability of these enzymes. It is clear that many experimental attributes of bromoperoxidases are not yet fully understood, yet these enzymes are undoubtedly involved in the biosynthesis of chiral halogens in natural products. By analogy with widely used enzymes such as lipases and esterases, bromoperoxidases from distinct biological sources likely have similar gross catalytic properties but may differ in the exact nature of their interaction with a specific substrate. This uniqueness has yet to be taken advantage of by those investigating these potentially highly useful enzymes. The role of enzymes in organic synthesis is rapidly growing in both research and industrial applications, as the versatility of enzymes and methodology for their use become apparent. Monographs dedicated to the practical use of enzymes in organic synthesis are published with regular updates, which include ‘‘recipes’’ and protocols for their use. One group of enzymes that are certain to gain greater use and interest in the near future are the bromoperoxidases.

ACKNOWLEDGMENTS The authors wish to gratefully acknowledge the financial support of the National Institutes of Health (GM-38040), the CSU Awards for Research, and the Tropical Technology Center (Okinawa, Japan).

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REFERENCES 1. G. W. Gribble, J. Nat. Prod., 55: 1353 (1992). 2. G. W. Gribble, Progress in the Chemistry of Organic Natural Products, Vol. 68 (W. Herz, et al., eds.), Springer, New York (1996). 3. E. D. Goldberg, The Sea, Vol. 2 (M. N. Hill, ed.), Wiley-Interscience, New York (1963). 4. K.-H. van Pee, Annu. Rev. Microbiology, 50: 375 (1996). 5. S. L. Neidleman and J. Geigert, Endeavour, 11: 1 (1987). 6. (a) S. L. Neidleman and J. Geigert, Biohalogenation: Principles, Basic Roles and Applications, Ellis Horwood Press (Wiley), New York (1986). (b) S. L. Neidleman and J. Geigert, Biochem. Soc. Symp., 48: 39 (1983). 7. R. F. Theiler, J. S. Siuda, and L. P. Hager, in Food-Drugs from the Sea (P. N. Kaul and C. J. Sindermann, eds.) Marine Technology Society, Norman, Okla. (1978). 8. A. Fukuzawa, M. Aye, Y. Takasugi, M. Nakamura, M. Tamura, and A. Murai, Chem. Lett., 2307 (1994). 9. C. A. Moore and R. K. Okuda, J. Nat. Toxins, 5: 295 (1996). 10. D. R. Morriss and L. P. Hager, J. Biol. Chem., 421: 1763 (1966). 11. A. Zaks and D. R. Dodds, J. Am. Chem. Soc., 117: 10419 (1995). 12. D. G. Baden and M. D. Corbett, Comp. Biochem. Physiol., 64B: 279 (1979). 13. Y. P. Chen, D. E. Lincoln, S. A. Woodin, and C. R. Lovell, J. Biol. Chem., 266: 23909 (1991). 14. D. J. Sheffield, T. Harry, A. J. Smith, and L. J. Rogers, Phytochem., 32: 21 (1993). 15. M. J. Clague and A. Butler, Adv. Inorg. Chem., 9: 219 (1994). 16. M. Andersson, V. Conte, F. Di Furia, and S. Moro, Tetrahedron Lett., 36: 2675 (1995). 17. W. D. Hewson and L. P. Hager, J. Phycol., 16: 340 (1980). 18. B. E. Krenn, H. Plat, and R. Wever, Biochim. Biophys. Acta, 912: 287 (1987). 19. H. Yu and J. W. Whittaker, Biochem. Biophys. Res. Comm., 160: 87 (1989). 20. H. Yamada, N. Itoh, S. Murakami, and Y. Izumi, Agric. Biol. Chem., 49: 2961 (1985). 21. M. Pedersen, Physiol. Planta, 37: 6 (1976). 22. T. J. Ahern, G. G. Allan, and D. G. Medcalf, Biochim. Biophys. Acta, 616: 329 (1980). 23. H. Vilter, Phytochem., 23: 1387 (1984). 24. H. S. Soedjak and A. Butler, Biochim. Biophys. Acta, 1079: 1 (1991). 25. J. A. Manthey and L. P. Hager, J. Biol. Chem., 260: 9654 (1985). 26. D. G. Baden and M. D. Corbett, Biochem. J., 187: 205 (1980). 27. R. Jannun and E. L. Coe, Comp. Biochem. Physiol., 88B: 917 (1987). 28. T. Higa, in Marine Natural Products: Chemical and Biological Perspectives, Vol. 5 (P. J. Scheuer, ed.), Academic Press, New York (1981). 29. H. S. Soedjak and A. Butler, Biochem., 29: 7974 (1990). 30. A. Butler and J. V. Walker, Chem. Rev., 93: 1937 (1993). 31. R. A. Tschirret-Guth and A. Butler, J. Am. Chem. Soc., 116: 411 (1994). 32. H. S. Soedjak, J. V. Walker, and A. Butler, Biochem., 34: 12689 (1995). 33. M. Shang, R. K. Okuda, and D. R. Worthen, Phytochem., 37: 307 (1994).

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34. 35. 36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50. 51. 52. 53. 54.

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D. J. Sheffield, T. R. Harry, A. J. Smith, and L. J. Rogers, Phytochem., 38: 1103 (1995). C. A. Moore and R. K. Okuda, unpublished results. J. Geigert, S. L. Neidleman, S. K. DeWitt, and D. J. Dalietos, Phytochem., 23: 287 (1984). S. L. Neidleman, W. F. Jr. Amon, and J. Geigert, U.S. Patent 4,247,641, Cetus Corp. (1981). N. Itoh, A. K. M. Q. Hasan, Y. Izumi, and H. Yamada, Eur. J. Biochem., 172: 477 (1988). R. S. Beissner, W. J. Guilford, R. M. Coates, and L. P. Hager, Biochem., 20: 3724 (1981). R. Theiler, J. C. Cook, L. P. Hager, and J. F. Siuda, Science, 202: 1094 (1978). B. Walter and K. Ballschmiter, Chemosphere, 22: 557 (1991). N. Itoh and M. Shinya, Marine Chem., 45: 95 (1994). R. M. Moore, M. Webb, R. Tokarcyzk, and R. Wever, J. Geophys. Res., 101(C9): 20,899 (1996). N. Itoh, Y. Izumi, and H. Yamada, Biochem., 26: 282 (1987). M. C. R. Franssen, J. D. Jansma, H. C. van der Plan, E. de Boer, and R. Wever, Bioorg. Chem., 16: 352 (1988). F. Lambert and G. Slegers, Appl. Radiat. Isot., 45: 11 (1994). M. Nieder and L. P. Hager, Arch. Biochem. Biophys., 240: 121 (1985). W. Fenical, J. Phycology, 11: 245 (1975). E. J. Allain, L. P. Hager, L. Deng, and E. N. Jacobsen, J. Am. Chem. Soc., 115: 4415 (1993). A. F. Dexter and L. P. Hager, J. Am. Chem. Soc., 117: 817 (1995). F. J. Lakner, K. P. Cain, and L. P. Hager, J. Am. Chem. Soc., 119: 443 (1997). N. Itoh, N. Morinaga, and A. Nomura, Biochim. Biophys. Acta, 1122: 189 (1992). S. J. Facey, F. Grob, L. C. Vining, K. Yang, and K. H. van Pee, Microbiology, 142: 657 (1996). F. Lambert and G. Slegers, Appl. Radiat. Isot., 45: 11 (1994).

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4 LC-Hyphenated Techniques in the Search for New Bioactive Plant Constituents Kurt Hostettmann, Maryse Hostettmann, Sylvain Rodriguez, and Jean-Luc Wolfender Universite´ de Lausanne, Lausanne, Switzerland

I. INTRODUCTION The use of medicinal plants for curing illnesses can be traced back over five millennia to written documents of the early civilizations in China, India, and the Near East, but it is doubtless an art as old as mankind. Even today, plants are a major and significant source of drugs for the majority of the world’s population. Although in industrialized countries medicinal plant research has faced cyclic fortune during the last few decades [1], nonetheless, substances derived from higher plants still constitute ca. 25% of prescribed medicines [2,3]. The plant kingdom represents an extraordinary reservoir of novel molecules. Of the estimated 400,000–500,000 plant species around the globe, only a small percentage has been investigated phytochemically and the fraction submitted to biological or pharmacological screening is even lower. Since plants may contain hundreds, or even thousands, of metabolites, there is currently a resurgence of interest in the vegetable kingdom as a possible source of new lead compounds for introduction into therapeutic screening programs. Furthermore, the rapid disappearance of tropical forests and other important areas of vegetation has added renewed pressure for the rapid isolation and identification of bioactive natural products. In general, the approach adopted to obtain an exploitable pure plant constit-

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uent involves interdisciplinary work in botany, pharmacognosy, pharmacology, chemistry, and toxicology and can be formulated as follows [1] (Fig. 1): Selection, collection, botanical identification, and preparation of plant material Extraction with suitable solvents and preliminary analysis Biological and pharmacological screening of crude extracts Chromatographic separation of pure bioactive constituents guided by bioassay Structure determination Analysis and pharmacological profile of pure compounds Toxicological testing Partial or total synthesis Preparation of derivatives for structure–activity relationships By following this approach alone, there is a very high risk of unnecessarily isolating bioactive, but known plant constituents. Furthermore, interesting lead compounds that do not exhibit the tested activity will simply be missed. In order to eliminate the time-consuming isolation of known constituents, hyphenated techniques such as liquid chromatography–mass spectrometry (LC/MS), LC– ultraviolet spectroscopy (LC/UV), and recently even LC–nuclear magnetic resonance (LC/NMR) imaging are being introduced at the earliest stage of separation to screen the crude extracts spectroscopically (Fig. 1) [4]. Indeed, efficient detection and rapid characterization of natural products play important roles as analyti-

Figure 1 Procedure for obtaining active principles from plants and use of LC hyphenated techniques as strategic analytical screening tools during the isolation process of a plant extract. LC, liquid chromatography.

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cal support in the work of phytochemists. This type of analysis is valuable for detecting compounds with interesting structural features and targeting their isolation. In this chapter, the role of the LC-hyphenated screening of the crude extracts used as a complement to the biological screening will be particularly emphasized.

II. HYPHENATION IN HIGH-PERFORMANCE LIQUID CHROMATOGRAPHY High-performance liquid chromatography (HPLC) is used routinely in phytochemistry to ‘‘pilot’’ the preparative isolation of natural products (optimization of the experimental conditions, checking of the different fractions throughout the separation) and to control the final purity of the isolated compounds [5–8]. For chemotaxonomic purposes, the botanical relationships among different species can be shown by chromatographic comparison of their chemical compositions. Comparison of chromatograms, used as fingerprints, of authentic samples and of unknowns permits identification of drugs and/or their adulteration. The selective detection of a given product in a complex mixture allows good quantitative measurement, as well as precise chemotaxonomic comparisons. Furthermore, in many applications, it may be necessary not only to detect but also to identify compounds in extracts. With conventional detection methods such as monochromatic UV detection, the identity of peaks can be confirmed only from their retention times by comparison with those of authentic samples. In order to get more information on the metabolites of interest, there is a need for a multiple detection system that takes advantage of chromatography as a separation method and spectroscopic techniques as detection and identification methods. At present, a number of important hyphenated techniques are available, the application of which depends upon the types of problems that have to be solved (Fig. 2). In our laboratory, for the screening of crude plant extracts, HPLC coupled with UV photodiode array detection (LC/DAD-UV) and LC/MS have been mainly used [4,9,10]. Recently, several studies have also shown that HPLC coupled with nuclear magnetic resonance (LC/NMR) is a very powerful complementary technique for on-line plant metabolite identification [11,12]. This new tool has been configured into our LC/UV/MS systems [13]. The use of several complementary detection techniques is necessary to obtain a complete picture of the extract composition. Indeed, a crude plant extract is a very complex mixture containing sometimes hundreds or thousands of different metabolites [1]. The chemical nature of these constituents varies considerably within a given extract, and the variability of the physicochemical and spectroscopic properties of these compounds causes numerous detection problems. Although different types of LC detectors such as UV, RI, fluorescence, electrochemical, and evaporative light scattering exist, none of these detectors permits within

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Figure 2 Summary of different coupling possibilities for chromatographic separation methods with spectroscopic techniques. GC, gas chromatography; SFC, supercritical fluid chromatography; HPLC, high-performance liquid chromatography; CE, capillary electrophoresis; TLC, thin layer chromatography; UV/VIS, ultraviolet/visible spectroscopy; FT-IR, Fourier transform infrared spectroscopy; MS, mass spectrometry; NMR, nuclear magnetic resonance.

the same analysis the detection of all the secondary metabolites encountered in a plant extract; each method has its own selectivity. A.

Liquid Chromatography/Ultraviolet Photodiode Array

For more than a decade HPLC coupled with UV photodiode array detection has been used by phytochemists for screening extracts [14–16]. The UV spectra of natural products often give useful information on the type of constituents and

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also, as in the case of polyphenols, on the oxidation pattern. New instruments allow storage of UV spectra of reference compounds in databases and computer matching can be realized automatically when screening for known constituents. The LC/DAD-UV technique has been used extensively for the screening of polyphenols [17,18], in combination with the postcolumn addition of classical UV shift reagents, allowing the determination of the positions of free hydroxyl groups on the polyphenol nucleus by comparison of the spectra obtained with each of the specific reagents [14,19]. The equipment used for this purpose is shown schematically in Fig. 3. Reagents are added to the effluent, leaving the HPLC column by means of a low-volume mixing tee. Diode array detection is then performed on the modified mobile phase. A weak base (sodium acetate) deprotonates only the more acidic phenolic groups, whereas a strong base (sodium methoxide, potassium hydroxide) reacts with all phenolic groups. The shift reagents AlCl 3 and H 3 BO 3 can also be employed and are compatible with the acidic acetonitrile–water eluent system. As disodium hydrogen phosphate is unsuitable as a weak base because of solubility problems in aqueous organic systems (for example, with an acetonitrile–water eluent), it can be replaced by sodium acetate (0.5 M ). However, this reagent is insufficient to deprotonate acidic phenolic groups in an aqueous solvent system, and NaOH (0.01 M ) may be added via a second HPLC pump to obtain a pH of 8 for the solvent exiting the HPLC column (Fig. 3). With the combination NaOAc-H 3 BO 3 it is not always possible to locate ortho-dihydroxyl groups in the acetonitrile–water system. This problem

Figure 3 Schematic representation of the experimental setup used for the postcolumn addition of UV shift reagents. UV, ultraviolet.

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can be solved by using AlCl3 in the acidified mobile phase (0.1% trifluoroacetic acid [TFA], pH 3). Similar shifts are obtained with AlCl 3-HCl in methanol, the reagent typically used in the characterization of flavonoids. By comparing online UV spectra with the addition of aluminum chloride at pH 7 and pH 3, it is possible to detect labile ortho-dihydroxyl complexes [14,18]. However, LC/DAD-UV is limited to the detection of UV-active constituents. The information recorded on-line is generally not sufficient for a complete identification, and other on-line complementary spectroscopic information is needed. B.

Liquid Chromatography/Mass Spectrometry

Although LC/UV has often been used for the analysis of crude plant extracts, LC/MS has been introduced more recently; its use is still not widespread in the phytochemical community [10]. At present, MS is one of the most sensitive methods of molecular analysis. Moreover, it has the potential to yield information on the molecular weight as well as on the structure of the analytes. Because of its high power of mass separation, very good selectivities can be obtained. For volatile, nonpolar compounds from essential oils, MS detection used in combination with gas chromatography (GC) separation has been applied successfully [20]. Indeed, GC/MS has become essential for applications in this field and has already been used routinely for two decades as the ideal GC detector [21]. The coupling between LC and MS has not been straightforward since normal operating conditions of a mass spectrometer (high vacuum, high temperatures, gas-phase operation, and low flow rates) are diametrically opposed to those used in HPLC, namely, liquid-phase operation, high pressures, high flow rates, and relatively low temperatures [22]. Because of the basic incompatibilities between HPLC and mass spectrometry (MS), on-line coupling of these instrumental techniques has been difficult to achieve. To cope with these different problems, a considerable number of LC/MS interfaces have been developed. These LC/MS interfaces must accomplish nebulization and vaporization of the liquid, ionization of the sample, removal of excess solvent vapor, and extraction of the ions into the mass analyzer. Thus, different techniques may involve variations in the methods by which these steps are accomplished. The most commonly used LC/MS interfaces are particle beam (PB) [23], thermospray (TSP) [24], atmospheric pressure chemical ionization (APCI) [25], and electrospray (ES) [26]. Each of these interfaces has its own characteristics and range of applications. With the appropriate LC/MS configuration, the analysis of small nonpolar molecules to very large polar molecules such as proteins is now possible. A number of LC/MS interfaces are suitable for the analysis of plant secondary metabolites. In our approach to LC/MS, mainly used for the HPLC screening

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of crude plant extracts, three interfaces, thermospray (TSP) [24], continuous flow fast-atom bombardment (FAB) (CF-FAB) [27], and electrospray (ES) [26], have been investigated [28]. They cover the ionization of relatively small nonpolar products (aglycones, 200 µ) to more polar molecules (glycosides, 2000 µ) (Table 1). In our laboratory, thermospray is the most widely used interface. 1. Thermospray Liquid Chromatography/Mass Spectroscopy The TSP interface [2] has most often been used for natural product analysis in crude plant extracts [10,29], and it has emerged as one of the most popular LC/ MS interfaces, although ES is also gaining more acceptance. Thermospray allows introduction of the liquid aqueous phase into the MS at a flow rate (about 1–2 mL/min) compatible with that usually employed for the standard HPLC conditions (organic solvents and aqueous mixtures in either isocratic or gradient mode). Thermospray operation provides mass spectra that are closely related to chemical ionization (CI solvent mediated) and is thus well suited for the analysis of moderately polar molecules in a mass range from 200 to 1000 µ or more in certain cases. In the TSP system, the LC effluent is partially vaporized and nebulized in the heated vaporizer probe to produce a supersonic jet of vapor containing a mist of fine droplets or particles. As the droplets travel at high velocity through the heated ion source, they continue to vaporize because of rapid heat input from the surrounding hot vapor. A portion of the vapor and ions produced in the ion source escapes into the vacuum system through a sampling cone, and the remainder of the excess vapor is pumped away by a mechanical vacuum pump [30] (Fig.

Table 1 Characteristics of the Liquid Chromatography/Mass Spectroscopy (LC/MS) interfaces: Thermospray (TSP), Continuous-Flow Fast-Atom Bombardment (CF-FAB), and Electrospray (ES) Interface

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TSP

CF-FAB

Flow Molecular weight Fragments

1–2 mL/min 200–800 µ Moderately polar

5–10 µL/min 500–2000 µ Polar

Liquid chromatography resolution Stability Operation

Good

Poor

Medium Many parameters to optimize

Poor Difficult to handle

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ES 1–1000 µL/min 200–500,000 µ Moderately polar and polar Excellent Good Easy, great influence of the modifiers

Figure 4 Schematic representation of a TSP–LC/MS interface. TSP, thermospray; LC, liquid chromatography; MS, mass spectrometry.

4). Four ionization modes exist with this interface. Besides chemical ionization initiated by conventional electron bombardment using a heated filament (‘‘filament-on’’ mode) or a discharge electrode (‘‘discharge-on’’ mode), a third ionization method, in which ammonium acetate or some other volatile buffer in the mobile phase plays a major role (‘‘thermospray buffer’’ ionization) is available; the fourth ionization mode is based on an ion evaporation mechanism. This latter mechanism may be operative in TSP buffer ionization mode as well [30]. The chemical process of ionization plays an important role and should always be taken into account when analyzing samples with LC/TSP-MS. Usually, as for CI, the values of the proton affinity of the analytes, the solvent, and the buffer should be considered. In our experience, for the analysis of plant metabolites (150–1000 µ, aglycones, mono- and diglycosides), the TSP operated mainly in the positive-ion mode in ‘‘thermospray buffer’’ ionization generally afforded MS spectra similar to those produced by (desorption chemical ionization (DCI) with NH 3 as reagent gas, positive-ion mode). In order to perform satisfactory LC/TSP-MS investigations of plant metabolites, or any other sample, different parameters should be optimized. Many interdependent experimental parameters are important in this respect: the mobile phase composition (amount and type of buffer and organic modifier), the solvent flow rate, the temperature of the vaporizer and the ion source, together with the geometry, the position, and the potential of the repeller electrode. The tuning of

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these parameters is of utmost importance in order to get high sensitivity and selectivity for a given type of analyte. Different articles have already emphasized the problems of thermospray optimization [31] and this particular aspect will not be further developed in detail here. C.

Liquid Chromatography/Nuclear Magnetic Resonance

Nuclear magnetic resonance (NMR) spectroscopy is by far the most powerful spectroscopic technique for obtaining detailed structural information about organic compounds in solution. Given a molecular mass, NMR spectroscopy can usually provide sufficient additional information to identify a completely unknown compound unambiguously [32]. Thus LC/NMR represents a potentially important complementary hyphenated technique to LC/UV and LC/MS. Proton-detected high-performance liquid chromatography/nuclear magnetic resonance (LC/ 1 H-NMR), despite being known for over 15 years [33], has not yet become a widely accepted technique, mainly because of its lack of sensitivity. However, the recent progress in pulse field gradients and solvent suppression, the improvement in probe technology, and the construction of high-field magnets have given a new impulse to this technique. Although LC coupling itself was rather straightforward compared to LC/ MS [30], one of the main problems of LC/NMR was the difficulty of observing analyte resonances in the presence of the much larger resonances of the mobile phase. This problem was compounded under LC conditions when more than one protonated solvent was used, and when solvent composition changed during gradient analysis. Furthermore, the continuous flow of sample in the detector coil complicated solvent suppression. These problems have now been overcome in part because of the development of fast, reliable, and powerful solvent suppression techniques such as the ‘‘water suppression enhanced through T 1 effect’’ technique (WET) [34], which produces high-quality spectra in both on-flow and stop-flow mode. These techniques consist of a combination of pulsed field gradients, shaped rf pulses, shifted laminar pulses, and selective 13 C decoupling and are much faster than classical presaturation techniques previously used [34]. Thus, in reversed-phase HPLC conditions, nondeuterated solvents such as MeOH or MeCN can be used, and water is replaced by D 2O. In contrast to conventional NMR probes, with LC/NMR the sample to be analyzed is not placed in a rotating 5-mm internal diameter (i.d.) tube. The mobile phase is flowing continuously through a nonrotating glass tube connected at both ends with HPLC tubing. The detection is made in most LC/NMR systems in flow cells of 60–180 µL with 2- to 4-mm i.d. placed in the active NMR coil region of a conventional probe body [32]. The choice of the volume of the flow cell is dependent on the sensitivity and/or the LC resolution required for each specific application.

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D.

Setup for Liquid Chromatography/Ultraviolet/Mass Spectroscopy and Liquid Chromatography/Nuclear Magnetic Resonance Analyses

In order to perform LC/UV/MS and LC/NMR analyses of crude plant extracts, a setup allowing the use of standard HPLC conditions (1 mL/min, gradient with aqueous solvent systems) with all the detectors has been used. An HPLC pump equipped with a gradient controller provides the eluent for the HPLC separation of the plant extracts (Fig. 5). A reversed-phase column (i.d. about 4 mm) is most often used. At the column outlet, the eluent passes through a photodiode array detector equipped with a high-pressure cell. For LC/MS, and with the aim of using the same columns without changing the chromatographic conditions, the buffer needed for the various LC/MS interfaces is added post column after the UV detector. For LC/TSP-MS, all the eluent enters the spectrometer, whereas for CF-FAB and ES, the eluent is split and the excess sample discarded (Fig. 5). Thus for LC/TSP-MS operation, postcolumn addition of the buffer needed for ‘‘TSP buffer’’ ionization is provided by an additional HPLC pump (usually ammonium acetate 0.5 M, 0.2 mL/min). The total eluent (1.2 mL/min) containing the buffer passes through the TSP interface, and the exhaust eluent is pumped away by a mechanical pump and trapped in a cold trap. For LC/CF-FAB-MS operation, the additional HPLC pump allows the postcolumn addition of the glycerol matrix needed for FAB ionization (usually

Figure 5 Schematic representation of the experimental setup used for LC/UV/MS (1) and LC/UV/NMR (2) analyses. LC, liquid chromatography; UV, ultraviolet; MS, mass spectroscopy; NMR, nuclear magnetic resonance.

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glycerol 50%, 0.2 mL/min). The viscous matrix is efficiently mixed into the eluent with a visco mixer. The mixture is then split with an accurate splitter (1/ 100), and only 10 µL/min of the total eluent enters the CF-FAB interface through a fused silica capillary. For LC/ES-MS operation, trifluoroacetic acid (TFA) or ammonium acetate is added into the eluent and the flow is split to leave ca. 200 µL/min flow rate entering the interface. For LC/NMR, no postcolumn addition is necessary and the water is replaced by D 2O. As with LC/MS, the separation is first monitored by a UV detector. In the on-line mode, all the eluent passes through the UV detector and then through the LC/NMR probe without splitting. In the stop-flow mode, the HPLC pump is stopped as soon as a peak of interest reaches the center of the NMR detection cell. A computer controlling the LC pump, the UV detector, and the NMR is used to synchronize this operation. The UV monitor is used to detect the peak and the HPLC is stopped after a delay corresponding to the time needed by this peak to reach the NMR cell. With such a setup, standard HPLC conditions for crude extract analysis (1 mL/min, 4 mm i.d. column) can be maintained without alteration. Furthermore, LC/DAD-UV detection is not affected by the buffer or the matrix used.

III. ROLE OF LIQUID CHROMATOGRAPHY/ULTRAVIOLET, LIQUID CHROMATOGRAPHY/MASS SPECTROSCOPY, AND LIQUID CHROMATOGRAPHY/NUCLEAR MAGNETIC RESONANCE AS SCREENING METHODS IN PHYTOCHEMICAL ANALYSIS When searching plant extracts for compounds with interesting properties, a multidimensional approach to their chromatographic analysis is of great significance. By combining HPLC with on-line UV, MS, and NMR, a large amount of preliminary information can be obtained about the constituents of an extract before isolation of the compounds of interest. In the case of polyphenols, for example, UV spectra recorded on-line give useful information (type of chromophore or pattern of substitution) complementary to that obtained with LC/MS and provide an initial assignment of the peak of interest at an early stage. When this information is not sufficient for a precise structural assignment, LC/NMR provides a useful complement for a full structural identification on-line. To illustrate this approach, examples of on-line LC/UV/MS, LC/MS/MS, and LC/NMR analyses of xanthones, flavones, and secoiridoids in crude extracts of plant species belonging to the Gentianaceae family will be discussed. Gentianaceae plants have indeed been extensively screened in our laboratory, especially in the search for novel antidepressive agents.

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A.

Search for Antidepressive Compounds from Gentianaceae

It is generally accepted that depression is characterized by low neurotransmitter concentration in the synapses. From a therapeutic viewpoint, there are two ways to increase the neurotransmitter concentration: slow down their recapture or inhibit their deamination. In this latter approach, a search for selective and reversible inhibitors of monoamine oxidase (MAO) has become of major importance [35,36]. Disturbances in monoamine oxidase levels have been reported in a series of disorders such as Parkinson’s disease, Huntington’s chorea, depression, anxiety, and senile dementias [37]. The enzyme monoamine oxidase (MAO) plays a key role in the regulation of certain physiological amines in the human body. It causes deactivation by deamination of neurotransmitters such as catecholamines and serotonin, and of endogenous amines such as tyramine. This enzyme exists as two isoenzymes, MAO-A and MAO-B, which exhibit different substrate specificities. Inhibitors of MAO, in particular of the A type isoenzyme, have potential as antidepressive drugs since they increase both noradrenaline and serotonin levels in the brain. There is current interest in the discovery of new reversible inhibitors of MAO [37] because existing drugs such as clorgyline (selectively inhibits MAO-A) and (⫺)-deprenyl (selectively inhibits MAO-B) are irreversible inhibitors and exhibit serious side effects. It is well known that plants contain inhibitors of MAO [38]. For example, the bark of the tropical liana Banisteriopsis caapi (Malpighiaceae) contains βcarboline alkaloids such as harmaline, a potent reversible inhibitor of MAO-A [39]. Infusions of St. John’s wort, Hypericum perforatum (Guttiferae), have been used for the treatment of melancholia in European folk medicine, and numerous preparations of this plant have now been commercialized for the management of different depressive states. The antidepressive activity of H. perforatum has been demonstrated in vivo [40], and it was originally thought that the anthrone dimer hypericin, which has shown MAO-A and MAO-B inhibitory activity in vitro [41], was responsible for this activity. However, more recent investigations point toward xanthones and flavonoids as the major source of the antidepressive activity since these have potent MAO inhibitory properties [42]. Investigations with extracts of Canscora decussata (Gentianaceae) [43] in vivo showed that the fractions containing xanthones had a stimulating effect on the central nervous system in mice, rats, and dogs [44]. A strong inhibition of monoamine oxidase (IMAO) by trisubstituted xanthones was observed by Suzuki and coworkers [45]. In fact, a number of xanthones that inhibit MAO have been identified from plant sources [46–48]. Of those that have been tested, the aglycones are, in general, more active than the corresponding glycosides. Xanthones with the substitution patterns 1-hydroxy-3,8-dimethoxy- and 1,3-dihydroxy-7,8-

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dimethoxy- showed pronounced activity. They were also selective inhibitors, being more effective against MAO-A than MAO-B. All 1,3,5,8-tetrasubstituted xanthones isolated from the aerial parts of Gentiana lactea (Gentianaceae) have been tested against monoamine oxidases from rat brain mitochondria. Two glycosides and the aglycone swerchirin (1,8-dihydroxy-3,5-dimethoxyxanthone) were inactive. Bellidifolin (1,5,8-trihydroxy-3-methoxyxanthone), on the other hand, inhibited MAO-A significantly. The activities of these xanthones were compared to those of specific synthetic inhibitors of MAO-A (clorgyline) and MAO-B (pargyline). The MAO-A activity of bellidifolin was of the same order of magnitude as that of pargyline, and 40 times less than that of clorgyline. It was also selective with a marked reduction in activity against MAO-B (IC 50 MAO-A/IC 50 MAOB ⫽ 0.001). Desmethylbellidifolin (1,3,5,8-tetrahydroxyxanthone) gave an inhibitory concentration (IC 50) against MAO-A that was approximately 10 times higher than that obtained with bellidifolin [48]. In a search for more active xanthones, other plants (especially of the families Gentianaceae, Guttiferae, and Polygalaceae) have been investigated. A comparison of the IMAO activity of some representative extracts showed, for example, that the dichloromethane extract of Chironia krebsii, a member of the Gentianaceae from Malawi, was much more active (100% IMAO-A at 15 µg/ mL) than that of Hypericum perforatum (18% IMAO-A at 15 µg/mL), which was of the same order of magnitude as that of the yellow gentian, Gentiana lutea (30% IMAO-A at 15 µg/mL) [49]. In order to find new xanthones, numerous extracts of Gentianaceae have been screened by LC-hyphenated techniques, and several examples of this research are described in the next section. B.

Liquid Chromatography/Ultraviolet/Mass Spectroscopy Analysis of the Aperitif Suze

One of the first gentian extracts to be analyzed by LC/UV/MS was not a crude plant extract but a beverage made with the roots of the yellow gentian G. lutea. Gentians contain bitter compounds belonging to the monoterpene glycosides (secoiridoids) and have been known since ancient time to stimulate appetite [50]. As it is known that yellow gentian also contains xanthones such as gentisin (1) that showed IMAO activity, it was of interest to check for the presence of 1 in the French aperitif called Suze. Direct injection of an aliquot of this beverage resulted in an LC/UV chromatogram with many different peaks, but only one having the characteristic xanthone UV spectrum (Fig. 6) [51]. As xanthones usually appear as aglycones or relatively small glycosides (mono- or diglycosides), the TSP interface was chosen for their ionization. After careful tuning of the TSP interface (especially the source and vaporizer temperatures), the peaks recorded by UV detection (254 nm) gave a clearly discernible MS response in the total ion current trace (TIC). The LC/TSP-MS spectrum of this peak exhibited a protonated mo-

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Figure 6 LC/UV analysis of the aperitif Suze  and UV and TSP-MS spectra of the only xanthone found in this beverage: gentisin (1). UV traces were recorded at 254 nm and UV spectra from 200 to 500 nm. HPLC: column, RP-18 NovaPak (4 µm, 150 ⫻ 3.9 mm i.d.); gradient, CH 3 CN-H 2 O (0.1% TFA) 5 :95 → 65: 35 in 50 min (1 mL/min), NH 4 OAc 0.5 M (0.2 mL/min post column). TSP: positive-ion mode; filament off; vaporizer 100°C; source 280°C. LC/UV, liquid chromatography/ultraviolet; TSP-MS, thermospray–mass spectrometry; HPLC, high-performance liquid chromatography; TFA, trifluoroacetic acid.

lecular [M ⫹ H] ⫹ ion at m/z 259. The molecular weight of this xanthone was thus 258 µ. As the mass of a bare xanthone nucleus is 196 µ, this indicated that the polyphenol was substituted by two hydroxyls and one methoxyl group. A comparison of the on-line UV spectra with those of trisubstituted xanthones from an in-house UV database confirmed that this compound was indeed gentisin, the main xanthone of the yellow gentian [52]. This example shows that for simple known compounds like xanthones, a precise identification can be performed online with the help of LC/UV and LC/MS, provided some information about the

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type of the constituents and their occurrence in given plant families is already available. C.

Liquid Chromatography/Ultraviolet/Mass Spectroscopy and Liquid Chromatography/Ultraviolet Analyses of Chironia krebsii Extracts with Postcolumn Addition of Ultraviolet Shift Reagents

The extracts of the roots of Chironia krebsii, a species from Africa, showed significant inhibition of MAO-A in vitro (CH 2Cl 2 ext. 100%; MeOH ext. 75% IMAO-A (15 µg/mL)). As mentioned, these activities were several orders of magnitude greater than those measured for H. perforatum extracts (CH 2Cl 2 ext. 18%; MeOH ext. 14% IMAO-A, 15 µg/mL) [49]. The LC/UV analysis of C. krebsii indicated that this plant was very rich in xanthones (see the structures in Fig. 7). The number of bands and the general aspects of the UV spectra of xanthones (UV spectra 4–15 in Fig. 8) allowed a first attribution of the type of oxygenation patterns encountered [51,53]. The constituents having a weaker chromophore (UV max. ca. 250 nm) were attributable to the secoiridoids swertiamarin (2) and sweroside (3), widespread bitter principles of the Gentianaceae. In order to get more information on the molecular weight, the molecular formula, and the sugar sequence (glycosides) of the xanthones, LC/TSP-MS analysis of the extract was carried out. The same LC conditions as used for the LC/UV/ MS analysis of the aperitif Suze were employed; LC/TSP-MS was performed in positive-ion mode with the filament off mode using ammonium acetate as buffer. The source was set at 280°C and the vaporizer at 100°C [18]. Under these conditions, the total ion current trace recorded with LC/TSP-MS showed ionization of all the peaks recorded in the UV trace of the extract (254 nm) (Fig. 8 and 9). Nevertheless, the total ion current responses for highly hydroxylated xanthones with high melting point temperatures (i.e., 11, Fig. 8) were found to be very weak in comparison to the UV trace of the corresponding peak, leading to a difficult MS detection [10]. The TSP mass spectra of the xanthone aglycones recorded on-line after HPLC separation exhibited only [M ⫹ H] ⫹ ions as the main peak (see TSP spectra of 15, Fig. 8, or 14 and 16, Fig. 9). Corresponding mass spectra of the xanthone glycosides usually showed two weak ions due to [M ⫹ H] ⫹ and [M ⫹ Na] ⫹ adducts, and a main peak for the protonated aglycone moiety [A ⫹ H] ⫹ (see TSP spectra of 9, Fig. 8, or 4 and 8, Fig. 9). In the case of diglycosides, successive losses of the monosaccharide units were marked by corresponding peaks in the spectrum. Thus, for the diglycosides of Chironia species, a loss of 132 µ was first observed, corresponding to a pentosyl residue, followed by a loss of 162 µ corresponding to a hexosyl moiety, leading to the aglycone ion [A ⫹ H] ⫹ (see

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Figure 7 Structure of the xanthones, flavones, and secoiridoids found in Chironia krebsii, Gentiana rhodantha, Halenia corniculata, and Swertia calycina.

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Figure 8 LC/UV and LC/TSP-MS analysis of the root methanolic extract of Chironia krebsii (Gentianaceae). UV spectra 2–3 are characteristic of secoiridoids. Peaks 4–11 are xanthone-O-glycosides; 12–15 are xanthone aglycones. LC/UV/MS: same conditions as for Fig. 6. LC/UV, liquid chromatography/ultraviolet; TSP, thermospray; MS, mass spectroscopy.

TSP spectra of 9, Fig. 8, or 4 and 8, Fig. 9). After subsequent isolation of these glycosides, the disaccharide was shown to be primeverose, a β-d-xylopyranosyl(1 → 6)-β-d-glucopyranoside disaccharide unit, often encountered in the Gentianaceae family [54]. Since MS detection allows the selective recording of each ion trace, it was possible to reconstruct the chromatogram of the ion m/z 657 and to obtain a precise assignment of the peak corresponding to compound 9 in the extract. Similarly, the specific ion trace at m/z 363 gave responses for the HPLC peaks of 9

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Figure 9 Summary of all the on-line spectral data obtained by LC/TSP-MS, LC/UV, and LC/UV with postcolumn addition of shift reagents for xanthones 4, 8, 14, and 16 from the root methanolic extract of Chironia krebsii (Gentianaceae). LC/UV/MS: same conditions as for Fig. 6. LC/UV/MS, liquid chromatography/ultraviolet/mass spectroscopy.

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with AlCl 3 reagent. In the case of 8, an important bathochromic shift was observed, characteristic of the chelated hydroxyl group at position C-1, whereas for 3 no such shift was recorded (Fig. 9: UV of 4 and 8). The primeverosyl moiety was thus attached at position C-1 in the case of 4 and at C-5 for 8. The corresponding aglycone of these two glycosides, 1,5-dihydroxy-3-methoxyxanthone (12) was also found as a slower eluting peak in the extract (Fig. 8), confirming the assignments of 4 and 8. Thus, the structures were established as 5-hydroxy-3-methoxy-1-O-primeverosylxanthone (4) and 1-hydroxy-3-methoxy5-O-primeverosylxanthone (8). In similar fashion, the structures of compounds 4–16 were also deduced (Fig. 7). Xanthones 4–16 were isolated [56] in order to test their IMAO activity. Several of them showed important selective and reversible inhibition of MAOA. The most active xanthone was 1,5-dihydroxy-3-methoxyxanthone (12) with an inhibitory activity of 0.04 µM (IC 50 MAO-A) [49]. The UV spectra of the pure products, fully identified, were registered in a UV database in order to permit a computer matching search against xanthones from other extracts and thus accelerate on-line identification of these compounds in other Gentianaceae species [57]. D.

Liquid Chromatography/Thermospray–Mass Spectroscopy and Liquid Chromatography/Continuous Flow–Fast-Atom Bombardment–Mass Spectroscopy of Gentiana rhodantha

Among the other Gentianaceae species screened by LC/UV/MS, Gentiana rhodantha from China presented a completely different extract composition profile from that found for C. krebsii. The LC/UV analysis of G. rhodantha showed the presence of only one predominant xanthone 17 and different secoiridoids. Compound 17 was easily identified as the widespread xanthone C-glycoside mangiferin (see structure in Fig. 7). The TSP-MS spectrum of 17 exhibited a pseudomolecular ion at m/z 423, characteristic fragments for C-glycosides at [M ⫹ H90] ⫹ and [M ⫹ H-120] ⫹, and a weak aglycone ion at m/z 261 (xanthone with four OH groups). The computer fit for the UV spectrum of 17 with our in-house UV spectral library allowed the definitive characterization of 17. With the help of the LC/TSP-MS spectra recorded on-line, chemotaxonomical considerations, and comparison with pure standards, the secoiridoids with retention times less than 10 min were also easily identified [58]; among them a very minor secoiridoid, sweroside (3)(MW 358), was found to be present. The slower eluting peaks 18 and 19 also exhibited the same characteristic UV spectra of secoiridoids (one band around 240 nm, Fig. 10). These compounds, which were less polar than common secoiridoids, were studied in more detail. The LC/ TSP-MS analysis of 18 and 19 in each case gave a spectrum identical to that

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(this ion is the main fragment of 9) and also of compound 15, a slower-running component of the extract (see ion traces in Fig. 8). According to the UV and TSP mass spectra of 15, this compound was readily identified as the corresponding free aglycone of 9. This information allowed the rapid identification of both aglycones and their corresponding glycosides simultaneously in a crude plant extract. In order to obtain more structural information on the position of the free hydroxyl groups on the xanthone nucleus, LC/UV with postcolumn addition of shift reagents was performed, using the setup described previously (see Fig. 3). The structural information obtained on-line by the combination of LC/UV, LC/ TSP-MS, and LC/UV with postcolumn addition of shift reagents is illustrated for four xanthones (4, 8, 14, and 16, Fig. 9). The two xanthone aglycones (14 and 16) exhibited almost identical UV spectra (Fig. 9). According to Kaldas [55] the UV spectra of 14 and 16 with four absorption maxima and a higher intensity for band II are characteristic for 1,3,7,8tetraoxygenated xanthones with a free hydroxyl at position C-1. The TSP-MS spectra recorded on-line permitted the molecular weight (MW) determination and the assignment of the number and type of substituents of 14 (MW 274: three OH, one OMe) and 16 (MW 302: one OH, three OMe, Fig. 9). The shifted UV spectra recorded on-line for 14 confirmed this compound to be 1,7,8-trihydroxy3-methoxyxanthone. Indeed, the presence of hydroxyl groups at positions C-1 and C-8 was characterized by the important bathochromic shift recorded with AlCl 3, and the ortho-dihydroxyl group at C-7 and C-8 was confirmed by the shift due to the complexation of boric acid. The position of the methoxyl at C-3 was confirmed by the absence of shift registered with the weak base NaOAc. The spectra of 16 with added KOH showed a significant decrease in band intensity and only a very small shift, indicating no free hydroxyl groups, with the exception of a chelated one. This was confirmed by the shift measured with AlCl 3. The spectra with added NaOAc and H 3BO 3 remained unchanged, confirming the structure of 16 to be 1-hydroxy-3,7,8-trimethoxyxanthone (Fig. 9). The two isomeric xanthone glycosides 4 and 8 also exhibited nearly identical UV spectra indicating the same oxygenation pattern (probably 1,3,5) [55]. The TSP-MS spectra of 4 and 8 were comparable: they both exhibited an [M ⫹ H] ⫹ pseudomolecular ion at m/z 553 with consecutive losses of 132 and 162 µ, leading to the same aglycone ion [A ⫹ H] ⫹ at m/z 259 (Fig. 9: TSP-MS of 4 and 8). The mass of the aglycone ion was characteristic of a xanthone with one methoxyl and two hydroxyl groups, and the presence of a disaccharide moiety consisting of a pentosyl and hexosyl subunits. The disaccharide was assumed to be primeverosyl since it is the only disaccharide of this mass that is known in the Gentianaceae family [54]. No shift was observed for either compound in the presence of weak base, indicating no free hydroxyl group at position C-3. The only difference between these two isomers was shown by the spectra recorded

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Figure 10 Combined TSP (a) and CF-FAB (b) LC/MS of the enriched n-BuOH fraction of the methanolic extract of Gentiana rhodantha (Gentianaceae). HPLC: column, RP-18 Novapak (4 µm, 150 ⫻ 3.9 mm i.d.); gradient, CH 3 CN-H 2 O (0.05% TFA) 5 :95 → 50: 50 in 30 min (0.9 mL/min). TSP (a): positive-ion mode; filament off; vaporizer, 90°C; source, 230°C; AcONH 4 0.5M (0.2 mL/min post column). CF-FAB (b): negative-ion mode; FAB tip 50°C; source 100°C; glycerol 50% (v/v) (0.15 mL/min post column); LC flow post column split 1:100. TSP, thermospray; CF-FAB, LC/MS, liquid chromatography; HPLC, high-performance liquid chromatography; TFA, trifluoroacetic acid.

obtained for 3; all exhibited an intense ion at 359 u and no ion at higher mass (Fig. 10). However, the chromatographic behavior was quite different for 18, 19, and 3. In order to obtain complementary information about these constituents, a second LC/MS analysis with CF-FAB was undertaken, using the same HPLC conditions. The total ion current recorded for the whole chromatogram showed a very important MS response for compounds 18 and 19, while the more polar metabolites were only weakly ionized (Fig. 10). The CF-FAB spectrum of 18 recorded on-line in negative-ion mode exhibited a very intense pseudomolecular

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ion [M ⫺ H] ⫺ at m/z 913 together with a weak ion at m/z 555 corresponding to the loss of a ‘‘swerosidelike’’ unit [M ⫺ H-358] ⫺ (CF-FAB spectrum of 18, Fig. 10b). This complementary information indicated clearly that the molecular weight of 18 was 914 u. For the same compound, only a fragment corresponding to a ‘‘sweroside-like’’ unit m/z 359 was recorded during the LC/TSP-MS analysis (TSP spectrum of 18, Fig. 10a). According to the different results obtained on-line for 18 in the HPLC screening of the extract of G. rhodantha, it was concluded that 18 was probably a type of moderately polar, large secoiridoid containing at least one unit very similar to 3. The CF-FAB spectrum of 19 exhibited an intense [M ⫺ H] ⫺ pseudomolecular ion at m/z 1629, indicating a molecular weight of 1630 for this compound. Fragments at m/z 1271, 913, and 555, respectively, showed the consecutive losses of ‘‘swerosidelike’’ units (⫺358 u). These on-line LC/MS results suggested that 19 was probably similar to 18 with two more ‘‘swerosidelike’’ units attached to it. After the LC/MS screening results, a targeted isolation of 18 and 19 was undertaken. A full structure determination of 18 with the help of one-dimensional (1D) and two-dimensional (2D) NMR experiments as well as with different chemical transformations showed that 18 consisted of two secoiridoid units linked together with a monoterpene unit through two ester groups (structure in Fig. 7) [58]. The full structure determination of 19 showed that this compound possessed two more secoiridoid units than 18, which were esterified on the free carboxylic functions of 18 [59]. These two compounds were found to be natural products of a new type. The structure determination of other closely related compounds from this extract is still in progress. This example illustrates the use of different LC/MS ionization techniques for targeting unknowns; LC/TSP-MS indicated that compounds 18 and 19 had common subunits (identical fragments m/z 359), and CF-FAB allowed the online molecular weight determination of all of these oligomeric compounds. The combination of both types of information has thus allowed the early recognition of this type of large secoiridoid glycoside. E.

Liquid Chromatography/Thermospray–Mass Spectroscopy and Liquid Chromatography/ Electrospray–Mass Spectroscopy of Halenia corniculata

As shown in the previous example, the use of different complementary MS ionization techniques is often important for unambiguous on-line determination of molecular weights. The analysis of another of the Gentianaceae species collected in Mongolia, Halenia corniculata, by TSP and LC/ES-MS also demonstrated the importance of an LC/MS analysis using two independent ionization methods for screening unknowns.

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The whole-plant MeOH extract of H. corniculata, which contains xanthones, flavones, and secoiridoid glycosides [57], was analyzed by both LC/TSPMS and LC/ES-MS. In the latter case, the analysis was carried out with TFA as buffer. Ionization of all the glycosides (20–30) found in the extract was observed in the negative ion (N.I.) LC/ES-MS (Fig. 11), but the response for the aglycones was very selective and most of them were not ionized [28]. The LC/ES-MS base peak trace showed a better chromatographic resolution than the corresponding LC/TSP-MS trace. The ES spectra of flavonoids 22, 23, and 30; xanthones 25– 29; as well as secoiridoid glycosides 20, 21, and 24 exhibited clearly discernible TFA anion adducts [M ⫹ CF 3 CO 2] ⫺, together with deprotonated ions [M ⫺ H] ⫺. No aglycone ions or other fragments were observed (see 22, Fig. 11). For polyphenols 22, 23, and 25–30, the positive ion (P.I.) TSP spectra recorded on-line exhibited [M ⫹ H] ⫹ ions together with base peak [A ⫹ H] ⫹ aglycone ions. In the case of diglycosides such as 22 (Fig. 11), the loss of the terminal sugar unit was also observed. This demonstrated that the spectra of a given compound such as 22 will differ considerably according to the ionization mode chosen. A good knowledge of the ionization process is thus necessary for a correct peak assignment [28]. In this example it was also shown that compound 24, a minor secoiridoid glycoside ester of the extract, was not ionized in TSP but in LC/ES-MS exhibited a deprotonated molecular ion [M-H] ⫺ at m/z 697 and a strong TFA adduct [M ⫹ CF 3 CO 2] ⫺ at m/z 811, indicating a molecular weight of 698 u (Fig. 12). The UV spectrum of 24 (λ max 219, 244, 297, 321 nm) differed from those usually encountered in Gentianaceae species extracts. The two UV bands, at 219 and 244 nm, together with the retention time (ca. 16 min), suggested that 24 was an iridoid-type compound. The two other maxima at 297 and 321 nm indicated the presence of an aromatic unit. No literature reference matched such a compound corresponding to this molecular weight. Thus, as in the case of G. rhodantha, a targeted isolation of 24 was undertaken [60]. Different hydrolyses of 24 followed by the HPLC of the degradation products showed the presence of a caffeoyl, a secoiridoid, and two hexosyl moieties. The structure was fully established by NMR experiments as 7-β-[4′-O-(β-d-glucopyranosyl)-trans-caffeoyloxy] sweroside, and this new natural product was named corniculoside [60]. F.

Liquid Chromatography/Ultraviolet/Mass Spectroscopy and Liquid Chromatography/Mass Spectroscopy/Mass Spectroscopy Analysis of the Methanolic Extract of Swertia calycina

Besides xanthones and secoiridoids, the LC/UV analysis of the methanolic extract of Swertia calycina, a Gentianaceae native of Rwanda in central Africa, revealed the presence of four flavonic constituents, 31–34 (Fig. 13) [61]. The TSP-MS spectra of these four flavonoids exhibited protonated molecular

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Figure 11 Combined LC/TSP-MS and LC/ES-MS of the MeOH extract of Halenia corniculata (Gentianaceae). HPLC: C18 NovaPak (4 µm, 150 ⫻ 3.9 mm i.d.); gradient, CH 3 CN-H 2O 5 :95 → 65:35 in 50 min (1 mL/min). Injection 200 µg. LC/TSP-MS (P.I.) and LC/ES-MS (N.I.) allowed the ionization of the different glycosides 20–30. Only the secoiridoid glycoside 24 was not ionized by TSP. As shown for the spectra of the flavonoid diglycoside 22, [M⫹H]⫹ ion and fragments (aglycone ion, sugar losses) were observed in LC/TSP-MS (P.I.), only [M⫺H] ⫺ and [M⫹CF 3 COO] ⫺ were visible in LC/ES-MS (N.I.). LC/TSP-MS, liquid chromatography/thermospray–mass spectroscopy; ES, electrospray; HPLC, high-performance liquid chromatography; P.I., positive ion; N.I., negative ion.

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Figure 12 LC/ES-MS analysis of the MeOH extract of Halenia corniculata (Gentianaceae) with the ES-MS and UV spectra of corniculoside (24). This compound was not recorded by LC/TSP-MS. LC: same conditions as for Fig. 6. LC, liquid chromatography; ES, electrospray; MS, mass spectroscopy; UV, ultraviolet; TSP, thermospray.

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Figure 13 (a) LC/TSP-MS and (b) LC/TSP-MS/MS spectra obtained on-line for isovitexin (33) in the methanolic extract of Swertia calycina. The LC/TSP-MS/MS spectrum of the ion m/z 313 (c) exhibits characteristic fragments for the substitution of the B-ring and for the glycosidation at C-6. LC/UV/MS: same conditions as for Fig. 6. LC/TSPMS, liquid chromatography/thermospray–mass spectroscopy; UV, ultraviolet.

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[M ⫹ H] ⫹ at m/z 449 (31), 463 (32), 433 (33), and 447 (34). Each spectrum also showed fragments [M ⫹ H ⫺ 90] ⫹ and [M ⫹ H ⫺ 120] ⫹ characteristic of the cleavage of C-glycoside moieties [38] (see TSP-MS spectrum of 33, Fig. 13A). Thus, 31–34 were all flavone C-glycosides, and according to their molecular weights, it was deduced that 31 bore four OH groups; 32, three OH groups and one OMe group; 33, three OH groups; and 34, two OH groups and one OMe group. The LC/TSP-MS/MS [62] analysis of the characteristic ions of these flavonoids provided more precise information about the fragmentation of the Cglycoside moieties and the localization of the substituents on the A- and B-rings. The LC/TSP-MS/MS was performed on the extract of S. calycina under the same LC and ionization conditions established for LC/TSP-MS. In the collision cell, argon was used for collisionally induced decomposition (CID). In a first analysis, the pseudomolecular ions m/z 449, 463, 433, and 447, corresponding to 31, 32, 33, and 34, respectively, were selected as parent ions and the daughter ions were recorded. The TSP-MS/MS spectra obtained on-line in the extract clearly showed for each flavonoid the characteristic [M ⫹ H ⫺ 120] ⫹ and [M ⫹ H ⫺ 150] ⫹ ions, due to the cleavage of the C-glycoside moieties [38], but the aglycone ion [A ⫹ H] ⫹ was not visible and further fragmentation of the aglycone moiety was not observed (see LC/TSP-MS/MS spectrum of 33, Fig. 13B). Thus, the LC/ TSP-MS/MS analysis of the flavonoid C-glycosides using the protonated pseudomolecular ions as parent ions did not provide information on the substitution sites on the A- and B-rings of the aglycones of these compounds. In contrast to Oglycosides that exhibit intense aglycone ions [A ⫹ H] ⫹ [62], the [A ⫹ H] ⫹ ions of C-glycosides are usually undetectable in the LC/TSP-MS spectra of flavonoids (see LC/TSP-MS spectrum of 33, Fig. 13A) and cannot be used for fragmentation studies. Thus, a second LC/TSP-MS/MS analysis of these constituents in the extract was performed by choosing the intense [M ⫹ H ⫺ 120] ⫹ ions as parent ions. In this case, a classic retro-Diels-Alder (RDA) fragmentation of the aglycone moiety of the flavonoids was observed [63]. For 31 and 32, a clearly discernible ion at m/z 137 was characteristic for a fragment of the B-ring bearing two hydroxyl groups, for 33 and 34, this fragment was recorded at m/z 121, indicating only one hydroxyl (see LC/TSP-MS/MS spectrum of 33, Fig. 13C). With this analysis it was also possible to ascertain the position of C glycosylation. As shown in Fig. 13, the LC/TSP-MS/MS spectrum of 33, obtained by choosing the [M ⫹ H ⫺ 120] ⫹ as parent ion, exhibited fragments at m/z 149 and 177 characteristic of 6-C-glycosylated flavones. Indeed, it is known that TSPMS/MS spectra of the [M ⫹ H ⫺ 120] ⫹ fragments of isomeric C-glycosylflavones show different specific daughter ions [64]. The A-C-ring fragments issued from RDA cleavages are only observable for C-6 positional isomers as in the case of 33. With these data, 33 was identified as isovitexin [65]. Similarly compounds 31, 32, and 34 were established as 6-C-glycosylflavones, and with additional computerized UV comparisons and retention time measurements were

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Figure 14 LC/UV and LC/TSP-MS analysis of the crude CH 2 Cl 2 extract of Swertia calycina. For each major peak, the single-ion LC/MS traces of the protonated molecular ions [M⫹H] ⫹ were displayed, together with the UV spectra obtained on-line. LC/ liquid chromatography/ultraviolet; TSP, thermospray; MS, mass spectroscopy.

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identified as isoorientin, swertiajaponin, and swertisin, respectively. All these flavonoids are well known in Swertia species and further isolation was not performed [61]. G.

Liquid Chromatography/Ultraviolet/Mass Spectroscopy and Liquid Chromatography–Nuclear Magnetic Resonance Analysis of the Dichloromethane Extract of Swertia calycina

The dichloromethane extract of Swertia calycina was strongly antifungal against Cladosporium cucumerinum and Candida albicans. Using the TLC bioautography assay [66], this activity was linked to a strong UV visible spot (R f ⫽ 0.38; petroleum ether/ethylacetate 1: 1). As some xanthones are known antifungal agents [67], one of these was suspected. The LC/UV chromatogram of the dichloromethane extract of S. calycina was simpler than the methanolic extract, and three main peaks (3, 35, and 16) were detected (Fig. 14). Compound 16 presented a UV spectrum with four ab-

Figure 15 Bidimensional LC/ 1H-NMR chromatogram of the crude CH 2Cl2 extract of Swertia calycina. Methoxyl groups and aromatic proton signals of 35 and 16 are clearly visible together with all the resonances of the monoterpene glycoside 3. The signal of HOD is negative and was continually shifted during the LC gradient. LC, liquid chromatography; NMR, nuclear magnetic resonance.

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sorption bands characteristic of a xanthone. The TSP-MS spectrum exhibited a strong protonated ion at 303 u, indicating a xanthone with a molecular weight of 302, and thus substituted with one hydroxyl and three methoxyl groups (Fig. 14). This information, together with the comparison with a UV spectral library, permitted the identification of 16 as decussatin, a xanthone widespread in the Gentianaceae family, also previously found in C. krebsii [56]. The on-line data obtained for 3 indicated the presence of a secoiridoid-type molecule with a molecular weight of 358 µ. The loss of 162 µ observable in the TSP spectrum was characteristic of the presence of a hexosyl moiety. These data suggested strongly that 3 was most probably sweroside, a well-known secoiridoid of the family [68]. The UV spectrum of 35 was not attributable to a common polyphenol of the Gentianaceae such as a flavone or xanthone. It was very weakly ionized in TSP (see the important background ions of the TSP spectrum in Fig. 16, for example, at m/z 231 and 200), but a protonated molecular ion was nevertheless found at m/z 189. This low molecular weight (188 u) and the UV spectrum suggested

Figure 16 Summary of all the spectroscopic data obtained on-line for naphthoquinone 35 in the CH 2 Cl 2 extract of Swertia calycina.

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that 35 could be a quinonic compound, but as no metabolite of this type was previously found in Gentianaceae, it was not possible to identify it on-line. In order to confirm these attributions and to obtain more structural information on-line, the same extract of S. calycina was submitted to an on-line LC/ 1 HNMR analysis on a 500-MHz instrument [13]. The same LC conditions as in the LC/UV/MS analysis were used except that the water of the LC gradient system was replaced by D 2O. However, the quantity of extract injected onto the column was increased to 1 mg to collect at least 20 µg for each peak of interest. For the suppression of the solvent signal of MeCN and its two 13 C satellites, as well as the residual HOD peak, WET solvent suppression [34] was run before each acquisition. In the gradient LC run, the solvent peaks change frequency during the course of the experiment. As a result, the solvent suppression must be continuously adjusted for optimal performance. To do this, a one-transientone-pulse experiment is used to find the solvent peaks prior to WET suppression (Scout Scan) [34]. As a result of this sequence, the transmitter is automatically adjusted to keep the biggest solvent peak at a constant frequency (MeCN), while the spectrometer is locked on D 2 O [13]. The on-line LC/NMR analysis of S. calycina provided 1 H-NMR spectra for all the major constituents. A plot of the retention time ( y axis) versus the NMR shifts (x axis) permitted the localization of the resonances of compounds 3, 35, and 16 (Fig. 15). On this unusual two-dimensional chromatogram, strong signals of aromatic methoxyl groups were observed around 4 ppm for 35 and 16. Xanthone 16 exhibited two pairs of aromatic protons, and the quinonic compound 35 presented five other low-field protons. The more polar secoiridoid 3 showed different signals between 3 and 6 ppm. The important trace starting from 4.8 ppm (at 0 min) and ending to 4 ppm (at 30 min) was due to the change of the chemical shift of the residual negative water (HOD) signal during the LC gradient. The traces between 1 and 2.6 ppm were due to residual MeCN signal and solvent impurities. The cross sections of this bidimensional plot along the x-axis produced single on-line LC/ 1 H-NMR spectra for each constituent, allowing a precise assignment of their specific resonances. Xanthone 16 exhibited three methoxyl resonances at 3.92, 3.93, and 3.95 ppm; a pair of meta-coupled aromatic protons (δ 6.55, d, J ⫽ 2.4 Hz, H-4; 6.41, d, J ⫽ 2.4 Hz, H-2) was indicative of a 1,3disubstituted A-ring. The B-ring protons exhibited a pair of ortho-coupled protons (δ 7.36, d, J ⫽ 9.2 Hz, H-5; 7.62, d, J ⫽ 9.2 Hz, H-6), suggesting a 1,3,5,6- or 1,3,7,8-substitution pattern for 16. These NMR data together with UV and MS information led to the conclusion that 16 was 1-hydroxy-3,7,8-trimethoxyxanthone (decussatin), as already suggested by the LC/UV/MS data. In the LC/ 1 H NMR spectrum of 35 (Fig. 16), two signals (δ 8.11, m, 2H: H-5,8; 7.89, m, 2H: H-6,7) were characteristic of four adjacent protons of an aromatic ring with two equivalent substituents. The low-field shift of the H-5,8

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signal indicated that these two protons were in peri positions of carbonyl functions, suggesting the presence of a naphthoquinone nucleus [69]. The strong bands recorded in the UV spectrum at 243, 248, 277, and 330 nm confirmed this deduction. The singlet at 6.35 ppm was attributed to H-3 and the remaining methoxyl group was thus at position C-2. With these on-line data and the molecular weight deduced from the LC/TSP-MS spectrum (MW 188), 35 was finally identified as 2-methoxy-1,4-naphthoquinone. As this was the first naphthoquinone to

Figure 17 Stop-flow WET-COSY spectrum of sweroside (3) in the CH 2 Cl 2 extract of Swertia calycina; 3 was kept 30 min in the cell for the total acquisition (8 scans ⫻ 220 increments). WET-COSY, water suppression enhanced through J, effect correlated spectroscopy.

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be reported in Gentianaceae, it was isolated and was found to be the compound responsible for the strong antifungal activity of the extract of S. calycina [61]. As indicated earlier, UV and MS data suggested that 3 was the secoiridoid sweroside. The on-line LC/ 1 H-NMR data indicated different resonances than those corresponding to this secoiridoid, but a rather poor signal-to-noise-ratio was obtained for this compound. In order to obtain clearer resonances, the LC/ 1 H-NMR spectrum of 3 was also recorded in the stop-flow mode. To perform this experiment, the LC flow was stopped when 3 was eluting into the LC/NMR flow cell. This was possible by following the separation of the extract by UV and synchronizing the LC pump to stop when the peak of interest entered the NMR detection cell (see setup in Fig. 5). In this mode, the analyte was maintained in the NMR for a greater number of transients and a better signal-to-noise ratio was obtained. All the resonances for 3 [70] were then easily identified. In order to prove the 1H-1 H connectivities for this secoiridoid, an 1 H, 1 H-correlated spectroscopy (COSY) spectrum was also acquired during the same stop-flow experiment (Fig. 17). This stop-flow WET-COSY experiment performed on the LC peak of 3 allowed assignment of all the 1 H-1 H correlations within the molecule. Only 35 min was needed to acquire and process this 2D spectrum. With these data, it was indeed possible to link all the protons of the glucosyl moiety of 3, starting from the anomeric proton (δ 4.84, d, J ⫽ 8.6 Hz, H-1′) and ending at the methylene H-6′a (δ 3.92) and H-6′b (δ 3.73) signals. Also, all the connectivities from the terminal methylene H-10 to H-7 progressing through the monoterpene skeleton were clearly assigned. These 1D and 2D LC/ 1H-NMR data, together with UV and MS information, allowed the identification of 3 as sweroside. This example showed that the LC/UV/MS and LC/NMR information obtained for S. calycina permitted a full structure identification of its main constituents.

IV. CONCLUSIONS The LC-hyphenated techniques are playing an increasingly important role as strategic tools to support phytochemical investigations. Indeed, these techniques provide a great deal of preliminary information about the content and nature of constituents of crude plant extracts. In certain cases, combination with a spectral library and precolumn or postcolumn derivatization allow structure determination on-line. This is very useful when large numbers of samples have to be processed because unnecessary isolation of compounds is avoided. Once the novelty or utility of a given constituent is established, it is then important to process the plant extracts in the usual manner, to obtain samples for full structure elucidation and biological or pharmacological testing. In this chapter, it has been shown that in the search for xanthones with potential monoamine oxidase–inhibitory activity, LC/MS and LC/UV analyses

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have been of great value for the preliminary screening of numerous species of Gentianaceae. These methods have been used not only to determine the presence of xanthones but also to provide an indication of their structures. The use of LC/UV and LC/MS has also aided the localization of new types of constituents such as secoiridoids of unusual molecular weight and helped to target their isolation. The introduction of complementary techniques such as postcolumn addition of UV shift reagents or MS/MS have extended the possibilities of on-line structural determination with LC/UV/MS. Analysis of crude plant extracts using LC/MS is, however, not straightforward because of the great variety of constituents. As has been shown, no interface allows an optimal ionization of all the metabolites within a single crude plant extract. Often, different ionization modes or different interfaces are necessary to obtain a complete picture of the extract composition. The recent introduction of LC/NMR for crude plant extract screening will probably allow another breakthrough in the on-line structural determination of natural products. This hyphenated method allows the recording of precious complementary on-line structure information when LC/UV/MS data are insufficient for an unambiguous peak identification. Indeed, LC/NMR has proved to be very effective in obtaining 1D spectra of both flowing and nonflowing samples, as well as stop-flow 2D spectra. However, compared with UV or MS, NMR remains a rather insensitive detection method. Thus, in the on-line mode, this technique is at present limited to the characterization of the major constituents of crude plant extracts. In order to obtain 1 H-NMR spectra of minor metabolites or to perform bidimensional experiments, the use of the stop-flow mode is necessary. With the full set of spectroscopic information obtained by LC/UV, LC/ MS, and LC/NMR, the phytochemist will be able to characterize the main constituents of a given plant rapidly and choose which metabolites are to be isolated for in-depth structural or pharmacological study. The chemical screening of extracts with such elaborate hyphenated techniques generates a huge amount of information. In order to rationalize this approach and to use it efficiently with high sample throughput, the next challenge will be to find a way to centralize all these data for rapid pattern recognition by reference to standard compounds. With such an analytical system, phytochemists will then be able to concentrate their efforts on finding new biological targets. This aspect remains the more difficult problem to solve when searching for new leads. Both biological and chemical screenings provide important information on the plant constituents, but these results will not be sufficient for the discovery of new potent drugs if no suitable pharmacological model exists. Thus, an important condition for success in the discovery of new bioactive plant constituents is the establishment of effective collaborations among botanists, phytochemists, and pharmacologists in order to realize all the steps involved, starting with plant material in the field and concluding with viable pharmacologically active compounds.

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Only in this manner can medicinal plants be investigated efficiently and new leads rapidly discovered.

ACKNOWLEDGMENT Financial support was provided by the Swiss National Science Foundation. Thanks are due to Dr. Wolf Hiller, Varian AG, Darmstadt, Germany, for the LC/NMR measurements and to Dr. Winfried Wagner-Redeker, Spectronex AG, Basel, Switzerland, for the LC/ES-MS experiments.

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47. O. Suzuki, Y. Katsumata, M. Oya, V. M. Chari, R. Klapfenberger, H. Wagner, and K. Hostettmann, Planta Med., 39: 19 (1980). 48. D. Schaufelberger and K. Hostettmann, Planta Med., 54: 219 (1988). 49. U. Thull, Monoamine oxidase inhibitors of natural and synthethic origin: Biological assay and 3D-OSAR, Thesis, Universite´ de Lausanne (1995). 50. H. Wagner and K. Muenzig-Vasirian, Dtsch. Apoth. Z., 115: 1233 (1975). 51. K. Hostettmann and M. Hostettmann, in Methods in Plant Biochemistry, (J. B. Harborne ed.), Academic Press, London, pp. 493 (1989). 52. G. G. Nikolaeva, V. I. Glyzin, M. S. Mladentseva, V. I. Sheichenko, and A. V. Patudin, Khim. Prir. Soedin., 1: 107 (1983). 53. A. A. Lins Mesquita, D. De Barros Correa, O. R. Gottlieb, and M. Taveira Magalhaes, Anal. Chim. Acta, 42: 311 (1968). 54. K. Hostettmann and H. Wagner, Phytochemistry, 16: 821 (1977). 55. M. Kaldas, Identification des compose´s polyphe´noliques dans Gentiana campestris L., Gentiana germanica Willd. et Gentiana ramosa Hegetschw, Thesis, Universite´ de Neuchaˆtel, Switzerland (1977). 56. J.-L. Wolfender, M. Hamburger, J. D. Msonthi and K. Hostettmann, Phytochemistry, 30: 3625 (1991). 57. S. Rodriguez, J.-L. Wolfender, G. Odontuya, O. Purev, and K. Hostettmann, Phytochemistry, 40: 1265 (1995). 58. W.-G. Ma, N. Fuzzati, J.-L. Wolfender, K. Hostettmann, and C. Yang, Helv. Chim. Acta, 77: 1660 (1994). 59. W. Ma, N. Fuzzati, J.-L. Wolfender, C.-R. Yang and K. Hostettmann, Phytochemistry, 43: 805 (1996). 60. S. Rodriguez, J.-L. Wolfender, G. Odontuya, O. Purev and K. Hostettmann, Helv. Chim. Acta, 79: 363 (1996). 61. S. Rodriguez, J.-L. Wolfender, E. Hakizamungu, and K. Hostettmann, Planta Med., 61: 362 (1995). 62. Y. Y. Lin, K. J. Ng, and S. Yang, J. Chromatogr., 629: 389 (1993). 63. T. J. Mabry and K. R. Markham, in The Flavonoids (J. B. Harborne, T. J. Mabry, and H. Mabry, eds.), Academic Press, New York, pp. 78 (1975). 64. G. Rath, A. Toure´, J.-L. Wolfender, and K. Hostettmann, Chromatographia, 41: 332 (1995). 65. K. Hostettmann, G. Bellmann, R. Tabacchi, and A. Jacot-Guillarmod, Helv. Chim. Acta, 56: 3050 (1973). 66. L. Rahalison, M. Hamburger, M. Monod, E. Frenk, and K. Hostettmann, Planta Med., 60: 41 (1994). 67. A. Marston, M. Hamburger, I. Sordat-Diserens, J. D. Msonthi, and K. Hostettmann, Phytochemistry, 33: 809 (1993). 68. H. Inouye, S. Ueda, and Y. Nakamura, Chem. Pharm. Bull., 18: 1856 (1970). 69. R. H. Thomson, Naturally Occurring Quinones, University Press, Cambridge (1987). 70. T. A. van Beek, P. P. Lankhorst, R. Verpoorte, and A. Baerheim Svendsen, Planta Med., 44: 30 (1982).

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5 Determination of the Absolute Configuration of Biologically Active Compounds by the Modified Mosher’s Method Takenori Kusumi Tokushima University, Tokushima, Japan

Ikuko I. Ohtani University of the Ryukyus, Okinawa, Japan

I. INTRODUCTION Most biologically active natural products have stereogenic centers and are obtained from their natural sources enantiomerically pure. In pharmaceutical drug development of naturally occurring and synthetic compounds, identification of the absolute configuration of the drug is crucial. In many cases, when a drug is chiral, only one enantiomer is effective, and sometimes the other enantiomer may be seriously hazardous [1]. Because of the continuous development of new spectroscopic techniques, unambiguous structure determination of natural and synthetic products has become relatively routine even in cases in which the molecular weight of a compound is greater than 1000 daltons. Few methods exist, however, to elucidate the absolute configuration of organic compounds; the most commonly employed nonchemical procedures are xray crystallography and the exciton chirality method [2]. The latter circular dichroic methodology has been most prominently developed by Nakanishi [3] and Harada [4] (see Chapter 5) over the past decade. We have been developing methodology to elucidate the absolute configuration of organic compounds using nuclear magnetic resonance (NMR) spectros-

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copy, generally considered the most important instrumental method in organic chemistry laboratories. Our methodology, which we have called the ‘‘modified Mosher’s method’’ [5], evolved from Mosher’s method [6] and uses Mosher’s reagent, 1-methoxy-1-phenyl-1-trifluoromethylacetic acid (MTPA), as a chiral derivatizing agent. A number of chemists have now applied this method to both natural and synthetic products, establishing it as highly reliable and effective for determining absolute configuration. This chapter introduces the concept and application of the modified Mosher’s method and lists numerous compounds whose absolute configurations were determined by this method. Since our initial reports on the modified Mosher’s method, several new, structurally similar chiral derivatizing reagents for NMR analysis have also been reported as tools to elucidate absolute configuration. All of these reagents consist of a chiral auxiliary with an aromatic group such as phenyl, 1- or 2-naphthyl [7,8,] binaphthyl [9], 2-anthryl [10], or 9-anthryl systems, which cause shifts of the protons by the anisotropic effect of the aromatic rings. Thus, for these compounds, we have proposed the term chiral anisotropic reagents [11]. Most of the chiral anisotropic reagents are applicable to organic compounds having a secondary alcohol moiety. Although most of these new chiral anisotropic reagents will be summarized elsewhere, N,N-dimethylphenylglycinamide (phenylglycine dimethylamide [PGDA]) and methyl phenylglycinate (phenylglycine methyl ester [PGME]), developed for the absolute configuration of carboxylic acids, are described in this chapter. II. MODIFIED MOSHER’S METHOD A.

Basic Principles of Mosher’s Method

By careful examination of the chemical shift differences observed between the diastereomeric pairs of (R)- and (S)-MTPA esters of chiral secondary alcohols, Mosher proposed [6] that in solution, the carbinyl proton, ester carbonyl, and trifluoromethyl groups of the MTPA moiety are oriented on the same plane (Fig. 1). PCILO calculations [12] and x-ray analysis [13] of the MTPA esters supported

Figure 1 Conformational model proposed by Mosher for MTPA esters. MTPA, 1-methoxy-1-phenyl-1-trifluoromethylacetic acid. (Source : Ref. 6.)

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this concept in a qualitative sense. As Mosher pointed out, when the MTPA ester exists in this conformation, resonances of ligand L 3 in the (S)-MTPA ester are upfield in the 1 H NMR spectrum relative to those of the (R)-MTPA ester as a result of the diamagnetic shielding effect of the benzene ring (Fig. 1). The reverse sense of nonequivalence is true for the 1 H resonances of ligand L 2, with those of the (R)-MTPA ester being upfield relative to those of the (S)-MTPA ester. Therefore, comparison of the chemical shifts of the proton resonances of L 2 and L 3 of the diastereomeric esters allows assignment of the absolute configuration of the secondary alcohol. When Mosher first put forward his analysis in the early 1970s however, the NMR instruments available were mostly 60–100 MHz in proton frequency, and the resolution and assignment of protons of complex organic molecules were practically impossible with such limited field strengths (14.1–23.6 kG). Consequently, the use of 19 F NMR [6,14] or a lanthanide-shift reagent to resolve the easily identified OMe singlets of the MTPA-esters [15] was often preferred to the original 1 H NMR method, which required identification of the usually overlapped protons bonded to the original alcohol (on L 2 and L 3). Since Mosher’s conformational analysis did not address the relative shielding of these resonances, unambiguous assignment of absolute stereochemistry was difficult (if not impossible!), and early applications of the MTPA ester method were primarily limited to determination of diastereomeric excess (enantiomeric excess of original alcohol) of enriched synthetic mixtures. The intrinsic drawback in these early applications of Mosher’s method for absolute stereochemistry assignment was that it usually depended on only two data points—the chemical shift difference of the two CF 3’s (19 F) or OMe’s (1 H) of the (R)- and (S)-MTPA esters—and not on the more difficulty assigned resonances of the substrate alcohol. The modified Mosher’s method, on the contrary, uses the chemical shifts of many protons as Mosher originally intended and thus is more reliable. B.

Basic Concept of the Modified Mosher’s Method

The basic concept of the modified Mosher’s method is essentially the same as Mosher proposed: the idealized conformation depicted in Fig. 2a accounts for the observed chemical shift differences in the MTPA esters of secondary alcohols. (For convenience, this conformation will be called the ideal conformation, and the plane defined by the carbonyl and the trifluoromethyl groups, and the carbinol proton H α in this conformation will be called the MTPA plane.) Because of the diamagnetic effect of the benzene ring, the NMR signals of H X ,Y,Z . . . of the (S)MTPA ester should appear upfield relative to those of the (R)-MTPA ester. The reverse should hold true for H A ,B ,C. . . Therefore, when ∆δ ⫽ δ S ⫺ δ R , protons on the right side of the MTPA plane must have positive values (∆δ ⬎ 0) and

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Figure 2 (a) MTPA plane of an MTPA ester; H A,B,C and H X,Y,Z are on the right and left sides of the plane, respectively, when viewed down the plane from the MTPA subunit. (b) Model A to determine the absolute configuration of secondary alcohols; Model A is the view of the MTPA ester drawn in (a) from the direction specified by the outlined arrow. MTPA, 1-methoxy-1-phenyl-1-trifluoromethylacetic acid. (Source: Ref. 5.)

protons on the left side of the plane should have negative values (∆δ ⬍ 0, Fig. 2B). Mosher’s original method can be extended as follows: (1) assign as many proton signals as possible of the (R)- and (S)-MTPA esters; (2) obtain ∆δ values for the protons; (3) place the protons with positive ∆δ on the right side, and those with negative ∆δ on the left side of Model A (Fig. 2b); (4) construct a molecular model of the compound in question and confirm that all the assigned protons with positive and negative ∆δ values are actually found on the right and left sides of the MTPA plane, respectively. The absolute values of ∆δ should be proportional to the distance from the MTPA moiety. When these conditions are all satisfied, Model A will indicate the correct absolute configuration of the compound. This modified Mosher’s method allows examination of chemical shift differences of as many protons as may be assigned by means of up-to-date NMR techniques including two-dimensional (2D) spectra such as H,H-Correlation Spectroscopy (H,H-COSY) and Homonuclear Hartmann-Hahn spectroscopy (HOHAHA) spectra. We feel that as this method relies on a larger number of

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data points (i.e., all the protons assignable by 2D NMR techniques), it is far more reliable than the previously mentioned variations of Mosher’s method using 19 F NMR or a lanthanide shift reagent.

1. Preparation of 1-Methoxy-1-Phenyl-1-Trifluoromethylacetic Acid Esters The MTPA esters of a secondary alcohol are easily prepared [5] by treatment of the alcohol (1) with (R)- or (S)-MTPA acid and 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide (EDC) and 4-dimethylaminopyridine (DMAP) in dichloromethane, or (2) with (R)- or (S)-MTPA chloride and DMAP/Et 3N in dichloromethane or pyridine. Usually the reaction is completed within 15 h at room temperature. In a few cases, the reaction rate of (R)- and (S)-MTPA acids or acid chlorides with the chiral alcohol differs significantly (kinetic resolution), and either diastereomer may be slow, or even difficult to obtain. Note the following: because of the nomenclature priority rules, the chloride of (R)-MTPA acid is (S)-MTPA chloride and the chloride of (S)-MTPA acid is (R)-MTPA chloride. The stereochemical correlation of MTPA acids and their chlorides is correctly shown in the scheme below.

When the amount of sample is limited, less than 1 mg of the sample per esterification is usually sufficient for assignment of the absolute configuration thanks to the high sensitivity of superconductive NMR instruments. One advantage of MTPA esters is that the MTPA moiety possesses ultraviolet (UV) absorbing phenyl chromophore. Combinatory use of preparative thin layer chromatography (TLC) (fluorescent under UV 254 nm) and HPLC successfully achieves purification of the MTPA esters.

2. Choice of Nuclear Magnetic Resonance Solvents for Recording the Spectra It is advisable to use nonaromatic solvents such as deuterochloroform or deuteromethanol for the modified Mosher’s method. Use of deuterobenzene (and probably other aromatic solvents such as deuteropyridine) often gives confusing results [5]. Interaction between the aromatic solvent and the phenyl group of MTPA may be the cause of this difficulty.

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3. Application to Natural Products or Their Simple Derivatives Having Secondary Hydroxyl Groups To confirm the validity of the present method, several compounds, including cholesterol, ergosterol, (⫺)-borneol, and (⫺)-menthol (the absolute configuration of which are known), were subjected to the method [5]. The assignments of the proton chemical shifts (all in CDCl 3) were accomplished using 1D and 2D (COSY, HOHAHA, NOESY [Nuclear Overhauser and Exchange Spectroscopy]) NMR techniques. As illustrated with (⫺)-menthol-(1), the ∆δ values (in parts per million [ppm]) were all systematically arranged so that the positive and negative nonequivalences were oriented on the right and left sides of the MTPA plane, respectively (Fig. 3). The absolute configuration predicted by this method was the same as known for the natural product. The results for the other three natural products were all consistent with the known absolute configurations. The modified Mosher’s method was subsequently applied [5] to several marine natural products, including sanadaol (2) [16] hydroxyacetyldictyolal (fukurinolal) (3) [17], cembranolide (4) [18], denticulatolide (5) [19], and crenulacetal B (6) [20], with unknown absolute configurations. These results are shown in Fig. 4. Without exception, systematic arrangement of positive and negative ∆δ values was observed for these compounds, and their absolute configurations were deduced as shown in the structures. For compounds 2 and 5, the absolute configurations determined by the present method have been confirmed by total synthesis [21] and X-ray crystallography [22], respectively. The modified Mosher’s method has now become an indispensable means to elucidate the absolute configuration of natural products, and additional examples using this method are continually being reported. In the following section are examples of compounds whose absolute configuration was determined by the modified Mosher’s method. As these examples demonstrate, the absolute config-

Figure 3 ∆δ Values (ppm) recorded for the MTPA esters of (⫺)-menthol (1). MTPA, 1-methoxy-1-phenyl-1-trifluoromethylacetic acid. (Source : Ref. 5.)

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Figure 4 ∆δ Values (ppm) recorded for the MTPA esters of marine terpenoids 2–6. MTPA, 1-methoxy-1-phenyl-1-trifluoromethylacetic acid. (Source: Ref. 5.)

uration of compounds that are not secondary alcohols yet possess functionality routinely converted into one with high stereoselectivity are also amenable to this method. For example, reduction of the aragupetrosine A (59) ketone group to the secondary alcohol with cyanoborohydride allowed absolute stereochemistry assignment by the modified Mosher’s method [23]. 4. Application to Natural Products Without a Secondary Hydroxyl Group When the compound in question has no secondary hydroxyl group, chemical introduction of the hydroxyl group provides the handle to use the modified Mosh-

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er’s method for the absolute configuration assignment. As illustrated in the previous section (Fig. 5), simple deacylation (e.g., 8, 31, 34, 44, 52, and 55) or reduction of a ketone (e.g., 59) may expose a latent secondary alcohol for subsequent formation of the MTPA esters. In addition, reduction of peroxides (e.g., 14, 15 and 47), epoxide or larger cyclic ether opening (e.g., 24, 41, and 63a,b), and an ozonation/reduction sequence (e.g., 10) have also been utilized to obtain the requisite secondary alcohol. Two less routine examples of such functionalizations are presented in this section.

Figure 5 Examples of natural and synthetic secondary alcohols whose absolute configurations were determined by the modified Mosher’s method.

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Figure 5 Continued

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Figure 5 Continued

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Figure 5 Continued

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Lobatriene (64) is a marine diterpene originally isolated from a soft coral [71]. The relative configuration of the substituents on ring A had been determined, but the stereochemical correlation of the substituents between rings A and B by NMR spectroscopy was not possible because of rapid rotation along the single bond connecting the rings. The absolute configurations of all the asymmetric centers were, therefore, determined by chemical transformations with subsequent application of the modified Mosher’s method, as shown in Fig. 6 [72]. Reductive cleavage of the allylic ether bond afforded 65, which possesses a secondary alcohol at C-17. The R-configuration at this carbon was deduced by the results detailed on the structure of the MTPA ester 65a. Ozonolysis of 65 gave methyl ketone 66 (1 mg), which was subjected to a Baeyer-Villiger reaction that proceeds with retention. Basic hydrolysis of the resulting acetate yielded the secondary alcohol 67 (ca. 500 µg). This was divided into two portions and treated with (R)and (S)-MTPA chloride, respectively. The ∆δ values obtained are detailed on

Figure 6 Chemical transformation of lobatriene (64) into secondary alcohols 65 and 67, and the ∆δ values (ppm) recorded for their MTPA esters 65a and 67a, respectively. MTPA, 1-methoxy-1-phenyl-1-trifluoromethylacetic acid. (Source: Ref. 72.)

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structure 67a. Beautifully systematic arrangement of positive and negative values was observed, and the S-configuration of the hydroxy group was easily established. Thus, the configuration of four asymmetric centers of 64 was determined as shown in Fig. 6. It should be noted that the sign of the ∆δ values of the methyl, ethyl, and isopropyl groups on the A-ring is negative despite its apparent location on the right side of the MTPA plane in 65a. One plausible explanation of this apparent inversion of the sign is that the long-side chain may exist in a twisted conformation, so that the alkyl substituents are actually oriented on the left side of the MTPA plane. Thus, application of the modified Mosher’s method to acyclic compounds must be interpreted with caution when nonequivalence of protons quite remote from the acylation site is observed. Another nonroutine example is isoclavukerin A (68), isolated from an Okinawan soft coral as a very volatile liquid [73]. The relative stereochemistry of 68 was deduced by comparison of the NMR data with those of clavukerin A, the C-2 epimer of 68 [74]. Fortunately isoclavukerin A autooxidized during storage in a refrigerator to give 69, which had a secondary hydroxyl group. Application of the modified Mosher’s method led to the absolute configuration of the secondary alcohol 69a, and thus of isoclavukerin A (Fig. 7). It is noteworthy that the benzoate 69b showed no split circular dichroism (CD), although it is an allyl benzoate [2,75]. Molecular models indicated that the dihedral angle between the benzoyloxy and the olefinic groups was 180°, which does not in principle give a Davydov-split Cotton effect [2,76]. Thus in such cases in which the exciton

Figure 7 Auto oxidation product 69 of isoclavukerin A (68) and ∆δ values (ppm) recorded for the MTPA esters of 69. MTPA, 1-methoxy-1-phenyl-1-trifluoromethylacetic acid. (Source : Ref. 73.)

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chirality method is not applicable, the modified Mosher’s method may be successfully applied. 5. Exceptions to the Rule of the Modified Mosher’s Method and Countermeasures a. Axial Alcohols. In our attempt to elucidate the absolute configuration of the marine triterpene sipholenol-A (70) [77] using the modified Mosher’s method, we noticed that ∆δ values of the protons on the A- and B-rings of the MTPA derivatives were irregularly arranged, as seen in Fig. 8 [78]. These data, of course, could not be used to determine the absolute configuration. However, this case illustrates another advantage of the present method: a self-examination mechanism based on systematic sense of nonequivalences to establish whether the observed data can be used or must be abandoned. Molecular models of 70 suggested that the OMTPA group exists in an axiallike orientation and was sterically compressed by the two axial protons on C-2 and C-7 (70′), which may compel the MTPA group to take a different conformation from the ideal one. Therefore, 70 was converted into the C4-epimer 71, in which the OH group exists in a sterically less crowded equatorial position, by oxidation of 70 (PDC [pyridinium dichromate] in CH 2 Cl 2) followed by reduction

Figure 8 The ∆δ values (ppm) recorded for the MTPA esters of sipholenol-A (70). MTPA, 1-methoxy-1-phenyl-1-trifluoromethylacetic acid. (Source: Ref. 78.)

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with sodium borohydride. (The Mitsunobu reaction did not work.) To our delight, the positive and negative ∆δ values of 71 were found to be systematically arranged, and that enabled us to determine the absolute configurations of 71, and thus of 70, as shown in the respective structures. The absolute configuration was subsequently verified by x-ray crystallography on the (S)-MTPA ester of 70 [79]. In order to confirm that steric interference may prevent adoption of the ideal conformation of the MTPA group in such cases with axial alcohols, and thereby cause irregular arrangement of ∆δ values, friedelan-3β-ol [72, R ⫽ H] was examined [5]. Because of the rigidity of the rings, the C3 hydroxyl group of 72 is axially oriented. The ∆δ values of the MTPA derivative of 72 (R ⫽ MTPA) are, as anticipated, irregularly distributed, in contrast to the results obtained for the C3 epimer 73 (Fig. 9), wherein the ∆δ values are completely in accord with the rule. These results suggest a general device to overcome the problem of the present method: if the ∆δ values of the MTPA derivatives of a secondary alcohol are irregularly arranged and cannot be used, epimerization of the hydroxyl group and subsequent application of the MTPA method may succeed. b. Anomalies Caused by the Anisotropy of Other π-Systems. As described, the modified Mosher’s method utilizes the differences in the corresponding proton chemical shifts between the diastereomers of (R)- and (S)-MTPA esters. If the conformation of the MTPA group of one diastereomer greatly differs from that of the other, as seen in the axial alcohol examples cited, the ∆δ values will not be systematically arranged about the MTPA plane. The question also arises, What

Figure 9 The ∆δ values (ppm) recorded for the MTPA esters of friedelan-3β-ol (72) with the axially oriented C3 alcohol, and those of the MTPA esters of its C3 epimer 73. MTPA, 1-methoxy-1-phenyl-1-trifluoromethylacetic acid. (Source : Ref. 5.)

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will be the consequences if the conformations of other functional groups differ in the diastereomers? During synthetic studies on tautomycin, the modified Mosher’s method was applied to intermediates 74a and 74b [80]. Ambiguous ∆δ values were observed for one of the α-protons of these two derivatives (∆δ ⫽ ⫺0.04 and 0, Fig. 10); however, other ∆δ values were systematically arranged [81]. To determine whether this irregularity of ∆δ values was common in compounds bearing a βaromatic ring, the modified Mosher’s method was applied to synthetic β-aromatic secondary alcohols, 75a–80a and 75b–80b. All the protons on the left and right sides of the MTPA plane possessed negative and positive ∆δ values, respectively, except these α-protons (H A and

Figure 10 Synthetic compounds 74a′ and 74b′ with irregular ∆δ values for one of the α-protons (shown in boldface) and other model compounds 75–80. (Source : Ref. 81.)

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Figure 11 Conformations of 74a′–80a′ and 74b′ and 80b′ deduced from NMR studies. NMR, nuclear magnetic resonance. (Source: Ref. 81.)

H B). Although these methylene protons should have positive ∆δ values, in many compounds they were negative or zero. The conformations of these esters deduced from their NMR properties (Fig. 11) were supported by molecular modeling studies of 74b′ (Fig. 12). Since the α-protons are located near the β-aromatic ring, these were anisotropically affected by both this β-aromatic ring as well as the benzene ring of the MTPA moiety. Even though a difference of the dihedral angles (H c-C-C-C1) in the predicted most stable conformations for 74b′ was not observed between the (S)- and (R)-MTPA esters (352° and 351°, respectively) by molecular mechanics calculation, subtle differences in the conformation around the β-carbon between (S)- and (R)-MTPA esters might cause these irregularities of the ∆δ values. A similar anomaly was observed in tanabalin [81], a natural product isolated from the dried flowers of the Brazilian medicinal plant Tanacetum balsamita, which has a secondary alcohol adjacent to a furan ring [82]. One of the C-11 methylene protons showed an anomalous positive ∆δ value thought to be caused by the furan ring anisotropy. The anomaly was ‘‘verified’’ by using model

Figure 12 Stable conformations predicted for 74b′ by molecular modeling. (Source: Ref. 81.)

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Figure 13 The ∆δ values (ppm) obtained for the MTPA esters of tanabalin 81a and model compound 82a. MTPA, 1-methoxy-1-phenyl-1-trifluoromethylacetic acid. (Source: Ref. 82.)

compound 82, which also showed the irregular ∆δ value of the α-proton denoted by a thick arrow (Fig. 13). Minale et al. [40] encountered similar anomalies in the structure determination of superstolide A that can be interpreted in the same manner: an irregular ∆δ value (denoted by the arrow) in both 28a and 28b (Fig. 14). This anomaly was apparently caused by the N-acetyl group anisotropy, the conformation of which may be slightly different between the (R)- and (S)-MTPA diastereomers, affecting the chemical shifts of the protons close to the carbonyl group. Since other ∆δ values were systematically arranged, their absolute configurations were safely assigned. In conclusion, the modified Mosher’s method can be applied to secondary alcohols possessing other π-systems, such as the aromatic rings and carbonyl groups that can fully or partially rotate along a single bond, in the vicinity of the hydroxy group. In such cases, however, one must be aware of the possible impact of this second anisotropy in generating irregular ∆δ values. 6. Applications in the Enantioselective Synthesis of Natural Products Biologically active natural products are primary targets of synthetic chemists. Most natural products exist as single enantiomers, and thus enantioselective synthesis is particularly important to organic chemists. When a chemical reaction yields only one enantiomer, the absolute configuration of the product can often

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Figure 14 The ∆δ values (ppm) reported by Minale for the MTPA esters 28a and 28b. MTPA, 1-methoxy-1-phenyl-1-trifluoromethylacetic acid. (Source : Ref. 40.)

be determined by using the modified Mosher’s method. Products obtained by ‘‘enantioselective’’ reactions, however, are commonly accompanied by minor amounts of the second enantiomer. In such cases, the absolute configuration of the major and minor products can be assigned by preparing only (S)- [or (R)-] MTPA esters of both enantiomers [83]. Fig. 15 shows why this is possible. [Bear in mind that the NMR spectrum of the (S)-MTPA ester of one product is identical with that of the (R)-MTPA ester of its enantiomer.] Esterification of the hypothetical product mixture consisting of the major enantiomer (R′) and its minor antipode (S′) with (S)-MTPA acid will give the S ⫺ R′ (major diastereomer) and S ⫺ S′ (minor diastereomer) esters, which are separated by chromatography. Because the 1 H NMR spectrum of S ⫺ S′ is identical with that of its enantiomer R ⫺ R′, subtraction of the chemical shifts of S ⫺ R′ from those of S ⫺ S′ (⫽ R ⫺ R′) will give ∆δ values for assigning the absolute configuration (R′) of the major product: [∆δ ⫽ δ (S)-MTPA ester ⫺ δ (R)-MTPA ester ⫽ δ S⫺R′ ⫺ δ R⫺R′ (⫽ δ S⫺S′)].

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Figure 15 Strategy for the determination of the absolute configuration of synthetic products obtained from enantioselective synthesis using the modified Mosher’s method.

In the total synthesis of (⫺)-dendrobine, Mori successfully used this idea [28]. (⫹)-Carveol [83] was converted to benzylamine 84, whose enantiomeric purity (ee) was found to be 90%, although the absolute configuration of the major enantiomer was unknown since either an S N2 or S N2′ mechanism could dominate in the Mitsunobu reaction (Fig. 16). The enantiomeric mixture of 84 and 84′ was transformed into allyl amine 85, and then into a tricyclic ketone, sodium borohydride reduction of which yielded 86 (plus its antipode). Conversion into the (S)-MTPA esters gave the two diastereomers, and ∆δ values (shown in 11) were calculated by subtraction of the chemical shifts (1H) of the major one from those of the minor. Because the (S)-MTPA esters were used, the absolute configuration of the major enantiomer of 86 was determined as shown in the structure. A similar application of the modified Mosher’s method has recently been reported by Fujimoto [38]. 7. Applications to Primary Amines Bonded to Methine Carbons The modified Mosher’s method is also applicable to the amines attached to tertiary carbons (methines). The MTPA amides may exist in the same conformation

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Figure 16 Synthetic pathway to the (⫺)-dendrobine intermediate 86, and ∆δ values (ppm) recorded for a mixture of the (S)-MTPA esters of 86 (ester 11) and its enantiomer. (S)-MTPA, (S)-1-methoxy-1-phenyl-1-trifluoroacetic acid. (Source: Ref. 28.)

as the MTPA esters; therefore, ∆δ ⫽ δ S ⫺ δ R can lead to the absolute configuration of the amines. Some examples using amino acid esters and amino alcohol derivatives are shown in Fig. 17 [84]. Other examples of the assignment of the absolute stereochemistry of aminecontaining natural and synthetic compounds [98–100] using the modified Mosher’s method are listed in Fig. 18 [85–87]. The modified Mosher’s method was also successful in assigning the absolute configuration of the primary aminecontaining subunit of fumonisin B 1 (32), as illustrated in Fig. 5 [44]. As these

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Figure 17 The ∆δ values (ppm) recorded for amino compounds 87–97. Proton assignments that could be interchanged lead to two possible values, as indicated by the second value in parentheses. Values for the methoxyl group of the MTPA subunit (all positive) are also given. MTPA, 1-methoxy-1-phenyl-1-trifluoroacetic acid. (Source: Ref. 84.)

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Figure 18 Examples of primary amines whose absolute configurations were determined by the modified Mosher’s method.

examples demonstrate, a large number of research groups have now found this procedure to be tremendously useful for the assignment of absolute stereochemistry for primary amines as well as secondary alcohols.

III. PHENYLGLYCINE DIMETHYLAMIDE AND PHENYLGLYCINE METHYL ESTER, CHIRAL ANISOTROPIC REAGENTS FOR THE ABSOLUTE CONFIGURATION DETERMINATION OF CARBOXYLIC ACIDS A.

Molecular Design for the Chiral Anisotropic Reagents

Applications of the modified Mosher’s method and other NMR reagents (chiral anisotropic reagents) are primarily restricted in their usage to secondary alcohols, and more recently to primary amines. By extending Mosher’s concept that comparison of the chemical shifts of diastereomeric derivatives of a chiral compound obtained from (R)- and (S)-derivatizing agents can lead to absolute configuration

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assignment, new chiral anisotropic reagents may be developed for the compounds possessing functional groups other than a secondary alcohol. In this context, we have elaborated new chiral anisotropic reagents, which allow determination of the absolute configuration of carboxylic acids, in which the carboxylic group is attached to a methine carbon [88]. The general molecular design for such chiral anisotropic reagents is shown in Fig. 19. A chiral carboxylic acid of type I is converted to acyl derivative II, where X can be any heteroatom. In order for the phenyl ring to cast its diamagnetic field effectively, and selectively, onto R 1 or R 2, coplanarity of C1 to Y5 in the dominant conformation is necessary. When X is NH, coplanarity from C1 to C4 may be achieved because of the planar structure as well as the preferred strans conformation of the amide group. If Y is also a polar group that would interact repulsively with the polar carbonyl group at C2 in a syn relationship, the C2 through Y5 orientation about the X3-C4 bond may also adopt a trans conformation. We chose the dimethylaminocarbonyl and methoxycarbonyl groups for Y, the former being preferable because of the enhanced dipole caused by the greater participation of the lone-pair electrons of the nitrogen atom with carbonyl resonance. Furthermore, hydrogen bonding between N 3H and C 5 ⫽ O,

Figure 19 Rationale to elucidate the absolute configuration of carboxylic acids by using PGDA and PGME. PGDA, phenylglycine dimethylamide; PGME, phenylglycine methyl ester. (Source : Ref. 88.)

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which will reinforce conformation IV, can be expected. It is equally important that the proton signals of the Z group (NMe 2 and OMe in these examples) do not obscure other 1 H NMR signals. The chiral anisotropic reagents thus designed are phenylglycine dimethylamide (PGDA) and phenylglycine methyl ester (PGME). If PGDA and PGME derivatives of a chiral carboxylic acid predominantly populate conformation IV, H X,Y,Z would be more shielded by the phenyl group of the (S)-PGDA (or PGME) moiety than those of the (R)-derivative. The reverse will be true for H A,B,C, which would be more shielded in the (R)-derivative. Like the modified Mosher’s method for secondary alcohols and amines, Model B, in which ∆δ ⫽ δ S ⫺ δ R , will show the correct absolute configuration of the carboxylic acid.

B.

Preparation of Phenylglycine Dimethylamide and Phenylglycine Methyl Ester

Although both enantiomers of PGME are commercially available, (R)-(⫺)-phenylglycine was used as starting material for both PGME and PGDA. Enantiomerically pure PGME {(R)-PGME hydrochloride; [α] D ⫺135°, (S)-PGME hydrochloride [α] D ⫹ 135°} was obtained by treating phenylglycine with thionyl chloride in methanol. The preparation of (R)-(⫺)-PGDA was also routine: NBoc-phenylglycine was condensed (PyBOP [benzotriazol-1-yloxytris(dimethylamino)phosphonium hexafluorophosphate], HOBT [1-hydroxybenzotriazole]) with dimethylamine to give the dimethylamide. The protecting group was removed under acidic condition (4 M HCl) to give PGDA hydrochloride {(R)PGDA hydrochloride; [α] D ⫺ 111°}. Partial epimerization of the phenylglycinate was observed in several attempts to prepare PGDA hydrochloride, so caution must be exercised to use only enantiomerically pure material for assignment of absolute stereochemistry, and especially for the determination of e.e. (enantiomeric excess).

C.

Applications of Phenylglycine Dimethylamide and Phenylglycine Methyl Ester to the Determination of Carboxylic Acid Absolute Stereochemistry

Prior to applying PGDA and PGME to carboxylic acids of unknown absolute stereochemistry, it was important to clarify whether derivatized amides actually had the expected conformation IV (Fig. 19). To this end, PGDA was condensed (PyBOP, HOBT) with 4-methylcyclohexanecarboxylic acid (101) to give crystalline 101a. An x-ray structure of 101a revealed that the diamide moiety actually exists in the expected conformation as shown in IV: C1 to Y-5 (NMe 2 in 101a)

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Figure 20 The ∆δ values (ppm) of PGDA derivatives of 4-methylcyclohexanecarboxylic acid 101a and (S)-2-methylbutanoic acid 102a. PGDA, phenylglycine dimethylamide. (Source: Ref. 88.)

were nearly coplanar, and, because the phenyl plane faces the methylcyclohexane residue, the anisotropy of the phenyl group was effectively cast over the methylcyclohexyl protons. The ∆δ values for this amide were calculated (Fig. 20): they are positive and negative on the right and left sides of the PGDA plane, respectively, as shown in 101a. Analogous results were also obtained with the PGDA derivative of (S)-2-methylbutanoic acid 102a. The absolute configuration de-

Figure 21 The ∆δ values (ppm) and NOEs supporting the proposed conformation of the PGME derivative of 4-methylcyclohexanecarboxylic acid 101c and the ∆δ values of the PGME derivative of (S)-2-methylbutanoic acid 102c. NOE [nuclear Overhauser effect], PGME, phenylglycine methyl ester. (Source : Ref. 88.)

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duced from Model B (Fig. 19), the conformation in agreement with that predicted by molecular mechanics calculations, was identical to the known absolute stereochemistry. With the promising results using PGDA as a chiral anisotropic reagent for the assignment of absolute configurations of chiral carboxylic acids obtained from the model studies discussed, PGME derivatives were subsequently examined for comparison. Condensation of (R)-(⫺)-PGME with 4-methylcyclohexanecarboxylic acid (101) gave amide 101c as an oil. Various NOE (nuclear Overhauser effect) experiments on this amide, particularly the NOE, which crossed the amide bond, suggest the conformation as shown (Fig. 21). The ∆δ values calculated for 101c are shown on the structure. Values having opposite signs are nicely arranged on the right and left sides of the PGME plane, respectively, in accord with the proposed conformation. To complete the comparison with PGDA, (R)- and (S)-PGME were condensed (PyBOP, HOBT) with (S)-2-methylbutanoic acid (102) and the ∆δ values recorded for the PGME derivative 102c (Fig. 21). The (S)-configuration assigned using Model B was identical to the known absolute configuration as also determined by using PGDA (Fig. 20, 102a). Although the ∆δ values for the PGME

Figure 22 The ∆δ values (ppm) recorded for PGME derivatives 103–106. PGME, phenylglycine methyl ester. (Source: Ref. 88.)

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amide 102c are smaller than those for the PGDA amide 102a as expected, they are still of practical magnitude for absolute stereochemistry assignment. Considering the ease of the preparation, the authors prefer PGME to PGDA, and thus for further derivatizations, PGME was subsequently applied to several carboxylic acids with known absolute configurations [88]. The results are summarized in 103–106 (Fig. 22). In all cases, the protons of ∆δ ⬎ 0 and ∆δ ⬍ 0 are on the right and left sides of the PGME plane, respectively, without exception, confirming the validity of the method. The absolute configurations predicted by the present method are identical with the known ones. It should be noted that PGME was applicable to rather complex compounds such as gibberellic acid (106). It was also found that the magnitude of the absolute values of ∆δ was proportional to the distance from the PGME moiety. The present method should be appropriate for absolute stereochemical assignment of natural products possessing a carboxyl group of type (I) as well as for primary alcohols 107 that can be routinely oxidized into carboxylic acid 108 (Eq. 1).

IV. CONCLUSIONS The number of applications appearing in the literature of the modified Mosher’s method to assign absolute stereochemistry is increasing dramatically, demonstrating the usefulness and reliability of the method. In principle, Professor Mosher’s original idea of using the magnetic anisotropy of an aromatic ring on a chiral reagent can be extended to a variety of functional groups. The use of PGME and PGDA derivatives is an example that shows that intense organic chemical considerations directed toward new chemical methodology for absolute stereochemistry determination can result in new chiral anisotropic reagents applicable to particular functional groups.

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Y. Jiao, T. Yoshihara, S. Ishikuri, H. Uchino, and A. Ichihara, Tetrahedron Lett., 37: 1039 (1996). J. Kobayashi, K. Yuasa, T. Kobayashi, T. Sasaki, and M. Tsuda, Tetrahedron, 52: 5745 (1996). J. Shin, Y. Seo, J.-R. Rho, E. Baek, B.-M. Kwon, T.-S. Feong, and S.-H. Bok, J. Org. Chem., 60: 7582 (1995). T. Ichiba, P. J. Scheuer, and M. Kelly-Borges, Tetrahedron, 51: 12195 (1995). C. M. Cerda-Garcia-Rojas and D. J. Faulkner, Tetrahedron, 51: 1087 (1995). T. Shibata, S. Kurihara, K. Yoda, and H. Haruyama, Tetrahedron, 51: 11999 (1995). L. Zeng, Z. Gu, X. Fang, P. E. Fanwick, C. Chang, D. L. Smith, and J. L. McLaughlin, Tetrahedron, 51: 2477 (1995). E. Palagiano, S. De Marino, L. Minale, R. Riccio, F. Zollo, M. Iorizzi, J. B. Carre´, C. Debitus, J. Provost, and L. Lucarain, Tetrahedron, 51: 3675 (1995). R. M. Rzasa, D. Romo, D. J. Stirling, J. W. Blunt, and M. H. G. Munro, Tetrahedron Lett., 36: 5307 (1995). M. Ojika, T. Nagoya, and K. Yamada, Tetrahedron Lett., 36: 7491 (1995). A. F. Barrero, E. Alvarez-Manzaneda, and A. Lara, Tetrahedron Lett., 36: 6347 (1995). P. A. Searle, R. K. Richter, and T. F. Molinski, J. Org. Chem., 61: 4073 (1996). L. Zeng, F.-E. Wu, Z. Gu, and J. L. McLaughlin, Tetrahedron Lett., 36: 5291 (1995). E. W. Schmidt and D. J. Faulkner, Tetrahedron Lett., 37: 3951 (1996). N. Sitachitta, M. Gadepalli, and B. S. Davidson, Tetrahedron, 52: 8073 (1996). Y. Guo, A. Madaio, E. Trivellone, G. Scognamiglio, and G. Cimino, Tetrahedron, 52: 8341 (1996). J.-L. Abad, J. Casas, F. Sa´nchez-Baeza, and A. Messeguer, J. Org. Chem., 60: 3648 (1995). R. W. Dunlop and R. J. Wells, Aust. J. Chem., 32: 1345 (1979). T. Kusumi, T. Hamada, M. O. Ishitsuka, I. Ohtani, and H. Kakisawa, J. Org. Chem., 57: 1033 (1992). T. Kusumi, T. Hamada, M. O. Ishitsuka, H. Ginda, and H. Kakisawa, Tetrahedron Lett., 33: 2019 (1992). M. Kobayashi, B. W. Son, M. Kido, Y. Kyogoku, and I. Kitagawa, Chem. Pharm. Bull., 31: 2160 (1983). N. Gonnella, K. Nakanishi, V. S. Martin, and K. B. Sharpless, J. Am. Chem. Soc., 104: 3775 (1982). N. Harada, S. L. Chen, and K. Nakanishi, J. Am. Chem. Soc., 97: 5345 (1975). (a) U. Shmueli, S. Carmely, A. Groweiss, and Y. Kashman, Tetrahedron Lett., 22: 709 (1981). (b) S. Carmely and Y. Kashman, J. Org. Chem., 48: 3517 (1983). I. Ohtani, T. Kusumi, Y. Kashman, and H. Kakisawa, J. Org. Chem., 56: 1296 (1991). Y. Inouye, I. Ohtani, T. Kusumi, Y. Kashman, and H. Kakisawa, Chem. Lett., 2073 (1990). A. Naganawa, Y. Ichikawa, and M. Isobe, Tetrahedron, 50: 8969 (1994). I. I. Ohtani, K. Hotta, Y. Ichikawa, and M. Isobe, Chem. Lett., 513 (1995). I. Kubo, V. Jamalamadaka, T. Kamikawa, K. Takahashi, K. Tabata, and T. Kusumi, Chem. Lett., 441 (1996).

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Modified 83. T. Kusumi, J. Syn. Org. Chem. Jpn., 51: 462 (1993). 84. T. Kusumi, T. Fukushima, I. Ohtani, and H. Kakisawa, Tetrahderon Lett., 32: 2939 (1991). 85. Y. Ichikawa, K. Tsuboi, and M. Isobe, J. Chem. Soc. Perkin Trans. 1, 2791 (1994). 86. T. Ohtsuka, Y. Itezono, N. Nakayama, A. Sakai, N. Shimma, M. Yanagisawa, K. Yokose, and H. Seto, Symposium Paper of the 35th Symposium on the Chemistry of Natural Products, Kyoto, Japan, pp. 306–313 (1993). 87. K. Iida, S. Fukuda, T. Tanaka, M. Hirama, S. Imajo, M. Ishiguro, K. Yoshida, and T. Otani, Tetrahedron Lett., 3: 4997 (1996). 88. Y. Nagai and T. Kusumi, Tetrahedron Lett., 36: 1853 (1995).

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6 Circular Dichroism Spectroscopy and the Absolute Stereochemistry of Biologically Active Compounds Nobuyuki Harada Tohoku University, Sendai, Japan

I. INTRODUCTION The theoretical calculation of circular dichroism (CD) spectra by the π-electron self-consistent field/configuration interaction/dipole velocity molecular orbital (SCF-CI-DV MO) method has become an important tool for the study of the absolute configuration of a variety of twisted π-electron systems including exciton-coupled systems [1–4]. We have assigned the absolute stereochemistry of (⫹)-1,8a-dihydro-3,8-dimethylazulene [(⫹)-1] isolated from a liverwort, a labile intermediate on the biosynthetic pathway to 1,4-dimethylazulene (2), as (8aS) by the application of this method to the calculation of the theoretical CD spectrum of the twisted tetraene system for comparison with that recorded for the natural product (Section III, Chart 1) [5,6]. In this case, we also succeeded in the experimental verification of the absolute configuration theoretically determined, by comparison of the CD spectrum of the natural product with those of synthetic chiral model compounds. We have also established the power of the π-electron SCF-CI-DV MO method for nonempirical determination of the absolute configuration of the more complicated, novel marine natural products of the halenaquinol family 18–27 with a new pentacyclic skeleton, isolated from tropical marine sponges (Section IV, Charts 2 and 3) [7,8]. In addition, we have confirmed that the absolute stereochemistry of the halenaquinol family, theoretically predicted, is correct by the first total synthesis of the chiral halenaquinols and halenaquinones [9–12]. The

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absolute stereochemistry of several related natural products was subsequently determined by the total synthesis of their natural enantiomers [12]. In addition to these natural products with twisted π-electron systems, the π-electron SCF-CI-DV MO method was successfully applied to the determination of another unusual class of natural products, the biflavones. Optically active biflavones are unique in the sense that they are chiral molecules devoid of stereogenic centers. One such chiral biflavone, (⫺)-4′,4⵮,7,7″-tetra-O-methylcupressuflavone [(⫺)-61], was isolated from the tropical plant Garcinia mangostana (Section V, Chart 4) [13]. This biflavone is composed of two flavone monomers linked at the 8- and 8′-positions; the monomeric subunits are almost perpendicular to each other with hindered rotation about the 8/8′-bond due to steric hindrance. Such molecules can exist as optically active compounds and belong to the class of chiral molecules known as atropisomers. Although the structure of 61 was reported in 1968 [14], the absolute stereochemistry remained undetermined for more than 20 years. Finally in 1992, we reported the determination of the absolute stereochemistry of the natural atropisomer as (aR) by theoretical calculation of the CD spectrum [13]. There are only two nonempirical methods for determining the absolute configuration of chiral organic compounds. One is the x-ray crystallographic Bijvoet method using the anomalous dispersion effect of heavy atoms within the molecule. The other is the theoretical CD method described in this chapter, which includes applications to the CD exciton chirality method. Although these two methods are based on totally different physical phenomena, they necessarily must give the same assignment of absolute configuration. In 1995, however, it was claimed that the absolute configuration of (⫺)-61 determined by x-ray crystallography disagreed with that obtained by the theoretical CD method [15]. This was a very interesting incident; however, the claim based on x-ray was later retracted [16,17]. Since then, we have succeeded in the total synthesis of (⫺)-61, confirming that the absolute configuration determined by the theoretical CD method was correct [18,19]. In this chapter, the theoretical calculation of CD spectra by the π-electron SCF-CI-DV MO method and its applications to the determination of the absolute stereochemistry of the biologically active compounds mentioned are described. The method has also been successfully applied to various chiral synthetic compounds with twisted π-electron systems for determining their absolute configurations [20–27].

II. COMPUTATIONAL METHODS A.

Molecular Structure

To calculate the CD spectra of chiral molecules with twisted π-electron systems, the molecular geometry and atomic coordinates of the most stable conformation

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are needed. The molecular geometries of chiral compounds with π-electron systems were optimized by using the molecular mechanics (MMP2) and/or MOPAC 93, AM1 programs [28,29]. The atomic coordinates obtained were then used for the next step: calculation of CD and ultraviolet (UV) spectra. B.

␲-Electron Self-Consistent Field/Configuration Interaction/Dipole Velocity Molecular Orbital Method: Numerical Calculation of Circular Dichroism and Ultraviolet Spectra

The CD and UV spectra of twisted, conjugated π-electron systems, including exciton-coupled systems, can be calculated by the π-electron SCF-CI-dipole velocity (DV) molecular orbital method (the computer program CD/NH-Sendai) [1–4,30]. In the dipole velocity method, the rotational strength Rba and dipole strength Dba are expressed as follows: Rba ⫽ 2(ϕa | ∇ |ϕb ) (ϕa | r ⫻ ∇ | ϕb )βM2 /(πσba )

(1)

Dba ⫽ 2(ϕa | ∇ | ϕb ) 2 βM2 /(πσba ) 2

(2)

where ∇ is the del operator, r is a distance vector, βM is the Bohr magneton, and σba is the excitation wave number of the transition a → b. The z axis components of the electric and magnetic transition moments are expressed, respectively, as (ϕa | ∇| ϕb ) z ⫽

冱 (C C ra

sb

⫺ Csa Crb ) 〈∇rs 〉 cos Zrs

(3)

bonds

(ϕa | r ⫻ ∇ | ϕb ) z ⫽

冱 (C

ra

Csb ⫺ Csa Crb )

bonds

〈∇rs〉 (Xrs cos Yrs ⫺ Yrs cos Xrs )

(4)

cos Zrs ⫽ (Zr ⫺ Zs )/Rrs

(5)

Xrs ⫽ (Xr ⫹ Xs )/2

(6)

where Cra is the coefficient of atomic orbital r in the wave function ϕa ; 〈∇rs 〉 is the expectation value of a dipole velocity vector ∇rs that is directed along the bond rs in the direction r → s; Xr, Yr, and Zr are the x, y, and z coordinates of an atom r, respectively; and Rrs is the interatomic distance between atoms r and s. In a similar way, the x and y components of the electric and magnetic transition moments were calculated. In the π-electron SCF-CI-DV MO calculation, the configuration interactions (CIs) among all singly excited states were included, and the following standard values of atomic orbital parameters were used: for aromatic carbon, ˚ ) ⫽ ⫺2.32 eV, W(C) ⫽ ⫺11.16 eV, (rr | rr) (C) ⫽ 11.13 eV, β(C ⫺ C, 1.388 A 7 ⫺1 ˚ 〈∇〉 (C ⫺ C, 1.388 A) ⫽ 4.70 ⫻ 10 cm ; for carbonyl oxygen, W(O)

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⫺17.28 eV, (rr | rr) (O) ⫽ 14.58 eV, β(C ⫽ O) ⫽ ⫺2.54 eV, 〈∇〉 (C ⫽ O) ⫽ 5.0 ⫻ 107 cm⫺1; for nitrogen, Z(N) ⫽ 2.0, W(N) ⫽ ⫺27.70 eV, (rr| rr)(N) ⫽ 17.44 eV, β(C ⫺ N) ⫽ ⫺1.899 eV, 〈∇〉(C ⫺ N) ⫽ 5.50 ⫻ 107 cm⫺1 [3,4]. The electric repulsion integral (rr | ss) was estimated by the Nishimoto–Mataga equation [31]. The resonance integral and del values were calculated by employing the following equations, respectively; ˚ )] β(C ⫺ C, 1.388 A ˚) β ⫽ [S/S (C ⫺ C, 1.388 A

(7)

˚ )/〈∇〉 (theor., 1.388 A ˚ )] 〈∇〉 (theor.) 〈∇〉 ⫽ [〈∇〉 (empir., 1.388 A

(8)

where S is the overlap integral. In the case of Method A, the component CD and UV bands were approximated by the Gaussian distribution [3]. ∆ε(σ) ⫽ ∑ ∆εk exp[⫺((σ ⫺ σk )/∆σ) 2 ] 2

ε(σ) ⫽ ∑ εk exp[⫺((σ ⫺ σk )/∆σ) ]

(9) (10)

where 2∆σ is the 1/e width of the bands. When the shape of the component CD and UV bands is much better approximated by the observed shape of the UV bands of model compounds than by the Gaussian distribution, the following equations (Method B) were adopted [3]: ∆ε(σ) ⫽ ∑ ∆εk f (σ ⫹ σo ⫺ σk )

(11)

ε(σ) ⫽ ∑ εk f (σ ⫹ σo ⫺ σk )

(12)

where f (σ) is the function describing the shape of component CD and UV bands and is taken from the observed UV spectrum of a model compound. For example, in the case of compounds containing naphthalene chromophores, the shape of the UV spectrum of naphthalene was adopted. On the other hand, when the observed CD and UV spectra are composed of many component bands, as in the case of the natural products discussed in this chapter, Method A was employed. The numerical calculations were carried out on a Sun S-4/10 Work Station and/or the NEC ACOS-3900 computer at the Computer Center of Tohoku University.

III. ABSOLUTE STEREOCHEMISTRY OF (ⴙ)-1,8aDIHYDRO-3,8-DIMETHYLAZULENE [(ⴙ)-1], A LABILE BIOSYNTHETIC PRECURSOR OF 1,4-DIMETHYLAZULENE [5,6] Considerable attention has focused on the chemistry of trinorsesquiterpenes with a 1,4-dimethylazulene skeleton isolated from marine soft corals and liverworts

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[32–36]. Takeda first isolated chiroptically active 1,8a-dihydro-3,8-dimethylazulene [(⫹)-1] (Chart 1), from the cell culture of the liverwort Calypogeia granulata Inoue, as a labile trinorsesquiterpenoid biosynthetic precursor of 1,4-dimethylazulene (2) [32]. The absolute stereochemistry of this intermediate is quite interesting from the viewpoint of the biosynthesis of trinorsesquiterpenes with 1,4-dimethylazulene skeleton. In fact, trinoranastreptene 3 [inflatene [35], also known as clavekerin B [36]], a probable biosynthetic precursor of (⫹)-1, was isolated from the same liverwort, and also from marine soft corals. Furthermore, Kitagawa and coworkers had established that the absolute structure of clavukerin A [(⫺)4], a related trinorsesquiterpene isolated from a marine soft coral, was (8S,8aS)(⫺)-3,8-dimethyl-1,2,6,7,8,8a-hexahydroazulene [33]. Since the π-electron system of 1 consists of a twisted, conjugated polyene chromophore, the problem of the absolute configuration is also interesting from the theoretical viewpoint. However, the absolute configuration of (⫹)-1 has remained undetermined because of its extreme instability and limited amounts of sample available. In the following, we discuss the absolute stereochemistry of (⫹)-1, as determined by theoretical calculation of the CD spectrum. In addition, we describe the experimental verification of the absolute configuration of (⫹)-1 by the synthesis of model compounds. The labile intermediate (⫹)-1 with a unique 1,8a-dihydroazulene skeleton

Chart 1 1,8a-Dihydro-3,8-dimethylazulene [(8aS)-(⫹)-1] and related compounds.

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showed very intense chiroptical activity, [α]D ⫹1165°, and intense CD Cotton effects (Fig. 1), suggesting a strongly twisted conjugated tetraene system. Therefore, it was reasonable to consider that the chiroptical activity of 1 is mainly due to the distortion of the π-electron chromophore. In order to predict the absolute configuration of (⫹)-1 theoretically, we performed the calculation of the CD curve of a model compound, 1,8a-dihydroazulene [(8aR)-9], on the basis of the π-electron framework approximation, using the SCF-CI-DV MO method [1–4]. The absolute configuration of 9 was arbitrarily chosen to be (8aR) for the calculation. The molecule has a very rigid skeleton in which the triene part in the sevenmembered ring is almost symmetric when reflected through the yz plane (Fig. 2). Therefore, it is also reasonable to consider that conjugation of the triene subunit with the additional double bound in the five-membered ring generates the chiroptical activity.

A.

Molecular Structure

Model compound (8aR)-9 has the following stereochemical features: The 1,8adihydroazulene ring skeleton is conformationally very rigid. The triene subunit

Figure 1 Circular dichroism and ultraviolet spectra of naturally occurring (8aS)-(⫹)1,8a-dihydro-3,8-dimethylazulene [(⫹)-1] in hexane. (Source: Ref. 5.)

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Figure 2 Stereochemistry of model compound (8aR)-1,8a-dihydroazulene [(8aR)-9] calculated by the MOPAC 93, AM1 program.

(3a-C/4-C/5-C/6-C/7-C/8-C) of the seven-membered ring is almost symmetric when reflected through the plane defined by 8a-C, 8a-H, and the midpoint between 5-C and 6-C. The additional double bond (2-C/3-C) in the five-membered ring is coplanar with the 3a-C/4-C double bond.

B.

Theoretical Determination of Absolute Stereochemistry by Calculation of Circular Dichroism Spectra

The calculated CD and UV curves of (8aR)-9 are illustrated in Fig. 3. The UV spectrum exhibited two allowed π → π* absorption bands: λmax 313 nm (ε 9900) and 219 nm (ε 27,300). The calculated values agree closely with the observed UV data of (⫹)-1: λmax 308.5 nm (ε 5400) and 227.5 nm (ε 25,600). Analysis of the calculations clarified that the two absorption bands at 313 and 219 nm consist of single π → π* transitions, respectively, and also revealed that, to a first approximation, the transition at 313 nm is polarized along the short axis of the molecule, whereas the transition at 219 nm is along the long axis. Therefore, the absorption band at 219 nm is stronger than that at 313 nm because the π-electron region along the long axis is larger than that along the short axis. The calculated values of the dipole strength are D ⫽ 12.6 ⫻ 10⫺36 and 24.4 ⫻ 10⫺36 cgs unit for the 313- and 219-nm transitions, respectively. In the calculated CD spectrum, these two transitions yielded negative and positive CD Cotton effects at longer and shorter wavelengths, respectively (Fig. 3): λext 313 nm (∆ε ⫺ 13.9) and 219 nm (∆ε ⫹ 46.2). These calculated values are in a good agreement with the observed data of (⫹)-1, though with reversed sign of the ∆ε values (Fig. 1): λext 314.0 nm (∆ε ⫹ 19.7) and 235.2 nm (∆ε ⫺ 47.4). The observed CD curve of (⫹)-1 showed a pattern characteristic of the 1,8a-dihydroazulene chromophore: the CD Cotton effect at longer wavelength is weaker than that at shorter wavelength, and the two Cotton effects are opposite in sign. It is evident that the observed CD spectral pattern characteristic of the

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Figure 3 Circular dichroism and ultraviolet curves of (8aR)-1,8a-dihydroazulene [(8aR)-9] calculated by the SCF-CI-DV MO method. SCF-CI-DV MO, self-consistent field/configuration interaction/dipole velocity molecular orbital. (Source: Ref. 5.)

1,8a-dihydroazulene system was well reproduced by the calculation when Figs. 1 and 3 are compared. The rotational strengths were calculated to be R ⫽ ⫺44.3 ⫻ 10⫺40 and ⫹103.3 ⫻ 10⫺40 cgs units for the 313- and 219-nm transitions of (8aR)-9, respectively. The observed CD curve of (⫹)-1 is almost a mirror image of that calculated for the model compound (8aR)-9, although the ∆ε values are slightly different. Accordingly, the absolute stereochemistry of the labile biosynthetic intermediate (⫹)-1 was theoretically predicted to be (8aS), as shown in structure 1. This conclusion was experimentally proved by the synthesis of model compounds, as described in the following section. The present results imply that the presumed precursor, trinoranastreptene 3, has the same absolute configuration at the bridgehead position as (⫹)-1. It should be noted that naturally occurring compounds (⫹)-1 and clavukerin A[(⫺)-4], partially hydrogenated azulene derivatives, have the same absolute configuration at the 8a position, regardless of the different sources.

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C.

Experimental Verification by the Synthesis of Model Compounds

In order to verify the absolute configuration of (⫹)-1 theoretically predicted by the SCF-CI-DV MO method, we synthesized model compounds, (1S,8aS)-(⫹)1,8a-dihydro-1-methoxy-8a-methylazulene [(⫹)-7] and (1S,8aS)-(⫹)-1,8a-dihydro-1-methoxy-6,8a-dimethylazulene [(⫹)-8], starting from the optically active Wieland-Miescher ketone (8aS)-(⫹)-10 (Scheme 1) [37–39]. These compounds were chosen as models since they resist oxidation to azulene, and the angular position 8a is substituted with a methyl group and thus not prone to epimerization. In addition, the methoxyl group makes the compounds less volatile, and hence the compounds can be easily handled.

Scheme 1 Synthesis of model compounds (⫹)-7 and (⫹)-8. (Source: Ref. 5.)

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Starting from the optically pure Wieland-Miescher ketone (8aS)-(⫹)-10 [39], the model compounds (1S,8aS)-(⫹)-7 and (1S,8aS)-(⫹)-8 were synthesized [5]. Thus, according to the procedure reported by Heathcock et al., the WielandMiescher ketone (8aS)-(⫹)-10 was converted to perhydroazulenone 11 (Scheme 1). The mixture of trans-fused ketone 11 and its cis-fused isomer was reduced to the corresponding glycols, then methylated to dimethyl ethers, from which a single stereoisomer (⫹)-12 was obtained as the major product by recrystallization from diethyl ether. Bromination of the ketal (⫹)-12 with PyHBr3 occurred exclusively at the C-7 position, yielding bromide (1S,3aR,4S,7R,8aS)-(⫹)-13. The position of the bromine atom in (⫹)-13 was determined by 1H NMR spectroscopy: H-7 appeared as a doublet of a doublet. Dehydrobromination of (⫹)-13 with t-BuOK and deketalization afforded enone 14. Successive eliminations of methanol and dehydrogenation with DDQ in the presence of a catalytic amount of p-TsOH yielded the desired trienone 15. Reduction of ketone 15 with LiAlH4 gave an epimeric mixture of alcohols 16; since these alcohols were extremely unstable, they were immediately used in the next reaction. For the purpose of dehydration of 16, various reaction conditions were examined, and iodine in benzene was finally found to be best. When a mixture of 16 and a catalytic amount of iodine in benzene was heated, the desired tetraene model compound (1S,8aS)-(⫹)-7 was obtained in good yield. Although (⫹)-7 was relatively unstable, it could be distilled in vacuo to give a faint yellow liquid: bp 35–45°C (0.067 kPa); [α]D ⫹ 393.3°. The other model compound (⫹)-8 was similarly synthesized: bp 60–70°C (0.029 kPa); [α]D ⫹ 323.8°. Since both of the dihydroazulenes 7 and 8 were unstable in pure form, they were stored in a freezer as dilute hexane solutions. The structure of (⫹)-7 was determined by 1H NMR studies including two-dimensional shift correlation and difference NOE spectra; all hydrogen atoms were fully assigned. The absolute configurations of these model compounds were also established by x-ray crystallographic analysis of synthetic intermediate (⫹)-13. The anomalous dispersion effect of the bromine atom led to the (1S,3aR,4S,7R,8aS) absolute configuration assignment as shown in Fig. 4. The CD and UV spectra of (1S,8aS)-(⫹)-7 are shown in Fig. 5; the UV spectrum exhibited a π → π* band of medium intensity at 324.3 nm (ε 6000) and an intense band at 223.2 nm (ε 23,700), which are characteristic of the 1,8adihydroazulene chromophore. In the region of these transitions, the CD spectrum showed a weak positive Cotton effect at λext 321.1 nm (∆ε ⫹5.7), and an intense negative Cotton effect at λext 221.3 nm (∆ε ⫺24.5). The other model compound (1S,8aS)-(⫹)-8 showed similar CD and UV spectra. The model compounds (⫹)-7 and (⫹)-8 have extra chirality due to the methoxyl group at the C-1 position. To exclude this extraneous stereogenic center, we synthesized the more ideal model compounds (8aS)-(⫹)-5 and (8aS)-(⫹)-

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Figure 4 ORTEP drawing of the absolute stereostructures of the two crystallographically independent molecules of (1S,3aR,4S,7R,8aS)-(⫹)-13 contained in an asymmetric unit. (Source: Ref. 6.)

6 starting from 11 following routine procedures (Scheme 2) [6]. The CD spectrum of (8aS)-(⫹)-5 showed more intense Cotton effects than that of (1S,8aS)-(⫹)-7 (Fig. 6); the CD spectrum of (8aS)-(⫹)-6 was similar to that of (8aS)-(⫹)-5. The CD spectra of model compounds (1S,8aS)-(⫹)-7, (1S,8aS)-(⫹)-8, (8aS)-(⫹)-5, and (8aS)-(⫹)-6 were similar in both sign and shape of the Cotton effects to that of dihydroazulene (⫹)-1. In the case of (8aS)-(⫹)-5, the CD intensity of the Cotton effects was also comparable to that of the natural product (⫹)1. Therefore, it was experimentally established that the natural dextrorotatory 1,8a-dihydro-3,8-dimethylazulene [(⫹)-1] had the (8aS) absolute configuration. The present results thus verified the theoretical determination of the absolute configuration of (⫹)-1 discussed previously. There were differences in the ∆ε values in the CD spectra of the natural product (⫹)-1 and model compounds (1S,8aS)-(⫹)-7 and (1S,8aS)-(⫹)-8, which may be due to the added chirality of the C-1 methoxyl substitution and/or to the difference in the positioning of the methyl groups. In fact, the methyl group at the C-6 position in (⫹)-6 diminished the ∆ε value, compared with that of (⫹)-5. Furthermore, it seems likely that the replacement of a hydrogen in the angular position 8a by a methyl group changes the molecular geometry to some extent and hence affects the CD activity. Nevertheless, the consistent pattern of

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Figure 5 Circular dichroism and ultraviolet curves of (1S,8aS)-(⫹)-1,8a-dihydro-1methoxy-8a-methylazulene [(⫹)-7] in EtOH. (Source: Ref. 5.)

a weak, long wavelength transition centered around 315–325 nm, with a stronger, oppositely signed band around 225 nm for these models, and the natural dihydroazulenes, along with the agreement of this pattern with the theoretical CD spectrum of (8aR)-9, indicated that the absolute stereoconfiguration of (⫹)-1 and related dihydroazulenes may be assigned on the basis of the theoretical calculations.

Scheme 2 Synthesis of model compounds (⫹)-5 and (⫹)-6. (Source: Ref. 6.)

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Figure 6 Circular dichroism and ultraviolet curves of (8aS)-(⫹)-1,8a-dihydro-8amethylazulene [(⫹)-5] in EtOH. (Source: Ref. 6.)

IV. ABSOLUTE STEREOCHEMISTRY OF THE HALENAQUINOL FAMILY, MARINE NATURAL PRODUCTS WITH A NOVEL PENTACYCLIC SKELETON, AS DETERMINED BY THE THEORETICAL CALCULATION OF CIRCULAR DICHROISM SPECTRA [7–12] In recent years, there have been many reports concerning isolation, structure determination, and biological activity studies of marine natural products. Many novel, biologically active compounds have been isolated from marine sponges. For example, Scheuer and coworkers isolated halenaquinone [(⫹)-18], an antibiotic with a novel pentacyclic skeleton, from the tropical marine sponge Xestospongia exigua collected off the Western Caroline Islands (Chart 2) [40]. The structure of (⫹)-18 was determined by x-ray crystallographic analysis. The absolute configuration, however, remained undetermined since the x-ray Bijvoet

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Chart 2 Halenaquinone [(12bS)-(⫹)-18] and its family of related marine natural products.

method for determination of the absolute configuration on the basis of the anomalous dispersion effect of heavy atoms could not be applied. Subsequent to Scheuer’s work, Kitagawa and coworkers isolated halenaquinol [(⫹)-19], the hydroquinone form of (⫹)-18, from the Okinawan sponge Xestospongia sapra, together with halenaquinol sulfate [(⫹)-20] [41]. Halenaquinol [(⫹)-19] was easily oxidized either at UV irradiation or through heating in air to give (⫹)-18. Furthermore, Nakamura and coworkers isolated xestoquinone [(⫹)-21], a powerful cardiotonic, from the same Okinawan sponge, X. sapra [42]. More recently, Schmitz and Bloor isolated a series of similar natural products, tetrahydroxestoquinol (23), the related dihydrofuran compound 24, adociaquinone A [(⫹)-25], adociaquinone B [(⫹)-26], and 3-ketoadociaquinone A (27), from a marine sponge, Adocia sp. collected in Truk Lagoon, in addition to (⫹)18 and (⫹)-21 (Chart 3) [43]. They also revealed that halenaquinone [(⫹)-18] and adociaquinone B [(⫹)-26] showed mild cytotoxicity. Harada and coworkers also reported (⫺)-prehalenaquinone [(⫺)-22], a putative biosynthetic precursor to (⫹)-18 and (⫹)-21 (Chart 2) [11]. Considering the interest in the halenaquinol family of marine natural products due to the physiological activity of these novel

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Chart 3 Other members of the halenaquinone family of marine compounds.

compounds, the determination of their absolute stereochemistries became an important problem. In this section, the application of the π-electron SCF-CI-DV MO method to calculate the CD spectra of the complex chromophore of the natural products of the halenaquinol family is described [7,8]. In the course of this work, we have also achieved the first total syntheses of (⫹)-halenaquinol [(⫹)-19], (⫹)halenaquinone [(⫹)-18], xestoquinol, (⫹)-xestoquinone [(⫹)-21], (⫺)-prehalenaquinone [(⫺)-22], (⫹)-adociaquinone A [(⫹)-25], and (⫹)-adociaquinone B [(⫹)-26] [9–12]. By these total syntheses, we experimentally confirmed that the absolute stereochemistry theoretically determined for these compounds with their unique, twisted π-electron system was correct. A.

An Attempt to Apply the Circular Dichroism Exciton Chirality Method for Determining the Absolute Configuration of the Halenaquinols

The CD exciton chirality method has been extensively applied to degenerate systems consisting of two identical chromophores such as dibenzoates, binaphthyls,

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bianthryls, and others [3]. In addition to such degenerate systems, the CD exciton chirality method is useful for determination of the absolute stereochemistry of nondegenerate systems that contain two different chromophores that experience exciton coupling. For example, Harada and Nakanishi determined the absolute configuration of chromomycin A3 by interpretation of the exciton coupling between the long-axis-polarized 1Bb transition of the naphthalene chromophore and the long-axis-polarized transition of the p-methoxybenzoate chromophore [3,44,45]. We considered that this method would be analogously applicable to the halenaquinols and related compounds. Specifically, we planned to synthesize benzoate derivative 32, then apply the CD exciton chirality method to the expected interaction between the naphthalene and benzoate chromophores (Scheme 3) [8]. Halenaquinol [(⫹)-19] was methylated in refluxing acetone with iodomethane in the presence of potassium carbonate with the exclusion of light to yield

Scheme 3 Strategy to apply the circular dichroism exciton chirality method.

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dimethyl ether (⫹)-28 as yellow needles (Scheme 3). To differentiate the two carbonyl groups at the 3- and 6-positions, dimethyl ether (⫹)-28 was selectively reduced with NaBH4 in the presence of CeCl3 ⋅ 7H2O, which catalyzes the regioselective 1,2-reduction of conjugated enones [46]. Keto-alcohol (⫹)-29 was obtained, and assignment of the C-3 stereocenter was secured by 1H NMR coupling constant data: J2β-H,3α-H ⫽ 8.0 Hz, J2α-H,3α-H ⫽ 8.0 Hz. The two large coupling constants indicate that the D-ring adopts a twisted half-chair conformation. This stereochemical assignment was subsequently confirmed by the NOE data of derivative 34 shown in Fig. 8 (Section IV.B). The alcohol was then protected as the t-butyldimethylsilyl ether (⫹)-30, followed by treatment with NaBH4 /CeCl3 ⋅ 7H2O in MeOH/CH2Cl2 to reduce the C-6 carbonyl group. However, the expected product, which was formulated as alcohol 31, was extremely unstable and could not be isolated, nor could it be trapped by benzoylation. Therefore, the preparation of naphthalene–benzoate compound 32, and hence the application of the CD exciton chirality method, was unsuccessful. B.

Attempted Application of the ␲-Electron SelfConsistent Field/Configuration Interaction/Dipole Velocity Molecular Orbital Method to Halenaquinol Dimethyl Ether [(ⴙ)-28]

We next attempted to apply the π-electron SCF-CI-DV MO method to halenaquinol dimethyl ether [(⫹)-28] to determine the absolute configuration of halenaquinol [(⫹)-19] since (⫹)-28 has a conjugated π-electron system composed of a naphthalene–ketone–furan–ketone chromophore that is twisted as a result of chiral center bearing the angular methyl group at the 12b-position [47]. Halenaquinol itself was not employed in this case because of its photochemical and thermal instability (even at 40°C). Furthermore, as a protecting group, a methyl ether group is better than others such as an acetate group, because the π-electron system of dimethyl ether (⫹)-28 containing only the lone pair of electrons of the ether oxygens is simpler than that of a diacetate. In the case of a diacetate, the πelectron system becomes complex as a result of the contribution of the ester carbonyls and their rotational conformation variability. Although we expected relatively intense CD Cotton effects for (⫹)-28, the CD spectrum showed only weak Cotton effects (Fig. 7). The weak intensity of the CD Cotton effects may be due to the existence of two carbonyl groups of strong electron-withdrawing nature, which makes the total π-electron system of (⫹)-28 less symmetric, and hence the electronic transitions more complex and weaker. Therefore, from the viewpoint of reliability of the determination, (⫹)28 was deemed unsuitable for the theoretical determination of the absolute configuration since it is rather difficult to discriminate small positive and negative ∆ε values. Although we actually carried out the calculation of the CD spectrum

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Figure 7 Circular dichroism spectrum of halenaquinol dimethyl ether [(12bS)-(⫹)-28] in EtOH derived from the natural sample of halenaquinol [(⫹)-19]. (Source: Ref. 47.)

of (⫹)-28, and the results suggested the 12bS absolute configuration for (⫹)-28 (as later shown to be correct), we don’t consider these results a convincing and unambiguous determination of the absolute configuration because of the small ∆ε values of CD Cotton effects. C.

Circular Dichroism Spectra of Naphthalene–Diene Derivatives with Twisted ␲-Electron Systems

As described, we could not isolate alcohol 31 because of its instability (Scheme 3) and therefore failed to determine the absolute configuration of the halenaquinols by application of the nondegenerate CD exciton chirality method. However, we were very happy to find that the reduction of ketone (⫹)-30 with NaBH4 / CeCl3 gave the elimination/addition products (⫺)-33 and (⫹)-34 instead of 31 (Scheme 4) [7,8]. It was presumed that (⫺)-33 and (⫹)-34 are derived from 31 by elimination of the hydroxyl group and subsequent addition of methanol at the 4-position. Furthermore, treatment of the reaction mixture with a catalytic amount of aqueous HCl following the borohydride reduction of (⫹)-30 gave trans-methoxy diene (⫺)-33 and cis-methoxy diene (⫹)-34 in moderate yields, 44% and 20%, respectively. The structures of acetal epimers (⫺)-33 and (⫹)-34 were determined on the basis of the spectroscopic data, with the relative stereochemistries unambiguously determined by the 1H NMR coupling constant data and NOE enhancement data (Fig. 8). It was very interesting to note that naphthalene–diene compounds (⫺)-33 and (⫹)-34 exhibited much stronger CD Cotton effects than other halenaquinol derivatives. For example, the UV spectrum of trans-methoxysilyl ether (⫺)-33

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Scheme 4 Synthesis of naphthalene–diene derivatives (⫺)-33 and (⫹)-34.

Figure 8 NOE and coupling constant data and conformation of naphthalene–diene derivative (⫹)-34. (Source: Ref. 8.)

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Figure 9 Observed circular dichroism and ultraviolet spectra of halenaquinol transmethoxydiene derivative (3R,4R,12bS)-(⫺)-33 in MeOH. (Source: Refs. 7 and 8.)

showed two intense π → π* bands (Fig. 9), a broad band at 324 nm (ε 27,000) with complex vibrational structure, and a sharp band at 218 nm (ε 42,000). In these corresponding regions, the CD spectrum of (⫺)-33 exhibited three major, intense Cotton effects: λext 338 nm (∆ε ⫹6.4), 301 nm (∆ε ⫺23.3), and 229 nm (∆ε ⫹40.9). The other naphthalene–diene compound (⫹)-34 also exhibited three major CD Cotton effects of similar intensity and of the same sign as those of (⫺)-33. These results clearly indicated that the CD Cotton effects originated mainly from the π-electron chromophore composed of the naphthalene–diene moiety, which is twisted by the angular methyl group at the 12b-position. Furthermore, the additional chiralities due to the silyloxy group at the 3-position and the methoxy group at the 4-position are only minor contributors to the CD Cotton

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effects. These observations were critical to our choice of a model compound for the theoretical study discussed in the next section. Thus, these naphthalene–diene compounds are ideal systems for the theoretical determination of the absolute stereochemistry by application of the π-electron SCF-CI-DV MO method. It is interesting to note that the orientation of the methoxyl group apparently dictated the sign of [α]D: negative for 33 and positive for 34. D.

Application of the ␲-Electron Self-Consistent Field/ Configuration Interaction/Dipole Velocity Molecular Orbital Method to Naphthalene–Diene Derivatives

As a model compound for the theoretical calculation of CD spectra, we adopted the molecule (12bS)-35, retaining the essential part of the π-electron system contained in the naphthalene-diene compounds 33 and 34 (Fig. 10) [7,8]. Of specific importance, in addition to the naphthalene and conjugated diene chromophores, the lone pair of electrons of the two methyl ether and furan ring oxygens were also included. The absolute configuration of 35 was arbitrarily chosen to be 12bS for the calculation. The molecular geometry of (12bS)-35 was calculated using the MOPAC 93, AM1 programs and is illustrated in Fig. 10. The molecular framework of this model compound is relatively rigid, and the terminal cyclohexene ring adopts a half-chair conformation. This conformation was supported by the 1H NMR coupling constant and NOE enhancement data of compounds (⫺)33 and (⫹)-34 (Fig. 8): the coupling constant J1α-H,2β-H ⫽ 12.0 Hz indicated the trans-diaxial relation between these protons. By contrast, J2β-H,3α-H ⫽ 8.0 Hz was smaller than J1α-H,2β-H and is equal to J2α-H,3α-H, indicating the half-chair conformation of the terminal cyclohexene ring. The C-6/C-5a double bond and naphthalene

Figure 10 Model compound (12bS)-35 and its stable conformation calculated by the MOPAC 93, AM1 programs.

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chromophore of (12bS)-35 constitute a clockwise helicity (dihedral angle of 5a6-6a-7: ⫹170°), whereas the conjugated diene moiety by itself constitutes a counterclockwise helicity (dihedral angle of 3a-12c-5a-6: ⫺167°). The helical sense of these two moieties is not changed even if the terminal cyclohexene ring adopts the higher-energy boat conformation. Thus, the sense of the twist of the conjugated π-electron system is governed solely by the chirality of the angular methyl group at the 12b position. Calculation of the theoretical CD and UV spectra of (12bS)-35 by the πelectron SCF-CI-DV MO method gave the curves illustrated in Fig. 11. The UV spectrum exhibited two intense π → π* bands: a broad band at 349 nm (ε 29,900)

Figure 11 Circular dichroism and ultraviolet curves of the model compound (12bS)35 calculated by the π-electron SCF-CI-DV MO, self-consistent field/configuration interaction/dipole velocity molecular orbital method. (Source: Refs. 7 and 8.)

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and a sharp band at 219 nm (ε 40,300). These calculated values agreed closely with the observed UV data of (⫺)-33: λmax 324 nm (ε 27,000) and 218 nm (ε 42,000) (Fig. 9). In the corresponding region, the calculation predicted three principal CD Cotton effects: a weak positive band at 378 nm (∆ε ⫹3.3), a negative band of medium intensity at 322 nm (∆ε ⫺22.4), and a positive intense one at 223 nm (∆ε ⫹35.5). These theoretically obtained CD bands were in a good agreement with the observed data of (⫺)-33: λext 338 nm (∆ε ⫹6.4), 301 nm (∆ε ⫺23.3), and 229 nm (∆ε ⫹40.9) (Fig. 9). The calculation thus reproduced the basic pattern of the observed CD and UV spectral curves, including the sign, position, intensity, and shape of the bands. Since the absolute configuration of the model compound 35 was fixed to be (12bS), comparison of the calculated and observed CD data leads to the conclusion that (⫺)-33 and other related naphthalene–diene compounds have the (12bS) absolute configuration. Accordingly, the absolute stereochemistry of halenaquinol [(⫹)-19] was theoretically determined to be (12bS). Since UV irradiation of (⫹)-19 in the presence of air gave halenaquinone [(⫹)-18,] and solvolysis of halenaquinol sulfate [(⫹)-20] quantitatively yielded halenaquinol [(⫹)-19], the absolute stereostructures of (⫹)-18 and (⫹)-20 were also established as (12bS), respectively.

E.

Circular Dichroic Power of a Twisted Naphthalene– Diene System

In the case of (8aS)-(⫹)-1,8a-dihydro-3,8-dimethylazulene [(⫹)-1], the composition of the apparent CD and UV bands was rather simple, because each of the apparent bands was composed of a single electronic transition. In contrast, the π-electron chromophores of the twisted naphthalene–diene systems 33–35 are complex and have no symmetric character [8]. Therefore, to confirm the applicability of the π-electron SCF-CI-DV MO method to such complex systems, it is important to analyze the composition of the apparent CD and UV bands theoretically obtained. As illustrated in Fig. 12, there are nine major electronic transitions that contribute to the CD and UV bands. The first and second electronic transitions, with weak positive rotational strengths at 374.5 and 351.6 nm, respectively, generate the weak positive Cotton effect at 378 nm (Fig. 12). The third electronic transition, with an intense negative rotational strength at 324.4 nm, results in the negative Cotton effect at 322 nm, and the sixth electronic transition, with a strong positive rotational strength, dominates the intense positive Cotton effect at 223 nm. The correspondence between the calculated CD rotational strengths and the observed CD Cotton effects in the spectra of (⫺)-33 and (⫹)-34 is quite evident. Therefore, this analysis using hypothetical model (12bS)-35 for comparison with (⫺)-33 and (⫹)-34 makes the theoretical determination of the absolute configurations of halenaquinol [(⫹)-19] and related compounds more reliable.

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Figure 12 Calculated rotational and dipole strengths of the transitions of the model compound (12bS)-35. (Source: Ref. 8.)

F.

The Synthetic Strategy for the First Total Synthesis of (ⴙ)-Halenaquinol [(ⴙ)-19] and (ⴙ)-Halenaquinone [(ⴙ)-18]

As discussed, we succeeded in the theoretical determination of the absolute stereochemistry of novel marine natural products of the halenaquinol family. It is quite natural that chemists, as the next step, want to prove experimentally the absolute configurations theoretically predicted. So, we undertook the synthesis of halenaquinol [(⫹)-19] and halenaquinone [(⫹)-18] in their natural enantiomeric forms in order to corroborate their structures by comparison of the CD spectra of the synthetic samples with that of the natural ones [9]. Our retrosynthetic strategy centered around the cycloaddition of a transient o-quinodimethane generated from benzocyclobutene 39 with enone 41 as the key, convergent step, forming 38 with the carbocyclic halenaquinol skeleton (Scheme

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Scheme 5 Retrosynthetic analysis for halenaquinol [(12bS)-(⫹)-19].

5). The naphthoquinone moiety of (12bS)-(⫹)-18, which can be reduced to halenaquinol [(⫹)-19], can be obtained by oxidative cleavage of the hydroquinone dimethyl ether (12bS)-(⫹)-28; the furan ring of (12bS)-(⫹)-28 was considered accessible by oxidation of triol 36. The diosphenol moiety of 36, would be obtained by the air oxidation of ketone 37 [48,49]. The tetracyclic skeleton of 37 would be constructed by the key Diels-Alder reaction between the transient oquinodimethane 40 (generated from benzocyclobutane 39) and enone 41. Further analysis for dienophile 41 suggested that this critical synthon could be derived from the Wieland-Miescher ketone 43 (Scheme 6), in which the onecarbon unit ultimately to become C-4 in 19 is introduced by application of Stork’s reductive hydroxymethylation procedure to 43 [50]. In this synthetic route, the absolute configuration of the bridgehead methyl group of optically active 43 is retained as that of the corresponding methyl group in the final product. Since the absolute configurations of halenaquinol and halenaquinone were predicted to be

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Scheme 6 Retrosynthesis of halenaquinol [(12bS)-(⫹)-19]; preparation of enone 41.

12bS as discussed, the synthesis of the natural enantiomeric forms of halenaquinol and halenaquinone should start from the (8aR)-(⫺) enantiomer of 43. Among several synthetic methods to prepare optically active 43, the most practical is the asymmetric cyclization of triketone 44 with a catalytic amount of optically active proline, which was independently discovered by Hajos’s and by Eder’s groups [39]. We achieved the first total synthesis of (⫹)-halenaquinol [(⫹)-19] and (⫹)halenaquinone [(⫹)-18] by following this strategy [9]. The carbonyl group at the 1-position of optically pure (8aR)-(⫺)-43 was selectively protected to give monoacetal (⫺)-45, which was then reductively hydroxymethylated according to the procedure of Stork [50] (Scheme 7): enone (⫺)-45 was reduced with lithium in liquid ammonia, and the resultant enolate was trapped as the trimethylsilyl ether 46. Regeneration of the enolate anion by treatment of 46 with methyllithium and then addition of gaseous formaldehyde gave keto alcohol (⫹)-42 as the sole stereoisomer in 82% overall yield from (⫺)-45. Keto alcohol (⫹)-42 was reduced with lithium tri-sec-butylborohydride (L-Selectride), producing cis-diol (⫹)-47 in 92% yield, which was then converted to ketodiol (⫹)-48 in 98% yield by treatment with aqueous p-toluenesulfonic acid ( p-TsOH). The relative stereochemistry of the 6(ax)-hydroxyl and 5(eq)-hydroxymethyl groups of (⫹)-48 was secured by the 1H NMR coupling constant data of its acetonide (⫹)-51 (Fig. 13). Formation of the tosylhydrazone of (⫹)-48, followed by treatment with methyllithium, gave olefin (⫺)-49 in quantitative yield. Next, the 1,3-diol of (⫺)49 was protected as the acetonide to give olefin (⫺)-50. Finally, allylic oxidation of acetonide olefin (⫺)-50 with CrO3 /3,5-dimethylpyrazole [51] gave conjugated

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Scheme 7 Synthesis of chiral dienophile 41.

Figure 13 1H nuclear magnetic resonance (NMR) coupling constant data of keto-acetonide (⫹)-51. (Source: Ref. 47.)

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enone (⫹)-41 in 63% yield, which was used as the dienophile in the Diels-Alder reaction with 40 generated in situ from 3,6-dimethoxybenzocyclobutene (39). Although dimethoxybenzocyclobutene 39 had been previously synthesized by photocycloaddition chemistry [52], we prepared it by pyrolysis of sulfone 55, which itself was prepared from 2,3-dimethyl-1,4-dimethoxybenzene [52] (Scheme 8) [9]. Bromination of 52 with N-bromosuccinimide (NBS), followed by treatment of the resulting dibromide 53 with sodium sulfide in aqueous ethanol, gave 54 in 70% yield. Oxidation of 54 with m-chloroperbenzoic acid (MCPBA) in dichloromethane afforded sulfone 55 in 89% yield. Various reaction conditions were examined for the thermal elimination of sulfur dioxide, and finally it was found that direct heating of solid sulfone 55 without solvent afforded the desired 3,6-dimethoxybenzocyclobutene (39) in moderate yield. Thus, crystals of 55 were pyrolyzed at 305–310°C in a muffle furnace under a stream of nitrogen to give 39 in 48% yield. The Diels-Alder reaction of o-quinodimethane 40 derived from 39 and (⫹)41 was achieved by heating a benzene solution of the two reactants in a sealed tube at 210–215°C for 20 h, giving tetrahydronaphthalene derivative (⫹)-38 in 33% yield (Scheme 9). Numerous reaction conditions were examined to increase the yield, but no further improvement was achieved. The 13C NMR spectrum of (⫹)-38 indicated that the product was composed of a single stereoisomer. However, the relative stereochemistry of the newly formed chiral centers was not investigated further since they were eliminated in the subsequent dehydrogenation. To dehydrogenate the tetrahydronaphthalene moiety of (⫹)-38, a benzene solution was refluxed with 2,3-dichloro-5,6-dicyano-1,4-benzoquinone (DDQ) to

Scheme 8 Synthesis of 3,6-dimethoxybenzocyclobutene (39).

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Scheme 9 Synthesis of halenaquinol derivative 56.

afford naphthalene derivative (⫺)-37 in 89% yield, which was then subjected to air oxidation in the presence of base [48]. Thus, oxygen was bubbled through a solution of (⫺)-37 and potassium t-butoxide in t-butyl alcohol for 5 h, and the mixture was worked up with aqueous ammonium chloride to give diosphenol 56 in 90% yield. The structure of 56 was secured by the 1H NMR (a sharp singlet at δ 7.60 ppm disappeared upon addition of D2O), UV spectra (a red shift and a hyperchromic effect of the longer-wavelength UV absorption band upon adding aqueous NaOH), along with high-resolution MS data. Deprotection of the acetonide group of 56 upon treatment with 60% aqueous acetic acid yielded triol 36, which was subjected to the next reaction without purification (Scheme 10). The oxidation of the primary and secondary hydroxyl groups of 36 and subsequent cyclization to form the furan ring were accomplished by treatment with dimethyl sulfoxide (DMSO) and 1,3-dicyclohexylcarbodiimide (DCC) in benzene in the presence of trifluoroacetic acid and pyridine (PTFA), giving the desired halenaquinol dimethyl ether [(12bS)-(⫹)-28] with the furan–diketone system, in 44% overall yield from (⫺)-37. All of the spectroscopic data of the synthetic sample of (⫹)-28 were identical with those of an authentic sample of (⫹)-28 prepared from natural halenaquinol. The dimethyl ether groups of (12bS)-(⫹)-28 were next oxidatively cleaved with cerium (IV) ammonium nitrate (CAN) in aqueous methanol, affording halenaquinone [(12bS)-(⫹)-18] in 45% yield (Scheme 10). The 1H NMR and UV spectra of the synthetic sample agreed with those of natural (⫹)-18. Finally, (12bS)-

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Scheme 10 Synthesis of halenaquinone [(12bS)-(⫹)-18] and halenaquinol [(12bS)(⫹)-19].

(⫹)-18 was reduced with aqueous sodium hydrosulfite in acetone to give halenaquinol [(12bS)-(⫹)-19] in near-quantitative yield. The 1H NMR spectrum of (12bS)-(⫹)-19 in DMSO-d6 exhibited two broad singlets at δ 9.6 and 9.8 that were due to the phenolic hydroxyl groups, which disappeared when D2O was added. The remaining 1H NMR peaks and UV spectrum curve were in good agreement with those of the natural sample. The first total synthesis of halenaquinol [(12bS)-(⫹)-19] and halenaquinone [(12bS)-(⫹)-18] with a novel polyketide skeleton had been thus accomplished.

G.

Circular Dichroism Spectrum of Halenaquinol Dimethyl Ether [(12bS)-(ⴙ)-28] and Experimental Proof of the Absolute Stereochemistry of the Halenaquinol Family

In the total synthesis of halenaquinol [(12bS)-(⫹)-19] and of halenaquinone [(12bS)-(⫹)-18], we started with the Wieland-Miescher ketone (8aR)-(⫺)-43, as presented earlier, and thus the synthetic sample of (⫹)-28 had the (12bS) absolute configuration. If the theoretical determination of the absolute stereochemistry of the halenaquinol family based on the π-electron SCF-CI DV MO method discussed in Section IV.D was correct, the CD spectrum of the synthetic sample must be identical to those of the authentic sample of (⫹)-28 derived from natural

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Figure 14 Circular dichroism spectrum of the synthetic sample of halenaquinol dimethyl ether [(12bS)-(⫹)-28] in EtOH. (Source: Ref. 47.)

halenaquinol. This was verified as shown in Figs. 7 and 14: the CD spectra of the synthetic and natural samples are identical. These results lead to the experimentally confirmed, unambiguous determination that the absolute stereochemistry of (⫹)-halenaquinol and (⫹)-halenaquinone is 12bS. In addition, these syntheses also proved that the theoretically predicted absolute configurations of halenaquinol compounds were correct.

H.

The First Total Synthesis of (ⴙ)-Xestoquinone [(ⴙ)-21] and Xestoquinol, and Their Absolute Stereochemistries

In similar fashion, we also achieved the first total synthesis of (⫹)-xestoquinone [(⫹)-21] and xestoquinol [10,11]. The CD spectrum of the synthetic sample of (12bS)-21 was identical to that of natural xestoquinone [(⫹)-21] [10]. Thus, beginning with Wieland-Miescher ketone (8aR)-(⫺)-43, synthetic (12bS)-xestoquinone [(⫹)-21] was prepared that was identical to natural xestoquinone [(⫹)-21], which must therefore also have the 12bS absolute configuration. Xestoquinone [(12bS)-(⫹)-21] was subsequently reduced to xestoquinol, so this hydroquinone also has the 12bS absolute configuration. The absolute stereochemistry of (⫹)xestoquinone and of xestoquinol was thus unambiguously determined. The synthetic sample of xestoquinone [(12bS)-(⫹)-21] was also converted into the adociaquinones A [(⫹)-25] and B [(⫹)-26] [12]. Therefore, the absolute configurations of these members of the halenaquinol family were also determined, as shown in Chart 3.

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I.

Efficient Synthesis of (ⴙ)-Halenaquinone [(ⴙ)-18] and (ⴙ)-Xestoquinone [(ⴙ)-21] via Prehalenaquinol Dimethyl Ether [(ⴚ)-57]

During the synthetic studies described, we found a new procedure to convert triol 36 into dihydrofuran (3S,3aS,12bS)-(⫺)-57 (Scheme 11). When 36 was treated with a limited amount of DMSO/DCC (5.0 equiv.)/PTFA (1.3 equiv.) in benzene at room temperature for 2.5 h, the formation of dihydrofuran-alcohol (⫺)-57 instead of furan-diketone (⫹)-28 resulted (75%). When (⫺)-57 was resubjected to the DMSO/DCC (8.6 equiv.)/PTFA (6.0 equiv.) oxidation at room temperature for 18 h, the final product (⫹)-28 was obtained. Therefore, dihydrofuran-alcohol (⫺)-57 is a synthetic intermediate of the Pfitzner-Moffatt oxidation of triol 36. When we found this unexpected product, we immediately considered that dihydrofuran alcohol (⫺)-57 could be a pivotal synthetic intermediate common to the halenaquinone and xestoquinone natural products. If dihydrofuran alcohol (⫺)-57 was dehydrated, the newly formed double bond should migrate into the five-membered ring to give xestoquinol dimethyl ether [(⫹)-58] (Scheme 12). Since the 3α-hydroxyl group and 3aβ-hydrogen of (⫺)-57 are in a trans-diaxial relationship, the configuration was ideal for dehydration. A solution of dihydrofuran alcohol (⫺)-57 in benzene was refluxed for 24 h in the presence of p-TsOH to yield xestoquinol dimethyl ether [(⫹)-58] as expected in 77% yield. Compound (⫹)-58 had previously been converted to xestoquinone [(⫹)-21] and xestoquinol

Scheme 11 Synthesis of dimethylhalenaquinol [(12bS)-(⫹)-28] from prehalenaquinone derivative (3S,3aS,12bS)-(⫺)-57.

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Scheme 12 Synthesis of xestoquinone [(12bS)-(⫹)-21] and xestoquinol [(12bS)-(⫹)59] from prehalenaquinone derivative (57).

[11]. This synthetic route is more efficient than our previous total synthesis of xestoquinone and xestoquinol. We have thus achieved the total syntheses of xestoquinol, halenaquinol, and related compounds from the common synthetic intermediate (⫺)-57. J. Synthesis and Absolute Stereochemistry of Prehalenaquinone and Prehalenaquinol, Putative Biosynthetic Precursors Common to Halenaquinone, Xestoquinone, and Related Natural Products The chemical conversion of dihydrofuran alcohol (⫺)-57 into xestoquinol, halenaquinol, and related compounds described poses a very attractive explanation for the biosynthesis of these natural products in marine sponges. That is, dihydrofuran–alcohol–quinone 22 and dihydrofuran–alcohol–hydroquinone 60 are putative biosynthetic precursors of both the halenaquinols and xestoquinols (Scheme 13). As in the case of the chemical conversion, dehydration of 22 or 60 would lead to xestoquinone [(⫹)-21] or xestoquinol [(⫹)-59], respectively, and oxidation of 22 or 60 would lead to halenaquinone [(⫹)-18] or halenaquinol [(⫹)-19]. The related compound, dihydrofuran–alcohol 24 (Chart 3), isolated by Schmitz [43], may be an earlier intermediate than precursors 22 and 60. Oxidation of the secondary alcohol group in the A-ring of 24 would give a diketone, which would be expected to tautomerize to 60; hydroquinone 60 could be easily oxidized to

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Scheme 13 Synthesis of prehalenaquinone [(3S,3aS,12bS)-(⫺)-22] and prehalenaquinol [(3S,3aS,12bS)-60].

quinone 22. Therefore, if the biosynthesis of halenaquinone, xestoquinone, and related compounds proceeded as postulated here, we thought biosynthetic precursors 22 and/or 60 may be found in marine sponges. To this end, samples of putative biosynthetic precursors (⫺)-22 and 60 were synthesized (Scheme 13). Treatment of dimethyl ether (⫺)-57 with ammonium cerium(IV) nitrate (CAN) afforded quinone (⫺)-22 in 79% yield, which was then quantitatively converted to hydroquinone 60 by treatment with sodium hydrosulfite. The structures of these compounds were confirmed by spectroscopic and physical data. From the synthetic route described, the absolute stereochemistry of (⫺)-22 and 60 must be (3S,3aS,12bS). The putative biosynthetic precursors to the halenaquinol family of natural products, (⫺)-22 and 60, were named prehalenaquinone and prehalenaquinol, respectively. We next sought to determine whether (⫺)-22 and 60 were actually present in the marine sponges from which the halenaquinols and xestoquinols were isolated. Using the synthetic samples of (⫺)-22 and 60 as standards, we analyzed by high-performance liquid chromatography (HPLC) the components of an extract of the Okinawan marine sponge Xestospongia sapra from which halenaquinone [(⫹)-18] and xestoquinone [(⫹)-21] had been isolated (HPLC conditions: silica gel, EtOA/hexane/MeOH 100 :100 :5, or ODS C18 column, 50% aqueous MeOH). Although no peak corresponding to prehalenaquinol (60) was detected, a peak corresponding to prehalenaquinone [(⫺)-22] was observed, and we subsequently succeeded in the isolation of (⫺)-22 from this source, as follows: The

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wet sponge (305 g) was treated with acetone, and the crude extract thus obtained upon removal of the solvent (6.5 g) was successively separated and purified by HPLC (silica gel, EtOAc, hexane/EtOAc 1: 3, and hexane/EtOAc/MeOH 100: 100 :2), giving (⫺)-22 (8.3 mg, 0.13% from the acetone extract). The spectral and physical data, including the CD data, of natural (⫺)-22 thus isolated were identical with those of synthetic (⫺)-22. These results imply that dihydrofuran– alcohol (⫺)-22 may be a biosynthetic precursor of the halenaquinone and xestoquinone families of compounds.

V.

ABSOLUTE STEREOCHEMISTRY OF A NATURAL ATROPISOMER, THE BIFLAVONE (ⴚ)-4′,4ⵯ,7,7″-TETRAO-METHYLCUPRESSUFLAVONE [(ⴚ)-61] [13,18,19]

The title biflavone was isolated from Garcinia mangostana L. [13] and was identified as optically active cupressuflavone tetramethyl ether [(⫺)-61] by a direct comparison of spectral data with those of the authentic sample isolated earlier from Araucaria cunninghamii and A. cookii (Chart 4) [14]. Compound 61 is a dimeric flavone as indicated by the MS spectral data ([M]⫹, m/z 594) and has a chiral C2-symmetric structure that exhibits an optical rotation of [α]D27 ⫺25.3°, with only seven signals in the 1H NMR spectrum, and 15 peaks in the 13C NMR spectrum. We established the dimeric structure with an 8-8″-linkage of 61 on the basis of detailed 1H and 13C NMR studies, including 1H NOESY, heteronuclear gated decoupled 13C NMR, 1H-13C COSY, and long-range 1H-13C COSY (COLOC) methods, and have achieved full assignment of all proton and carbon

Chart 4 Natural biflavone atropisomer (aR)-4′,4⵮,7,7″-tetra-O-methylcupressuflavone [(⫺)-(aR)-61], and monomer 62.

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signals, leading to the structure as shown in Chart 4. The absolute stereochemistry of (⫺)-61, however, had never been determined. A.

Ultraviolet and Circular Dichroism Spectra of the Natural Atropisomer of Biflavone (ⴚ)-61

The UV spectrum of biflavone (⫺)-61 exhibited three intense π → π* bands at 324.2 nm, 273.0 nm, and 225.8 nm. The two UV bands at 324.2 nm and 273.0 nm were accompanied by bisignate CD Cotton effects, respectively (Fig. 15). For the first UV band at 324.2 nm, positive and negative CD Cotton effects were observed at 362.0 and 326.2 nm, respectively; for the second UV band at 273.0 nm, a negative CD shoulder around 300 nm and a positive Cotton effect at 267.5

Figure 15 Circular dichroism and ultraviolet spectra of biflavone (⫺)-4′,4⵮,7,7″-tetraO-methylcupressuflavone [(aR)-(⫺)-61] in EtOH. (Source: Ref. 13.)

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nm were seen. Therefore, these Cotton effects seemed to originate from exciton coupling between the two flavone chromophores. To determine absolute stereochemistry by application of the CD exciton chirality method, the direction and position of the transition moments in the molecule must be determined. From a qualitative viewpoint, the UV bands at 324.2 and 273.0 nm may be assigned to the intrachromophoric charge transfer, and the long-axis-polarized-transitions of the p-methoxycinnamoyl and p-methoxybenzoyl chromophores, respectively. On the basis of such a simple assignment of the transition moments, the S configuration had once been deduced for biflavone (⫺)-61 [53]. However, the actual flavone chromophore is a composite of these two moieties. Furthermore, it is difficult to predict the exciton chirality between the long axes of the two p-methoxycinnamoyl subunits, even if the transition at 324.2 nm may be ascribed to the p-methoxycinnamoyl moiety, because the two transition moments incline toward the inside of the molecule. It is thus not simple to deduce the absolute stereochemistry of such a complex system by application of the CD exciton chirality method. In such a case, the theoretical calculation of CD and UV spectra by means of the π-electron SCF-CI-DV MO method is extremely useful for determination of the absolute stereochemistry. B.

Theoretical Calculation of the Ultraviolet and Circular Dichroism Spectra of the Natural Atropisomer of Biflavone (ⴚ)-61 and Nonempirical Determination of Absolute Stereochemistry

We adopted the biflavone 61 with the (aR) absolute configuration as the model compound for the calculation of the CD and UV spectra. The atomic coordinates of (aR)-61 were obtained by the molecular mechanics calculation using the MMP2 program (Fig. 16), and the CD and UV spectral curves of (aR)-61 were calculated by use of the π-electron SCF-CI-DV MO method [1–4]. The calculated UV spectrum exhibited three π → π* bands: a broad and intense band at λmax 322.6 nm (ε 66,200), a weak band appearing as a shoulder around 270 nm, and an intense band at 226.8 nm (ε 78,300) (Fig. 17). These calculated values agreed well with the observed UV data of (⫺)-61: λmax 324.2 nm (ε 40,900), 273.0 nm (ε 41,400), and 225.8 nm (ε 51,800) (Fig. 15 and 17). The basic pattern of the UV spectral curve of biflavone 61 was thus well reproduced by the theoretical calculation, with the only differences being in the relatively intensities of these bands. The CD spectral curve for (aR)-61 was similarly calculated to give CD Cotton effects as shown in Fig. 17. In the region of the first UV band, a positive Cotton effect at λext 359.7 nm (∆ε ⫹28.6) and a negative one at λext 317.5 nm (∆ε ⫺45.0) were predicted. For the second UV band, the calculation gave a nega-

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Figure 16 Stereoscopic view of the stable conformation of biflavone, (aR)-4′,4⵮,7,7″tetra-O-methylcupressuflavone [(aR)-(⫺)-61] calculated by molecular mechanics. (Source: Ref. 13.)

tive Cotton effect that appeared as a shoulder around 290 nm and a positive one at λext 263.2 nm (∆ε ⫹21.7). These calculated CD bands were in excellent agreement with those observed in the CD spectrum of (⫺)-61, including sign, intensity, and position of the Cotton effects (compare Figs. 15 and 17): λext 362.0 nm (∆ε ⫹25.6), 326.2 (∆ε ⫺54.4), a negative shoulder around 300 nm, and 267.5 (∆ε ⫹21.3). Since the calculation was performed for the enantiomer with the aR configuration, the absolute stereochemistry of natural biflavone, (⫺)-4′,4⵮,7,7″-tetraO-methylcupressuflavone [(⫺)-61], was thus determined to be aR (or M helicity) in a nonempirical manner. C.

Circular Dichroic Power Due to the Atropisomerism of Biflavone (aR)-61

The calculated CD and UV spectral data of (aR)-61 were analyzed in detail to clarify the mechanism of CD and UV activity. As shown in Fig. 18, there are 10 π → π* transitions above 250 nm. The electric transitions 1, 4, 7, and 10 are polarized along the C2-symmetrical axis, and those of transitions 2, 3, 8, and 9 are polarized perpendicular to the axis. In the region of 400–300 nm, electric

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Figure 17 Circular dichroism and ultraviolet spectral curves of biflavone, (aR)4′,4⵮,7,7″-tetra-O-methylcupressuflavone [(aR)-(⫺)-61] calculated by the π-electron SCFCI-DV MO method. SCF-CI-DV MO, self-consistent field/configuration interaction/ dipole velocity molecular orbital. (Source: Ref. 13.)

transitions 1, 2, 3, and 4 make dominant contributions to CD and UV spectra. The analysis of the numerical calculation further clarifies that transitions 1 and 2 were, to a first approximation, exciton-coupled partners, as seen from the opposite signs of their rotational strengths. These transitions are composed of the transition a of the monomeric flavone chromophore 62 (Fig. 19). The transition moment of the monomeric transition a is almost parallel to the long axis of the p-methoxycinnamoyl moiety. For determination of the nature of the transition a, the molecular orbital distribution was investigated. The dominant excitation configuration contained in transition a is HOMO (No. 12) → LUMO (No. 13), illustrated in Fig. 20. The HOMO is mainly distrib-

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Figure 18 Rotational and dipole strengths of the transitions of biflavone, (aR)4′,4⵮,7,7″-tetra-O-methylcupressuflavone [(aR)-(⫺)-61], calculated by the π-electron SCF-CI-DV MO method. Transitions 1, 4, 7, and 10 are polarized along the C2-symmetrical axis, whereas the transition moments of 2, 3, 8, and 9 are perpendicular to the axis. SCF-CI-DV MO, self-consistent field/configuration interaction/dipole velocity molecular orbital. (Source: Ref. 13.)

uted in the p-methoxycinnamoyl subunit. Therefore, it may be concluded that the transitions 1 and 2 are composed of exciton coupling between the long axis– polarized transitions of two p-methoxycinnamoyl chromophores. However, the transitions 1 and 2 deviate from pure exciton coupling because the rotational strength of transition 1 (R ⫽ ⫹681 ⫻ 10⫺40 cgs unit) is larger than that of transition 2 (R ⫽ ⫺525 ⫻ 10⫺40 cgs unit). In the ideal case of exciton coupling, the rotational strengths of the two exciton transitions should be opposite in sign but equal in absolute value to each other. The imbalance of rotational strengths produces the positive Cotton effect at 359.7 nm.

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Figure 19 Polarization and relative intensity of dipole strength of monomeric flavone 62 calculated by the π-electron SCF-CI-DV MO method. SCF-CI-DV MO, self-consistent field/configuration interaction/dipole velocity molecular orbital. (Source: Ref. 13.)

It was found that transitions of 3 and 4 were also exciton-coupled partners. These transitions are composed of the monomeric transition b, the transition moment of which inclines as shown in Fig. 19. The major excitation configuration of transition b is MO (No. 10) → LUMO (No. 13), where MO (No. 10) is delocalized over the entire monomeric flavone subunit (Fig. 20). Thus, it is difficult to assign transition b to a specific part of the chromophore. Since the rotational strength of the transition No. 3 (R ⫽ ⫺1041 ⫻ 10⫺40 cgs unit) is larger than that of its exciton-coupled partner, transition No. 4 (R ⫽ ⫹899 ⫻ 10⫺40 cgs unit), summation of the two rotational strengths gives rise to the negative Cotton effect at 317.5 nm. It was thus clarified that the bisignate Cotton effects at 359.7 and 317.5 nm originate from the composition of two pairs of exciton couplings: one pair originates from transitions 1 and 2, and the second pair derives from transi-

Figure 20 Molecular orbital distribution of monomeric flavone 62 calculated by the π-electron SCF-CI-DV MO method. SCF-CI-DV MO, self-consistent field/configuration interaction/dipole velocity molecular orbital (Source: Ref. 13.)

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tions 3 and 4. Therefore, it became obvious that the CD exciton chirality method could not be applied in a straightforward way to these Cotton effects. For determination of the absolute stereochemistry of biflavone (⫺)-61, the theoretical calculation of the CD curve was thus needed. The analysis of the negative CD shoulder at 290 nm and the positive Cotton effect at 263.2 nm is a little simpler than the case of the Cotton effects at 359.7 and 317.5 nm. Transitions 9 and 10, which are exciton-coupled partners, are responsible for these Cotton effects (Fig. 18). Transitions 9 and 10 are composed of the monomeric flavone transition e, which is polarized along the x axis (Fig. 19). The main excitation configuration of transition e is MO (No. 10) → LUMO (No. 13) (Fig. 20). Since transition e is perpendicular to the 8-8″ linkage bond, and the two component transitions e of exciton coupling constitute a counterclockwise twist for the (aR) absolute configuration, exciton Cotton effects of negative chirality would be expected. In accordance with this expectation, the CD calculations gave a negative CD shoulder at 290 nm and a positive Cotton effect at 263.2 nm. The analysis of CD Cotton effects around 290–260 nm thus also supported the (aR) absolute configuration of biflavone (⫺)-61.

D.

Designation of the Enantiomers of the Biflavone 61 by Their Circular Dichroism Data

Traditionally, enantiomers are designated by the sign of optical rotation, [α]D. For example, the designation (aR)-(⫺)-61 means that the enantiomer of 61 with negative rotation has the (aR) absolute configuration. However, it is also well known that [α]D values are dependent on the sample concentration and solvent used. In some extreme cases, the sign of [α]D is known to be inverted by changing solvents [54], as shown in Table 1; biflavone 61 is such a case. The [α]D of the natural product 61 first measured in methanol showed a negative rotation [13]. Later it was reported by Lin et al. that the [α]D of natural 61 is positive when measured in MeOH and EtOH [15]. Therefore, the use of the [α]D value to define a specific enantiomer has caused confusion. We have proposed the use of CD data to designate enantiomers in addition to [α]D values in such cases of ambiguity [55]. For example, the designation ‘‘[CD(⫹)362.0]-(aR)-61’’ means the enantiomer of 61 showing a positive CD Cotton effect at 362.0 nm has the (aR)-absolute configuration. The importance of including the sign of an invariant optical property becomes apparent when the absolute stereochemistry of a chiral molecule is unknown. This method is very useful in the case of (⫺)-61, because the CD spectrum of 61 always exhibits a positive CD Cotton effect around 362.0 nm irrespective of the solvent, as shown in Table 1. Therefore, the natural biflavone is unambiguously defined as [CD(⫹)362.0]-(aR)-61. To maintain consistency with the previous [α]D designa-

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Table 1 Solvent Dependence of [α]D and CD Data of Biflavone 4′,4⵮,7,7″-tetra-OMethylcupressuflavone {[CD(⫹)362.0]-(aR)-(⫺)-61}a Solvent Natural sample MeOH EtOH Synthetic sample MeOH

EtOH

[α] D ⫺25.3°(c 0.3) b

λext 362.0 nm (∆ε ⫹25.6), 326.2 (⫺54.4), 267.5 (⫹21.3)b ⫺11.3°(c 0.261) (lit. ⫹1°(c 0.2)) c ⫹43.9°(c 0.259)

(lit. ⫹77°(c 0.2))c CHCl3

CD data

⫹77.0°(c 0.257)

λext 362.6 nm (∆ε ⫹22.0), 326.4 (⫺51.3), 268.6 (⫹19.4), 226.8 (⫹19.4), 216.6 (⫺4.5), 207.4 (⫹11.6) λext 363.0 nm (∆ε ⫹25.5), 327.0 (⫺55.3), 269.2 (⫹21.3), 226.4 (⫹9.1), 216.8 (⫺2.6), 208.0 (⫹11.8) (lit. λext 360 nm (∆ε ⫹28.1), 324 (⫺66.4), 267 (⫹24.8))c λext 361.0 nm (∆ε ⫹31.5), 326.2 (⫺64.0), 269.8 (⫹24.7)

c: concentration. a From Ref. 18. The sign of optical rotation was taken from the [α]D value of the natural sample in MeOH. b From Ref. 13. c From Ref. 15.

tion, we adopt both methods here; the natural product (⫺)-61 is thus defined as [CD(⫹)362.0]-(aR)-(⫺)-4′,4⵮,7,7″-tetra-O-methylcupressuflavone. Since it is also often useful to include P and M designations for these enantiomers, the (aR) enantiomer has M helicity and the (aR) isomer has P helicity. E.

Total Synthesis of [CD(ⴙ)362.0]-(aR )-(ⴚ)-4′,4ⵯ,7,7″Tetra-O-Methylcupressuflavone [(aR )-(ⴚ)-61]

1. Synthesis of (⫾)-3,3′-Diacetyl-4,4′,6,6′Tetramethoxybiphenyl-2,2′-diol [65] by the Solid-State Phenol Coupling Reaction As a chiral synthetic building block for the total synthesis of (⫺)-61, we selected 3,3′-diacetyl-4,4′,6,6′-tetramethoxybiphenyl-2,2′-diol (65) (Scheme 14). To begin the synthesis of biphenyldiol 65, phloroacetophenone (63) [56] was selectively methylated with dimethyl sulfate and K2CO3 in acetone, giving 2,4-dimethylphloroacetophenone (64) in 92% yield [57]. Unfortunately, the phenol coupling

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Scheme 14 Synthesis and resolution of biphenyl (⫾)-65.

reaction of 64 using typical oxidants (FeCl3, (t-Bu)2O2, and others) in various solvents was completely unsuccessful. We subsequently turned to the solid-state reaction conditions reported by Toda and coworkers [58]: heating a mixture of phenol 64 and FeCl3 (1.0 eq.) to 50°C for 12 h produced the desired coupled product 65 in 27% yield (Scheme 14). When phenol 64 was heated with excess FeCl3 (5.2 eq.) at 50°C for only 5.5 h, aryl chloride 66 was obtained in 33% yield, but no desired coupled product 65 was detected. We explored the reaction conditions further and found that the solid-state phenol coupling reaction in the presence of FeCl3 /silica gel [59] gave the best yield of biphenyldiol 65. When a mixture of phenol 64 and 2.8 equivalent

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of FeCl3 /silica gel (1:2) was heated at 43–45°C for 6 days, the desired coupled product (⫾)-65 was produced in 81% yield. 2. Enantioresolution of (⫾)-3,3′-Diacetyl-4,4′,6,6′Tetramethoxybiphenyl-2,2′-diol as the bis(Camphanate) Diesters 68 and X-Ray Crystallographic Analysis Resolution of (⫾)-65 using the recently developed chiral dichlorophthalic acid method [60–62] was unsuccessful, presumably because of the steric hindrance around the phenol groups of 65, which prevented acylation. However, application of the camphanate method to 65 (Scheme 14) using (1S)-(⫺)-camphanic acid chloride [63] and 4-dimethylaminopyridine in pyridine (refluxed for 3 days) yielded a mixture of diastereomeric diesters 68a and 68b (ca. 700 mg), which were separated by HPLC on silica gel (CH3CN/CHCl3 1 :19, separation factor α ⫽ 1.18, resolution factor Rs ⫽ 1.62) to give the faster-eluting (⫹)-68a (42%) followed by (⫺)-68b (46%). Recrystallization of diester 68b from hexane/EtOAc (1:1) gave large, colorless single crystals suitable for x-ray diffraction: orthorhombic, space group P212121; R ⫽ 0.0974 and Rw ⫽ 0.1061 [19]. It was found that the crystal contained EtOAc molecules as crystalline solvent, the crystal structure of which deviated from the ideal structure of EtOAc. Therefore, the position of hydrogen atoms could not be determined and the final R value remained at 9.74%. Nevertheless, the absolute stereochemistry of the biphenyl portion of diester 68b could be unambiguously assigned as (aR) by the internal reference method using the known absolute configuration of the camphanate ester moieties. It was difficult to determine the structure of the EtOAc molecules contained as the crystalline solvent, and hence the R value remained at a high level. The main reason for this difficulty was presumably the high volatility of EtOAc (bp 76.8°C), which may vaporize during the x-ray diffraction experiment. To eliminate this problem and obtain a more precise molecular structure, we attempted to replace EtOAc with another, higher-boiling ester solvent yet with a molecular size still similar to that of EtOAc such as propyl acetate (PrOAc, bp 101.6°C). Although PrOAc was not incorporated as a crystalline solvent, it was exciting to find that the large prismatic crystals obtained by recrystallization from PrOAc were composed of 68b only and were single crystals suitable for x-ray diffraction. A single crystal of 68b obtained by recrystallization from propyl acetate was subjected to x-ray analysis: colorless prism, monoclinic, space group P21; R ⫽ 0.0346 and Rw ⫽ 0.0409 [18]. The absolute stereochemistry of the biphenyl moiety of diester 68b could be unambiguously determined to be (aR) by the internal reference method using the known absolute configuration of the camphanate ester moieties (Fig. 21). The absolute configuration of diester 68a was therefore determined to be (aS).

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Figure 21 ORTEP drawing of diester (aR)-(⫺)-68b. The atoms are drawn as 50% probability ellipsoids. (Source: Refs. 18 and 19.)

3. Completion of the Synthesis of Biflavone [CD(⫹)362.0](aR)-(⫺)-61 Completion of the enantioselective total synthesis of the natural enantiomer of biflavone 61 then commenced with chiral biphenyldiol (aR)-65 obtained from diester (⫺)-68b after acidic hydrolysis (Scheme 15). When diester 68b was treated with aqueous LiOH in EtOH, deacetyl products were obtained instead of 65. Therefore, diester (⫺)-68b was heated in aqueous 6 M HCl/EtOH to 78°C for 18 h, affording optically active biphenyldiol 65 (64% yield), whose CD spectrum showed a negative CD Cotton effect at 301.2 nm. Although we had expected to obtain enantiopure biphenyldiol (aR)-65, it was found that 65 had partially racemized during the acid-catalyzed hydrolysis (79% ee). The enantiomeric excess of biphenyldiol 65 was determined by 1H NMR spectroscopy, using the chiral shift reagent europium tris[3-(trifluoromethylhydroxymethylene)-(⫹)-camphorate] [Eu(tfc)3]. Biphenyldiol (aR)-(⫺)-65 was next converted to bichalcone (aR)-(⫺)-69 (Scheme 15) by heating (70–75°C) with 4-anisaldehyde in 10% aqueous KOH/

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Scheme 15

Synthesis of biflavone [CD(⫹)362.0]-(aR)-(⫺)-61.

EtOH overnight, giving (aR)-(⫺)-69 in 65% yield. Bichalcone (aR)-(⫺)-69 was then cyclized with iodine in the presence of a catalytic amount of H2SO4 in DMSO [31] to afford 4′,4⵮,5,5″,7,7″-hexa-O-methylcupressuflavone [(aR)-(⫹)-70] in 52% yield. Finally, hexamethyl derivative (aR)-(⫹)-70 was selectively demethylated with BCl3 in CH2Cl2, giving the desired natural product (aR)-(⫺)-61 in 80% yield (78% ee). In the course of this work it was noticed that racemic biflavone 61 was barely soluble in MeOH. Therefore, the enantiomerically enriched biflavone (aR)-61 (78% ee) was recrystallized from methanol, providing enantiomerically pure biflavone (aR)-61. The enantiopurity was confirmed by 1H NMR spec-

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troscopy after conversion to the (S)-MTPA ester (aR)-71. The physical data of 100% ee biflavone 61 were mp 147–149°C (MeOH), [α]D22 ⫺11.3° (c 0.261, MeOH), and CD (EtOH) λext 363.0 nm (∆ε ⫹25.5). 4. Experimental Verification of the Absolute Stereochemistry of Biflavone [CD(⫹)362.0]-(aR)-(⫺)-61 Theoretically Determined by the Total Synthesis of Its Natural Enantiomer As discussed in the examples of (⫹)-1,8a-dihydro-3,8-dimethylazulene [(⫹)-1] and the chiral halenaquinol compounds, the theoretically predicted absolute ste-

Figure 22 Circular dichroism and ultraviolet spectra of the synthetic sample of biflavone,4′,4⵮,7,7″-tetra-O-methylcupressuflavone {[CD(⫹)362.0]-(aR)-(⫺)-61]} in EtOH. (Source: Ref. 18.)

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reochemistry of [CD(⫹)362.0]-(aR)-(⫺)-4′,4⵮,7,7″-tetra-O-methylcupressuflavone [(aR)-(⫺)-61] was confirmed by enantioselective total synthesis. The case of biflavone [CD(⫹)362.0]-(aR)-(⫺)-61 was rendered more intriguing, though, since it had once been claimed that the absolute configuration of (⫺)-61 determined by x-ray crystallography disagreed with our theoretical prediction discussed earlier [15]. However, this claim was later retracted [16,17]. The CD and UV spectra of synthetic biflavone (aR)-(⫺)-61 are illustrated in Fig. 22, and the CD curve is identical to that of the natural sample of 61 shown in Fig. 15. Therefore, the absolute stereochemistry of the natural atropisomer, [CD(⫹)362.0]-(⫺)-4′,4⵮,7,7″-tetra-O-methylcupressuflavone [(⫺)-61], was determined to be (aR). This conclusion is consistent with our previous theoretical determination of the absolute stereochemistry of biflavone [CD(⫹)362.0]-(⫺)61 by the molecular orbital calculation of CD spectra.

VI. CONCLUSIONS We have established the absolute stereostructures of natural, chiral dihydroazulenes, novel marine natural products of the halenaquinol family with a new pentacyclic skeleton, and the natural atropisomeric biflavone (⫺)-61 by theoretical calculation of CD spectra using the π-electron SCF-CI-DV MO method. The calculated CD data were in excellent agreement with the observed CD data. We have also clarified the mechanism of their CD Cotton effects by analyzing their circular dichroic power, which made the theoretical determination of the absolute stereostructures more reliable. The absolute stereostructures of natural products theoretically predicted were experimentally confirmed by the total syntheses of these natural products and pertinent model compounds. This theoretical methodology has thus become a promising tool for the determination of the absolute stereochemistry of various optically active compounds, including many natural products with twisted π-electron systems.

REFERENCES 1. A. Moscowitz, Tetrahedron, 13: 48 (1961). 2. (a) C. M. Kemp and S. F. Mason, Tetrahedron, 22: 629 (1966). (b) A. Brown, C. M. Kemp, and S. F. Mason, J. Chem. Soc. A, 751 (1971). 3. N. Harada and K. Nakanishi, Circular Dichroic Spectroscopy: Exciton Coupling in Organic Stereochemistry, University Science Books, Mill Valley, Calif., and Oxford University Press, Oxford (1983). 4. N. Harada, in Circular Dichroism, Principles and Applications (K. Nakanishi, N. Berova, and R. W. Woody, eds), VCH Publishers, New York, p. 335 (1994).

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5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33.

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N. Harada, J. Kohori, H. Uda, K. Nakanishi, and R. Takeda, J. Am. Chem. Soc., 107: 423 (1985). N. Harada, J. Kohori, H. Uda, and K. Toriumi, J. Org. Chem., 54: 1820 (1989). M. Kobayashi, N. Shimizu, I. Kitagawa, Y. Kyogoku, N. Harada, and H. Uda, Tetrahedron Lett., 26: 3833 (1985). N. Harada, H. Uda, M. Kobayashi, N. Shimizu, and I. Kitagawa, J. Am. Chem. Soc., 111: 5668 (1989). N. Harada, T. Sugioka, Y. Ando, H. Uda, and T. Kuriki, J. Am. Chem. Soc., 110: 8483 (1988). N. Harada, T. Sugioka, H. Uda, and T. Kuriki, J. Org. Chem., 55: 3158 (1990). N. Harada, T. Sugioka, H. Uda, T. Kuriki, M. Kobayashi, and I. Kitagawa, J. Org. Chem., 59: 6606 (1994). N. Harada, T. Sugioka, T. Soutome, N. Hiyoshi, H. Uda, and T. Kuriki, Tetrahedron: Asymmetry, 6: 375 (1995). N. Harada, H. Ono, H. Uda, M. Parveen, N. U.-D. Khan, B. Achari, and P. K. Dutta, J. Am. Chem. Soc., 114: 7687 (1992). M. Ilyas, J. N. Usmani, S. P. Bhatnagar, M. Ilyas, W. Rahman, and A. Pelter, Tetrahedron Lett., 5515 (1968). F.-J. Zhang, G.-Q. Lin, and Q.-C. Huang, J. Org. Chem., 60: 6427 (1995). F.-J. Zhang, G.-Q. Lin, and Q.-C. Huang, J. Org. Chem., 61: 5700 (1996). G.-Q. Lin and M. Zhong, Tetrahedron Lett., 38: 1087 (1997). H.-Y. Li, T. Nehira, M. Hagiwara, and N. Harada, J. Org. Chem., 62: 7222 (1997). N. Harada, H.-Y. Li, T. Nehira, and M. Hagiwara, Enantiomer, 2: 353 (1997). N. Harada, H. Uda, T. Nozoe, Y. Okamoto, H. Wakabayashi, and S. Ishikawa, J. Am. Chem. Soc., 109: 1661 (1987). N. Harada, J. Iwabuchi, Y. Yokota, and H. Uda, Croatica Chem. Acta, 62: 267 (1989). D. Gargiulo, F. Derguini, N. Berova, K. Nakanishi, and N. Harada, J. Am. Chem. Soc., 113: 7046 (1991). N. Berova, D. Gargiulo, F. Derguini, K. Nakanishi, and N. Harada, J. Am. Chem. Soc., 115: 4769 (1993). N. Harada, N. Hiyoshi, and K. Naemura, Recl. Trav. Chim. Pays-Bas, 114: 157 (1995). N. Harada, A. Saito, N. Koumura, H. Uda, B. de Lange, W. F. Jager, H. Wynberg, and B. L. Feringa, J. Am. Chem. Soc., 119: 7241 (1997). N. Harada, A. Saito, N. Koumura, D. C. Roe, W. F. Jager, R. W. J. Zijlstra, B. de Lange, and B. L. Feringa, J. Am. Chem. Soc., 119: 7249 (1997). N. Harada, N. Koumura, and B. L. Feringa, J. Am. Chem. Soc., 119: 7256 (1997). N. L. Allinger, QCPE, 11: 318 (1976); QCPE, Program No. 318. MOPAC 93: J. J. P. Stewart, Fujitsu Limited, Tokyo, Japan (1993). N. Harada, the computer program for CD and UV calculation, CD/NH-Sendai (1978). N. Mataga and K. Nishimoto, Z. Physik. Chem., 13: 140 (1957). R. Takeda and K. Katoh, J. Am. Chem. Soc., 105: 4056 (1983). M. Kobayashi, B. W. Son, M. Kido, Y. Kyogoku, and I. Kitagawa, Chem. Pharm. Bull., 31: 2160 (1983).

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34. B. F. Bowden, J. C. Coll, and D. M. Tapiolas, Aust. J. Chem., 36: 211 (1983). 35. R. R. Izac, W. Fenical, and J. M. Wright, Tetrahedron Lett., 25: 1325 (1984). 36. M. Kobayashi, B. W. Son, Y. Kyogoku, and I. Kitagawa, Chem. Pharm. Bull., 32: 1667 (1984). 37. N. Harada, T. Sugioka, H. Uda, and T. Kuriki, Synthesis, 53 (1990). 38. N. Harada, T. Sugioka, H. Uda, and T. Kuriki, Collect. Czech. Chem. Commun., 57: 1459 (1992). 39. (a) Z. G. Hajos and D. R. Parrish, J. Org. Chem., 39: 1615 (1974). (b) U. Eder, G. Sauer, and R. Wiechert, Angew. Chem., 83: 492 (1971); Angew. Chem. Int. Ed. Engl., 10: 496 (1971). (c) J. Gutzwiller, P. Buchschacher, and A. Furst, Synthesis, 167 (1977). (d) P. Buchschacher and A. Furst, Org. Synth., 63: 37 (1986). (e) P. Buchschacher, A. Furst, and J. Gutzwiller, Org. Synth. Coll. 7: 368 (1990). 40. D. M. Roll, P. J. Scheuer, G. K. Matsumoto, and J. Clardy, J. Am. Chem. Soc., 105: 6177 (1983). 41. M. Kobayashi, N. Shimizu, Y. Kyogoku, and I. Kitagawa, Chem. Pharm. Bull., 33: 1305 (1985). 42. H. Nakamura, J. Kobayashi, M. Kobayashi, Y. Ohizumi, and Y. Hirata, Chem. Lett., 713 (1985). 43. F. J. Schmitz and S. J. Bloor, J. Org. Chem., 53: 3922 (1988). 44. N. Harada, K. Nakanishi, and S. Tatsuoka, J. Am. Chem. Soc., 91: 5896 (1969). 45. N. Harada and K. Nakanishi, Accounts Chem. Res., 5: 257 (1972). 46. A. L. Gemal and J. L. Luche, J. Am. Chem. Soc., 103: 5454 (1981). 47. N. Harada and T. Sugioka, in Studies in Natural Products Chemistry (A. Rahman, ed.), Elsevier, Amsterdam, p. 33 (1995). 48. L. F. Fieser and M. Fieser, Reagents for Organic Synthesis, Vol. 1, John Wiley & Sons, New York, p. 921 (1967). 49. A. C. Baille and R. H. Thomson, J. Chem. Soc. (C), 2184 (1966). 50. G. Stork and P. D’Angelo, J. Am. Chem. Soc., 96: 7114 (1974). 51. W. G. Salmond, M. A. Barta, and J. L. Havens, J. Org. Chem., 43: 2057 (1978). 52. M. Oda and Y. Kanao, Chem. Lett., 37 (1981). 53. M. Parveen, N. U. Khan, B. Achari, P. K. Dutta, Abstracts of Papers, 3rd International Symposium on Flavonoids in Biology and Medicine, Singapore, Abstract p. 22 (Nov. 13–17, 1989). 54. (a) D. H. S. Horn and Y. Y. Pretorius, J. Chem. Soc., 1460 (1954). (b) A. Horeau, Tetrahedron Lett., 3121 (1969). (c) Y. Kumata, J. Furukawa, and T. Fueno, Bull. Chem. Soc. Jpn., 43: 3920 (1970). (d) D. Parker Chem. Rev., 91: 1441 (1991). 55. (a) N. Harada, J. Iwabuchi, Y. Yokota, H. Uda, Y. Okamoto, H. Yuki, and Y. Kawada, J. Chem. Soc. Perkin. Trans. 1, 1845 (1985). (b) N. Harada, Enantiomer, 1: 81 (1996). 56. K. C. Gulati, S. R. Seth, and K. Venkataraman, in Organic Syntheses (A. H. Blatt, ed), John Wiley & Sons, New York, Vol. II, p. 522 (1966). 57. K. Nakazawa, Chem. Pharm. Bull., 10: 1032 (1962). 58. F. Toda, K. Tanaka, and S. Iwata, J. Org. Chem., 54: 3007 (1989). 59. (a) K. Keinan and Y. Mazur, J. Org. Chem., 43: 1020 (1978). (b) T. C. Jempty, L. L. Miller, and Y. Mazur, J. Org. Chem., 45: 751 (1980).

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60.

(a) N. Harada, N. Hiyoshi, V. P. Vassilev, and T. Hayashi, Chirality, 9: 623 (1997). (b) N. Harada, V. P. Vassilev, and N. Hiyoshi, Enantiomer, 2: 123 (1997). 61. (a) N. Harada, T. Soutome, S. Murai, and H. Uda, Tetrahedron Asymmetry, 4: 1755 (1993). (b) N. Harada, T. Hattori, T. Suzuki, A. Okamura, H. Ono, S. Miyano, and H. Uda, Tetrahedron Asymmetry, 4: 1789 (1993). (c) N. Harada, T. Soutome, T. Nehira, H. Uda, S. Oi, A. Okamura, and S. Miyano, J. Am. Chem. Soc., 115: 7547 (1993). (d) T. Hattori, N. Harada, S. Oi, H. Abe, and S. Miyano, Tetrahedron Asymmetry, 6: 1043 (1995). (e) H. Hagiwara, T. Okamoto, N. Harada, and H. Uda, Tetrahedron, 51: 9891 (1995). 62. (a) N. Harada, T. Nehira, T. Soutome, N. Hiyoshi, and F. Kido, Enantiomer, 1: 35 (1996). (b) N. Harada, N. Koumura, and M. Robillard, Enantiomer, 2: 303 (1997). (c) H. Nemoto, J. Miyata, K. Fukumoto, H. Ehara, N. Harada, H. Uekusa, and Y. Ohashi, Enantiomer, 2: 127 (1997). 63. W. L. Meyer, A. P. Lobo, and R. N. McCarty, J. Org. Chem., 32: 1754 (1967).

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7 Recent Applications of Circular Dichroism to Carbohydrate Conformational Analysis and Direct Determination of Drug Levels Jesu´s Trujillo Va´zquez Universidad de La Laguna, Tenerife, Spain

This chapter is intended to show that circular dichroism (CD), although a wellrecognized spectroscopic technique, is still being developed, and that new applications are yet to be discovered. In spite of the existence of many useful empirical rules and of the nonempirical CD exciton chirality method for determination of absolute configuration and conformational analysis, this technique remains to be developed fully. Two recent CD applications are described: (1) First is the conformational analysis of carbohydrates, exemplified by the CD and 1 H-nuclear magnetic resonance (1 H-NMR) study of the rotational population dependence of the hydroxymethyl group in alkyl glucopyranosides on the aglycon and its absolute configuration, a clear correlation between the rotamer distributions and the stereoelectronic exo-anomeric effect being observed. As a consequence of these results the absolute configuration of secondary alcohols can be determined by either CD or 1 H NMR, as a single enantiomer is necessary for this purpose; (2) The direct determination of drug levels in pharmaceutical formulations (oral suspensions, injections, and capsules), as well as in human biological fluids (urine and serum), with specific reference to the β-lactam antibiotics, is described.

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I.

INTRODUCTION

Circular dichroism (CD) is a powerful technique that has been mainly used for the determination of absolute configuration of a great number of compounds of both natural and synthetic origin. This technique has its origins in the 1960s through important contributions by scientists such as C. Djerassi, P. Crabbe´, and G. Snatzke, who introduced many effective empirical rules for absolute configuration determinations of organic compounds [1]. Further developments came with the exciton chirality method [2], mainly through the seminal scientific contributions of N. Harada and K. Nakanishi. This method, which allows the determination of absolute configurations in a nonempirical way, offers great analytical potential and, in my opinion, establishes a second era for CD. Applications of circular dichroism have also been widely used in the conformational studies in solution of small to medium size molecules, as well as macromolecules, especially proteins [3]. The development of the CD exciton chirality method [2] has extended the utility of this technique for determining the absolute configuration of organic compounds. The high sensitivity and straightforward spectral interpretation of this method, as a consequence of the general validity of the pairwise additivity in exciton-coupled systems [4], have not been fully exploited. CD is still the technique that is least known and used by most organic chemists. The CD exciton chirality method is based on the interaction through space of the electric transition moments of two chromophores, which gives rise to an excited state split into two energy levels. Excitations to these levels lead to a CD spectrum with two Cotton effects of opposite signs, namely, to a ‘‘split’’ CD curve [2]. The chiral environment of the two chromophores determines the sign of the Cotton effects; the sign of the exciton chirality is that of the first Cotton effect, the one at longer wavelength. Furthermore, the existence of an additivity relation in multiple-chromophoric systems [4] allows an easy spectral interpretation of this type of compound, since the observed CD spectrum is the sum of CDs arising from all pairwise interactions present in the system: namely, the CD spectrum of a system composed of three interacting chromophores XYZ is the sum of the CDs arising from X/Y, X/Z, and Y/Z interactions. The general validity of the pairwise additivity in exciton-coupled systems is valid independently of whether the interacting chromophores are the same or not [4]. In the belief that CD is a spectroscopic technique that offers a series of analytical advantages over other techniques, our group led the research to apply the CD exciton chirality method to the conformational analysis of carbohydrates, namely, to study the rotational population dependence of the hydroxymethyl group in alkyl glycopyranosides on the structural nature of the aglycon and its absolute configuration.

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CD has proved to be an extremely sensitive technique to carry out conformational analysis, and many studies have been performed using this approach [1].

II. CARBOHYDRATE CONFORMATIONAL ANALYSIS Recognition of the biological importance of carbohydrates has increased dramatically over the last two decades, as a result of the fact that they possess many biological functions. In cells, carbohydrates are normally attached to proteins or lipids, forming glycoproteins or glycolipids, and it has been clearly demonstrated that the carbohydrate moiety has an enormous effect on the properties of these glycoconjugates. Glycoproteins are involved in many processes, such as cell surface recognition, immune defense, cell growth, and cell differentiation [5]. As a consequence of the extremely important biological role of glycoconjugates, and taking into account that their oligosaccharides normally define the biological function of these compounds, any contribution to a better knowledge of the conformational properties of these sugar moieties would help to understand how these macromolecules interact with other biomolecules. Most of the oligosaccharides involved in glycoconjugates are made up of hexopyranoses. The overall conformation is determined by the torsion angles (φ and ψ) around the glycosidic linkages [6]. When (1–6) linkages are involved, the corresponding torsion angle about C5–C6 bonds (ω), the preferred rotamers around the C5–C6 bond being the gauche-gauche (gg), the gauche-trans (gt), and the trans-gauche (tg) (Fig. 1) must be considered.

Figure 1 Torsion angles about the glycosidic linkage and around the C5–C6 bond (top); Newman projections of the gg, gt, and tg rotamers around the C5–C6 bond (bottom).

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The conformational analysis of carbohydrates is not a new application of CD, as may be inferred from the title. The CD exciton chirality method has already been successfully used in the conformational study of the 2-(N-acetylp-bromobenzamido) group in 2-amino-2-deoxy-galactopyranosides [7]; NMR and CD techniques are a good tandem for this type of study. An attempt will be made later to demonstrate this last point, by showing the CD and 1 H NMR study of the rotational population dependence of the hydroxymethyl group in alkyl glucopyranosides on the structural nature of the aglycon and its absolute configuration. Furthermore, the study revealed a clear correlation between the rotamer distributions and the stereoelectronic exo-anomeric effect and also revealed that as a consequence of these results the absolute configuration of secondary alcohols can be determined, as a single enantiomer is necessary for this purpose. The studies on the additivity principle in circular dichroism performed with hexopyranosides have shown that the rotational population of the hydroxymethyl group at C6 depends mainly on the configuration of the substituent at C4 (axial or equatorial) [8], and also on the bulkiness of this substituent [4a]. Recently, we have reported, on the basis of 1 H NMR and CD data [9], that the rotational population of the hydroxymethyl group in chiral and nonchiral alkyl β-d-glucopyranosides also depends on the aglycon and its absolute configuration. It was observed that the population of the gg and gt rotamers decreased and increased, respectively, as the pK a of the aglycon increased, whereas the tg population remained almost constant. Fig. 2 shows the easily interpretable CD spectra of different nonchiral alkyl 2,3-bis-O-( p-bromobenzoyl)-4,6-bis-O-( p-methoxycinnamoyl)-β-d-glucopyranosides. This bichromophoric system is composed of two homo interactions, one benzoate–benzoate (2B/3B), centered about the p-bromobenzoate λmax 245 nm, and one cinnamate–cinnamate (4C/6C), centered about the p-methoxycinnamate λ max 311 nm, as well as four hetero interactions (2B/4C, 2B/6C, 3B/4C, 3B/6C), which cover both the benzoate and cinnamate regions and are weaker in intensity than the homo interactions. The spectral differences were located in the cinnamate-cinnamate coupling region centered about the cinnamate λ max 311 nm, as can be observed in the dotted box in Fig. 2 (left) or in its expanded box in the same figure (right). The intensities of the first and second Cotton effects of the CD spectra of the glucosylated nonchiral alcohols 1a–e gradually decreased from methyl (A C value 36.6) [10], to ethyl (35.4), to iso-propyl (31.3), to cyclohexyl (31.3), and to tert-butyl (23.6) glucopyranoside derivatives. These differences in the CD curves clearly indicated that only the interactions involving the chromophore at the 6 position are significantly affected, mainly the homo cinnamate– cinnamate (4C/6C) pairwise interaction, and, therefore, that the rotamer distributions have changed.

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Figure 2 Circular dichroism spectra (CH3CN) of model alkyl 2,3-bis-O-( p-bromobenzoyl)-4,6-bis-O-( p-methoxycinnamoyl)-β-d-glucopyranosides: 400- to 200-nm region (left), and 350- to 264-nm region (right).

Moreover, these spectral differences are only consistent with a gradual decrease and an increase in the population of the gg and gt rotamers, respectively, as can be deduced by analyzing the pairwise interactions involved in these compounds. Fig. 3 shows for the glucopyranosyl system those pairwise interactions involving the chromophore at the 6 position, the 2/6, the 3/6, and the 4/6 interaction, for the three rotamers. As can be observed, those interactions involving the gg rotamer have a net positive contribution; those involving the gt rotamer have a net negative contribution; and those interactions involving the tg rotamer have a nil contribution. Therefore, the general decreases in the CD Cotton effects can be explained by a decrease in the population of the gg rotamer (net positive contribution) and an increase in the population of the gt rotamer (net negative contribution). The other three existing pairwise interactions, the positive 2/3, the negative 3/4, and the nil 2/4, having constant intensities and signs, do not affect this CD interpretation. Furthermore, the striking increase in the first positive and second negative Cotton effects observed on these spectra by lowering the temperature (MeOH, ⫺80°C) can only be explained by an increase in the rotational population of the energetically favored gg rotamer. Rotamer distribution is normally determined by 1 H NMR analysis of the coupling constants of the prochiral protons at C6 [11], which are assigned on

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Figure 3 Pairwise interactions involving the chromophore at the 6 position in each of the three stable rotamers. (Source : Ref. 9.)

the basis of their chemical shifts. That is, the coupling constant values of the usually well-differentiated doublet of doublets for H6R and H6S are substituted in empirical equations to calculate the rotamer distribution [12]. With the exception of the cyclohexyl derivative 1d, a doublet, rather than the usual clearly differentiated doublet of doublets for each proton at C6, was obtained for the bichromophoric compounds. Therefore, their rotamer distributions could not be calculated by means of 1 H NMR coupling constants. However, the coupling constant J H5,H6, which gradually increased from methyl (3.9 Hz), to ethyl (4.1 Hz), to iso-propyl (4.4 Hz), and to tert-butyl (4.6 Hz) glucopyranoside derivatives (1a–c,e), corroborated the increase in the gt population. Since the results show that the gt population increases as the pK a of the aglycon increases, and it is accepted that the stereoelectronic exo-anomeric effect [13] increases with increasing ease for charge delocalization from the aglycon to the anomeric carbon [14], this stereoelectronic effect must be responsible for the rotational population dependence of the hydroxymethyl group on the aglycon, because for these low-molecular-size alcohols the existence of nonbonded interactions with the chromophore at C6 cannot be expected. This correlation was confirmed by the CD and NMR data comparison of

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the methyl and the acetyl tetra-O-p-bromobenzoyl-β-d-glucopyranosides, compounds 2a and 2b, respectively (Scheme 1). The spectrosopic data (Table 1) showed a higher gg population for the acetyl glucopyranoside derivative than that of the methyl glucopyranoside 2a, in agreement with the nil exo-anomeric effect reported for the acetyl glucopyranoside 2b [15]. Rotational studies performed with nonchiral alkyl α-d-glucopyranosides [16] and β-d- and α-d-galactopyranosides [17] have revealed that the gt population also increases as the pK a of the aglycon increases; this behavior seems to be general in hexopyranosides. Spectroscopic analysis of chiral alkyl 2,3,4,6-tetrakis-O-( p-bromobenzoyl)-β-d-glucopyranosides 2c–h (Scheme 1) showed another correlation, that is, between the rotational population of the hydroxymethyl group and the absolute configuration of the aglycon. Higher CD A values and smaller J H5,H6R coupling constants were obtained for the R-alkyl glucopyranosides than for their S-alkyl glucopyranoside counterparts. CD spectra of these glucosylated secondary alcohols showed a general decrease in the positive first and negative second Cotton effects with respect to those of the methyl derivative 2a (Table 1), as occurred with the glucosylated

Scheme 1 Structures of model alkyl β-d-glucopyranosides.

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Table 1 Circular Dichroism Data (CH 3CN), J H5,H6 Coupling Constants (CDCl 3, 400 MHz), and Calculated Rotameric Populations (Percentage) around the C5-C6 Bond for Model 2,3,4,6-Tetrakis-O-(p-bromobenzoyl)-β-d-glucopyranosides 2a–h Compd 2a 2b 2c 2d 2e 2f 2g 2h

Aglycon

Cl′

∆ε at 250/232 nm

A Value

J H5,H6R

J H5,H6S

P gg

P gt

P tg

Methyl Acetyl (⫺)-2-Octyl (⫹)-2-Octyl (⫺)-Menthyl (⫹)-Menthyl (⫺)-Bornyl (⫹)-Bornyl

— — R S R S R S

23.7/–6.2 26.2/–5.1 18.6/–5.0 17.8/–4.6 17.7/–4.1 9.5/–0.5 17.2/–4.8 16.8/–3.3

29.9 31.3 23.6 22.4 21.8 10.0 22.0 20.1

4.7 4.7 5.1 5.3 5.2 6.1 5.2 5.4

3.4 3.0 3.6 3.6 3.6 3.4 3.7 3.6

57 59 53 50 51 45 51 50

26 28 29 32 31 40 30 32

17 13 18 18 18 15 19 18

nonchiral secondary alcohols 1c–d with respect to 1a. Moreover, CD spectra of those stereoisomers having an R absolute configuration at the aglyconic carbon (C1′) (2c, 2e, and 2g) showed higher amplitudes (A values) than those obtained from stereoisomers with the opposite absolute configuration (2d, 2f, and 2h). Fig. 4 shows the CD spectra of methyl, (⫹)- and (⫺)-2-octyl, and (⫹)- and (⫺)menthyl 2,3,4,6-tetrakis-O-( p-bromobenzoyl)-β-d-glucopyranosides (2a,c–f, respectively); their spectral differences are analyzed on the basis of their pairwise interactions (Fig. 3). In agreement with CD data, analysis of the 1 H NMR coupling constants

Figure 4 Circular dichroism spectrum (CH 3CN) of methyl, (⫹)- and (⫺)-2-octyl (left), and (⫹)- and (⫺)-menthyl (right) 2,3,4,6-tetrakis-O-( p-bromobenzoyl)-β-d-glucopyranosides. (Source : Ref. 9.)

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J H5,H6 of these chiral alkyl glucopyranosides showed smaller J H5,H6R coupling constants for those stereoisomers having an R absolute configuration at the aglyconic carbon than for their glucopyranoside counterparts (Table 1). Circular dichroism and 1 H NMR data comparison indicated the existence of an excellent correlation between the magnitudes of the rotamer populations obtained by 1 H NMR and the CD A values. Furthermore, analysis of these spectroscopic data established that those stereoisomers having an R absolute configuration at the aglyconic carbon possess higher gg and smaller gt populations than their stereoisomers with the opposite absolute configuration. The differences between each pair of stereoisomers may be due to steric interactions, since the pK a of the enantiomers bonded is the same. There are two possible explanations: (1) The first is the existence of nonbonded interactions between the aglycon and the chromophore at C6. In agreement with the rotational dependence of the torsion angle Ψ on the structure of the aglycon [6] higher nonbonded interactions can be expected between aglycons having the bulkiest substituent syn to O5 (2c,e, and 2g) and the chromophore at C6 in the gt conformation than for those stereoisomers having the opposite configuration at the aglyconic carbon (2d, 2f, and 2h) (Fig. 5). (2) There exists a more stabilizing exoanomeric effect for the diastereoisomers with an S absolute configuration at the aglyconic carbon, those with a higher gt population. Since the study performed with the glucosylated nonchiral alcohols revealed that a higher gt population correlates with a higher value of the exo-anomeric effect, these diastereoisomers may have a more favored disposition of the orbitals involved in this stereoelectronic effect [13] than their corresponding diastereoisomers. Chiral alkyl galactopyranosides [17] have shown behavior similar to that of alkyl glucopyranosides.

Figure 5 Schematic representation of the rotational dependence of the torsion angle ψ on the structure of the aglycone, and corresponding Newman projections.

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We can conclude from these experiments that the different intensities of the CD spectra and values of the 1 H NMR coupling constants J H5,H6R and J H5,H6S of alkyl glycopyranosides are mainly due to different values of the stereoelectronic exo-anomeric effect. As a consequence of these findings, two methods for the absolute configuration determination of secondary alcohols were proposed, one by applying CD [9,18] and the other by means of 1 H NMR [18]. As can be observed in Tables 1 and 2, a simple comparison of the intensities of the CD curves of the corresponding diastereoisomeric alkyl 2,3,4,6-tetrakisO-benzoyl- or 2,3,4,6-tetrakis-O-( p-bromobenzoyl)-β-d-glucopyranosides shows that compounds of R absolute configuration at the aglyconic C-atom exhibit a higher A value, or rather, a higher intensity of the Cotton effect at longer wavelength, than their corresponding diastereomers with S configuration at the same stereogenic C1′. The ∆ε values shown for both glucopyranosides of testosterone were obtained after subtraction of the overlapping π → π* transition of the enone system at 234 nm (⫹7.7).

Table 2 Circular Dichroism Data (CH 3CN) and 1 H Nuclear Magnetic Resonance Coupling Constants JH5,H6 (CDCl 3, 400 MHz) for 2,3,4,6-Tetrakis-O-Benzoyl-βGlucopyranosides of Secondary Alcohols Entry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18

Alcohol (⫺)-2-Octanol (⫹)-2-Octanol (⫺)-Menthol (⫹)-Menthol (⫺)-Neomenthol (⫹)-Neomenthol (⫺)-Borneol (⫹)-Borneol Cholesterol Cholesterol Cholestanol Cholestanol Testosterone Testosterone Dimethyl d-malate Dimethyl l-malate (⫺)-Methyl 3-hydroxy butyrate (⫹)-Methyl 3-hydroxy butyrate

Source: Ref. 9.

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Glucose series

Cl′

∆ε 233/220 nm

A Value

J H5,H6R

J H5,H6S

D D D D D D D D D L D L D L D D D

R S R S R S R S S S S S S S R S R

7.3/–0.9 6.3/–0.6 5.8 4.2 6.6 6.2 8.7 7.5/–0.3 7.4 ⫺7.8 6.2/–0.9 ⫺7.2 8.0 ⫺8.1 8.0 5.8 10.8

8.2 6.9 5.8 4.2 6.6 6.2 8.7 7.8 7.4 ⫺7.8 7.1 ⫺7.2 8.0 ⫺8.1 8.0 5.8 10.8

5.6 5.8 5.7 6.8 5.7 5.9 5.7 5.8 5.9 6.1 6.0 6.1 5.6 4.7 5.6 5.0 5.2

3.3 3.2 3.3 3.1 3.3 3.3 3.6 3.2 3.3 3.1 3.4 3.2 3.1 — 3.5 3.2 3.3

D

S

6.2/–0.8

7.0

5.6

3.2

In order to apply this method it is not necessary to have both enantiomers, since for CD spectra comparison the CD spectrum of the unavailable diasteromer can be obtained by glycoside formation of the available enantiomer with the easily prepared 2,3,4,6-tetrakis-O-benzoyl- or 2,3,4,6-tetrakis-O-( p-bromobenzoyl)-α-l-glucopyranosyl bromide and multiplying its CD spectrum by ⫺1 (Fig. 6). The 1 H NMR data obtained from the glucosylated secondary alcohols also allow the absolute configuration of these secondary alcohols to be determined in two different ways: (1) in agreement with a higher gt population for stereoisomers with an S absolute configuration at the aglyconic carbon, a higher J H5,H6R coupling constant (CDCl 3) has been observed for these stereoisomers than for their corresponding diastereomers (Tables 1 and 2). However, the 2,3,4,6-tetrakis-O-benzoylglucopyranosides of cholesterol, cholestanol, and dimethyl malate (Table 2) did not exhibit that behavior, probably because of the absence of a substituent at the β-positions in the first two cases, and of the appearance of new anisotropic effects from the carbonyl groups in the last case. (2) The dramatic shifts in the aglycon 1 H NMR peaks, upon tetra-O-benzoyl-β-glycosylation of secondary alcohols, as a consequence of anisotropic effects and glycosylation-induced 1 H-

Figure 6 Calculation of the circular dichroism curve of an ‘‘available’’ S-alkyl 2,3,4,6tetrakis-O-( p-bromobenzoyl)-β-d-glucopyranoside (dotted line) by multiplying by ⫺1 the CD curve (dashed line) of its R-alkyl β-l-glucopyranoside derivative resulting from bonding the available enantiomer (R) to the 2,3,4,6-tetrakis-O-( p-bromobenzoyl)-α-l-glucopyranosyl bromide. (Source : Ref. 9.)

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Figure 7 Configurational correlation models for secondary alcoholic (a) β-d- and (b) β-l-glucopyranosides. (Source : Ref. 18.)

NMR shifts (Fig. 7), constituted the basis of a new method to determine the absolute configuration of secondary alcohols [18]. The differences between the proton chemical shifts of the d-glucosylated derivative and the free alcohol (∆δ ⫽ δ D ⫺ δ ROH ) or, more significantly, between their chemical shifts in the d- and lglucosylated derivatives (∆δ ⫽ δD ⫺ δL) are characteristic of the absolute configuration of the secondary chiral alcohol. Moreover, this method involves the use of one enantiomer and generally a single derivatization is sufficient. Fig. 8 shows for (⫺)-menthol (in hertz, at 500 MHz, 25°C, CDCl 3) the chemical shift differences (δD ⫺ δL) obtained by applying our method, on the basis of the tetra-O-benzoylglucosylation, as well as those (δ S ⫺ δ R) acquired by the advanced Mosher’s method [19]. The larger values obtained by the former can make it a suitable alternative method to determine the absolute configuration of secondary alcohols. III. DIRECT DETERMINATION OF DRUGS In order to be CD-active, a compound must fulfill two requisites: it must be optically active and absorb electromagnetic light in the region under study. This

Figure 8 Comparison of the δ S ⫺ δ R and the δ D ⫺ δ L values of the MTPA [19d] (left) and the tetrakis-O-benzoyl-glucopyranoside (right) derivatives of (⫺)-menthol, respectively. MTPA, 1-methoxy-1-phenyl-1-trifluoromethylacetic acid. (Source: Ref. 18.)

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is due to the fact that two physical requirements (ellipticity and absorbance) are measured simultaneously. Although this limits the applicability of CD, it also confers on this technique a high degree of analytical selectivity, allowing an optically active absorbing drug in a mixture to be analyzed directly. A.

Direct Determination of Drugs in Pharmaceutical Formulations

A clear example of the advantage of the high selectivity of CD can be found in the pharmaceutical field [20,21], in which an optically active absorbing drug can be directly analyzed from its pharmaceutical preparations, with additives not interfering with the CD measurement [22], as well as in biological fluids (urine and serum). It is interesting to note that although the chiral drugs greatly outnumber achiral ones, CD has only been used infrequently in pharmaceutical development [20,21]. Thus, we have reported a method for the discrimination and accurate and precise determination of cephalosporins in pure form, as well as for direct determination of cephalosporins in commercial oral suspensions, injections, and capsules [23]. The study, performed with 15 commercial cephalosporins in common clinical use (Scheme 2), first revealed sufficient CD spectral dissimilarities to discriminate among the cephalosporin homologues and classify these antibiotics, on the basis of the wavelengths of their Cotton effects, in five spectroscopic groups (Fig. 9 and Table 3). Furthermore, distinguishing between the β-lactam antibiotics, penicillins [22], cephalosporins, and cephamycins (R 3 ⫽ OCH 3) on the basis of their CD spectral data was found to be straightforward [23]. The observed CD spectral dissimilarities come from red shifts of the two transitions (at 260 and at 230 nm) of the 3-cephem chromophore, the basic skeleton of cephalosporin antibiotics [24], produced by the substituents directly attached to it (R 2 and R 3), and/or the overlapping of new transitions originating from the chromophores present in R 1 and R 2. The validity of the method for the determination of cephalosporins by CD was confirmed by analysis of the variance (ANOVA). This is a statistical analysis technique that allows us to overcome the ambiguity that the estimation of significant differences represents when more than one comparison is performed. Table 4 shows the slopes and the intercepts of the equations of the regression lines [25] calculated for one member of each spectroscopic group, as well as the excellent correlation coefficients and values of the root mean square of the ANOVA error term, S θc, for these antibiotics. The acceptable limit of quantitation was found to be 5 µg/mL, the overall accuracies for cefadroxil, cefapirin, cefamandole, cefoxitin, and ceftriaxone were 99.9, 100.1, 101.1, 100.1, and 100.1, with precisions of 1.10, 1.46, 1.69, 1.19, and 1.66, respectively. The method was then applied to the direct determination of cephalosporins in pharmaceutical formulations. The samples were dissolved and CD measured

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Scheme 2 Structures of the cephalosporins analyzed.

directly, without performing any derivatization, filtration, or chromatographic separation steps. The CD spectral patterns of the analyzed drugs were identical to those of the standard solutions, showing the noninterference of additives, since these are non-CD-active. As can be observed in Table 5, the mean amount estimated in each case, by using either the CD or the HPLC [26] technique, is in good agreement with the amount claimed on the label. Therefore, CD can be of

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Figure 9 Circular dichroism spectra for one member of each group: cephradine (first group), cephalotin (second group), cephazolin (third group), cefoxitin (fourth group), and ceftriaxone (fifth group). (Source: Ref. 23.)

Table 3 Circular Dichroism Data of the Commercial Cephalosporins Analyzed Group First

Second

Third

Fourth Fifth

Name Cefadroxil Cefoperazone Cephalexin Cephradine Cefotaxime Cephalotin Cefuroxime Cefaclor Cefapirin Cefsulodin Cefamandole Cephazolin Cefoxitin Moxalactam Ceftriaxone

Source: Ref. 23.

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λ ext /∆ε

306/–0.6 296/–1.1

296/–1.6

λ ext /∆ε

λ ext /∆ε

255/9.2 255/3.1 255/12.1 255/10.6 258/10.5 258/12.4 258/11.4 259/11.0 259/14.6 263/6.3 263/6.6 263/7.4 262/17.2 265/8.0 284/–5.2

211/–16.3 223/–17.4 223/–18.3 221/–14.4 228/–18.3 228/–17.4 227/–16.4 228/–15.8 229/–17.5 231/–16.9 230/–16.9 228/–16.6 238/–18.7 239/–28.8 236/–17.4

λ ext /∆ε 207/–17.7 202/–15.2

215/3.5

∆ε ⫽ 0 235, 202 238 237 237 289, 242, 206 242 296, 242, 207 242 242, 208 247, 210 248 285, 247 250, 222, 208 255, 215 206

Table 4 Parameters of Calibration Graphs for One Member of Each Circular Dichroism Spectroscopic Group in Distilled Water Compound

Range (µg/mL)

n

CD λ ext (nm)

Cefadroxil Cefapirin Cefamandole Cefoxitin Ceftriaxone

1.0–80.0 1.0–60.0 5.0–80.0 1.0–80.0 5.0–40.0

30 25 24 28 24

255 259 263 262 236

Source: Ref. 23.

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Slope (m) 0.8575 0.9960 0.4228 1.3209 ⫺0.9968

⫾ ⫾ ⫾ ⫾ ⫾

0.0016 0.0047 0.020 0.0051 0.0065

Intercept (z) ⫺0.0656 0.2014 ⫺0.0521 ⫺0.1915 0.3867

⫾ ⫾ ⫾ ⫾ ⫾

0.060 0.1583 0.0912 0.2281 0.1642

Correlation coefficient (r)

Sθc

0.9999 0.9998 0.9997 0.9998 0.9995

0.265 0.537 0.277 0.806 0.410

Table 5 Determination of Cephalosporins in Pharmaceutical Formulations

Dosage form Oral suspension (n ⫽ 3) Capsules (n ⫽ 4) Injection (n ⫽ 5)

Amount of labeled claim (g/unit) 3.0 35.0 0.5 1.0

(Cefadroxil) (Saccharose) (Cefadroxil) (Cefoxitin)

Mean amount estimated ⫾ SD (%) a CD

HPLC

104.5 ⫾ 2.13

99.2 ⫾ 2.45

104.8 ⫾ 1.48 103.1 ⫾ 1.24

102.8 ⫾ 3.26 97.93 ⫾ 0.74

a

CD, circular dichroism; HPLC, high-performance liquid chromatography. Source: Ref. 23.

great utility for the direct determination of drugs in pharmaceutical preparations; its principal advantages, besides those mentioned, are quickness and simplicity. In addition, the analysis is inexpensive and can be performed in a few minutes; all these properties are very practical for quality control laboratories. Moreover, the spectroscopic classification facilitates the discrimination of cephalosporin homologues and allows their correct identification in most cases. We wish to emphasize that although we have used the β-lactam antibiotics as model compounds to prove that CD is a suitable technique for the direct determination of these drugs in pharmaceutical formulations, this result can be extended to any optically active absorbing drug. B.

Direct Determination of Drugs in Human Biological Fluids (Urine and Serum)

Since any research that, directly or indirectly, contributes to health improvement is of great interest, we decided to perform an exhaustive study of the applicability of CD to the quantitative determination of optically active absorbing drugs in biological fluids [27]. The high selectivity of this technique should allow CDactive drugs to be determined among other non-CD active ones, since practitioners usually prescribe several drugs simultaneously. 1. Human Urine At the time the study that follows [27] was performed, only two reports had been published for the determination of drugs in human biological fluids by CD, one describing the determination of tetracycline in urine [28] and the other the determination of tri- and tetra-(hydroxyethyl)-rutosides in urine and serum [29]. In collaboration with the urological department of the Hospital Universita-

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rio de Canarias, over 200 urine samples from 10 healthy adult subjects and 51 hospitalized patients were analyzed by CD. The standard dilution used in this work for CD analysis of drugs in urine was prepared by diluting 20 µL of urine in a 5-mL calibrated flask with distilled water: a 250-fold dilution. The CD spectra of the urine samples belonging to healthy volunteers and to some patients under the administration of multiple nonoptically active absorbing drugs showed no Cotton effects in the 400- to 200-nm range. This dilution offers the best chance of eliminating any possibility of interference from the urine metabolites. Therefore, any Cotton effect that appears in this region under the specified conditions must be due to the presence of urinary proteins and/or CD-active drugs in the urine. The optically active absorbing drugs vitamin C, cephalexin, cefoxitin, ampicillin, and amoxicillin were detected in the CD spectra of the urine samples of the patients under treatment with one of these drugs. Urinary proteins were also detected in the CD spectra of the urine samples of 31 of 51 patients, whose routine urinalysis had already revealed significant amounts of proteins (15–300 mg/dL). In many cases the patients under treatment with one β-lactam antibiotic were under multiple-drug administration. None of the drugs shown in Table 6 interfered with the CD analyses of the β-lactam antibiotics. These drugs are nonabsorbing and/or nonoptically active or possess an ellipticity too low to be detected in our standard dilution. Interference was only observed in those cases in which the patients were also under treatment with vitamin C, an optically active absorbing compound. Therefore, vitamin C must be avoided when another CDactive drug is analyzed. The pattern of the CD spectra of the urine samples depends on the presence

Table 6 Noncircular Dichroism–Active Drugs in Urine Antimicrobial agents

Hypnotics and sedatives Antidepressants Analgesics–antipyretics Nonsteroidal antiinflammatory agents Methylxanthine bronchodilators Coronary vasodilators Antihypertensive agents Antiemetics Antihistamines Source : Ref. 27.

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Tobramycin, gentamycin, netilmycin, norfloxacin, pipemidic acid, ciprofloxacin, cotrimoxazole Alprazolam, triazolam Maprotiline hydrochloride Acetaminophen, amidopyrine Acetylsalicylic acid, diclofenac Theophylline Nitroglycerin, isosorbide dinitrate, nifedipine Prazosin Metoclopramide Astemizole, ranitidine

of proteins and/or a CD-active drug; thus, (1) the CD spectra of those proteinfree samples containing a CD-active drug were superimposable with those of the standard dilutions [30] after matching the concentrations (Fig. 10); (2) the CD spectra of those samples containing only urinary proteins exhibited clear proteintype CD spectra [31], namely, a broad negative Cotton effect with two extrema (around 220 and 209 nm) superimposable in most cases with that of human albumin [30], the main human urinary protein (Fig. 11); (3) the CD spectra of those samples containing, in addition to proteins, a CD-active drug exhibited ‘‘complex’’ CD spectra identical to those of the standard dilution in the range 400– 250 nm, although with modified patterns below 250 nm, as a consequence of the overlapping of Cotton effects of the urinary proteins on those of the CD-active drugs (Figs. 12 and 13). The direct determination of β-lactam antibiotics in human urine by CD was validated by using as model drugs the antibiotics cephalexin, cefoxitin, and ampicillin. The linearity of the present method was confirmed by ANOVA of the linear regression line equations of these drugs (Table 7) [25]. In addition, the validation study performed confirmed the accuracy and precision of the method; the overall accuracy for ampicillin, cefoxitin, and cephalexin was 100.5%, 99.6%, and 100.1%, with precision of 1.69, 1.18, and 1.68, respectively. In the case of urinary proteins it was assumed that the ellipticity of each

Figure 10 Circular dichroism (CD) spectrum of standard CD-active drugs: cefoxitin (6.98 µg/mL), cephalexin (9.32 µg/mL), amoxicillin (9.91 µg/mL), and ampicillin (8.50 µg/mL). (Source : Ref. 30.)

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Figure 11 Circular dichroism spectrum of standard human albumin (10.48 µg/mL). (Source : Ref. 27.)

Figure 12 Circular dichroism spectra of the urine of three patients under cefoxitin therapy (250 times dilutions): without proteins, solid line (23.25 µg/mL of cefoxitin); with proteins, dashed line (15.29 µg/mL of cefoxitin, and 11.19 µg/mL of proteins); dotted line, (7.48 µg/mL of cefoxitin, and 17.16 µg/mL of proteins). (Source: Ref. 27.)

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Table 7

Parameters of Calibration Graphs for Model Drugs and Human Albumin in Urine

Compound

Range (µg/mL)

n

CD λ ext (nm)

Ampicillin Cefoxitin Cephalexin Albumin

1.0–40.0 1.0–80.0 1.0–50.0 1.0–50.0

25 24 25 25

234 262 255 220

Source: Ref. 27.

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Slope (m) 0.9582 1.1500 1.1766 ⫺1.5407

⫾ ⫾ ⫾ ⫾

0.0043 0.0036 0.0067 0.0046

Intercept (z) ⫺0.1769 ⫺0.2219 ⫺0.0717 ⫺0.3232

⫾ ⫾ ⫾ ⫾

0.0888 0.1671 0.2000 0.1229

Correlation coefficient (r)

Sθc

0.9998 0.9999 0.9996 0.9999

0.288 0.534 0.634 0.391

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C

Figure 13 Complex circular dichroism (CD) spectrum of the urine of a patient under ampicillin therapy (250 times dilutions) (solid line), its estimated protein CD curve (dotted line) (2.54 µg/mL), and its estimated ampicillin CD spectrum (dashed line) (14.55 µg/ mL). (Source: Ref. 27.)

urinary protein was identical with that of the main human urinary protein, human albumin. Similarly to the β-lactam antibiotics, the analysis of the variance, (ANOVA) of the linear regression of the calibration line carried out with human albumin at 220 and 209 nm confirmed the linearity of the method (Table 7); the statistical analyses showed precise and accurate values for both wavelengths, although slightly better for the one measured at 220 nm. The direct quantitative determination of a CD-active drug in protein-free urine samples from type-1 CD spectra or the determination of the total urinary proteins in urine samples lacking CD-active drugs from type-2 CD spectra is achieved in a straightforward manner by measuring in each case the ellipticity angle (mdeg) at the selected wavelength and using the corresponding regression line equation (Table 7). The procedure to determine a CD-active drug from a ‘‘complex’’ type-3 CD spectrum depends on the wavelength of the first Cotton effect of the drug to be analyzed: (1) If the drug exhibits a Cotton effect above 250 nm, its determination can be performed by direct measurement of the ellipticity at the wavelength of this Cotton effect, regardless of the presence of proteins (Fig. 12). This can also be determined by CD spectral subtraction of the CD-active drug contribution to the ‘‘complex’’ CD spectrum; (2) the simultaneous determination of a

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CD-active drug, having only Cotton effects below 250 nm, and the total urinary proteins required to perform a CD spectral subtraction, due to the overlapping of their Cotton effects. In this case it is necessary to carry out a regression fit using the shapes of the drug and the albumin spectra (Fig. 13). The individual regression line equations are then used to determine their concentration. CD analyses carried out in the presence of proteins did not exhibit, under our standard dilution, overlapping of the very weak signals, from aromatic residues of proteins [3], and/or extrinsic Cotton effects from drug–protein binding [32]. Therefore, the quantitation of CD-active drugs is not interfered with by the possible existence of the Cotton effects mentioned. Fig. 14 shows how the wavelength of the first Cotton effect of ‘‘complex’’ CD spectra of in-house mixtures of cefoxitin (80, 60, 40, 20, 5, and 1 µg/mL) and a constant concentration of albumin (48 µg/mL) remained unaltered at 262 nm, the wavelength of the first Cotton effect of cefoxitin. Furthermore, CD spectra shown in Fig. 15 exhibit a constant ellipticity at this wavelength independent of the concentration of urinary proteins. These ‘‘complex’’ CD spectra become clear protein-type spectra (Fig. 11) by removal of the corresponding CD drug contribution, so the total urinary proteins (or human albumin for in-house mixtures) can be determined by measuring at 220 nm the resulting ellipticity angle (Table 8).

Figure 14 Complex circular dichroism spectra, from top to bottom, of in-house mixtures of albumin (48 µg/mL) and cefoxitin (80, 60, 40, 20, 5, and 1 µg/mL), respectively [30]. (Source: Ref. 27.)

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Figure 15 Circular dichroism spectra of three in-house samples with a constant concentration of cefoxitin (20 µg/mL): without albumin (solid line); with albumin: 24 µg/mL (dashed line) and 48 µg/mL (dotted line) [30]. (Source: Ref. 27.)

The present method for the direct determination of a CD-active drug in human urine was also confirmed by performing a comparative analysis by means of CD and HPLC. As can be observed in Table 9, the concordance between the values obtained by these techniques was excellent. The present method can be very useful to perform pharmacokinetic studies

Table 8 Determinations of in-House Lactam–Albumin Mixtures Determined concentration (µg/mL)

Entry

Compound

Drug

Albumin

Drug

Albumin

1 2 3 4 5 6

Cefoxitin

20.00 20.00 20.00 20.00 20.00 20.00

24.00 48.00 24.00 48.00 24.00 48.00

19.98 20.05 19.78 19.87 20.04 20.51

24.10 48.19 24.14 48.51 24.63 48.93

Cephalexin Ampicillin

Source : Ref. 27.

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Table 9 Comparative Analysis by HPLC and CD Spectroscopy a Determined concentration (mg/mL) Entry

Compound

CD

HPLC

1 2 3 4 5 6 7 8 9

Ampicillin

14.58 12.23 17.12 2.22 19.18 23.15 11.86 17.20 17.78

13.75 12.61 16.62 2.30 20.48 23.32 11.85 16.11 17.88

Cefoxitin

Cephalexin

a

HPLC, high-performance liquid chromatography; CD, circular dichroism. Source: Ref. 27.

of optically active absorbing drugs. As can be observed in Fig. 16, a clear gradual decrease in the intensity of the Cotton effects of the urine samples along a 6-h period was obtained from a patient receiving intravenously (i.v.) the first dose of 1 g of cefoxitin. The corresponding CD hourly determinations were in excellent agreement with those obtained from HPLC, showing the applicability of this method to monitor drugs. The total amounts of cefoxitin recovered from the first up to the sixth hour were 349.4, 227.1, 113.4, 74.1, 59.7, and 39.3 mg, respectively. 2. Human Serum The most complex and important of the biological fluids is blood, and thus once the study in urine was completed, the possibility of applying CD to establish a method to determine CD-active drugs in this biological fluid was analyzed [33]. Since the concentration of proteins in human serum or plasma is very high, it was necessary to dilute the serum sample to 1 : 1000, in order to have a suitable voltage in the CD photomultiplier (below 460 V). Consequently, the direct determination of a CD-active drug is not possible from these very dilute solutions; deproteinization of the serum sample is necessary before CD analysis. The precipitation of the serum/plasma proteins by means of acetonitrile proved to be an excellent way to perform CD analysis of drugs. The method

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Figure 16 Circular dichroism spectra recorded along a 6-h period from the urine samples of a patient under cefoxitin therapy (250 times dilution). From the first up to the sixth hour the following concentrations of cefoxitin were determined: 53.54, 37.68, 25.11, 21.10, 14.88, and 7.44 µg/mL, respectively. (Source: Ref. 27.)

consists of: (1) adding 3 mL of CH 3CN to 1 mL of serum; (2) centrifuging the mixed solution at 5000 rpm for 15 min; (3) measuring the CD spectrum of the protein-free supernatant (about 3.5 mL). Following this method, the CD spectrum of several drug-free serum samples showed no Cotton effect in the 400- to 200-nm range, clearly showing the total removal of the proteins, although important absorptions below 250 nm remained. Nonchiral absorbing compounds, such as creatinine and uric acid, must be responsible for these absoptions. Therefore, only drugs having Cotton effects at wavelengths above 250 nm can be measured by the procedure. On the other hand, the CD spectra of protein-free supernatants containing different cephalosporins (Fig. 17), all having Cotton effects above 250 nm, showed Cotton effects in agreement (⫾2 nm) with those of the standard solutions in the 400- to 250-nm range. Therefore, the determination of CD-active drugs in human serum/plasma samples could be easily performed by measuring the ellipticity angle of their first Cotton effects and replacing their values in the corresponding regression line equations. Currently we are performing successfully the validation and recovery studies of the present method [33]. It can be concluded that CD is a valid alternative spectroscopic technique to perform, in a simple, precise, and accurate manner, the quantitative determina-

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Figure 17 Circular dichroism spectra of the resulting protein-free serum samples containing 19.1 µg/mL of cephalotin, 16.7 µg/mL of cephalexin, and 19.0 µg/mL of ceftriaxone (10-mm cylindrical quartz cell).

tion of drugs in biological fluids. Its high degree of selectivity, together with the fact that it is quick and economical, could make CD a common technique in clinical laboratories in the near future.

ACKNOWLEDGMENTS I wish to thank all the persons who worked out the results described: Drs. J. I. Padro´n, E. Q. Morales, P. Gorta´zar, M. Ravina, and Mrs. M. Trujillo. Support of this work by the Direccio´n General de Investigacio´n Cienti´fica y Te´cnica (DGICYT), Ministerio de Educacio´n y Ciencia (Spain), through grant PB930559, and by the Gobierno de Canarias is gratefully acknowledged.

REFERENCES AND NOTES 1. (a) F. Ciardelli and P. Salvadori, eds., Fundamental Aspects and Recent Developments in Optical Rotatory Dispersion and Circular Dichroism, Heyden & Son, London (1973). (b) K. Nakanishi, N. Berova, and R. W. Woody, eds., Circular Dichoism, Principles and Applications, VCH Publishers, New York (1994). 2. (a) N. Harada and K. Nakanishi, Circular Dichroic Spectroscopy-Exciton Coupling in Organic Stereochemistry, University Science Books, Mill Valley, Calif. (1983). (b) Ref. 1b, p. 361–398. 3. C. D. Fasman, ed., Circular Dichroism and the Conformational Analysis of Biomolecules, Plenum Press, New York, p. 25–157 (1996). 4. (a) W. T. Wiesler, J. T. Va´zquez, and K. Nakanishi, J. Am. Chem. Soc., 109: 5586

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5.

6. 7. 8. 9. 10.

11.

12.

13.

14. 15. 16. 17. 18. 19.

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(1987). (b) J. T. Va´zquez, W. T. Wiesler, and K. Nakanishi, Carbohydr. Res., 176: 175 (1988). (a) M. L. Phillips, E. Nudelman, F. C. A. Gaeta, M. Perez, A. K. Shingai, S. Hakomori, and J. C. Paulson, Science, 250: 1130 (1990). (b) L. A. Lasky, Science 258: 964 (1992). (c) R. A. Dwek, Chem. Rev., 96: 683 (1996). R. U. Lemieux and S. Koto, Tetrahedron 30: 1933 (1974). L-C. Lo, N. Berova, K. Nakanishi, E. Q. Morales, and J. T. Va´zquez, Tetrahedron Asymmetry, 4: 321 (1993). H.-W. Liu and K. Nakanishi, J. Am. Chem. Soc., 104: 1178 (1982). E. Q. Morales, J. I. Padro´n, M. Trujillo, and J. T. Va´zquez, J. Org. Chem., 60: 2537 (1995). We define A C and A B values as the amplitudes of split CD Cotton effects in the cinnamate–cinnamate and in the benzoate–benzoate coupling regions centered about the cinnamate λ max 311 nm and about the benzoate λ max 245 nm, respectively. (a) G. D. Wu, A. S. Serianni, and R. Barker, J. Org. Chem., 48: 1750 (1983). (b) Y. Nishida, H. Ohrui, and H. Meguro, Tetrahedron Lett., 25: 1575 (1984). (c) H. Ohrui, Y. Nishida, M. Watanabe, H. Hori, and H. Meguro, Tetrahedron Lett., 26: 3251 (1985). (d) H. Hori, Y. Nishida, H. Ohrui, H. Meguro, and J. Uzawa, Tetrahedron Lett., 29: 4457 (1988). (e) Y. Nishida, H. Hori, H. Ohrui, H. Meguro, J. Uzawa, D. Reimer, V. Sinwell, and H. Paulsen, Tetrahedron Lett., 29: 4461 (1988). (f) K. Bock, and J. Duus, J. Carbohydr. Chem., 13: 513 (1994). The following set of three equations is used to calculate the ratio of P gg , P gt , and P tg : 1.3 P gg ⫹ 2.7 P gt ⫹ 11.7 P tg ⫽ J H5,H6S ; 1.3 P gg ⫹ 11.5 P gt ⫹ 5.8 P tg ⫽ J H5,H6R ; P gg ⫹ P gt ⫹ P tg ⫽ 1. See Ref. 11. The stereoelectronic exo-anomeric effect is the preference for the gauche [(sc): synclinal] conformation about the glycosidic C-OR bond of sugar derivatives. (a) R. U. Lemieux, A. A. Pavia, J. C. Martin, and K. A. Watanabe, Can. J. Chem., 47: 4427 (1969). (b) Deslongchamps, Stereoelectronic Effects in Organic Chemistry (J. E. Baldwin, ed.), Organic Chemistry Series, Vol. I, Pergamon Press, Oxford (1983). (c) A. J. Kirby, The Anomeric Effect and Related Stereoelectronic Effects at Oxygen (K. Hafner, C. W. Rees, B. M. Trost, J. M. Lehn, P. R. Schleyer, and R. Zahradnik, eds.), Reactivity and Structure Concepts in Organic Chemistry, Vol. 15, Springer-Verlag, Berlin (1983). (d) The Anomeric Effect and Associated Stereoelectronic Effects (G.R.J. Thatcher, ed.) ACS Symposium Series 539, Washington, D.C. (1993). J.-P. Praly and R. U. Lemieux, Can J. Chem., 65: 213 (1987). A. Cosse´-Barbi, D. G. Watson, and J. E. Dubois, Tetrahedron Lett., 30: 163 (1989). J. I. Padro´n and J. T. Va´zquez, Chirality, 9: 626 (1997). J. I. Padro´n, E. Q. Morales, and J. T. Va´zquez, J. Org. Chem., 63: 8247 (1998). M. Trujillo, E. Q. Morales, and J. T. Va´zquez, J. Org. Chem., 59: 6637 (1994). (a) J. A. Dale and H. S. Mosher, J. Am. Chem. Soc., 95: 512 (1973). (b) G. R. Sullivan, J. A. Dale, and H. S. Mosher, J. Org. Chem., 38: 2143 (1973). (c) T. Kusumi, I. Ohtani, Y. Inouye, and H. Kakisawa, Tetrahedron Lett., 29: 4731 (1988). (d) I. Ohtani, T. Kusumi, O. M. Ishitsuka, and H. Kakisawa, Tetrahedron Lett., 30: 3147 (1989). (e) I. Ohtani, T. Kusumi, Y. Kashman, and H. Kakisawa, J. Org. Chem., 56: 1296 (1991). (f) I. Ohtani, T. Kusumi, Y. Kashman, and H. Kakisawa,

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20.

21. 22. 23. 24. 25.

26. 27. 28. 29. 30.

31. 32. 33.

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J. Am. Chem. Soc., 113, 4092 (1991). (g) T. Pehk, E. Lippmaa, M. Lopp, A. Paju, B. C. Borer, and R. J. K. Taylor, Tetrahedron: Asymmetry, 4: 1527 (1993). (a) N. Purdie, and K. A. Swallows, Anal. Chem., 61: 77A (1989). (b) N. Purdie, Analytical Applications of Circular Dichroism in Techniques and Instrumentation in Analytical Chemistry, Vol. 14 (N. Purdie and H. G. Brittain, eds.), Elsevier Science B. V., Amsterdam, p. 241–278. (1994). A. Gergely, J. Pharm. Biomed. Anal., 7: 523 (1989). N. Purdie and K. A. Swallows, Anal. Chem., 59: 1349 (1987). P. Gorta´zar, and J. T. Va´zquez, J. Pharm. Sci., 83: 1204 (1994). R. Nagarajan and D. O. Spry, J. Am. Chem. Soc., 93: 2310 (1971). The regression line equations of the linear relationship between the ellipticity angle and the concentration of the antibiotic were defined as θ ⫽ mc ⫹ z, where θ is the ellipticity angle (millidegree [mdeg]) at the selected wavelength, c is the concentration (µg/mL), m is the slope of the fitted line, and z is the θ intercept of the regression line. A. M. Brisson and J. B. Fourtillan, J. Chromatogr., 223: 393 (1981). P. Gorta´zar, M. Ravina, and J. T. Va´zquez, J. Pharm. Sci., 84: 1316 (1995). J. M. Bowen, and N. Purdie, J. Pharm. Sci., 71: 836 (1982). G. Jung, M. Ottnad, and W. Voelter, Eur. J. Drug Metab. Pharmacokinet., 3: 131 (1977). Standard solutions were prepared by dissolving the accurately weighed compound in calibrated flasks with control human urine. Measured volumes of the standard solutions were diluted into calibrated flasks with human urine and 20 µL of the resulting solutions diluted into 5-mL calibrated flasks with distilled water. For CD measurement a 10-mm cylindrical quartz cell was used. For further details see Ref. 27. See, for example, Ref. 1a, pp. 352–372. I. Sjo¨holm and T. Sjo¨din, Biochem. Pharmacol., 21: 3041 (1972). P. Gorta´zar, A. Roe¨n, and J. T. Va´zquez, Chirality, 10: 507 (1998).

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8 Furan-Terminated Cationic π-Cyclizations in the Synthesis of Natural Products Steven P. Tanis Pharmacia & Upjohn, Inc., Kalamazoo, Michigan

I. INTRODUCTION Approximately 23 years ago the sesquiterpene warburganal (1) was isolated from the East African medicinal plant Warburgia ugandensis [1], and we had the opportunity to embark on a synthesis prior to the publication of the structure of 1. Our initial approach constructed the sesquiterpene framework through a DielsAlder cycloaddition of 1-vinyl-2,6,6-trimethyl-1-cyclohexene with dimethyl acetylenedicarboxylate, as shown in Scheme 1 [2]. The Diels-Alder adduct 2 (83%) resisted the development of the desired trans-ring fusion via catalytic hydrogenation, affording instead only cis-fused products. While we were wrestling with this problem, which was eventually solved [2], we considered alternatives that might directly construct the desired trans-fused sesquiterpene skeleton. The cationic π-cyclization, which has been widely utilized in the construction of polycyclic ring systems, has been the object of intense study since the early 1950s [3]. Initial forays into this arena demonstrated the syntheses of fused ring terpenoid-type systems, and later efforts demonstrated the construction of spiro and bridged ring carbocyclic systems [4]. These studies have demonstrated that a wide variety of initiating functions (e.g., epoxides, allylic alcohols, enones, olefins, carbinolamides) and terminating moieties (e.g., aromatic rings, acety-

Dedicated to Professor Koji Nakanishi on the occasion of his 70th birthday.

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Scheme 1 Approach to warburganal 1.

lenes, allylsilanes, allenes, olefins) can be incorporated into the cyclization substrate to lead to terpenoids and alkaloids. In the synthesis of simple fused ring systems, such as the perhydronaphthalene moiety of warburganal (1), this method has produced the target 4,4,8a-trimethyl-trans-fused skeleton as shown in Eq. (1) [5]. The issue to be considered prior to examining a cationic π-cyclization for the preparation of 1 was the identity of the 1,4-dialdehyde equivalent as the terminator function for the cyclization.

(1)

In principle, the 1,4-dialdehyde unit found in warburganal (1) could be prepared by the hydrolysis of a furanoid precursor. This suggested the possibility of terminating a cyclization such as that depicted in Eq. (1) with the π-excessive aromatic ring of furan. The natural propensity of furan to suffer electrophilic aromatic substitution at an available α-position (Fig. 1, Path a, X ⫽ H) dictates that a blocking group X (Fig. 1, Path b, X ⫽ Me, TMS, SPh, etc.) be employed in order to have access to the desired target substitution pattern. When we were considering the chemistry of Fig. 1, we were faced with a paucity of precedent in the literature. We surmised that it was likely due to the relative inaccessibility of suitable substrate [6], the questionable nucleophilic character of the furyl residue relative to more standard terminator functions [7], and the increased acid liability of the more highly substituted product compared with the starting material [7]. As a result, careful choice of both the initiating

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Figure 1 A furan-based approach to the synthesis of warburganal. (Source: Ref. 1.)

moiety and the reaction conditions would be required if the projected chemistry were to be successful. Our interest in developing substituted furans as nucleophilic synthons in annulative processes stemmed from the variety of useful functional groupings that might be realized from the relatively unreactive furyl nucleus. As mentioned (Fig. 1), a furan can serve as the operational equivalent of a 1,4-dialdehyde; Fig. 2 indicates additional acyclic, heterocyclic, and carbocyclic subunits that can be derived from a furan after standard chemical manipulations.

Figure 2 Furan functional equivalencies.

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Scheme 2 Possible disconnections to provide linearly fused-, spirocyclic-, and bridgedring-containing compounds.

With a variety of potentially sensitive functionalities that might be derived from the furan unit, we designed the connections shown in Scheme 2 to provide access to linearly fused-, spirocyclic-, and bridged-ring-containing systems. For the electronically favored furan 3-to-2 closure, we can consider the reaction of the hypothetical furan 7 with a variety of doubly reactive synthons (8–10). The interaction of a reactive side chain nucleophile/electrophile of 7 with 8–10 will provide a coupling product; subsequent activation of the nascent-electron-deficient center of 8–10, followed by aromatic substitution, could provide fused(11), spirocyclic- (12), and bridged- (13) ring systems. Manipulation of the furan nucleus (Fig. 2) and other residual functionality would provide complex intermediates for natural products synthesis. The regiochemistry of furan termination of the reactions of Scheme 2 could be altered by connecting the furyl tether to the furan 2-position. Success in this electronically less favored cyclization paradigm would lead to the isolation of the compounds of Scheme 2, regioreversed with respect to the furan residue.

II. LINEARLY FUSED TERPENOID COMPOUNDS (SCHEME 2, PATH A) A.

Epoxide Initiated Furan 3-to-2 Cyclizations

The epoxide moiety, among other groups, has been widely employed as an initiator function for cationic π-cyclizations [8]. Workers in the field have employed

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a wide variety of Lewis acids to initiate the cyclization sequence, and when this is coupled with the ease of precursor olefin epoxidation or introduction intact, it suggested the epoxide as our first cyclization initiator. The cyclization substrates that were examined were designed to permit entry into five-, six-, or seven-membered ring systems. In order to prevent ambiguity in the choice of a given ring system available from a given oxirane, the epoxide function was biased to favor one mode of C-O bond polarization over the alternative bond [8]. We also studied the placement of the initiating function within the forming cycle (endocyclic) or outside the forming cycle (exocyclic) [9]. On the basis of the work of Baldwin, we anticipated a successful cyclization for 5-, 6-, and 7-exo systems, although only 6-endo should proceed readily. The epoxy furan substrates selected for this study were constructed as shown in Eq. (2). A Grignard reagent 14, prepared from 3-chloromethylfuran, was coupled with a haloalkene, in the presence of a catalyst, to give furyl-olefins 15, which were epoxidized with m-chloroperoxybenzoic acid (MCPBA) to afford furyl-epoxides 16 [10]. Although the furan moiety is known to be susceptible to oxidation, the relative rates of furan vs. olefin attack as a function of the degree of substitution had not been reported. The compounds of Table 1 were prepared by the chemistry of Eq. (2).

(2) As shown in Table 1, the coupling reactions proceed smoothly when 14 was reacted with alkyl and allylic halides (runs 2–5, Li2CuCl4 as catalyst) [7a,11]. The synthesis of olefin 17, utilizing a vinyl halide, required anhydrous FeCl3 as a catalyst [11]. Furyl-olefins 17–21 were submitted to standard epoxidation conditions (1.05 eq. MCPBA, CH2Cl2, 0°C) to furnish good yields of epoxides 22 (86%), 24 (85%), and 26 (81%), all derived from trisubstituted olefins. Furyl-olefin 18 led to epoxide 23 in only 25% yield, and the epoxidation of olefin 20 provided none of the expected epoxide 25. Closer examination of run 2 indicated that epoxide 23 was accompanied by unreacted 18 (23%) and the anhydride derived from 18 (37%). Olefin 20 led only to anhydride and recovered 20. The three remaining epoxides required in quantity for the study were prepared as shown in Eq. (3) and (4). 3-Furylmethyllithium [7a,12] [Eq. (3)] was reacted with epoxy iodides 27 and 28 [hexamethylphosphorus triamide (HMPA), ⫺25°C] [13] to give epoxy furans 25 (73%) and 29 (68%), respectively. Monosubstituted epoxide 30 was prepared in 37% yield from a coupling of 3-furylmethyllithium and a protected iododiol, as shown in Eq. (4).

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Table 1 Synthesis and Oxidation of 3-Furyl Olefins [Eq. (2)] Run

Olefin

Catalyst

Fulyl-alkene (yield)

Furyl-epoxide (yield)

(3)

(4)

1. Cyclization Studies Relatively potent Lewis acids such as boron trifluoride etherate are often selected to catalyze epoxy olefin cyclizations [8]. Given the acid lability of the starting furans and the increased acid sensitivity of the products, the choice of Lewis acid should have a profound effect on the partitioning of the reaction between a fruitful cyclization and acid-mediated decomposition. Six Lewis acids were utilized

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the initial study [7a], BF3⋅OEt2 [8], EtAlCl2 [14], Et2AlCl [14], Al2O3 [13], Ti(OiPr)3Cl [15], and ZnI2 [16]. The choice of Lewis acid was dictated by (1) the ability to modify the potency of a group of Lewis acids with a common metal center readily and (2) the possibility of moderating the Brønsted acidity of the medium through the choice of the Lewis acid. Adventitious protic acid might be scavenged by a Lewis acid possessing a metal–carbon bond releasing an alkane; alternatively with the proper choice of metal, the product metal–alcohol complex should be a much weaker protic acid compared to a BF3 –alcohol complex. The substrate epoxy furans were exposed to boron trifluoride etherate (0.3 eq.) in methylene chloride at ⫺25°C as the standard cyclization conditions (Table 2). Five-membered ring precursors 22, 23, and 30, when treated with BF3⋅OEt2, afforded only allylic alcohols 32 (62%), 34 (53%), and 36 (49%), respectively. Only epoxy furans 24 and 25 gave appreciable quantities of cyclized products, 37 (47%) and 39 (30%), respectively. The majority of the material isolated from the BF3⋅OEt2-mediated cyclizations consisted of the depicted allylic alcohols, and the mass balances were poor (ca. 60% or less). The poor yields of cyclized materials and low mass balances are in agreement with the surmises regarding the Lewis/Brønsted acidity requirements for these reactions. The aluminum-based Lewis acids, EtAlCl2, Et2AlCl, and Al2O3, provided better mass balance (ca. 70% or more), and variable amounts of cyclized products. The treatment of 22, 23, and 30, five-membered ring precursors, with either EtAlCl2 or Et2AlCl led to good yields of allylic alcohols. Only furans 24 (6endo), 25 (6-exo), and 26 (7-endo) gave any cyclized material (10%–22%) with EtAlCl2 or Et2AlCl. Alumina (Al2O3) provided only allylic alcohol for all epoxy furans except 24, which furnished a 32% yield of 37 and a 51% yield of allylic alcohol 38. The modification of the Lewis acid to provide a protonolyzable M-C bond, thus reducing Brønsted acidity of the medium, did lead to improved mass balance; however, elimination was the dominant path with the alkylaluminum halides. Further modification of the aluminum-centered Lewis acid to alumina also provided improved mass balance but little cyclization. The next most Lewis acidic compound in the series Ti(Oi-Pr)3Cl [15] proved to be an efficient and useful promoter of epoxy furan cyclization. As before, five-membered-ring precursor oxiranes 22, 23, and 30 did not lead to desired cyclized products, with elimination products 32 (80%) and 34 (72%) coming from 22 and 23, respectively. The monosubstituted epoxide 30 could not be induced to react, even after exposure to 3 eq. of Ti(Oi-Pr)3Cl for 24 h at room temperature. Similar treatment of epoxy furan 24 led to the formation of the desired cyclized adduct 37 in 78% yield, with no elimination product observed. 6-exo-Epoxide 25 and 7-endo-precursor 26 afforded cyclized adducts 39 (89%) and 41 (87%), respectively, with the latter accompanied by 8% of allylic alcohol 42. Even 7-exo-precursor epoxide 29 gave a respectable yield of cyclized product 43 (36%) when treated with Ti(Oi-Pr)3Cl. Epoxides 22, 23, and 30, five-membered ring precursors, were next exposed

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Table 2 Initial Cyclization Studies, Effect of Ring Size, Epoxide Placement, and Lewis Acid Furyl-epoxide

Product(s)

22 BF3-OEt2 (0.3 eq.) Et2AlCl (2 eq.) Al2O3 Ti(OiPr)3Cl (3 eq.) ZnI2 (3 eq.)

31 0% 0% 0% 0% 0%

32 62% 85% 83% 80% 76%

BF3-OEt2 (0.3 eq.) Et2AlCl (2 eq.) Ti(OiPr)3Cl (3 eq.) ZnI2 (3 eq.)

33 0% 0% 0% 0%

34 53% 85% 72% 70%

35 0% 0%

36 49% 78%

23

30 BF3-OEt2 (0.3 eq.) Et2AlCl (2 eq.) Ti(OiPr)3Cl (3 eq.) ZnI2 (3 eq.)

25%

44%

BF3-OEt2 (0.3 eq.) EtAlCl2 (2 eq.) Et2AlCl (2 eq.) Al2O3 Ti(OiPr)3Cl (3 eq.) ZnI2 (3 eq.)

37 47% 16% 22% 32% 78% 71%

38 0% 57% 49% 51% 0% 0%

24

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no reaction

Table 2 Continued Furyl-epoxide

Product(s)

25 BF3-OEt2 (0.3 eq.) EtAlCl2 (2 eq.) Et2AlCl (2 eq.) Al2O3 Ti(OiPr)3Cl (3 eq.) ZnI2 (3 eq.)

39 30% 0% 10% 0% 89% 70%

40 10% 73% 70% 81% 0% 0%

BF3-OEt2 (0.3 eq.) EtAlCl2 (2 eq.) Et2AlCl (2 eq.) Al2O3 Ti(OiPr)3Cl (3 eq.) ZnI2 (3 eq.)

41 0% 0% 10% 0% 87% 88%

42 41% 76% 69% 83% 8% 9%

BF3-OEt2 (0.3 eq.) EtAlCl2 (2 eq.) Et2AlCl (2 eq.) Al2O3 Ti(OiPr)3Cl (3 eq.) ZnI2 (3 eq.)

43 10% 0% 0% 0% 36% 23%

44 12% 64% 73% 79% 47% 52%

26

29

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to the final Lewis acid in this initial study ZnI2 (3 eq. CH2Cl2, room temperature). Allylic alcohols 32 (76%) and 34 (70%) were produced from 22 and 23, respectively; however, furyl-epoxide 30 gave a mixture of the five-membered ring target 35 (25%) and allylic alcohol 36 (44%). Epoxides 24, 25, 26, and 30 afforded modest to excellent yields of cyclic products. The results presented in Table 2 suggested that the furan- (3-to-2 closure) terminated/epoxide-initiated cationic π-cyclization is useful for the construction of six- and seven-membered rings. Good to excellent yields can be realized with a judicious choice of Lewis acid. However, closure to form five-membered rings is difficult. 2. Syntheses of Pallescensin A and Aphidicolin A more rigorous test of the epoxy furan cyclization would require the formation of two or more rings, with the development of stereochemistry and/or transmission of chirality. Pallescensin-A (45) [7a,17,18] was selected as the initial test of the former, and a formal total synthesis of aphidicolin (46) [19] would be used to illustrate the latter.

Scheme 3 presents the total synthesis of (⫾)-pallescensin A (45). 6,7-Epoxygeranyl chloride [10,20] was coupled with 3-furylmethylmagnesium chloride to give 7,8-epoxydendrolasin (47) in 79% yield [10]. Cyclization with BF3⋅OEt2 gave 3 β-hydroxypallescensin A (48) in 47% yield [21]. As expected from the results of Table 2, triisopropoxytitanium chloride and zinc iodide afford 48 in higher yields, 62% and 65%, respectively, and the reaction mixtures were cleaner and less complex. Compound 48 is smoothly converted to pallescensin A (45), as described by Nasipuri and Das [21]. (⫹)-Aphidicolin (46), a diterpene tetraol produced by the mold Cephalosporium aphidacola Petch [19a,b], has provoked interest as a synthetic target over the past 20 years [22]. (⫹)-Aphidicolin (46) has been associated with a range of biological activities, including antiviral [19c,d], antitumor [19e], and antileukemic [19f], activity as well as activity as a reversible inhibitor of deoxyri-

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Scheme 3 The synthesis of (⫾)-pallescensin A (45).

bonucleic acid (DNA) polymerase-α [19g]. McMurry [23] has reported the synthesis of (⫾)-46 from diketone 49. We envisioned accomplishing a synthesis of (⫾)-49 and (⫺)-49 [24] via furan-terminated cationic π-cyclization, as shown in Scheme 4. The chemistry outlined in Scheme 4 would exploit a furan-terminated cationic-π-cyclization to establish the carbon framework of 49 rapidly with the correct relative and absolute configuration created at carbons 4, 5, and 10; a furan to dione conversion would provide 49. The potential availability of the modified epoxygeranyl chloride precursor to furan 51 in optically pure form from a Sharpless asymmetric epoxidation [25] is an additional advantage to the route depicted. In the event, geraniol was converted to the related benzoate (Scheme 5), which readily underwent allylic hydroxylation (SeO2, TBHP) [26] to give alcohol 52 (70%). Sharpless asymmetric epoxidation of hydroxybenzoate 52 gave (⫺)53 in 71% yield [25a]. The optical purity of (⫺)-53 was judged to be ⱖ95% ee after an examination of (⫺)-53 by nuclear magnetic resonance (NMR) in the presence of Eu(hfpc)3 [25a] and high-performance liquid chromatography (HPLC) analysis of the related Mosher ester [27]. Alternatively, (⫺)-53 could be prepared in 83% yield and ⱖ95% ee via the Sharpless catalytic asymmetric epoxidation protocol [25b]. We considered a number of blocking groups with which to protect the free hydroxyl group of (⫺)-53 during the halogenation/ coupling/cyclization steps. Initially we selected a t-butyldimethylsilyl (TBDMS) moiety for this function. This proved to be somewhat difficult at the cyclization

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Scheme 4 A furan-terminated cationic π-cyclization approach to aphidicolin (46).

Scheme 5 The formal total synthesis of (⫾)-aphidicolin (46).

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step, when widely variable yields of products were obtained. The required group must be smaller than the TBDMS group, survive the chlorination/coupling/cyclization steps, yet be readily removed to facilitate a chelation-directed reduction of the C-3 ketone. We selected a benzyl ether to block the free hydroxyl of (⫺)53. Toward that end (⫺)-53 was treated with sodium hydride (NaH) and benzyl bromide in tetrahydrofuran (THF) with added n-Bu4NI to provide (⫺)-54. Benzoate hydrolysis (NaOMe, MeOH, n-Bu4NI) furnished the related alcohol, which was immediately converted to the allylic chloride (⫹)-55 (n-BuLi, p-TsCl, LiCl) [28] in 84% yield for the two steps. Chloride (⫹)-55 was smoothly coupled with (3-furyl)methylmagnesium chloride, giving (⫺)-56 in 79% yield. In studies described in Table 2 and Scheme 3, ZnI2 and Ti(Oi-Pr)3Cl stood out as the Lewis acids of choice for epoxy furan cationic π-cyclizations. In the present case, exposing the more highly oxygenated (⫺)-56 to ZnI2 and Ti(Oi-Pr)3Cl led to highly variable (0%–49%) yields of (⫹)-57. After considerable experimentation it was discovered that treatment of (⫺)-56 with BF3⋅OEt2 (6 eq.) and Et3N (3 eq.) in CH2Cl2 /PhH/hexanes (1: 1 :1) at ⫺78°C gave an excellent 72% yield of (⫹)57. The successful completion of the synthesis then required the inversion of stereochemistry at the C-3 position. Alcohol (⫹)-57 was oxidized under Swern conditions [29] to afford ketone (⫹)-58 in 97% yield, which then had to be selectively reduced. The reduction of (⫹)-58, with L-Selectride, was examined with and without added metal salts. It was assumed that the proper choice of metal salt would conspire to provide a chelated intermediate, which would be selectively reduced [30]. Precomplexation of (⫹)-58 with ZnI2, MgBr2, Ti(Oi-Pr)3Cl, or Ti(Oi-Pr)4 followed by the addition of L-Selectride (⫺78°C, Fig. 3) gave 3α:3β alcohols 63 and 64 in ratios ranging

Figure 3 Reduction selectivity of (⫾)-58 with L-Selectride in the presence and absence of added metal salts.

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from 1.2 : 1 to 8.5 :1. The optimal conditions (Fig. 3, entry d) employed 2 equivalents of MgBr2 ⋅ OEt2 and provided a very respectable 8.5 : 1 ratio of 63 and 64 in 95% combined yield. With optimal reducing conditions in hand, the reduction of (⫹)-58 on a larger scale was examined as shown in Scheme 5. Ketone (⫹)-58 was treated with MgBr2 ⋅ OEt2 and L-Selectride to give an 8.5 : 1 mixture of alcohols 63 and 64 (Fig. 3). Reductive cleavage of the benzyl ether, as described by Kutney [31], led to (⫹)-59 in 89% yield, as an 8.5 :1 mixture at C-3. The mixture was readily separated after the conversion of the cis-diol 63 to the related acetonide (⫹)-60 (acetone, H⫹; 93%). Furan (⫹)-60 was brominated at the available α-position (NBS, DMF) [32], the bromide was immediately subjected to metal–halogen exchange (n-BuLi), and the lithiofuran was alkylated (MeI) to provide 61 in 65% overall yield. Furan oxidation with MCPBA [24,33] furnished the ene-dione (⫺)62 (97%), which was hydrogenated (H2 /Pd-C) to give the (⫺)-49 (96%), completing the formal total synthesis of (⫺)-aphidicolin (46) in 16 steps and 10.7% overall yield from geraniol.

B.

Allylic Alcohol–Initiated Furan 3-to-2 Cyclizations

To expand the usefulness of furan-terminated cationic π-cyclizations in the synthesis of terpenoids, we wished to extend the scope of cyclization initiators to allylic alcohols, enones, and acid derivatives. In the case of allylic alcohols, the nature of the species shown in Scheme 2, Path A, could be altered to facilitate selectivity in the initial bond formation. The utilization of a vinyl epoxide [Eq. (5)] [34] or the enol ether of an α,β-epoxy ketone [Eq. (6)] [34a] as bis-electrophilic synthons would provide selectivity, as the second electrophilic center would result after the initial SN2′-like addition of an organometallic reagent to 65 [35], 66 [34a,36], and 67 [37] [Eq. (7)] [34a].

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(7) Initial attempts to construct linearly fused terpenoid precursors via furan 3-to-2 terminated cationic π-cyclization utilized simple variants of 66 and 67 prepared from cyclohexenone. Scheme 6 illustrates the syntheses of exocyclic allylic alcohol cyclization precursors from 66 (n ⫽ 1, Scheme 2, Path A). Cyclohexenone was converted to the spirovinyl epoxide 66 by the addition of methylthiomethyl lithium to cyclohexenone, to give the related alcohol (89%), methylation of sulfur (MeI, 99%), and epoxide closure (KOt-Bu, 91%) [34a,36]. The Grignard reagents derived from 2-(3-furyl)-1-bromoethane and 3-(3-furyl)-1-bromopropane were treated with CuCN followed by 66 to give the SN2′ addition products 68 (m ⫽ 2, 56%) and 69 (m ⫽ 3, 58%), respectively. For anticipated ease of ionization, alcohols 68 and 69 were converted to their respective secondary allylic alcohols 70 (75%) and 71 (66%), respectively, via pyridinium chlorochromate (PCC) oxidation and MeLi addition to the derived enals. The synthesis of endocyclic allylic alcohol cyclization substrates from vinyl-epoxide 67 (n ⫽ 1) is presented in Scheme 7. The Grignard reagents derived from 2-(3-furyl)-1-bromoethane and 3-(3-furyl)-1-bromopropane were treated with CuCN followed by enol-ether epoxide 67 [37] to give SN2′ addition products. Acid treatment produced enones 72 (m ⫽ 2, 75%) and 73 (m ⫽ 3, 72%), the products of enol ether hydrolysis and dehydration. Methyl lithium addition then afforded the endocyclic tertiary allylic alcohol cyclization substrates 74 (90%)

Scheme 6 The synthesis of linearly fused–allylic alcohol substrates—I.

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Scheme 7 The synthesis of linearly fused–allylic alcohol substrates—II.

and 75 (90%), respectively. The inclusion of only six- and seven-membered precursor chain lengths (m ⫽ 2, 3) in the compounds of Schemes 6 and 7 was based upon the results presented in Table 2. 1. Cyclization Studies Allylic alcohols have been extensively employed as initiators in cationic π-cyclizations [3], and the reaction conditions that have been employed generally involve a protic acid of reasonable strength in a solvent in which it is soluble. Of the many conditions reported in the literature, the two-phase mixture of anhydrous formic acid and cyclohexane [38] was selected for this study as the mildest method for initiating the cyclization of allylic alcohols 70, 71, 74, and 75 (Scheme 8). Alcohols 70 and 71, designed to form six- and seven-membered rings, respectively, from an allylic alcohol with the hydroxyl center exocyclic to the existing ring, were separately exposed to formic acid in cyclohexane for 20 min at room temperature. Alcohols 70 and 71 smoothly cyclized to give the six-membered target 76 and the seven-membered target 77 in 68% and 61% yields, respectively, as mixtures of exo-ethylidene double-bond isomers. The endocyclic allylic alcohols 74 and 75 were next treated with formic acid in cyclohexane to give tricyclic furans 77 (73%) and 78 (56%), respectively. The studies of Scheme 8 firmly established allylic alcohols as suitable initiators in furan-terminated cationic πcyclizations. Next we examined the cyclization of enals 79 and 80, which were prepared as intermediates from the oxidation of 68 and 69, as previously illustrated in Scheme 6. Enals 79 and 80 [Eq. (8)] were treated with BF3⋅OEt2, SnCl4, TiCl4, EtAlCl2, Et2AlCl, MgBr2⋅OEt2, ZnI2, and Ti(Oi-Pr)3Cl, in a variety of solvents at various temperatures, to no avail. Extensive decomposition was observed when BF3⋅OEt2, SnCl4, TiCl4, EtAlCl2, and Et2AlCl were employed; the use of MgBr2⋅OEt2, ZnI2, and Ti(Oi-Pr)3Cl led to recovered starting materials. Acylative-type enal cyclizations similar to those reported by Andersen [39], Marshall [40], and Harding [41]

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Scheme 8 Linearly fused–allylic alcohol initiated–furan terminated cyclizations.

were also examined. The treatment of 79 and 80 with either Ac2O/HClO4 /EtOAc or (CF3CO)2 O/CF3CO2H resulted in a facile and high-yield acylation of the furyl nucleus at the 2-position. Protic acid treatment (HCOOH, c-C6H12) of 79 and 80 led to recovered starting material or, after extended treatment, extensive decomposition.

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C.

Acylium Ion–Initiated Furan 3-to-2 Closure: The Syntheses of (ⴞ)- and (ⴚ)-Fastigilin-C

1. First-Generation Approach The cytotoxic, antineoplastic helenanolide (⫺)-fastigilin C (81) [42] was an attractive target for total synthesis via furan-terminated cationic π-cyclization, as

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illustrated in Eq. (9). The butyrolactone moiety of 81 would be derived from the furan of acyl-furan 82, which would be derived from cyclopentanone 83, which possessed a cyclization initiator (acylium ion precursor) and the terminator furan. An attractive benefit, in addition to the regiospecific protolactone introduction, of incorporating a furan into dione 82 was the possibility of routine control of stereochemistry about the periphery of the bicyclo[5.3.0]decane by inducing the normally flexible seven-membered B-ring to adopt a well-defined chairlike conformation [43].

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Syntheses of cyclopentanones such as 83 require control of two exocyclic stereocenters and concomitant trans-addition of the elements of propionate and 3-furaldehyde to the 3- and 2-positions, respectively, of 2-methyl-2-cyclopentenone [44]. Our first-generation approach to (⫾)-81 is shown in Scheme 9 [44]. Mukaiyama has described a trityl salt–catalyzed, silicon transfer, tandem conjugate addition–aldol condensation sequence [45] to form trans-2,3-disubstituted cyclopentanones with predictable exocyclic stereochemistry. Such a proto-

Scheme 9 First-generation approach to (⫾)-fastigilin C (81).

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col was ideally suited for the synthesis of the target cyclopentanone 83. Toward that end, t-butyl thiopropionate was converted to the related TBDMS-enol ether (TBDMSOTf, i-PrNEt2) [46] 84 (80%, Z/E ⬎95: 5), which was combined with 2-methyl-2-cyclopentenone (CH2Cl2, ⫺95°C bath), and the resulting mixture was treated with 5 mol% of Ph3CSbCl6. After 20 min, 3-furaldehyde was added and the mixture was warmed to room temperature overnight to give a mixture of 83 and pro-C-6-iso-83 (73%, 6 :1), which was difficult to separate. The ratio of 83 to pro-C-6-iso-83 is in agreement with the reports of Mukaiyama [45] and is temperature-dependent, falling to 2.5 :1 at ⫺80°C. The stereochemical outcome has been rationalized [45,47] as being the result of consecutive conjugate addition and aldol reactions, which proceed in a trans-fashion across the cyclopentenone 2and 3-positions, through an open transition state. The B-ring closure was readily accomplished by exposing 83 to Hg(OTFA)2 (2 eq., anhydrous CH3CN, room temperature) [48] to afford dione 82 (R ⫽ TBDMS) in 65% yield. The fifth of the seven B-ring stereocenters of fastigilin C (81) was selectively introduced by reduction of the 9-ketone with NaBH4 (EtOH) to provide the 9β-alcohol 85 (99%) as a single stereoisomer. This alcohol was protected as the related 2-(trimethylsilyl)ethoxymethyl ether (SEM), giving 86 (99%). At this juncture it was necessary to consider the completion of the endeavor. That is, it would be necessary to develop the A-ring double bond prior to the development of the fragile butyrolactone moiety, to avoid the difficulties encountered by Lansbury et al. [49]. This strategy would require an A-ring enone protection before furan-to-butyrolactone elaboration, followed at some point by a deblocking. The advantages offered by the Scheme 9 route, (1) robust intermediates and (2) a well-defined B-ring conformation that aids in B-ring development, could not outweigh the problems to be overcome, which were (1) when and how to develop the A-ring and (2) difficulties in transforming the chemistry of Scheme 9, a route to (⫾)-81, to access the natural optical antipode (⫺)-81. 2. The Synthesis of (⫾)-Fastigilin C[(⫾)-81] The introduction of an A-ring 2,3-double bond surrogate, which would obviate the anticipated end-game problems of incorporating this unit of unsaturation and also provide for the development of a route to (⫺)-81, was examined next. This might be achieved by substituting 4-methoxy-2-methyl-2-cyclopentenone into the Michael-aldol chemistry of Scheme 9. The synthesis of (⫾)-fastigilin C [(⫾)81] is shown in Scheme 10 [47]. Racemic 4-methoxy-2-methyl-2-cyclopentenone [(⫾)-87] [50] was reacted with the TBDMS-enol ether of t-butyl thiopropionate and 3-furaldehyde, mediated by trityl hexachloroantimonate in CH2Cl2, at ⫺22°C for the Michael reaction, followed by cooling to ⫺78°C for the silicon transfer aldol condensation (addition of 3-furaldehyde). This protocol gave (⫾)-88 in 90% yield, uncontaminated by stereoisomers at any of the derived stereogenic centers.

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Scheme 10 The synthesis of (⫾)-fastigilin C [(⫾)-81].

The relative orientation of the methoxyl group of the cyclopentanone, to the exocyclic stereocenters, could not be ascertained spectroscopically; therefore we elected to continue the synthesis and determine the relative orientation when a suitable crystalline derivative was obtained. Initial attempts to cyclize (⫾)-88 with Hg(OTFA)2 gave only a 12% yield of (⫾)-89, with the bulk of the recovered material corresponding to the carboxylic acid equivalent of (⫾)-88. Exposing (⫾)-88 to mercuric triflate/N,N-dimethylaniline complex [51], with a less nucleophilic triflate counteranion, afforded an excellent 96% yield of the target bicyclo[5.3.0]decane (⫾)-89. The reduction of (⫾)-89 was modified to employ Luche conditions (NaBH4, MeOH, CeCl3 ⋅ 7H2O, ⫺78°C [52], after sodium borohydride in ethanol afforded mixtures. This modification led to the isolation of (⫾)-90 in 93% yield as a single stereoisomer, establishing the fifth stereocenter about the seven-membered B-ring. The C-9-OH was protected as the corresponding TBDMS-ether, giving (⫾)-91 (99%). Numerous attempts to block the 4-ketone of (⫾)-91 as the related

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dioxolane/ketal were unsuccessful. However, (⫾)-91 was readily reduced with diisobutylaluminum hydride (DIBAL) to the corresponding 4α-alcohol (⫾)-92 (97%). Alcohol (⫾)-92 was nicely crystalline and was submitted to single-crystal x-ray analysis, which established the relative configuration in this series to be that shown for (⫾)-92 in Scheme 10. The x-ray stereostructure of (⫾)-92 (Fig. 4) indicated that the C-2-OMe relative orientation is that expected from a steric controlled approach of the Michael nucleophile in the Mukaiyama conjugate addition–aldol protocol [53] and established (S)-4-methoxy-2-methyl-2-cyclopentenone as the requisite starting material for the synthesis of (⫺)-81. Alcohol (⫾)-92 was protected as the corresponding methoxyethoxymethyl (MEM) ether (MEM-Cl, i-Pr2NEt) to furnish (⫾)-93 (97%), which was desilylated (n-Bu4NF, THF), giving (⫾)-94 (98%). Furan-diol 94 was selectively monosilylated (TBDMSCl, imidazole) to give (⫾)-95 (98%), and the 6-OH was protected as the readily removable ethoxyethyl ether analogue (⫾)-96 (96%). The furan unit of (⫾)-96 was smoothly silylated (i. n-BuLi; ii. Me3SiCl), yielding (⫾)⫹97 (95% overall) after careful ethoxyethyl ether cleavage ( p-TsOH). As desired, silylfuran (⫾)-97 was readily oxidized (CH3CO3H) [54] to give butenolide (⫾)-98 (87%), setting the stage for the crucial hydroxyl-directed hydrogenation required to set the required stereochemistry at C-7 and C-8. After examining the Wilkinson catalyst [(Ph3P)3RhCl] [55], Crabtree’s catalyst [Ir(COD)py(P(Cy)3)PF6] [55,56], and [Rh(NBD)(DIPHOS-4)]BF 4 [55,57], it was discovered that the cationic rhodium catalyst readily reduced (⫾)-98, at 1000 psi, affording (⫾)-99 in 82% yield. Lactone (⫾)-99 was converted to αmethylene lactone (⫾)-100 (83%) via the procedure of Lansbury et al. [49], carboxylation followed by treatment with Eschenmoser’s salt. Lactone (⫾)-100 was treated with the symmetrical anhydride derived from 3-methylcrotonic acid [4dimethylaminopyridine (DMAP), Et3N], and the mixture was heated in refluxing xylenes to provide senecioate ester (⫾)-101 (92%). Zinc bromide treatment of (⫾)-101 smoothly and selectively deprotected the C-4-α-oxygen function, leading to alcohol (⫾)-102 (89%). Oxidation, to the 4-one, and α-elimination to furnish the 2,3-double bond, remained before a total synthesis of (⫾)-81 was achieved. Pyridinium chlorochromate (PCC) oxidation of (⫾)-102 gave β-methoxyketone (⫾)-103 (81%), setting the stage for double-bond introduction via

Figure 4 The relative stereochemistry of (⫾)-92.

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Scheme 11 The preparation of (S)-(⫺)-4-hydroxy-2-methyl-2-cyclopentenone.

β-elimination. Ketone (⫾)-103 was heated to reflux in ether with Amberlyst15 to produce (⫾)-fastigilin C [(⫾)-81, 96%], a product of β-elimination and deprotection of the 9-OH. The sequence from 4-methoxy-2-methyl-2-cyclopentenone (⫾)-87 to (⫾)-81 was accomplished in 17 steps and an overall yield of 24.6%. 3. The Synthesis of (⫺)-Fastigilin C [(⫺)-81] The synthesis of (⫺)-81 required a source (S)-(⫹)-4-methoxy-2-methyl-2-cyclopentenone (⫹)-87. An enzymatic resolution of (⫾)-4-hydroxy-2-methyl-2-cyclopentenone [(⫾)-104], shown in Scheme 11, was developed to provide gram quantities of (S)-(⫹)-87. Racemic 4-hydroxy-2-methyl-2-cyclopentenone [(⫾)-104] [50] and β,β,βtrifluoroethyl butyrate [58] were dissolved in ether, and the resulting mixture was treated with porcine pancreatic lipase (PPL) [59]. After 6 days the reaction had proceeded to 51% conversion, providing (S)-(⫺)-104 in 43% yield and 68% ee, and butyrate (R)-105 (44%) after chromatography. The butyrate (R)-105 was cleaved to the alcohol (R)-104, as described by Wong [59] (guanidine, MeOH) (78%, 46% ee), and the stereocenter was inverted (Ph3P, diethylazodicarboxylate, HCO2H, MeOH, Al2O3) [59], furnishing an additional quantity of (S)-alcohol (S)(⫺)-104 (combined 66% yield, 60% ee). The combined (S)-(⫺)-104 was again

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exposed to β,β,β-trifluoroethyl butyrate and PPL in ether to afford (S)-(⫺)-4hydroxy-2-methyl-2-cyclopentenone [(S)-(⫺)-104] in 52% isolated yield and ⱖ98% ee. The alcohol was easily converted to the target (S)-(⫹)-4-methoxy-2methyl-2-cyclopentenone (⫹)-87 as outlined in Eq. (10).

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The chemistry of Scheme 10 was then undertaken, on a large scale, starting with ca. 50 g of (S)-(⫹)-87. The 17-step sequence from (S)-(⫹)-87 to (⫺)-fastigilin C [(⫺)-81] was accomplished in 14% overall yield.

III. SPIROCYCLIC TERPENOID COMPOUNDS (SCHEME 2, PATH B) A.

Allylic Alcohol– and Enone-Initiated Furan 3-to-2 and Furan 2-to-3 Cyclizations

As has been previously discussed, allylic alcohols and enones can serve as initiators for cationic π-cyclizations. These moieties can be prepared by nucleophilic addition of an organometallic reagent to vinyl epoxides as was illustrated in Eqs. (5), (6), and (7). Those equations defined by the disconnection illustrated in Scheme 2, Path A, for the preparation of linearly fused ring-containing compounds by equating vinyl epoxides with a cycloalkyl dication [Eq. (7)], wherein the electron-deficient centers reside on adjacent carbons, are realized sequentially, the first as a result of the SN2′-type reactivity of a vinyl epoxide, and the second as a consequence of the ionization of the allylic alcohol produced from the initial addition. This paradigm ensures regiochemical integrity in the annulation relative to resident markers. In order to prepare spirocyclic compounds by Path B of Scheme 2, one must alter the placement of the electron-deficient centers, placing the first exocyclic, which will allow closure via furan termination at the tertiary pro-spiro center. The addition of an organometallic reagent to an exo-vinyl cycloalkyl epoxide [Eq. (11)] would result in the preparation of an allylic alcohol by SN2′ addition [34,60]. Subsequent acid treatment would reveal the second electron-deficient center, thus equating an exo-vinyl cycloalkyl epoxide with the dication 9 [Eq. (12)].

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(11)

(12)

To examine the utility of allylic alcohols and enones in the synthesis of spirocyclic systems via furan 3-to-2 cyclizations, a variety of exo-methylene 2,3epoxycycloalkanes were prepared [34a,60] and reacted with (3-furyl)alkyl Grignard reagents 104a–c in the presence of CuCN (Scheme 12). This reaction afforded the target allylic alcohol cyclization substrates 105a–e in 59%–82% yields. The desired related enones were smoothly prepared via PCC oxidation (CH2Cl2) of 105a–e to give enones 106a–e (72%–87%). The cyclizations of furyl-allylic alcohols 105a–e were attempted with HCOOH/cyclohexane by analogy to the chemistry of Scheme 8. In the event

Scheme 12 Spirocyclic furan 3-to-2 cyclization precursor synthesis.

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Scheme 13

Allylic alcohol–initiated spirocyclic furan 3-to-2 cyclization.

(Scheme 13), alcohols 105a and 105b were exposed to the two-phase mixture of HCOOH/cyclohexane to give spiro[4,5]decane 107a (58%) and spiro[4,6]undecane 107b (53%), respectively. Similarly, alcohols 105c–e were treated with HCOOH/cyclohexane to furnish the formate of 105c (84%), spiro[5,5]undecane 107d (72%), and spiro[5,6]dodecane 107e (58%), respectively. The enones 106a–e, produced as illustrated in Scheme 12, were dissolved in cyclohexane and treated with formic acid. Of the five substrates shown in Scheme 14, only enones 106a and 106d, leading to spiro[4,5]decane 108a (72%) and spiro[5,5]dodecane 108d (66%), respectively, provided any cyclized products. Enones 106b,c and 106e were recovered unchanged. Compounds 106b,c

Scheme 14

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Enone-initiated spirocyclic furan 3-to-2 cyclization.

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Scheme 15 Spiro-ring formation, furan-2-to-3-closure.

and 106e also proved resistant to cyclization under a wide variety of other reaction conditions including Lewis acids and acylating agents. In order to examine whether the formation of a six-membered ring is the necessary factor in a successful enone-initiated/furan-terminated cyclization, the furan 2-to-3 closure related to the chemistry of Scheme 13 was examined. The Grignard reagent derived from 3-(2-furyl)-1-bromopropane was added to the isobutyl enol ether of 2-methyl-1,3-cyclohexanedione (Scheme 15), [60] to give enone 109 (85%) after acidic workup. Enone 109 was exposed to a variety of Brønsted and Lewis acids, as well as acylating agents, to no avail. The target spiro[5,5]undecane furan 110, regioisomer of 108d, was not observed. This relatively less favored (electronically) furan 2-to-3 cyclization [61] was observed when allylic alcohol 111, prepared from 109 in 93% yield (NaBH4, CeCl3), was treated with formic acid/cyclohexane, affording olefin 111 (68%).

IV.

BRIDGED-RING-CONTAINING TERPENOID COMPOUNDS (SCHEME 2, PATH C)

A simple drawing defining a possible disconnection for the synthesis of bridged, ring-containing compounds is presented in Scheme 2, Path C. An existing ring would present a reactive electrophilic/nucleophilic center to interact with a furancontaining moiety possessing a reactive nucleophilic/electrophilic center, thereby creating a ring system with a tethered furan. Cyclization to form the bridged system would result when an electron-deficient center was developed at a noncontiguous carbon center. Three strategies were considered to develop this reactivity

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pattern: (1) create the second reactive center as a result of the initial bond construction; (2) utilize differentially reactive electrophilic centers and maximize selectivity [such as was described in the syntheses of pallescensin A (45) and aphidicolin (46)]; and (3) perform the first bond construction, then introduce the desired electrophilic center. Nakafuran 9 (112), a fish antifeedant isolated from the marine sponge Dysedea fragilis and the sponge predacious nudibranchs Hypselodoris godeffroyana and Chromodoris maridadilus [62], was selected as the test case for bridgedring synthesis via furan-terminated cationic π-cyclization. A retrosynthesis of 112, presented in Scheme 16, suggests that the bicyclo[4.3.1]decane skeleton would be prepared from the readily available 3-furylmethyl dianion (discussed previously) and a highly substituted dication. The synthesis of (⫾)-nakafuran-9 [112] is presented in Scheme 17 [34a]. The Grignard reagent derived from 3-chloromethylfuran was coupled with the exo-methylene vinyl epoxide, prepared from 2-methyl-2-cyclohexenone, in the presence of CuCN to give allylic alcohol 113 in 62% yield, thus establishing the C-4, C-5 bond of 112. Oxidation (PCC, 89%) and treatment of the derived enone with MeCu ⋅ BF3 [63] introduced the C-6 methyl group, providing ketone 114 (70%) as a 60 :40 mixture at pro-C-7, in 62% overall yield from 113. The second electrophilic center needed for closure at C-10 was easily introduced as the enone via selenylation [64] of the kinetic enolate, followed by oxidation and elimination (H2O2, Et3N) of the selenoxide-yielding enone 115 (72%). It was discovered that triethylamine was a useful addend to the oxidation mixture as it provided a sink for the phenylseleninic acid produced in the elimination. In its absence, enone 115 was produced admixed with the desired cyclization product 116, albeit in greatly reduced yields. Cyclization of 115 was effected with HCOOH-c-C6H12, affording the desired bicyclo[4.3.1]decanone 116 in excellent yield (79%) as a 60:40 mixture at C-7. A methyl equivalent and double bond were simultaneously introduced via a Wittig olefination of 116 using the conditions of Conia and Limasset [65] (Ph3P⫹CH3 I⫺, K-t-amylate) to give 117 (80%) as a 60:40 mixture at C-7. After considerable experimentation [34a] it was found that exposure of 117 to a solution of p-toluenesulfonic acid in refluxing benzene

Scheme 16

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Retrosynthetic analysis of nakafuran 9 (112).

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Scheme 17 The synthesis of (⫾)-Nakafuran-9 (112).

for 16 min provided a 95:5 mixture of nakafuran-9 (112) and ∆8,9-isonakafuran9 (118) in 80% yield.

V.

ALKALOID SYNTHESIS VIA FURAN-TERMINATED CATIONIC ␲-CYCLIZATION

The successful investigations involving the use of furan-terminated cyclizations in the synthesis of carbocycles (discussed previously) led to the suggestion of creating azacycles under the same, or similar, conditions. Alkaloid natural products should be easily and efficiently accessed through a furan-terminated cationic π-cyclization by employing a nitrogen-containing cationic initiator such as an Nacyliminium ion. Much of the chemistry surrounding the synthesis, reactivity, and utility of the acyliminium ion has been very well documented in the pioneering work of Speckamp and coworkers [66]. On the bases of this work and the work of others, such as Chamberlin [67a] and Evans [67b], it was assumed that the ease of synthesis, and high reactivity of acyliminium ions were a good match for a furan terminator. A variety of linearly fused, spirocyclic, and bridged azacycles could be synthesized by simply altering the placement of the furan tether

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Figure 5 An approach to alkaloid synthesis via furan-terminated cationic-π-cyclization.

on the N-acyliminium ion precursor as illustrated in Fig. 5. The first foray into alkaloid synthesis via N-acyliminium ion furan-terminated cyclization [68] was directed to the preparation of the spirolactam intermediate 119 in Kishi’s synthesis of perhydrohistrionicotoxin (120) [69].

The Kishi spiropiperidine 119 [69] was approached by the spirocyclic disconnection presented in Fig. 5. For the present application it was necessary to alter the substrate from the illustrated carbinolamide-3-substituted furan to a 3-(5ethyl-2-furyl)propyl carbinolamide in order to provide a proper ketone–side chain location. In the event (Scheme 18), the Grignard reagent prepared from 1-bromo3-(5-ethyl-2-furyl)propane was added to the iodomagnesium salt of glutarimide [67b] to furnish the related carbinolamide 121 in excellent crude yield. Without

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Scheme 18 A formal total synthesis of (⫾)-perhydrohistrionicotoxin 120.

purification 121 was cyclized (HCOOH/c-C6H12) to afford spiropiperidine 122 in 72% overall yield. The furan ring was oxidatively cleaved with MCPBA [24,33], yielding 123 (70%) after reduction (H2-Pd/C; EtOAc, aq. HOAc) of the relatively unstable ene-dione. The completion of the synthesis requires dione differentiation and removal of the unwanted side chain oxygen. Ketone 123 was treated under Noyori kinetic ketalization conditions [70] (TMSS(CH2)2STMS, TMSOTf ) to lead, almost exclusively, to the side chain thioketal 124 (67%, ⱖ98 :2). Reductive cleavage was accomplished with Raney nickel in refluxing ethanol, leading to the target spiropiperidine 119 in 78% yield. The chemistry of Scheme 18 describes a concise (6 steps, 26%) formal total synthesis of perhydrohistrionicotoxin [120] and illustrates the utility of the furanterminated cationic π-cyclization protocol in alkaloid synthesis. The application of furan-terminated cationic π-cyclizations to the synthesis of linearly fused- and bridged-ring-containing alkaloids is currently under study. These results will reported in due course.

VI.

CONCLUSIONS

The furan-terminated cationic π-cyclization has been developed as a useful reaction sequence for the construction of appropriate linearly fused-, spirocyclic-, and bridged-ring-containing terpenoids. Through the careful selection of nonBrønsted acidic reaction conditions, high yields of cyclized products have been realized with epoxides, allylic alcohols, enones, and thioesters as initiating func-

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tions. Ring size restrictions, for the forming cycle, have been discovered, and the cyclization has been demonstrated to be somewhat sensitive to the position of attachment of the furan to the tether (3 vs. 2). Asymmetry transfer from the cyclizing chain has been realized, and the use of exogenous chirality to induce asymmetry has been demonstrated. Finally, alkaloid synthesis, through the agency of an acyliminium ion initiator, has been demonstrated in a limited sense (spirocycles), with much work remaining to bring this area to the level of understanding enjoyed in the terpene field.

ACKNOWLEDGMENT I wish to thank those people who made all this possible; the grad students, postdocs, and coworkers at Michigan State University, the Upjohn Co., and Pharmacia and Upjohn Inc., who did the overwhelming majority of the work mentioned. Sincere thanks go to Yu-Hwey Chuang, Mark Collins, Melissa Deaton, Lisa Dixon, Dave Head, Paul Herrinton, Mark McMills, Tim Parker, and Ed Robinson. Thanks also to the Camille and Henry Dreyfus Foundation, the National Institutes of Health, and the Upjohn Postdoctoral Research Scholar Program for financial support.

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24. (a) S. P. Tanis, Y.-H. Chuang, and D. B. Head, Tetrahedron Lett., 26: 6147 (1985). (b) S. P. Tanis, Y.-H. Chuang, and D. B. Head, J. Org. Chem., 53: 4929 (1988). 25. (a) Stoichiometric asymmetric epoxidation: T. Katsuki and K. B. Sharpless, J. Am. Chem. Soc., 102: 5974 (1980). (b) Catalytic asymmetric epoxidation: R. M. Hanson and K. B. Sharpless, J. Org. Chem., 51: 1922 (1986). 26. K. B. Sharpless and M. A. Umbreit, J. Am. Chem. Soc., 99: 5526 (1977). 27. J. A. Dale, D. L. Dull, and H. S. Mosher, J. Org. Chem., 34: 2543 (1969). 28. G. Stork, P. A. Grieco, and M. Gregson, Org. Synth., 54: 68 (1975). 29. A. Mancuso, and D. Swern, J. Org. Chem., 43: 2480 (1978). 30. For some examples of diastereoselective reductions of 3-alkoxy- and 3-hydroxyketones see: (a) D. A. Evans and K. T. Chapman, Tetrahedron Lett., 27: 5939 (1986). (b) T. Oishi, New Synthetic Methodology and Functionally Interesting Compounds. Proceedings of the 3rd International Kyoto Conference on New Aspects of Organic Chemistry (Z-i. Yoshida, ed.), Elsevier, Amsterdam, p. 81 (1986). (c) K. Narasaka and F.-C. Pai, Tetrahedron, 40: 2233 (1984). 31. J. P. Kutney, N. Abduraham, C. Gletsos, P. LeQuesne, E. Piers, and I. Vlattas, J. Am. Chem. Soc., 92: 1727 (1970). 32. R. H. Mitchell, Y. H. Lai, and R. V. Williams, J. Org. Chem., 44: 4733 (1979). 33. P. D. Williams and E. LeGoff, J. Org. Chem., 46: 4143 (1981). (b) P. D. Williams, and E. LeGoff, Tetrahedron Lett., 26: 1367 (1985). 34. (a) S. P. Tanis, and P. M. Herrinton, J. Org. Chem., 50: 3988 (1985). (b) J. P. Marino and H. Abe, J. Am. Chem. Soc., 103: 2907 (1981). (c) E. Ziegler and M. A. Cady, J. Org. Chem., 46: 122 (1981). 35. For a synthesis and reactions of 65 when n ⫽ 1 see: S. Chang, R. M. Heid, and E. N. Jacobsen, Tetrahedron Lett., 35: 669 (1994). 36. For a synthesis and reactions of 66 when n ⫽ 1 see ref. 34a, and S. P. Tanis, M. C. McMills, and P. M. Herrinton, J. Org. Chem., 50: 5887 (1985). 37. For syntheses and reactions of 67 when n ⫽ 1 see: (a) J. P. Marino, and J. C. Jaen, J. Am. Chem. Soc., 104: 3165 (1982). (b) P. A. Wender, J. M. Erhardt, and L. Letendre, J. Am. Chem. Soc., 103: 2114 (1981). 38. See for example: ref. 34a, and E.-J. Brunke, J.-J. Hammerschmidt, and H. Struwe, Tetrahedron, 37: 1033 (1981). 39. (a) N. H. Andersen and H. Uh, Tetrahedron Lett., 2079 (1973). (b) N. H. Andersen, D. W. Ladner, and A. L. Moore, Synth. Commun., 8: 437 (1978). 40. J. A. Marshall, and P. G. M. Wuts, J. Org. Chem., 42: 1794 (1977). 41. (a) J. L. Cooper, and K. E. Harding, Tetrahedron Lett., 3321 (1977). (b) K. E. Harding, J. L. Cooper, P. M. Puckett, and J. D. Ryan, J. Org. Chem., 43: 4363 (1978). 42. (a) W. Herz, S. Rajappa, S. K. Roy, J. J. Schmid, and R. N. Mirrington, Tetrahedron, 22: 1907 (1966). (b) G. R. Pettit, C. H. Herald, D. Gust, D. L. Herald, and L. D. Vanell, J. Org. Chem., 43: 1092 (1978). 43. E. S. Glazer, R. Knorr, and J. D. Roberts J. Am. Chem. Soc., 94: 6026 (1972). 44. S. P. Tanis, M. C. McMills, T. A. Scahill, and D. A. Kloosterman, Tetrahedron Lett., 31: 1977 (1990). 45. (a) T. Mukaiyama, M. Tamura, and S. Kobayashi, Chemistry Lett., 743 (1987). (b) T. Mukaiyama, M. Tamura, and S. Kobayashi, Chemistry Lett., 1817 (1986). (c) T.

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9 Chemistry and Biology of Semisynthetic Avermectins Timothy A. Blizzard Merck Research Laboratories, Rahway, New Jersey

I. INTRODUCTION The avermectins are a family of macrocyclic natural products discovered at Merck in the late 1970s with useful anthelmintic and pesticidal activities [1–3]. After their discovery, a medicinal chemistry program was initiated to discover analogues with improved activity against a broader spectrum of parasites, an improved safety profile, and/or increased stability. The primary fermentation product, avermectin B1 (1) (Fig. 1), was isolated from the fermentation of Streptomyces avermitilis as a mixture of two components. The major a component (ⱖ80%) contains a sec-butyl side chain at C-25, whereas the minor b component (ⱕ20%) has an isopropyl group at this carbon. Although the components can be separated by high-performance liquid chromatography (HPLC) [4], in production this is not normally done since the a and b isomers have essentially identical biological activities. Thus, all compounds referred to in this article are actually mixtures of a and b isomers, but for the sake of clarity only the a component is shown [i.e., the structure (1) shown for avermectin B1 is actually the structure of avermectin B1a]. Avermectin B1, also known as abamectin, is currently used as an agricultural pesticide. Ivermectin (2) [5], the semisynthetic 22,23-dihydro analogue of avermectin B1, is a widely used anthelmintic agent in animal health [1]. Ivermectin also plays an important role in efforts to control onchocerciasis, a serious human health problem in some parts of the world. Since the avermectins not only are complex natural products with interesting structures but also have major economic importance, there has been considerable interest in their chemical modification and total synthesis [6]. The field of avermectin chemistry has

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Figure 1 Structures of avermectin B1 (1) and ivermectin (2).

been widely reviewed [1–4,6] and will not be comprehensively reviewed again here. This article will describe only work in the late stages of the medicinal chemistry program with which I was involved, with reference to other work only as necessary to place our work in context.

II. CHEMISTRY A.

Avermectin Epoxides

1. Avermectin B1 8,9-Epoxide Avermectin B1 8,9-epoxide (3) (Fig. 2) is a highly active anthelmintic that was synthesized at Merck as a potential agricultural pesticide [7]. Although 3 is more stable than 1 because of the elimination of the photosensitive diene, the presence of a potentially reactive epoxide functionality still raised concerns about the chemical stability of 3. In order to address these concerns we explored the reactivity of 3 with various nucleophiles. Not surprisingly, we found that the epoxide reacted readily with strong nucleophiles such as thiophenoxide [8]. This result was not of great importance by itself; however, the fact that the thiophenol adduct 4 crystallized from methanol turned out to be highly significant. When we began our studies on 3, the epoxide stereochemistry had not been unambiguously established. X-ray diffraction analysis of the crystals of 4 allowed the definitive assignment of the stereochemistry of 4 as 8S,9R. Since 4 is derived from 3 with inversion at C-9, then 3 must be the α-epoxide. Although 4 proved to be useful for structural determination, epoxide-opened analogues of 3, including 4, were uniformly inactive (see Section III.B for a discussion of bioactivity).

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Figure 2 Epoxide-opening reactions of avermectin B1 8,9-epoxide (3). (Source: Adapted from Ref. 8.)

2. Avermectin B1 3,4-Epoxide Although disappointed by the inactivity of epoxide-opened derivatives of 3, we were encouraged by the activity of intact 3 and decided to explore the possibility of preparing additional avermectin epoxides. Since 3 had been prepared by utilizing a hydroxyl-directed epoxidation, we felt that it might be possible to apply similar chemistry to epoxidize the 3,4-double bond of 1. In fact, once the C-7 hydroxyl group was blocked as the trimethylsilyl (TMS) ether 6 (Fig. 3) by persilylation of 1 with bis(trimethylsily)trifluoroacetamide (BSTFA) followed by selective hydrolysis of the secondary TMS ethers, the C-5 hydroxyl group efficiently directed epoxidation of the 3,4 double bond. The β-3,4-epoxide 7 was thus readily prepared [9]. As observed with the 8,9-epoxide, the 3,4-epoxide 7 also reacted readily with strong nucleophiles to afford inactive ring-opened adducts (e.g., 8). In another par-

Figure 3 Synthesis of avermectin B1 3,4-epoxide (7) and reaction of 7 with thiophenol. (Source: Adapted from Ref. 9.)

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allel with the 8,9-epoxide series, we were able to use the epoxide-opened analogue 8 to determine the stereochemistry of the epoxide, this time by careful analysis of nuclear magnetic resonance (NMR) coupling constants [9]. In the 1H NMR spectrum of 8, the coupling constant J2,3 was found to be ⬇4 Hz, consistent with an α orientation of the phenylthiolate group at C-3 (the corresponding isomer with a β phenylthio substituent at C-3 should have J2,3 ⬇ 9 Hz). Since 8 is derived from 7 with inversion at C-3, 7 must be the β-epoxide, a conclusion that is also consistent with the mechanism of the hydroxyl-directed epoxidation used to prepare 7. Although both epoxides turned out to be somewhat of a disappointment as starting materials for the synthesis of active avermectin analogues, the epoxide work did result in some interesting chemistry. One especially interesting rearrangement of the 8,9-oxide is discussed in the next section. B.

Reactions of Natural Avermectins

1. Fragmentations and Rearrangements Some of the most interesting avermectin chemistry involved unexpected fragmentation or rearrangement reactions. An example from earlier work with avermectins is shown in Fig. 4 [10]. Treatment of unprotected avermectin B1 8,9-oxide 3 with p-toluenesulfonic acid in wet THF afforded the expected diol 5 (Fig. 2). However, under the same reaction conditions, the corresponding TMS ether 9 afforded the unexpected rearrangement product 10 (60%–70%). Since the two secondary TMS ethers of 9 are cleaved almost immediately under these condi-

Figure 4 Acid-catalyzed rearrangement of avermectin B1 8,9-epoxide 9. (Source: Adapted from Ref. 10.)

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tions whereas the more hindered tertiary silyl ether at C-7 survives for several hours, it is likely that the C-7 OTMS group somehow facilitates the rearrangement, possibly via silyl transfer to the epoxide oxygen. In addition to the epoxide work described, we were interested in exploring the chemistry of avermectin analogues wherein the macrocyclic lactone had been cleaved. Treatment of avermectin B1 (1) with methanolic diazabicycloundecene (DBU) resulted in an unexpected fragmentation reaction (Fig. 5) [10]. Along with the expected ring-opened product 11, we isolated a small amount of fragmentation product 12 in which a significant portion of the molecule had been removed. Unfortunately, the destruction occurred in that part of the molecule that is most important for biological activity and 12 was totally inactive. Another novel fragmentation reaction was observed during our effort to synthesize 13-epi-avermectins [11]. Glycosylation of the protected 13-epi-avermectin B2a aglycone 13, prepared from avermectin B2a aglycone by inversion at C-13 [12], using conditions similar to those employed by Nicolaou et al. in their partial synthesis of avermectin B1 [13] resulted in only low yields of the desired glycoside. We hypothesized that the free hydroxyl group at C-7 was interfering in the glycosylation reaction so we prepared the 5,7,23-tris-protected aglycone 14 by persilylation of 13 with BSTFA followed by selective hydrolysis of the secondary TMS ether at C-13 [11]. We were pleased to find that glycosylation of 14 did, in fact, cleanly and rapidly afford one major product under the same reaction conditions. However, the product was not the desired glycoside, but was instead aldehyde 15 (Fig. 6). This unexpected fragmentation product is probably formed by a vinylogous fragmentation/elimination reaction similar to the Grob fragmentation [14–16]. Additional experiments established that the glycosyl do-

Figure 5 Base-catalyzed fragmentation of avermectin B1 (1). (Source: Adapted from Ref. 10.)

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Figure 6 Fragmentation of 13-epi-avermectin B2 aglycones 13 and 14. (Source: Adapted from Ref. 11.)

nor, in this case a glycosyl fluoride, was not involved in the reaction. However, both SnCl2 and AgClO4 were required, for reasons that we do not currently understand. Furthermore, the reaction was independent of C-13 stereochemistry, but was greatly accelerated by the presence of a TMS ether at C-7, possibly as a result of relief of steric strain. It was subsequently shown that 13 also produced 15 under the glycosidation conditions. Recall that we had previously observed a similar accelerating effect of a 7-OTMS ether in the rearrangement reaction of avermectin B1-8,9-oxide derivative 9 described. 2. Disaccharide Excision Although the fragmentation and rearrangement reactions discussed previously are potentially useful as sources of intermediates for avermectin synthesis, we did not utilize them for that purpose. We did, however, apply the concept of using a natural avermectin as a source of synthetic intermediates when we required a source of the avermectin disaccharide. By modifying the multistep procedure developed by Hanessian et al. [17], we were able to prepare the desired disaccharide 17 successfully from bis-silylated avermectin B1 (16) on a multigram scale in 57% yield via a one-pot procedure (Fig. 7) [18]. Reaction of disaccharide 17 with diethylaminosulfur trifluoride (DAST) provided the glycosyl fluoride 18. This convenient preparation of large quantities of the disaccharide proved to be essential in our later synthetic studies, as described later. C.

Avermectins with Modified Glycosidic Linkages

The availability of the avermectin disaccharide enabled us to pursue several interesting avenues of research that had previously been inaccessible. For example,

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Figure 7 Preparation of the avermectin disaccharide 18. (Source: Adapted from Ref. 18.)

we were able to study the effect of modifying the linkage between the avermectin aglycone and the disaccharide. Starting with ivermectin aglycone 19, we introduced a variety of spaces (e.g., the ethylene glycol spacer shown in Fig. 8), then attached the disaccharide to the new spacer to afford, after deprotection, ‘‘spacermectin’’ analogues such as 23 and 24 [19]. Interestingly, some of the ‘‘spacermectins’’ had activities approaching those of the natural compounds.

D.

Stereoisomers of Natural Avermectins

1. 13-epi-Avermectins In addition to allowing synthesis of the ‘‘spacermectins,’’ the efficient disaccharide preparation enabled us to undertake the considerably more challenging synthesis of 13-epi-avermectins. It was known that epimerization at C-2 of an avermectin results in substantial loss of activity [1], so we were interested in exploring the effect of epimerization at other stereocenters. We therefore prepared several 13-epi-avermectins, including 13-epi-avermectin B1 (30) and 1′,13-bis-epi-avermectin B1 (31), by inversion at C-13 of the corresponding aglycone followed by glycosylation with the avermectin disaccharide [12]. Although the synthesis of these analogues, outlined in Fig. 9, was relatively straightforward, it was during this work that we discovered the interesting fragmentation reaction discussed earlier. We were pleased to find that the 13-epi-avermectins not only were as

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Figure 8 Synthesis of ivermectin analogues, the ‘‘spacermectins’’ 23 and 24, with a spacer between the aglycone and the disaccharide. (Source: Adapted from Ref. 19.)

active as the natural isomers, but also had a substantially better therapeutic index, which was due to their reduced mammalian toxicity [12]. 2. 19-epi-Avermectin B1 (36) Encouraged by the outstanding activity of the 13-epi-avermectins,we decided to undertake the synthesis of 19-epi-avermectin B1 (36) (Fig. 10). The synthesis of 36 was complicated somewhat by the propensity of the avermectin C-3,4 double bond to move into conjugation with the lactone carbonyl under basic conditions. Thus, although formation of the seco-acid 32 and inversion of the C-19 stereochemistry via Mitsunobu lactonization to give 33 are relatively straightforward, reestablishment of the C-2 stereocenter is more difficult. After considerable experimentation, we were able to accomplish the deconjugation successfully by

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Figure 9 Synthesis of 13-epi-avermectin B1 (30). (Source: Adapted from Ref. 12.)

using a procedure based on methods previously developed by Hanessian et al. [20,21] and refined by Danishefsky et al. [22] in their avermectin total-synthesis studies. Thus, quenching of the vinylogous enolate formed upon deprotonation of 7-OTMS-derivative 34 with the highly hindered pivalic acid gave the deconjugated penultimate intermediate 35. Subsequent deprotection of 35 completed our synthesis of 19-epi-avermectin B1 (36) [23]. A synthesis of the related 19-epiavermectin A1 via a very similar route was independently accomplished by Hanessian et al. [24]. Unfortunately, 36 turned out to be substantially less active than avermectin B1.

III. BIOLOGY Although much interesting chemistry was discovered during this avermectin project, our primary interest was in the biological activity of the novel avermectin analogues that we prepared. The next section describes some of our efforts to measure biological activity and discusses the interesting structure–activity relationships that were uncovered.

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Figure 10

A.

Synthesis of 19-epi-avermectin B1 (36). (Source: Adapted from Ref. 23.)

Development of a Brine Shrimp Assay for Avermectin Analogues

The success of any medicinal chemistry project depends on the timely availability of biological data for each new analogue prepared by the medicinal chemists. The avermectin project was not exceptional in this regard. What was exceptional about this project was that we were able to develop and implement an assay that could be performed by chemists with no biological training or facilities. Since the use of brine shrimp larvae as an assay for general toxicity is well known [25–28], it seemed likely that we might be able to use a similar assay to evaluate new avermectin analogues. A moderate amount of experimentation soon led to a simple, highly reproducible assay that employed brine shrimp larvae to evaluate new avermectin analogues rapidly [29]. As we had hoped, the assay was generally predictive of activity in our more elaborate and time-consuming biological assays and thus provided valuable feedback on a very timely basis. This proved most

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helpful in developing structure–activity relationships that greatly aided our selection of subsequent synthetic targets.

B.

Structure–Activity Relationships of Semisynthetic Avermectins

The brine shrimp assay results for the avermectin derivatives discussed in this article are summarized in Table 1. The data in Table 1 allow several conclusions to be drawn regarding which structural features of the avermectins are necessary for optimal biological activity. For example, epimerization of the stereocenter at C-13 results in compound 30, which is nearly as active as avermectin B1 (1) (compare items 1 and 10). However, epimerization at C-2 (Item 13), C-19 (Item 12), or C-1′ (compare Items 10 and 11) leads to analogues that are much less active. Also, epoxidation of the double bonds at C-3,4 (Item 6) or C-8,9 (Item 3) results in little, if any, loss of activity, but opening the resulting epoxide with a nucleophile causes substantial loss of activity. The complete inactivity of 7OTMS-avermectin B1 (6) (Item 5) demonstrates the importance of the C-7 hydroxyl group, the moderate activity of the ‘‘spacermectin’’ 23 (Item 9) suggests some flexibility in the exact spatial orientation of the disaccharide. The brine shrimp assay thus allowed us quickly to delineate useful structure–activity rela-

Table 1 Activity of Avermectins vs. Brine Shrimp Larvae Compound

IC100 (µg/mL)a

Avermectin B1 (1) Ivermectin (2) Avermectin B1 8,9-epoxide (3) 8-SC6H5,-9-OH-8,9-H2-Avermectin B1 (4) 7-OTMS-Avermectin B1 (6) Avermectin B1 3,4-epoxide (7) 3-SC6H5,4-OH-3,4-H2-Avermectin B1 (8) Avermectin B2 fragmentation product (15) Spacermectin (23) 13-epi-Avermectin B1 (30) 1′,13-bis-epi-Avermectin B1 (31) 19-epi-Avermectin B1 (36) 2-epi-Avermectin B1

335 430 650 ⬎55,500 ⬎55,500 430 ⬎55,500 ⬎55,500 1,730 870 10,415 13,900 ⬎55,500

Item 1 2 3 4 5 6 7 8 9 10 11 12 13 a

IC100 ⫽ minimum concentration at which 100% of brine shrimp are immobile. OTMS ⫽ o-trimethylsilyl.

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tionships that were, in general, confirmed by activity in our more traditional assays.

IV.

CONCLUSIONS

The most enjoyable aspects of the avermectin project were the rich diversity of the chemistry and the multidisciplinary nature of the project, which incorporated elements of both chemistry and biology. In this respect, the avermectin project at Merck resembled many of the projects in Professor Nakanishi’s lab at Columbia. In both cases, effective collaboration with scientists in other fields was not only critical to success, but also a key element in making the project a lot more fun for all concerned.

ACKNOWLEDGMENTS I would like to thank Professor Koji Nakanishi for a highly educational and very enjoyable postdoctoral experience. My years in his laboratory were the best possible preparation for the interdisciplinary nature of research in the pharmaceutical industry. I am also indebted to the many excellent scientists at Merck who were involved in the avermectin project, especially Dr. Helmut Mrozik and Ms. Gaye Margiatto.

REFERENCES 1. 2. 3.

4.

5.

6. 7. 8.

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W. C. Campbell, ed., Ivermectin and Abamectin, Springer-Verlag, New York (1989). H. G. Davies and R. H. Green, Nat. Prod. Rept., 3: 87 (1986). T. Blizzard, M. H. Fisher, H. Mrozik, and T. L. Shih, in Recent Progress in the Chemical Synthesis of Antibiotics (G. Lukacs and M. Ohno, eds.), Springer-Verlag, Berlin, pp. 65–102 (1990). T. W. Miller and V. P. Gullo, in Natural Products Isolation—Journal of Chromatography Library Series, Vol. 43 (G. H. Wagman and R. Cooper, eds.) Elsevier, Amsterdam, chapter 9, pp. 347–376 (1989). J. C. Chabala, H. Mrozik, R. L. Tolman, P. Eskola, A. Lusi, L. Peterson, M. Woods, M. H. Fisher, W. C. Campbell, J. R. Egerton, and D. A Ostlind, J. Med. Chem., 23: 1134 (1980). T. Blizzard, Org. Prep. Proc. Int., 26: 617 (1994). H. Mrozik, U.S. Patent 4,530,921 (1985). T. A. Blizzard, H. Mrozik, and M. H. Fisher, Bioorg. Med. Chem. Lett., 3: 2093 (1993).

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9. T. A. Blizzard, H. Mrozik, F. A. Preiser, and M. H. Fisher, Tetrahedron Lett., 31: 4965 (1990). 10. T. A. Blizzard, H. Mrozik, and M. H. Fisher, Tetrahedron Lett., 29: 3163 (1988). 11. T. A. Blizzard, G. Margiatto, H. Mrozik, and M. H. Fisher, J. Org. Chem., 58: 3201 (1993). 12. T. A. Blizzard, G. Margiatto, H. Mrozik, W. L. Shoop, R. A. Frankshun, and M. H. Fisher, J. Med. Chem., 35: 3873 (1992). 13. K. C. Nicolaou, R. E. Dolle, D. P. Papahatjis, and J. L. Randall, J. Am. Chem. Soc., 106: 4189 (1984). 14. C. A. Grob, Angew. Chem. Int. Ed. Eng., 8: 535 (1969). 15. M. Ochiai, T. Ukita, S. Iwaki, Y. Nagao, and E. Fujita, J. Org. Chem., 54: 4832 (1989). 16. H. B. Henbest and B. B. Millward, J. Chem. Soc., 3575 (1960). 17. S. Hanessian, A. Ugolini, P. J. Hodges, and D. Dube, Tetrahedron Lett., 27: 2699 (1986). 18. T. Blizzard, G. Marino, H. Mrozik, and M. H. Fisher, J. Org. Chem., 54: 1756 (1989). 19. T. Blizzard, G. Margiatto, B. Linn, H. Mrozik, and M. H. Fisher, Bioorg. Med. Chem. Lett., 1: 369 (1991). 20. S. Hanessian, A. Ugolini, D. Dube, P. J. Hodges, and C. Andre, J. Am. Chem. Soc., 108: 2776 (1986). 21. S. Hanessian, D. Dube, and P. J. Hodges, J. Am. Chem. Soc., 109: 7063 (1987). 22. S. J. Danishefsky, D. M. Armistead, F. E. Wincott, H. G. Selnick, and R. Hungate, J. Am. Chem. Soc., 111: 2967 (1989). 23. T. Blizzard, L. Bostrom, G. Margiatto, H. Mrozik, and M. Fisher, Tetrahedron Lett., 32: 2723 (1991). 24. S. Hanessian and P. Chemla, Tetrahedron Lett., 32: 2719 (1991). 25. A. S. Michael, C. G. Thompson, and M. Abramovitz, Science, 123: 464 (1956). 26. J. Harwig and P. M. Scott, Appl. Microbiol., 21: 1011 (1971). 27. M. G. Prior, Can. J. Comp. Med., 43: 352 (1979). 28. B. N. Meyer, N. R. Ferrigni, J. E. Putnam, L. B. Jacobsen, D. E. Nichols, and J. L. McLaughlin, Planta Med., 45: 31 (1982). 29. T. A. Blizzard, C. L. Ruby, H. Mrozik, F. A. Preiser, and M. H. Fisher, J. Antibiotics, 42: 1304 (1989).

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10 Chemical and Biological Approaches to Molecular Diversity: Applications to Drug Discovery Harold V. Meyers New Chemical Entities, Inc., Framingham, Massachusetts

I. INTRODUCTION Despite recent scientific advances in the pharmaceutical industry, drug discovery costs have steadily increased while the rate of introduction of useful pharmaceuticals has decreased. The average cost for introducing a new drug entity into the marketplace has recently been estimated at more than $400 million, nearly a third of which goes to the preclinical tasks of discovery and optimization of lead chemical compounds. The synthetic preparation of each novel molecule in the traditional serial fashion alone has been estimated to cost between $5,000 and $10,000. From the economic viewpoint, there is clearly room for significant improvement in the discovery process. An exciting advance in the industry, the ability to screen hundreds of thousands to millions of compounds in relatively short time frames, has been made possible with the recent advent of automated, high-throughput screening (HTS) technologies. Such capacity often exceeds the supply of available compounds. Compound libraries typically used in mass screening consist of either historical collections synthesized in drug programs, or natural products collections. Although historical collections can number in the hundreds of thousands of compounds at large pharmaceutical firms, they often have limited structural diversity (e.g., many β-lactams, steroids), whereas natural product collections can be disadvantageous because of their structural complexity, which renders making synthetic analogues to generate drug candidates potentially very difficult.

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During the last decade a new source of compounds has arisen with the advent of combinatorial chemistry, which has the effect of providing large numbers of compounds for biological testing at a greatly reduced cost. In conjunction with HTS, combinatorial chemistry can enhance the speed to market of a drug by reducing development timelines. For the development of novel and proprietary drug candidates, combinatorial chemistry enables many unique chemical structures to be synthesized rapidly, thereby allowing for broader patent coverage around lead compounds. This can diminish the reliance on ‘‘me too’’ drug strategies of many companies, whereby clinical compounds are derived from typically conservative structural variations of competitors’ compounds having narrow patent coverage. As the marketplace rewards innovation and places a premium on novel, efficacious drugs, being the first to market with a novel compound can provide enormous financial rewards. Furthermore, by substantially reducing the period for drug candidate selection, the useful patent life of a product can be significantly extended. For a big-selling drug (e.g., $1 billion/year) this can translate into hundreds of millions of dollars of additional sales [1]. The bottom line is, combinatorial chemistry accelerates the drug discovery process in a more economical manner. This chapter will provide an overview of combinatorial chemistry methodologies as applied to drug discovery, concluding with an assessment of future directions.

II. SYNTHESIS AND SCREENING OF COMBINATORIAL LIBRARIES This section discusses some of the key biological and chemical methodologies available for the creation of molecular diversity. In particular, the molecular biological generation of peptide and protein libraries by phage display techniques is discussed; then an overview of chemical strategies and methods developed for the combinatorial synthesis of peptide and small molecule libraries used in drug discovery and lead optimization processes is presented. Several good review articles describing molecular biological and chemical methodologies are available for a more comprehensive discussion of these topics [2–6]. A.

Biological Approaches

Several techniques for creating large libraries of filamentous phage clones displaying unique peptides or proteins on their protein coat surface have been developed [7–11]. By inserting fully random cassettes of synthetic oligonucleotides into targeted loci, tens of millions of mutant strains can be isolated [12–15]. Each unique peptide is structurally encoded by its respective single-stranded viral deoxyribonucleic acid (DNA) sequence and can be expressed as copies as a fusion

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peptide with either the minor-coat protein (pIII) or the major-coat protein (pVIII), respectively. Phage display methods have been widely used in the area of epitope mapping, peptide drug discovery, and protein engineering to study structure/function relationships. As phage libraries are large enough to preclude analysis of individual mutants, population selection schemes are necessary. In the ‘‘colony-lift’’ technique, a polymeric membrane is placed on a high-density clone colony and lifted off to adsorb the expressed macromolecules that mirror their location on the growth colony. A labeled protein or antibody that binds to the membrane then reveals the location of the clone, which can be further analyzed [16]. An affinity purification technique termed biopanning [8] allows the affinity selection of peptides from phage display libraries by biological targets (ligates). A biotinylated ligate is incubated with the phage library in solution, and then a ‘‘panning’’ step captures the phage-bound ligate with a streptavidin-coated polystyrene Petri dish. Alternatively the library is panned using an immobilized ligate that has been coated directly in the Petri dish [11]. The unbound phage is washed away, and the bound phage can be eluted and propagated in Escherichia coli for DNA sequencing to reveal the identity of active peptides. Often several iterations of biopanning are performed in a selection/amplification process to find high-affinity peptides to the target(s) of interest. An application of phage display peptide libraries, ‘‘epitope libraries,’’ have been used in mapping the specificity of antibodies. Peptides that mimic antibodybinding determinants through continuous [9,10] and discontinuous epitopes [17], the latter composed of adjacent residues in a folded or conformationally dependent structure but distant in primary sequence, have been discovered. Epitope mapping enables ligands to be found for antibodies whose specificity is not known in advance. The screening of phage peptide libraries against receptors of pharmaceutical interest has been an active area of research. Conformationally constrained peptide libraries, e.g., loops derived from cysteine disulfide formation, have also been used to find higher-affinity ligands to receptors such as gpIIb/IIIa [18], as cyclic peptides provide better leads for drug development. The application of phage display libraries in the field of protein engineering has made possible the study of structure/function relationships, for example, with human growth hormone [19] and alkaline phosphatase [20]. The proper folding of protein domains on the phage surface has been demonstrated to afford functional receptors with respect to binding [19] and catalytically active enzymes [20]. The construction of numerous variants of a parental protein by phage display techniques has allowed the alteration of catalytic properties of enzymes and binding properties of proteins, as exemplified with potent protein inhibitors of human neutrophil elastase [21], through iterative affinity selection/amplification methods against selected targets (protein biopanning), an area of research that has

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come to be known as ‘‘directed evolution.’’ Applied evolution methods will undoubtedly improve industrial processes that utilize enzymes, among other things. In the antibody engineering area, the development of customized antibodies, especially completely humanized antibodies, with high affinities and selectivities holds considerable promise for their use as therapeutic agents. In a process that mimics an immune response for high-affinity antibody selection in vivo, ‘‘in vitro affinity maturation’’ utilizes the introduction of sequence variation and recombination of both light and heavy chains as well as mutagenesis in the complementarity determining regions (CDRs) to find enhanced binding with an antigen. This selection process may be repeated with an enriched antibody pool until suitable candidates are isolated [22]. Other applications include screening for catalytic and metal coordinating antibodies [23]. Limitations of phage display techniques for drug discovery purposes include the use of only naturally occurring amino acids, and little is known about how the phage environment affects molecular recognition events. These described techniques are suitable to isolate clones from cDNA or genomic libraries of up to ⬃107 clones, but are not practical for larger libraries, which are better suited for drug discovery projects (107 –109 clones). Toward that end more powerful biological peptide display methods have recently been described. One method involves the display of peptides on plasmids, an alternative scheme that links the peptide and DNA by the DNA-binding protein LacI [24]. An in vitro system that displays nascent peptides in polysome complexes has been very recently described [25]; it has the potential to create polysome libraries of 1014 –1015 members. Perhaps the biggest drawback of utilizing biological display methods for drug discovery is that if affords peptides. It is well known in the pharmaceutical industry that peptides are not generally well suited as drugs because of in vivo instability and lack of oral absorption. Furthermore, converting a peptide lead into a peptide mimic is far more difficult than identifying the peptide lead, since a general solution for designing pharmaceutically useful, orally available peptide mimics has yet to be developed [26]. These drawbacks have provided the impetus for the pharmaceutical industry to apply combinatorial chemistry methods to the synthesis of small-molecule, nonpeptidyl libraries for drug discovery. B.

Chemical Approaches

Early chemical combinatorial efforts were focused on synthesizing libraries of peptides [27], followed by the progression to peptidelike oligomeric libraries [28], and most recently to small-molecule, nonpeptidyl libraries [29,30]. The use of solid-phase chemistry, originally developed for peptide synthesis [31] and later amply demonstrated for organic synthesis [32–34], has experienced a renaissance of late spurred by combinatorial chemistry. Solid-phase synthesis precludes often

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tedious reaction workup and purifications typically performed after each synthetic step. These advantages are amplified when constructing even relatively small libraries of hundreds of compounds. In addition, reactions can be driven to completion on solid supports by employing a large excess of reagents. Hence, many combinatorial approaches rely on solid-phase chemistry, but isolation and purification techniques recently applied to parallel solution synthesis are enhancing the utility of this approach (discussed later). The key issues that differentiate the various combinatorial approaches, then, involve the use of solid-phase or solution chemistry, the synthesis of individual or mixtures of compounds, the size of the library and application of encoding strategies, the extent of automation for various syntheses, purification, and analytical tasks, and the screening of libraries, on or off solid supports. Regardless of the approach, it is desirable to have each compound in a library present in a roughly equimolar concentration for screening purposes. What follows is a description of some key combinatorial chemistry technologies and their inherent advantages and disadvantages. 1. Synthesis of Mixtures Combinatorial libraries can be synthesized and screened as mixtures. The synthesis of large mixtures of compounds (105 –108) was made possible by employing the split synthesis approach on solid supports. a. Split Synthesis (Split-Pool, Split-Mix, or Portion Mixing). Furka first demonstrated the split-mix strategy on solid supports to synthesize large mixtures of peptides in equimolar quantities rapidly [35]. This one bead/one unique peptide approach is schematically represented (Fig. 1). A solid support material is split into equal-sized portions and placed in separate reaction vessels. Each portion is coupled with excess building blocks (e.g., unique amino acid derivatives), which provide uniform coupling since competition between reactants is eliminated. The coupled resins are then pooled together into one reaction vessel for the removal of common protecting groups. The resin is partitioned again into separate pools, each containing mixtures of unique peptides, for coupling new reactants. This iterative split-couple-mix strategy theoretically enables libraries of oligomers of Xn members to be assembled, where X is the number of monomers, amino acids in this example, in the basis set and n is the length of the oligomer. A pure statistical distribution of sequences results. When screening split-pool libraries on solid supports, bioactive compounds can be physically isolated through incubation with a tagged ligate. Polymeric beads possessing peptides bound to a ligate have been identified by visual inspection, physical removal, and microsequencing of the active peptides [36]. Alternatively, the supported peptides are cleaved into solution for screening. When screening mixtures of compounds in solution from split-synthesis libraries, sev-

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Figure 1 The split-pool method: the shaded circles represent the beads (solid support) and the lettered squares are individual chemical building blocks, for example, individual amino acids.

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eral deconvolution methods for determining the structures of compounds possessing the most bioactivity have been developed. These include ‘‘iterative’’ or ‘‘recursive’’ deconvolution methods involving resynthesis of bioactive pools [37,38], methods that preclude the resynthesis of bioactive pools such as ‘‘positional scanning’’ [39] and ‘‘orthogonal’’ libraries [40], and encoded methods (discussed late). It has been demonstrated, however, that increasingly larger pool sizes (ca. 100 or more) result in higher rates of false-positive and false-negative bioactives in screens. The testing of mixtures creates the potential for compounds to interact with one another, leading to possible increased measured potencies through synergistic effects (false-positive) or decreased or antagonistic effects (false-negative) on the measured potencies as compared with those of pure compounds. Also, as the number of compounds in a library increases, the concentration of each compound typically decreases to maintain solubility of all members, and that can preclude detecting members with moderate activities. Nevertheless, split synthesis is generally considered one of the most powerful tools for lead discovery, as the large number of compounds it provides should statistically offer better chances of uncovering lead compounds against biological targets of interest. Encoding strategies. The assembly of very large libraries (e.g., 106 –108) generally results in minute quantities of compound (e.g., 10⫺9 –10⫺15 mole), necessitating an encoding scheme to identify structurally any component of interest from a biological assay. Several methods that utilize molecular tagging systems to encode structures in combinatorial libraries have been developed. In these approaches a molecular ‘‘tag’’ is attached to the solid support and used as an identifier for each monomer used in split synthesis. Hence, an orthogonal and compatible solid-phase synthesis scheme is needed for both the library members and their respective tags. One method utilizes single-stranded oligonucleotides to encode each library member, wherein a unique oligonucleotide sequence encodes each library member. Oligonucleotides have the advantage of allowing their amplification through polymerase chain reaction (PCR) for delineating their sequence by DNA sequencing methods [41–43]. The limited stability of oligonucleotides, however, can preclude the application of some chemistries and hence the accessibility of certain compound classes. In another method, natural amino acids are used to construct peptide tags for a library of peptide-based oligomers (‘‘binding’’ strands), whereby bioactive constituents in a library are decoded through Edman degradation [44]. The binding strand can be cleaved from the solid support for solution assays prior to decoding. A potential limitation of this approach is that peptide tags can potentially interact with biological targets. The use of halocarbon derivatives as molecular tags in a binary encoding scheme was more recently demonstrated; in this method each tag is photolytically cleaved and identified by electron capture capillary gas chromatography [45]. Unlike the oligonucleotide and peptide encoding approaches, which record the order of as-

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sembly of chemical building blocks in the tag sequence, the binary approach uses uniquely defined mixtures of tags to represent each building block of the binding strand at a particular synthetic step. Recent reports on the use of radiofrequency-encoded tags preclude the need for molecular tags altogether. In these methods a microchip capable of receiving, storing, or emitting radiofrequency signals is encapsulated with the synthesis resin in a polymeric vessel to record each unique synthesis site with a unique signal. Radiofrequency encoding is an easy and rapid method for decoding chemical structures of interest and does not interfere with any chemical steps in the library synthesis [46,47]. b. Mixture Synthesis. In this approach an equimolar mixture of reactants is placed in a reaction vessel with a single substrate theoretically to generate an equimolar mixture of products. This strategy was employed in an early peptide library synthesis [27]. Although this is the most direct method to construct mixtures, a limitation of this approach is that reactivity is highly dependent on the structure of reactants, so equimolarity will only be achieved if reactants have comparable reactivity with the substrate. An equimolar mixture of products can also result in principle from reacting 1 equivalent of an equimolar mixture of reactants with a substrate; however, this requires very efficient coupling reactions to take place [48]. 2. Multiple Simultaneous Synthesis (Parallel, Discrete, Array, or Spatially Separate Synthesis) The multiple simultaneous synthesis approach ideally constructs one target compound per reaction vessel, and therefore is not very practical for libraries containing more than 104 –105 members. Although more labor-intensive than split synthesis, automation can largely diminish the tedious, repetitive nature of constructing these libraries. Parallel synthesis libraries are useful tools for lead discovery, especially if they are well designed and of good quality, and are the method of choice in the drug industry for optimizing lead compound identification. Structure/activity data are more reliable on individual compounds than on pools of compounds, and the screening of single, structurally defined molecules has a proven track record in the industry. Moreover, the need for encoding strategies is precluded as micromolar quantities of each compound are available from many parallel synthesis methods, enabling routine spectroscopic techniques to be applied directly to obtain structural information. As little as 10 micromoles of each compound is sufficient to perform multiple HTS assays over a period of several years. a. Pin Technology. Geysen first demonstrated the multiple simultaneous synthesis of peptides on polyacrylic acid–grafted polyethylene pins arranged in a microtiter plate format, which allows the construction of 96 unique peptides per

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plate [49]. This technique utilizes conventional peptide synthesis methods and has been used to construct thousands of peptides within several weeks. The peptides can be screened on the pins, enabling them to be reused for additional tests, or they can be cleaved and screened in solution [50]. b. Tea Bag Method. In this version of multiple peptide synthesis, porous polypropylene containers, dubbed ‘‘tea bags,’’ are used to enclose the synthesis beads [51]. Each bag contains a unique peptide resin, which is immersed in individual solutions of activated amino acid derivatives to effect coupling reactions, while washing and deprotection steps are all carried out by pooling the bags together, then portioning the bags for subsequent coupling steps. Cleavage of the peptides from their solid support provides a unique, soluble peptide from each individual bag. This technique has been partially automated to permit the synthesis of up to 150 different peptides [52], and sufficient quantities of material can be generated for purification and complete characterization (approx. 0.5 mmol). c. Spatially Addressable, Light-Directed Synthesis. A method combining solid-phase synthesis with photolithography has been developed for the parallel synthesis of more than 100,000 discrete peptides [53]. This was accomplished by anchoring amine-blocked alkyl groups on a silicon chip with photolabile protecting groups. By manipulating a photolithographic mask, specific regions of the chip are deblocked by light, exposing the reactive amine functionality at select locations for subsequent coupling reactions with activated amino acids. The ‘‘address’’ or location of the compound on the chip, determined by the pattern of masks and sequence of reactants, reveals its structure. A limitation of this approach is that the library must be screened on the silicon chip. This requires the use of a tagged, soluble ligate, precluding certain receptors and enzyme targets. The screening of any compound on a solid support also introduces inherent uncertainty regarding the validity of molecular recognition events as compared to its soluble counterpart since the density of compounds on the support, the support itself, and other factors may perturb relevant solution conformations, binding kinetics, and other critical factors. d. Solution Phase Synthesis. Methods for parallel solution phase synthesis have recently been reported [54,55]. These methods essentially rely upon classic aqueous acid and aqueous base extractions for purification of intermediates and final products with selective unmasking of acidic and basic functional groups within the intermediates and final products to control solubility. The major limitations to performing solution phase parallel synthesis have been the isolation and purification of the products of large libraries. The use of parallel purification techniques such as liquid/liquid extraction or solid-phase extraction, however, makes solution phase parallel synthesis an attractive alternative to solid-phase synthesis. The numerous solid supported reagents and catalysts for organic syn-

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thesis that are available [56] and scavenging resins for selective removal of products or excess reagents in solution will strengthen the utility of solution phase synthesis, especially for multistep sequences. Multicomponent condensation reactions like the Ugi reaction [57] have also been demonstrated in parallel synthesis to generate relatively complex molecules [58]. The advantages of performing parallel synthesis in solution include the larger scales possible for performing reactions, which provide more of each compound. In addition, the development or utilization of functionalized resins and of cleavable linker groups, the attachment and cleavage of substrates, and any capping steps are precluded. Solution phase parallel manipulations are readily automated as well [59]. In a related process, a soluble polymer support for conducting synthesis in solution has been reported [60]. Precipitation of the polymer after each synthetic transformation for washing provides the advantages of both solution and solidphase synthesis. e. Sphinx Approach. An approach taken at Sphinx was to create large numbers of small, nonpeptidyl molecules in a parallel format as a general discovery tool. The concept of a ‘‘universal library’’ has driven our efforts in this field (discussed late). The libraries are synthesized on solid supports but are made available in solution for screening and are generated in sufficient quantities (milligrams) for use in multiple screens. The initial parallel synthesis demonstration at Sphinx involved the preparation of several hundred substituted phenols in good yields and high purity levels [61]. A simple, economical apparatus for multiple simultaneous synthesis was constructed and successfully utilized for the rapid preparation of organic molecules via multistep resin-based procedures (Fig. 2). A 96-well format was chosen,

Figure 2 Reaction plate and clamp assembly (side view).

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as used in numerous biological applications, including automated highthroughput screens. Plate synthesis was carried out on cross-linked polystyrene resin placed in sterile polypropylene deep well plates, which were modified for filtration by drilling a small hole in the bottom of each well, then placing a porous polyethylene frit into the bottom of each well. An aluminum plate clamp was made as a two-piece assembly, consisting of a solid base clamp fitted with four removable corner stainless steel studs and a frame clamp that fits atop the plate and is secured with wing nuts. A Viton gasket was utilized on the base clamp to prevent leakage of well contents. A vacuum plenum can be used to facilitate the filtration of resin washings and isolation of solution libraries. This is accomplished by removing the modified plate from the clamp assembly after the synthetic transformation and placing it atop the vacuum plenum designed to accommodate the plate. The well caps are then removed and the vacuum source connected to an outlet in the plenum floor. The vacuum plenum is deep enough to place a rack of 96 microdilution tubes beneath the plate to collect the individual product solutions. The inexpensive equipment and limited human resources required for this effort illustrate the economic and productivity advantages of this technology. Universal library concept. Biological macromolecules (e.g., receptor, enzyme, antibody) recognize binding substrates or ligands through a number of precise physicochemical interactions. These interactions can be divided into a number of different parameters: size, hydrogen bonding ability, hydrophobic interactions, dipole interactions, and others. To explore multiparameter space rapidly, the Sphinx libraries were designed to orient groups responsible for these binding interactions at unique locations in space through a scaffolding approach. A major challenge was to select a class of target molecules of sufficient generality to allow for wide structural variations, preferably utilizing well-precedented synthetic methodologies. The biphenyl scaffold met these criteria and was chosen as the basic structure for a class of target molecules that allow for facile introduction of three or four functional groups in a large number of spatial arrangements (Fig. 3). The use of aromatic templates such as the biphenyl moiety also enables the size, shape, and other properties of these molecules to be changed readily by incorporating different linker groups between the aromatic rings while varying

Figure 3 General structure for a universal library using a biphenyl scaffold.

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their substitution patterns, and derivatizing these scaffolds with a diverse set of functional groups using related sequences of chemical reactions. A large number of compounds prepared around each scaffold can thus explore unique sizes, shapes, and volumes. A collection of such libraries will represent a ‘‘universal’’ library designed to explore multiparameter space (‘‘diversity’’ space) incrementally and will result in the effective identification of chemical leads for any biological target of interest. An initial report on the solid-phase synthesis of functionalized biphenyls, which utilizes Mitsunobu chemistry to introduce functional groups via ether bond formation, has appeared recently [62]. The universal library is being generated using a double combinatorial strategy, a novel approach developed at Sphinx (Fig. 4). In this scheme functional groups are introduced onto the first scaffold building block (the first phenyl ring) in the first combinatorial cycle bound to a solid support. Then, the second scaffold building block (another phenyl ring) is covalently attached and the second round of functional group introduction takes place. The final target molecule is cleaved from the solid support to afford the desired product in solution and available for screening. The key step in the biphenyl library synthesis was a Stille coupling to attach the second phenyl scaffold, thereby forming the biphenyl structure (Fig. 5). This double combinatorial approach enables large numbers of highly functionalized, low-molecular-weight molecules to be assembled rapidly from a small collection of building blocks.

Figure 4 Synthetic strategy for the ‘‘double combinatorial’’ approach to the biphenyl library.

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Figure 5 Solid-phase Stille coupling to complete biphenyl scaffold.

III. CONCLUSIONS The field of chemical diversity generation is experiencing enormous growth within the pharmaceutical industry. Almost every major pharmaceutical and biotechnology company has initiated, collaborated on, or acquired an effort in this area. Successes have been disclosed in both the lead generation and optimization of potential drugs, and an increase in the number of clinical candidates arising from combinatorial chemistry can be expected. Improved methods are evolving for the design, synthesis, and analysis of combinatorial libraries, particularly in the realm of small organic molecules. Solution chemistry will continue to be adapted to solid supports to expand the repertoire of solid-phase reactions. Eventually one can expect that most solution reactions will have a solid-phase counterpart, and some solid-phase chemistry having no solution phase counterpart will develop. New types of solid supports, including organic polymers and inorganic materials, are being developed for organic synthesis to, among other things, increase loading capacities and product yields. In the field of natural products, recent demonstrations of combinatorial analoging may provide a resurgence for natural product screening in the pharmaceutical industry. More active research in the areas of combinatorial biosynthesis to generate diverse and structurally complex libraries, particularly through the genetic manipulation of polyketide biosynthetic pathways [63–65], can also be expected. Advances in automation and instrumentation are also rapidly evolving for the synthesis of chemical libraries. Commercial vendors now provide robotic workstations for partially or fully integrated synthesis solutions, albeit at substantial cost in some cases, but more practical and affordable robotic systems are being developed for eventual use by each and every bench chemist. Parallel, highthroughput purification and spectroscopic analysis and detection techniques (e.g., high-performance liquid chromatography–mass spectrometry [HPLC-MS], highperformance liquid chromatography–nuclear magnetic resonance [HPLC-NMR];

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see Chapter 4) under development will eventually provide high-quality, wellcharacterized libraries. A trend toward better integration of synthesis and screening, whereby both chemical and biological methods of diversity generation can be coupled directly to screening protocols, is emerging as well. Furthermore, progress will likely drive combinatorial synthesis and screening toward miniaturization. New computational methods will also provide a more complete understanding of ‘‘diversity’’ space, what it is and how it is measured, for more effective drug design. For example, the desirable physicochemical properties of known drugs, such as bioavailability, can be computationally culled to design ‘‘druglike’’ libraries of compounds. Empirically derived structure/activity data can also be used in computational analyses to optimize the properties of lead compounds more rapidly. Genetic algorithms are also being applied in library design strategies to uncover potent compounds more efficiently through iterative search and selection protocols of ‘‘virtual libraries’’ to determine which subset populations to synthesize [58,66]. As an explosion of chemical, physical, biological, and computational data is resulting from combinatorial libraries, it will be critical to manage, integrate, and analyze these data effectively and efficiently to maintain a competitive edge. Advances in the field of genomics, in which genetic information derived from high throughput sequencing and analysis (‘‘bioinformatics’’), will likely result in an enormous number of new and relevant biological targets for screening. Combinatorial libraries will be useful in not only identifying new drug candidates, but evaluating and validating the physiological relevance of new targets in a much shorter time frame than traditional discovery protocols have demonstrated. With the rapid growth in the chemical generation of molecular diversity, combinatorial libraries will significantly decrease the time and cost of discovering novel therapeutic agents for multiple disease states. The development of combinatorial chemistry methods and their application in the discovery process represents one of the most significant paradigm shifts in the pharmaceutical industry in decades and has become an important and indispensable tool.

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4. M. A. Gallop, R. W. Barrett, W. J. Dower, S. P. A. Fodor, and E. M. Gordon, J. Med. Chem., 37: 1233 (1994). 5. E. M. Gordon, R. W. Barrett, W. J. Dower, S. P. A. Fodor, and M. A. Gallop, J. Med. Chem., 37: 1385 (1994). 6. L. A. Thompson and J. A. Ellman, Chem. Rev., 96: 555 (1996). 7. G. P. Smith, Science, 228: 1315 (1985). 8. S. F. Parmley and G. P. Smith, Gene, 73: 305 (1988). 9. S. Cwirla, E. A. Peters, R. W. Barrett, and W. J. Dower, Proc. Natl. Acad. Sci. USA, 87: 6378 (1990). 10. J. K. Scott and G. P. Smith, Science, 249: 386 (1990). 11. J. J. Devlin, L. C. Panganiban, and P. E. Devlin, Science, 249: 404 (1990). 12. M. D. Matteucci and H. L. Heyneker, Nucleic Acids Res., 11: 3113 (1983). 13. J. A. Wells, M. Vasser, and D. B. Powers, Gene, 34: 315 (1985). 14. M. S. Z. Horwitz and L. A. Loeb, Proc. Natl. Acad. Sci. USA, 83: 7405 (1986). 15. J. F. Reidhar-Olsen and R. T. Sauer, Science, 241: 53 (1988). 16. R. A. Young and R. W. Davis, Science, 222: 778 (1983). 17. M. Balass, Y. Heldman, S. Cabilly, D. Givol, E. Katchalski-Katzir, and S. Fuchs, Proc. Natl. Acad. Sci. USA, 90: 10638 (1993). 18. K. T. O’Neil, R. H. Hoess, S. A. Jackson, N. S. Ramachandran, S. A. Mousa, and W. F. DeGrado, Proteins Struct. Funct. Genet., 14: 509 (1992). 19. S. Bass, R. Green, and J. A. Wells, Proteins Struct. Funct. Genet., 8: 309 (1990). 20. J. McCafferty, R. H. Jackson, and D. J. Chiswell, Prot. Eng., 4: 955 (1991). 21. B. L. Roberts, W. Markland, A. C. Ley, R. B. Kent, D. W. White, S. K. Guterman, and R. C. Lander, Proc. Natl. Acad. Sci. USA, 89: 2429 (1992). 22. E. Soderlind, A. C. Simonsson, and C. A. K. Borrebaeck, Immunol. Rev., 130: 109 (1992). 23. C. Barbas, J. Bain, D. Hoekstra, and R. Lerner, Proc. Natl. Acad. Sci. USA, 89: 4452 (1992). 24. M. G. Cull, J. F. Miller, and P. J. Schatz, Proc. Natl. Acad. Sci. USA, 89: 1865 (1992). 25. L. C. Mattheakis, R. R. Bhatt, and W. J. Dower, Proc. Natl. Acad. Sci. USA, 92: 9022 (1994). 26. A. Giannis, and T. Kolter, Angew. Chem. Int. Ed. Engl., 32: 1244 (1993). 27. H. M. Geysen, S. J. Rodda, and T. J. Mason, Mol. Immunol., 23: 709 (1986). 28. R. J. Simon, R. S. Kania, R. N. Zuckerman, V. D. Huebner, D. A. Jewell, S. Banville, S. Ng, L. Wang, S. Rosenberg, C. K. Marlowe, D. C. Spellmeyer, A. D. Frankel, D. V. Santi, F. E. Cohen, and P. A. Bartlett, Proc. Natl. Acad. Sci. USA, 89: 9367 (1992). 29. B. A. Bunin and J. A. Ellman, J. Am. Chem. Soc., 114: 10997 (1992). 30. S. H. DeWitt, J. S. Kiely, C. J. Stankovic, M. C. Schroeder, D. M. R. Cody, and M. R. Pavia, Proc. Natl. Acad. Sci. USA, 90: 6909 (1993). 31. R. B. Merrifield, J. Am. Chem. Soc., 85: 2149 (1963). 32. F. Camps, J. Cartells, and J. Pi, Anales Quim., 70: 848 (1974). 33. H. Rapoport and J. I. Crowley, Acc. Chem. Res., 9: 135 (1976). 34. C. C. Leznoff, Acc. Chem. Res., 11: 327 (1978).

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A. Furka, F. Sebestyen, M. Asgedom, and G. Dibo, Int. J. Pept. Protein Res., 37: 487 (1991). K. S. Lam, S. E. Salmon, E. M. Hersh, V. J. Hruby, W. M. Kazmierski, and R. J. Knapp, Nature, 354: 82 (1991). C. T. Dooley, N. N. Chung, B. C. Wilkes, P. W. Schiller, J. M. Bidlack, G. W. Pasternak, and R. A. Houghten, Science, 266: 2019 (1994). E. Erb, K. D. Janda, and S. Brenner, Proc. Natl. Acad. Sci. USA, 91: 11422 (1994). C. T. Dooley and R. A. Houghten, Life Sci., 52: 1509 (1993). B. Deprez, X. Williard, L. Bourel, H. Coste, F. Hyafil, and A. Tartar, J. Am. Chem. Soc., 117: 5405 (1995). S. Brenner, and R. A. Lerner, Proc. Natl Acad. Sci. USA, 89: 5381 (1992). J. Nielsen, S. Brenner, and K. D. Janda, J. Am. Chem. Soc., 115: 9812 (1993). M. C. Needels, D. G. Jones, E. H. Tate, G. L. Heinkel, L. M. Kochersperger, W. J. Dower, R. W. Barrett, and M. A. Gallop, Proc. Natl. Acad. Sci. USA, 90: 10700 (1993). J. M. Kerr, S. C. Banville, and R. N. Zuckerman, J. Am. Chem. Soc., 115: 2529 (1993). M. H. J. Ohlmeyer, R. N. Swanson, L. W. Dillard, J. C. Reader, G. Asouline, R. Kobayashi, M. Wigler, and W. C. Still, Proc. Natl. Acad. Sci. USA, 90: 10922 (1993). K. C. Nicolau, X. Y. Xiao, Z. Parandoosh, A. Senyei, and M. P. Nova, Angew. Chem. Int. Ed. Eng., 34: 2289 (1995). E. J. Moran, S. Sarshar, J. F. Cargill, M. M. Shahbaz, A. Lio, A. M. M. Mjalli, and R. W. Armstrong, J. Am. Chem. Soc., 117: 10787 (1995). T. Carell, E. A. Wintner, A. Bashirhashemi, and J. Rebek, Angew. Chem. Int. Ed. Engl., 33: 2059 (1994). H. M. Geysen, R. H. Meleon, and S. J. Barteling, Proc. Natl. Acad. Sci. USA, 81: 3998 (1984). A. M. Bray, N. J. Maeji, and H. M. Geysen, Tetrahedron Lett., 31: 5811 (1990). R. A. Houghten, Proc. Natl. Acad. Sci. USA, 82: 5131 (1985). A. G. Beck-Sickinger, H. Durr, and G. Jung, Pept. Res., 4: 88 (1991). S. P. A. Fodor, J. L. Read, M. C. Pirrung, L. Stryer, A. T. Lu, and D. Solas, Science, 251: 767 (1991). D. L. Boger, C. M. Tarby, P. L. Myers, and L. H. Caporale, J. Am. Chem. Soc., 118: 2109 (1996). S. Cheng, D. D. Comer, J. P. Williams, P. L. Myers, and D. L. Boger, J. Am. Chem. Soc., 118: 2567 (1996). J. J. Parlow, Tetrahedron Lett., 36: 1395 (1995). I. Ugi, A. Domling, W. Horl, Endeavour, 18: 115 (1994). L. Weber, S. Wallbaum, C. Broger, and K. Gubernator, Angew. Chem. Int. Ed. Engl., 34: 2280 (1995). P. L. Myers, Proceedings of the 13th International Symposium of Laboratory Automation Robotics, 1995: 235 (1995). H. Han, M. M. Wolfe, S. Brenner, and K. D. Janda, Proc. Natl. Acad. Sci. USA, 92: 6419 (1995).

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61. H. V. Meyers, G. J. Dilley, T. L. Durgin, T. S. Powers, N. A. Winssinger, H. Zhu, and M. R. Pavia, Molec. Div., 1: 13 (1995). 62. M. R. Pavia, M. P. Cohen, G. J. Dilley, G. R. Dubuc, T. L. Durgin, F. W. Forman, M. E. Hediger, G. Milot, T. S. Powers, I. Sucholeiki, S. Zhou, and D. G. Hangauer, Bioorg. Med. Chem., 4: 659 (1996). 63. J. Rohr, Angew. Chem. Int. Ed. Engl., 24: 881 (1995). 64. R. McDaniel, C. M. Kao, H. Fu., P. Hevezi, C. Gustafsson, M. Betlach, G. Ashley, D. E. Cane, C. Khosla, J. Am. Chem. Soc., 119: 4309 (1997). 65. C. M. Kao, M. McPherson, R. N. McDaniel, H. Fu, D. E. Cane, C. Khosla, J. Am. Chem. Soc., 119: 11339 (1997). 66. R. P. Sheridan and S. K. Kearsley, J. Chem. Inf. Comput. Sci., 35: 310 (1995).

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11 Imidazoline Receptors and Their Endogenous Ligands Colin J. Barrow The University of Melbourne, Parkville, Victoria, Australia

Ian F. Musgrave Prince Henry’s Institute for Medical Research, Clayton, Victoria, Australia

I. INTRODUCTION The development of clonidine as an antihypertensive agent over 20 years ago opened new vistas in receptor research with the elucidation of multiple α-adrenoceptor subtypes (α1A-D, α2A-D). Paradoxically it is now accepted that clonidine also defines distinct, nonadrenergic receptors or sites that recognize its imidazoline structure [1–3]. These sites have been previously termed imidazoline preferring sites, imidazoline-guanidinium receptive sites (IGRSs), or nonadrenergic imidazoline-binding sites (NAIBSs) and are now designated as imidazoline receptors [1–3]. Imidazoline receptors appear to be of physiological importance [1–3], but whether these sites are receptors in the pharmacological sense remains to be determined.

II. PHARMACOLOGY AND FUNCTION OF IMIDAZOLINE RECEPTORS Imidazoline receptors are classified principally by their ligand binding characteristics. Like α2-adrenoceptors, imidazoline receptors have high affinity for compounds with an imidazoline [e.g., clonidine (3) and idazoxan (10)], oxazoline

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[e.g., rilmenidine (8)], or guanido structure [e.g., guanabenz (13), see Table 2 for structures] [4–7]. Because of this, it was originally thought that imidazolinebinding sites were a subtype of the α2-adrenoceptors. However, as imidazolinebinding sites have negligible affinity for phenylethylamines such as epinephrine and norepinephrine, or for α2-adrenoceptor ligands that are structurally unrelated to imidazolines, such as yohimbine and BHT 933 [4–7] this is now considered unlikely. Furthermore, imidazoline receptors have been shown to be physically distinct from α2-adrenoceptors [8], and it is now established that the imidazoline receptors represent distinct entities [3]. Imidazoline receptors can be subdivided into at least two types of imidazoline-binding sites. These are the I1-sites, which have a high affinity for clonidine [4,9], and I2-sites, which have a high affinity for idazoxan [10] (see Table 1 for rank orders of potency; see also Table 2). Both the I1- and the I2-sites may be further subdivided on the basis of drug affinity. There is also at least one further site, the so-called I3 binding site [13]. In addition, some imidazoline-preferring, nonadrenergic sites that cannot be classified as I1-imidazoline-binding sites or I2imidazoline-binding sites exist and so are collectively called non-I1, non-I2-sites [13,14].

A.

I1-Imidazoline-Binding Sites

The I1-imidazoline-binding sites are characterized by having nanomolar affinity for clonidine (3), naphazoline (2), cirazoline (4), and idazoxan (10), with clonidine having a higher affinity than idazoxan [7,15–18] (see Table 2). Moxonidine (7) and rilmenidine (8), recently developed antihypertensives, are relatively selective for the I1-imidazoline-binding sites over I2-imidazoline-binding sites and α2adrenoceptors, although the exact degree of selectivity varies in different reports [2,15,17]. There may be more than one form of I1-imidazoline-binding site; for example, human I1-imidazoline-binding sites differ from bovine sites in having lower affinity for guanabenz (13) and moxonidine [2,15–18]. The binding of ligands to the I1-imidazoline-binding sites is strongly inhibited by cations such as K⫹ and tetraethyl ammonium (TEA) [7,15] (see Table 1). I1-Imidazoline-binding sites are mostly distributed in the brain (corpus striatum, hippocampus) and the brain stem (medulla oblongata, rostral ventrolateral medulla) [2,11]. I1-Sites have also been found in the prostate [19] and the kidney [16], in both cases localized to the epithelium. Virtually nothing is known about the structure of I1-imidazoline-binding sites. They are present in the plasma membrane, and although some investigators have reported that [3H]-clonidine binding to these sites is modulated by guanosine triphosphate (GTP) suggesting a G-protein-linked receptor [19,20], to date there is no consensus [7,15,21]. Furthermore, binding of ligands to I1-sites does not

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Table 1 Classification of Imidazoline Receptorsa

Selectivity

Cation sensitivity Location G-protein linked Subtypes Structure Tissue location Action

a

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References

Various, often idazoxan ⱖ clonidine

6,7,14,74,77, 80,81

Unknown

7,15,77,81

Unknown Unknown

7,40,80,82 77,78,79,81, 82 7,77,80,81, 82 41 2,11,12,16,19, 14,37,78,81 29,30,31,33, 34,44,71

I2-Imidazoline-binding sites

Cirazoline ⫽ clonidine ⫽ naphazoline ⬎ idazoxan ⬎ phentolamine ⬎⬎⬎ adrenaline 4-AP ⬎ CsCl ⫽ TEA ⬎ RbCl ⬎ KCl ⬎ LiCl ⬎ NaCl Plasma membrane Unknown

Cirazoline ⫽ idazoxan ⬎⬎ clonidine ⬎ naphazoline ⫽ phentolamine ⬎⬎⬎ adrenaline 4-AP ⬎ CsCl ⬎ TEA ⬎ RbCl ⬎ KCl ⬎ NaCl ⬎ LiCl Outer mitochondrial membrane No

Probably

A,B

Probably a number of sites

Unknown Brain stem, basal ganglia, hippocampus, prostate, kidney Modulation of blood pressure Modulation of ocular pressure? Modulation of gastric secretion? Modulation of renal secretion?

Monoamine oxidase, other? Adipose tissue, liver, placenta

Unknown Brain, stomach

Growth factor–like effects?

Unknown, modulation of gastric secretion?

TEA, tetraethyl ammonium.

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Non-I1, non-I2-imidazolinebinding sites

I1-Imidazoline-binding sites

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Table 2 Structure/Affinity Relationships for Ligands at I1-Imidazole-Binding Sitesa Compound

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Ki vs. I1

Ki vs. I2 Ki vs. α2A

Reference

Oxymetazoline (1)

6.2 19,500

Naphazoline (2)

8.4

1,451

14.3

7,53,77

Clonidine (3)

14.8

272

8.7

7,53,77

Cirazoline (4)

17.0

4.3

193

BU 224 (5)

41.5

20.8

4,020

Efaroxan (6)

52.4 44,800

Moxonidine (7)

55

Rilmenidine (8)

59.2

65.1

2-BFI (9)

66.7

9.0

36,600

Idazoxan (10)

103

Phentolamine (11)

148

1,424

Agmatine (12)

127

74,440

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4.1

5.9

Structure

9.8

150

36.2

8,490

53,80

7,53,77

53

53

53,83

53

53

12.2

7,53,77

51.3

7,53,77

46,980

53

Table 2 Continued Compound

Ki vs. I1

Guanabenz (13)

35,100

Amiloride (14)

a b

⬎10,000

Ki vs. I2 Ki vs. α2A

Structure

Reference

12.9

2.0

53,80

30b

n.d

7,77

Ki Values are nanomolar. Amiloride distinguishes between two forms of the I2-imidazoline, binding site, one with a high affinity for amiloride (I2A) and one with a low affinity (⬎1000 nM: I2B); n.d., Not determined.

activate G-protein-mediated signal transduction pathways such as adenylate cyclase or phospholipase C [22–25]. Other possible targets are the ion channels. However, imidazolines do not appear to inhibit voltage operated calcium channels or nonselective cation channels in the neuronal model, rat pheochromocytoma (PC12) cells [24], or nonselective cation channels in a nonneuronal model [26]. Ligand-gated ion channels, which are important in modulation of neuronal activity, have been shown to be inhibited by imidazolines [24,25,27], in particular the nicotinic acetylcholine receptor [24,25], the 5HT3 receptor [27], and the adenosine triphosphate– (ATP)-dependent K⫹ channel [28]. However, the profile of potency of imidazoline ligands against the latter two channels suggests a non-I1, non-I2 site [27,28]. The I1-imidazoline-binding site has been implicated in the hypertension produced by clonidine and other imidazolines. When injected into the brain stem baroreflex center; the rostral ventrolateral medulla (RVLM) of the cat [29], rat, and rabbit [30,31]; imidazolines produced hypotension through a nonadrenergic mechanism [29–31]. This observation suggests an I1-imidazoline-binding sitemediated effect on the basis of the relative potencies of the imidazolines, and the efficacy of the relatively I1-site-selective agents moxonidine and rilmenidine [1–4]. The actual mechanism by which I1-imidazoline-binding sites act is unclear. However, there is a reduction in the activity of catecholaminergic neurons in the brain stem when imidazoline antihypertensives are injected [31,32]. The I1-site has also been implicated in modulation of gastric acid secretion [33], modulation of intraocular pressure via central nervous system (CNS) action [34], and modulation of electrolyte secretion in the rat kidney [35]. However, full pharmacological characterization of these actions has not been performed.

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B.

I2-Imidazoline-Binding Sites

I2-Imidazoline-binding sites have a distinct binding profile, with idazoxan and cirazoline having nanomolar affinity for these sites, whereas clonidine and moxonidine are much weaker, typically having micromolar affinity (see Tables 1 and 2). Relatively selective ligands for I2-imidazoline-binding sites include 2-(2benzofuranyl)-2-imidazoline (2-BFI) (9) and 2-(4,5-dihydroimidaz-2-yl-quinoline) (BU224) (5) [36]. The I2-imidazoline-binding sites can be further subdivided on the basis of their affinity for amiloride (14) into I2A-sites and I2B-sites. Similar to I1-imidazoline-binding sites, the I2-imidazoline-binding sites are sensitive to cations (Table 1). The I2-imidazoline-binding sites are widespread, found in astrocytes, kidney, liver, adrenal medulla, adipocytes, placenta, and urethra [37]. There is considerable detailed information about the structure of the I2sites. These sites are proteins of molecular weight 60–70 kDa, insensitive to GTP, present in the outer mitochondrial membrane, and physically distinct from the α2-adrenoceptor [10,38–40]. The molecular size and mitochondrial location suggest that I2-sites could be located on monoamine oxidases. Recently it has been demonstrated that cloned monoamine oxidases (MAOs) A and B do indeed possess I2-sites [41], and that the two forms of imidazoline-binding protein correspond to MAO-A and MAO-B. Whether all I2-imidazoline binding sites represent MAO remains to be determined. In contrast to that for the I1-imidazoline-binding sites, no definitive physiological role for I2-sites has yet been determined. Binding of imidazolines to monoamine oxidase does inhibit these enzymes, and thus they may have a role in modulating mitochondrial function [42]. Prolonged incubation of tissues with imidazolines also down-regulates I2-imidazoline-binding sites, suggesting that they are under physiological control and therefore have a functional role [43]. Furthermore, idazoxan also acts as an antiproliferative agent in vascular smooth muscle [44]. This action occurs presumably through I2-imidazoline-binding sites, since the potency of compounds as antiproliferative agents correlates with their affinity at the I2-imidazoline-binding site in blood vessels [44].

III. THE SEARCH FOR ENDOGENOUS IMIDAZOLINE RECEPTOR LIGANDS A.

Extraction and Isolation of Clonidine-Displacing Substance

More than 10 years of research has been devoted to the search for endogenous ligands for imidazoline receptors. As early as 1984 an endogenous compound that competes for [3H]-clonidine binding was partially purified from both bovine and rat brain [45]. This compound, labeled ‘‘clonidine-displacing substance’’

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(CDS), was determined to be a heat-stable, methanol-soluble, nonpeptidic small molecule. The molecular weight of CDS was estimated at 500 daltons using size exclusion chromatography [45,46], and at 588 Da using plasma desorption mass spectrometry [47]. More recently, CDS has been partially purified from human plasma [48], human cerebrospinal fluid [49], and bovine lung [50]. However, it has not been unambiguously determined whether the CDS compound or compounds responsible for the observed biological activities are the same in each case. There has been considerable disagreement in the literature on both the physicochemical and the biological properties of CDS, possibly because of the range of extraction procedures, assay systems, and purification strategies employed by the various groups in obtaining CDS [4,45,46,50–52]. For example, partially purified CDS initially isolated from bovine brain by Atlas and Burstein, eluted from reversed-phase chromatography with 39%–45% propanol in 10 mM ammonium bicarbonate [45], indicated a hydrophobic molecule. In contrast, a recent investigation found that neither lung nor brain CDS was retained on reversed-phase chromatography, with all activity eluting with water, suggesting a hydrophilic compound [50]. Atlas and Burstein initially obtained CDS extracts from bovine brain by homogenizing at pH 7.7 in aqueous solution, boiling the supernatant for 15 min to coagulate proteins, centrifuging and removing solvent, then extracting the residue with methanol. These authors reported that CDS is pH- and heat-stable. A recent study, however, indicated that CDS degraded in brain homogenates at high temperature and that 50% of the CDS activity was lost over 3 h when methanol extracts were diluted with HEPES-magnesium-EGTA assay medium at pH 7.7 [53]. These inconsistent properties reported for CDS indicate that multiple CDS compounds are probably present in brain extracts. The presence of multiple CDS compounds together with possible instability could be responsible for the varying biological activities observed for extracts reputedly containing CDS. The initial detection and partial purification of an endogenous CDS were performed by monitoring binding to α2-adrenergic receptors rather than imidazoline receptors [44]. The major reasons for this were the availability of sensitive assays for α2-adrenoceptors, the uncertainty as to the existence of specific imidazoline receptors, and the activity of CDS, like that of clonidine, at both α2-adrenergic and imidazoline receptors. However, the recently isolated CDS, agmatine (12), is considerably more potent at the I1-receptor than at the α2-adrenergic receptor [53], indicating that bioassay directed isolation using a single binding assay to detect CDS activity could be misleading. Thus, if CDS activity in brain extract is due to multiple compounds with differing receptor selectivity, then multiple binding assays should be used during fractionation to distinguish activities due to different CDS molecules. Using a single binding assay for monitoring isolation makes it impossible to determine whether activity spread during frac-

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tionation is due to a single compound running poorly or the separation of multiple active components. Use of single concentrations of extract in binding assays during CDS isolation was also recently reported to be misleading in that some fractions with potent activity at a single concentration showed steep binding curves when multiple concentrations were examined [50], suggesting that the activity was an artefact or, alternatively, that the receptors exhibit strong positive cooperativity. Thus, it is strongly advisable to monitor binding at multiple concentrations during fractionation. In addition, compounds that have CDS binding activity do not necessarily have corresponding functional activity, indicating that fractionation should also be monitored at least periodically using an appropriate functional assay. For example, agmatine, recently isolated as an endogenous CDS with both imidazoline and α2-adrenergic receptor-binding activity, does not appear to be functionally active at the α2-receptor (see Section V.C). It should also be noted that functional activity reported for methanolic brain extracts containing CDS activity may be misleading. For example, norepinephrine present in crude methanolic bovine brain extracts may be responsible for the reported ability of CDS to inhibit electrically evoked contraction of the rat isolated vas deferens by activation of α2adrenoceptors [50,54]. The levels of norepinephrine in the crude methanolic extract of brain appear to be too low to account for all the CDS binding activity but could account for the observed functional activity in a system with a high receptor reserve where even low concentrations of norepinephrine could produce a response [50]. Any effort to isolate new endogenous CDS compounds needs to eliminate the possibility that activity is due to known contaminants such as monovalent cations, histamine, catecholamines, and the reported CDS agmatine. In addition, once a semipurified CDS has been obtained, then a variety of assays need to be performed to confirm the presence of both binding and functional activity. Further purification should be monitored using full binding curves and functional activity, at least periodically, to eliminate false-positive results. However, not until purified CDS is isolated and structurally characterized will the biological properties of endogenous CDS compounds be unambiguously determined. B.

Biological Activity of Clonidine-Displacing Substance

In addition to displacing [3H]-clonidine from various membrane preparations, CDS has been shown to have functional effects when applied to several preparations. Microinjection of partially purified CDS derived from brain into the RVLM of the rat or cat modifies arterial pressure [55–57]. However, the results are not consistent: either an increase [56,57] or a decrease [55] in arterial pressure may be produced. It is not clear whether the variations are due to differences in the CDS produced or the presence of contaminants such as biogenic amines and K⫹ [50]. Contaminants may not be necessarily responsible for the hypertensive

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fect, as a nonhypertensive dose of the CDS was able to inhibit the hypotensive effect of clonidine microinjected into the RVLM of rats and cats [57] and the hypotensive effects of rilmenidine microinjected into the RVLM in rabbits [58]. These results are consistent with an action at a nonadrenergic site in the brain stem. Partially purified CDS also has functional effects that may be due to an action at α2-adrenoceptors. For example, brain-derived CDS blocks electrically stimulated contraction of rat vas deferens [54]. This effect was reversed by yohimbine and thus is consistent with an agonist action at α2-adrenoceptors [54]. In this CDS preparation a possible contribution from norepinephrine cannot be excluded, but in other preparations such contamination is unlikely. For example, brain-derived CDS antagonizes the epinephrine-induced aggregation of human platelets but is unable to stimulate aggregation by itself [59]. Thus, blockade of aggregation is unlikely to be due to contaminating biogenic amines. If there was a significant amount of norepinephrine/epinephrine present, then aggregation should have been seen, or epinephrine-induced agregation potentiated. Brain-derived CDS can also contract urethral smooth muscle but has no effect on vascular smooth muscle [60]. This is a nonadrenergic effect that could not be blocked by antagonists to muscarinic, serotonergic, angiotensin II, or bradykinin receptors [60]. Furthermore, as the contractile effect was not seen in vascular smooth muscle, this eliminates the possibility that cations such as K⫹ are responsible [60]. In contrast, plasma-derived CDS was able to produce concentration-dependent contraction of rat aortic rings [61]. This appears to be due to an action at α2-adrenoceptors as the effect was inhibited by the α2-adrenoceptor antagonist rauwolscine, but not by the α1-adrenoceptor antagonist prazosin [61]. These results suggest that the effect of the CDS is not due to contaminating biogenic amines, which should have activated α1-adrenoceptors, or cations such as K⫹, which would cause depolarization. Thus, despite a wide variety of preparation procedures and biological sources, in some cases CDS has actions that can be attributed to effects at α2adrenoceptors, independent of contaminating biogenic amines or cations. Whether or not there is more than one CDS or whether other non-CDS material in the partially purified extracts contributes to activity is unclear from these results.

IV. AGMATINE: THE FIRST ENDOGENOUS CLONIDINE-DISPLACING SUBSTANCE A.

Isolation and Identification of Agmatine

Recently agmatine (12), the decarboxylation product of arginine, was isolated as the first endogenous CDS compound from bovine brain [62]. Although agmatine does not appear to have all the biological properties attributed to CDS, it clearly

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binds to imidazoline receptors and has some functional activity [62]. Agmatine was isolated from bovine brain with a bioassay-directed isolation strategy, using displacement of [3H]-p-aminoclonidine from rat cerebral cortical membranes to track activity. The extraction method began with homogenization of fresh brain in chilled water, then centrifugation. Protein was then precipitated from the supernatant with 70% ethanol, then the extract was concentrated, centrifuged, and passed through a Dowex 50 (H⫹) column eluting with 3N hydrochloric acid [62]. This method prevents the temperature extremes employed by Atlas and Burstein in their initial isolation and identification of a CDS [45–47], which appears not to be agmatine [63,64]. After ion-exchange chromatography, agmatine was partially purified by size-exclusion chromatography, followed by reversed-phase C18 high-performance liquid chromatography (HPLC), but because of the high polarity of this compound and lack of an ultraviolet (UV) chromophore, attempts to purify agmatine further were unsuccessful. Agmatine was eventually isolated as its 9fluorenylmethyl chloroformate (FMOC) derivative. Structural confirmation of this CDS compound was performed using electrospray mass spectroscopy and direct comparison with authentic agmatine [62]. During the decomposition of dead tissue, polyamines such as agmatine are formed from amino acids by the action of bacterial decarboxylases. Indeed, detection of agmatine has previously been used as an indicator of the freshness of fish [65]. Therefore, when agmatine was initially isolated in minor quantities from bovine brain there was concern that it may be a bacterial contaminant rather than an endogenous compound. However, arginine decarboxylase activity was shown to be present in the brain by measuring the conversion of l-[14C]arginine to 14CO2 [62]. The presence of arginine decarboxylase activity in rat brain indicated that agmatine is indeed a compound endogenous to mammalian brain, and therefore a true endogenous CDS. B.

Distribution of Agmatine in Mammalian Tissue

In a more recent study, agmatine levels in a variety of rat tissues were measured by a standardized method consisting of tissue homogenization, derivatization, extraction on a C18 cartridge, and HPLC chromatographic analysis with fluorescence detection [66]. Agmatine was found in all tissues examined with the greatest concentration in the stomach (Table 3). It is interesting to note that agmatine was present in relatively low concentration in brain tissue (2.4 ng/g) and that agmatine concentrations in the lung were approximately 5 times higher than in the brain, whereas those in the stomach were approximately 20 times higher than in the brain [66]. Singh et al. recently reported the presence of a noncatecholamine CDS in methanolic extracts of bovine lung in a threefold higher concentration than that found in bovine brain [50]. However, it was concluded that this

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Table 3 Distribution of Agmatine in Various Tissues of Adult Male Sprague-Dawley Rats Tissue Stomach Aorta Small intestine Large intestine Spleen Lung Vas deferens Adrenal gland Kidney Heart Liver Skeletal muscle Brain Testes Plasma

Wet weight (ng/g)a 71.00 57.41 55.35 27.86 17.38 10.23 9.45 6.97 6.45 6.45 5.63 5.30 2.40 2.04 0.45

⫾ ⫾ ⫾ ⫾ ⫾ ⫾ ⫾ ⫾ ⫾ ⫾ ⫾ ⫾ ⫾ ⫾ ⫾

10.33 12.74 9.39 6.73 3.17 2.82 2.08 3.29 1.40 0.79 0.87 0.72 0.60 0.22 0.05

a

Concentrations are expressed in wet weight (ng/g). Data are averaged from either four or five experiments. Source: Ref. 66.

CDS was not agmatine, again indicating the presence of multiple CDS compounds in multiple tissue types [50]. Thus, agmatine and other CDS compounds appear to be widely distributed in mammalian tissue. Agmatine present in the stomach appears to be contained in endothelium and vascular smooth muscle and so may play a role in modulating gastric secretion [66]. However, there is substantial variation in the quantity of agmatine in individual animals [66], greater than 10-fold in some cases, raising concern about a functional role for agmatine. Furthermore, the imidazoline-binding site in the stomach appears to be a non-I1, non-I2-site [14].

V.

BIOLOGICAL EVALUATION OF AGMATINE

A.

Cardiovascular Effects of Agmatine

The effect of agmatine on sympathetic cardiovascular regulation in conscious rabbits was recently characterized [67]. Agmatine injected intracisternally caused central sympathoexcitation by increasing renal sympathetic nerve activity, plasma

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catecholamine concentrations, and arterial blood pressure [67]. However, a decrease rather than an increase in heart rate was observed. It is interesting to note that extracts containing CDS have been reported to cause either an increase [56,57] or a decrease [55] in blood pressure (see Section III.B), illustrating the difficulties of working with semipurified material. Agmatine could have been the active compound in the studies reporting an increase in blood pressure, but in the study reporting a decrease in blood pressure, a second compound would have to be responsible. The mechanism by which agmatine exerts these cardiovascular effects is unclear, although mediation via imidazoline receptors is a possibility. Agmatine is able to pass and block the intrinsic-ion channel of the nicotinic receptor [68]. However, this is unlikely to play a role in sympathoexcitation by agmatine as high millimolar concentrations of agmatine are required to block the nicotinic receptor [68]. It is also unlikely that agmatine acts via α2-adrenoceptors for a number of reasons. First, agmatine does not appear to be functionally active at α2 adrenoceptors [53,67]; that is, it is neither an antagonist nor an agonist (see later discussion). Second, intracisternally injected agmatine causes bradycardia, in contrast to intracisternally injected yohimbine which is known to act via blockade of α2-adrenoceptors [67]. It has been suggested that the I1-receptor agonist clonidine acts on I1-receptors in the RVLM to produce its sympathoinhibitory effect [69]. Thus, it is possible that the sympathoexcitatory effects of agmatine are due to antagonist effects at I1-receptors. However, a recent report confirming the sympathoexcitatory activity of agmatine injected intracisternally showed that agmatine, unlike clonidine, does not alter arterial pressure when microinjected into the RVLM [69]. Iontophoresis of agmatine onto defined vasomotor neurons in the RVLM was also without effect [69]. Therefore, although agmatine has a high affinity for I1-receptors [53], its cardiovascular effects elicited by intracisternal injection are probably not modulated by these receptors. Also, because inhibition of the RVLM vasomotor neurons represents the major mechanism by which clonidine produces its hypotensive effects [69], agmatine and clonidine appear to have quite different mechanisms of cardiovascular action. In contrast to central administration of agmatine, intravenous administration of agmatine into rats produces sympathoinhibition [69] with a fall in arterial blood pressure [69,70]. The mechanisms for the hypotensive actions of peripherally administered agmatine are still unclear, but blockade of ganglionic transmission and vasodilatation from a direct action on vascular smooth muscle may be involved [69,70]. B.

Gastrointestinal Function of Agmatine

The relationship between hypertension and gastrointestinal disorders, such as gastric ulcers, can be investigated by examining the actions of known antihyper-

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tensive drugs on gastric function or response to injury. For example, the antihypertensive drug clonidine has been shown to decrease gastric acid secretions while enhancing gastric adherent mucus levels [33]. The identity of the receptors through which clonidine produces its gastrointestinal actions is unknown. However, the I1-imidazoline receptor agonist moxonidine is both a potent inhibitor of basal gastric acid secretion and significantly raised intragastric pH, indicating that I1-imidazoline receptors may be involved in gastrointestinal function [33]. Agmatine, if an endogenous ligand for the imidazoline receptors, should have similar gastrointestinal effects to those of I1-imidazoline receptor agonists such as moxonidine. Agmatine was recently tested for effects on gastrointestinal function and on experimental gastric mucosal injury and was found to augment gastric acid and pepsin secretion, decrease gastric adherent mucus, and worsen experimental gastric mucosal injury consequent to stress [71]. These effects are opposite to the gastrointestinal effects observed for the I1-site agonist moxonidine [33]. The observed ‘‘inverse agonist’’ activity of agmatine at imidazoline receptors could be a result of the involvement of different subtypes of the imidazoline receptor at the same site. Alternatively, agmatine may modulate gastrointestinal function via novel receptor subtypes [14,71]. C.

Interaction of Agmatine with ␣2-Adrenoceptors and Imidazoline-Binding Sites

Agmatine has been shown to produce a concentration-dependent inhibition of [3H]-clonidine binding to both rat and bovine cerebral cortex membranes with a Ki of approximately 10 µM [18,62,72]. However, in functional studies agmatine failed to act as either an α2-adrenoceptor antagonist or an agonist in a wide variety of tissues [67,72,73]. For example, agmatine did not inhibit electrically evoked contractions of the rat vas deferens [72], in contrast to activity originally described for partially purified CDS [54] and the action of the α2-adrenoceptor agonist clonidine [72]. This indicates that in studies with CDS-containing extracts, agmatine may be responsible for the activity detected in the radioligand binding assay, but other compounds such as catecholamines [74] or a second CDS compound must be responsible for the observed functional activity. Furthermore, at concentrations of 100 µM, agmatine did not shift the concentration response curve to clonidine in the rat vas deferens even though a 10fold shift was expected [72], suggesting that agmatine is not an α2-adrenoceptor antagonist. The same results were seen separately with both clonidine and the selective α2-adrenoceptor agonist UK 14304 in rat thoracic aorta, rat cerebral cortical slices, porcine palmar veins, and guinea pig ileum [72]. Similar results have also been seen in rabbit occipital cortex slices [67] or rat locus ceruleus slices [73]. Thus, agmatine would appear to be neither an agonist nor an antago-

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Table 4 Comparison of Clonidine-Displacing Substance and Agmatine Inhibition Constants at α2-Adrenergic and Imidazoline-Binding Sites from Human and Bovine Tissuesa Binding sites Human Human Human Human Human

α2A α2B α2C I2b I1

Bovine I1

CDS IC50 (Diluted ⫻ 10⫺3) 51 48 20 23 49

⫾ ⫾ ⫾ ⫾ ⫾

4 3 1 1 2

9⫾2

Agmatine Ki (nM) 46,980 ⫾ 2,600 164,400 ⫾ 21,412 26,300 ⫾ 3,969 74,400 ⫾ 14,152 Highb 33 ⫾ 19 Low 284,600 ⫾ 37,903 High 127 ⫾ 33 Low 275,500 ⫾ 51,900

a

CDS, clonidine-displacing substance. Human and bovine I1-binding sites have both a high- and a low-affinity site for agmatine. Source: Ref. 51.

b

nist at α2-adrenoceptors, even though it binds to these receptors. However, it may be that agmatine is a much weaker antagonist than its Ki value would suggest. Jurkiewitz et al. [74] found that agmatine acted as an α2-adrenoceptor antagonist in the rat vas deferens, but only at concentrations of 0.3 and 1 mM. Because crude methanolic bovine brain extracts show CDS activity consistent with agonist activity at α2-adrenoceptors, and this activity is attributable to neither agmatine nor histamine, a second CDS compound is probably present in these extracts [75]. Furthermore, direct comparison of relative potencies of CDS-containing brain extract and agmatine at all three human α2-adrenergic receptor subtypes and at human imidazoline sites suggests that CDS and agmatine are not equivalent (Table 4). Crude bovine CDS was potent at all sites, although a 2.5- to 6-fold higher potency for bovine imidazoline I1-sites was observed when compared with all human sites, indicating some selectivity of CDS for I1-sites [53]. Agmatine, in contrast to CDS, bound to two sites, one with high affinity (33% of all sites) and one with micromolar affinity (Table 4). The reason for the apparent presence of both high- and low-affinity binding sites for agmatine at I1-receptors is still under investigation [53]. Again in contrast to CDS, agmatine has a much lower affinity for the α2B-adrenoceptor vs. the α2A-adrenoceptor (Table 4). The differences between the potencies of methanolic CDS extract from brain and agmatine further suggest that at least one component other than agmatine is present in the CDS extract. However, the high affinity of agmatine for a subset of I1-sites still supports agmatine as a true endogenous ligand for I1-sites [53].

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VI. RECENT PROGRESS IN THE SEARCH FOR ENDOGENOUS CLONIDINE-DISPLACING SUBSTANCES The purification and characterization of endogenous CDSs other than agmatine remain elusive. Major reasons for this include the absence of a definitive biological action for CDS, the presence of multiple CDSs and biologically active contaminants, and the limited abundance of CDS in tissue. A recent study has shown that tissue other than brain contains CDS activity [75, see also 44]. As previously noted, bovine lung is reported to have approximately three times the CDS activity found in the brain [75]. Although it has not been confirmed that the same CDSs are responsible for activity observed in different tissue types, it appears that known contaminants such as monovalent cations, histamine, catecholamines, and the known CDS agmatine cannot explain all observed binding activity found in either lung or brain extracts. After eliminating activity due to these contaminants it appears that the remaining CDS activity in both lung and brain samples is due to an unidentified compound with agonist activity at α2-adrenoceptors [72]. Chemical properties of the unidentified CDSs from bovine lung and brain indicate that both tissues contain the same CDSs [50]. Both lung and brain CDS are poorly soluble in nonpolar solvents at room temperature and are retained on cation exchange columns, indicating that the CDS from both tissues has a positive charge. Furthermore, CDS was recovered from similar fractions when both lung and brain extracts were chromatographed using size-exclusion chromatography [50]. The authors of this study found that neither lung nor brain CDS is retained on C18 chromatography, a result contrasting with previous work, when CDS from brain was strongly retained on C18 material and only eluted with 35% propanol/ water [46,47]. In our group we have recently attempted to simplify the search for new endogenous CDS by (1) developing a large-scale extraction method to overcome the difficulties in obtaining enough CDS material for structure determination; (2) determining whether CDS activity in lung is due to the same compound that is responsible for CDS activity in brain; (3) developing an HPLC detection method for CDS; and (4) obtaining structural information for a CDS other than agmatine. In the following section we detail some of these results [76]. A.

Development of a Large-Scale Extraction Method

The standard method typically used for extraction of CDS activity from tissue is difficult to scale up to the kilogram quantities needed to obtain enough CDS for nuclear magnetic resonance (NMR) studies, for several reasons. First, the method uses time-consuming homogenization and filtration of the homoge-

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nate. Second, large quantities of aqueous filtrate then need to be freeze-dried. Finally, the water extraction procedure exposes CDS to enzymatic degradation, and the use of boiling water could potentially degrade heat-sensitive CDS molecules. We have modified the standard extraction method by replacing the timeconsuming filtration with centrifugation and eliminating heating, water extraction, and use of a homogenizer. We have thus developed an extraction method that can be routinely used for extraction of CDS activity from a variety of tissues on a kilogram scale. This method involves lyophilization of wet tissue followed by extraction of inert fats by blending with hexane. After centrifugation to remove the hexane solubles the tissue is blended with methanol and the methanol-insoluble material removed by centrifugation. The methanol is removed by rotary evaporation to yield a CDS extract. Quantitative comparison of this method with the previously used standard extraction method has shown that considerably more CDS activity was extracted from both lung and brain tissue by using our optimized method.

B.

Development of a Large-Scale Clonidine-Displacing Substance Isolation Method

During the isolation procedure, CDS activity was monitored by using an α2adrenoceptor radioligand membrane binding assay, with displacement of [3H]clonidine. Naphazoline was used to calculate nonspecific binding. Clonidine, norepinephrine, and yohimbine were used as positive controls [45,46]. Methanol extracts were preabsorbed onto C18 reversed-phase material and passed through a C18 column, eluting first with water and subsequently with increasing ratios of methanol/water. In addition to ease of scale-up, using C18 chromatography in the first purification step retained the majority of CDS activity on the stationary phase after elution with water, so cations responsible for nonspecific activity were readily removed. As reported previously [50], CDS activity was retained on a cation exchange column, indicating that CDS is positively charged. We found that CDS activity was enriched after eluting the partially purified C18 fraction from an anion exchange column, probably as a result of retention of inactive anionic compounds. Therefore, an anion exchange step was incorporated after the C18 fractionation. Subsequent size-exclusion chromatography, using Biogel P2, gave good enrichment of CDS activity in fractions corresponding to a molecular weight of less than 600 Da. Final purification of a new CDS was achieved on HPLC using a C18 reversed-phase semipreparative column, eluting with 0.1% triflouroacetic acid in water (Fig. 1). Fifty micrograms of pure CDS material was obtained from 2.2 Kg of wet lung [76].

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Figure 1 Strategy for the isolation of CDS from both bovine lung and brain. CDS, clonidine-displacing substance.

C.

Development of a High-Performance Liquid Chromatography Detection Method for Clonidine-Displacing Substance

Using a variety of acidic methanol/water or acetonitrile/water gradients it was found that the majority of CDS activity eluted from C18 reversed-phase chromatography with the solvent front. However, using isocratic elution with 0.1% triflouroacetic acid in water, the majority of CDS activity eluted in a sharp band. The CDS activity was associated with a single peak with a UV maximum at 280 nm (Fig. 2). Application of the HPLC method to the detection of the new CDS compound in extracts of bovine lung and brain indicates that this compound is present in both tissues and is approximately threefold more abundant by weight in lung than in brain. Semipreparative isolation and bioassay of this peak from the initial extract confirmed its CDS activity and indicated that the compound was responsible for approximately 50% of the CDS activity observed in both lung and brain. Norepinephrine, epinephrine, histamine, and agmatine were eliminated as being responsible for the observed CDS activity by comparing the retention times and UV profile of each with those observed for the new CDS.

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Figure 2 HPLC profile with UV spectrum of purified CDS. HPLC, high-performance liquid chromatography; UV, Ultraviolet; CDS, clonidine-displacing substance.

D.

Partial Structure Determination of a New Clonidine-Displacing Substance

Using tandem mass spectrometry–mass spectrometry (MSMS) and 1H nuclear magnetic resonance (NMR) spectroscopy, a partial structure for the new CDS was obtained. Electrospray MS indicated a molecular weight of 275 Da and MSMS showed two consecutive losses of 84 Da, which were not readily assignable. The 1H NMR spectrum in dimethyl sulfoxide-d6 (DMSO-d6) indicated the presence of a sugar moiety and a heteroaromatic group. The 1H NMR and UV data were consistent with the presence of a substituted imidazole ring. Although a final structure has not been assigned, a probable partial structure (15) shows a close relationship to guanosine (16). However, the new CDS was shown not to be guanosine or its mono-, di-, or triphosphate analogues, by comparison of HPLC retention times, MS, NMR spectra, and clonidine-displacement binding data with those of authentic samples.

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It has not been determined whether the new CDS was the same as the original compound partially purified by Atlas et al. [45], or one of the other partially purified CDSs [72,75]. However, it is clear that at least one CDS other than agmatine exists in both brain and lung tissue. Full structure determination is needed to unravel the relationship between the various CDS activities being observed by a variety of groups and also to allow full biological characterization of the proposed endogenous CDS.

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12 Oxidoredox Suppression of Fungal Infections by Novel Pharmacophores Valeria Balogh-Nair The City College of the City University, New York, New York

I. INTRODUCTION Over the last decade, the incidence of fungal infections has increased dramatically. This can be attributed to drug resistance, the emergence of new pathogens and resurgence of old ones, and the lack of effective therapeutics. As patients become severely immunocompromised because of underlying diseases such as leukemia or acquired immunodeficiency syndrome (AIDS), and as immune suppression is becoming routine in cancer and organ transplantation patients, opportunistic pathogens have begun preying on a growing population, causing lifethreatening systemic fungal infections. Candida species, the ordinarily harmless denizens of the gastrointestinal and genitourinary tracts, are now becoming one of the largest threats in hospitals. Infections by Candida albicans and other opportunistic fungi are often the most devastating in AIDS patients [1]. Pneumocystis carinii pneumonia [2] affects ⬎70% of AIDS patients and is one of the most common lethal infections among them. Cryptococcus neoformans causes potentially fatal meningitis [3] in 7%–10% of AIDS patients and Candida albicans causes oral and esophageal candidiasis [4] in ⬎70% of them. In this chapter, focus will be on the development of novel oxidoredox pharmacophores and their effectiveness against the three selected pathogens, C. albicans, C. neoformans, and P. carinii. This chapter is dedicated to the memory of Professor M. S. Ramachandran Nair.

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Because fungi are eukaryotes, the search for antifungals selectively detrimental to the fungi, but safe to the host, is difficult. Investigations into the modes of action of existing antifungals have led to a better understanding of the biochemical characteristics of the fungal cell membrane and have provided a target for drug design. Advances in antifungal therapy have been dominated for many years by a single large class of synthetic compounds, which have as their common feature an azole nucleus. Fluconazole [5], like other N-substituted azoles, inhibits fungal sterol biosynthesis, resulting in the accumulation of 14-methyl fungal sterols and ultimately in damage to the fungal cell membrane. However, the unhindered nitrogen lone pair in fluconazole interacts not only with the heme of cytochrome P-450–linked monooxygenase, which catalyzes lanosterol C-14 demethylation, inhibiting fungal sterol biosynthesis, it also interferes with steroid metabolism in humans. Although possessing structures entirely different from the azoles, amphotericin B and nystatin are prominent representatives of polyene macrolide antibiotics. For more than 30 years, amphotericin B has been the preeminent drug for the treatment of serious systemic fungal infections [6]. The steroldependent ion channel formation in membranes, favoring however slightly the ergosterol-rich fungal membranes, is responsible for its potent activity. However, because of its only slight preference to disrupt fungal membranes rather than mammalian ones, its therapeutic value is greatly diminished. Although it can be lifesaving, its side effects tend to make amphotericin B an agent of last resort. Among the opportunistic infections associated with AIDS, cryptococcosis can be suppressed with fluconazole, after primary treatment with amphotericin B (or preferably with flucytosine). About 50% of the AIDS patients do not survive treatment with flucytosine, and mortality rate during therapy is 33% and 40% for amphotericin B and fluconazole, respectively. Mucocutaneous candidiasis in human immunodeficiency virus– (HIV)-infected patients can be treated with fluconazole, but its continued use to prevent recurrence leads to both clinical treatment failure and antifungal resistance, especially in highly immunodeficient patients. Pneumocystis carinii pneumonia (PCP) responds to antiprotozoal agents; therefore P. carinii has been considered by experts to be a protozoan. More recently, however, on the basis of phylogenetic classification of its ribosomal ribonucleic acid (RNA), it has been reclassified as a fungus [7]. The most effective drug regimen used against PCP infections is a combination of trimethoprim/ sulfamethoxazole [8,9]. For about 20% of patients, for whom this combination of a sulfa drug with a dihydrofolate reductase fails, requiring a switch to the alternate drug pentamidine, the prognosis is poor, with a high mortality rate. These examples illustrate the shortcomings of the antifungals in use and demonstrate the acute need for novel, safer, and more effective drugs. The main approaches to the development of better antifungal therapies include high-throughput screening programs for novel antifungal antibiotics, screening of synthetic combinatorial libraries for new leads, and rational drug

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design to identify lead compounds based on three-dimensional understanding of drug receptor interactions. Another approach, the subject of this review, is hypothesis-driven drug development. In this approach, unlike those that led to the development of azoles and polyene antifungals, the fungal cell membrane is not the target for drug design. The types of drugs sought are intended to modulate or mimic the body’s own defense mechanisms against fungal pathogens. We report herein on the discovery of novel oxidoredox pharmacophores and their antifungal activities.

II. THE MOLECULAR BASIS OF NATURAL DEFENSES AGAINST FUNGAL INFECTIONS A.

Macrophage-Derived Defense Molecules

Until the late 1980s, nitric oxide (•NO), a highly toxic gaseous free radical, was considered just one of the environmental pollutants, a destroyer of ozone, and a suspected carcinogen. Therefore, the first reports [10,11] identifying it with the elusive endothelial relaxing factor [12], a vasodilatory messenger, were greeted by many with skepticism. However, by 1992, •NO had become the ‘‘Molecule of the Year,’’ emerging as an essential and unifying thread linking neuroscience, physiology, and immunology [13]. At present, •NO is recognized as a major messenger molecule in the cardiovascular and nervous systems, and as a major effector molecule released by murine macrophages and other cells after immunological activation [14]. As an effector molecule, •NO has a role in the immune system that is quite different from its function in either neurons or blood vessels. The •NO released by macrophages is cytotoxic to intracellular microorganisms, pathogens such as fungi and helminths, and tumor cells. It is in this role that • NO, highly reactive and freely diffusing through cell membranes, participates in the host defense mechanism of eukaryotes. However, since •NO is toxic, its release by macrophages to control microorganisms, pathogens, and tumors is also linked to tissue destruction that occurs in a number of diseases, including arthritis, diabetes, septic shock, transplant rejection, and multiple sclerosis [15]. Control of the synthesis of •NO, catalyzed by nitric oxide synthases in vivo, thus becomes a key issue in regulating its activity and all of its physiological functions. Immune-stimulated macrophages express inducible nitric oxide synthases (iNOSs). These homodimeric enzymes contain a reductase domain similar to P450 reductase, and a P-450-like oxygenase domain that contains heme, H4biopterin, and a binding site for l-arginine. These two domains interact to catalyze the second step in the conversion of l-arginine to citrulline and •NO, as shown in Scheme 1 [16]. There is a consensus that when acting as a signal transducer, through oxidation of thiols, hemes, Fe clusters, and nonheme prosthetic groups, •NO regulates

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Scheme 1 Oxidative conversion of l-arginine to •NO and citrulline catalyzed by iNOS.

the target proteins that evoke the functional response. It is also understood that coordination to the enzyme-bound ferrous heme by •NO triggers guanylate cyclase-catalyzed generation of cyclic guanosine monophosphate (GMP) from guanosine triphosphate (GTP). The increases in intracellular second messenger cyclic GMP concentrations are then responsible for the cellular responses, such as vascular smooth muscle contraction. However, there is no consensus concerning the role of •NO in the immunological response leading to the microbicidal effects and accompanying tissue injury. Along with the mechanisms of action associated with •NO itself, it has been postulated that redox-related forms of •NO play a major role. Beckman et al. [17] were the first to point out that under pathophysiological conditions, both •NO and superoxide anion (O •⫺ 2 ) are produced by macrophages at high rates. These can react to form the more potent oxidant peroxynitrite (ONOO⫺), the conjugate base of peroxynitrous acid. Peroxynitrite is known to react with a variety of biologically important molecules. Nitric oxide or O •⫺ 2 reacts only slowly with the iron–sulfur cluster in mitochondrial aconitase, a major target of oxidant-mediated toxicity in cells, but peroxynitrite rapidly inactivates it [18]. Peroxynitrite oxidizes sulfhydryls, ascorbate, α-tocopherol, and lipids. Nitration of phenolic compounds, such as tyrosine, in a metal-catalyzed process also can occur. Therefore, peroxynitrite is considered sufficiently reactive to mediate nitric oxide–dependent microbial killing. During macrophage activation, its precursors •NO and O •⫺ 2 are formed simultaneously, in high concentrations, and peroxynitrite is formed at almost diffusion-controlled rates (6.7 ⫻ 10 9 M⫺1s⫺1). This indicates that significant concentrations of peroxynitrite should be attainable in vivo [19,20]. The observation that superoxide dismutase (SOD), a known quencher of superoxide, enhances •NO levels in vivo, and that nitrotyrosine is detected in biological fluids that produce high levels of • NO, casts peroxynitrite in a key role. However, these lines of evidence attributing to peroxynitrite a key role in vivo are considered by some as circumstantial.

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Other interpretations [14] have been advanced, relegating peroxynitrite to a less important role, if any. Thus, it has been suggested that enhancement of •NO levels by SOD may not be due to the ability of SOD to dismutate superoxide; SOD could oxidatively convert a reduced metabolite of •NO back to •NO instead. Alternate mechanisms for nitrotyrosine formation that do not necessarily require the presence of peroxynitrite in biological fluids have been proposed. That peroxynitrite is not the microbicidal species of phagocytes, but only an obligatory intermediate for forming other cellular oxidants, was proposed on the ground that its rapid reaction with CO 2 makes its existence in physiological fluids unlikely [21]. An intrinsic difficulty with these, as well as with many other lines of evidence for and against peroxynitrite, is that they are derived mainly from in vitro studies. Progress in the development of methods to detect and identify highly reactive intermediates in complex biological matrices has been relatively slow, limiting advances in the in vivo characterization of radical reaction pathways in these media. Sensors that can readily detect 10 ⫺20 mol of •NO, suitable for measuring intracellular concentrations, are available [22], but these •NO-specific sensors do not allow concomitant monitoring of other reactive species, sources, or targets of •NO under physiological conditions. Further studies are required to establish whether nitrosyl complexes are as important in cell defense as the reaction of • NO with superoxide radical. B.

Neutrophil-Derived Defense Molecules

Neutrophils constitute 50% to 70% of the total white blood cells in humans. Together with the macrophages, they play a critical role in the immune response against microorganisms. When triggered, reduced nicotinamide-adenine dinucleotide phosphate (NADPH) oxidase in the plasma membrane of neutrophils generates a superoxide-derived pool of ‘‘reactive oxygen derivatives’’ to serve as a first line of defense. A second line of defense consists of microbicidal–cytotoxic peptides, defensins [23], discharged from the neutrophils’ cytoplasmic granules into the phagocytic vacuole and into the extracellular space. The focus of the following discussion is on the ‘‘reactive oxygen derivatives’’ because they are important in developing novel first-line therapies. Unlike in the case of macrophages, in which the existence of iNOS is firmly established, in neutrophils the occurrence of a nitric oxide–generating system has been controversial. Depending on the assay used, it has been shown that neutrophils can or cannot release •NO [24–30]. Further, there are differences in the amounts of •NO released by neutrophils from different sources. It was argued that human neutrophils produce substantially more O •⫺ 2 than do rat neutrophils and, therefore, the resultant amount of •NO release in human neutrophils is low. To rationalize the low levels of •NO release in human neutrophils, others invoked its rapid reactions with active oxygen species or with myeloperoxidase, preclud-

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ing its detection. Careful assessment of the available data led us to conclude that although •NO release might occur in neutrophils, at present there is no evidence that neutrophil-derived •NO plays a role in the cytotoxic activity of these cells. It is an alternative system, with myeloperoxidase cast in a central role, that sustains the cytotoxic activity against both normal and tumor cells and may contribute to inflammatory tissue destruction by neutrophils. At the start of World War I, solely on the basis of screening of compounds for bactericidal action, and without awareness of their modes of action, HOCl and chloramines were employed extensively as microbicides to prevent infection of wounds. These were the very compounds that are at present held responsible for the microbicidal action of neutrophils. With many of the crucial mechanistic details still not well understood, HOCl and chloramines are produced in a complex sequence of events, as follows [31]: The NADPH oxidase in stimulated neutrophils produces superoxide anion. The bulk of the superoxide anion generated rapidly dismutates to yield hydrogen peroxide. Almost all of the hydrogen peroxide produced is used up by the large amounts of myeloperoxidase discharged from the neutrophil (up to 5% of the dry weight of the cell) to oxidize halides or thiocyanate ions. Because at most sites in vivo the plasma concentration of chloride is more than one thousand times that of other halides and thiocyanate, hypochlorous acid (HOCl) is the major oxidation product. Although the production of HOCl by human neutrophils can be sustained for up to 3h, HOCl alone is not the only important oxidant produced by neutrophils. Rapid oxidation of endogenous primary and secondary amines by HOCl under physiological conditions yields a plethora of N-Cl derivatives, the chloramines [32]; amides are also oxidized, yielding chloramides. The chloramines are less powerful, but longlived oxidizing agents, retaining the oxidizing equivalentce of HOCl. Their release is associated with the slow oxidation of microbial sulfhydryls, thereby enhancing and prolonging the microbicidal effect of neutrophils. The cytotoxicity of chloramines appears to be modulated by their overall structures, rather than by the N-Cl moiety alone. The balance between formation of lipophilic and hydrophilic N-Cl derivatives may regulate the biocidal activities of neutrophils [33]. About 30% to 40% of the endogenous amines in neutrophils consists of the βamino acid, taurine. It is converted by the myeloperoxidase/H 2 O 2 /Cl⫺ system into the hydrophilic taurine chloramine, a milder oxidant than HOCl. Thus, taurine chloramine formation could attenuate the biocidal activity of HOCl, protecting the cells from autolysis. The tissue damage accompanying the release of oxidants has been correlated with activation of latent metalloproteinases contained in the neutrophil granules [31]. These proteolytic enzymes cause tissue damage by catalyzing the hydrolytic degradation of the extracellular matrix. However, it appears that neutrophils have both oxidative and nonoxidative paths for activation of their metalloproteinases, and it has been found that the HOCl/enzyme ratio must fall

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in a narrow range for oxidative activation to occur [34]. Therefore, it is doubtful whether effective regulation of metalloproteinases by HOCl could occur in vivo, where various HOCl scavengers abound.

III. THE OXIDOREDOX HYPOTHESIS OF THE ANTIFUNGAL PHARMACOPHORES From the 1980s onward, the number of antimicrobial compounds in preclinical and clinical development has declined and there is now a shortage of truly novel antimicrobials. In the face of emerging multidrug-resistant microbes, new avenues need to be explored to develop more effective weapons against the threat of microbe-borne infectious diseases. The rapidly developing field known as ‘‘hostdefense peptide’’ biology, by exploring how various life forms employ antimicrobial substances, is offering new hope to solve the problem of microbial resistance [35]. The natural host-defense peptides, e.g., cecropins, defensins, tachyplasmins, magainins, and analogues derived from them, which kill microbes by permeabilizing their cell membranes, but not those of the host’s cell, are a new class of drugs exploiting innate immunity. However, these host-defense peptides are not immune to the common problems associated with peptide drug development, e.g., stability, delivery, and manufacturing costs, rendering their pharmaceutical development difficult. To address the acute need for better antimicrobials, we chose to develop drugs based on the understanding of host-defense processes occurring in macrophages and neutrophils. However, instead of seeking to enhance the drug potential of natural, peptidic defense molecules, entirely novel, nonpeptidic, and fully synthetic pharmacophores were sought. Considering the nature of the reactive species involved in the cytotoxic flux, we hypothesized that simple organic functionalities with oxidoredox properties could modulate microbicidal activity. An example of this modulation is seen with taurine, which is present in large amounts in neutrophils and which is chlorinated to the N-chloro derivative by the neutrophil’s HOCl. Since the chloramine product is a more gentle oxidant than HOCl, it attenuates the microbicidal activity. To attempt modulation of biocidal activity, five different functionalities, but each with oxidoredox properties, were selected as potential pharmacophores against the opportunistic fungal pathogens, Candida albicans, Cryptococcus neoformans, and Pneumocystis carinii: Oxaziridines, by virtue of the properties of their strained ring, are extensively employed in organic synthesis as oxygen transfer agents. Sulfonyloxaziridines are among the most effective oxidizing agents currently available for chiral epoxidations. Nitrones are versatile building blocks employed in constructing heterocycles via 1,3-additions of the nitrone dipole; they form oxaziridines by electrocyclic ring closure, and the oxaziridines can then be deoxygenated by various reducing agents. Mac-

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rocyclic polyamines could be converted to N-chloro compounds in neutrophils or could undergo N-oxidation, and metallomacrocycles could form metal oxo complexes or form adducts with •NO. Currently we are exploring the effect of these pharmacophores on selected fungal pathogens.

IV.

THE OXIDOREDOX CHEMISTRY OF THE NOVEL PHARMACOPHORES

The controlled release of active oxygen species in an exact position in space in an organism and at the exact time is critical for health. Since both high and low levels of these small but vital molecules can be pathogenic, in an attempt to strike a balance between utilization and elimination of them, drug development efforts included the use of extracellular forms of SODs, synthetic SOD mimetics, •NO releasers and scavengers, and a broad range of antioxidant compounds [36]. The radical scavenging function of SODs appears to protect biological macromolecules from oxidative damage. These ubiquitous metalloenzymes have a redox active cofactor to catalyze the one-electron redox cycle required for oxygen dismutation: (1) The use of extracellular forms of SOD enzyme to inhibit oxygen radical– mediated cell injury met with limited success, and its efficacy is controversial [37]. Thus, SOD alone or in combination with catalase has been reported to afford protection from ischemia/reperfusion injury, but recent studies in humans showed inconsistent results, perhaps because protection occurs only in a narrow range of doses. This controversy about the efficiency of SOD enzyme emphasized the need for low-molecular-weight antioxidants, the SOD mimetics. Both in vitro and in vivo studies with nonpeptidic SOD mimetics indicated that they perform better than SOD in inhibiting tissue injury. However, because most SOD mimetics catalyze the removal of superoxide anion alone without affecting the removal of hydrogen peroxide, they offer limited protection from neutrophil-mediated injury [38]. Since both elevated and reduced levels of •NO (necessary for physiological function) result in diseased states, depending on the site/process affected, both • NO donors and •NO quenchers have therapeutic potential. The ever growing family of compounds designed to control biological concentrations of •NO includes the well-known sodium nitroprusside and the sydnonimines that release • NO. Recently, a very interesting •NO releaser, possibly a useful source of •NO in vivo was reported by Lerner’s group [39]. In an antibody-catalyzed retro DielsAlder reaction, nitroxyl is released from an anthracene-HNO adduct; the released

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nitroxyl is then oxidized to •NO in the presence of SOD. Most efforts to reduce • NO levels were in situations in which abnormally high levels of •NO produce severe clinical problems, such as endotoxic shock, responsible for a high death rate in surgical emergency units. Inhibitors of iNOS should have high therapeutic potential to control •NO levels in vivo. The difficulty with approaches to reduce • NO levels that use arginine analogues is that the majority of these studies were carried out using partially purified iNOS, or cells in culture; hence these must be interpreted with great caution. The amino acid moiety is not a prerequisite for either potency or selectivity. Thus, isothioureas, bisisothioureas, and a recently reported amidine derivative showed various degrees of selectivity for human iNOS [40]. However, the major problem is that for a therapeutic application, isoform-selective inhibitors are a prerequisite, and the active sites’ structures in these proteins are not yet known with sufficient precision to permit rational drug design. The strategy proposed here to employ oxidizing agents as potential therapeutics to treat systemic fungal infections can be viewed as problematic because of the concern that the damage they may cause to the host will outweigh the benefits of their microbicidal effects. However, careful selection of the oxidoredox functionalities may allow striking a balance between tissue destruction and microbicidal activity. The rationale for selection of each type of oxidoredox pharmacophore is outlined below. A.

Sulfonyloxaziridines

Sulfonyloxaziridines, known for two decades, are among the most versatile in the repertory of oxidizing agents [41]. They are best known as the reagents of choice for chiral oxidations, but it is other aspects of their chemical characteristics that work in their favor in the context of pharmacophore development. Because the oxygen in sulfonyloxaziridines is more electrophilic than is the oxygen in oxaziridines, sulfonyloxaziridines, with few exceptions, are more reactive to nucleophiles than are oxaziridines. Oxygen transfer by sulfonyloxaziridines (Scheme 2) involves S N 2-type displacement of the sulfimine, facilitated by a relatively weak oxygen–nitrogen bond and by the favorable enthalpy of the C N π bond formed, with a slight bias in favor of a planar rather than a spiro transition state. Their reactivity is modulated by the substituents attached to the carbon and

Scheme 2

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nitrogen, those attached to the nitrogen having the greater effect. However, in the oxygen transfer reactions of interest here, the dominant factor appears to be steric, not electronic. Thus, although most sulfonyloxaziridines are highly reactive with nucleophiles, camphorsulfonyloxaziridine does not react with amines. Sulfonyloxaziridines rapidly oxidize sulfides to sulfoxides, convert disulfides to thiosulfinates at room temperature, and oxidize thiols via sulfenic acid intermediates [42]. Tertiary amines are oxidized to N-oxides, secondary amines to hydroxylamines and nitrones, and primary amines to nitroso compounds (Scheme 3) [43]. Of particular interest is the sulfonyloxaziridines’ enhanced reactivity to biologically important nitrogen and sulfur nucleophiles. We hypothesized that these reactions may be important for modulation of the oxidoredox species in vivo and that they might be equally relevant to the pharmacological action of sulfonyloxaziridines. In humans, the hepatic flavooxygenase plays a crucial role in the detoxification of natural and xenobiotic amines and sulfides by N and S oxidations; thus N-oxides are apparently the straightforward metabolic reaction products of tertiary amines. It is interesting that the N-oxide metabolites that sulfonyloxaziridines might produce, were they to react with biological amines, are the same as the metabolites formed in vivo. However, the enzymatic reactions are reversible by reductases that are able to deoxygenate the N-oxide metabolites back to the amines. If sulfonyloxaziridines react with biological secondary amines to form hydroxylamines and/or nitrones, these products too might act as oxidoredox modulators (as discussed in the next section). In model reactions, when two equivalents of sulfonyloxaziridine are employed to oxidize secondary amines, nitrones are produced almost exclusively. Nitrones are known metabolites. Human hepatic flavomonooxygenases produce them in detoxifying hydroxylamines (Scheme 4).

Scheme 3 Oxidation of sulfur and nitrogen nucleophiles by arylsulfonyloxaziridines, or by flavoenzyme mixed function oxidase. (Source: From Ref. 43.)

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Scheme 4 Conversion of hydroxylamines to nitrones in enzymatic and model reactions. (Source: From Ref. 43.)

Since sulfonyloxaziridines transfer an oxygen atom to the same types of substrates as enzyme systems do, they should be excellent oxygenase mimics. Marked interest in the mechanisms of biochemical reactions of peroxidases, catalases, and cytochrome P-450s that modulate oxidant levels in vivo spawned numerous studies on model systems in which various oxidants interact with the metal centers of heme and nonheme prosthetic groups. This led to a consensus that a chemical process similar to the ‘‘peroxide shunt’’ of cytochrome P-450 enzymes takes place when peroxidases react with H 2 O 2, alkylhydroperoxides, or percarboxylic acids. In each case, the key intermediate is an enzyme-bound ironoxo(IV)-porphyrin π-cation radical species [44,45] containing an Fe(IV) O unit at the iron center. In horseradish peroxidase, this iron-oxo species can react with a number of electron donors, including amines and sulfides, before returning to the Fe(III) state. However, in the case of chloroperoxidase and the myeloperoxidase of the neutrophils, this iron-oxo species reacts with chloride to yield HOCl. Thus, if sulfonyloxaziridines were competent to generate iron-oxo intermediates, these in turn would increase the levels of the oxidizing species of neutrophils. Unfortunately, most of the model studies were focused on the interaction of H 2 O 2 and other peroxides with the Fe(III) or Mn(III) centers in porphyrins [46] or employed various other auxiliary oxidants in combination with metal centers, but these did not include oxaziridines. Yuan and Bruice [47], however, reported studies in which a sulfonyloxaziridine was employed as an oxene transfer agent to manganese (III) tetraphenylporphyrin chloride in the catalytic epoxidation of alkenes. This lends support to our hypothesis that the sulfonyloxaziridine pharmacophore may act by forming an oxene, which serves to regulate the levels of biocidal oxidoredox species. Considering the remarkable reactivities of sulfonyl-

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Scheme 5

oxaziridines, together with their antifungal properties reported here, it is anticipated that studies in progress will help to determine whether oxo-transfer reactions contribute to the antifungal effects or whether other mechanisms operate. Since both photochemical and thermal rearrangements of oxaziridines yield isomeric amides, it is tempting to speculate whether sulfonyloxaziridines undergo similar types of rearrangements to form the corresponding sulfonylamides. A precedent for this could be the isolation of N-(p-chlorophenylsulfonyl)acetamide by Davis et al. [48] from an unsuccessful attempt to oxidize a sulfonylimine to the sulfonyloxaziridine (Scheme 5). Although the major first-pass metabolites of the sulfonyloxaziridine functionality are likely to originate from the sulfonylimine, formed by loss of the active oxygen, the occurrence of rearrangements to sulfamide-containing compounds in vivo should lead to metabolites with interesting pharmacological properties [49]. B.

Oxaziridines

Despite an idea advanced in the early seventies that the critical oxygenating species of flavin oxygenases [50] is an oxaziridine, before our studies [51] there were no efforts to explore the pharmacophore potential of the oxaziridine functionality. These oxygenases are unique in that they carry out the only known non-metalion-requiring oxygen activation reactions in biological systems. They bind and activate molecular oxygen, ultimately transferring one oxygen atom to substrate and releasing the second as water. On the basis of mechanistic studies of these enzymes, in 1974 Orf and Dolphin [50] proposed an oxaziridine intermediate as the monooxygenating species, derived by rearrangement of an initial intermediate, 4α-hydroperoxyflavin (Scheme 6). Working with nonenzymatic models, Rastetter et al. [52] proposed that a nitroxyl radical derived from Dolphin’s oxaziridine (Scheme 6) is a viable candidate for the oxygenating species. However, many other mechanisms were proposed later, not involving the oxaziridine, and these received more attention [43]. The lack of interest in possible biological roles for oxaziridines can be attributed to the fact that at the time of the mechanistic proposals discussed not much was known about the chemical properties of of oxaziridines. Despite the substantial number of papers published since then, many details in the mechanism

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Scheme 6 Oxaziridine [50] and nitroxyl radical [52], the putative oxygenating species in flavin monooxygenases.

of oxygen transfer from structures where the active site oxygen is part of a threemembered ring, e.g., in metal peroxides, dioxiranes, oxaziridines, and sulfonyloxaziridines, remain obscure. The similarity in the active site structures, however, suggests that they may have a common mechanism of oxygen transfer, an S N2type displacement by the nucleophilic substrate (Z) on the electrophilic oxygen atom (Scheme 7). Kinetic studies of deoxygenation of oxaziridines and sulfonyloxaziridines [53], epoxidation of alkenes by sulfonyloxaziridines [54], and theoretical calculations on the transfer of oxygen from oxaziridines to ethylene [55] support this mechanism. Oxaziridines are active oxygen compounds. They transfer their ring oxygen atom to tertiary phosphines and oxidize HI, but they are considered to be poor reagents to oxidize sulfides to sulfoxides, or to epoxidize alkenes. However, their reactivity is strongly dependent on the substitution pattern of the oxaziridine. Thus, a bis-oxaziridine oxidized thiacycloalkanes to sulfoxides in only 5%–7% yields, but an N-tert-butyloxaziridine converted dimethylsulfide to the sulfoxide quantitatively [41]. An oxaziridine substituted with electron withdrawing groups, 2-(trifluoromethyl)-3,3-difluorooxaziridine, epoxidized alkenes under extremely mild conditions (⫺50°C, ⬃1 h), and perfluorodialkyloxaziridines performed hydroxylation of unactivated tertiary aliphatic C-H bonds at room temperature, in high yields [56,57]. Moreover, aza-aromatic N-oxides, when irradiated with ultraviolet (UV) light, transfer their oxygen atom. It was proposed that oxaziridines, formed by photoisomerization of the N-oxides, are the oxygen transfer agents,

Scheme 7 S N2-type mechanism of oxygen transfer.

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and the loss of oxygen is facilitated by rearomatization of the heterocycle [58]. Oxaziridines react with ferrous salts, leading to products resulting from homolytic cleavage of the oxaziridine ring. Their broad range of reactivities and their potential to react with nitrogen and sulfur nucleophiles, ubiquitous in biological systems, make oxaziridines attractive candidates for modulating the oxidoredox processes occurring in phagocytic cells.

C.

Nitrones

Nitrones could directly modulate the levels of oxidoredox species important in the biocidal action of macrophages and neutrophils by preferential trapping of one or more of the biocidal species. However, it is more likely that a complex series of events takes place, in which nitrone-derived radicals and/or nitronederived hydroxylamines are major contributors to the biocidal effects: 1.

Nitrones, by undergoing ring closure to the corresponding oxaziridines, could act as precursors of these efficient oxygen transfer agents discussed in Section IV.B.

2.

Nitrones are well known spin traps [59]. Depending on their structure, they can react with C-, O-, N-, and S-centered radicals, including those produced by macrophages and neutrophils, to modulate their in vivo concentrations (Scheme 8). Recently, efficient spin trapping of O-, C-, and S-centered radicals and peroxynitrite, but not superoxide, was reported when using 2H-imidazole-1-oxides as spin traps [60]. Because the radical trapping reactions of nitrones yield stable nitroxyl radicals, these radicals might participate in controlling the levels of oxidoredox species present in vivo by serving as traps for short-lived radicals. On the other hand, stable nitroxyl radicals are SOD and catalase mimics,

Scheme 8

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Scheme 9

and hence they are potential agents of cytoprotection against postischemic reperfusion injury [61]. Heteroaromatic N-oxides, some reactions of which resemble those of nitrones, are currently being evaluated as hypoxia-selective cytotoxins in the clinical treatment of solid tumors [62]. It is believed that the tumor selectivity is due to both tumor hypoxia and the expression of high levels of reductive enzymes. It has been suggested that nontoxic, free radical reduction products of the heteroaromatic N-oxides are involved in the mechanism of cytotoxicity. Further, it was proposed that these radicals are toxic to the hypoxic cells, but the drug, D, is restored in the well-oxygenated tissues (Scheme 9). However, no mechanism was proposed to explain how the N-oxides would inflict damage on the target, except that drug is likely to abstract a hydrogen from the sugar residue of DNA, generating a sugar radical. 3. We hypothesized that various reductants that are abundant in biological systems could convert the nitrones to nitroxyl radicals and to hydroxylamines (Scheme 10). The hydroxylamines produced by reduction of

Scheme 10

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4.

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nitrones could react with superoxide radicals, and with the peroxynitrite from macrophages. In support of this, very recently new spin traps, TEMPONE-H and CP-H, containing hydroxylamine moieties were reported to trap efficiently both superoxide radical and peroxynitrite, yielding nitroxyl radicals [63]. VBN-3, a bisnitrone, is highly active against P. carinii, both in vitro and in vivo. Among the several mechanisms by which VBN-3 could work in vivo, two pathways are shown in Scheme 11. According to pathway a, reductive activation would yield a diradical. This could release two •OH/mol of VBN-3, and cause the antifungal effect. The antifungal properties of heteroaromatic N-oxides were attributed to their ability to produce hydroxyl radicals that are known DNA cleavers, causing oxidative damage to the sugar–phosphate backbone of DNA [64]. The DNA damage may also occur as a result of H• abstraction by the diradical initially formed via pathway a. According to pathway b, oxidative activation would yield the nitronyl nitroxide, a diradical. Model experiments showed this radical can be obtained by slow air oxidation of VBN-3, as a purple, crystalline, and stable material. In vivo, oxidants of neutrophils, for example, H 2 O 2, could accomplish this step. The nitronyl nitroxide should readily react with •NO, since other nitronyl nitroxides have been employed to detect •NO in air [65]. The reaction with •NO would yield the imino nitroxide. The imino nitrogen in imino nitroxides has a pronounced basic character and can be reversibly protonated; the nitroxide is reducible, for example, by ascorbate or SH groups, to the corresponding hydroxylamine. Reduction by SH groups, however, is still controversial. The nitronyl nitroxide could also be derived from pathway a, by dehydration/oxidation steps (Scheme 11). The bond energy for the OH bond in hydroxylamines is relatively low (⬃70 kcal/mol), and therefore reoxidation to the nitroxides can be done with a variety of oxidants. In vivo, the hydroxylamines could be reoxidized to nitroxides, for example, by flavomonooxygenase (FlOOH), by a metal-catalyzed reaction that proceeds through superoxide, or by myeloperoxidase (MPO) oxidants of the neutrophils. Simple nitroxide spin traps, in which the nitroxide is contained in five-membered and six-membered rings, protect mammalian cells from oxidative damage; hence they appear to function as SOD mimics. Although the SOD activities of the nitroxides tested to date are orders of magnitude lower than that of SOD, their low toxicity and high cell permeability might permit employing them at higher concentrations to enhance their SOD activity. It is interesting to speculate whether the nitroxides derived from VBN-3 types of compounds (Scheme 11) might also confer SOD activity on them.

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Scheme 11 VBN-3.

D.

Pathways postulated for (a) reductive and (b) oxidative activation of

Macrocyclic Amines

The rationale for selecting macrocyclic amines as potential microbicidal pharmacophores is based on consideration of the reactions of amines with the potent oxidants produced by macrophages and neutrophils. The macrocyclic structure, as carrier of the amine functionalities, was chosen because such structures can

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incorporate several amine moieties within the same molecule, thereby providing high local concentrations of the pharmacophore, as in the ‘‘respiratory burst’’ characterized by high local concentrations of the reactive species. Further, we envisaged that the macrocyclic structures will facilitate partition of the compounds into membranes, and we have conducted preliminary toxicity studies to select macrocyclic frameworks devoid of toxicity to human cell lines. The mechanisms of amine oxidation by peroxynitrite or peroxynitrous acid have been controversial. Peroxynitrous acid is both a one- and a two-electron oxidant, but two-electron oxidations (that is, oxygen transfer) of amines by HOONO have been observed experimentally [66].

From the limited set of macrocyclic amines tested to date, it is not likely that N-oxidation of tertary amines confers antifungal activity. Another possibility is that biologically abundant metal nitrosyl complexes acting as electrophilic nitrosating agents transfer NO⫹, thus converting the amines to N-nitrosamines. However, the biological relevance of NO⫹ under physiological conditions has been disputed [67]. An alternate, and most likely explanation for the mode of action of macrocyclic amines is that they function as prodrugs; that is, they are converted by the HOCl of neutrophils into N-Cl derivatives, potent and longlasting oxidizing agents. It is interesting to note that two marine alkaloids, papuamine and haliclondiamine, containing unusual 13-membered macrocyclic rings encompassing 1,3-diaminopropane moieties, display significant antifungal activity against C. albicans. However, the recent total synthesis of papuamine by Weinreb’s group [68] required 16 steps, whereas the macrocyclic imines reported here, highly active against C. albicans, can be obtained from commercially available starting materials, in 2 steps, in ⬎70% overall yields.

E.

Metallomacrocycles

The rationale for employing metallomacrocycles as antifungals was based on consideration of the reactivities of their biological counterparts, as follows: 1.

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Peroxidases contribute to the modulation of the physiological levels of microbicidal oxidants. They have iron-porphyrin cofactors, and when activated, their active site structures include the FeIV ⫽ O unit responsible for oxygen atom transfer to substrates. The electron needed to complete the oxygen’s octet is drawn from the porphyrin, to yield porphyrin⫹•, or from an active site amino acid residue, to yield residue⫹•

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(Scheme 12). For example, horseradish peroxidase and catalase contain porphyrin⫹•, but cytochrome c peroxidase contains residue⫹•. Besides these well-characterized oxo-metal species, numerous catalytic systems mimicking the oxygen transfer step have been described. The macrophages’ iNOSs, akin to the P-450 enzymes, are also well characterized. They have their heme coordinated to the apoprotein through an axial thiolate ligand. The molecular events involved in •NO production and ligand release from the metalloporphyrin-NO complexes in iNOS are known to a great extent. In contrast, although a large amount of information on the parameters of activation of the neutrophil NADPH oxidase complex has accumulated during the past few years, the heme active site structure of its O •⫺ 2 generating redox core, flavocytochrome b, remains elusive. However, an interesting observation was made in connection with the activation of the NADPH oxidase. It was reported that the endogenous oxidative activity of macrophages can be enhanced by peroxidases [69]. Thus, when horseradish peroxidase was added to macrophages, it enhanced their phagocytic and oxidative activity, even though some of the peroxidase added was lost by proteolysis. This implied that the heme of the peroxidase might be transferable to the cytochrome b of NADPH oxidase, to activate the latter. This raised the question of whether synthetic metallomacrocycles might be employed, instead of the endogenous hemes, in drug design to enhance the microbicidal effects of macrophages and neutrophils. 2. Further impetus to test whether synthetic metallomacrocycles might have microbicidal activity was provided by consideration of the high reactivity of certain hemoproteins to •NO. Reaction with protein cysteine residues yields nitrosothiol adducts, and reaction with vicinal thiols generates disulfide bonds. Reaction with a ferrous heme center, as in deoxymyoglobin, yields Fe 2⫹-N O, that is, nitrosomyoglobin. When • NO reacts with the superoxide form of bound oxygen, Fe 2⫹ Oδ⫹-Oδ⫺, as in oxymyoglobin, •NO diffuses into the heme pocket first, then reacts

Scheme 12

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rapidly to give Fe 3⫹-OH2, that is, metmyoglobin and nitrate [70] (Scheme 13). Ferrihemes also react with •NO. Thus, metmyoglobin, ferricytochrome c, and catalase bind •NO reversibly, and methemoglobin reacts with it irreversibly to yield nitrosohemoglobin [71]. The reactivity of ferriheme nitrosyl adducts can be rationalized in terms of the charge transfer from the •NO to the metal to give a structure best described as Fe II ⋅ ⋅ ⋅ ⋅ NO⫹. This makes the nitrosyl more electrophilic, and susceptible to attack by a variety of biological nucleophiles. Other important reactions of •NO with hemes relevant to the antifungal effect of metallomacrocycles are as follows: Binding of excess •NO to the heme center of iNOS deactivates the enzyme, thereby providing an unusual example of a self-regulating enzyme whose products deactivate it. Binding of •NO to the soluble guanylate cyclase’s b type heme activates this receptor to produce cGMP [72]. Removal of the heme results in loss of ability of •NO to activate the enzyme, whereas reconstitution of the enzyme with its heme partially restores the ability of • NO to activate it. Ignarro, in the early 1980s, demonstrated that purified heme-deficient soluble guanylate cyclase can be activated by preformed nitrosyl–heme complexes, and also by protoporphyrin IX [73]. He proposed that •NO activates soluble guanylate cyclase by binding directly to the heme to form a 5-coordinate nitrosyl-heme complex [74]. On the basis of the preceding considerations, it is likely that addition of surrogate prosthetic groups might activate the otherwise latent microbicide-producing enzymes, such as NADPH oxidase and iNOS, to spur production of endogenous oxidants. However, it is also possible that by forming surrogate heme–oxidant complexes, better oxidants,

Scheme 13

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that is, microbices, become available in vivo. The synthetic metallomacrocycle, VBN-10, is highly active against P. carinii, both in vitro and in vivo. Mode of action studies will establish whether its activity is related to the formation of metal-oxo or metal-nitrosyl complexes in vivo. 3. Remarkable findings about •NO binding to hemoglobin were recently reported by Stamler’s group [75]. They demonstrated that S-nitrosothiol formation endows hemoglobin with hitherto unsuspected allosteric and electronic properties, which enable it to control tissue oxygenation levels. Specifically, when oxygen is bound to heme iron, this binding promotes S-nitrosylation of cysteineβ93 by •NO. Deoxygenation triggers an allosteric transition from the oxygenated (R) state to the deoxygenated (T) state and concomitant release of •NO. The R state of the protein contracts blood vessels; the T state relaxes them. Thus, hemoglobin senses the oxygen gradient in tissues via conformational changes and ensures that blood flow is in line with oxygen requirements. In hypoxic tissues •NO is released, but if oxygen supply exceeds demand, the •NO is retained by the hemoglobin in the R structure with the net effect of reducing blood flow. These newly discovered properties of hemoglobin will have far reaching therapeutic applications, not limited to the control of blood pressure. The mode of action of any drug candidate having structure enabling it to affect •NO levels must take into account the •NO shuttle of hemoglobin. This is especially relevant to those novel pharmacophores we are developing whose mode of action might be tied to •NO levels, or to levels of oxidants connected to it. It will be important to evaluate how the high, local concentrations of •NO, or other oxidants produced by macrophages and neutrophils in response to pathogen attack, fit the new image of the • NO shuttle.

V.

THE SYNTHESIS OF ANTIFUNGAL DRUGS CONTAINING OXIDOREDOX PHARMACOPHORES

Of the five pharmacophores selected for synthesis, at least one compound in each class showed high activity against P. carinii, except the macrocyclic polyamines, which showed activity only against Cryptococcus neoformans and Candida albicans. Further, we recently established, in the case of the metallomacrocycle pharmacophore VBN-10, active against P. carinii, that the in vitro activity closely correlates with in vivo efficacy. Here we summarize the syntheses of the lead compounds and their respective activities:

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Scheme 14 a, Amberlyst 15/toluene/reflux, 16 h; 47%; b, m-CPBA/aq. NaHCO3CH 2 Cl2 /BTEAC, 0°–2°C; 92%.

A.

Synthesis of Sulfonyloxaziridines

Finding novel structural classes, hitherto not employed as therapeutic agents, is usually difficult and seldom attempted. We overcame this hurdle by synthesizing the lead compounds, a sulfonyloxaziridine podand, VBN-1, and the macrobicyclic oxaziridine, VBN-2 (see Section V.B). However, since these are truly new structural leads and thus have not been employed in any drug development, it is necessary to determine which types of molecular structures are best as vehicles for delivery of the active pharmacophore. Podand VBN-1 contains two sulfonyloxaziridine moieties; it was obtained in 92% yield by m-CPBA (meta-chloroperbenzoic acid) oxidation of the corresponding sulfonimine [76] in the presence of a phase transfer catalyst [77] (Scheme 14). It is a stable white solid that can easily be obtained on ⬎10 g scale. The two oxaziridine moieties in VBN-1 have identical geometry, as ascertained by spectral data. In vitro, VBN-1 was active against P. carinii, at a concentration of 2.1 µM, comparable to that of pentamidine. In vivo testing in mice indicates it has no gross toxicity at 50 mg/kg/day intraperitoneally the highest concentration tested to date. B.

Synthesis of Oxaziridines

Since oxaziridine as a pharmacophore was unknown prior to our studies, in the first attempt to establish its usefulness against P. carinii we prepared a compound containing several oxaziridine units, so as to have as many active oxygens as possible delivered per mole. We were aware of the formidable difficulties, since one can theoretically obtain a large number of isomers. To gain some control over the stereochemistry we chose a rigid macrocyclic framework as carrier for the oxaziridine groups. The hexaimino macrobicyclic precursor of VBN-2, which the spectral and x-ray crystallographic analysis showed to contain six trans-imines [78], was deemed to be a good candidate. Inspection of its structure generated with MacroModel and those of two related hexaimino macrobicycles allowed us to visualize the likely mode of attack of the peracid on these imines. Even though the initial trans-geometry of the imines is essential, this alone was not expected to provide stereocontrol over the production of the oxaziridines, since in nonmac-

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rocyclic analogues oxaziridine ratios obtained were independent of the composition of the initial imine mixtures [79]. However, if one considers a BaeyerVilliger-type mechanism in which the peracid adds to the imine to form the tetrahedral intermediate, which then affords the oxaziridine by nucleophilic attack of the nitrogen on the electron-deficient oxygen atom, then blocking attack by the peracid on the C N bond from the ‘‘endo’’ direction by the macrocycle’s framework should favor the production of only one oxaziridine diastereomer. Examination of the structure of the hexaimino macrobicycle indicates this might be the case. Further, it was reasonable to assume that N-O bond formation to yield the oxaziridine versus C-N bond rotation will be strongly influenced by the macrocyclic framework. The molecular modeling studies employed to select the structure of a macrobicyclic imine precursor were also helpful in the interpretation of the nuclear magnetic resonance (NMR) spectra of the macrocycles. For example, in the hexaimino macrobicycle prepared from 1,3-benzenedicarboxaldehyde, one group of aromatic protons appeared at an unusually high field (5.3 ppm). We used MacroModel to explain this observation and to confirm the structure. It showed that in the energy-minimized conformation, the molecule is highly symmetric. The high-field proton on each aromatic ring points directly to the center of an another aromatic ring, thus explaining this shielding (Scheme 15). We synthesized macrobicyclic oxaziridine, VBN-2, by m-CPBA oxidation of a hexaimino macrocycle in the presence of BTEAC (benzyltriethylammonium chloride), a phase transfer catalyst [51]. Starting material, the macrocyclic Schiff base, was obtained by template-free [3 ⫹ 2] condensation of terephthaldicarbox-

Scheme 15 Energy-minimized conformation of a hexaimino macrobicycle containing 1,3-disubstituted aromatic rings.

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aldehyde with tris(2-aminoethyl) amine in excellent yield (88%, Scheme 16). Two other groups have also synthesized this hexaimino macrocycle, using the template-free route, in yields of 50% [80] and 80% [81], respectively. Selecting the suitable solvent was critical; in CH 3CN as solvent, oligomers/polymers were obtained instead of the desired [3 ⫹ 2] adduct [51,80]. The structure of the metalfree macrocyclic Schiff base was determined by x-ray crystallography [78], which showed a trans-arrangement for all the imine functions, a divergent (uncoordinat˚ diameter. Oxidation ing) arrangement of the N donors, and a tiny cavity of 4A of the hexaimino macrocycle, using Davis’s method [82], gave VBN-2, the first member of a new family of macrocycles carrying oxaziridine groups. VBN-2 transfers six oxygen atoms to triphenylphosphine quantitatively. Molecular modeling using MacroModel along with 1 H and 13 C NMR spectroscopic analysis [51] indicated that the molecule is highly symmetric and that all oxaziridine moieties have E geometry. The 1 H nmr of the VBN-2 consists of two sharp singlets in the low-field region, indicating that all aromatic protons are equivalent, as are all six oxaziridine protons; the methylene protons are well resolved. Oxaziridine isomers have diagnostic values for the oxaziridine proton in the E isomer at δ 4.6–4.7 ppm, whereas in the Z isomers this proton is deshielded by the nitrogen lone pair to δ 5.3–5.4 ppm. Conformational motions are limited in VBN-2, as shown by the well-resolved methylene signals. Thus, in VBN-2 all oxaziridines are E, and hence control of stereochemistry by using the macrocycle is evident. A related macrobicyclic Schiff base that has a larger cavity size and that, because of one additional methylene group in each bridge, is more flexible than the precursor of VBN-2 gave a complex mixture of products, from which no oxaziridine could be isolated. There are now many examples that apply a similar control of threedimensional (3D) space, via macrocycles or molecular recognition constructs, to achieve desired stereochemistry. An interesting example is an extended aromatic

Scheme 16 a, Anhydrous EtOH, reflux/3 h/N2; 88%, b, m-CPBA/CHCl3 /aq. NaHCO3 /BTEAC/0°–2°C, 6 h; 25%.

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shelf (naphthyl group) that acts as an impenetrable barrier to the approach of the reagent from the bottom of the structure [83]. Toxicity studies in human cell lines, CEM-SS and Jurkat, were reported recently [77]. VBN-2, with an in vitro activity of 1.5 µM against P. carinii, is close in activity to pentamidine, thus demonstrating the potential of the novel oxaziridine pharmacophore (Table 1, Sec. VI). The Indiana University culture method [84] demonstrated that VBN-2 at 2 µg/ml decreased growth by 50% compared to that of untreated cultures, suggesting that VBN-2 may be effective against P. carinii in vivo. C.

Synthesis of Nitrones

During our studies on retinoids, we synthesized highly conjugated open chain nitrones [85] and found that this conversion reduced the teratogenicity of retinoids [86,87]. However, we decided to try a less conjugated molecule containing two nitrone groups and an aromatic ring to employ as a pharmacophore against P. carinii. Bis-(amidine oxides) were very attractive as target structures for testing the nitrone pharmacophore. Nitrones are known to react with the biologically important radicals, which we believe to be particularly relevant to the inhibition • of P. carinii (e.g., •OH, •OOH, O ⫺• 2 , and NO), and are thereby themselves converted to more stable aminoxyl (nitroxyl) radicals. These aminoxyls elicit heightened interest as potential therapeutic agents, spin traps, and synthetic intermediates [88]. The aminoxyl radicals generated from the reaction of nitrones are in themselves of considerable interest because their drug potential is enhanced, as evidenced by the observation that they are typically quite nontoxic in vivo [59]. Nitronyl nitroxides were also of interest because they react specifically with nitric oxide [89], an important product of iNOS in macrophages, to yield imino nitroxyl radicals. We prepared the target nitrones, VBN-3 and VBN-4, and nitronyl nitroxides, VBN-3-ox and VBN-4-ox, by condensation of 2,6-pyridinedicarboxaldehydes and 1,4-benzenedicarboxaldehydes with 2,3-bis(hydroxylamino)-2,3dimethylbutane sulfate followed by oxidation of the condensation products according to the procedure of Ullman [90,91]. In the case of 2,6-pyridinedicarboxaldehyde, yellow crystals of two products, VBN-3 and VBN-3′, were obtained in 85% and 5% yields, respectively. The structures of both were ascertained by UV, infrared (IR), 1 H, and 13 C NMR spectra and by MS. Interestingly, these conformers do not interconvert in solution at room temperature and can be readily distinguished by the low-field region 1 H NMR spectra. In VBN-3′ only one of the pyridine protons (C-5H) shows the typical deshielding (δ 9.5 ppm) [92] by the nitrone oxygen, whereas both 3 and 5 protons are deshielded in VBN3 (δ 9.44 ppm). In VBN-3′ the pyridine-3-proton appears at δ 8.21 ppm.

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Bisnitrones VBN-3 and VBN-4, containing the pyridine and 1,4-disubstituted benzene rings, respectively, showed both stability and nontoxicity to Jurkat cell lines. Podand VBN-4 was inactive against P. carinii; it self-associates in solution [93], and this stacking may be the cause of its lack of activity. But VBN-3 showed excellent activity. At the lowest concentration tested, 2.8 µM (1 µg/mL), 79.4% inhibition was observed after 24 h, and after 48 h no live P. carinii could be detected (Table 1, Sec. VI). In vivo tests of this nitrone in mice are in progress at the National Institute of Allergy and Infectious Diseases (NIAID) (Table 4). D.

Synthesis of Macrocyclic Amines and Metallomacrocycles

In order to assess the potential of the target metallomacrocycles as oxidoredox pharmacophores against opportunistic infections (OIs), a prerequisite was the development of synthetic methods by which these macrocycles can be obtained metal-free (template-free) from easily accessible starting materials in reasonable yields. However, traditionally, to prevent formation of oligomeric/polymeric products, the majority of the macrocycles that interest us were obtained via template-directed syntheses. For the planned purpose, the resulting template–ligand complexes would have only limited usefulness in developing efficacious agents against the OIs because the presence of the template would render modification of the structures difficult. Although the ‘‘macrocyclic effect’’ [94,95] would confer enhanced stability to the template–ligand complex, it would at the same time preclude many of the further reactions intended to enhance in vivo efficacy of the macrocyclic structure. Exchange of the template and attempts to dissociate it have met with only limited success. In the few early cases when the macrocyclic rings were obtained template-free, without recourse to a metal template or high dilution techniques, some special circumstances were present. Examples include hydrogen bonding to reduce lone-pair repulsions that would otherwise be dominant within the macrocyclic ring, and successful high yield (⬎90%) syntheses of macrocycles from sterically rigid precursors, which by promoting intramolecular

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Scheme 17 Synthesis of metal-free macrocyclic imines by [2 ⫹ 2] condensations. (Source: From Ref. 77.)

hydrogen bond formation favored ring closure. Lehn was among the first in 1987 [96] to report efficient syntheses of macrocyclic Schiff bases via template-free [3 ⫹ 2] condensations. Since that time, diverse needs for metal-free macrocyclic ligands led to the development of methodologies to obtain them by simple [2 ⫹ 2] and [3 ⫹ 2] macrocyclizations. We synthesized the Schiff base macrocycles, such as the macrobicyclic imine precursor of VBN-2, and a series of iminomacrocycles, such as VBN-5 to VBN-9, and the macrocyclic amide precursor of VBN10 via these nontemplate syntheses [51,77]. These, together with the syntheses of aminomacrocycles screened for antifungal activity against Candida albicans and Cryptococcus neoformans, result to date in over 50 macrocycles made in template-free manner. The success of these syntheses (yields ⬎ 50%), in which we obtained the metal-free macrocycles by direct [3 ⫹ 2] and [2 ⫹ 2] condensations (step a, Scheme 16; Scheme 17), depends on the careful selection of inher-

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ently rigid reactants for condensations. This reduces the conformational degrees of freedom in the reactants/intermediates, thereby optimizing entropy factors that promote cyclization. Further, the target macrocycles are designed with rigid aromatic ‘‘head groups,’’ and some of them, e.g., VBN-5 to VBN-8, also contain diphenylmethane ‘‘hinges’’ that contribute to the observed stability of these Schiff base macrocycles. The precursor of Schiff base macrocycle VBN-2 and imino macrocycles VBN-5-9 were synthesized in high yields (⬎80%), template-free, via [3 ⫹ 2] and [2 ⫹ 2] condensations [51,77]. However, it is important to notice that experimental conditions will vary, depending on the type of macrocyclic structure that contains the Schiff base groups. Thus, Lehn’s group obtained the macrobicyclic hexaimino Schiff bases by simply adding dropwise an acetonitrile solution of the dialdehydes to a solution of tren, [N(CH 2 CH 2NH 2 )3 ], in CH 3 CN and stirring the solution at room temperature for 15 min, when ‘‘a precipitate of almost pure macrobicycle hexaimine was formed’’ [96]. On the other hand, Menif and Martell report [97] a complex NMR spectrum (possibly due to Schiff base isomers) when a tetraimino macrocycle (unlike Lehn’s macrobicycle) was prepared (in 72% yield) using Lehn’s reaction conditions, but longer reaction times. Nelson reports synthesis of Schiff base macrobicyles via direct [3 ⫹ 2] condensations in refluxing alcohol (⬃60% yields), but in this same series of compounds that contain aromatic ‘‘head groups’’ a thiophene-containing one could not be obtained template-free, and in the solvents employed condensation of furan-dicaroxaldehyde with 2,6-bis(aminomethyl)pyridine did not give the macrocyclic [2 ⫹ 2] product [98]. These authors indicate the importance of the solvent employed; thus a trispyrrole BISTREN macrocycle could not be obtained when CH 3 CN was used as solvent. A polymer formed instead. However, with MeOH as solvent a good yield of this cryptand was obtained [98]. In 1993 Smith et al. [99] reported the direct [3 ⫹ 2] synthesis in 50% yield of a hexaimino macrobicyclic Schiff base from condensation of tris(2-aminoethyl)amine (tren) with an aldehyde in the absence of a template ion. This macrobicycle is similar to the precursor macrocycle of VBN-2 (Scheme 16) but is much more flexible. They caution that the ‘‘essential element of this reaction appears to be the slow addition of glyoxal to tren at low temperatures (0°C). Higher temperatures and faster addition rates lead to variable yields, ranging from 10–30%.’’ All these reports demonstrate the interest in template-free syntheses, and the importance of the appropriate reaction conditions. We have synthesized over 19 macrocyclic Schiff bases by [2 ⫹ 2] and 3 macrobicyclic ones by [3 ⫹ 2] condensations, template-free, and in high yields. Trans geometry of the Schiff bases was demonstrated by NMR spectroscopy. The same observation has been reported for similar macrocyclic imines based on x-ray crystallographic and NMR analyses [100,101]. The large ring size (26 and 28 atoms) can easily accommodate trans-double bonds. This versatile methodology was used to afford macrocycles containing different functionalities

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in their framework; thus macrocyclic imines were obtained by condensation of various dialdehydes with diamines and macrocyclic amides by condensation of the diamines with diacid chlorides. In addition, a wide array of ‘‘spacer-arms’’ (the R groups) can be introduced prior to the condensation step, or they can be subsequently attached to the macrocycle. A further built-in feature of this synthetic approach is that it not only allows introduction of functionalized spacer arms but also can be used to obtain other macrocyclic skeletons; e.g., the imino macrocycles can be converted to macrocycles containing amine, oxaziridine, or nitrone groups. Moreover, the headgroups (shaded boxes in Scheme 17) can be varied by changing the carbonyl component. The imino macrocycle VBN-5 with the pyridine head group was toxic to both CEM-SS and Jurkat cell lines. In contrast, macrocycles VBN-6-8 containing furan and 1,3- and 1,4-disubstituted phenyl head groups and VBN-9 were not. The furan headgroup in VBN-6 demonstrated a cell viability of 100%. Cyanoborohydride reduction of the metal-free macrocyclic imines afforded a series of aminomacrocycles in high yields. The ease of the two-step synthesis, yields of ⬎50% in the [2 ⫹ 2] cyclization and almost quantitative yields in the reduction step, as well as the facile derivatization of the spacer-arms, make these macrocycles attractive target compounds. Nineteen aminomacrocycles, obtained by cyanoborohydride reduction of the macrocyclic imines, were screened for antifungal activity [102]. Among these, compound 4 showed outstanding activity against Cryptococcus neoformans at a minimum inhibitory concentration (MIC ⱕ 0.5 µM); compound 11 was fungistatic against Candida albicans at a MIC ⱕ 0.5 µM and also showed modest fungistatic and fungicidal activity against Cryptococcus neoformans strains 1 and 2 (Tables 2 and 3). The metallomacrocycle, VBN-10 (Scheme 18), was obtained by applying the [2 ⫹ 2] condensation procedure developed by us [51]. Thus, in a typical procedure for the preparation of macrocyclic amides (e.g., VBN-10, R ⫽ C 10 H 21), 2,2′-methylenebis(4-nitrophenyl decyl ether) dissolved in aqueous EtOH-THF (ethanol tetrahydrofuran), was reduced with Zn/NH 4 Cl. To the dried, crude reduction product dissolved in absolute EtOH was added the diacid chloride, and

Scheme 18

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Spacer-armed metallomacrocyclic amides.

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the mixture was stirred at rt for 16 h under inert atmosphere. The precipitate of the macrocycle was washed with ethanol and was dried to obtain the macrocyclic amides in yields of 50%–55% (unoptimized). The nickel complex, VBN-10, was obtained by complexation of the deprotonated macrocycle in THF with NiCl2 ⋅ 6H 2 O, 12 h, rt. Complexation with nickel resulted in a new red-shifted absorption band and a new peak in the FAB-MS (fast atom bombardment-mass spectrum). The metallomacrocycle VBN-10 was not toxic to CEM-SS and Jurkat cells and showed potent in vitro activity against P. carinii, causing inhibition of 89.9% and 97.4% (24/48 h) at a concentration of 0.7 µM (Table 1, Sec. VI). Its in vivo efficacy is comparable to that of trimethoprim/sulfamethoxazole (TMP/SMX) (Table 4).

VI.

IN VITRO AND IN VIVO STUDIES WITH DRUGS CONTAINING NOVEL PHARMACOPHORES DIRECTED AGAINST OPPORTUNISTIC INFECTIONS

Assessment of the efficacy profile of therapeutic agents presently in clinical use against OIs indicates an acute need to develop better agents. This can be accomplished either by improving the efficiency of the known therapeutic agents or by obtaining novel types of lead compounds and their analogues by syntheses. We developed such lead compounds on the basis of the oxidoredox hypothesis outlined in the foregoing. The biological test results against the fungal pathogens P. carinii, C. albicans, and C. neoformans are summarized in the following: A.

Activities Against Pneumocystis carinii

In vitro assays against P. carinii were carried out by the NIAID contractor’s laboratory, following the protocol described by Chen and Cushion [103]. A total of 14 compounds were tested against P. carinii isolates from Brown Norway rat lungs with severe infection. The activity of four of these compounds, VBN-1, VBN-2, VBN-3, and VBN-10, 2.1 µM, ⱕ1.5 µM, ⱕ2.8 µM, and 0.7 µM, respectively, was comparable to that of pentamidine, 1.7 µM, the positive control (Table 1). For VBN-2 and VBN-3, no end points were determined (concentrations less than 1.7 and 2.8 µM were not tested); thus the activity of VBN-2 and VBN-3 may well exceed that of pentamidine in vitro. Compounds VBN-4–9 and VBN11–14 showed no appreciable activity against P. carinii. None of the active compounds was toxic to human (Jurkat and CEM-SS) cell lines tested [77]. There are many compounds active against P. carinii, in vitro. These activities, however, do not necessarily predict in vivo efficacy. The rationale for our high expectations of in vivo activity in the case of compounds VBN-1, -2, -3, and -10 is that despite their drastically different structures con-

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Table 1 Evaluation of VBN-1–VBN-14 Compounds Against Pneumocystis carinii a Compound

Untreated

1 µg/mL (% Decrease)

Pentamidine VBN-1 VBN-2

61.6 14.9 12.2 17.5 17.5 17.5 38.5 78.7 78.7 78.7 63.7 63.7 33.3 (0) 30.4 30.4 30.4 30.4

0.6 (99.5) 1.7 µM b 1.3 (91.3) 2.1 µM b ⋅⋅⋅⋅⋅ 3.1 (82.3)/24 h 1.5 µM b ⋅⋅⋅⋅⋅ 3.6 (79.4)/24 h 2.8 µM b 1.0 (97.4) 0.7 µM b 67.2 (14.6) 56.7 (28.0) 77.0 (2.2) 46.3 (27.2) 48.1 (24.5) 33.3 (0) 25.1 (17.4) 30.0 (0) 31.3 (0) 30.4 (0)

VBN-3 VBN-10 VBN-4 VBN-5 VBN-6 VBN-7 VBN-8 VBN-9 VBN-11 VBN-12 VBN-13 VBN-14

10 µg/mL (% Decrease)

100 µg/mL (% Decrease)

0.2 (98.7) 0.2 (98.7) ⋅⋅⋅⋅⋅ ⋅⋅⋅⋅⋅ 2.9 (83.3)/24 h 0.5 (97.3)/24 h ⋅⋅⋅⋅⋅ ⋅⋅⋅⋅⋅ 0.7 (96.0)/24 h 1.5 (95.1)/24 h 1.4 (96.4) 0.5 (96.4) 55.6 (29.4) 0.9 (99.9) 61.8 (21.5) 0.7 (99.1) 58.9 (25.2) 11.1 (85.9) 42.4 (33.3) 0.5 (99.2) 31.4 (50.7) 0.4 (99.4) 33.3 (0) 33.3 (0) 28.8 (15.3) 16.2 (46.7) 28.3 (6.9) 0.9 (97.0) 32.0 (0) 18.3 (39.9) 28.6 (5.9) 5.6 (81.6)

a

ATP ⫻ 10 ⫺9 /10 8 M/10 8 nuclei (ATP decrease %). All results are after 48 h, 24 h results are also given for VBN-2 and VBN-3, where ATP % was not detectable after 48 h. ⋅ ⋅ ⋅ ⋅ ⋅, ATP % below the level of detection at 48 h. ATP, adenosine triphosphate. b For the first four (active) compounds, micromolar (µM) values given are equivalent to the concentrations at 1 µg/mL.

taining unlike functionalities that render them chemically very different, each of these compounds is able to cause oxidative damage to P. carinii; i.e., the only common feature of these compounds, which are structurally so widely dissimilar, e.g., VBN-10, molecular weight of 1412, a metallomacrocycle, and VBN-3, a relatively small, molecular weight (MW) ⬍ 400 aromatic entity, is that all are oxidizing agents. This gives credence to our hypothesis, the basis of our design of these compounds, namely, that P. carinii is susceptible to oxidative damage. The in vivo studies discussed in Section VI.C confirmed this. B.

Activities Against Cryptococcus neoformans and Candida albicans

During the course of the synthesis of oxidoredox pharmacophores, we encountered several compounds that, although inactive against P. carinii, were active against C. neoformans and C. albicans. Among the 14 compounds tested initially

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Table 2 In Vitro Assays Against Cryptococcus neoformans Strains 1 and 2 MIC 48/72 h (µg/mL) Analogue 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

MLC 48/72 h (µg/mL)

Formula

1

2

1

2

C 28 H 38N6O4 C 32 H 46N6O4 C 28 H 46N6O4 C 32 H 54N6O4 C 34 H 50N6O4 C 28 H 38N6 C 40 H 46N6 C 30 H 42N6 C 44 H 54N6 C 32 H44N4O6 C 28 H 46N6 C 46 H 50N6 C 40 H 38N6 C 44 H 46N6 C 44 H 44N4O 2 C 44 H 48N8 C 26 H 32N4O6 C 16 H 26N4O 2 C 18 H 30N4O4

5/⬎5 6.25/⬎6.25 5/⬎5 ⱕ0.3/ⱕ0.3 ⬎5/⫺ ⬎5/⫺ 5/⬎5 ⬎5/⫺ 5/5 ⬎5/⫺ 0.6/1.25 5/⬎5 5/⬎5 ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺

5/5 6.25/6.25 5/5 ⱕ0.3/0.6 5/5 ⬎5/⫺ 2.5/2.5 5/⬎5 2.5/2.5 5/⬎5 1.25/1.25 1.25/2.5 ⱕ0.3/2.5 5/5 2.5/2.5 1.25/2.5 ⬎5/⫺ ⬎5/⫺ ⬎5/⫺

⬎5/⫺ 6.25/⬎6.25 ⬎5/⫺ ⱕ0.3/0.6 ⫺/⫺ ⫺/⫺ 5/⬎5 ⫺/⫺ ⫺/⫺ ⬎5/⫺ 1.25/2.5 ⬎5/⫺ ⬎5/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺

⬎5/⫺ 6.25/⫺ ⬎5/⫺ 0.6/2.5 ⬎5/⫺ ⫺/⫺ ⬎5/⫺ ⫺/⫺ 5/5 ⬎5/⫺ 1.25/1.25 ⬎5/⫺ ⬎5/⫺ 5/⬎5 2.5/⬎5 5/⬎5 ⫺/⫺ ⫺/⫺ ⫺/⫺

(data not shown), one of the macrocyclic compounds showed encouraging activity in in vitro tests. Since this active compound could be synthesized in two easy steps, and in high yields, using the methodology we developed, we prepared 19 analogues and tested them against these fungal species. The results of the assays of these analogues, compounds 1–19, are shown in Tables 2 and 3. Compound 4 (Table 2) showed activity against C. neoformans strains 1 and 2 at a minimum inhibitory concentration (MIC) ⱕ0.5 µM (ⱕ0.3 µg/ml). This level of activity compares favorably with that of fluconazole, the positive control, especially, since an end point was not established; i.e., the lowest concentration tested was 0.5 µM. Compound 11 (Table 3) was found to be fungistatic against C. albicans strains A–E at a MIC ⱕ 0.5 µM (ⱕ0.3 µg/mL) and also showed both fungistatic and fungicidal activity, though more modest, against C. neoformans strains 1 and 2 at a MIC of 1.3–2.6 µM (0.6–1.25 µg/mL) (Table 2). Because of the high levels of in vitro activity, the syntheses of both compounds 4 and 11 were scaled-up to gram amounts for in vivo assays on mice. Initial toxicity studies are encouraging, since all mice survived for at least 1 month when daily doses of 15 mg/kg of compound 11 and 10 mg/kg of compound 4 were given

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Table 3 In Vitro Assays Against Candida albicans Strains A–E MIC 24/48 (µg/mL) No. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19

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Formula

A

C 28 H 38N6O4 ⬎5/⫺ C 32 H 46N6O4 6.25/⬎6.25 ⬎5/⫺ C 28 H 46N6O4 ⬎5/⫺ C 32 H 54N6O4 ⬎5/⫺ C 34 H 50N6O4 ⬎5/⫺ C 28 H 38N6 ⬎5/⫺ C 40 H 46N6 ⬎5/⫺ C 30 H 42N6 ⬎6.5/⫺ C 44 H 54N6 ⬎5/⫺ C 32 H 44N6O6 ⱕ0.3/5 C 28 H 46N6 C 40 H 50N6 ⬎5/⫺ 5/⬎5 C 40 H 38N6 5/⬎5 C 44 H 46N6 ⬎5/⫺ C 44 H 44N4O 2 C 44 H 48N8 5/⬎5 ⬎5/⫺ C 26 H 32N4O6 ⬎5/⫺ C 16 H 26N4O 2 C 18 H 30N4O4 ⬎5/⫺

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MLC 24/48 (µg/mL)

B

C

D

E

A

B

C

D

E

⬎5/⫺ ⬎6.5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎6.5/⫺ ⬎5/⫺ ⱕ0.3/5 ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ 5/⬎5 ⬎5/⫺ ⬎5/⫺ ⬎5/⫺

⬎5/⫺ 6.25/⬎6.25 ⬎5/⫺ 5/⬎5 ⬎5/⫺ 5/⬎5 5/⬎5 5/⬎5 ⬎6.5/⬎6.5 ⬎5/⫺ ⱕ0.3/5 5/⬎5 5/⬎5 5/⬎5 5/⬎5 5/⬎5 5/⬎5 5/⬎5 5/⬎5

⬎5/⫺ 6.25/⬎6.25 ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎6.5/⫺ ⬎5/⫺ ⱕ0.3/5 ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ 5/⬎5 ⬎5/⫺ ⬎5/⫺ ⬎5/⫺

⬎5/⫺ 6.25/⬎6.25 ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎6.5/⫺ ⬎5/⫺ ⱕ0.3/5 ⬎5/⫺ ⬎5/⫺ 5/⬎5 ⬎5/⫺ ⱕ0.3/⬎5 ⬎5/⫺ ⬎5/⫺ ⬎5/⫺

⫺/⫺ ⬎6.5/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⬎5/⫺ ⫺/⫺ ⬎5/⫺ ⫺/⫺ ⫺/⫺ ⬎5/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺

⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ 5/5 ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⬎5/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺

⫺/⫺ ⬎6.5/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⬎6.5/⫺ ⫺/⫺ 5/⬎5 ⫺/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⬎5/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺

⫺/⫺ ⬎6.5/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ 5/⬎5 ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⬎5/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺

⫺/⫺ ⬎6.5/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺ ⬎5/⫺ ⫺/⫺ ⫺/⫺ ⬎5/⫺ ⫺/⫺ ⬎5/⫺ ⫺/⫺ ⫺/⫺ ⫺/⫺

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intraperitoneally. Should these structural leads we have identified prove successful in the ongoing in vivo studies, subsequent drug development steps starting with the establishment of pharmacokinetical parameters; e.g., clearance, distribution, bioavailability, will be initiated.

C.

In Vivo Studies of VBN-1, VBN-3, and VBN-10 Against Pneumocystis carinii

In vivo studies were carried out at the NIAID contractor’s laboratory according to well-established protocols. Thus, male C3HeB/FeJ mice, 29 g average weight, were given dexamethasone and a cephradine antibiotic to initiate their latent P. carinii infections, then were submitted to drug therapy at 7 weeks into the immunosuppression therapy. Stock solutions of VBN-1, VBN-2, and VBN-10 were made in dimethyl sulfoxide (DMSO) and were diluted in sterile water. The mice were given daily doses intraperitoneally. The results obtained to date are summarized in Table 4. The efficacy of VBN-10 is comparable to that of TMP-SMX, and it had no gross toxicity up to 50 mg/kg. The table shows efficacy for VBN10 at 50 → 20 mg/kg/day but no activity for VBN-1 and VBN-3 at this dosage. Considering that VBN-1 and VBN-3 even at high concentrations are not toxic to human cell lines, higher dosage levels of VBN-1 and VBN-3 were tested. These in vivo studies now in progress indicate activity at a dosage of 150 mg/ kg/day.

VII.

CONCLUSIONS

The existing drug regimens to treat opportunistic infections, devastating to the immunocompromised, are of only limited use because of their high toxic effects and the pathogens’ ability to develop resistance to them readily. The oxidoredox hypothesis advanced here serves as foundation for a novel therapeutic approach to develop clinically useful alternatives to prevent and cure OIs. Four novel, oxidoredox pharmacophores were selected and incorporated into molecules having disparate structures, to yield four types of lead compounds. The synthesis of the lead compounds entailed development of methodologies that use very few steps; the key steps are template-free [2 ⫹ 2] and [3 ⫹ 2] macrocyclizations. Template-free [3 ⫹ 2] cyclization and subsequent oxidation afforded a novel class of macrocycles having oxaziridine moieties. The in vitro and in vivo antifungal activities of each of the four types of leads indicate the high therapeutic potential of the novel pharmacophores. Thus, the feasibility of generating therapeutic agents based on the oxidoredox hypothesis is demonstrated.

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Table 4 In Vivo Studies of VBN-1, VBN-3, and VBN-10 Against Pneumocystis carinii in Mice Treatment Control steroid

Cyst count (median)

P value

—

2.4 ⫻ 10 6 (2.23 ⫻ 104–1.83 ⫻ 10 7)b 2.62 ⫻ 10 6 (3.57 ⫻ 10 5 –1.23 ⫻ 10 7) 1.701 ⫻ 10 6 (1.34 ⫻ 10 5 –5.90 ⫻ 10 6) 4.15 ⫻ 10 6 (1.56 ⫻ 10 5 –1.09 ⫻ 10 7) 7.37 ⫻ 10 5 (6.70 ⫻ 10 4 –1.54 ⫻ 10 7) 5.18 ⫻ 10 5 (6.92 ⫻ 10 5 –1.30 ⫻ 10 7) 3.27 ⫻ 10 6 (7.82 ⫻ 10 5 –8.15 ⫻ 10 6) 3.90 ⫻ 10 6 (4.47 ⫻ 10 4 –1.23 ⫻ 10 7) 1.65 ⫻ 10 6 (2.23 ⫻ 10 4 –5.85 ⫻ 10 6) 2.23 ⫻ 10 4 (2.23 ⫻ 10 4 –3.57 ⫻ 10 5) 2.23 ⫻ 10 4 (2.23 ⫻ 10 4 –7.15 ⫻ 10 5) 1.34 ⫻ 10 5 (2.23 ⫻ 10 4 –2.55 ⫻ 10 6) 3.13 ⫻ 10 6 (2.23 ⫻ 10 4 –5.00 ⫻ 10 6) 2.23 ⫻ 10 4 (2.23 ⫻ 10 4 –2.23 ⫻ 10 4) 2.23 ⫻ 104 (2.23 ⫻ 10 4 –2.23 ⫻ 10 4)

—

VBN-1

0.1 mg/kg/d IP

VBN-1

10 mg/kg/d IP

VBN-1

50 mg/kg/d IP

VBN-3

0.1 mg/kg/d IP

VBN-3

10 mg/kg/d IP

VBN-3

50 mg/kg/d IP

VBN-10

0.1 mg/kg/d IP

VBN-10

10 mg/kg/d IP

VBN-10

50 → 20 mg/kg/d IP

Pentamidine

10 mg/kg/3 ⫻ wk IM

Atovaquone

100 mg/kg/d PO

Primaquine/clindamycin

2/225 mg/kg/d PO

TMP/SMXd

50/250 mg/kg/d PO

Control Normal a

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Dose regimen a

—

N/S N/S N/S N/S N/S N/S N/S N/S ⬍0.001 ⬍0.05 N/S N/S ⬍0.01 —

Cyst count (geometric mean) 6.13 ⫾ 0.8c (1.35 ⫻ 10 6)d 6.42 ⫾ 0.50 (2.63 ⫻ 10 5) 6.21 ⫾ 0.46 (1.62 ⫻ 10 6) 6.43 ⫾ 0.49 (2.69 ⫻ 10 6) 5.89 ⫾ 0.60 (7.76 ⫻ 10 5) 6.63 ⫾ 0.34 (4.27 ⫻ 10 6) 6.50 ⫾ 0.32 (3.16 ⫻ 10 6) 6.29 ⫾ 0.77 (1.95 ⫻ 10 6) 5.94 ⫾ 0.84 (8.71 ⫻ 10 5) 4.52 ⫾ 0.37 (3.31 ⫻ 10 4) 4.86 ⫾ 0.67 (7.24 ⫻ 10 4) 5.23 ⫾ 0.97 (1.70 ⫻ 10 5) 6.11 ⫾ 0.82 (1.29 ⫻ 10 6) 4.35 ⫾ 0.00 (2.23 ⫻ 10 4) 4.35 ⫾ 0.00 (2.23 ⫻ 10 4)

P value — N/S N/S N/S N/S N/S N/S N/S N/S ⬍0.001 ⬍0.001 ⬍0.05 N/S ⬍0.001 —

IP, intraperioteneal; IM, intramuscular PO, oral; N/S, not significant; TMP/SMX, trimethoprim/sulfamethoxazole, the positive control. b Range. c Mean ⫾ 1 SD. d Antilog.

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ACKNOWLEDGMENTS The author thanks Drs. Nancy A. Roth and Ronald J. Roth for more than the critical reading of the manuscript. She would also like to express her appreciation for the contributions of her coworkers. Financial support from NIH, National Institute of Allergy and Infectious Diseases grant ROI AI39418, is gratefully acknowledged.

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13 A Mechanistic Analysis of CEO Bond Cleavage Events with a Comparison to 3,6-Dideoxysugar Formation David A. Johnson and Hung-wen Liu University of Minnesota, Minneapolis, Minnesota

I. INTRODUCTION The making and breaking of carbon–oxygen bonds are ubiquitous in biological systems by virtue of the organic nature of life. Most processes from energy metabolism to deoxyribonucleic acid (DNA) synthesis involve the formation and/or cleavage of CEO bonds. Understandably, there is a wide variation in the structures of the molecules that undergo this transformation. As expected, a diverse array of mechanisms has evolved to facilitate the CEO bond scission for these different molecules. However, most of the reaction mechanisms appear to display a characteristic feature in that the reaction is initiated by the cleavage of a CEH bond before the target CEO bond is broken. For these enzyme systems, the manner in which this critical CEH bond is labilized during catalysis is a defining property of the mechanism. Some of the enzymes can catalyze the reaction using binding activation without assistance from any cofactor, such as enoyl–coenzyme A (enoyl-CoA) hydratase. Other enzymes, such as diol dehydrase and ribonucleotide reductase, require a highly reactive radical-generating cofactor to break the CEH bond and provide the necessary driving force for the reaction. Various other cofactors are used in different mechanisms, including those that initiate a CEO bond cleavage before a CEH bond is broken. One particular enzyme, cytidine diphosphate-6-deoxy-l-threo-d-glycero-

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4-hexulose-3-dehydrase (E 1), depends on pyridoxamine 5′-phosphate (PMP), an inorganic [2Fe-2S] center, and a separate reduced nicotinamide adenine dinucleotide– (NADH) dependent reductase (E 3, formerly known as CDP-6-deoxy-∆ 3,4glucoseen reductase) to remove the C-3 hydroxyl of CDP-6-deoxy-l-threo-dglycero-4-hexulose irreversibly, leading to the biosynthesis of ascarylose, a 3,6-dideoxy sugar [1]. Abstraction of the pro-S hydrogen at C-4′ of the PMP– substrate complex initiates this deoxygenation. Interestingly, the labile CEH bond that promotes the catalysis in this reaction is part of the cofactor skeleton instead of the substrate. Additionally, the fact that radical intermediates are involved in this transformation makes E 1 the only member in a novel class of hydrolases. With emphasis on the cofactor requirements, this review summarizes the catalytic mechanisms of several characterized enzymes involved with CEO bond cleavage events, whether the reaction is irreversible or not. These systems are later used in a comparative analysis to highlight the singular nature of E 1. To narrow the focus, irreversible CEO bond formations, such as hydroxylations and oxygenations, and hydrolysis reactions (replacement of one CEO bond with another) are not discussed here. For the sake of discussion, each enzyme is placed in a ‘‘mechanistic class,’’ which is defined by the cofactor requirements of the deoxygenation reaction. Although enzyme systems representative of each general mechanistic class are presented as examples, it is not the intent of this review to list exhaustively all enzymes that would fit into each class.

II. GENERAL MECHANISMS OF CEO BOND TRANSFORMATIONS A.

Reactions Catalyzed by Enzymes Without Cofactors

1. Enoyl–CoA Hydratase Enoyl–CoA hydratase (crotonase, EC 4.2.1.17) is a member of the hydratase/ isomerase superfamily [2]. As shown in Fig. 1, it catalyzes the reversible hydration of a ∆ 2,3-unsaturated enoyl-CoA substrate to the corresponding 3-hydroxyacyl-CoA product [3]. This reaction is the second step in the β-oxidation pathway of fatty acid metabolism and is also an important reaction in the catabolism of branched-chain amino acids. Early studies (Fig. 1a) revealed that bond formation/ cleavage at the α- and β-positions proceeds in a concerted manner with a syn stereochemistry [4,5]. In an attempt to explain the capability of this enzyme to abstract the α-H, which has a pK a of 16–20 in free solution, a stepwise process via an enolic intermediate (Fig. 1b) has also been suggested [6,7]. However, more recent studies provided further evidence supporting the concerted mechanism by demonstrating that the 3-hydroxyl leaving group is necessary for cleavage of the

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Figure 1 (a) Concerted model for the β-elimination catalyzed by enoyl-CoA hydratase. Evidence supports the assignment of a glutamate residue as the active-site amino acid responsible for both protonation and deprotonation. (b) Stepwise model for the reaction. In this model, an acidic residue protonates the carbonyl and lowers the pK a of the αproton so a single basic residue can both abstract the proton and protonate the leaving hydroxyl.

CEH bond to occur [8]. In the hydration direction, the driving force of this catalysis has been attributed to the capability of crotonase to polarize the π-electrons of the α,β-unsaturated double bond, thus enhancing the electrophilicity of the β-carbon [9]. Consequently, the corresponding active-site acid/base pair plays a ‘‘push–pull’’ role in the reverse (dehydration) reaction and promotes the α-H abstraction and the β-OH cleavage. Sequence alignments with other members of this family implicated a highly conserved glutamate (Glu) residue (Glu164 in rat liver enzyme) as the catalytic residue. In fact, substitution of this residue with glutamine (Gln) led to a dramatic decrease of k cat by more than 100,000-fold [10]. Since the K m remained essentially unaffected by the mutation, it was concluded that Glu164 assumes the role as the active-site base to abstract the α-proton. A similar conclusion was reached in studying the enoyl-CoA hydratase activity associated with the large α-subunit of the multienzyme complex (79 kDa) of fatty acid oxidation of Escherichia coli [11–14]. Again, a glutamate residue (Glu139) was identified as the catalytic residue by site-directed mutagenesis [15]. Although additional work is necessary to elucidate the mechanistic details, sufficient data have been collected to establish that the hydratase activity in this multifunctional protein proceeds through a general acid–general base mechanism with no required cofactors.

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2. β-Hydroxydecanoylthioester Dehydrase A similar reaction can be found in the fatty acid biosynthetic pathway (Fig. 2). This reversible dehydration in the anaerobic pathway of unsaturated fatty acid biosynthesis in bacteria is catalyzed by β-hydroxydecanoylthioester dehydrase (EC 4.2.1.60, encoded by fabA), which is a tetrameric protein and contains no cofactors [16]. Its catalysis consists of two parts: the dehydration of (R)-3-hydroxydecanoyl-ACP (acyl carrier protein) (1) to (E )-2-decenoyl-ACP (2), and the subsequent isomerization to (Z )-3-decenoyl-ACP (3). Extensive studies have shown that the dehydration is a syn process [17], whereas the isomerization is a suprafacial allylic rearrangement [18]. Together, these results strongly imply an active-site topology that includes a single, acidic/basic residue [19], which is responsible for deprotonation and protonation at specific sites. Mechanism-based inactivation and site-directed mutagenesis experiments have identified His70 (numbering of the E. coli enzyme) as an essential active-site base [20–23]. The participation of a histidine residue (His70) in catalysis was further supported by the newly released x-ray structure of the E. coli enzyme [24], however, the xray data implicate the additional participation of an aspartate (Asp84′), a cysteine (Cys80), and a glycine (Gly79) in the reaction mechanism. It is proposed that Asp84′ acts as a second base and both Cys80 and Gly79 assist the stabilization of the anionic intermediate via hydrogen bonds. Future experiments will be aimed at exploring these proposals. By comparative analysis, the role of His70 in the E. coli enzyme is assumed by His878 in the multifunctional rat fatty acid synthase, which is a 272-kDa homodimeric enzyme [25,26]. A mutation in which alanine (Ala) replaced His878 completely abolished the dehydrase activity, although none of the other activities in the multifunctional mutant enzyme was significantly affected [25]. In contrast to the catalytic residue His70 in the E. coli dehydrase, His878 is not involved in any isomerization, and all 3-hydroxyacyl-S-ACP intermediates are converted to their corresponding 2-enoyl thioesters, which are subsequently hydrogenated by the enoyl reductase activity of the enzyme. Therefore, this rat synthase generates only saturated fatty acids.

Figure 2 Fatty acid biosynthetic pathway involving β-hydroxydecanoylthioester dehydrase. Both the dehydration and isomerization steps are depicted. ACP, acyl carrier protein.

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3. 3-Dehydroquinase Enzymes with no cofactor requirements often recruit amino acid residues other than general acids or bases to participate in the catalytic mechanism, and 3-dehydroquinase (DHQase, EC 4.2.1.10) is a good example. This enzyme catalyzes the dehydration of 3-dehydroquinic acid (4) to 3-dehydroshikimic acid (5) in the third step of the shikimate pathway, leading to the biosynthesis of aromatic metabolites in microorganisms, fungi, and plants [27,28]. There are two distinct classes of DHQase, designated type I and type II, and they exhibit different structural properties and mechanisms [29–31]. Chemical modification and differential peptide mapping studies revealed that the type I enzyme, exemplified by the E. coli enzyme, catalyzes a syn elimination of a water molecule [32] via an imine intermediate linked to a conserved lysine residue (Lys170) [33,34], and it uses a conserved histidine residue (His143) as a general base [35]. Subsequent experiments with site-directed metagenesis of the monofunctional, homodimeric E. coli enzyme [36] confirmed these conclusions, and the deduced mechanism is illustrated in Fig. 3. The covalent imine serves as an electron sink to stabilize the carbanion intermediate after the pro-R hydrogen is abstracted from the C-2 position. As a result of a proposed distortion of the carbocyclic ring of dehydroquinate upon formation of the Schiff base [32], the critical CEH bond is apparently reactive enough to be abstracted by His143. Interestingly, this histidine residue

Figure 3 Mechanism for 3-dehydroquinase (DHQase) from the third step of the shikimate pathway in Escherichia coli.

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may also play a key role in the formation and breakdown of the imine intermediate [36]. Unlike the type I enzymes, for the type II enzymes not much is known about the mechanism. However, it has been established that the catalysis does not involve an imine intermediate [29], and the elimination is antistereospecific [32,37]. Additionally, a recent study has identified a hyperactive arginine that may be involved in substrate binding [38]. The crystal structure of a type II DHQase has been determined [31], and when more information is available, a detailed comparison of the type I and type II enzyme mechanisms should yield useful insight.

B.

Reactions Catalyzed by Divalent Metal Ion–Dependent Enzymes

1. Carbonic Anhydrase Although some enzymes such as those discussed previously are able to activate the necessary CEH and/or CEO bonds, leading to dehydration without cofactor assistance, many enzymes must rely on various cofactors to accomplish their catalyses. The polarizing effect of a Lewis acid metal on the leaving group of a substrate is commonly utilized by biological systems to facilitate eliminations, and many CEO bond cleavage reactions catalyzed by divalent metal-containing enzymes are known. The reversible hydration of CO 2 catalyzed by carbonic anhydrase (CA, EC 4.2.1.1) is a good example. This zinc protein is expressed in almost all living organisms, including bacteria, plants, and animals, and it is a critical participant in a range of physiological processes [39,40]. Mammalian CAs are the best known, and there are at least seven different isozymes, designated CA I through VII [40]. The currently available structural and kinetic data support a similar active-site topology and a common mechanism (Fig. 4) for these CA isozymes. X-ray data indicate that the active site consists of a cleft with a zinc ion bound at the bottom [41]. In mammalian CA, the zinc ion is tetrahedrally coordinated to three histidine imidazoles and to a water molecule, which exists in the ionized, hydroxyl form with a pK a around 7 [42]. A threonine (Thr) and a Glu in the active site participate in a hydrogen bonding network with the Zn-OH ⫺ (6), and this structural feature is conserved in all nonplant CA isozymes. This network forcibly orients one of the hydroxyl lone electron pairs toward the CO 2 substrate, which is in a hydrophobic pocket [43,44]. The lone pair adds to CO 2 and forms a Zn-coordinated bicarbonate (7), which is rapidly replaced with H 2O (Fig. 4). Loss of one water proton to the solvent regenerates the Zn-OH ⫺, and this rate-limiting proton transfer is mediated by a His residue [42,45] and two water molecules [46], which together form an intramolecular proton shuttle. Whether the equilibrium favors hydration or dehydration depends significantly

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Figure 4 Schematic representation of the reversible hydration of CO 2 catalyzed by carbonic anhydrase. A histidine shuttle (not shown) is involved in the transfer of a proton between the solvent buffer and the catalytic ZnOH ⫺.

on the specific isozyme and tissue type [40], though both the forward and reverse reactions proceed through the highly reactive Zn-OH ⫺. In spinach CA, and perhaps in other plant CAs, extended x-ray absorption fine structure (EXAFS) studies have shown that the coordination sphere for the zinc ion includes at least one sulfur ligand [47]. Though the exact ligand structure is not yet known, plant CAs apparently have a quite different active-site organization than mammalian CAs. Regardless of the structural differences, the kinetics of spinach CA are consistent with the mammalian zinc-hydroxide mechanism, including a rate-determining proton transfer step that requires the participation of buffer for maximum efficiency [47]. Despite the lack of conclusive evidence for a proton-shuttle system in the spinach CA [47], it appears that a similar mechanism is shared by all known CAs. Further structural studies of the convergently evolved mammalian and plant enzymes will be instrumental in developing a deeper understanding of the catalysis of CA. Sly and Hu [40] summarize the information on this enzyme more comprehensively. 2. Enolase and Related Enzymes Enolase (EC 4.2.1.11), an enzyme in the glycolytic and gluconeogenesis pathways, catalyzes the reversible dehydration of 2-phospho-d-glycerate (8) (2-PGA, Fig. 5) [48]. Its catalysis requires two divalent cations, one of which binds at the high-affinity ‘‘conformational’’ site (site I); the other binds at the lower-affinity ‘‘catalytic’’ site (site II) [49]. Although the natural cofactor Mg 2⫹ provides the highest activity [49], Mn 2⫹, Zn 2⫹, or other divalent metal ions also fulfill the requirement [50,51]. At high concentrations, a metal cation apparently binds at a third site and inhibits the enzyme [52,53]. The overall catalytic cycle includes a minimum of eight steps [53], though the following discussion pertains only to the deoxygenation steps. Isotope exchange experiments [54–57] and analogue

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Figure 5 Reaction catalyzed by enolase. Though there is disagreement on the identity of the base, isotopic exchange experiments support the intermediacy of the aci-carboxylate (see text for details). 2-PGA, 2-phospho-d-glycerate.

studies [57–59] have supported a mechanism (Fig. 5) in which an active-site base abstracts the α-proton, then the enolate intermediate (depicted as the acicarboxylate 9 in Fig. 5) undergoes vinylogous antielimination [60] of the 3-hydroxyl to give the product 10. Although this general, stepwise mechanism is broadly accepted, some details such as the identities of the active-site base and the metal ligands remain disputed. One of the three contrasting proposals on the catalytic machinery of enolase is derived from the analysis of crystals of the yeast enzyme soaked with the natural substrate 2-PGA [61] and with the inhibitor phosphonoacetohydroxamate (PhAH) [62]. In their proposal, Lebioda and Stec contend that a water molecule serves as the catalytic base in conjunction with Glu168 and Glu211, and subsequent studies on the E211Q mutant supported this ‘‘charge shuttle’’ mechanism [63]. These studies also indicated that the 3-OH of the substrate is ligated to metal I. Janin and coworkers, judging from the x-ray structure obtained from lobster enolase-Mg 2⫹ crystals soaked with the inhibitor phosphoglycolate, proposed a similar role for metal I in the lobster enzyme [64]. However, the lobster enolase data suggested that His157, instead of water, is likely the catalytic base. Rayment, Reed, and their coworkers presented a third proposal, which is based on x-ray analysis of yeast enolase cocrystallized with Mg 2⫹ and PhAH [65]. In this proposal, Lys345 is the active-site base and metal I coordinates to the C-1 carboxylate. Recently, they cocrystallized yeast enolase with the natural substrate 2-PGA and confirmed their early conclusions regarding the metal coordination (octahedral) and active-site base assignment (Lys345) [66]. More importantly, both metal sites were resolved in this work, and they were found to be separated ˚ , in contrast to a distance of 6–9 A ˚ estimated by nuclear by a distance of ⬃4.2 A magnetic resonance (NMR) spectroscopy [67]. Furthermore, the 3-OH of 2-PGA appears to interact with Glu211 instead of with metal I, as in the other two proposals. Studies with K345A, E168Q, and E211Q mutants provided additional support for the role of Lys345 as the catalytic base and for the involvement of Glu211

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in 3-OH activation [56]. Notably, the relative positions of Lys345 and Glu211 in the Reed and Rayment proposal are consistent with the antistereochemistry of the elimination reaction. The x-ray crystal structure obtained by Larsen et al. [66] also revealed that both essential divalent cations interact with the carboxylate of the substrate/ product. Together with Lys396, they appear to promote the redistribution of the negative charge developed during the formation of the aci-carboxylate intermediate. Thus, a relatively weak active-site base, most likely assisted by significant substrate orientation by the enzyme [68], can abstract a nonacidic (pK a ⬃ 28–32) proton and expel a poor leaving group. Another example of this class of hydrolase is galactonate dehydratase, which depends on Mg 2⫹ [69] and appears to utilize His285 as the active-site base [70]. The Mn 2⫹-dependent imidazole glycerol phosphate dehydratase [71] may also be related to this mechanistic class of hydrolase. The properties of the latter enzymes remain to be further characterized. C.

Reactions Catalyzed by Enzymes Containing Iron–Sulfur Clusters

1. Aconitase Iron–sulfur clusters are common redox cofactors in biological systems. However, cases are known in which this metal center serves as a Lewis acid in enzymic reactions. For example, aconitase, otherwise known as citrate(isocitrate) hydrolase (EC 4.2.1.3), catalyzes the interconversion of citrate (11) and isocitrate (13) in the Krebs cycle through the dehydrated intermediate, cis-aconitate (12) (Fig. 6). This mitochondrial enzyme (83 kDa) contains a [4Fe-4S] center [72– 74], which is in the inactive [3Fe-4S]⫹ form when aconitase is purified aerobically [75,76]. Reduction in the presence of Fe2⫹ will convert the enzyme to the active [4Fe-4S] 2⫹ state (Fig. 7) [75]. Additional reduction to the electron paramagnetic resonance (EPR)-active [4Fe-4S] ⫹ state causes a ⬃60% drop in activity relative to the [4Fe-4S] 2⫹ state [77], presumably due to the decreased Lewis acidity of the reduced [4Fe-4S] ⫹ center. Mo¨ssbauer spectroscopic analyses on enzyme re-

Figure 6 General mechanism of the reaction catalyzed by aconitase.

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Figure 7 Interconversion between the inactive and active forms of the iron–sulfur cluster of aconitase. The labile iron atom is designated as Fe a . (Source: Adapted from Ref. 72.)

constituted with 57 Fe revealed a single labile iron site, which is called Fe a [75,78]. Protein ligands of the nonlabile iron atoms are cysteine residues [79–82], and the labile iron is ligated to a hydroxyl ion, which becomes protonated to water when substrate is bound [81–84]. Results gathered from x-ray analysis [82], mutation experiments [85], and inhibitor studies [86] support a general mechanism (amino acid residue numbering for porcine heart aconitase), that starts with substrate’s binding to the enzyme and causes the active-site cleft to close. Then Arg452 and Arg580 act in concert with His101 and His147 to bind and orient the α- and γ-carboxyl groups of citrate (Fig. 8). At the same time, both the β-carboxylate and β-hydroxyl of citrate coordinate to Fe a, thus expanding the coordination sphere to octahedral, and the ironbound hydroxyl group becomes protonated to form water. The proton to be abstracted is aligned with serine (Ser642), which is in an oxyanion hole (Arg644) and consequently deprotonated as a result of a significant lowering of pK a. Anionic Ser642 accepts the proton from the substrate and eliminates the hydroxide via a carbanion intermediate [86] to form cis-aconitate. The positive charge on

Figure 8 Binding of citrate to the [4Fe-4S] cluster of aconitase and partial active-site structure. (Source: Adapted from Ref. 74.)

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Fe a also assists in the dissociation of the hydroxyl group from the substrate. As shown in Fig. 8, cis-aconitate, with its γ-carboxyl bound to Arg580, then flips 180° as a result of a conformational change [87]. Such a flip switches the positions of the α- and β-carbons with respect to Fe a and allows addition of water onto the α-carbon. The overall reaction is readily reversible. 2. Fumarate Hydratase Fumarate hydratase (fumarase, EC 4.2.1.2) catalyzes the reversible hydration of fumarate (14) to (S)-malate (15) in the citric acid cycle (Fig. 9). The mammalian enzyme is extremely efficient and catalyzes the reaction with an apparent secondorder rate constant (k cat /K m ⫽ 2.4 ⫻ 10 8) approaching the diffusion limit [88,89]. Unlike fumarases of mammals, yeast, and Bacillis subtilis (class II), fumarase A from E. coli (class I) is an iron–sulfur-containing hydrolase [90]. The active fumarase A is a homodimer (60 kDa per monomer), and each subunit contains a [4Fe-4S] cluster that can be oxidized to the inactive [3Fe-4S] state in the same fashion as shown in Fig. 7 for aconitase [90]. The reduced [4Fe-4S] form of fumarase A is much less active (10- to 50-fold) than the oxidized [4Fe-4S] 2⫹ form, and substrate binding dramatically affects the [4Fe-4S] ⫹ EPR signal. These results are used as evidence to support a catalytic mechanism for fumarase that parallels the one for aconitase [90]. Other class I fumarases may include E. coli fumarase B [91], Euglena gracilis fumarase [92], and Rhodobacter capsulatus fumarase [93]. It should be noted that class II fumarases bear little homology to fumarase A and fumarase B from E. coli [94,95], and they have no cofactor requirements, so these other enzymes do not function by the same mechanism.

Figure 9 Postulated mechanism of the reaction catalyzed by fumarase. There is no evidence for a sixth ligand, but a water molecule is bound as a sixth ligand in the related enzyme aconitase [82]. Thus, a question mark designates a possible bound water molecule in fumarase. (Source: Adapted from Ref. 90.)

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3. Dihydroxy-Acid Dehydratase Dihydroxy-acid dehydratase (DHAD, EC 4.2.1.9) catalyzes the dehydration of 2,3-dihydroxycarboxylic acids 16 to the corresponding 2-keto acids 17, a key step in the branched-chain amino acid synthesis of valine and isoleucine (Fig. 10). The E. coli enzyme is a 125-kDa homodimeric protein containing a [4Fe4S] cluster per monomer [96]. This DHAD is sensitive to both O 2 and superoxide [96–98]. However, in marked contrast to that in aconitase, the in vitro activity of the inactivated enzyme could not be restored by incubation with Fe 2⫹ and reducing agent [96], apparently because the Fe-S cluster completely degrades upon oxidation. Conversely, in vivo studies demonstrated that the inactivation was reversible [99]. Resonance Raman spectroscopy established the presence of a single iron, which lacks cysteinyl coordination in the [4Fe-4S] cluster [96]. This iron is postulated to be directly involved in the reaction as with aconitase. As opposed to the E. coli enzyme, DHAD from spinach leaves (homodimer, 63 kDa per monomer) is not inactivated by O2, and it has a [2Fe-2S] center instead of a [4Fe-4S] center [100]. The EPR spectra of this unique Fe-S dehydratase (g ave ⫽ 1.90) are characteristic of Rieske Fe-S proteins, even though the redox potential of the [2Fe-2S] center (⫺470 mV) more closely resembles that of planttype ferredoxins. This dual nature of the spinach DHAD [2Fe-2S] center is reflected in its reaction with substrate. When substrate is added to the reduced protein, the EPR signal transiently shifts so the g ave approaches that of other planttype ferredoxins (gave ⫽ 1.95). After the substrate is consumed, the EPR spectrum reverts to the one similar to Rieske proteins. Reduction of spinach DHAD decreases the activity 6-fold. These observations indicate the direct involvement of the Fe-S center in the reaction, probably as a Lewis acid, to facilitate the departure of the β-hydroxyl group. Consistent with this suggestion, substrate analogue stud-

Figure 10 Schematic for the reaction catalyzed by dihydroxyacid dehydratase. Note that a single enzyme is involved in the synthesis of two different amino acids, depending on the identity of the R group.

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ies support the development of a partial positive charge at β-C [101]. The substrate must have the R-configuration at α-C, but DHAD surprisingly accommodates either the 2R,3R or 2R,3S isomer with nearly equal efficiency. Solvent hydrogen incorporation at the β-position proceeds with retention of configuration [102]. Because DHAD from all sources examined thus far also requires a divalent metal ion for activity [101], it would be interesting to find out whether the mechanism of DHAD bears any similarity to those discussed in Section II.B as well as to that of aconitase. 4.

L-Serine

Dehydratase

Recently, an l-serine dehydratase (EC 4.2.1.13) involved in both the biosynthesis and metabolism of serine was purified from Peptostreptococcus asaccharolyticus and shown to be a heterodimer of 55 kDa (α, 30 kDa; β, 25 kDa) containing a [3Fe-4S] center [103,104]. The enzyme could be inactivated by exposure to air and reactivated by incubation with Fe 2⫹ under anaerobic conditions, presumably by regenerating an oxidized [4Fe-4S] 2⫹ center. Though EPR spectroscopic experiments confirmed the presence of an oxidized [3Fe-4S] ⫹ center, direct evidence for a [4Fe-4S] center is still lacking [104]. Instead, the purported [4Fe-4S] 2⫹ center was inferred by titration of the anaerobically purified enzyme with K 3Fe(CN) 6 and observation of a successively larger [3Fe-4S] ⫹ EPR signal, which correlated with a decrease in activity. Interaction of the substrate with the Fe-S center was also inferred from the observation that l-serine could both shield the [3Fe-4S] ⫹ center from buffer-dependent changes in the EPR signal and prevent oxidative loss of activity when the active enzyme is exposed to air. This evidence has been used to propose a mechanism in which the labile iron of the [4Fe-4S] 2⫹ center binds the hydroxyl and carboxyl groups of serine [104]. This iron atom would thus serve as a Lewis acid and facilitate loss of the hydroxyl group, an otherwise poor leaving group (Fig. 11). Such a mechanism is vastly different from that for most l-serine dehydratases from nonbacterial sources, which utilize PLP as the cofactor in the irreversible deamination of serine [105]. In the latter case, the α-H and not the β-hydroxyl group is activated during catalysis. D.

Reactions Catalyzed by Enzymes Containing Organic Cofactors

1. Pyridoxal 5′-Phosphate–Dependent Dehydratases: L-Serine Dehydratase and L-Threonine Dehydratase Many enzymes that catalyze β-elimination reactions are pyridoxal 5′-phosphate– (PLP)-dependent [106,107]. Two such PLP-dependent enzymes that catalyze the β-elimination of a hydroxyl group are l-serine dehydratase (EC 4.2.1.13) and l-threonine dehydratase (EC 4.2.1.16), which irreversibly convert l-serine and

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Figure 11 Postulated mechanism for the reaction catalyzed by the iron–sulfur-containing l-serine dehydratase from Peptostreptococcus asaccharolyticus. (Source: Adapted from Ref. 104.)

l-threonine to pyruvate and α-ketobutyrate, respectively. Although all the lthreonine dehydratases from different sources have thus far proved to be PLPdependent, l-serine dehydratases from Peptostreptococcus asaccharolyticus [103] and Clostridium propionicum [108] depend instead on an iron–sulfur center (discussed in Section II.C.4). The role of the PLP coenzyme in these hydrolases is to facilitate the C-H bond cleavage at α-C of the substrate by delocalizing the resulting anion throughout the pyridine ring. These electrons are then used to drive the subsequent elimination of the β-hydroxyl group. As depicted in Fig. 12, the reaction is initiated by the formation of Schiff base 18 between PLP and the amino group of the substrate. Abstraction of the α-proton generates resonance-stabilized intermediate 19. Electron pushing in this transient quinonoid species expels the β-hydroxyl group and leads to the formation of aminoacrylate intermediate 20. The ⑀-amino group of the active-site lysine then adds to the C-4′ position of the PLP moiety (21), followed by the release of the corresponding imine. Hydrolysis of the resulting imine yields the 2-keto acid product 22 and completes the catalytic cycle. The large body of evidence supporting this mechanism has been summarized by Miles [106]. Tryptophan synthase (EC 4.2.1.20) utilizes a similar mechanism [106,109], though the reaction is a β-replacement of the hydroxyl group with an indole moiety instead of a hydrogen. 2. Nicotinamide-Adenine-Dinucleotide-Dependent Dehydratases: TDP-Glucose 4,6-Dehydratase Temporarily converting a hydroxyl group to a keto function via intramolecular oxidation–reduction is another strategy adopted by biological systems for activating an adjacent CEH bond in CEO bond cleavage reactions. For example, this

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CEO Bond

Figure 12 Scheme for the dehydration of l-serine (R ⫽ H) and l-threonine (R ⫽ CH 3) catalyzed by PLP-dependent l-serine and l-threonine dehydratase. Note that these enzymes from most sources will act on both l-serine and l-threonine with varying degrees of efficiency. PLP, pyridoxal 5′-phosphate. (Source: Adapted from Ref. 106.)

interesting deoxygenation method is utilized by nucleotidyl diphosphohexose 4,6-dehydratases, which transform nucleotidyl diphosphohexoses into the corresponding 4-keto-6-deoxy sugar derivatives that are the precursors of many unusual sugars [110,111]. The best-studied enzyme within this family is TDP-dglucose 4,6-dehydratase (80-kDa homodimer, EC 4.2.1.46) isolated from E. coli, and this enzyme contains 1 NAD ⫹ per dimer [112]. Notably, recent studies of CDP-d-glucose 4,6-dehydratase, a closely related enzyme from Yersinia pseudotuberculosis, have shown that it has a higher affinity for NADH than for NAD ⫹ [113]. Since two cofactor binding sites were found in all these enzymes, yet only one NAD ⫹ per dimer was generally detected, it is possible that the other coenzyme binding site is occupied by NADH.

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As illustrated in Fig. 13, the reaction consists of three catalytically discrete steps: step I, oxidation of nucleotidyl 5′-diphosphohexose 23 to the corresponding 4-ketohexose 24; step II, C5/C6 dehydration to a 4-keto-∆5,6-glucoseen intermediate 25; and step III, reduction at C-6 to give the resulting 4-keto-6-deoxyhexose product 26. This net intramolecular oxidation–reduction has been shown to involve an internal hydrogen transfer from C-4 of the substrate 23 to C-6 of the product 26, and the enzyme-bound NAD ⫹ apparently functions as a hydride carrier in this catalysis [114,115]. Thus, there are actually two C-H cleavage events involved in this catalysis. The first CEH bond rupture forms the keto group, which activates the hydrogen at C-5 for the second C-H bond cleavage, triggering the elimination of the 6-OH. The displacement of the hydroxyl group at C-6 by the hydrogen from C-4 occurs with net inversion, the hydrogen transfer to NAD ⫹ is ‘‘si-face’’-specific, and the dehydration from C-5 and C-6 is a syn elimination [116–119]. These results indicate that sugar 4,6-dehydratases belong to a small group of enzymes in which the pyridine nucleotide coenzyme is a catalytic prosthetic group, contrary to most other nicotinamide-dependent enzymes, in which NAD ⫹ or NADP ⫹ functions as a cosubstrate [110]. In addition to the sugar 4,6-

Figure 13 Catalytic cycle for the dehydration of nucleotidyl diphosphohexoses. The E od label is meant as a general reference to all enzymes that catalyze this type of reaction. The NDP at C-1 reflects the alternate nucleotidyl sugar substrates that are utilized by the different dehydratases, NDP, nucleotidyl 5′-diphospho-. (Source: Adapted from Ref. 113.)

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dehydratases, this unique group of catalysts includes a few other enzymes, such as 3-dehydroquinate synthase (EC 4.6.1.3) [120,121] and 2-deoxy-scyllo-inosose synthase [122]. 3. Flavin Dependent Dehydratases a. (R)-2-Hydroxyglutaryl-Coenzyme A Dehydratase. Another interesting class of enzymes that catalyze the elimination of water using redox chemical processes is the hydroxy acyl-CoA dehydratases from Clostridium and related bacteria. These oxygen-sensitive enzymes are able to dehydrate a wide range of hydroxyacyl-CoA derivatives in the process of anaerobic fermentation. One such enzyme, isolated from Acidaminococcus fermentans, catalyzes the reversible syn dehydration of (R)-2-hydroxyglutaryl-CoA (27) to (E )-glutaconyl-CoA (28) in the glutamate fermentation pathway (Fig. 14) [123]. This reaction is of considerable interest since it involves the cleavage of a nonactivated C-H bond at C-3. The active heterodimer (α, 54 kDa; β, 42 kDa) is oxygen-sensitive, and it contains 4 mol nonheme iron, 4 mol inorganic sulfur, 0.3 mol reduced riboflavin, and 1 mol reduced flavin mononucleotide (FMN) [124]. An additional adenosine triphosphatic– (ATP)- and Mg 2⫹-dependent Fe-S protein (homodimer, 54 kDa per monomer) is required in catalytic amounts to activate the dehydratase reductively. The in vitro assays used Ti(III) citrate as the reductant, though NADH is suspected to be the electron source in vivo, possibly via an electron-relay pathway involving a diaphoraselike enzyme that can transfer single electrons [123]. Titanium (III) citrate delivers a low-potential electron (E°′ ⫽ ⫺600 mV) that is apparently recycled for many turnovers, suggesting a radical mechanism. The observation that oxidants, including 2-nitrophenol, 3-nitrophenol, 4-nitrophenol, 4-nitrobenzoate, and chloramphenicol, temporarily inhibit the dehydration reaction until these compounds are consumed also supports the idea of a radical mechanism [124]. As shown in Fig. 14a, the mechanism may include a ketyl intermediate 29 that eliminates the hydroxyl group with the assistance of a [4Fe-4S] center. It has been demonstrated that α-hydroxyketones undergo single-electron chemical reactions in the presence of a strong one-electron donor to eliminate hydroxyl groups [125]. The resulting enoxy radical 30 could be reduced to give 31, which then releases the 3-H as a hydride to regenerate the reduced flavin coenzyme and concomitantly afford the product 28. However, it is more likely that enoxy radical intermediate 30 releases the 3-H as a proton to give 32, followed by an electron loss to form glutaconyl-CoA 28 (Fig. 14b). The proposed recycling of a single, low-potential electron is mechanistically appealing and may be involved in the catalysis of other related enzymes. Two possible examples are (R)-2-hydroxyglutaryl-CoA dehydratase from Fusobacterium nucleatum [126] and (R)-lactyl-CoA dehydratase (EC 4.2.1.54) from Clostridium propionicum [127], both of which are iron–sulfur flavoenzymes in-

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volved in the anaerobic fermentation of glutamate and alanine, respectively. Studies with EPR spectroscopy have characterized a [4Fe-4S] and a [3Fe-3/4S] center in the latter enzyme [128], and the dehydration using an alternate substrate, (R)2-hydroxybutyryl-CoA, was established to be a syn elimination [129]. Further studies may reveal that these enzymes operate through a common mechanism, in which a C-H bond is broken after the C-O bond cleavage has occurred, as opposed to the mechanism of 4-hydroxybutyryl-CoA dehydratase discussed later and most other enzymes reviewed here. b. 4-Hydroxybutyryl-Coenzyme A Dehydratase. This enzyme is 4-hydroxybutyryl-CoA dehydratase from C. aminobutyricum (homotetramer, 56-kDa per subunit), which converts 4-hydroxybutyryl-CoA (33) to crotonyl-CoA [34] in the metabolism of γ-aminobutyrate (Fig. 15) [130,131]. Stoichiometric quantities of iron and sulfur as well as flavin adenine dinucleotide (FAD) have been found in the enzyme [132], and EPR spectroscopic studies indicated the presence of an oxygen-sensitive [4Fe-4S] cluster that is required for activity [133]. Interestingly, this [4Fe-4S] cluster decomposes to a species other than a [3Fe-4S] center upon air inactivation, in contrast to the cluster in aconitase (Section II.C.1). Thus, the [4Fe4S] cluster may serve in a structural capacity instead of, or in addition to, acting as a Lewis acid. Other studies have revealed that the reduced enzyme is inactive [132], so the flavin cofactor must be oxidized before catalysis can be initiated. These data support a mechanism consisting of three steps: dehydrogenation, dehydration, and isomerization. As shown in Fig. 15a, the reaction could be initiated by an α-proton abstraction followed by a β-hydride transfer to reduce the active-site bound FAD and give 4-hydroxycrotonyl-CoA (35). The reducing equivalents would be subsequently used in a nucleophilic attack at the α-carbon, and the 4-hydroxyl could be eliminated possibly with assistance from the [4Fe4S] center, as is the case with aconitase. The resulting vinylacetyl-CoA (36) is then isomerized to crotonyl-CoA (34). Alternatively, a mechanism that includes an oxidized ketyl intermediate could be involved [134]. In this proposed radical mechanism, depicted in Fig. 15b, the intermediate enolate 37 could be oxidized by the FAD to enoxy radical 38, which would deprotonate to a ketyl radical anion 39, expel the 4-OH (with or without assistance from a [4Fe-4S] cluster), and go through dienoxy radical

Figure 14 Proposed mechanisms for the reversible dehydration of 2-hydroxyglutaryl-CoA involving loss of either (a) a hydride or (b) a proton from the substrate. The participation of the activase is illustrated in (b), though it would play a similar role in (a) as well. The possible coordination of the substrate to a [4Fe-4S] center is represented by ‘‘Fe’’ in (a), and it is shown with more detail in (b). CoA, coenzyme A; FMN, flavin mononucleotide; ATP, adenosine triphosphate; ADP, adenosine diphosphate. (Source: Adapted from Ref. 124.)

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Figure 15 Scheme for the possible (a) anionic and (b) radical mechanisms for the reversible dehydration of 4-hydroxybutyryl-CoA. To reduce congestion, the [4Fe-4S] cluster is represented as ‘‘Fe’’ in (b). FAD, flavin adenine dinucleotide.

40 before regenerating the FAD cofactor and completing the catalysis. Support for the latter hypothesis derives from two primary sources: (1) There is a close relationship with the (R)-2-hydroxyglutaryl-CoA dehydratase, as discussed in the previous section, and (2) the FAD of 4-hydroxyglutaryl-CoA dehydratase is readily reduced to the neutral semiquinone whereas reduction to the hydroquinone appears to be kinetically hindered [133]. Though the details of this mecha-

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nism still need to be elaborated and verified, it seems clear that the flavin coenzyme directly participates in the deoxygenation reaction, and it needs to be oxidized to do so. c. Chorismate Synthase. Another deoxygenation in which the cleavage of a C-O bond may occur prior to the rupture of a C-H bond is catalyzed by chorismate synthase (EC 4.6.1.4) in the seventh step of the shikimate pathway (also see Section II.A.3). The enzyme from all known sources depends on a reduced flavin coenzyme in the conversion of 5-enolpyruvylshikimate 3-phosphate (EPSP) (41) to chorismate (42) [135–139], as shown in Fig. 16. Although this reduced flavin is often generated by an alternative pathway, the enzyme from some sources is bifunctional and can provide the reduced flavin via an additional diaphoraselike activity using NADPH as the cosubstrate [28]. After the catalytic cycle, the flavin cofactor remains reduced, so there is no net change in the oxidation state of the enzyme. Though the precise role of the flavin is unknown, studies using flavin and

Figure 16 Possible nonconcerted pathways for the chorismate synthase reaction. Current data cannot distinguish between a cationic (top) and a radical (bottom) mechanism. (Source: Adapted from Ref. 158.)

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substrate analogues [140,141] and ultraviolet–visible (UV-Vis) spectrophotometry [142,143] indicated that this prosthetic group may be chemically, not just structurally, involved in turnover. The net 1,4-elimination of the 3-phosphate and the 6-H R proceeds with antistereochemistry [144–146]. Model system studies [147,148] and molecular orbital calculations [149,150] suggested a preference for syn stereochemistry in concerted 1,4-eliminations. Thus, it has been proposed that the chorismate synthase reaction is nonconcerted [151,152], and kinetic isotope effect studies are consistent with this proposal [153]. However, evidence supporting a nonconcerted mechanism is not conclusive [152,154–158], so other possibilities must still be considered. Interestingly, the observation of a neutral flavin semiquinone upon incubation with the competitive inhibitor (6R)-6-fluoroEPSP suggested the likely involvement of radical intermediates [159]. Although single-turnover kinetics did not detect the neutral semiquinone radical, a different flavin intermediate was found to form and decay in a kinetically competent fashion [156–158]. The spectral features of this flavin intermediate may be attributable in part to an anionic semiquinone radical [158]. Thus, these biophysical studies support a nonconcerted mechanism that includes an allylic radical intermediate, though the current data are unable to distinguish this mechanism from other nonconcerted models (Fig. 16). Clearly, a more detailed investigation is necessary to characterize further the nature of the flavin intermediate, determine its role in the catalysis, and discriminate between an ionic and a radical mechanism.

E.

Protein-Radical-Dependent Dehydratases

1. Diol Dehydrases Though the aforementioned flavin-dependent enzymes may include a free-radical intermediate in the reaction pathway, none of those reactions requires the formation of a protein radical as part of the catalysis. However, there are a few systems in which the generation of a protein radical appears to be a prerequisite to abstract a hydrogen atom from the substrate before deoxygenation can occur. The reactions mediated by diol dehydrases and glycerol dehydrases are two likely examples [160]. These multimeric enzymes (⬃230 kDa) catalyze the irreversible dehydration of vicinal diols into corresponding aldehydes or ketones [161] in the anaerobic fermentation of glycerol and other glycols in several bacterial organisms [Table I of Ref. 162]. Adenosylcobalamin (AdoCbl) is a required cofactor of these enzymes [163], and the adenosyl radical [164] that is generated by the homolytic cleavage of the reactive Co-C bond of AdoCbl [165] is the initiator of the reaction. A monovalent cation, such as K ⫹, NH 4⫹, Tl ⫹, or Rb ⫹, is also required to assist in the binding of AdoCbl to the enzyme [166,167]. In the cases with methylcobalamin-dependent methionine synthase [168] and methylmalonyl-

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CoA mutase [169], the axial dimethylbenzimidazole (DMB) ligand of AdoCbl is displaced by a histidine residue of the enzyme, and this ligand substitution initiates the formation of a charge-relay system that seems to be important for modulating the Co-C bond strength [170]. However, analysis of the deduced amino acid sequence of diol dehydrase fails to locate the fingerprint sequences characteristic of histidine ligation to cobalt [170], so the mechanism whereby this protein affects the Co-C bond strength is not yet clear. The AdoCbl-catalyzed dehydration of vicinal diols is mechanistically quite similar to most other AdoCbl-dependent rearrangements [161,171]. The overall mechanism is depicted in Fig. 17, and the reaction is initiated by the substrateinduced homolytic cleavage of the Co-C bond of AdoCbl into a 5′-deoxyadenosyl radical and cob(II)alamin. The adenosyl radical abstracts a hydrogen atom from a protein residue (shown in Fig. 17 as a cysteine), and the resulting protein radical abstracts a C-1 hydrogen atom (H a) from the substrate 43 to generate 44. Substrate radical 44 rearranges to the product radical 45, which is converted to gem-diol 46 by receiving H a from the protein residue. Subsequent dehydration leads to the formation of product 47. Release of the product shifts the equilibrium so the AdoCbl cofactor returns to the associated form, thus completing the catalytic cycle. Signals arising from low-spin Co(II) (g ⫽ 2.2–2.4) and an organic radical (g ⫽ 1.95 and 2.04) have been detected by EPR [172–174], and both species were later shown to be formed at a kinetically competent rate [175]. The fact that hydrogen isotope exchange occurs readily between C-5′ of the adenosyl moiety and C-1 of the substrate corroborates the intermediacy of the adenosyl radical [176–180]. Both the hydrogen migration and the hydroxyl elimination are stereospecific [181]. The stereochemistry at C-2 dictates which hydrogen atom at C-1 undergoes migration, with the 1-H S being abstracted from (S)-1,2-propanediol and the 1-H R from the (R)-substrate [182]. The subsequent hydrogen atom rebound to replace the migrating hydroxyl group proceeds with inversion of the C-2 configuration [181,182]. During the migration of the hydrogen atom, there is no exchange with the solvent [183]. The involvement of a protein radical in diol dehydratase is implicated mainly by an observed isotope effect of 125 for the transfer of 3 H from cofactor to product [163]. In order to explain the large k H /k T isotope effect, an additional step to generate a protein radical intermediate appeared to be necessary [184]. In fact, the intermediacy of a protein radical has been suggested for many vitamin B 12 –dependent reactions. The observations of a critical thiol residue in diol dehydrase [185], of a possible protein-based radical in AdoCbl-dependent ethanolamine ammonia lyase [186,187], and of a kinetically competent thiyl radical intermediate in the AdoCbl-dependent ribonucleotide reductase [188] all support the assignment of this putative protein radical as a thiyl radical, and this proposal is reflected in Fig. 17. Although the rearrangement mechanisms have not been

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Figure 17 Proposed mechanism of the diol dehydrase reaction. Note that the AdoCbl is predominantly in the associated form until the substrate binds. Though 1,2-propanediol is illustrated as the substrate in this scheme, several other glycols are recognized by this enzyme (Table I of Ref. 162). The stereochemistry as depicted may be altered by the configuration at the C-2 position (see text for details). Ado, adenosyl moiety; [Co], cobalamin.

fully elucidated and the protein-based radical has not been directly detected in diol dehydrase, the participation of the adenosyl radical in the reaction is conclusive. However, a reactive protein radical may actually be responsible for abstracting a relatively inert C-H bond of the substrate to initiate the deoxygenation.

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2. Ribonucleotide Reductase Of the many deoxygenation reactions that have been characterized, the conversion of ribonucleotides to 2′-deoxyribonucleotides is perhaps the most prominent because of its role in deoxyribonucleic acid (DNA) biosynthesis. Because of the sheer importance of the reaction it catalyzes, ribonucleotide reductase (RNR) has been studied extensively over the years, and the mechanism is now well characterized. Ribonucleotide reductase utilizes a stable protein radical to catalyze the irreversible, reductive replacement of the 2′-hydroxyl group with hydrogen in ribonucleotides. Interestingly, at least three classes (I–III) of RNR have been discovered, and each employs different cofactors and protein radical–generating mechanisms, though the subsequent catalytic steps are the same. In all known forms of RNR, radical-mediated abstraction of a nonacidic hydrogen atom precedes C-O bond cleavage, similar to most anionic deoxygenation mechanisms. Several reviews summarizing the work that has been done on this enzyme are available [189–195], and information pertaining to cofactor requirements and the mechanism is briefly presented. Class I ribonucleotide reductases are characterized by a binuclear high-spin Fe(III) complex and a tyrosyl radical, and these enzymes aerobically deoxygenate ribonucleotide diphosphates. The 258-kDa (α 2β 2: α 2 usually known as R1 and β 2 as R2) protein from Escherichia coli is the best studied member of this class [196–202]. Crystal data of R2 confirmed the presence of a binuclear µ-oxo bridged iron cluster that is close enough to stabilize the tyrosine–122 (Tyr122) radical (Fig. 18) [203]. Scavenging of this Tyr122 radical by hydroxyurea [204] or by the substrate analogue 2′-deoxy-2′-mercaptouridine 5′-diphosphate [205] leads to irreversible inactivation of the enzyme. Extensive study of the spontaneous assembly of the iron cluster and the Tyr122 radical in the presence of Fe(II) and O 2 revealed the involvement of a diferric Fe(III)/Fe(IV) species and a neutral tryptophan–48 (Trp48) radical [206–211]. The Trp48 occupies a site in a hydrophobic region of R2 that is believed to interact with R1 [212]. It may also be part of an uncharacterized electron relay, involving His118, Asp237, and Trp48, that transfers the radical from Tyr122 in R2 to a critical protein residue ˚ ) where the substrate binds and reacts. This protein in R1 (a distance of 35 A residue is proposed to be Cys439 on the basis of mutagenesis studies [213] and of sequence comparisons with AdoCbl-dependent RNR from Lactobacillus leichmannii, in which a kinetically competent thiyl radical (Cys408) was characterized by EPR spectroscopy [188]. As shown in Fig. 19, deoxygenation is initiated by the abstraction of a hydrogen atom from the 3′-position of the ribose ring by the thiyl radical. The 2′-hydroxyl group is then protonated by one cysteine of a redox-active thiol pair (Cys225 and Cys462) and is lost as water, resulting in a resonance-stabilized cation radical intermediate. Though such a substrate-based radical has not yet

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Figure 18 Representation of the µ-oxo diferric iron cluster and the neighboring tyrosyl radical in class I RNR (Escherichia coli). The bridging oxide anion derives from molecular oxygen. The tyrosyl radical is stabilized by the iron cluster. (Source: Adapted from Ref. 195.)

been detected, a possible nucleotide-based radical was recently observed during studies with RNR and the time-dependent inactivators (E )- and (Z )-2′-fluoromethylene-2′-deoxycytidine 5′-diphosphate [214]. The redox-active thiol pair reduces the cation and protonates C-2′, becoming a disulfide in the process. Participation of this thiol pair in the reaction has been demonstrated by mutagenesis studies [215–217]. The more recent observations of a [XN ⋅ S CysR1] radical upon inactivation with 2′-azido-2′-deoxyuridine 5′-diphosphate [218] and of a perthiyl radical upon inactivation with 2′-deoxy-2′-mercaptouridine 5′-diphosphate [205] provide initial evidence for the intriguing possibility that reduction of the substrate intermediate by the redox-active thiol pair may occur via one-electron chemistry involving thiyl radicals, as opposed to a two-electron hydride transfer. After the cation radical intermediate is reduced, the same hydrogen atom abstracted by the thiyl radical is returned to the 3′-position to form the product deoxyribonucleotide. A biological thiol reducing agent, such as dithiothreitol, thioredoxin, or glutaredoxin, reduces the redox-active disulfide and completes the catalytic cycle. It is important to note that once the Cys439 thiyl radical is generated, the remaining catalytic steps are identical for all known classes of ribonucleotide reductases. Class II RNR depends instead on AdoCbl for its ability to reduce ribonucleotide triphosphates [219]. The best characterized member of this class is from L. leichmannii, and it is active under either aerobic or anaerobic conditions. This enzyme (82-kDa monomer) promotes the homolytic cleavage of the inherently

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Figure 19 Proposed mechanism of the reduction of ribonucleotides catalyzed by ribonucleotide reductase. The method for generating the catalytic thiyl radical depends on the class of RNR (see text). Amino acid numbering for the Escherichia coli class I enzyme is shown, though enzymes from class II and III utilize the same mechanism. RNR, ribonucleotide reductase. (Source: Adapted from Refs. 189 and 195.)

reactive CoEC bond of AdoCbl to form cob(II)alamin and the 5′-deoxyadenosyl radical. As for diol dehydrase (Section II.E.1), it is not understood how the RNR induces the homolytic cleavage of the CoEC bond [170], though a nitrogen has been identified as a ligand to cobalt [220]. The 5′-deoxyadenosyl radical then abstracts a hydrogen atom from Cys408 [221] to form a thiyl radical, which has recently been detected and characterized [188,222]. Subsequent ribose reduction steps are identical to those described previously (Fig. 19). Unlike the preceding two classes of RNR, a ribonucleotide triphosphate reductase (heterodimer, 160-kDa α2, 35-kDa β2) produced by E. coli grown under anaerobic conditions has been shown to have an absolute requirement of S-adenosylmethionine (AdoMet) for activity [223]. Similar to AdoMet-dependent pyruvate-formate lyase [224], this class III RNR uses an iron-dependent ‘‘activase,’’ AdoMet, NADPH, flavodoxin, and flavodoxin reductase to generate a proteinbased radical under anaerobic conditions [225], though in RNR the activase is actually part of the complete reductase complex [226]. It appears that the two

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subunits of the small protein are held together by a [4Fe-4S] center with one oxygen-sensitive, labile iron [226,227], and this small dimer forms a tight complex with the large dimer much as with class I RNR. On the basis of mechanistic work with pyruvate-formate lyase [228–230], the small dimer probably interacts with AdoMet and induces the homolytic scission to form both methionine and a 5′-deoxyadenosyl radical, which subsequently generates the stable protein radical in the large dimer. Experiments have localized this protein radical to Gly681 [231,232], which presumably takes the place of the tyrosine radical in class I RNR and generates the catalytic thiyl radical [233]. Thereafter, reduction of the ribose moiety and deoxygenation proceed as described (Fig. 19), except class III RNR utilizes electrons from formate instead of thioredoxin to regenerate the redox-active thiol pair [234].

III. A NOVEL CEO BOND CLEAVAGE EVENT A.

Background

Sugars are most commonly associated with processes involved in the storage and generation of the energy necessary to sustain life, such as glycogen synthesis and glycolysis. The most notable exception to the use of sugars as metabolic vehicles is the ribose moiety of DNA and ribonucleic acid (RNA), in which the sugar plays a definitive structural role by providing the scaffolding for all biological genetic material and the messages needed to translate it. The only difference between DNA and RNA molecules is the substitution of a hydrogen for the 2′hydroxyl group of the ribose moiety, and this structural difference has been directly linked to the greater stability of DNA over RNA. In fact, such substitutions generally enhance the stability of sugars, as reflected by the large number of structural roles that have been assigned to deoxysugars [235]. Although in most cases the precise functions of deoxysugars have not yet been fully elucidated, it appears that incorporation of a deoxysugar as a structural component provides a facile means of modifying the surface properties of a compound or organism. One relevant example is the incorporation of 3,6-dideoxysugars into the lipopolysaccharide (LPS) of some gram-negative bacteria [236–238], where these deoxysugars confer immunogenic properties to the bacteria [236,239]. Among the five known 3,6-dideoxyhexoses—ascarylose, abequose, paratose, tyvelose, and colitose—the biosynthesis of ascarylose by Yersinia pseudotuberculosis is the most thoroughly studied [111]. The proposed biosynthetic sequence of 3,6-dideoxysugars is exemplified by the formation of cytidine 5′-diphospho– (CDP)-ascarylose (48), as shown in Fig. 20. Initiation of the pathway proceeds with the coupling of α-d-glucose1-phosphate (49) and cytidine triphosphate (CTP) by α-d-glucose-1-phosphate

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Figure 20 Overview of the biosynthetic pathway for CDP-ascarylose. CDP, cytidine 5′-diphosphate.

cytidylyltransferase (E p) to give CDP-d-glucose (50). Subsequent steps start with an irreversible intramolecular oxidation–reduction catalyzed by the NAD ⫹-dependent CDP-d-glucose 4,6-dehydratase (Eod ), which uses a well-known mechanism, as described in Section II.D.2. The resulting product, CDP-6-deoxyl-threo-d-glycero-4-hexulose (51), is converted to 3,6-dideoxy-d-glycero-dglycero-4-hexulose (54) in two consecutive steps mediated by the actions of CDP-6-deoxy-l-threo-d-glycero-4-hexulose-3-dehydrase (E1) and E1 reductase (E3) [240,241]. The final steps are an epimerization catalyzed by CDP-3,6-dideoxy-d-glycero-d-glycero-4-hexulose-5-epimerase (Eep), which inverts the configuration at C-5, followed by stereospecific reduction at C-4 catalyzed by CDP-3,6-dideoxy-d-glycero-l-glycero-4-hexulose-4-reductase (Ered ) to give CDP-ascarylose (48). There is little doubt that the most intriguing step of this pathway is the C-3 deoxygenation catalyzed by E1 and E 3. This CEO bond cleavage event is particularly interesting since this is the first example of a pyridox-

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amine 5′-phosphate– (PMP)-mediated dehydration, and the reaction involves the formation of a radical intermediate, which will be discussed further along with other mechanistic aspects in the following sections. 1. CDP-6-Deoxy-L-Threo-D-Glycero-4-hexulose-3Dehydrase (E1) Enzyme E 1 is a red–brown protein consisting of two identical subunits (49 kDa per monomer). Both radiometric [242] and fluorometric [243] quantification revealed one equivalent of PMP per monomer for full activity. Analytical assays detected stoichiometric amounts of iron and sulfur, and EPR analysis of the reduced enzyme confirmed the presence of an adrenodoxin/putidaredoxin-type [2Fe-2S] center (g ⫽ 2.007, 1.950, and 1.930) [244]. It should be noted that purified E 1 is a mixture of apo- and holo-enzyme, so exogenous PMP, iron, and sulfur are required to reconstitute its activity fully. Incubation of the enzyme with substrate revealed that the PMP of E 1 forms a Schiff base with the C-4 keto group of the substrate [245]. Then, His220 abstracts the pro-S hydrogen from the C-4′position of the PMP in the Schiff base complex and triggers the expulsion of the C-3 hydroxyl group of the substrate to give PMP-∆ 3,4-glucoseen complex 52 (Fig. 20) [243]. Although intermediate 52 has never been isolated, it is reversibly hydrated on the si face of the conjugated imine, as demonstrated by isotopic incorporation with [18 O]H 2O [245]. Also, replacement of the C-3 hydroxyl group by a solvent hydrogen in the E 1 –E 3 coupled reaction proceeds with retention of configuration [246]. Together, these observations imply that the overall catalysis in the active site of E 1 is most likely a suprafacial process occurring on the si face of the PMP–substrate complex, as is the case with most other coenzyme B 6 –dependent enzymes [247]. In fact, sequence alignments demonstrated a clear relationship between E 1 and other PLP/PMP enzymes [243,248]. One significant difference highlighted by the comparisons is the replacement of a highly conserved lysine by a histidine at position 220 in E 1. This single replacement may be responsible for converting a PLP-dependent transaminase to a PMP-dependent dehydrase. The preceding results clearly showed that E 1 behaves as a normal coenzyme B 6-dependent enzyme that catalyzes a dehydration reaction. However, the C-3 deoxygenation product is formed only after NADH and E 3 are added to the reaction mixture. Other reductases, such as diaphorase and methane monooxygenase (MMO) reductase, are also able to generate small amounts of product [245]. Removal of the [2Fe-2S] center of E 1 impaired its ability to promote the final product formation, though E 1(apoFeS) could still abstract the C-4′ hydrogen of PMP [244]. This requirement of the [2Fe-2S] center, which is an obligatory oneelectron carrier, for product formation strongly implicated the intermediacy of free radicals in the reaction. An EPR analysis of the chemically reduced E 1 sub-

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strate complex confirmed the presence of a Lorentzian-type absorption (g ⬇ 2), which displayed no detectable hyperfine splitting [249]. The participation of the PMP cofactor in a deoxygenation is unique, but the involvement of a radical mechanism truly places E1 in a class by itself. 2. CDP-6-Deoxy-L-Threo-D-Glycero-4-hexulose-3-Dehydrase Reductase (E 3) CDP-6-Deoxy-l-threo-d-glycero-4-hexulose-3-dehydrase reductase (E 3) is a monomeric (36-kDa) protein of a red–brown color, and it contains 1 mol of FAD as revealed by high-performance liquid chromatography (HPLC) and UV-Vis analysis [250]. Iron and sulfur quantification combined with UV-Vis and EPR spectroscopy indicated the presence of a plant-type [2Fe-2S] center. Sequence alignments established a close relationship between E 3 and other iron–sulfur flavoproteins in the ferredoxin-NADP ⫹ reductase (FNR) family [251]. Like other members in its family, E 3 will transfer reducing equivalents from NADH to a variety of one-electron acceptors, including O 2, with varying degrees of efficiency [252]. Removal of the [2Fe-2S] center impairs the ability of E 3 to catalyze final product formation in the presence of E 1 and substrate, but the FAD of E 3(apoFeS) remains functional as a two-electron/one-electron switch in the reduction of O 2 to H 2O 2 [250]. Therefore, the FAD can operate independently of the iron–sulfur center when E 3 acts as a NADH oxidase, but the electron-transfer relay from NADH to substrate during sugar reduction clearly includes both FAD and the [2Fe-2S] center. Stopped-flow spectroscopic experiments confirmed the independent nature of each cofactor by showing that FAD is reduced by NADH at a similar rate in both E 3(apoFeS) and holo-E 3 [253]. After the FAD is reduced to the hydroquinone form, the subsequent electron transfer to the [2Fe-2S] center was found to be pH-dependent. At pH 7, the equilibrium favors the hydroquinone and oxidized [2Fe-2S] 2⫹ state of the two-electron reduced enzyme, whereas at pH 10 the flavin semiquinone radical and reduced [2Fe-2S] ⫹ state are favored. Spectroelectrochemical studies substantiated this pH dependence by showing that the redox potentials of both the FAD and the [2Fe-2S] center change with respect to pH [254]. Importantly, the midpoint potential of the FAD was dramatically altered by pH, changing from ⫺212 mV at pH 7.5 to ⫺273 mV at pH 8.4 versus a shift of ⫺257 to ⫺279 mV for the [2Fe-2S] center under the same conditions. The ionizable group responsible for the pH dependence is estimated to have a pK a of 7.3 by the stopped-flow studies [253], and N-1 of the flavin is likely the protic position in terms of work with other flavoenzymes [255]. Overall, the proposed electron transport sequence is consistent with the role of E 3 as a 2e ⫺ /1e ⫺ switch, and the evidence provides compelling support for a radical mechanism in the C-3 deoxygenation of 51 (Fig. 20). In light of the fact that PMP–glucoseen adduct

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52 is the ultimate acceptor receiving electrons from E 3, the catalytic role of E 3 in the biosynthesis of ascarylose clearly constitutes a novel example of biological deoxygenation. B.

Mechanism of the Coupled Deoxygenation Reaction

On the basis of the aforementioned evidence, a proposed mechanism of electron transfer in the E 1 –E 3 coupled reaction is presented in Fig. 21. First, NADH binds to E 3 and forms a charge-transfer complex with the oxidized FAD (Int-I) [253]. Chemical modification studies have identified Cys296 as a possible residue that participates in the stabilization of this charge-transfer complex [251]. The bound NADH then transfers a hydride and reduces the FAD to the hydroquinone form (Int-II). Subsequently, electrons are shuttled one at a time from the hydroquinone through the [2Fe-2S] centers of both E 3 and E 1 to reduce the PMP–glucoseen intermediate bound in the E 1 active site as a Schiff base (Int-III to Int-VI). Such

Figure 21 Proposed electron-transfer pathway in the E 1 –E 3 coupled reaction. FAD, flavin adenine dinucleotide; NAD⫹, nicotinamide-adenine dinucleotide; NADH, reduced NAD⫹; Int, intermediate. (Source: Adapted from Ref. 256.)

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a one-electron transfer process can be expected to produce transient radicals, and two kinetically competent organic radicals formed during this process have been observed by using both stopped-flow spectrophotometry and freeze-quench EPR spectroscopy [253,256]. The first radical is clearly the flavin semiquinone discussed in the previous section; the nature of the second radical is less well characterized. Nonetheless, the available evidence supports the assignment of the second radical as a phenoxyl radical having its unpaired spin localized mainly on the C-3 oxygen of the PMP in the PMP–substrate Schiff base [256]. After the second electron transfer, the resulting two-electron reduced Schiff base is hydrolyzed to release the product and end one catalytic cycle (Int-VI to Int-VII). It should be pointed out that during the first turnover two additional NADH molecules are needed to prime the enzyme complex for steady-state catalysis (cycling among Int-V, Int-VI, and Int-VII). Such priming is common for redox systems like P-450 enzymes and dioxygenases.

C.

Comparative Analysis of the Catalytic Role of Each Cofactor

1. The FAD Cofactor In the FNR family of reductases, the flavin cofactor generally serves as a switch between one- and two-electron chemistry [257]. As a member of this family, E 3 acts in the same manner by shuttling electrons from NADH, a known two-electron donor, to the E 1 and E 3 [2Fe-2S] centers, which are single-electron acceptors. However, the flavin in other enzyme systems involved with deoxygenation reactions mentioned in this review appears to act directly as a catalytic redox center rather than as part of an electron shuttle. For example, the enzyme 4-hydroxybutyryl-CoA dehydratase (Section II.D.3.a) utilizes an FAD cofactor to store either a hydride or a single electron from the substrate transiently, but the cofactor is regenerated during the catalytic cycle with no net change in oxidation state. Similarly, other enzymes in anaerobic fermentation (Section II.D.3.b) and chorismate synthase (Section II.D.3.c) all leave the flavin cofactor in the same oxidation state at the end of each turnover as in the beginning. On the other hand, one equivalent of NADH must be consumed by E 3 to reduce the FAD for another round of turnover, as the reducing equivalents are not catalytically recycled. In this role, the function of E 3 more closely resembles that of the thioredoxin reductase and/or glutaredoxin reductase that participates in the regeneration of the redox-active thiol pair in RNR at the end of each catalytic cycle. However, E 3 mediates an electron transfer to reduce the PMP–glucoseen intermediate 52 to give the product 54, not to regenerate E 1 after 54 has formed (Fig. 20). Thus, there is sufficient evidence to indicate that the roles of the flavin cofactor in E 3 and other flavin-dependent hydrolases are markedly different.

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2. The [2Fe-2S] Center Cofactors Primarily, the metal cofactors in deoxygenation reactions catalyzed by enzymes like carbonic anhydrase and enolase (Section II.B) serve as Lewis acids to polarize the C-O bond and facilitate the departure of the hydroxyl group. The labile iron atom in the [4Fe-4S] center of the aconitase family of enzymes (Section II.C) acts in an identical manner. Whether the [4Fe-4S] clusters of the bacterial anaerobic fermentation enzymes (Section II.D.3) participate as a Lewis acid or as an important structural element (or both) remains to be established. Nonetheless, it is clear that these cofactors are not involved with electron transfer in the reaction. A notable exception to this group of oxygen-sensitive iron–sulfur enzymes is the [2Fe-2S] center, containing spinach dihydroxyacid dehydratase (Section II.C.2), which can function in the presence of oxygen. However, the catalytic role of this [2Fe-2S] center seems similar to that of the iron–sulfur center of aconitase. On the contrary, the [2Fe-2S] centers of E 1 and E 3 transfer electrons in a redox reaction (Section III.A). Removing this cofactor does not significantly affect the rate at which E 1 forms the dehydrated glucoseen intermediate 52 (Fig. 20), so the [2Fe-2S] center does not appear to act as a Lewis acid to facilitate loss of the hydroxyl group. Although the function of the iron–sulfur clusters of E 1 and E 3 as part of an electron relay is well precedented, the synergetic involvement of the [2Fe-2S] centers and PMP in redox chemical processes distinguishes this deoxygenation from other C-O bond cleavage events mediated by iron–sulfur centers. 3. The Pyridoxamine 5′-Phosphate Cofactor The B 6 coenzymes play an essential role in a wide array of biological transformations that include deamination, transamination, decarboxylation, racemization, βand γ-elimination/substitution, transsulfurization, among others. In these reactions, the coenzyme exists in the PLP form, except transaminases, which also utilize PMP as a result of the interconversion between these two forms of the coenzyme during catalysis. Significantly, E 1 is the best characterized enzyme that catalyzes a reaction using PMP without oscillating with PLP. Though the mechanistic characterization is in the early stages, a PMP-dependent dehydratase in the biosynthetic pathway of the nucleoside antibiotic blasticidin S may also be a member of this class [258]. Other hydrolases that employ coenzyme B 6 as a cofactor, such as serine and threonine dehydratase (Section II.D.1), are all PLPdependent, and their reactions involve the formation and stabilization of an αanion typical for β-eliminations catalyzed by this class of enzymes. The fact that the PMP cofactor in E 1 directly participates not only in the β-elimination but also in the subsequent electron transfer reduction via a radical mechanism clearly places E 1 into a class of its own.

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It is noteworthy that the mechanism of lysine-2,3-aminomutase, another coenzyme B 6 –dependent enzyme, also includes an organic radical intermediate [259–261]. In this reaction (Fig. 22), homolytic cleavage of AdoMet generates a 5′-deoxyadenosyl radical that abstracts a hydrogen atom from the lysine substrate (55) and initiates the radical rearrangment via the PLP cofactor to give 56. Though this reaction demonstrates that a B 6 coenzyme can participate in radical reactions, it is quite different from the reaction catalyzed by E 1. Rather than being generated by a free radical–mediated hydrogen atom abstraction of the substrate, the radical in E 1 –E 3 catalysis is generated by direct single-electron reduction of the highly conjugated cofactor–substrate complex 52 (detailed in Fig. 23). Presumably, the PMP–glucoseen Schiff base 52 tautomerizes to the quinone methide species 57 before the facile reduction of this reactive quinone methide

Figure 22 Proposed mechanism for the radical-mediated rearrangement catalyzed by lysine-2,3-aminomutase. (Source: Adapted from Ref. 259.)

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Figure 23 Detailed electron-transfer mechanism proposed for the E 1 –E 3 coupled reaction. (Source: Adapted from Ref. 256.)

[262,263]. Some related enzymes involved in the biosynthesis of deoxysugars other than ascarylose [111] have been speculated to take advantage of this inherent redox capability to catalyze similar deoxygenations.

IV.

CONCLUSIONS

In this review, we have illustrated the unique nature of the C-O bond cleavage event catalyzed by the E 1 –E 3 coupled enzyme system. These enzymes recruit several cofactors to achieve a common biological transformation, albeit via a novel mechanism. The PMP cofactor is directly responsible for the reversible dehydration catalyzed by E 1. Electrons are then transferred from the biological reducing agent NADH through an FAD cofactor and two [2Fe-2S] centers of E 1 and E 3 to drive the deoxygenation to completion and regenerate PMP. This

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generation of a deoxysugar via radical intermediates is reminiscent of the ribonucleotide reductase reaction (Section II.E.2). However, the details of the mechanisms are quite distinct. Whereas a catalytic protein radical directly oxidizes the substrate to create a substrate radical in RNR, no radical seems to form on a protein residue in E 1, and the transient radical is generated by single-electron reduction instead of oxidation. Interestingly, the characteristics of the proposed quinone methide that undergoes single-electron chemical processes is somewhat analogous to the catalytic properties of topa quinone, tryptophan tryptophylquinone, and pyrroloquinoline quinone [264]. This analogy opens the possibility that B 6 cofactors may also exhibit chemistry characteristic of these newly discovered quinone coenzymes. With a novel mechanism and these intriguing hypotheses, E 1 may be the prototype for a new class of coenzyme B 6 –dependent enzymes that use an alternate chemistry to catalyze transformations not normally expected for this cofactor. This review also summarizes the current knowledge on characterized enzymes that carry out deoxygenation reactions. The timeliness of this effort is evident in the large number of recent advances in our understanding of these important catalysts. This transformation takes place in such diverse biological processes as energy metabolism, DNA replication, and drug biosynthesis. Not surprisingly, for the remarkable breadth of compounds that undergo deoxygenation, several mechanistic themes have evolved to achieve this transformation. The various themes, ranging from no cofactor requirements to protein radical involvement, seem to be finely tuned toward the reactivity of the substrates, though transition-state stabilization by binding in the active site unquestionably plays an important role in enzyme efficiency [89]. Unfortunately, it is difficult to establish a meaningful correlation between the reactivity of the substrate and the possible mechanism of a given deoxygenation because of the variability in active-site topology and a lack of detailed thermodynamic information on all the reactions. This difficulty is highlighted by the fact that even enzymes that catalyze the same reaction, such as deoxynucleotide biosynthesis (Section II.E.2), may exhibit subtle and sometimes glaring mechanistic differences. These differences are a testament to the evolutionary diversity of biological CEO bond cleavage events, and they will continue to challenge and inspire the ingenuity of enzymologists well into the future. As more enzyme systems become characterized, it will be interesting to see what other novel deoxygenations may be discovered.

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237. L. Kenne and B. Lindberg, in The Polysaccharides, Vol 2. (G. O. Aspinall, ed.), Academic Press, New York, p. 287 (1983). 238. C. R. H. Raetz, Annu. Rev. Biochem., 59: 129 (1990). 239. O. Lu¨deritz, A. M. Staub, and O. Westphal, Bacteriol. Rev., 30: 192 (1966). 240. P. A. Rubenstein and J. L. Strominger, J. Biol. Chem., 249: 3776 (1974). 241. P. A. Rubenstein and J. L. Strominger, J. Biol. Chem., 249: 3782 (1974). 242. T. M. Weigel, L.-d. Liu, and H.-w. Liu, Biochemistry, 31: 2129 (1992). 243. Y. Lei, O. Ploux, and H.-w. Liu, Biochemistry, 34: 4643 (1995). 244. J. S. Thorson and H.-w. Liu, J. Am. Chem. Soc., 115: 7539 (1993). 245. T. M. Weigel, V. P. Miller, and H.-w. Liu, Biochemistry, 31: 2140 (1992). 246. P. A. Pieper, Z. Guo, and H.-w. Liu, J. Am. Chem. Soc., 117: 5158 (1995). 247. M. M. Palcic and H. G. Floss, in Vitamin B6 Pyridoxal Phosphate, Chemical, Biochemical, and Medical Aspects, Vol. I, Part A (D. Dolphin, R. Poulson, and O. Avramovic, eds.) Wiley Intersciece, New York, p. 25 (1986). 248. J. S. Thorson, S. F. Lo, H.-w. Liu, and C. R. Hutchinson, J. Am. Chem. Soc., 115: 6993 (1993). 249. J. S. Thorson, and H.-w. Liu, J. Am. Chem. Soc., 115: 12177 (1993). 250. V. P. Miller, J. S. Thorson, O. Ploux, S. F. Lo, and H.-w. Liu, Biochemistry, 32: 11934 (1993). 251. O. Ploux, Y. Lei, K. Vatanen, and H.-w. Liu, Biochemistry, 34: 4159 (1995). 252. S. F. Lo, V. P. Miller, Y. Lei, J. S. Thorson, H.-w. Liu, and J. L. Schottel, J. Bacteriol., 176: 460 (1994). 253. G. T. Gassner, D. A. Johnson, H.-w. Liu, and D. P. Ballou, Biochemistry, 35: 7752 (1996). 254. K. D. Kim, P. A. Pieper, H.-w. Liu, and M. T. Stankovich, Biochemistry, 35: 7879 (1996). 255. F. Mu¨ller, J. Vervoort, C. P. M. van Mierlo, S. G. Mayhew, W. J. H. van Berkel, and A. Bacher, in Flavins and Flavoproteins (D. E. Edmundson, and D. B. McCormick, eds.), Walter de Gruyter, Berlin, p. 261 (1987). 256. D. A. Johnson, G. T. Gassner, V. Bandarian, F. J. Ruzicka, D. P. Ballou, G. H. Reed, and H.-w. Liu, Biochemistry, 35: 15846 (1996). 257. P. A. Karplus, M. J. Daniels, and J. R. Herriott, Science, 251: 60 (1991). 258. S. J. Gould and J. Guo, J. Am. Chem. Soc., 114: 10176 (1992). 259. M. D. Ballinger, P. A. Frey, and G. H. Reed, Biochemistry, 31: 10782 (1992). 260. M. D. Ballinger, P. A. Frey, and G. H. Reed, Biochemistry, 34: 10086 (1995). 261. W. Wu, K. W. Lieder, G. H. Reed, and P. A. Frey, Biochemistry, 34: 10532 (1995). 262. H.-U. Wagner and R. Gompper, in The Chemistry of the Quinonoid Compounds (S. Patai, ed.) John Wiley & Sons, New York, p. 1145 (1974). 263. K. Karabelas and H. W. Moore, J. Am. Chem. Soc., 112: 5372 (1990). 264. J. P. Klinman and D. Mu, Annu. Rev. Biochem., 63: 299 (1994).

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14 The Molecular Mechanism of Amyloidosis in Alzheimer’s Disease Michael G. Zagorski Case Western Reserve University, Cleveland, Ohio

I. INTRODUCTION Amyloidosis is the process in which normally innocuous, soluble proteins deposit as insoluble amyloid fibrils, resulting in organ dysfunction and death [1]. This condition is generally a disease of aging and accompanies many widespread medical ailments such as cancer, rheumatoid arthritis, Alzheimer’s disease (AD), chronic renal dialysis, familial amyloid polyneuropathy, spongiform encephalopathies, maturity-onset diabetes, and systemic amyloidosis. The amyloid deposits are characterized by distinct tinctorial properties and fibrils of 7–10 nm in diameter, in which the primary components are aggregated proteins with antiparallel cross-β-pleated sheet structures [2,3]. In AD, the major protein constituent of the amyloid deposits is the ‘‘βpeptide,’’ which varies in length and consists of 39–43 amino acids. There are also other proteins (presently 15 known) that share the ability to form insoluble amyloid fibrils, which include the amyloid A fragment, amylin, cystatin C, transthyretin, prion, and related fragments. The amyloid proteins and their deposits tend to be organ- and disease-specific. For example, the protein amylin is localized on the pancreas of patients with maturity-onset diabetes whereas the β-peptide deposits mainly in the brain of AD patients. Except for their ability to aggregate into β-sheet amyloid fibrils, the majority of these proteins do not share any similarities in their primary amino acid sequence or their presumed biological functions. Most experts agree that the production of amyloid is harmful and may play an important role in the disease processes. During amyloidosis, monomeric proteins bind together, by way of hydrogen or electrostatic bonding interactions,

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producing dimeric, trimeric, tetrameric, and larger oligomeric species. Eventually, a critical aggregated size that ultimately precipitates as amyloid is reached. It is also thought that the mechanism of amyloidosis for all amyloid proteins involves a common pathway [4,5]. One idea, which was formulated by us, invokes the conversion of soluble, nontoxic α-helical structures into toxic, oligomeric β-sheet structures. These transformations occur with the prion protein [6] and the amyloid β-peptide of AD [7,8]. The β-peptide results from the processing of a larger amyloid precursor protein (APP) [9–11] (Fig. 1). The biological functions of both the APP and the β-peptide are currently unknown, although they are believed to play roles in neuronal homeostasis, cell adhesion, G-protein coupling, and/or oxidative stress. In humans, soluble β-peptide is ubiquitous in biological fluids [12–15], and AD patients also have larger quantities of the insoluble β-peptide as amyloid [16– 18]. Extensive genetic and biochemical studies with early-onset AD cases have identified mutations in the APP gene [11,19], and the genes for other proteins

Figure 1 Overview of the formation of β-peptide from the amyloid precursor protein (APP) that contains 695 residues. The amino acid sequences of the β-(1–28) and β(1–42) peptides are shown. Depending upon conditions, in solution the β-peptide exists in distinct conformations, whereas in the amyloid deposit only the oligomeric β-pleated sheet structure is present.

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called the presenilins, which are larger proteins (greater than 400 residues) containing six to nine transmembrane domains [11,20–22]. All of the mutations support a role for amyloid production in AD, particularly the ‘‘longer’’ 42-residue β(1–42) peptide rather than the ‘‘shorter’’ 40-residue β-(1–40) peptide. Selective processing in familial AD patients with APP and presenilin mutations results in the favored production and deposition of the β-(1–42) rather than the more soluble, shorter 40-residue β-(1–40) peptide [23–26]. Additional research supports the release of the β-(1–42) by a specific protease [27]. Together, these results, along with other biophysical studies [7,28,29], indicate that less soluble β-(1– 42) may be the actual culprit in initiating amyloid formation in AD. The levels of β-peptide aggregation correlate with the neurotoxicity to cortical and neuronal cell cultures [30–32], and there is a direct association between the accumulation of β-peptide into amyloid and the severity of dementia in AD [33–35]. In AD, a major goal of research is to uncover a therapeutic procedure that keeps β-peptide in solution long enough to be degraded/excreted and thus prevents it from precipitating into amyloid. Once deposited as dense amyloid plaque cores, the surrounding nerve cells are dystrophic or dead, and the β-peptide becomes highly resistant to further proteolysis [36–38]. Thus, a basic understanding of the molecular mechanisms of β-amyloidosis, particularly in regard to unraveling the structures of the early-folded and soluble intermediates, may be critical to the development of an efficacious therapeutic procedure to prevent AD. The purpose of this review is to highlight certain aspects of our research work with the amyloid β-peptide of AD. This project was started in early 1990 during my appointment at the Suntory Institute of Bioorganic Research (SUNBOR) in Osaka, Japan, and is now being continued in my research group at Case Western Reserve University (CWRU). At the time of its inception, very little was known about the amyloidosis of the β-peptide. The lack of information mainly resulted from the difficulty in studying the β-peptide, which is due to its high insolubility and rapid aggregational properties. Once produced in the brain, it was thought, the β-peptide rapidly aggregates into insoluble β-pleated sheet structures, the probable structure in amyloid plaques [39–41]. Over the past few years, however, much has been learned about the biochemical mechanisms of β-peptide production and aggregation into amyloid. Despite the obstacles with handling the β-peptide, the work we painstakingly began at SUNBOR uncovered many important properties of its aggregation. This research was carried out with excellent colleagues, and later at CWRU with many outstanding graduate students and other collaborators who are acknowledged at the end of this article. Most notably, our initial work at SUNBOR established that, under certain solution conditions, the synthetic β-peptides can exist in monomeric (nonaggregated) α-helical conformations. However, if the solution conditions are appropriately changed, the β-peptide can rearrange (α-helix → β-sheet) and even-

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tually precipitate to produce insoluble amyloidlike deposits [7]. Our unique approach, by carefully studying the solution conformations of synthetic β-peptides, uncovered these new properties, whereas at that time studies on natural β-peptides isolated from amyloid plaques in other laboratories around the world failed to detect these important features. These results raised the possibility that β-peptide may exist in vivo in a soluble, nonaggregated state and that an understanding of the mechanisms and the intrinsic solution conditions for the α-helix → β-sheet conversion should be sought. In fact, in our initial report in [7] we proposed that the β-peptide may normally exist in human biological fluids in a soluble form, and that the longer 42-residue peptide results from an abnormal proteolysis. Significantly, both of these propositions were later shown to be correct in AD patients [23–26]. As mentioned before, familial mutations for early-onset AD cases have greater amyloid production, which results from the favored production and deposition of the β-(1-42) rather than the more soluble, shorter 40-residue β-(1-40) peptide [23–26]. Thus, our chemical and structural work illustrated an important point in biomedical research: detailed basic knowledge of the solution chemical properties and the structures can facilitate parallel pharmacological studies. On the basis of biophysical studies with synthetic β-peptides, x-ray fibril diffraction established that the major conformation in the amyloid plaques is an oligomeric, antiparallel cross-β-pleated sheet structure [39–41]. In regions of the brain free of the amyloid plaque, synchrotron Fourier transform infrared microspectroscopy showed that the β-peptide can fold into α-helical or random coil structures [42]. In solution and before precipitation as amyloid, the β-peptide can adopt α-helix, random coil, β-turn, and/or β-sheet structures, in relative ratios that are strongly dependent on the solution conditions [7,40,43–46]. The α-helical and β-sheet/β-turn structures are monomeric, whereas the β-sheet structure in solution is oligomeric, is toxic [31,32,47] and is a precursor to that found in the amyloid plaque. Our work at SUNBOR was the first to demonstrate that the solution conformations vary with pH and solvent [7,48], as later confirmed by other laboratories [45,49,50]. The spectroscopic techniques we employ include circular dichroism (CD), nuclear magnetic resonance (NMR), and infrared (IR) spectroscopy. A major portion of our initial studies used the β-(1–28) peptide (Fig. 1), which contains residues 1–28 and is an appropriate structural model for the complete β-(1–42) peptide. Although the β-(1–28) peptide is incapable of adhering to the surface of preexisting plaques [51,52], it produces soluble monomeric α-helical structures [7,50] as seen in the brain [42], and also plaquelike oligomeric β-sheet structures, similar to those found in natural amyloid plaques [40,43]. The hydrophobic 29–42 region increases the rate of aggregation and βsheet production [44,45,48,53] and greatly contributes to the insolubility of the peptide in aqueous solution. Because of space limitations, this review will highlight only specific aspects of our research with the amyloid β-peptide. The topics were selected to provide

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a framework of our approach to elucidating the mechanisms of β-amyloidosis. Our studies have focused on examining the solution-state of the β-peptide, before it begins to precipitate as amyloid. The selected sections are summarized as follows: 1. The extraordinary sensitivity of the β-peptide solution structures (αhelix, β-sheet, random coil) and aggregational states to the environmental conditions such as pH, temperature, and solvent hydrophobicity 2. Our approach to provide reproducible starting points for biophysical studies 3. Progress toward obtaining the three-dimensional solution structure of the physiologically important β-(1–42) peptide 4. Using nicotine as an example, research for the design of β-amyloid inhibitors that may eventually serve as drug targets 5. Proposed biological mechanisms for β-amyloidosis that take into account the in vitro chemical studies

II. SOLUTION CONDITIONS AND THEIR IMPACT ON THE STRUCTURE A.

Peptide Preparation and Characterization

All peptides were either purchased from commercial sources or prepared in our own laboratory on solid phase using conventional t-Boc or Fmoc strategies [54]. Because of their widespread use in research, the amyloid β-peptides are sold by many commercial laboratories. We find it most economical to purchase unpurified peptide batches and purify them in our own laboratory by standard reversephase high-performance liquid chromatography (HPLC) using acetonitrile, water, and trifluoroacetic acid (TFA) solutions for elution [48,55]. By purifying the peptides ourselves, we remove the inconsistencies with varying purity levels from different commercial sources. Peptide identities and purity levels are always verified by mass spectrometry and NMR spectroscopy. An important protocol with our current studies is to ensure that the β-peptide adopts a well-defined, monomeric state before starting the biophysical measurements. A major difficulty with solution studies of the amyloid peptides, particularly the more insoluble β-(1–42) peptide, relates to the ease with which they aggregate and precipitate. Although it has been established that the soluble βsheet structure of the β-peptide is neurotoxic [31,32,56], there still exists considerable disagreement about the levels of neurotoxicity [31,57–59]. In fact, several groups still report that synthetic β-peptides show either trophic or toxic responses on neurons in vitro. The reasons for these discrepancies are believed to arise in part from differences in the aggregation states and the structures of the β-peptides.

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Different commercially prepared batches of high-performance liquid chromatography–(HPLC)-purified β-(1–42) peptides can have different starting aggregation states and structures [60], which will then in turn affect their solubility, aggregation rates, biological activities in solution [31,57–59], and ability to reproduce biophysical measurements. The reproducibility of the presumably neurotoxic aggregated β-sheet structure is dependent on many factors, particularly the peptide concentration, ionic strength, and solvent polarity [29,44,48,61]. For example, the aggregation rate is extremely rapid in aqueous acetonitrile solutions, such as those used for HPLC purification of the peptide [62]. The longer the β-(1–42) peptide remains in aqueous acetonitrile solution, the more likely it will become an aggregated β-sheet structure. To overcome the complications described, we developed a simple pretreatment protocol that involves sonicating the dry peptide in neat TFA solution before biophysical measurements [55]. The TFA breaks up the preaggregated peptides and affords monomeric random coil structures. This method ensures that different batches of the purified β-peptides will provide reproducible starting points for biophysical and neurotoxicity studies. B.

Circular Dichroism Studies: Effect of Solvent and pH

Because of the structural instability of the β-peptide, before starting the more time-consuming NMR measurements, we typically first conduct thorough CD studies. This enables us to explore rapidly the influences of solvent, temperature, pH, and peptide concentration on the secondary structures. The CD technique is fairly reliable for measuring peptide secondary structure [63,64], and, unlike NMR, it is easy to use, very sensitive, and capable of working with peptide concentrations down to the micromolar (µM ) range. A remarkable feature of the β-peptides is that the relative ratios of secondary structures are greatly influenced by the solvent hydrophobicity and the pH [7,8,29,44,45,48,49,62,65,66]. In earlier work we showed that, in aqueous trifluoroethanol (TFE) solutions, the β-(1–28), β-(1–39), and β-(1–42) peptides all adopt monomeric α-helical structures at low and high pH, as shown by the intense, negative CD bands at 208 and 222 nm [7,48]. However, at intermediate pH (4–7) an oligomeric β-structure (the probable structure in plaques) predominates, as shown by the weak, broad negative bands at about 214 nm. The CD spectra were analyzed using standard poly-l-lysine curves [63,64] and a graphical summary of the results for the β-(1–28) peptide in 60% TFE is shown in Fig. 2A. These experiments were performed by recording CD spectra for two identical peptide solutions at high and low pH, followed by a stepwise increase or decrease of the pH. To prevent errors that can arise from secondary structural changes over time, CD spectra were recorded within 10 min after each pH alteration.

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Figure 2 Graphs showing the variation of secondary structures of the β-(1–28) peptide with pH in 60% TFE (a) and water alone (b) from CD data; the β-sheet structure is favored at midrange pH (4–7), particularly in 60% TFE solution. TFE, trifluoroethyl; CD, circular dichroism; —䊐—, % helix; —䉫—, % sheet; —䊊—, % random coil.

Trifluoroethanol is a structure-promoting solvent that encourages intramolecular hydrogen bonding [67–69]. In solutions containing up to 80% TFE, a synthetic peptide that comprises residues 29–42 [β-(29–42)] remained in a β-sheet conformation, indicating that this segment may direct folding of the complete β-(1– 42) peptide to produce the β-pleated sheet structure found in amyloid plaques. As expected, the secondary structures for the completely hydrophobic β-(29–42) peptide were not altered with pH. Under equivalent conditions, we found that an α-helical structure is more easily formed in the shorter β-(1–39) than in β-(1– 42), indicating that β-(1–42) forms a more stable β-sheet structure than does β(1–39). The β-(1–28) peptide at pH 2.8 and 7.4 in 60% TFE solution is 58% and 47% α-helix, respectively, with the remainder being mostly random coil (Fig. 2A). The conformation was independent of peptide concentration throughout the 1.7- to 3000-µM range; this finding, together with those of other NMR studies [70], establishes that the α-helical structure is monomeric. These solutions were stable over time, and no changes in the relative amounts of α-helix and random coil were observed over a 2-month period. Within the pH 4–7 range, however, mixtures of β-sheet, α-helix, and random coil coexist in solution. Moreover, for peptide solutions kept at pH 4–7, the relative proportions of α-helix, β-sheet, and random coil structures are both time- and concentration-dependent. At pH 4–7, the amount of β-sheet structure increases over time and is accompanied by

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precipitation as an amyloidlike β-pleated sheet deposit. The relative proportion of β-sheet structure increases at higher peptide concentrations, consistent with its highly aggregated state in the amyloid plaques of AD brain. In water with or without TFE, the β-sheet structure is preferred at pH 4– 7, whereas the random coil and α-helical conformations are favored at pH 1–4 and 7–10. A summary of the secondary structures observed for the β-(1–28) peptide in water solution as a function of pH without any cosolvents is shown in Fig. 2B. Analysis of the CD spectra using standard poly-l-lysine curves showed that, for all pH values, 80%–90% of the major conformation was random coil, except at pH 6.0, where approximately 52% β-sheet and 42% random coil conformations coexist (Fig. 2B). The presence of an isodichroic point in the CD data indicates that only two conformations (random coil and β-sheet) are in equilibrium. By contrast, there are two isodichroic points for the CD data in 60% TFE [48]. At pH 4–7 in 60% TFE, the α-helical content drops abruptly and the β-sheet content rises, whereas the random coil content remains relatively unchanged. These data suggest that, in TFE solution at midrange pH, an α-helix → β-sheet conversion occurs directly without a random coil intermediate. However, in TFE solution at low and high pH little, if any, β-sheet structure exists, and instead α-helix and random coil structures coexist. If an α-helix → random coil → β-sheet transformation occurred at pH 4–7, then an increase in random coil structure should occur. Related α-helix (soluble, membranebound) → β-sheet (insoluble, aqueous solution) conversions of other peptides and proteins, including the scrapie prion proteins, are well known [6,71]. C.

Circular Dichroism Studies: Effect of Micelles and pH

To examine the effect of a lipid environment on the solution structures of the βpeptide, additional CD studies were undertaken in micellular solution. Micelles adequately mimic a membranelike or bilayer environment and are frequently employed in the structural studies of peptides and proteins [66,72,73]. In earlier work we proposed that the β-peptide adopts an α-helical structure in lipid solutions [48,74], which may be pertinent to the native structure when bound to lipoproteins and albumin in human plasma [75]. The present studies were undertaken to explore the effect of pH and the surface charge of micelles on the solution structures of the β-(1–28) peptide. Shown in Fig. 3 are graphs summarizing the percentage secondary structures with negatively charged, positively charged, and neutral micelles at different pH values. To prevent possible structural perturbations over time, the experiments were performed by recording CD spectra for two identical peptide solutions at high and low pH, followed by a stepwise increase or decrease of the pH. The results establish that the α-helical structure is produced in the physiological pH range with either the negatively charged sodium dodecyl sulfate (SDS) or the

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positively charged dodecyltrimethylammonium chloride (DTAC) micelle. The chemical structures of these and other micelles are presented in Fig. 4. The zwitterionic, neutral dodecylphosphocholine (DPC) micelle does not produce significant α-helical structure above pH 4. For the CD spectra obtained in all three micelles, there are mixtures of two conformations, random coil and α-helix. This interpretation is consistent with the presence of isodichroic points, together with α-helical and random coil bands observed in the CD spectra (data not shown). In 20-mM SDS solution, which is above the critical micelle concentration of 8 mM, the peptide adopts predominantly α-helical structure at low to weakly basic pH (Fig. 3), as shown in the CD spectra by a strong positive band at 195 nm and negative bands at 208 and 222 nm. The α-helical content remains relatively constant within two ranges, pH 2–4 and 7–9, which is approximately 80% and 50% α-helix, respectively, with the remaining structure being random coil. At pH 10 and 11 the peptide adopts mostly random coil structures, as demonstrated by the presence of the intense negative band at 198 nm. No β-sheet structure exists in any of the micelle solutions, regardless of pH. These data are in stark contrast to the results in TFE–water solution, especially at pH 4–7, where an αhelix → β-sheet conversion occurs. One possibility is that the histidine side chains of the β-peptide are shielded from the solvent by the micelles, as these residues are critical to stabilizing the β-sheet structure [70,76]. These studies establish that membranelike media with a charged surface stabilize the soluble α-helical conformation of the β-peptide in the physiological pH range. D.

Nuclear Magnetic Resonance Studies

Another analytical technique we use with the amyloid β-peptides is NMR spectroscopy [77,78]. In comparison to the more qualitative techniques such as CD and IR spectroscopy, the NMR approach has the advantage of being able to determine specifically where a particular secondary structure is located within the primary sequence. Furthermore, for drug design the NMR method can provide valuable information about protein dynamics and specific data about the individual amino acid side chains that bind to a particular ligand [79,80]. With sufficient distance and dihedral angle constraints, the NMR approach is also capable of rendering a complete three-dimensional structure. However, for solution NMR applications, a general requirement is that the protein predominantly adopts one well defined and sufficiently stable structure usually for a period of several days and at relatively high concentrations (up to several milli-molar). This stability is mandatory when long, multidimensional data acquisitions may be run over a period of several days. These necessities have greatly restricted NMR studies of the amyloid β-peptides. In water solution, freshly dissolved monomeric β-(1– 42) peptide adopts a mixture of rapidly interconverting α-helix, random coil, and aggregating β-sheet structures. This rapidly equilibrating mixture causes severe

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Figure 4 Chemical structures of the sodium dodecyl sulfate (SDS), dodecyl phosphocholine (DPC), hexadecyl-N-methylpiperidium (HMP), myristyltrimethylammonium (MTMA), dodecyltrimethylammonium chloride (DTAC), N-tetradecyl-N, N-dimethyl-3ammonio-1-propanesulfonate (Z 3–14), Congo red (CR), (S)-(⫺)-nicotine, and (S)-(⫺)cotinine. Depending on whether the compound has a net positive or negative charge, the counterions are either sodium or bromide.

line broadening and prevents the detection of well-resolved NMR signals. Additional serious problems are the subsequent aggregation and precipitation, particularly with the longer β-(1–42) peptide, that readily occur at the higher peptide concentrations required for NMR spectroscopy. It is possible, however, to perform NMR studies of fragments that are less likely to aggregate such as the β(1–28), β-(12–28), β-(10–35), and β-(25–35) in water solution, or with the complete β-(1–42) peptide in water solution containing TFE or micelles [61,70,74, 81–83]. One of the most useful NMR parameters is the nuclear Overhauser effect

Figure 3 Graphs depicting the variation of random coil and α-helical secondary structures of the β-(1–28) peptide with pH in SDS, DPC, and DTAC micelle solutions from CD data. The chemical structures of the micelles are shown in Fig. 4. The charged SDS and DTAC micelles stabilize the α-helix at physiological pH and β-sheet structure was not observed in any micelle solution. SDS, sodium dodecyl sulfate; DPC, dodecyl phosphocholinc; DTAC, dodecyl trimethylammonium chloride; DPC, dodecyl phosphocholine; CD, circular dichroism.

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(NOE), which provides through-space information between protons that are ˚ of each other and are dipolar-coupled [84]. The pattern of interresidue within 5 A NOEs and the magnitude of another parameter, the amide (NH) to αH J-coupling constant, together permit an accurate representation of the secondary structure of proteins [77,78,85,86]. Another useful marker for secondary structures relies on the αH chemical shifts, with α-helices showing upfield shifts relative to βsheets [87,88]. The major secondary structural elements in proteins are α-helices, turns, parallel β-sheets, and antiparallel β-sheets. The NOE patterns, backbone J-coupling constants, and αH chemical shifts are distinct for these structures. The α-helix has smaller NH-αH J-coupling constants (⬍6.0 Hz), relative to β-sheets (⬎7.0 Hz). In addition, the α-helix can be identified by strong NN(i, i ⫹ 1), αN(i, i ⫹ 3), and αβ(i, i ⫹ 3) NOEs, whereas β-sheets are characterized by very strong αN(i, i ⫹ 1) NOEs and the absence of any other NOEs among the backbone protons. The antiparallel β-sheet exhibits NOEs between αH protons on adjacent strands whereas these are replaced by NOEs between NH and αH protons on adjacent stands for the parallel β-sheet. There are also differences in NH exchange rates, in which the first four residues in helices, the first three residues in a 310 helix, and every second residue in the peripheral strands of β-sheets have greater exchange rates than neighboring NHs. Similarly, there are characteristic patterns of NOEs, spin–spin couplings, and amide proton exchange rates for turns of types I, II, I′, and II′, and half-turns derived from type II tight turns. The first NMR studies were performed with the β-(1–28) peptide and these involved two-dimensional (2D) NMR studies below pH 3 in 60% TFE-d3, [7,70]. According to earlier CD work, under these conditions the β-(1–28) peptide should fold into a largely α-helical structure (Fig. 2) that is stable and amenable to long-term 2D NMR data acquisitions. This initial work represented the first high-resolution NMR data obtained with an amyloid peptide in solution. Subsequent studies done in our laboratory used perdeuterated-DPC (DPC-d38) and perdeuterated-SDS (SDS-d25) solutions [74] under conditions that also maintained stable α-helices (Section III.C). Analysis of the 2D NOESY data showed numerous NN(i, i ⫹ 1), αN(i, i ⫹ 3), and αβ(i, i ⫹ 3) NOEs that established a predominantly α-helical structure, in accordance with the CD data (Fig. 2 and 3). As shown in Fig. 5, the three-dimensional NMR structure in 60% TFE-d3 contained right-handed α-helices (residues 2–11 and 13–27), connected by a bend-centered at Val12 [74]. The bend is in agreement with the observed periodic nature of the αH chemical shifts [87,88] and also with the NMR-derived NH temperature coefficients. At 25°C and below pH 3, nearly identical NOE data were seen in aqueous solutions containing TFE-d3, DPC-d38, or SDS-d25, suggesting that similar tertiary α-helical structures are present under these conditions. However, there are differences in the stabilities of the two α-helical segments (residues 2–11 and 13–27) with temperature. The NH temperature coefficients in SDS-d25 solution showed that both α-helical segments are stable up to 50°C, whereas in 60% TFE-

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Figure 5 The final 95 structures for the β-(1–28) peptide superpositioned using the backbone atoms (pH 3, 60% TFE solution). The N-terminus is shown at the top and the C-terminus is shown at the bottom. The side chains of Val12 and Phe20 are labeled to illustrate their location on the same face of the helix TFE, trifluoroethyl. (Source: From Ref. 74.)

d3 and DPC-d38 solutions, the smaller α-helix (residues 2–11) rapidly unfolds above 25°C with the other helix (residues 13–27) unfolding above 30°C. Both α-helices unfold to random coil structures and refold back to α-helices with cooling. The NH temperature coefficients were determined by recording NOESY data at 10, 25, 35, 40, and 50°C. The random pattern of coefficients suggests that the peptide is not buried within the hydrophobic interior of the SDS-d25 micelle, but instead is located at the surface near the lipid/water interface. These studies demonstrate that the negatively charged SDS micelle provides an environment most conducive to α-helix formation and that this stability is not due to the location of the peptide within the hydrophobic interior of the SDS micelle. To evaluate further the effect of pH on the loss of α-helical and the formation of β-sheet structure, we are currently determining the dissociation constants (pKa) of the aspartic acid (Asp), glutamic acid (Glu), histidine (His), and tyrosine (Tyr) residues under different solution conditions. We have used the β-(1–28)

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peptide as a model for the longer β-(1–42) peptide, since the remaining 29–42 region is devoid of polar and charged amino acid residues. The respective chemical shifts for the 2H, 4H; 3H, 5H; γCH2s, and βCH2 signals of the His; Tyr; Glu; and Asp residues at different pH values are being determined by a combination of 1D and 2D NMR spectroscopy. Preliminary analysis demonstrates that the Glu and His residues have larger pKa values in SDS solution, compared with 60% TFE, DPC, and water alone. This result may partly account for the greater stability of the α-helix in SDS solution. Complete loss of the helix occurs above pH 9, which corresponds to the pKa of two His residues. Below pH 9 the His side chains are predominantly positively charged, that charge should bind them more strongly with the negatively charged SDS surface. A complete summary of these results is presented in a forthcoming paper [89]. With a relatively clear understanding of the effects of solvent, pH, temperature, and lipids on the structures and aggregational properties of the β-(1–28) peptide, we next examined the more difficult, longer, and naturally occurring 42residue β-(1–42) peptide. It was felt that the 1–28 region should function in an analogous manner in both peptides, since the remaining 29–42 region is completely hydrophobic and can be considered a distinct subregion. As it happens, our studies with the β-(1–28) did indeed provide sufficient background knowledge that enabled us to examine the more important β-(1–42) peptide. The β(1–42) peptide can be extremely insoluble and act as a seed that nucleates amyloid formation with the other shorter amyloid peptides [23,28,29]. In addition, the β-(1–42) peptide is the predominant protein component in cerebrovascular amyloid and plaque core deposits [90,91], and it is thought that the longer peptide may be the actual culprit behind amyloid deposition in AD [24,90–92]. Unfortunately, despite its biological importance to AD, biophysical and biological studies have primarily used the shorter peptides since the β-(1–42) is difficult to dissolve and aggregates/precipitates too rapidly. Before beginning NMR studies with the longer peptide, we first conducted CD studies. However, unlike the β-(1–28) peptide, the β-(1–42) peptide in water solution above pH 7 forms an aggregated β-sheet structure and rapidly precipitates at concentrations above 100 µM. Therefore, NMR studies in water solution at physiological pH are clearly precluded. When the CD studies were repeated with 1 molar-equivalent of SDS pH 7.2, the CD spectra obtained at 0, 24, 48, 96, and 120 h were nearly identical, consistent with a predominantly α-helical structure, and significantly, no precipitation was seen. These results suggest that the SDS micelle promotes formation of a predominantly α-helical structure, which is probably similar to that observed for the β-(1–28) peptide. The stability and lack of precipitation of the β-(1–42) peptide provided by the SDS micelle suggest an amenable system for detailed structural analysis by high-resolution NMR methods. This ability of SDS to prevent aggregation is analogous to other recent work, in which the micelle, hexadecyl-N-methylpiperidinium bromide

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(HMP), inhibited aggregation of the β-(1–40) peptide [93]. The chemical structures of HMP, plus other micelles and compounds that are β-amyloid inhibitors, are shown in Fig. 4. These include myristyltrimethylammonium (MTMA), Ntetradecyl-N,N-dimethyl-3-ammonio-1-propanesulfonate (Z 3–14), Congo red (CR) [94], and (S)-(⫺)-nicotine [95]. The first three compounds were selected from a screening of 4300 compounds as β-amyloidosis inhibitors [93]; both HMP and MTMA inhibited β-(1–40) aggregation whereas Z 3–14 showed weak inhibition. The NMR characterization of the β-(1–42) peptide in SDS-d25 solution has proceeded very well. In fact, aqueous solutions of the β-(1–42) at relatively high concentrations (up to 2 mM ) are stable for several months. This is important for multiple 2D NMR data acquisitions that can require several days. The SDS-d25 concentrations were significantly higher than those for the β-(1–28) peptide. Typically, the SDS-d25 concentrations are 80–150 mM, so that the peptide/micelle ratio is always 1 :1; with no salt, the average SDS aggregation number is 62 and the critical micelle concentration is 8 mM [73]. With the exception of minor chemical shift differences for some residues at the N-terminus, 1D NMR spectra obtained at protein concentrations of 1.4, 0.94, 0.56, and 0.16 mM were virtually identical. Therefore, it appears that aggregation does not proceed above the level of a monomer within the concentration range 1.4–0.16 mM. Additional studies at lower peptide concentrations are currently being explored in our laboratory. A contour plot of the complete 2D NOESY spectrum of the β-(1–42) peptide in SDS-d25-H2O solution at pH 7.2 is shown in Fig. 6. The NOESY spectrum ˚. reveals through-space connectivity among protons separated by less than 5 A The numerous well-resolved cross peaks demonstrate that the β-(1–42) peptide is folded, and numerous sequential NN(i, i ⫹ 1) NOEs between adjacent amide protons (NHs) are present (Fig. 7). The NOESY spectra obtained with suppression of the H2O signal by presaturation and with the ‘‘Jump and Return’’ pulse sequence [96] were almost identical in the NH–NH region. This demonstrates that NH signals are not lost by presaturation of the water signal, which can lead to saturation transfer. In particular, strong NN(i, i ⫹ 1) NOEs are observed within the Tyr10–Val24 and Lys28–Ala42 segments. These and other medium-range backbone NOEs, together with the upfield chemical shift positions for the αH signals, are consistent with α-helical structure within these regions [77,87]. The complete proton NMR spectrum has been assigned and further efforts for building the three-dimensional structure are currently under way. Shown in Fig. 6 is a hypothetical model for the association of the β-(1–42) peptide with the SDS micelle. Although not shown, side-chain binding interactions between the positively charged side chains of the β-(1–42) and the negatively charged SDS surface are most likely occurring. These interactions stabilize the α-helix and prevent formation of the aggregated β-sheet, which is the predominant structure in water without SDS. The location of the peptide at the micelle surface,

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Figure 6 The complete 2D NOESY spectrum (600 MHz, mixing time 270 ms) for the β-(1–42) peptide (1.4 mM ) in H2O :D2O (9:1) containing SDS-d25 (80 mM ) at pH 7.2 and 20°C. The H2O signal was suppressed by presaturation. The proposed secondary structure of the β-(1–42) peptide based on the NMR chemical shift, NOE, and NH-temperature coefficient data is shown. The monomeric peptide appears to remain on the surface at the lipid–water interface of the SDS micelle and does not become embedded into the hydrophobic interior. NOESY, nuclear overhauser effect; SDS, sodium dodecyl sulfate; NMR, nuclear magnetic resonance.

rather than the hydrophobic interior, is consistent with the amide-NH temperature coefficients and the rapid deuterium exchange rates. The irregular pattern of the NH-temperature coefficients and the rapid NH-exchange rates are consistent with the peptide being located on the surface of the SDS micelle. If residues were buried within the hydrophobic interior of the micelle, then significantly lower temperature coefficients and NH-exchange rates should be observed. Overall, the secondary structure is defined by an extended structure (residues 1–9), α-helix (residues 10–24), loop (residues 25–27), and α-helix (residues 28–42). The presence of a loop within the Gly25–Ser26–Asn27 segment is supported by the absence of any major backbone NOEs and the downfield chemical shift of the Asn27 αH. Interestingly, the location of this loop is nearly identical to a reverse turn suggested for the β-sheet structure of a β-(10–43) peptide, in which the turn was proposed to be within the Ser26–Asn27–Lys28–Gly29 sequence [44]. Recent NMR studies of a β-(10–35)-CONH2 fragment suggests a turn–strand– turn motif between His13 and Val24 in aqueous solution [51], whereas the β(1–40) peptide forms two α-helical segments between Gln15–Asp23 and lle31– Met35 in 30% TFE-d3 solution [81].

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Figure 7 An expanded NOESY region for the β-(1–42) showing the NH-NH connectivities, as well as other peaks for NOE interactions between the aromatic side chains and the backbone NH. NOESY, nuclear overhauser effect.

III. NICOTINE INHIBITION OF ␤-PEPTIDE FIBRIL FORMATION An intense area of research in AD involves identifying potential inhibitors that either slow down or prevent the precipitation of the β-peptide into amyloid. The recent reports of an inverse relationship between the risk of AD and cigarette smoking [97–102] inspired us to investigate the effects of nicotine and cotinine (structures in Fig. 4) on the solution conformations and aggregational properties of the β-peptide. Nicotine is a major component of cigarette smoke that is metabo-

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lized and excreted as cotinine. Using CD and ultraviolet spectroscopic techniques, we recently showed that nicotine inhibits amyloid formation by the β-(1–42) peptide [95]. Cotinine slows down amyloid formation, but to a lesser extent than nicotine. Although it is not yet known whether a similar nicotine inhibition to amyloidosis occurs in the human brain, these studies may facilitate the design of less toxic and more efficient amyloid inhibitors that are structurally related to nicotine. In this review, we summarize unpublished molecular modeling studies that were not included in the original manuscript [95]. The molecular modeling studies provide a more detailed representation of the nicotine inhibition to βamyloidosis. Our previous NMR studies demonstrated that nicotine binds to the 1–28 peptide region when folded in an α-helical conformation [95]. The β-(1–28) peptide is an appropriate structural model for the complete β-(1–42) peptide, since it produces soluble monomeric α-helical structures [7,50], as well as plaquelike oligomeric β-sheet structures, similar to those found in natural amyloid plaques [40,43]. The hydrophobic 29–42 region increases the rate of aggregation and βsheet production [44,45,48,53] but should not affect the ability of nicotine to bind to the β-peptide. Nicotine is a weak base with pKa of 7.9 [103], and at blood pH of 7.4, nicotine is about 69% ionized (protonated) and 31% nonprotonated. Therefore, if binding does indeed occur, nicotine would be more likely to bind to the polar, hydrophilic 1–28 peptide region, rather than the completely hydrophobic 29–42 region (Fig. 1). On the basis of chemical shift data, the binding between nicotine and the β-(1–28) peptide primarily involves the N-CH3 and 5′CH2 pyrrolidine moieties of nicotine and the His residues of the peptide. The binding is in fairly rapid exchange, as shown by single averaged NMR peaks. A summary of the bound chemical shifts and intermolecular NOEs between nicotine and the peptide region is shown in Table 1. The modeling studies used the NMR data, molecular dynamics, and energy minimization protocols. On the basis of the bound NMR chemical shift data (Table 1), we envisioned a molecule of nicotine bound to the α-helix of the β-peptide, as depicted in Fig. 8. In the α-helical structure, the side chains of His13 and Lys16 are proximate [74] and could conceivably accommodate nicotine. The possibility that the Lys16–ζNH⫹3 is tied up with an intramolecular hydrogen bond to the N3 of His13 was ruled out on the basis of previously published NMR data [70], which showed that the Lys16–ζNH⫹3 is solvent exposed. Besides the bound chemical shift data that support this model, another attractive feature for this proposed binding is the complementary charge distribution between nicotine and the peptide. At pH 7.2 the nicotine pyrrole nitrogen (N1′) will be predominantly protonated (pKa 7.9), whereas the pyridine N1 is not protonated (pKa 3.12) [104]. For the β-peptide, at pH 7.2 the side chain N3 of His13 is not protonated (pKa 6.2–7.0) whereas the Lys–ζNH⫹3 remains protonated up to pH 10–11 [105].

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Table 1 Chemical Shifts, NOEs, and Distances in the Complex of Nicotine and the β-(1–28) Peptide

NMR resonance Peptide His13-2H His13-4H Lys16-ζNH⫹3 Nicotine 2H 6H 4H 5H 2H′ 5-α-CH′ 5-β-CH′ 3CH′2 4CH′2 N-CH3

Intermolecular NOEs His13-4H to 4CH′2 Tyr10-2, 6H to 5-β-CH′ Va118-βH to 2H′ Tyr10-βH to 5-β-CH′ Va112-γCH3 to 5-α-CH′ a

Bound chemical shift (ppm)a ⫺0.22

Minimization and ˚ )b distance restraints (A 6–10 Above the pyridine ring

⫺0.06 0.00 0.10 0.07 0.07 0.03 0.07 0.24 0.31 0.02 0.05 ⫺0.50 0.19

6–10 In plane of His13 ring 6–10 In plane of His13 ring

6–10 In plane of His13 ring 3–5 3–5 3–5 3–5 3–5

Starting distance ˚ )c (A

Final distance ˚ )c (A

10.07

6.84

12.66 12.10

9.94 8.90

11.64

8.69

11.64

7.96

8.97

6.05

˚ A ˚ A ˚ A ˚ A ˚ A

Obtained by subtracting the chemical shifts in nicotine and the peptide from those seen in the complex. The negative shifts are upfield. b For the upfield or downfield shifts, distance restraints refer to a proton located either above or in the plane, respectively, of the nicotine–pyridine and peptide–His13 aromatic rings. Pseudoatoms were used to represent the three equivalent NH⫹3 and CH3 protons. c The starting and final distances, before and after molecular dynamics plus several cycles of energy minimization. For the 2H of His13, the distances refer to its location relative to the center of the nicotine–pyridine aromatic ring and for the Lys16-ζNH⫹3 relative to the N1 of nicotine. NOE, nuclear overhauser effect; NMR, nuclear magnetic resonance.

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One molecule of nicotine was annealed and energy minimized to its lowest energy conformation in water solution at pH 7.2. The nicotine–peptide complex was generated by manually docking the minimized (S)-nicotine molecule to the NMR-derived averaged, energy-minimized β-(1–28) α-helical structure [74]. Pseudoatoms for the five atoms of the imidazole ring of His13 and for the six atoms of the pyridine ring of nicotine were created. The docking was performed in a manner consistent with the diagram in Fig. 8 and the bound chemical shift data (Table 1), which were interpreted as resulting from ring current effects [106]. The N-CH3 of nicotine displays two NMR signals when mixed with peptide: one signal appears downfield from the original unbound shift by 0.19 ppm, and the other is upfield by 0.50 ppm in an approximate 2 :1 ratio, respectively. The two signals probably represent two different orientations of the pyrrole ring when bound to the His residues of the peptide. For the purpose of modeling, we used the more abundant downfield signal.

Figure 8 Proposed structure of the (S)-(⫺)-nicotine/β-(1–28) peptide complex, when the peptide folds as an α-helix, with the N-terminus at the top and the C-terminus at the bottom. The schematic diagram proposes the His13-N3 and Lys16-ζNH⫹3 binding to the pyrrole H-N1′⫹ and N1 of nicotine, respectively.

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The upfield shift of the His 13-2H results from its location above or below the nicotine–pyridine ring; the downfield shifts of the N-CH3, and 5CH′2 protons of nicotine were explained by their location in the His13 imidazole ring plane. Accordingly, the N-CH3 and 5CH′2 protons were positioned to the periphery of the His13 ring, and the His13-2H proton was placed above the pyridine ring (Table 1). The possibility that other aromatic rings in the peptide, such as those of the phenylalanine and tyrosine residues, contribute to the binding and ring current shifts was ruled out, since the NMR peaks for these rings do not undergo ˚ was created for chemical shift changes in the mixture. A restraint file of 6–10 A the preceding protons on the basis of well-known methods that use ring current shifts to relate the position of a proton relative to an aromatic ring [106,107]. For bound chemical shifts less than 0.5 ppm, the distances from the ring center ˚ ) and all bound chemical shifts are less than 0.31 ppm (Table are larger (6–10 A 1). These restraints kept the N-CH3, and 5CH′2 protons within the deshielding plane of the His13 ring throughout the course of the dynamics and minimizations. However, to prevent movement of the His-2H away from the shielding region ˚ restraints between the 2H and of the nicotine–pyridine ring, individual 6–10 A each heavy atom of the pyridine ring were required. To provide a more realistic binding environment and stabilize the charges, the SOAK Assembly operation in INSIGHT II (Biosym, Inc.) was utilized to solvate the nicotine–peptide complex. This involved building a two-layer solvent sphere, which contained 336 and 186 water molecules in the inner and outer ˚ with the spheres, respectively. The diameter of the complete sphere was 21 A ˚ inner sphere taking up 15 A To prevent evaporation of the water during the dynamics, the water molecules in the outer sphere were tethered, whereas the water molecules in the inner sphere were allowed to move freely. Thus, the outer layer provided a boundary and prevented escape of the water molecules in the inner sphere. The starting nicotine–peptide complex was placed in the center of the inner sphere, so that the Val12–Leu17 region completely occupied the sphere (Fig. 9A). The SOAK operation automatically repositioned those water molecules that overlapped with the atoms of the complex. To reduce computation time, a ˚ The cutoff distance defines the minimum range cutoff distance was set to 25 A for atoms that will be considered in the calculation. The atoms that were outside this region (amino acid residues 1–11 and 18–28) were kept fixed. Within the Val12–Leu17 segment, the backbone atoms were tethered and the side chains were completely free to move throughout the dynamics and minimizations. The only distance restrictions within the Val12–Leu17 region involved the 6– 10– ˚ restraints placed on four hydrogen atoms: His13-2H, 5-α-CH′, 5-β-CH′, and to A the pseudoatom for the N-CH3, as shown in Table 1. These restraints were made on the basis of the bound chemical shifts and ring current effects (discussed previously). The computations began with 100 steps of steepest descent energy minimization, followed by molecular dynamics at 300 K for 1 ps, and finally 6000

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(a)

(b)

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steps of conjugate gradient energy minimization. A globally enforced dielectric constant of 1.0, cvff force field, and a template force with a force constant of ˚ 2 were employed. A simple harmonic potential was used for the bond 1000 kcal/A energies, and the cross terms were omitted from the energy expression. Distances are provided for selected protons in nicotine and the peptide in Table 1. The final total energy of the solvated nicotine–peptide complex was ⫺5219 kcal/mole. The main adverse factor that contributed to the total energy were the nonbonded repulsion energies from the fixed atoms in amino acid residues 1–11 and 18– 28. Longer molecular dynamic runs did not produce significantly lower total energies, demonstrating that the final nicotine–peptide complex was not trapped in a local energy minimum. Other annealing methods are currently being explored for obtaining lower energies. An expanded view of the β-peptide/nicotine complex is shown in Fig. 9B, and distances between selected protons are provided in Table 1. As shown, all final distances are substantially shorter, indicating that nicotine and the peptide move closer in space. If an unfavorable interaction were present, then greater distance would be seen. In particular, both the His13-2H and 5-β-CH′ have significantly shorter distances than their initial distances, and these NMR signals likewise show the largest changes in NMR chemical shifts with binding. The ˚; distance between the nicotine-N1 and the Lys16–ζNH⫹3 is reduced by 3.20 A ˚ additionally, the labile nicotine H-N1′ hydrogen is less than 3 A from the N3 of His13. These results support the proposed binding mode shown in Fig. 8, in which nicotine binds to the Lys16-ζNH3⫹ and the His13-N3 by hydrogen bonding. A similar nicotine binding motif may occur with other proteins. In human plasma, 5% nicotine becomes bound to albumin and several serum lipoproteins [108], and in vitro nicotine binds to peptide segments of the nicotinic acetylcholine receptor (AChR). The AChR is a well-characterized protein that produces rapid ligand-gated ion channels [109]. Although the structure of the binding sites for the lipoproteins, albumin, and AChR are not completely characterized, several His and Lys residues are believed to occupy important locations. The His and Lys residues occupy α-helical sections in the α-bungarotoxin binding site of AChR, where nicotine also becomes bound. Chemical modifications or replace-

Figure 9 The three-dimensional model of the nicotine–peptide complex (a and b) after molecular dynamics and energy minimizations. (a) The NMR-derived averaged, energyminimized β-(1–28) α-helical structure is shown with the His13, His14, and Lys16 side chains, and the backbone as a ribbon. The water molecules that solvate the complex are shown in dark gray (a). (b) Expanded view that corresponds to the region within the box in (a) shows the His-2H positioned above the nicotine–pyridine ring, consistent with the NMR data. The modeling studies also suggest that the Lys16-ζNH3⫹ may bind to the nicotine-N1. NMR, nuclear magnetic resonance. (Source: From Ref. 74.)

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ment of the His residues results in ion channel impairment and a decreased affinity for nicotine binding [110–112]. Substitution of Lys185 in a peptide segment corresponding to residues 181–200 of the Torpedo alpha 1 AChR protein reduced the binding of nicotine to the peptide [113]. Thus, the model of nicotine binding shown in Figs. 8 and 9 may be general with other proteins. The model shown in Figs. 8 and 9 may assist in the development of nicotine analogues to retard amyloidosis by preventing an α-helix → β-sheet conformational transformation that is important in the pathogenesis of Alzheimer’s disease. Further studies with additional nicotine analogues are required to support this binding mode. Potential analogues could be synthesized with electron donating substituents in the 4 position of the pyridine ring of nicotine, since this would increase the basicity of the N-1 nitrogen and perhaps strengthen the interaction with Lys 16-ζNH⫹3 . Interestingly, an analogous structural motif may be involved in the binding of the β-peptide to transthyretin, a normal protein component of plasma [114]. A structural model for the complex of transthyretin and the αhelical structure of the β-peptide showed that the side chain of Lys16 forms part of a positive potential binding surface that makes contact with a negative potential surface of transthyretin.

IV.

MECHANISM OF AMYLOIDOSIS OF THE ␤-PEPTIDE

Despite the importance to AD, the underlying mechanisms for the accumulation of the β-peptide into insoluble amyloid remain largely unknown. A popular concept involves an ‘‘amyloid-initiated-cascade’’ phenomenon, in which altered production, removal, and aggregation of the amyloid β-peptide initiate a sequence of events that leads to neuronal death [11,38,115]. It should be kept in mind that other lesions such as the neurofibrillary tangles are also abundant in AD brains, and it may happen that the tangles are more important in the pathogenesis of the disease [116]. The tangles are intracellular deposits consisting of twisted filaments of the cytoskeletal tau protein. However, since the majority of genetic and biological data support a critical role for amyloid formation in AD, the ‘‘amyloidinitiated-cascade’’ hypothesis appears to be the most promising model for drug discovery [11,117]. A hypothetical model for β-amyloidosis is outlined in Scheme 1. This mechanism attempts to take into account the results of our work, as well as work from other laboratories. The various pathways shown in the scheme connect the in vitro biophysical studies to a natural situation that may exist in the brain. The model emphasizes a conformationally driven mechanism, in which the three major solution structures of the β-peptide coexist in equilibrium: random coil (monomeric), α-helix (monomeric), and β-sheet (oligomeric). The aggregated β-sheet

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Scheme 1

is the predominant structural motif in the amyloid plaques. When the aggregation reaches a critical mass, the β-sheet structure precipitates as amyloid, and reconversion back to soluble random coil or α-helical structures is no longer possible [48,62,118]. Recent data suggest that soluble β-sheet dimers are toxic [47], with the larger amyloid plaques that form later further disrupting neuronal circuitry. During β-sheet formation, the β-peptide-induced neurotoxicity increases [31,32], suggesting that the processes involved in amyloid plaque formation are critical in the pathogenesis of AD. The mechanisms whereby the β-sheet structure exerts its neurotoxicity remain unknown [119]. Once released by proteolysis of APP, the β-peptide would be expected initially to adopt a monomeric, random coil conformation. This interpretation is based on the known physiological concentrations of the β-peptide in cerebrospinal fluid, which are in the nano- to picomolar range [91,120]. At these low concentrations, the production of aggregated β-peptide as amyloid is very unlikely [28,29,48,66] since the threshold concentrations required for generation of amyloidlike oligomeric β-sheet structure are much higher 50–350 µM. Presumably, the random coil structure is biologically inactive and would not be involved in specific binding processes to other macromolecules. This suggests that, in order

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to produce amyloid deposits, localized regions of the brain must have much higher peptide concentrations, or alternatively some change in the brain microenvironment must occur to promote β-aggregation. Under different environmental conditions, the β-peptide can fold into another structure dominated by β-sheet and β-turns [51,52,121]. Since this structure adheres to the surface of preexisting plaques, it is probably the immediate precursor to the aggregated β-sheet structure. Conversion of this presumably monomeric structure to aggregated β-sheet structure can occur with reductions in brain pH [122] or by the attachment of the β-peptide to larger macromolecules such as ApoE [123–127], metals, or small amounts of preaggregated peptide that can act as a seed for precipitation [28]. Some of the other components that induce aggregation include glycosaminoglycans (GAGs) [128,129], zinc [130,131], Apo-E [123,125,132,133], glycation [134,135], or (non-Aβ protein component precursor (NACP) [136]. These processes occur during normal aging that can be accompanied by changes of brain microenvironments, including β-peptide concentrations, pH, and membrane integrity. The binding between heparan sulfate proteoglycans and the β-peptide is extremely rapid below pH 7 [128,129,137], and analysis of the precipitates of β-peptide containing sulfate ions or heparan showed that the sulfate moieties of the GAG chains promote extensive lateral aggregation of β-peptides [138]. The heparan sulfate proteoglycans are components in the amyloid plaques. It was proposed that the interactions of GAGs and the β-peptide may stabilize one of the secondary structures and/or accelerate the aggregation properties of the βpeptide. Unpublished data from our laboratory established that the β-sheet structure reacts with heparin to produce an amyloidlike precipitate. Related studies with other peptides have shown that different GAGs can markedly influence their solution conformations and folding properties [139–141], in that heparin promoted formation of a β-sheet structure rather than an α-helical structure. Molecular modeling and NMR studies showed that the His14 and Lys16 side chains, which both reside within the presumed heparan sulfate binding motif [138] are critical to the binding interaction of the GAGs and the β-peptide. At physiological pH, both the α-helix and random coil structures do not bind to heparin (Scheme 1). The general consensus is that amyloid fibril formation is promoted by the binding of components to the β-sheet structure. The components that promote or retard aggregation and toxicity of the β-peptide have recently been summarized [18]. If the β-peptide encounters a hydrophobic environment, similar to TFE or micelles, then an α-helical structure should form. This situation may occur in human plasma when the β-peptide becomes attached to lipoproteins, albumin, ApoJ, or transthyretin [75,114,142]. It is thought that the β-peptide injures cells by a surface membrane effect [9,143] and membrane-bound β-peptide that is closely associated with GM1 ganglioside forms a catalytic seed for amyloid for-

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mation [144,145]. Moreover, a recent report using synchrotron Fourier transform infrared (FTIR) microspectroscopy showed that significant quantities of α-helical structure of the β-peptide are present in the brain gray matter [42]. In all probability, the α-helical structure seen in the brain should be identical with the NMR structure seen with the synthetic β-(1–42) peptide in SDS. As mentioned before, the SDS micelle adequately mimics a biologically relevant, membranelike environment and is extensively employed in biophysical investigations of peptide and protein structures [72,73]. The NMR structure of the β-(1–42) is defined by an extended structure (residues 1–9), α-helix (residues 10–24), loop (residues 25– 27), and α-helix (residues 28–42). Most likely, this structure is nontoxic, since it is monomeric, ordered, and very soluble [146]. The binding interactions of the α-helical structure with other macromolecules such as lipoproteins are probably desirable, since they prevent the β-peptide from aggregating into a β-sheet structure that is toxic and precipitates as amyloid. We believe that the α-helical structure is an appropriate target for the design of amyloid inhibitors. The α-helical structure is monomeric and stable, thus suitable for modern structure-based drug design methods [147]. As shown with our work on the nicotine inhibition to amyloidosis [95], the NMR work established that nicotine binds to the His residues of the α-helical structure, which provides a rationale for the loss of β-sheet production and precipitation. It should be pointed out that evidence to support such a binding interaction in the brain is not yet available. However, if the binding occurs, then a reduction in the αhelix → β-sheet conversion should occur, which would prevent or slow down amyloidosis. Related α-helix → β-sheet conversions for other peptides and proteins [148,149], including peptide segments of the prion proteins [150,151], are well documented. In water solution, freshly prepared, monomeric β-(1–42) peptide adopts mixtures of α-helix, random coil, and β-sheet structures that are in equilibrium. The precipitation drives the equilibrium toward β-sheet structure, so once precipitation has begun the α-helical structure will rearrange to the oligomeric β-sheet structure [α-helix → β-sheet (solution) → β-sheet (precipitate)]. Once plaque formation begins, the oligomeric β-pleated sheet structure that forms is resistant to further proteolysis and turnover [37,152]. It may happen that the monomeric β-sheet/β-turn and/or the early soluble β-sheet structures that form before the plaque are likewise resistant to proteolysis, similar to soluble peptide segments of the amyloid forming prion proteins [151].

V.

CONCLUSIONS

There are currently no effective treatments for AD, and the current drug development process is often cumbersome, taking an average 9 years for approval [153]. The methods discussed in this review, notably the solution NMR technique, can

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provide critical experimental data to confirm that a newly designed compound binds to a protein receptor [79,80], in a manner consistent with pure computational approaches [147,154]. The NMR data may be an important prerequisite to the further development of additional compounds. In closing, we believe that the approaches discussed in this review, which involve combined molecular modeling and NMR, will assist in the design of less toxic analogues to prevent βamyloidosis in AD.

ACKNOWLEDGMENTS I would like to thank the many excellent colleagues who completed the studies shown in this review. These include Colin Barrow, Takashi Iwashita, and Vassil Vassilev from the Suntory Institute for Bioorganic Research, Osaka, Japan, and likewise Keith Marcinowski, Joseph Talafous, Haiyan Shao, Kan Ma, Arthur Salomon, Shu-chuan Jao, Charlene Keane, Erin Clancy, Jing Yang and Rob Friedland from Case Western Reserve University, Cleveland, Ohio. The work was supported in part by grants from the American Health Assistance Foundation, the Suntory Institute for Bioorganic Research, the National Institutes of Aging (AG-08992-06 and AG-14363-01), American Federation of Aging Research, Philip Morris, Inc., Gliatech, Inc., Pfizer, Inc., Smokeless Tobacco Agency, and a Faculty Scholars Award from the Alzheimer’s Association (FSA-94-040). The 600-MHz NMR spectrometer was purchased with funds provided by the National Science Foundation, the National Institutes of Health, and the state of Ohio. I would also like to thank Anita Hong (Anaspec, Inc.), Larry Sayre, and Witold Surewicz for helpful discussions. Finally, I would like especially to thank Koji Nakanishi for his sound advice, particularly his support and encouragement throughout my stay in his groups in both New York and Japan.

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15 Bacteriorhodopsin Structure/ Function Studies: Use of the Demethyl Retinal Analogues for Probing of the Arg82Ala Mutant Rosalie K. Crouch, Donald R. Menick, and Yan Feng Medical University of South Carolina, Charleston, South Carolina

Rajni Govindjee and Thomas G. Ebrey University of Illinois at Urbana-Champaign, Urbana, Illinois

I. INTRODUCTION Bacteriorhodopsin (bR), the only protein in the purple membrane of the archaebacterium Halobacterium salinarium, is a retinal-based, light-transducing pigment [1]. The dark adapted form contains roughly equivalent amounts of the alltrans and 13-cis isomers of retinal (1, Fig. 1), the aldehyde of vitamin A, as its chromophore [2]. The protein functions as a light driven transmembrane proton pump and is of quite some interest for its potential to harvest light energy. The discovery that bR with its seven transmembrane α-helices has the same spatial arrangement as the large class of biologically important G-protein receptors has further increased interest in this protein. In addition, the protein is easily grown and purified, making it readily available for a wide range of spectral and physiological studies. The photocycle of all-trans bR [WT (wild type)/1a] initiates with the isomerization of all-trans retinal to the 13-cis isomer and consists of a sequence of at least five steps (Fig. 2). The 13-cis pigment [WT/1] is also photochemically active and undergoes a distinct photocycle [3] in which proton translocation is not observed [4]. The photocycle of the all-trans isomer has been studied in the

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Figure 1 Structures of retinal analogues: 1, R 1 ⫽ R2 ⫽ methyl, in retinal; 2, R 1 ⫽ hydrogen, R 2 ⫽ methyl, in 9-demethylretinal; 3, R 1 ⫽ methyl, R 2 ⫽ hydrogen, in 13demethylretinal; 4, R 1 ⫽ R 2 ⫽ hydrogen, in 9,13-demethylretinal.

Figure 2 Photocycle of bR showing the main intermediates. bR, bacteriorhodopsin.

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greatest detail as it is the one associated with the transport of protons. A complication of these photocycles is that the photointermediates themselves undergo photoreversible reactions [5]. The exact role of the chromophore in the proton translocation process is still not completely defined although it is clear that the chromophore in bR and in its various photointermediates is capable of lightinduced isomerization that results in changes in the protein conformation. Little is currently understood of the chromophore–protein interactions inducing these conformational changes. There are two analogue-type approaches that can be taken to probe the structure–function questions regarding this protein: the chromophore can be derivatized or the protein can be mutated (see Fig. 3 for a general scheme). The native chromophore of bR is retinal (1, vitamin A aldehyde), which is a small molecule (molecular weight [MW] ⬍ 300) containing a conjugated polyene system. This native chromophore can be easily replaced with a suitably derivatized retinal [6], and bR has been shown to accept a wide variety of retinal analogues (Fig. 4). The binding site is selective for the polyene chain as pigments cannot be formed with analogues that have significant additional bulk along this chain. However, the portion of the binding site that accommodates the methylated cyclohexyl ring is quite flexible as pigments have been formed with a number of ringderivatized analogues. The approach of replacing the native chromophore with retinal derivatives has been extensively utilized by Nakanishi and his group, both at Columbia and later in his students’ own laboratories [7–9].

Figure 3 Schematic of two approaches to modification of bacteriorhodopsin for structure/function studies.

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Figure 4 Schematic of structural requirements of retinal analogues for pigment formations with bR. bR, bacteriorhodopsin.

9-Demethylretinal (2) has previously been reported to form a bR pigment that has a blue shifted absorption maximum (20 nm) [10,11]. The wild-type pigment containing 2 (WT/2) appeared to function normally with the same proton pumping action as the native pigment. Using a mutant bR in which the tryptophan 182 (Trp 182) was mutated to a phenylalanine, Yamazaki et al. [12] have shown that there is a strong interaction of the 9-methyl group with this residue. Thus, FTIR studies show that the L intermediate is diminished and M formation is delayed. These data indicate that the 9-methyl is located near Trp 182 at this step of the photocycle. A stable bR pigment has also been reported to form with 13-demethylretinal (WT/3). Ga¨rtner et al. [10,13] found that 85% of the chromophore was in the 13-cis conformation in WT/3, in contrast to the pigment with 2, which had an isomeric ratio close to that for the native bR (WT). These investigators concluded that the 13-methyl of retinal is critical for the chromophore isomerization. This analogue therefore provides an excellent tool for the study of systems involving the 13-cis conformation. Proton pumping has been found to be reduced to about 20% that of the native pigment [14]. Removal of methyls from both the 9- and 13-positions generated a pigment that resembled the 2 pigment in absorption properties [11]. Site-directed mutagenesis is a tool that has been extensively explored in the study of bR. Numerous mutants have been made, and the study of the biophysical properties of these mutants has led to much of our current understanding of the

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structure and function of bR [15–17]. The arginine at position-82 (R82) has been found to be key to both the absorption and proton pumping properties of bR. This amino acid has been found to stabilize the ionized state of aspartate at position 85, the proton acceptor and part of the counterion to the Schiff base. The aspartate 85 is deprotonated in purple membrane, but decreasing the pH results in the formation of blue membrane and protonation of D85 [18,19]. Replacing the arginine with the neutral residue alanine (R82A) strongly affects the shift in the pKa of the purple to blue transition, shifting it from 2.6 in WT to 7.2–7.5 in the mutant [20,21]. The result is that the mutant R82A is inactive at pH ⬎ 6. We have combined the use of site-directed mutagenesis and regeneration with retinal derivatives to examine structure/function relationships in the resulting pigments. The chromophore has been altered by deletion of the retinal side chain methyls, one of which is known to affect the isomeric composition of the chromophore in the pigment. The mutant studied is R82A, in which the critical charged amino acid arginine has been mutated to a neutral residue alanine. In each case, the analogue pigment has been compared to its corresponding pigment with either the native chromophore or the native protein.

II. MATERIALS AND METHODS A.

Synthesis of Retinals

The demethyl retinals [22] (all-trans isomers) were synthesized as outlined in Fig. 5. This synthetic route is similar to those used for the syntheses of numerous retinal analogues. Diethyl (3-cyano-2-methylprop-2-enyl)phosphonate was prepared from 1-chloro-3-cyano-2-methylpropene by heating with triethylphosphite. Sodium bis(dimethylsilyl)amide in dry tetrahydrofuran (Aldrich) was used as the base and diisobutylaluminumhydride (DiBal; Aldrich) was used as the reducing agent. The products were purified by TLC (5% ethyl acetate/hexane) and/or HPLC)(Varian 5000 gradient system, Altech Econosphere Silica SU column, 250 ⫻ 6 mm, 0.9% ethylacetate, 0.1% methanol/99% hexane). The retinals were characterized by nuclear magnetic resonance data from a Varian VXR400 spectrometer with deuterated chloroform as solvent. The analogues were used as soon as possible after preparation but, when necessary, were stored at ⫺70°C under argon. The purity was confirmed by HPLC and absorption spectra before use. All experiments with retinals were performed under dim red light. B.

Bacterial Strains and Growth Conditions

Escherichia coli BMH71-18 mutL [23] was used for site-directed mutagenesis, E. coli JM101 was used to propagate mutant phages, and competent E. coli Nova Blue (Novagen, Madison, WI) was used for transformation of the pT7Blue vector

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(Novagen, Madison, WI) containing polymerase chain reaction–(PCR)-generated fragments. All E. coli strains were grown in 2X YT or LB medium [24]. The wild-type and R82A bop genes were expressed in Halobacterium salinarium IV8 [25] (a gift of Richard Needleman), which contains a stable ISH1 insert in the bop gene [26]. C.

Site-Directed Mutagenesis of bR

Oligonucleotide-directed, site-specific mutagenesis was performed essentially as described by Menick [27]. The 2.7-kb BamHI-HindIII bop gene insert was restricted from the H. salinarium shuttle vector pMC-1 (a generous gift of R. Needleman) and ligated into M13mp19. The Arg82 was replaced with Ala by using the mutagenic primer 5′CTGG GCG GCG TAC GCT GAC 3′, which contained the indicated two-nucleotide mismatch. Mutants were initially screened by colony blot hybridization and then plaque purified, and the mutation was verified by dideoxy sequencing [28]. The entire coding region of the bop gene was sequenced to ensure that no additional mutations had occurred. The BamHI-HindIII bop gene insert was restricted from the M13mp19RF and religated into pMC-1. D.

Transformation of Halobacterium salinarium IV-8

The R82A bR mutant and wild-type control were transformed into the H. salinarium strain IV-8 containing ISH1 insert within the bop gene as described [21,29]. The spheroplasts were pelleted by centrifugation, resuspended in growth medium (plus 15% sucrose) [25], and incubated at 37°C for 48 h. Aliquots were plated on growth medium agar plates with 15% sucrose and 5 mg/mL mevinolin (a generous gift of Merck Sharp & Dohme). Several independent clones were picked (after 2 weeks of growth) and struck out on growth medium plates; single colony isolates were grown in 2 L of grown medium to characterize each bR mutant. E.

Southern and Sequence Analysis of the R82A Mutant

Southern blot analysis was carried out as described [24]. Genomic deoxyribonucleic acids (DNAs) from H. salinarium cells IV-8 and IV-8 cells transformed with wild-type or the R82A bop mutant were digested with PstI, transferred to Genescreen Plus membranes, and hybridized with a 32 P-labeled 500-bp KpnI bop gene fragment. Additionally, the R82A bop gene was amplified by polymerase chain reaction from the R82A transformed IV clones. The PCR was performed with 1 mL of sheared H. salinarium genomic DNA as template as previously described [30]. The 1150-bp PCR product was gel purified and sequenced.

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F.

Isolation of Purple Membrane

Purple membrane was also isolated from cultures of H. salinarium by the method of Becker and Cassim [31]. Purple membrane was prepared from the two sources (cultures of H. salinarium IV-8 transformed with wild-type and R82A bop) as described [32] and purified with an additional sucrose gradient (30%–60% sucrose, 100,000 g, 17 h). G.

Pigment Regeneration

The native and transformed pigments were bleached according to the method of Tokunaga et al. [6]. Pigments were regenerated by additions of the all-trans retinals in ethanol (⬍1% vol) to bleached membranes (50 mM Tris-HCl, pH 8.8) at 25°C. Pigment formation was measured after 24 h by absorption changes with bleached membrane as reference. H.

Spectroscopic Methods

Absorption spectra were recorded in 5-mm quartz cuvettes thermostatted at 20°C using a Cary-Aviv 14 spectrophotometer (Aviv Associates, Lakewood, NJ) by digital measurement with 1-nm steps and a spectral bandwidth of 1 nm. Kinetic measurements of dark adaptation were done at 580 nm using the kinetic mode of the spectrophotometer. A homemade cryostat was used for low-temperature measurements [33]. Flash-induced absorption changes were measured on a kinetic spectrophotometer [34,35]. The actinic pulse was from a Quanta Ray DCR11 Nd:YAG laser (532 nm, 7 ns, 5–10 mJ/pulse; Spectra Physics, Mountain View, CA). Flashinduced pH changes were measured at pH 7–7.2 using the pH-sensitive dye pyranine.

III. RESULTS AND DISCUSSION The retinal analogues were synthesized as outlined in Fig. 5. This scheme is quite similar to those used in other laboratories for the syntheses of various retinal analogues and relies primarily on Horner-Emmons olefination for chain elongation. Yields of the various steps were between 40% and 65%, but the products were seldom isolated other than for an initial characterization, because of the lability of these polyene compounds. Stable pigments were formed between bacteriorhodopsin and the trans isomer of all three retinal analogues 2–4. The absorption maxima of these WT pigments were similar to those reported by Ga¨rtner et al. [10] (Table 1). Pigments

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Figure 5 Synthesis of retinal analogues. Route 1, synthesis of 9-demethylretinal (2); route 2, synthesis of 13-demethylretinal (3); route 3, synthesis of 9,13-demethylretinal (4).

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Table 1 Absorption of Pigment Analogues a Absorption maximum (nm)

Native WT Transformed WT R82A a

Retinal (1)

2

3

4

558 558 555

540 540 537

569 567 566

539 538 539

All pigments were regenerated with the all-trans isomer of the respective retinal. Pigment solutions were dark adapted for 1 h before measurement. Spectra were recorded in 50-mM-Tris-HCl, pH 8.8 at 24°C. Values ⫾2 nm. WT, wild type.

were formed with both bR apoprotein generated from the normal purple membrane purified from cultures of the halobacteria and from protein of H. salinarium IV-8 transformed with the WT bop gene. The transformed and native pigments had identical absorption properties. The data presented here are from pigments regenerated with the apoprotein isolated from the native pigment. The pigments formed with the apoprotein of R82A and the three demethyl retinals showed absorption maxima at 537 nm (2), 566 nm (3), and 539 nm (4), respectively (Table 1). The pigments were reasonably stable in the presence of 10-mM hydroxylamine, pH 7.5, with only modest degradation noted for the pigments formed with 4 (⬍25% over 2 h). The deletion of the 9-methyl group shifted the absorption maxima in both of the pigments, whereas the removal of the 13-methyl and the mutation at the 82-position had little effect on the absorption properties. The R82A pigment has a pK a for the purple to blue transition of 8.7 in 25% glycerol/water (Fig. 6). For the R82A/3, the pK a was shifted to 8.1. The origin of this shift can be attributed to the finding that for the 3 pigment most of the chromophore is probably in the 13-cis conformation and the pK a of the 13-cis form of the pigment is lower than that for the all-trans form [36]. The photocycle intermediates were detected by monitoring the flash-induced absorption changes at 410 nm, 540 nm, and 600 nm. The WT changes (Fig. 7a) represent predominantly M (410 nm) and O (640 nm) intermediates as well as the recovery of the pigment (540 nm). The WT/2 bR converted to a 410nm M intermediate and a 600-nm species, as a result of the O intermediate, after 20 ms (Fig. 7c). The formation of the M intermediate is in agreement with the results of Yamazaki et al. [12]. Excitation of WT/3 produced only a small amount of the 410-nm M intermediate species (Fig. 7e). The half-time of decay observed for this species was about 3 s, whereas the decay of M in the WT bR is 10 ms. The predominant conversion of WT/3 was to a 600-nm species that is much less in WT/2 (Fig. 7c). It has been proposed that this 600-nm species, which is different from the

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Figure 6 Fraction of blue membrane as a function of pH in dark-adapted R82A (2) and R82A/3 (1).

O intermediate of the trans cycle, arises from the 13-cis cycle. [7,12] This identification of the photoproduct and a small amount of M is consistent with the fact that the 13-cis isomer accounts for at least 80% of the retinal in the binding site of 3 pigment. Furthermore, upon the illumination of WT/4 pigment, the M intermediate species from the trans cycle was quite small and only an unusual, long-lived 600-nm species was found (Fig. 7g). These results of flash-induced spectroscopic experiments are consistent with the conclusion that the 13-position of retinal is important for the stability of the all-trans configuration of retinal in the binding site [7]. In R82A, the light-induced absorption change at 410 nm decayed much faster than the absorption at 580 nm recovered, and an O intermediate was not observed (Fig. 7b). Interestingly, a rapidly forming 600-nm species is observed in all three R82A/2–4 pigments (Fig. 7d, f, and h), indicating an increase in the proportion of the 13-cis isomer in the pigment. The R82A/3 and R82A/4 pigments have particularly long-lived 600-nm species. The flash-induced proton changes were measured by the transient absorption changes of the pH indicator pyranine at pH 7.2. The fast (⬍1 ms) decrease in dye absorbance observed in WT corresponds to release of protons, and the subsequent restoration of absorbance (rise time 12 ms) corresponds to the uptake of protons (Fig. 8a) [11]. The signals from all three WT/2–4 pigments were ⬍20% that of WT (Fig. 8b, c, and d). The proton signal from R82A was about 40% that of WT, and the order of proton release and uptake is reversed

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Figure 7 Flash-induced absorption changes: a, WT; b, R82A; c, WT/2; d, R82A/2; e, WT/3; f, R82A/3; g, WT/4; h, R82A/4. Measuring wavelengths as marked, λ actinic 532 nm, temperature 20°C, 0.1 M NaCl, pH 7.

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Figure 8 Flash-induced absorption changes of the pH-sensitive dye pyranine showing proton release/uptake: a, WT; b, WT/2; c, WT/3; d, WT/4; e, R82A; f, R82A/4. Measuring wavelengths at 458 nm with other conditions as in Fig. 4.

compared to that of the WT, as observed previously (Fig. 8e) [11]. No absorption change of the dye in R82A/3 was observed (Fig. 8f), demonstrating that the combination of the deletion of the 13-methyl and the removal of the charged amino acid at position-82 prevents the light-induced movement of protons for this pigment. IV.

CONCLUSIONS

In conclusion, the use of analogues of retinal lacking the 9- and/or 13-methyl in combination with the R82A mutation has allowed us to examine the photocycle

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and function of pigments in which the 13-cis to all-trans isomeric ratio is expected to be increased. The 9-methyl is critical to maintaining the absorption maximum of bR, and the environment of interaction that causes this shift has not been altered in this mutant. The photocycles of the pigments are affected in that the R82A/2 shows an inhibition of intermediate M formation, and all the R82A/2– 4 pigments show the 600-nm species. Proton pumping is eliminated in the R82A/ 3 pigment. These results support previous findings on the importance of the Arg82 residue to the function of bR and illustrate the value of the combination of the two analogue approaches in probing the complicated chromophore–protein interactions in the proton pumping process.

ACKNOWLEDGMENTS This work was supported by NIH (EY04939 and GM52023), DOE (95ER20171), and Research to Prevent Blindness, Inc. We thank Dr. R. Needleman for providing the shuttle vector containing the bop gene, pMC-1, and bR-defective H. salinarium strain, IV-8. We particularly thank Dr. Koji Nakanishi, without whose initial guidance and enthusiasm this collaboration would not have existed, and therefore these studies would not have been accomplished.

REFERENCES 1. D. Oesterhelt and W. Stoeckenius, Nature, 233NB: 149 (1971). 2. For recent reviews, (a) T. G. Ebrey, Thermodynamics of Membrane Receptors and Channels; (M. Jackson, ed.), CRC Press, Boca Raton, Fla. pp 353–386. (1993). (b) J. K. Lanyi and G. Varo, Israel J. Chem., 35: 365 (1996). 3. C. Gergely, C. Ganea and G. Varo, Biophys. J., 67: 855 (1994). 4. A. Fahr and E. Bamberg, FEBS Lett., 140: 251 (1982). 5. S. P. Balashov, Isr. J. Chem., 35: 429 (1996). 6. F. Tokunaga, R. Govindjee, T. G. Ebrey, and R. K. Crouch, Biophys. J., 19: 191 (1977). 7. W. Humphrey, I. Logunov, K. Schulten, and M. Sheves, Biochemistry, 33: 3668 (1994). 8. Y. Feng, D. R. Menick, B. M. Katz, C. J. Beischel, E. S. Hazard, S. Misra, T. G. Ebrey and R. K. Crouch, Biochemistry, 33: 11624 (1994). 9. For a recent review: K. Nakanishi and R. K. Crouch, Isr. J. Chem., 35: 253 (1996). 10. W. Ga¨rtner, P. Towner, H. Hopf and D. Oesterhelt, Biochemistry, 22: 2637 (1983). 11. M. M. Szweykowska, A. D. Broek, J. Lugtenburg, R. L. van der Bend and P. W. M. van Dijek, Recl. Trav. Chim. Pays-Bas. 102: 42 (1983). 12. Y. Yamazaki, J. Sasaki, M. Hatanaka, H. Kandori, A. Maeda, R. Needleman, Shinada, K. Yoshihara, L. S. Brown, and J. K. Lanyi, Biochemistry, 34: 577 (1995). 13. H.-W. Trissl and W. Ga¨rtner, Biochemistry, 26: 751 (1987).

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16 Autonomous Genomes David G. Lynn The University of Chicago, Chicago, Illinois

I. INTRODUCTION In 1995, the first complete genomic sequence was published [1], and since then several others have become available. Of particular importance to the bioorganic chemist, this list includes Escherichia coli [2] and Bacillus subtilis [3], the organisms whose proteins and molecular genetics have been used so extensively to define and characterize the chemical reactions of biology. These primary sequence data will certainly extend and enrich our understanding of the reactions that drive these organisms, but it also opens an entirely new set of questions. It is likely that no scientific discipline will benefit more directly from this knowledge than bioorganic chemistry and chemical biology. Just as the structure and the synthesis of the steroid nucleus placed human reproduction and hormone physiological processes at the atomic level, and the structure of proteins revealed the exquisite sophistication of the mechanisms of biological catalysts, the structure of the genome, the molecule that codes for the entire organism, places the most central questions of biology on an atomic scale. More than just understanding the molecular machines of biology, it is now possible to consider how chromosomes function: to wonder about the logic for the organization and positions of genes that are functionally interdependent, the function of intervening sequences that tie the coding regions together, the reason for the positioning of the origin and terminus of replication, the functional significance of G-C-rich domains; the positions, frequencies, and function of repeating sequences; and the organization and utility of coding redundancy. With data on multiple genomes, strategies for organization and stability can be compared and the rules governing success may be understood. Questions as to how

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genome organization differs across a genus, a family, a kingdom; how an entire genome responds to major environmental changes; how stages and specific rates of molecular evolution occur across the genome can be experimentally addressed. The genome is a far larger structure than a steroid or an enzyme, and different methods are needed for its analysis. In this chapter I would like to consider several approaches that we have explored to understand genome function. These approaches will be presented in the context of the minimal structure for an autonomous genome [4–9]. The small genomes were the first to be sequenced, and these are already providing new glimpses of theoretical evolutionary genomics and the rooting of evolutionary trees [6–9]. But the term autonomous is slippery in a biological context. For example, Morowitz [4] suggested that the smallest cell is likely to contain the minimal genome and, making estimates of average macromolecule volume, proposed that Mycoplasma genitalium, with a 0.3 mmdiameter, could encode only 600 proteins. This genome sequence was published in 1995 and contains 469 genes [10], putting a lower limit on the number of catalysts needed for an autonomous cellular entity. But M. genitalium is a parasite and contains, for example, very few genes necessary for the synthesis of its amino acids and nucleic acid bases. Its autonomy, then, depends very much on the environment in which the genome functions. Once the environment becomes equivocal, then a virus, a plasmid, or a viroid becomes an autonomous genome within the host cell. At the molecular level, then, the minimal structure for genome function is context-dependent and is a chemical issue. Although it is within biology that the term genome has evolved and must be understood, it is within the limits of the chemical environment that the reactions necessary for autonomy must be defined. Much insight will be gained over the next few years as genome sequences are compared, but just as with the steroid nucleus, it is within the context of the construction of new structures that build on the operational principles uncovered within these genomes that we will be able to advance beyond what exists today and to explore the limits of what is autonomous and what is chemically possible.

II. TOP-DOWN GENOME REDUCTION Maniloff [6] argued that small autonomous genomes could have arisen along two independent paths, which he referred to as ‘‘top-down’’ and ‘‘bottom-up.’’ The bottom-up path is essentially an origin of life argument, to which I will return. The top-down path requires the evolutionary development of symbiotic, parasitic, or other interdependent associations such that genomic information can be reduced. The biosphere today is a tightly woven network of interdependent organisms. Plants can utilize CO 2, O 2, and NO 2, inorganic materials, and photons for energy, seemingly functioning in the absence of other organisms. There are many

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associations, cellular organelles, micorrhizal/plant symbioses, lichens, legume/ rhizobium symbioses, insect vectored pollination strategies, etc., in which interdependence appears to have evolved. In many of these systems there is an obvious driver for the association, but the actual evolutionary pathway and the molecular and mechanistic changes that allowed for its development are less clear. The parasitic angiosperms provide a situation that may allow for a molecular level description of the traits that led to this loss of genome autonomy and of the resulting evolutionary consequences. Searcy and MacInnis proposed three general developmental phases for the molecular evolution of the parasitic plants [11,12]. The first consisted of the development of the specialized organ that forms the physiological bridge to the host—the haustorium. Once formed, the subsequent events are of specialization and adaptation. These events include the loss of biochemical pathways and morphological structures that become redundant with host attachment, and the accrual of more complex adaptations specific to an obligate parasite, such as host specialization and mechanisms to overcome host defenses. Analyses of the nonphotosynthetic holoparasite Epifagus virginiana have provided striking support for evolutionary reduction [13,14]. The plastid deoxyribonucleic acid (DNA) (ptDNA) of this organism was found to be approximately a third the size of a nonparasitic relative, Nicotiana tabacum, and essentially all of the plastid encoded photosynthetic genes, the reduced nicotinamide-adenine dinucleotide phosphate (NADPH) dehydrogenase genes, all four ribonucleic acid (RNA) polymerase genes, 13 of the 30 plastid encoded tRNAs, and 6 of the 22 plastic encoded ribosomal protein genes were either missing completely or present as pseudogenes [15,16]. Accelerated rates of molecular evolution of both the remaining plastid genes and the rDNA of these parasites have also been documented and explained by molecular population genetic models of mutation, selection, and drift [17,18]. Significant biochemical evidence also exists for highly specialized processes for host selection. A component exuded from the roots of sorghum seedlings was shown to induce Striga asiatica seed germination and was the first characterized host-derived stimulant, the sorghum xenognosin for Striga spp. germination (SXSg) [19,20]. This component, however, was not sufficient to explain the distance regulation observed for S. asiatica seed germination in culture, and mathematical models of host commitment uncovered an additional component, which ironically proved to be produced via the same biosynthetic pathway as SXSg [21,22]. Several other examples of host specialization have been documented [23], and, more recently, mechanisms for overcoming host resistance have been detected [24–27]. By Searcy’s model, the development of the host attachment organ was the critical first step leading to the subsequent genomic adaptations. Current estimates are that 1% of all flowering plants are parasitic [13,28], broadly distributed across

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at least 16 families [29], suggesting that the parasitic strategy, driven by haustorial development, was repeatedly discovered throughout angiosperm evolution [23]. The identification of 2,6-dimethoxy-p-benzoquinone (DMBQ) as a natural sorghum xenognosin for Striga haustoria (SXSh) that was both necessary and sufficient for haustorial induction in several of the parasitic Scrophulariaceae [20,30] suggested that oxidative release from host cell walls provided an important signaling strategy [31]. It has now been shown that the S. asiatica root meristem constitutively produces H 2O 2 (Fig. 1). This oxidant has been hypothesized to serve as a cosubstrate, along with host cell wall phenols, for apoplastic localized host peroxidases to generate the xenognostic signals [32]. The released quinones mediate host cell surface detection (Fig. 2). This strategy would provide an effective means for the parasite to identify a viable host and to respond in a distancedependent manner to the host root surface [33]. H 2O 2 has been shown to be produced at the cell wall during pathogen invasion [34], and two pathways are known for its production. Apoplastic peroxidases [35,36] generate H 2O 2 via an H 2O 2-independent oxidation of a number of substrates, including NADH, NADPH, thiols, and certain phenols [36–40]. At pres-

Figure 1 Light micrograph of a 2-day-old Striga asiatica seedling that has been stained with pyrogallol. The red coloration [shown here as a darkening at the root meristem (see arrow)], of the root meristem highlights the H 2O 2 production. (Source: Ref. 32.)

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Figure 2 The proposed mechanism for haustorial induction involves a four-step autocatalytic cycle: (a) H 2O 2 is produced at the parasite root meristem and diffuses to the host, (b) the H 2O 2 serves as a substrate for host apoplastic peroxidases that oxidatively remove wall bound phenolics at kpx[H 2O 2][PhOH], (c) some percentage of these phenols are oxidized to quinones that diffuse to the parasite, and (d) a parasite oxidoreductase uses the quinones to produce H 2O 2 at k et[Q]. The parasite oxidoreductase is coupled to a signal transduction chain that induces haustorial development.

ent there is no evidence for the exudation of these more easily oxidized substrates to the parasite cell surface. Evidence for a membrane-bound NADPH oxidase complex, homologous to the human neutrophil oxidative burst machinery, has also been found in plants [41–44]. In that regard, SXSh quinones were recently shown to be detected via a benzoquinone-dependent oxidoreductase [45,46]. In terms of the observed redox range (Fig. 3), the terminal acceptor could be O 2, via a cytochrome-mediated reduction. It may be, therefore, that signal detection is coupled with oxidative signal release through the production of H 2O 2 in an autocatalytic host detection mechanism. It has been suggested that a mutation in the oxidative burst machinery controlling H 2O 2 production in a dicotyledonous plant has made it possible for the parasite to exploit monocot cell surfaces via a dicot defense pathway. A good defense is converted into an offensive host

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Figure 3 The active haustorial inducers fall within a narrow first half-wave reductive potential window, between 20 and ⫺280 mV, consistent with their role as redox carriers. (Source: Ref. 46.)

detection strategy [32]. Such a gain of function mutation could have easily evolved and played a critical role in the development of parasitism. Further analysis of the oxidoreductase genes responsible for H 2O 2 production in Striga should allow the model to be evaluated. The study of Striga spp. makes possible a structure/activity analysis of the parasitic genome. By defining the elements that lead to parasitism, a functional understanding for the process of genome reduction can be obtained. Through comparisons with other related parasites, both the rate of gene loss and the resulting changes in genomic organization may be understood. Analogous ‘‘compare and contrast’’ approaches have taught us much about the forces that drive biology, and it is clear that these analyses have now moved to an atomic level. In our case, the genomic reorganization that occurs between parasitic and nonparasitic relatives provides glimpses into the structure/function analysis of their genome.

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III. BOTTOM-UP GENOME CONSTRUCTION The soil bacterium Agrobacterium tumefaciens can harbor an extrachromosomal genome, the Ti plasmid, which is responsible for coupling the bacterium and a plant cell into a disease state known as the crown gall tumor. An elaborate metabolic interface is developed with the bacterium, which results in the oncogenic transformation of a plant cell host [47–49]. Together the two organisms are coordinated to ensure the survival of the Ti plasmid. Fig. 4, presented as a timeline, outlines the signaling events that coordinate this interface. In the left panel, the first signal is generated from a dividing compe-

Figure 4 The timeline for the infection process of Agrobacterium tumefaciens. Across the top, the oncogenetically transformed plant cell displays tumorigenic growth. In the left-hand panel, the gene transfer event is induced by specific signaling molecules (represented by phenol) that originate from the plant cell. The T-DNA is transferred after activation of the vir regulon of the Ti plasmid (pTiA6). In the middle panel, octopine is produced by the transformed plant cell and activates both occ and traR expression. In the righthand panel, CF, a factor whose biosynthetic genes are constitutively expressed from Ti, together with TraR, activates conjugal transfer of the Ti plasmid.

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Figure 5 Comparisons of the two gene transfer events in Agrobacterium spp. The oncogenetic transfer (a) integrates multiple inputs, of which the phenol signal is most critical as the initiation event. Conjugal transfer (b) shows an autocatalytic amplification step to induce the machinery for gene transfer. The genetically encoded machinery for both transfer processes is homologous.

tent plant cell and recognized via the activation of the virulence regulon of the Ti plasmid. A histidine two-component sensor kinase and response regulator are constitutively expressed from the regulon and integrate information from monosaccharide, phenol, and acidic pH to control expression of the remaining genes (Fig. 5). The virulence regulon encodes the machinery that catalyzes the excision of the T-DNA, its transfer into the plant cell, and targeting aspects of its incorporation into the genome of the plant cell. The center panel shows the result of incorporation of the T-DNA. Driving the prokaryotic genes for critical steps in the biosynthesis of auxin and cytokinin behind eukaryotic promoters leads to the uncontrolled growth of the plant cell. In addition to the oncogenes, an expressed dehydrogenase catalyzes the reductive amination of pyruvate with arginine to generate octopine, a substance not metabolized by the plant, but that can serve as the sole carbon and nitrogen source for A. tumefaciens that harbor the catabolic genes of the Ti plasmid. Octopine accumulation activates both the expression of the octopine catabolism regulon (occ) and the regulatory protein, TraR. TraR and a homoserine lactone adduct autoinducer, AI, constitutively biosynthesized from

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S-adenosyl methionine and the acyl-CoA keto ester by gene products of the Ti [50], are required to activate the expression of the tra regulon [51]. This regulon controls transfer of the entire Ti plasmid, as shown in the right-hand panel via an autocatalytic amplification of the machinery for conjugal transfer (Fig. 5). Given the homology that exists between the two gene transfer machines shown in Fig. 5, the process has little specificity with respect to recipient cell and transfers genes to both prokaryotic and eukaryotic recipients. Thus, the Ti plasmid, the original tumor inducing principle of Armin Braun, successfully orchestrates an interface between eukaryotic and prokaryotic cells and, once the economic advantage of opine production is engineered, delivers itself to any cell capable of exploiting the advantage. The only requirements for the host cell are the replication and expression of Ti. Ti is therefore an autonomous genome, but its autonomy is context-dependent. The eukaryotic host plant range is very broad, virtually for any dicotyledonous plant, and as such A. tumefaciens provides a general vector for higher plant transformation [47]. The bacterial host, however, is restricted to A. tumefaciens. In a bottom-up approach, Ti can be explored as a scaffold on which increased capability can be added to make a more autonomous genome. Bacterial host range competence may be limited by several factors, but a central one involves the initial step of establishing the interface with the plant, activation of the VirA/VirG signal transduction chain. Two different experimental approaches have suggested that VirA and VirG alone are not sufficient for signal transmission. First, analysis of the known inducing structures led to a proposed mechanistic-model for induction [52]. From this model a series of specific irreversible inhibitors were prepared and developed into affinity reagents. These affinity reagents did not label VirA, but did label a series of low-molecular-weight chromosomally encoded proteins in vivo within a time frame consistent with the physiological inhibition. Therefore, VirA was proposed not to be the xenognostic receptor [53,54]. More recently, A. tumefaciens mutants have been selected for increased sensitivity to the signal molecule. These mutants show an enhanced response to specific methoxy geometric isomers of the designed inducers. This trait has been localized to the chromosomal background [55]. The identification of chromosomal loci that are critical for signal perception suggests that A. tumefaciens maintains some parental control over the promiscuous Ti plasmid. Defining those loci should provide an understanding of the origin of this apparent control and further raise the possibility of cloning these genes back into Ti to extend its bacterial host range. Plasmids represent mobile elements whose molecular genetic manipulation is now highly developed, and methods are available to extend their capability. They were among the first genetic elements to be synthetically engineered and that synthetic potential is still growing. The advantage with Ti is the great flexibility of the DNA transfer complexes, and increased replication autonomy repre-

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sents a biotechnological potential that will allow for the molecular entraining of other organisms. Such synthetic approaches, which share a conceptual link with the synthesis of alternate functional groups on the steroid nucleus, should ultimately extend to the construction of larger increasingly autonomous genomes.

IV.

EXTENDING THE PRINCIPLES TO OTHER SKELETONS

As the requirements of a functioning genome are understood, it becomes possible to develop rationally, as has been the case with steroid hormone action, different architectures capable of genome function. Conceptually, the storage and transfer of molecular information, copying, reading, and translation, are all template-directed polymerization reactions. In biological systems these reactions are mediated by a complex array of catalysts, and kinetic control of both the rate and the fidelity of the information transfer is critical. However, in a primordial bottomup approach to genome construction, species capable of such catalysis would not be present and must have evolved from thermodynamically controlled selfassociations. Several model templates have shown features of self-replication, including nucleotide-based dimers and oligomers [56–64], peptides [65,66], and micelles [67]. The goal of many of these studies has been to demonstrate exponential growth of the encoded information, and this effort has seen limited success because template turnover in replication is limited by product dissociation. In existing biological systems separate catalysts evolved that use chemical energy to drive template dissociation prior to subsequent rounds of replication and, in this way, achieve exponential growth. The evolution of catalysts is therefore central, and in considering possible biopolymer catalysts, both DNA and RNA are candidates [68–73]. The range of reactions catalyzed by the nucleic acid polymers is however, more limited than that of proteins for at least two reasons. First, the narrower range of side chain functional groups and the more specific associations of the existing side chains in Watson–Crick and Hoogsteen pairing motifs limit their catalytic availability. Second, the hydration requirements of the polyanionic phosphate backbone reduce its ability to pack into rigid structures. Hence, binding site definition, substrate binding and release, as well as large-scale and rapid conformational rearrangements important for allostery in proteins are more limited in nucleic acids [74]. The thermodynamic stability of Watson–Crick duplexes is significant, and we have harnessed this energy to drive reversible imine condensation [75,76]. Under reducing conditions, the imine is trapped as the amine, and the base sequence information encoded within a DNA heximer is irreversibly transferred with good fidelity (Fig. 6). By defining the thermodynamic stability of the amine and imine linkage [77,78], conditions were developed in which product inhibition

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Figure 6 The thermodynamic and catalytic cycles for translating DNA information into the altered backbone product. The substrates are labeled with 15 N to allow the assignment of the equilibrium constants, K, for each step. DNA, deoxyribonucleic acid. (Source: Ref. 79.)

of template turnover was reduced [79] such that the template efficiently catalyzed the amplification of the encoded information. This opportunity to have multiple turnovers on the template allows for the information to be transferred accurately and amplified many times with catalytic amounts of template. This ligation reaction gives a different product backbone from the template and, in that way, has characteristics of a translation process. To extend this process to replication would require the amine product to catalyze the construction of the template as shown in Fig. 7. The ethyl amine linkage is more hydrophobic than the phosphate diester, and its reduced hydration and increased flexibility

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Figure 7 Proposed two-step replication process in which the phosphate backbone polymer catalyzes the synthesis of the amine backbone and the amine backbone polymer catalyzes the synthesis of the phosphate backbone.

should enable polymers containing such linkages to fold and pack in a way that is more like that of proteins. The basic amine in the linkage should expand the catalytic competence of the polymer much as the 2′-OH does in RNA [68,74]. Therefore it may be possible to reduce the central dogma of biology, DNA to RNA to protein, to a two-step process as shown in Fig. 7. The ligation efficiency of the reaction outlined in Fig. 6 has allowed us to consider new approaches to template-directed synthesis, even a synthetic approach that extends the architectural limits of the bottom-up construction of a genome. Now the term autonomy becomes more relevant because this genome would be synthetic. The extension of the ligation reaction to template-directed polymerizations would be necessary, and eventually, for the template to be autonomous, it must encode the capability of catalyzing a ‘‘metabolism’’ sufficient for monomer generation [80–82]. Therefore, the central question becomes how the required diversity of catalysts might be produced from the template. The principles inherent in the imine coupling reactions can be extended to other coupling reactions, giving different functionality in the backbone of the translation product. For example, DNA has recently been shown to direct the condensation of peptide nucleic acid amides under dehydrating conditions [83,84]. These products have new backbones and represent new translation products that could be selected for new catalytic function. In other words, one template could be translated into many different products in which the base sequence is the same, but the different

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backbone increases the diversity of catalytic function. In the future, the definition of autonomy will be solely in the hands of the chemist who defines the environment, i.e., the access to raw materials and the energy source.

V.

CONCLUSIONS

Chemistry is as much a philosophical approach to problem solving as it is a scientific discipline. As our understanding of biology increasingly approaches an atomic scale, chemistry’s problem solving approaches become more generally applicable. Clearly genome structure has radically increased the convergence of chemistry and biology, and it is the contributions of the fathers of bioorganic chemistry, including Koji Nakanishi, to whom this volume is dedicated, who have developed the methods and the ideas that have made this convergence possible. At the center of chemistry is the ability to create new structures synthetically and change composition in a rational, definable way. As a result of the phenomenal synthetic advances that have occurred, notably since the 1960s, the current methodology boasts incredible diversity. The physical laws that drive life must operate within the structural limits of molecular frameworks and environmental conditions, and, as I have tried to develop here, this synthetic flexibility will allow those limits to be expanded continually.

ACKNOWLEDGMENTS The ideas presented here have grown from discussions and experiments with very talented coworkers, most of whom are referenced here. I am grateful for a longstanding and valuable collaboration with Andy Binns, University of Pennsylvania, and support from the NIH, NSF, DOE, Novartis, and the Rockefeller Foundation. Most specifically, I acknowledge Koji Nakanishi for establishing a breadth of inquiry and an enthusiasm for learning that have so broadly impacted science, and for the personal support that has made these investigations possible.

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17 Stereochemical Considerations of Immunoglobulin Heavy Chain Enhancer Activation Barbara S. Nikolajczyk* and Ranjan Sen Brandeis University, Waltham, Massachusetts

I. INTRODUCTION Immunoglobulin secreting cells, or B cells, differentiate in the bone marrow from a common blood cell precursor. This developmental process requires changes in gene expression, one example of which is the B cell–specific expression of the immunoglobulin µ heavy chain gene (IgH). We have focused on dissecting the details of transcriptional activation of the IgH gene to gain insight into the mechanisms responsible for B cell development. B lymphocyte–specific expression of the IgH gene is activated by the µ enhancer, a deoxyribonucleic acid (DNA) regulatory sequence located in the J HC µ intron. The µ enhancer is activated at early stages of B cell differentiation and participates in targeting the IgH locus for V(D)J recombination close to the time of B lineage precursor cell commitment [1,2]. The µ enhancer is hypothesized to increase accessibility of the recombinase machinery to the µ locus during V(D)J recombination. Later in B cell differentiation, after recombination has positioned a V H promoter close to the C µ locus, the enhancer is required for expression of the functionally rearranged µ gene as demonstrated by numerous transfection and transgenic assays. Furthermore, in transgenic assays, the µ enhancer is sufficient to activate a reporter gene in lymphocytes, but not in other hematopoi-

* Current affiliation: Boston University Medical School, Boston, Massachusetts.

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etic cell types, indicating its autonomous cell-type specificity [3–5]. Our objective is to understand the molecular mechanisms of µ enhancer function. We have previously defined a core region of the µ enhancer that is active in B cells [6]. This region contains three elements—µA, µE3, and µB—all of which are simultaneously required for transcriptional activation. The µA and µB elements bind ETS domain transcription factors; µE3 binds several widely expressed leucine zipper-containing basic helix–loop–helix proteins (bHLH-zip). The ETS domain proteins are a family of DNA binding factors that have amino acid homology in the DNA binding domain, the ETS domain. Similarly, bHLHzip transcription factors are homologous in the bHLH-zip domains, which are responsible for the dimerization and DNA-binding properties of this protein family. Several ETS proteins can bind µA [6–9], and the µB site binds PU.1, an ETS protein that is highly expressed in B cells and macrophages [10]. None of the ets or bHLH-zip family members relevant to µ enhancer activation are restricted to pre-B and B cells where IgH genes are expressed. The lack of correlation between the tissue distribution of enhancer binding proteins and cells in which the enhancer is active led us to propose that tissue specificity of the enhancer is achieved by a combinatorial mechanism [6]. Specifically, we proposed that µ enhancer activation is B cell–specific because the appropriate combination of transcription factors is present only in B cells and this combination must bind enhancer DNA in the appropriate three-dimensional arrangement to form a transcriptionally active nucleoprotein complex. Consistent with this idea, multimerization of µA, µE3, or µB does not reconstitute a regulatory element that activates transcription in B cells [11–13]. These observations suggest that the properties of this tripartite enhancer domain are determined by the appropriate juxtaposition of the three sites.

II. A PROPOSED MECHANISM FOR ␮ ENHANCER ACTIVATION IN B CELLS A.

␮A–␮B Spacing Is Critical for Enhancer Activity in B Cells

Our previous analysis demonstrated that the µA and µB sites are essential for activation of the murine µ heavy chain gene enhancer [6]. Examination of the corresponding regions of the murine and human IgH loci showed that both µA and µB sites were highly conserved (Fig. 1). Interestingly, the spacing between the µA and µB motifs is evolutionarily conserved; however, we found that the µE3 site defined in the murine enhancer was absent in the human enhancer. Preservation of µA–µB spacing even in the absence of a recognizable E box suggested that correct spatial orientation of these two sites was critical for enhancer function.

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Figure 1 Comparison of murine and human µ enhancer sequences. Note high conservation of µA and µB and lack of µE3 conservation. Spacing between µA and µB is identical.

To test the hypothesis that the organization of the µA and µB sites was critical for enhancer activity in B cells, we constructed mutated µ enhancers with either 4 or 10 base pairs inserted between the µE3 and µB sites (Fig. 2) and tested these mutations in DNA binding and functional assays. Insertion of 4 or 10 bp rotates the µB site approximately 145° or 350°, respectively, relative to the µA or µE3 sites, assuming a typical DNA helical periodicity of 10.5 bp. Protein binding to the three sites was not affected in the rotation mutations. However, transcription activation in B cells was significantly impaired in both the ⫹4

Figure 2 Precise alignment of µA and µB is necessary for µ enhancer function in B cells. Insertion of one-half ([⫹4] construct) or one full ([⫹10] construct) helical turn of DNA between µA and µB inactivates the enhancer in B cells, despite restoration of helical alignment in the [⫹10] construct. Activity of the control [⫹18] construct excludes trivial effects due to changing enhancer–promoter distance.

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and ⫹10 mutations (Fig. 2). These observations suggest that appropriate spacing between the µA and µB sites is critical for enhancer activity. B.

Comparison of Orientation Mutated Enhancers in B Cells and Macrophages

The µA and µB sites that bind ETS domain proteins are nonpalindromic. This ‘‘directionality’’ is depicted in Figs. 2 and 3 as single-headed arrows. In contrast, the intervening µE3 element is palindromic and therefore shown as a doubleheaded arrow. To investigate further the spatial constraints of a functional µA/ µB motif, we tested enhancers in which the relative orientations of the µA and µB sites had been altered. Mutated enhancers F2-5, as indicated in Fig. 3, contained changes in the relative orientation of the µA, µE3, and µB sites alone or in combination. In the wild-type enhancer (F1), the core GGAA of the µA site is 11 bp 5′ to the core CATGTG of µE3, and the core GGAA of the µB site is 19 bp 3′ to the µE3 core binding site (spacing counts are based on centers of µA

Figure 3 Precise alignment of µA and µB is necessary for µ enhancer activity in B cells, but not macrophages. Any change in µA–µB orientation (F2–F4) inactivates the enhancer in B cells. The macrophage µ enhancer nucleoprotein complex can retain function despite changes in µA–µB orientation (F2–F3). Therefore, a given array of ets sites can define a tissue-specific enhancer activated by the combination of DNA-binding proteins present in a particular cell type.

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and µB that are between the G and A of the GGAA cores, and the center of µE3 between the middle TG). Enhancer F2 contains a 14-bp spacing between the µE3 and µB cores, and the µB GGAA is located on the noncoding DNA strand, whereas F3 contains a 19-bp spacing between the µE3 and µB cores, and µB is again located on the noncoding DNA strand. In F4, µA is moved to the noncoding DNA strand and the distance between the core GGAA of µA and the core CATGTG of µE3 is conserved at 11 bp, resulting in the helical realignment of µA relative to µB. In F5 we moved the µE3 site further 3′ by flipping the DNA between µA and µB. Because µE3 is partially palindromic, inversion of the site should not affect the stereochemistry of protein binding to DNA. Enhancer F5 places the µA and µE3 cores 14 bp apart and the µE3 and µB cores 16 bp apart. In vitro binding analyses with mutated enhancers and purified proteins indicated that the mutations had not inadvertently affected protein binding. To test the hypothesis of B cell–specific combinatorial activation of the µ enhancer directly, we tested activity of F1–F5 in the hematopoietic cell types B cells and macrophages. The wild-type enhancer F1 was active in both S194 B cells and RAW 264.7 macrophages (Fig. 3). Functional tests of F2–F5 probed the mechanism that activates the wild-type enhancer in the two cell types. A change in the orientation of µA (e.g., F4) or µB (e.g., F2, F3, F4) inactivated the enhancer in B cells. However, when the orientation and spacing between µA and µB were maintained, the enhancer was active in B cells even if the spacing between the µE3 and µA or µB sites was altered (F5). In contrast, the F2 and F3 constructs, in which the orientation of the µB site is changed, were active in RAW 264.7 cells (Fig. 3). Enhancer F4, in which the µA–µE3 distance and relative orientations of µA and µB are altered, was inactive in macrophages. The F5 enhancer was about 50% active in macrophages, indicating that the moderate modification in µE3 alignment relative to µA and µB was not critical for enhancer activity in the context of appropriately arranged µA and µB sites. Overall, we conclude that proper juxtaposition of µA and µB is critical for µ enhancer activation in B cell lines, but not in macrophages. The results of the orientation mutated enhancers strongly suggest that the mechanism underlying activation of the minimal µ enhancer differs in B cells versus macrophage. These observations highlight the tissue-specific regulatory possibilities that can be achieved by appropriately juxtaposing a combination of ETS domain protein binding sites. C.

␮ Enhancer Deoxyribonucleic Acid Flexure Is Increased by Protein Binding

The studies described showed that changes in either the spacing or the orientation of the µA and µB sites drastically reduced enhancer activity in B cells, but did not affect protein–DNA interactions. Clearly, recruiting the proteins to the DNA is not sufficient to activate transcription; rather, a stereochemically precise super-

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complex of the three proteins must be assembled appropriately on the DNA. To examine one structural component of such a complex, we investigated whether the proteins that bound to the µA, µE3, and µB elements affected DNA structure. We assayed two categories of changes in DNA structure that are known to result from DNA–protein interactions: (1) an increase in DNA flexure and (2) the induction of a directed bend in the DNA. Deoxyribonucleic acid flexure refers to a distortion in the DNA that brings the ends of the DNA molecule closer to each other, but not in any fixed orientation. One way to visualize it is a bend in the DNA in which one ‘‘half ’’ of the molecule can take on several positions on part of the surface of a cone (Fig. 4). A directed bend is one in which the ‘‘half’’ of the molecule takes up a unique position on the surface of such a cone (bold arrow). Protein-induced changes in DNA flexure were determined by circular permutation assays. These assays were carried out using DNA fragments where the µ enhancer was located at different positions relative to the end of the DNA. Increased flexure was detected as reduced mobility of the DNA/protein complex in nondenaturing acrylamide gels. Two proteins, PU.1 and TFE3, scored positive in this assay, increasing DNA flexure by 48° and 40°, respectively. By comparison, Ets-1, a µA binding protein, had no effect on DNA structure in these analyses.

Figure 4 Model of how a DNA binding protein can alter DNA structure. A DNA binding protein that increases DNA flexure can result in numerous conformations indicated by thin and dotted arrows. A DNA binding protein that induces a directed DNA bend locks the DNA into a particular structure, represented by the bold arrow.

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D.

Only PU.1 Protein Induces a Directed Bend in ␮ Enhancer Deoxyribonucleic Acid

Although circular permutation analysis provides evidence for protein-induced DNA distortion, this assay more accurately reflects changes in DNA flexure that result from protein–DNA interactions. Alternatively, anomalous migrations in circular permutation assays may result from the nonglobular conformations of some DNA binding proteins [14]. To distinguish between DNA distortion and directed DNA bending, we performed phasing analysis. For phasing analysis, the µ enhancer fragment was cloned adjacent to three poly A tracts [14,15]. Each poly A tract confers an intrinsic DNA bend of 18° for an overall 54° minor groove directed DNA bend. This bend serves as an internal standard against which protein-induced bending in the adjacent DNA fragment can be compared. The µ enhancer was cloned at five different distances (designated the spacer length and ranging from 29 to 41 bp) from the A 5 tracts to position it at various helical positions relative to the intrinsic minor groove bend (Fig. 5A). Protein-induced bends directed toward the minor groove reinforce the intrinsic A tract bend when the protein binding site is positioned at integral (n) helical turns from the A tracts. In contrast, major groove bends reinforce the intrinsic bend when the protein binding site is positioned on the opposite helical face (n ⫹ 5). If a DNA binding protein induces only a change in flexure, the location of the binding site with respect to the A tracts does not affect DNA bending as assayed by nucleoprotein complex migration. The EMSA (electrophoretic mobility shift assay) analysis demonstrates bend reinforcement by decreased mobility of the DNA–protein complex. We performed phasing analysis with the DNA binding domain of PU.1 and Ets-1 [ETS (PU.1) and ETS (Ets-1) respectively], and a naturally occurring splice variant of TFE3, TFE3S (Fig. 5B). Differences in protein–DNA complex mobility were apparent for ETS(PU.1) but not for ETS(Ets-1) or TFE3S, indicating that only ETS(PU.1) induced a directed DNA bend. Results with full-length PU.1 were similar to results with ETS(PU.1) (data not shown). Phasing analysis further allowed us to calculate the degree of ETS(PU.1)-induced DNA bending, as described by Thompson and Landy [16] and modified by Kerpolla and Curran [14,15,17]. The best fit cosine curve for ETS(PU.1) (Fig. 5C) indicated that PU.1 bent µ enhancer DNA to an angle of 24° after correction for the slight bend (1°) contributed by the probe alone. Direction of the ETS(PU.1)-induced DNA bend can also be ascertained from the phasing analysis once the center of the bend is determined. Although circular permutation results indicated the center of the ETS(PU.1)-induced flexure lies within the µB element, the precise bend center of this nonpalindromic site is not discernible by this method. However, PU.1 binding to the µB site (5′TTCCCCAAA-3′) induces a strong deoxyribonuclease (DNAase) I–hypersensi-

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Figure 5A Phasing analysis DNA fragments. The helical positioning of µ enhancer protein binding sites relative to an internal control bend (A 5 tracts) was altered by increasing spacer DNA from 29 to 41 bp. Directed DNA bending was interpreted relative to the internal control bend. DNA, deoxyribonucleic acid.

tive site in the noncoding strand between the C and A residues opposite the T-C junction of the coding strand, suggesting protein induced distortion of the phosphodiester backbone [18]. We approximated that this hypersensitive site is the center of the PU.1-induced DNA bend and calculated the lengths between µB and the poly A tract bend centers with reference to the position of the hypersensitive site. The ETS(PU.1)-induced DNA bend reinforced the poly A tract bend, resulting in the slowest mobility nucleoprotein complex when the µB site was positioned 36 bp (3.4 helical turns) from the poly A tract. Because ETS(PU.1) reinforced the minor groove-directed poly A tract bend when the µB element was on the opposite side of the DNA helix, we propose ETS(PU.1) bends µ enhancer DNA toward the major groove (Fig. 6).

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Figure 5B PU.1 induces a directed bend in µ enhancer DNA. EMSA analysis demonstrates the ETS (PU.1) nucleoprotein complex migration changes as the helical alignment (determined by spacer length) between µB and the A 5 tracts changes. ETS (Ets-1) and TFE3S-µ enhancer complex migration is independent of spacer length. ETS (PU.1) or ETS(Ets-1)-DNA complexes are bracketed; TFE3S-DNA complex is demarcated by an arrow. ETS (PU.1), and ETS (Ets-1) refer to the ETS domains of the transcription factors PU.1 and Ets-1, respectively. EMSA ⫽ electrophoretic mobility shift assay.

Figure 5C The best-fit cosine curve of ETS (PU.1)–µ enhancer nucleoprotein complexes from Figure 5B. Analysis indicated ETS (PU.1) induces a 24° directed bend in the µ enhancer DNA. ETS (PU.1) ⫽ the ETS domain of PU.1.

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Figure 6 Model: a functional nucleoprotein complex on the µ enhancer. We hypothesize specific interfaces must be formed by protein–protein interactions of Ets-1, PU.1, and TFE3 to result in B cell–specific enhancer activity. Appropriate alignment and DNA structure of µA, µE3, and µB forms a scaffold for this active complex. One possible mechanism for activity is that Ets-1/PU.1 interfaces are needed to recruit transcriptional coactivator(s).

III. DISCUSSION To facilitate mechanistic analysis of the lymphoid-specific immunoglobulin µ heavy chain gene enhancer, we previously described a minimal enhancer that contains three sequence elements: µA, µE3, and µB [6]. Here we have summarized the results of experiments that show that precise spatial organization of the sites, and consequently the three-dimensional nucleoprotein structure formed with enhancer DNA, is necessary for B cell–specific transcriptional enhancement. Furthermore, one of the three minimal enhancer binding proteins induces a directed bend in the DNA that may facilitate the formation of the functional stereospecific nucleoprotein complex. Comparison of the activity of orientation mutated enhancers in B cells and macrophages showed that the requirements differed greatly in the two cell types. For example, altering the head-to-tail organization of the µA and µB elements abrogates transcriptional activity in B cells but results in relatively little change in macrophage activity (i.e., Fig. 3, F2, 3, and 4). Note that in F1 the ‘‘arrowhead’’ of µA points toward µB, whereas in F4, the arrowhead of µA points away from µB. Inactivity of F4 demonstrates that proper 5′-3′ organization of µA and µB is insufficient for enhancer activity, and the further stereochemical considerations constrain the combinatorial use of ETS protein binding sites in both cell types. These results are reminiscent of the properties of hormone receptor binding

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sites, where the spacing of half sites has been shown to affect the transcriptional outcome [19,20]. In our interpretation of the PU.1 phasing data, we approximated the center of the bend to the site of a strong PU.1 induced DNAase 1–hypersensitive site between residues 5 and 6 on the noncoding strand of the µB site, 5′-TATTTGGGGAAGGGAA-3′. We have found that the three adenosine residues complementary to the three contiguous thymidines in the µB site score strongly in methylation interference assays [18]. Since the methyl group of methyl adenosines fall in the minor groove, these observations suggest that PU.1 is located over the minor groove in the vicinity of the PU.1 induced bend. Taken together with the phasing analysis that shows the induced bend is toward the major groove, our observations are consistent with the DNA’s being bent away from the bound PU.1 protein. It remains to be determined how the PU.1-induced DNA bend contributes to the formation of the three-protein DNA complex.

CONCLUSIONS Our data suggest that precise stereochemistry of the µ enhancer nucleoprotein complex is critical for B cell–specific immunoglobulin heavy chain gene expression. The relative positioning of ETS domain proteins and the PU.1-induced DNA bend determines the nature and extent of both protein–DNA and protein–protein contacts within the multicomponent complex. We suggest that these contacts define an active enhancer only in B cells because of the coexpression of a unique combination of transcriptional activators.

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