Alkaloids: Chemical and Biological Perspectives, Volume 13, Volume 13

Alkaloids: Chemical and Biological Perspectives

Related Titles of Interest Books GAWLEY & AUBE Principles of Asymmetric Synthesis GRIBBLE & GILCHRIST Progress in Heterocyclic Chemistry, Volume 10 SESSLER & WEGHORN Expanded Contracted and Isomeric Porphyrins PELLETIER Alkaloids: Chemical Alkaloids: Chemical Alkaloids: Chemical Alkaloids: Chemical

& & & &

Biological Biological Biological Biological

Perspectives, Perspectives, Perspectives, Perspectives,

Volume Volume Volume Volume

9 10 11 12

WONG & WHITESIDES Enzymes in Synthetic Organic Chemistry Major Reference Works BARTON, NAKANISHI, METH-COHN Comprehensive Natural Products Chemistry KATRITZKY & REES Comprehensive Heterocyclic Chemistry I CD-Rom KATRITZKY, REES & SCRIVEN Comprehensive Heterocyclic Chemistry II Journals Bioorganic & Medicinal Chemistry Bioorganic & Medicinal Chemistry Letters Carbohydrate Research Heterocycles (distributed by Elsevier) Phytochemistry Tetrahedron Tetrahedron Asymmetry Tetrahedron Letters

Full details of all Elsevier Science publications, and a free specimen copy of any Elsevier Science journal, are available on request from your nearest Elsevier Science office.

ALKALOIDS: CHEMICAL AND BIOLOGICAL PERSPECTIVES Volume Thirteen

Edited by

S. WILLIAM PELLETIER Institute for Natural Products Research and Department of Chemistry The University of Georgia, Athens

1999

PERGAMON An Imprint of Elsevier Science Amsterdam - Lausanne - New York - Oxford - Shannon - Singapore - Tokyo

Elsevier Science Ltd The Boulevard, Langford Lane Kidlington, Oxford O X 5 1GB, U . K . © 1999 Elsevier Science Ltd. All rights reserved. This work is protected under copyright by Elsevier Science Ltd, and the following terms and conditions apply to to its use: Photocopying Single photocopies of single chapters may be made for personal use as allowed by national copyright laws. Permission of the publisher and payment of a fee is required for all other photocopying, including multiple or systematic copying, copying for advertising or promotional purposes, resale, and all forms of document delivery. Special rates are available for educational institutions that wish to make photocopies for non-profit educational classroom use. Permissions may be sought directly from Elsevier Science Rights & Permissions Department, PO Box 800, Oxford OX5 IDX, UK; phone: ( + 44) 1865 843830, fax: ( + 44) 1865 853333, e-mail: [email protected]. You may also contact Rights & Permissions directly through Elsevier's home page (http://www.elsevier.nl), selecting first 'Customer Support', then 'General Information', then 'Permissions Query Form'. In the USA, users may clear permissions and make payments through the Copyright Clearance Center, Inc., 222 Rosewood Drive, Danvers, MA 01923, USA; phone: (978) 7508400, fax: (978) 7504744, and in the UK through the Copyright Licensing Agency Rapid Clearance Service (CLARCS), 90 Tottenham Court Road, London WIP OLP, UK; phone: ( + 44) 171 436 5931; fax: ( + 44) 171 436 3986. Other countries may have a local reprographic rights agency for payments. Derivative Works Tables of contents may be reproduced for internal circulation, but permission of Elsevier Science is required for external resale or distribution of such material. Permission of the publisher is required for all other derivative works, including compilations and translations. Electronic Storage or Usage Permission of the publisher is required to store or use electronically any material contained in this work, including any chapter or part of a chapter. Contact the pubHsher at the address indicated. Except as outlined above, no part of this work may be reproduced, stored in a retrieval system or transmitted in any form or by any means, electronic, mechanical, photocopying, recording or otherwise, without prior written permission of the publisher. Address permissions requests to: Elsevier Science Rights & Permissions Department, at the mail, fax and e-mail addresses noted above. Notice No responsibility is assumed by the Publisher for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions or ideas contained in the material herein. Because of rapid advances in the medical sciences, in particular, independent verification of diagnosis and drug dosages should be made. First edition 1999 Library of Congress Cataloging in Publication Data A catalog record from the Library of Congress has been applied for. British Library Cataloguing in Publication Data A catalogue record from the British Library has been applied for. ISBN: 0-08-043403-7 Transferred to digital printing 2005 Printed and bound bv Antony Rowe Ltd, Eastbourne

Dedicated to the memory of

Ernst Spath (1886-1946) Ernst Spath was bom in Bam, a small village in Austro-Hungarian Europe. After completion of the doctorate degree (1910), he was on the faculty of Vienna University for over thirty years where he maintained a large group of research students. He attained a wide reputation for his researches on the chemistry of plant substances; the study of alkaloids, was of special interest to Spath. He made important contributions to the chemistry of alkaloids: the tetrahydroisoquinolines (papaverine, a-methylnorlaudanosine, anhalidine, anhalonine, carnegine), quinolines (galipoline, cusparine), aporphines (bulbocarpine, A^-methyllaurotetanin), benzophenanthridines (chelidonine, chelerythrine, sanguinarine), amaryllidaceae (tazettine), quinolizidine (cytisine), etc. He investigated the pyridine and piperidine alkaloids such as nicotine, anabasine, and conhydrine. Spath also worked on other pharmacologiclly important alkaloids: ephedrine, conessine, mescaline, harmine, harmaline, and calycanthine. Besides the alkaloids, he explored other classes of natural products such as coumarins, flavonoids, lignans, and saponins. Spath had a love for Vienna which he adopted as his home. On September 30, 1946 while attending the celebration of the Swiss Society of Natural Sciences, he succumbed to a heart attack and passed away soon after his sixtieth birthday.

B. S. Joshi

This Page Intentionally Left Blank

Contributors Uffe Anthoni, Marine Chemistry Section, Department of Chemistry, University of Copenhagen, Universitetsparken 5, DK-2100, Copenhagen, DENMARK. Todd A. Blythe, Vertex Pharmaceuticals, 130 Waverly Street, Cambridge, Massachusetts 02139-4242, U.S.A. Carsten Christopherson, Marine Chemistry Section, Department of Chemistry, University of Copenhagen, Universitetsparken 5, DK-2100, Copenhagen, DENMARK. John W. Daly, Laboratory of Bioorganic Chemistry, National Institutes of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland 20892, U.S.A. Ying Dong, Department of Chemistry and Bamett Institute, Northeastern University, 360 Huntington Avenue, Boston, Massachusetts 02115-5096, U.S.A. H. Martin Garraffo, Laboratory of Bioorganic Chemistry, National Institutes of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland 20892, U.S.A. Donna M. lula. Department of Chemistry, State University of New York at Stoney Brook, Stoney Brook, New York 11794, U.S.A. Balawant S. Joshi, Institute for Natural Products Research, Chemistry Building, The University of Georgia, Athens, Georgia 30602-2556, U.S.A. Philip W. LeQuesne, Department of Chemistry and Barnett Institute, Northeastern University, 360 Huntington Avenue, Boston, Massachusetts 02115-5096, U.S.A. Per Halfdan Nielson, Marine Chemistry Section, Department of Chemistry, University of Copenhagen, Universitetsparken 5, DK-2100, Copenhagen, DENMARK. Iwao Ojima, Department of Chemistry, State University of New York at Stoney Brook, Stoney Brook, New York 11794, U.S.A. S. William Pelletier, Institute for Natural Products Research and Department of Chemistry, The University of Georgia, Athens, Georgia 30602-2556, U.S.A. Thomas F. Spande, Laboratory of Bioorganic Chemistry, National Institutes of Diabetes and Digestive and Kidney Diseases, National Institutes of Health, Bethesda, Maryland 20892, U.S.A.

This Page Intentionally Left Blank

Preface Volume 13 of this series presents five timely reviews of research on alkaloids. Chapter 1, by John W. Daly, H. Martin Garraffo and Thomas F. Spande is a comprehensive update and supplement to the chapter, "Amphibian Alkaloids: Chemistry, Pharmacology, and Biology," written by Daly and Spande for volume 4 of Alkaloids: Chemical and Biological Perspectives (1986). This chapter presents newer developments in the chemistry and biology of alkaloids from amphibian skin. It provides a synopsis and tabulation of the hundreds of alkaloids that have been detected, with an emphasis on occurrence, structure, dietary origins, and biological activity. Also included are references to synthetic work that has appeared since the 1986 review. Chapter 2, by Uffe Anthoni, Carsten Christophersen and Per Halfdan Nielson reviews alkaloids containing the 1,2, 3, 3a, 8, 8a - hexahydropyrrolo [2,3b] indole ring system and the cyclotryptamines. This chapter provides an exhaustive list of available structures. Moreover, the chemical and biological structures have been evaluated critically so as to identify existing errors and expose regularities in appearance or biological function. In addition attention is drawn to the possible implications of the accumulated knowledge related to the synthesis, occurrence, and biochemistry of this class of alkaloids. Chapter 3 by Philip W. Le Quesne, Ying Dong and Todd A. Blythe summarizes recent work on alkaloids containing the comparatively non - basic pyrrole ring system. Over the past fifteen years there has been a dramatic increase in the number of pyrrole alkaloids identified. This chapter treats isolation, structure elucidation, biological activity, and selected chemical syntheses of certain pyrrole alkaloids. Chapter 4 by Balawant S. Joshi and S. William Pelletier surveys recent developments in the chemistry of diterpenoid and norditerpenoid alkaloids occurring in Aconitum, Delphinium and Consolida genera of the Ranunculaceae family. These plants have been used in tradition Chinese and Indian medicine as cardiotonics, febrifuges, sedatives, and anti - rheumatics. Because recent studies have shown that diterpenoid alkaloids are the active constituents responsible for the medicinal properties of these plants, great interest in these alkaloids has developed. Chapter 5 by Iwao Ojima and Donna M. lula focuses on transition metal - catalyzed carbonylations as efficient and novel approaches to the construction of piperidine, izidine and quinazoline alkaloids, which occur in great numbers in nature. Because of their diverse biological activities, these compounds have attracted the attention of synthetic, medicinal, pharmaceutical, and organic

X

Preface

chemists. The alkaloids, as synthetic targets, have contributed to growth and development of modem organic syntheses. Each chapter in this volume has been reviewed by at least one specialist in the field. The editor thanks these reviewers for the important contributions they have made to this volume. Indexes for both subjects and organisms are provided. The editor invites prospective contributors to write to him about topics for review in future volumes of this series. S. WiUiam Pelletier Athens, Georgia August 10,1998

Contents of Previous Volumes Volume 1 1. The Nature and Definition of an Alkaloid S. William Pelletier 2. Arthropod Alkaloids: Distribution, Functions, and Chemistry Tappey H. Jones and Murray S. Blum

33

3. Biosynthesis and Metabolism of the Tobacco Alkaloids Edward Leete

85

4. The Toxicology and Pharmacology of Diterpenoid Alkaloids M. H. Benn and John M. Jacyno 5. A Chemotaxonomic Investigation of the Plant Families of Apocynaceae, Loganiaceae, and Rubiaceae by Their Indole Alkaloid Content M. Volkan KisabUrek, Anthony J.M. Leeuwenberg, and Manfred Hesse

153

211

Volume 2 1. Some Uses of X-ray Diffraction in Alkaloid Chemistry Janet Finer-Moore, Edward Arnold, and Jon Clardy 2. The Imidazole Alkaloids Richark K. Hill 3. Quinolizidine Alkaloids of the Leguminosae: Structural Types, Analyses, Chemotaxonomy, and Biological Properties A. Douglas Kinghom arid Manuel F. Balandrin

49

105

4. Chemistry and Pharmacology of Maytansinoid Alkaloids Cecil R. Smith, Jr. and Richard G. Powell 5. l^C and Proton NMR Shift Assignments and Physical Constants of Ci9-Diterpenoid Alkaloids S. William Pelletier, Naresh V. Mody, Balawant S. Joshi, and Lee C. Schramm

149

xii

Contents of Previous Volumes

Volume 3 1. The Pyridine and Piperidine Alkaloids: Chemistry and Pharmacology GaborB. Fodor and Brenda Colasanti 2. The Indolosesquiterpene Alkaloids of the Annonaceae Peter G. Waterman

1

91

3. Cyclopeptide Alkaloids Madeleine M. Joullie and Ruth F. Nutt

113

4. Cannabis Alkaloids Mahmoud A. ElSohly

169

5. Synthesis of Lycopodium Alkaloids Todd A. Blumenkopf and Clayton H. Heathcock

185

6. The Synthesis of Indolizidine and Quinolizidine Alkaloids of Tylophora, Cryptocarya, Ipomoea, Elaeocarpus, and Related Species R. B. Herbert

241

1. Recent Advances in the Total Synthesis of Pentacyclic Aspidosperma Alkaloids Larry E. Overman and Michael Sworin

275

Volume 4 1. Amphibian Alkaloids: Chemistry, Pharmacology and Biology John W. Daly and Thomas F. Spande 2. Marine Alkaloids and Related Compounds William Fenical 3. The Dimeric Alkaloids of the Rutaceae Derived by Diels-Alder Addition Peter G. Watermann 4. Teratology of Steroidal Alkaloids Richard F. Keeler

275

331

389

Contents of Previous Volumes

Volume 5 1. The Chemistry and Biochemistry of Simple Indolizidine and Related Polyhydroxy Alkaloids Alan D. Elbein and Russell J. Molyneux 2. Structure and Synthesis of Phenanthroindiolizidine Alkaloids and Some Related Compounds Emery Gellert

55

3. The Aporphinoid Alkaloids of the Annonaceae Andre Cave, Michel Leboeuf, Peter G. Waterman

133

4. The Thalictrum Alkaloids: Chemistry and Pharmacology Paul L Schijf, Jr.

271

5. Synthesis of Chephalotaxine Alkaloids Tomas Hudlicky, Lawrence D. Kwart, and Josephine W. Reed

639

Volume 6 1. Chemistry, Biology and Therapeutics of the Mitomycins William A. Remers and Robert T. Dorr 2. Alkaloids of Tabemaemontana Species Teris A. van Beek and Marian A.J.T. van Gessel 3. Advances in Alkaloid Total Synthesis via Iminium Ions, a-Aminocarbanions and a-Aminoradicals David J. Hart 4. The Biosynthesis of Protoberberine Alkaloids Christopher W. W. Beecher and William J. Kelleher 5. Quinoline, Acridone and Quinazoline Alkaloids: Chemistry, Biosynthesis and Biological Properties Michael F. Grundon

75

227

297

339

xiv

Contents of Previous Volumes

Volume 7 1. Homoerythrina and Related Alkaloids /. Ralph C. Bick and Sirichai Panichanum 2. Carbon-13 NMR Spectroscopy of Steroidal Alkaloids Pawan K. Agrawal, Santosh K. Srivastava, and William Gaffield 3. Carbon-13 and Proton NMR Shift Assignments and Physical Constants of Norditerpenoid Alkaloids S. William Pelletier and Balawant S. Joshi

1

43

297

Volume 8 1. Curare Norman G. Bisset 2. Alkaloid Chemistry and Feeding Specificity of Insect Herbivores James A. Saunders, Nichole R. O'Neill, and John T. Romero

151

3. Recent Advances in the Synthesis of Yohimbine Alkaloids Ellen W. Baxter and Patrick S. Mariano

197

4. The Loline Group of Pyrrolizidine Alkaloids Richard G. Powell and Richard J. Petroski

320

Contents of Previous Volumes

Volume 9 1. Taxol M.E. Wall and M. C. Want 2. The Synthesis of Macroline Related Sarpagine Alkaloids Linda K. Hamaker and James M. Cook

23

3. Erythrina Alkaloids Amrik Singh Chawla and Vijay K. Kapoor

85

4. Chemistry, Biology and Chemoecology of the Pyrrolizidine Alkaloids Thomas Hartmann and Ludger Witte 5. Alkaloids from Cell Cultures of Aspidosperma Quebracho-Bianco P. ObitZy J. StockigU L. A. Mendonza, N, Aimi and S.-i. Sakai 6. Fumonisins Richard G. Powell and Ronald D. Plattner

155

235

247

Volume 10 1. Alkaloids from Australian Flora /. R. C. Bick 2. Pyridine and Piperidine Alkaloids: An Update Marilyn J. Schneider

155

3. 3-Alkylpiperidine Alkaloids Isolated from Marine Sponges in the Order Haplosclerida Raymond J. Andersen, Rob W. M. Van Soest and Fangming Kong

301

4. p-Carboline and Isoquinoline Alkaloids from Marine Organisms BillJ. Baker

357

xvi

Contents of Previous Volumes

Volume 11 1.

The Thalictrum Alkaloids: Chemistry and Pharmacology (1985 - 1995) Paul L Schiff, Jr.

2.

Taxine Giovanni Appendino

3.

The Alkaloids of South American Menispermaceae Mary D. Menachery

4.

The Chemistry and Biological Activity of Calystegines and Related A^c>rtropane Alkaloids Russell J. Molyneux, Robert J, Nash, and Naoki Asano

5.

Polyhydroxylated Alkaloids that Inhibit Glycosidases Robert J. Nash, Naoki Asano, and Alison A. Watson

Volume 12 1.

Acronycine-type Alkaloids: Chemistry and Biology Frangois Tillequin, Sylvie Michel, and Alexios-IJandros Skaltsounis

2.

Solanum Steroid Alkaloids — an Update Helmut Ripperger

3.

Synthesis and Structure-Activity Studies of Lissoclinum Peptide Alkaloids Peter Wipf

4.

Pyroglutamate as a Chiral Template for the Synthesis of Alkaloids Michael B. Smith

5.

Analysis of Alkaloids by Capillary Electrophoresis and Capillary Electrophoresis — Electrospray Mass Spectrometry Joachim Stockigt, Matthias linger, Detlef Stockigt, and Detlev Belder

6.

Oxidation of Anthelmentic Marcofortine A, an Indole Alkaloid Byung H. Lee, Michael F. Clothier, and Gabe I Komis

Contents 1. Alkaloids from Amphibian Skins John W. Daly, H. Martin Garraffo and Thomas I;. Spande

1

2.

Naturally Occurring Cyclotryptophans and Cyclotryptamines Uffe Anthoni, Carsten Christophersenand Per Halfdan Nielson

163

3.

Recent Research on Pyrrole Alkaloids Philip W. LeQuesne, Ying Dong and Todd A. Blythe

237

4.

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids Balawant S. Joshi and S. William Pelletier

289

5.

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids by Means of Transition Metal Catalyzed Carbonylations Iwao Ojima and Donna M. I d a

37 1

Subject Index

413

Organism Index

425

xvii

This Page Intentionally Left Blank

Chapter One

Alkaloids from Amphibian Skins John W. Daly, H. Martin Garraffo and Thomas F. Spande Laboratory of Bioorganic Chemistry National Institute of Diabetes and Digestive and Kidney Diseases National Institutes of Health Bethesda, MD 20892 CONTENTS 1. INTRODUCTION

2

2. STEROIDAL ALKALOIDS 2.1. Samandarines 2.2. Batrachotoxins

3 3 6

3. BICYCLIC ALKALOIDS 3.1. Pumiliotoxins and Allopumiliotoxins 3.1.1. 8-Deoxypumiliotoxins and other Pumihotoxin Congeners 3.2. Homopumihotoxins 3.2.1. Homopumiliotoxin Congeners 3.3. Histrionicotoxins 3.4. Decahydroquinolines 3.5. Pyrrolizidines (azabicyclo[3.3.0]octanes) 3.6. Indolizidines (azabicyclo[4.3.0]nonanes) 3.6.1. 3,5-Disubstituted Indolizidines 3.6.2. 5,8-Disubstituted Indolizidines 3.6.3. 5,6,8-Trisubstituted Indolizidines 3.7. Quinolizidines (azabicyclo[4.4.0]decanes) 3.7.1. 4,6-Disubstituted Quinolizidines 3.7.2 . 1,4-Disubstituted Quinolizidines 3.8. Azabicyclo[5.3.0]decanes

10 12 23 25 28 31 39 48 53 53 61 67 71 71 72 76

4. TRICYCLIC ALKALOIDS 4.1. Gephyrotoxins 4.2. Cyclopenta[6]quinolizidines 4.3. Coccinellines 4.4. Spiropyrrolizidines

77 78 80 83 85

5. MONOCYCLIC ALKALOIDS 5.1. Pyrrolidines 5.2. Piperidines

89 89 92

6. PYRIDINE ALKALOIDS 6.1. Epibatidine

95 96

2

J. W. Daly, H. M. Garraffo and T. F. Spande

6.2. Noranabasamine

103

7. INDOLE ALKALOIDS 7.L Pseudophrynamines 7.2. Chimonanthine/Calycanthine

104 104 108

8. SUMMARY

109

APPENDIX

113

REFERENCES

147

1.

INTRODUCTION

A remarkably diverse array of biologically active compounds occurs in amphibian skin [1]. These include biogenic amines and derivatives thereof, peptides, proteins, bufadienolides, tetrodotoxins, and numerous lipophilic alkaloids. Such toxic/noxious compounds are sequestered in granular glands of skin where they serve under duress as a passive chemical defense against predators. Some of the peptides may, in addition, serve to protect amphibian skin from bacterial or protozoan infections [2]. While most of these biologically active compounds found in amphibian skin are presumably produced by the amphibian, the tetrodotoxins and most of the lipophilic alkaloids apparently have another source. The tetrodotoxins probably are formed by a symbiotic microorganism [3,4], while the lipophilic alkaloids with the exception of the steroidal samandarines appear to be derived without metaboUc change from dietary arthropods [5-8]. Over five hundred lipophilic alkaloids have been discovered in amphibian skin and with the exception of less than thirty, none have been detected elsewhere in Nature. The few that have been found elsewhere occur in ants, beetles, and millipedes, all of which are known prey items for the amphibians that contain skin alkaloids. We have previously reviewed the chemistry and biology of alkaloids of amphibian skin first in 1982 [9], then in 1986 [10] and again m 1993 [11]. Synthetic efforts directed towards alkaloids of amphibian skin have been reviewed in detail in 1982-3 [9,12] and in 1986 [10]. The present chapter will attempt to cover further developments in chemistry and biology of alkaloids from amphibian skin and, thereby, provide a current synopsis and tabulation of the hundreds of alkaloids that have been detected with an emphasis on the structures, occurrence, possible dietary origins and biological activity. Some structures in the present review are, as yet, tentative, or incompletely defined and require further research. Most of these alkaloids have been detected only as trace constituents in skin extracts. The occurrence among amphibian genera of some twenty classes of lipophilic alkaloids is presented in Table 1 in the Appendix. The unique structures of these lipophilic alkaloids have provided and still provide a challenge for chemical synthesis. The current chapter will cite references to synthetic work that has appeared since our 1986 review. All of the lipophilic alkaloids, except the samandarines and batrachotoxins, that have been detected in frog/toad skin

Alkaloids from Amphibian Skins

extracts are tabulated in the Appendix using a code designation based on use of the molecular weight and an identifying letter(s) in bold face.

2.

STEROIDAL ALKALOIDS

Two classes of steroidal alkaloids, the samandarines and the batrachotoxins, have been characterized from amphibian skin or skin glands. The samandarines have a modified steroidal A-ring containing a ring-nitrogen and in most cases an oxygen-bridge that forms an oxazolidine ring. They are known only from salamanders of the Eurasian genus Salamandra, where they occur in parotoid glands. They are apparently synthesized by the salamander. Samandarines are potent local anesthetics. The batrachotoxins have a homomorpholine ring at the steroidal CD-ring junction, a 3,9-hemiketal bridge and, for the two most toxic alkaloids, a 20p-2,4-dialkylpyrrole-3-carboxylate moiety. They are known in amphibians only from dendrobatid frogs of the neotropical genus Phyllobates, where they occur only in skin. It appears that batrachotoxins of frog skin are probably derived unchanged from dietary arthropods, but the identity of such batrachotoxin-containing arthropods remains shrouded in mystery. One of the highly toxic alkaloids of this class, homobatrachotoxin, has now been discovered in skin/feathers of New Guinean birds of the genus Pitohui. Batrachotoxins cause permanent opening of voltage-dependent sodium channels of nerve and muscle and represent a powerful tool for the study of such channels.

2.1.

Samandarines

The salamanders of Europe have been considered to be poisonous since ancient times and in 1886 the toxic principles from the brilliant black and yellow fire salamander were reported to be alkaloidal [see ref. 10 for a review of the early literature]. It remained for CI. Schopf and colleagues to isolate the major alkaloids from the parotoid glands of fire salamanders {Salamandra salamandra, Salamandridae) and to determine their structures. The studies began in the 1930's using classical methods of chemical conversions and IR and UV spectral analyses. Later, X-ray crystallographic analyses played a major role. By 1961 the structures of all nine naturally occurring samandarines (Figure 1) had been determined and in-depth reviews were provided first by Schopf [13] and later by his colleague G. Habermehl [14,15]. Most of the samandarines contain an oxazolidine ring, which is responsible for a pair of diagnostic infrared absorbances in the region of 830 to 875 cm"^ and for major mass spectral ions of C4H8NO"^ (m/z 86) and C4H7NO^ (m/z 85). A brief summary with references on the physical (m.p., optical rotation) and spectral (MS, IR and NMR) properties of samandarines has been provided [11].

J. W. Daly, H. M. Garraffo and T. F. Spande

(+)-Samandarine * O-Acetylsamandarine *

R= H (-)-Samandarone R = COCH3

OCOCH3

HN , 0 1

I H

(+)-Samandaridine

N'

I

I H

Cycloneosamandione *

Isocycloneosamandaridine Figure 1. Structures of samandarines. Isocycloneosamandaridine was originally referred to as cycloneosamandaridine [18] in what proved to be an incorrect structural analogy to cycloneosamandione. * Absolute configuration as shown. Synthesis. The syntheses of the samandarines have been reviewed most recently in 1986 [10]. The first such alkaloid to be synthesized was samandarone in 1967 by Hara and Oka [16,17]. Three of the natural samandarine alkaloids, namely samandinine, samandenone and isocycloneosamandaridine, have to our knowledge not been synthesized. The name

Alkaloids from Amphibian Skins

5

isocycloneosamandaridine was proposed [10] as more appropriate for a natural alkaloid, originally called cycloneosamandaridine and postulated to be either a C-19 carbinolamine corresponding to cycloneosamandione or an isomer with a C-6 carbinolamine [18]. Synthetic cycloneosamandaridine (C-19 carbinolamine) was not identical with the natural alkaloid [19]. The name cycloneosamandaridine is appropriately retained for the synthetic material, which like cycloneosamandione has a C-19 carbinolamine, while the name isocycloneosamandaridine is more appropriate for the natural alkaloid, presumably the C-6 carbinolamine [10,11]. A synthetic alkaloid with a 17p-hydroxyl group instead of the 16P-hydroxyl group of samandarine, has not been detected in Nature; it was referred to in our earlier reviews [10,11] as the Hara-Oka alkaloid. In our 1986 review certain references pertinent to synthetic efforts in the samandarine field were omitted: i) The synthesis of samane (deoxysamanine) and 17(i-hydroxysamane [20], neither of which have been detected in Nature. Samane had been obtained as a degradation product of cycloneosamandione [21]. ii) A stereoselective synthesis of the samandarine nucleus [22]. We are not aware of any further synthetic work in this area, since our review in 1986 [10]. Occurrence. The samandarines represent the first example of "animal alkaloids" and are known only from the two species of the genus Salamandra, namely the fire salamander {Salamandra salamandra) from Europe, northwestern Africa, and southwestern Asia as far as the Iraq-Iran border and the alpine salamander {Salamandra atra) from mountainous regions of central and southern Europe. The parotoid glands of each fire salamander contain about 20 mg of samandarine as the major alkaloid, while the glands of each alpine salamander contain about 5 mg. A closely related salamander of Italy, Salamandrina terdigitata, has not been investigated because of strict animal protection laws. The Australian myobatrachid frog, once proposed to contain samandarine alkaloids [23], was found instead [24, 25] to contain pumiliotoxins (see Section 3.1) and pseudophrynamines (see Section 7.1). There has been an erroneous report on the occurrence of the 17p-hydroxy isomer of samandarine (the Hara-Oka alkaloid) from the giant Japanese salamander, Cryptohranchus maximus, as discussed in our 1986 review [10]. The fire salamanders appear to synthesize the samandarine alkaloids, since there was no change in alkaloid content of parotoid glands over three generations reared in captivity (G. Habermehl, personal communication, 1989). Activity. Samandarines are highly toxic alkaloids with an injected lethal dose in mouse being about 70 jig [see refs. 10,11]. The toxicity is presumably due to very potent local anesthetic activity. Remarkably, the fire salamander is sensitive to the toxic effects of samandarine. Samandarines have antimicrobial activity, but only at millimolar concentrations. We are not aware of fiirther studies on the biological activity of samandarines since our reviews in 1986 and 1993 [10,11].

6

2.2.

J. W. Daly, H. M. Garraffo and T. F. Spande

Batrachotoxins

Brightly colored frogs of the Pacific coast of Colombia have been used, probably for centuries, to poison blow-darts used in hunting. The first report on this practice, namely the skewering of frogs and application of secretion from the frog skin to the tips of the blowdart, was in 1825 [see refs. 10,26,27 for early literature]. Efforts to establish the chemical nature of the toxic principles were rather unsuccessfiil until Marki and Witkop in 1963 reported that the toxic principles were alkaloidal [26]. Three species of frogs (Phyllobates aurotaenia, P. hicolor and P. terribilis, family Dendrobatidae) are now known to be used by Choco Indians of western Colombia to poison blow-darts; all contain the same alkaloids [27]. Successful isolation and structure elucidation of the alkaloids began in the mid-sixties with methanol extracts from thousands of skins from Phyllobates aurotaenia, a common, widely distributed, poison-dart frog in lowland western Colombia. Three major alkaloids, which were finally given the names batrachotoxin, homobatrachotoxin and batrachotoxinin A, were obtained. The structures are shown in Figure 2. A trace alkaloid, pseudobatrachotoxin, was unstable and converted to batrachotoxinin A at room temperature. X-ray crystallographic analysis of the 20P-p-bromobenzoate of batrachotoxinin A revealed the structure of the steroidal P-pregnane moiety [28]. Batrachotoxinin A was 3a,9a-epoxy14P, 18p-(epoxyethano-A^-methylimino)-5P-pregna-7,16-diene-3 p, 11 a,20p-triol. The stereodiagram in the initial publication [28] depicted the wrong enantiomer with the 20Sconfiguration. The correct stereodiagram with the 20R-configuration for batrachotoxinin A was presented in 1972 [29]. Unfortunately, the original incorrect 20a designation has persisted in the literature until now (Y. Kishi, personal communication, 1998). In addition, the synthetic enantiomer of 7,8-dihydrobatrachotoxinin A was designated as 20p when in actuality it was 20a as depicted in the stereodiagram of reference 29. The presence of a pyrrole moiety, indicated by prior observation of positive Ehrlich color reactions for batrachotoxin and homobatrachotoxin [30], followed by detailed NMR spectral analyses and NMR comparisons with ethyl dimethylpyrrole-3-carboxylates led to the structures of the highly toxic batrachotoxin and homobatrachotoxin [31, see refs. 10,11 for detailed reviews on structure elucidation]. Mass spectral analyses had been confounded by a failure to detect the molecular ion of batrachotoxin; direct probe analysis afforded an apparent molecular ion of m/z 399, due to pyrolytic elimination of the pyrrole carboxylate. Major fragment ions of CVHQNO:'' (m/z 139), C6H9N'^ (m/z 95) and C6H8N-' (m/z 94) for batrachotoxin and C8H11NO2'' (m/z 153), C7H8N02'' (m/z 138), CyHnN^ (m/z 109) and C^HgN^ (m/z 94) for homobatrachotoxin, all of which derive from the pyrrole carboxylate moiety, are diagnostic. Another major fragment ion, C4HioNO"^ (m/z 88), occurs for all three alkaloids, and derives from the homomorpholine ring bridging the steroidal CD-ring junction. Congeners of batrachotoxin and homobatrachotoxin with a 4P-hydroxyl group were later isolated from Phyllobates terribilis [32].

Alkaloids from Amphibian Skins

R= H

(-)-Batrachotoxinln A

rN>ni: R=

jr~\ CH3

(-)-Batrachotoxin *

N

H

Homobatrachotoxin

R= '2'^5

N H

Figure 2. Structures of batrachotoxins. Two minor congeners are 4P-hydroxybatrachotoxin and 4p-hydroxyhomobatrachotoxin. * Absolute configuration as shown. The stereodiagrams initially reported [28] were depicted unintentionally for the wrong enantiomer of batrachotoxinin A. Thus, the batrachotoxins have a 20R configuration, i.e., a 20P ester substituent, rather than the 20S that has been perpetuated in the literature [9-12]. A brief summary with references on the physical (optical rotation) and spectral (UV, IR, MS and NMR) properties of batrachotoxins has been provided [11]. The MS, proton NMR, and UV spectra of the batrachotoxins have been published [31]. An IR spectrum, probably of a mixture of batrachotoxin and homobatrachotoxin, has been published [26]. Reaction of batrachotoxins with methanol during purification has yielded 3-0-methyl derivatives, while reaction of acetone with the pyrrole moiety has yielded "dimers" [32]. Synthesis. The synthesis of batrachotoxinin A was accomplished by H. Wehrli and colleagues in 1972 [33, see also reviews in refs. 9-12]. Batrachotoxin had earher been prepared by acylation of natural batrachotoxinin A [31]. 7,8-Dihydrobatrachotoxinin A was also prepared by Wehrli's group and later by others [see ref 10 for review of synthetic efforts until 1986]. The synthesis of several derivatives of 7,8-dihydrobatrachotoxinin A and their biological activity appeared in 1988 [34]. A chiral synthesis of the AB ring system of batrachotoxins utilizing a carbene-mediated ring expansion of a chiral hydrindane was reported in 1993 [35]. In 1994, a stereoselective intramolecular Diels-Alder reaction was used to produce a functionalized steroid suitable for elaboration of the A-D rings of batrachotoxin [36]. Further elaboration was accomplished when the precursor steroid was converted, via a "trapped" Michael-addition product containing the necessary homomorpholine ring bridging the CD ring-junction, to a steroid with the A-D ring-system of batrachotoxinin A [37]. The conversion of this steroid to batrachotoxinin A has now been completed (Y. Kishi, personal communication, 1997).

8

J. W. Daly, H. M. Garraffo and T. F. Spande

Occurrence. The batrachotoxins are steroidal alkaloids that are unique in several structural features, in particular the 3,9-hemiketal function, the homomorpholine bridge and the 2,4dialkylpyrrole-3-carboxylate moiety. Until recently, batrachotoxins were known only from frogs of the genus Phyllobates of the neotropical family Dendrobatidae. Only the three Colombian species {P. terrihilis, P. hicolor and P. aurotaenia) out of the five species in the genus have levels of toxin in skin high enough to be used to poison blow-darts and only these three species deserve to be called poison-dart frogs. The highest levels occur in skin of P. terribilis, which contains about 500 |lg batrachotoxin, 300 |ig homobatrachotoxin and 200 )Lig batrachotoxinin A per frog [1,27]. Skins of the less toxic P. bicolor and P. aurotaenia contain about 20 |ig batrachotoxin, 10 |ig homobatrachotoxin, and 50 |ig batrachotoxinin A per frog. The two remaining Phyllobates species, both from Central America, contain very low levels of batrachotoxins. Skin of the Costa Rican species P. vittatus has only about 0,2 jig batrachotoxin, 0.2 |ig homobatrachotoxin and 2 L | Lg batrachotoxinin A per frog. Most populations of the Panamanian-Costa Rican species P. lugubris do not contain detectable amounts of batrachotoxins assessed either by toxicity or the sensitive Ehrlich color reaction. One population of P. lugubris from Panama, however, did contain trace levels of batrachotoxin, homobatrachotoxin, and batrachotoxinin A. Batrachotoxins have not been detected from frogs of other dendrobatid genera (Dendrobates, Epipedobates, and Minyobates) that contain monocyclic, bicyclic and tricyclic alkaloids in their skin. Levels of batrachotoxins in skin of wild-caught P. terribilis, maintained for years in terraria on fruit flies and crickets, slowly declined [38]. Furthermore, offspring of wild-caught P. terribilis raised in terraria did not contain detectable levels of batrachotoxins. At present the most likely explanation of these observations is that the Phyllobates species are dependent on a dietary source for batrachotoxins and that they have developed or over-expressed an efficient uptake system for accumulation of batrachotoxins and other alkaloids into the granular (poison) secretory glands of their skin. Such glands are thought to be the storage site for most, if not all, of the secondary metabolites present at elevated levels in skin of amphibians [39 and ref therein]. Captive-reared P. bicolor, which have no batrachotoxin in skin, were able to accumulate batrachotoxinin A, as provided on alkaloid-dusted fruit flies, into skin [7]. Other alkaloids provided on fruit flies were also accumulated into skin. Batrachotoxinin A was still present seven weeks after cessation of such feeding experiments, but no conversion to batrachotoxin or homobatrachotoxin had occurred. Both wild-caught and captive-raised frogs of the genus Phyllobates have sodium channels that are insensitive to batrachotoxin as reported for P. aurotaenia and P. terribilis [38,40]. Thus the frogs of this genus have evolved a batrachotoxin-resistant sodium channel, allowing them to eat a putative batrachotoxin-containing prey item. The channel retains sensitivity to veratridine, a plant alkaloid that activates sodium channels. Dendrobatid frogs from the genus Dendrobates have normal batrachotoxin-sensitive sodium channels [38] and such frogs rejected as "distasteful" fruit flies dusted with batrachotoxinin A [unpublished observation, see ref 7].

Alkaloids from Amphibian Skins

The nature of what small arthropod might be a dietary source of batrachotoxins is a complete mystery. The largest frog of the genus, Phyllobates terribilis (40 mm snout-vent length), has the highest levels of batrachotoxins, while the smallest frog of the genus, P. lugubris (21 mm snout-vent length), usually has no detectable amounts of batrachotoxins. Possibly, the putative batrachotoxin-containing prey is too large for the smallest species to eat. Frogs of the dendrobatid genus Phyllobates do eat larger prey items than non-batrachotoxin-containing frogs of the genus Dendrobates. However, it should be noted that the Colombian P. aurotaenia (28 mm snout-vent length), which has levels of skin batrachotoxins 100-fold higher than the Costa Rican P. vitattus, is not significantly larger than P. vittatus (26 mm snout-vent length). Other explanations are possible, including perhaps a ready availability of the putative prey item in the rain forest of western Colombia, where the three very toxic species occur, compared with a low availability in quite different habitats in Central America, where the two least toxic or even non-toxic species occur. Another possibility is that the alkaloid-accumulating system is expressed to differing extents in the five species, or may even have been virtually lost in the Central American species P. vittatus and P, lugubris. One of the batrachotoxins has recently been discovered in skin and feathers of New Guinean birds of the genus Pitohui (Muscicapidae) [41]. Only homobatrachotoxin was detected in the initial study, but one out of many subsequent samples had, in addition to homobatrachotoxin, trace amounts of batrachotoxin (J. Dumbacher, personal communication, 1997). Levels of homobatrachotoxin in skin and feathers were highest at about 20 |ig per bird in the hooded pitohui (Pitohui dichrous), about 10-fold lower per bird in the variable pitohui {Pitohui kirhocephalus) and even lower in the rusty pitohui {Pitohui ferrugineus) [41]. Levels of homobatrachotoxin in skin/feathers of the black pitohui {Pitohui nigrescens) were stated to be similar to those in the rusty pitohui [42]. In further studies, it was found that levels of homobatrachotoxins in skin/feathers of hooded pitohui varied greatly with the site of collection and that birds from certain areas in Papua New Guinea were virtually nontoxic [42]. The question as to whether homobatrachotoxin is synthesized by the birds, or whether there is a dietary source remains unanswered. Such birds have not yet been raised in captivity. Birds of the genus Pitohui subsist mainly on seeds and arthropods; in intial studies homobatrachotoxin has not been detected in seeds and other food items of the birds in Papua New Guinea (unpublished results with J. Dumbacher). Certain related New Guinean birds of the family Muscicapidae do not appear to be toxic (J. Dumbacher, personal communication, 1997). Activity. Batrachotoxins are extremely toxic with a lethal injected dose for batrachotoxin or homobatrachotoxin being less than 100 ng for mice [see ref 10]. Batrachotoxinin A is about 500-fold less toxic, but is still nearly as toxic as strychnine. The site of action for batrachotoxins is the voltage-dependent sodium channel of nerve and muscle [43]. Batrachotoxin appears to bind to an open form of the sodium channel, preventing the closing of the channel. The resultant massive influx of sodium depolarizes membranes of nerve and muscle, blocking their function. The development of a radioligand, batrachotoxinin A 20 p-

9

10

J. W. Daly, H. M. Garraffo and T. F. Spande

[^H]benzoate [44], has facilitated the study of other sodium channel activators, such as veratridine, and the study of the allosteric effects of a variety of agents, including local anesthetics. In 1985 batrachotoxin was reported to partially inhibit the activation of calcium channels elicited by a dihydropyridine [45]. The biological activity of batrachotoxin was reviewed in detail in 1986 [10]; a few of the many articles on batrachotoxin appearing in the next seven years were cited in 1993 [11]. In the four years since the 1993 review, there have been over fifty articles on the biological activity of batrachotoxin, but documentation is beyond the scope of this review. The use of batrachotoxinin A 20p-[^H]benzoate as a radioligand for the study of effects of local anesthetics, anticonvulsants and other drugs on voltage-dependent sodium channels was reviewed in 1992 [46]. A batrachotoxinin A 20p-o-azidobenzoate was introduced in 1993 as a photoaffmity probe for the batrachotoxin-binding site on the sodium channel [47] and subsequently one transmembrane segment of the sodium channel was identified as the major site of photoaffinity labeling with batrachotoxinin A 20(J-[^H]o-azidobenzoate [48]. A single site mutation conferred batrachotoxin-insensitivity to a rat muscle sodium channel expressed in cultured cells; veratridine-sensitivity was retained [48a].

3.

BICYCLIC ALKALOIDS

A variety of bicyclic alkaloids, ranging from the structurally novel pumiliotoxins, allopumiliotoxins, homopumiliotoxins and histrionicotoxins to relatively simple disubstituted decahydroquinolines and "izidines" (pyrrolizidines, indolizidines, quinolizidines and azabicyclo[5.3.0]decanes), have been characterized from amphibian skin, mainly from neotropical dendrobatid frogs of the genera Dendrobates, Epipedobates, Minyobates and Phyllobates, but also from subtropical/temperate South American bufonid toads of the genus Melanophryniscus, from Madagascan mantelline frogs of the genus Mantella, and from Australian myobatrachid frogs of the genus Pseudophryne. During the initial years, nearly a hundred bicyclic alkaloids were detected by GC-MS techniques and in 1978 a code system was introduced to designate these alkaloids, using in bold face the nominal molecular weight of the alkaloid with an identifying letter or letters to distinguish it from other alkaloids of the same nominal molecular weight [49]. To designate various stereoisomers, prefixes, such as cis, trans, epi, and iso, and primes (e.g. A', A") have been used. For certain izidines, the configurations of hydrogens on chiral carbons were indicated relative to the hydrogen on the chiral carbon at the lowest ring position, as in indolizidine 5Z,9£'-223AB, where the reference hydrogen is at C-3 (see Section 3.6.1). The occurrence of the various classes of bicyclic alkaloids in amphibian skin is presented in Table 1 in the Appendix. Structures of bicyclic alkaloids are presented (Figures 3,5,7,9,10,11,13,15,17,19, 21,23,25,27,29) along with structures in which stereochemistry and/or the nature or position of unsaturated moieties, hydroxyl groups and keto groups are not yet defined. Certain structures, some of which have been proposed in the literature, are labeled "tentative", when alternative structures cannot be excluded.

Alkaloids from Amphibian Skins

11

The pumiliotoxins have a unique 6-alkyHdene-8-hydroxy-8-methyl-l-azabicyclo[4.3.0] nonane structure, while the allopurhiHotoxins differ only in the additional presence of a 7hydroxyl group. The alkylidene side-chain varies from relatively simple to complex. Isoprenoid elements are usually present. Pumiliotoxins/allopumiliotoxins occur in skin of frogs from all the above mentioned dendrobatid, bufonid, mantelline and myobatrachid genera. Over forty pumiliotoxins/allopumiliotoxins have been detected. Presumably, the frogs sequester the pumiliotoxins/allopumiliotoxins into skin glands from unknown dietary arthropods. Since the pumiliotoxin/allopumiliotoxin-containing frogs have a world-wide distribution in tropical and subtropical regions, such dietary arthropods must also have a wide distribution. The identity of such putative pumiliotoxin/allopumiliotoxin-containing arthropods remains a mystery. Pumiliotoxins/allopumiliotoxins have cardiotonic and myotonic activity, probably due to positive modulatory effects on the open-time of sodium channels, and perhaps also to effects on release of intracellular calcium. The effects are highly dependent on structure. The homopumiliotoxins are 7-alkylidene-9-hydroxy-9-methyl-l-azabicyclo[4.4.0] decanes and, thus, are closely related in structure to the pumiliotoxins/allopumiliotoxins, but differ in having a quinolizidine rather than an indolizidine ring. The homopumiliotoxins, of which eleven have been detected, do not occur as commonly as pumiliotoxins/allopumiliotoxins in dendrobatid, mantelline and bufonid anurans and have not been detected in myobatrachid frogs. Presumably, like other alkaloids found in anuran skin, the homopumiliotoxins are sequestered unchanged from a dietary source. Like the pumiliotoxins/allopumiliotoxins, the homopumiliotoxins probably will prove to be positive modulators of sodium channels, but have not been available in sufficient quantities for pharmacological study. The histrionicotoxins have a unique 2,7-disubstituted azaspiro[5.5]undecanol structure. The sixteen histrionicotoxins detected in skin extracts from neotropical dendrobatid frogs have the same relative configuration and differ only in the length and degree of unsaturation of the two side-chains. In addition to internal and terminal double bonds, terminal allenic and acetylenic moieties occur in the side-chains. Histrionicotoxins have only been detected in skin extracts of the neotropical dendrobatid frogs. A dietary source for histrionicotoxins is, as yet, unknown, but it is suspected that the source of the histrionicotoxins, like the 2,5disubstituted decahydroquinolines that often accompany them in skin samples, will prove to be myrmicine ants. The histrionicotoxin-containing ants may be limited to the New World tropics, since only the neotropical dendrobatids contain histrionicotoxins. The histrionicotoxins could derive biosynthetically from a precursor with a linear carbon chain. The histrionicotoxins represent the first class of amphibian skin alkaloids shown to act as noncompetitive blockers of nicotinic receptor-channels. The decahydroquinolines, like most of the bicyclic alkaloids of amphibian skin, are disubstituted. The substituents vary in chain-length and degree of unsaturation. The stereochemistry at the ring junction {cis or trans) and the relative stereochemistry at the four chiral centers are highly variable. Vapor-phase FTIR spectra are diagnostic for stereochemistry. The dietary source for the forty some decahydroquinolines that have been detected in amphibian skins appears to be myrmicine ants, based on the recent discovery of

12

J. W. Daly, H. M. Garraffo and T. F. Spande

2,5-disubstituted decahydroquinolines in ants. The decahydroquinolines could derive biosynthetically from a precursor with a linear carbon chain. The decahydroquinolines are noncompetitive blockers of nicotinic receptor-channels. The remaining bicyclic alkaloids of amphibian skin are the so-called "izidines", which include the following classes: 3,5-disubstituted pyrrolizidines, 3,5-disubstituted indolizidines, 5,8-disubstituted indolizidines, 5,6,8-trisubstituted indolizidines, 4,6-disubstituted quinolizidines, 1,4-disubstituted quinolizidines and a novel new class of izidines consisting of 3,5-disubstituted azabicyclo[5.3.0]decanes. Over one-hundred izidine alkaloids have been detected in extracts of amphibian skin. The substituents in izidines vary in chainlength, degree of unsaturation and in the presence or absence of hydroxyl or keto groups. The relative stereochemistry at the chiral centers is variable in each class. EI-MS and vaporphase FTIR spectra are diagnostic. Myrmicine ants probably are the dietary source of 3,5disubstituted pyrrolizidines, 3,5-disubstituted indolizidines and 4,6-disubstituted quinolizidines. Possible dietary sources for 5,8-disubstituted indolizidines, 5,6,8-trisubstituted indolizidines, 1,4-disubstituted quinolizidines and 3,5-disubstituted azabicyclo[5.3.0]decanes have not yet been discovered. The 3,5-disubstituted pyrrolizidines, the 3,5-disubstituted indolizidines, the 4,6-disubstituted quinolizidines and the azabicyclodecanes could derive biosynthetically from a precursor with a linear carbon chain, while the remaining three izidine classes could not. Several of the izidine alkaloids have been shown to be noncompetitive blockers of nicotinic receptor-channels. Tentative structures for bicyclic alkaloids, in particular the izidine alkaloids, were often proposed primarily based on gas chromatography coupled with EI-MS, NH3-CI-MS and ND3-CI-MS, where data can be obtained from nanogram amounts of an alkaloid present in a mixture of dozens of other alkaloids. The technique of collision-activated NH3-CI-MS/MS analysis is now proving useftil particularly for certain izidine alkaloids (see Section 3.8). Microchemical procedures on alkaloid mixtures, such as perhydrogenation, base-catalyzed deuterium exchange, acetylation, etc., have proven useful. In recent years, vapor-phase FTIR spectral analysis has complemented the MS analyses, providing data relevant to the nature of functional groups, including terminal, conjugated and internal double bonds, acetylenes and allenes, hydroxyls and carbonyls, and ring stereochemistry, in particular cis versus trans ring junctions in decahydroquinolines and relative configuration of substituted carbons adjacent to nitrogen in decahydroquinolines and izidines. Vapor-phase FTIR spectra can be obtained from less than a microgram of a relatively volatile alkaloid present in mixtures of dozens of alkaloids. Isolation of 100 to 500 |ig of an alkaloid now provides sufficient material for detailed NMR spectral analysis. Synthesis of diastereomers or enantiomers can provide final verification of structure and relative or absolute stereochemistry.

3.1.

Pumiliotoxins and AUopumiliotoxins

Pumiliotoxin A and pumiliotoxin B were isolated in the mid-sixties as the major toxic alkaloids from skin extracts from twenty specimens of a population of a small dendrobatid

Alkaloids from Amphibian Skins

13

frog, Dendrobatespumilio, found in abundance on Isla Bastimentos, Panama [50]. Further collections yielded larger amounts of these pumiliotoxins. Analysis of the EI-MS and NMR spectra indicated that pumiliotoxin A and B differed only in the terminal portion of the side-chain, being -CH=CCH3CHOHCH2CH3 for pumiliotoxin A (307A) and -CH=CCH3CHOHCHOHCH3 for pumiliotoxin B (323A). A crystalline salt could not be obtained and the ring system remained incompletely defined, until X-ray analysis revealed the structure of a simpler congener, pumiliotoxin 251D, isolated in the late seventies from another dendrobatid frog, Epipedobates tricolor, of western Ecuador [51], Structures of pumiliotoxins A and B and of other pumiliotoxins and allopumiliotoxins were then deduced through analysis of NMR spectra in relationship to the NMR spectral assignments for pumiliotoxin 251D [52]. The structures of the allopumiliotoxins, namely pumiliotoxins with an additional ring hydroxyl group, were also deduced. Alkaloids had been assigned to the pumiliotoxin class based on prominent EI-MS fragment ions at m/z 166 and 70 or to the allopumiliotoxin class based on EI-MS fragment ions at m/z 182 and 70 (see below). The relative configurations of the side-chain hydroxyl groups of pumiliotoxin B were deduced by NMR model studies [53] and then the absolute configuration was defined by comparison of a dihydroxypentanone derived from pumiliotoxin B with that derived from (-)-tartaric acid [54]. A detailed account of the early structure elucidation of pumiliotoxins and allopumiliotoxins has been provided [10]. Structures and tentative structures for pumiliotoxins and allopumiliotoxins are shown in Figure 3. The absolute configuration is known for several pumiliotoxins/allopumiliotoxins, based either on X-ray crystallography (251D) or on enantioselective synthesis (267A, 307A, 323A, 339A and 339B). It has been assumed to be the same for the others, the structures of which are based mainly on MS and vapor-phase FTIR analysis (see caption to Figure 3). Some structures are tentative, in particular for alkaloids for which satisfactory vapor-phase FTIR spectra were not obtained. Structures for several alkaloids (207B, 293E, 297B, 305A, 307D, 307E, 325B, 341B, 353A, 357) that are currently assigned to the pumiliotoxin/ allopumiliotoxin class are not proposed because of insufficient data (see tabulation in Appendix). An erythro diastereomer of pumiliotoxin B (323 A) occurred as a minor alkaloid in skin extracts of an Australian myobatrachid frog and has been designated erythro-2i>2ZA [55]. The vapor-phase FTIR spectra of pumiliotoxins and allopumiliotoxins have proven to be of diagnostic value. Vapor-phase FTIR spectra for 251D and 267A are presented in Figure 4. Vapor-phase FTIR spectra for 209F, 225F, 267C, 305B, 307A, 307B, 307F», 307G', 307H, 309A, 323A, erythro-323A, 323B and 325A' have been published [25,56,57]. The FTIR spectra show a sharp hydroxyl absorption at about 3544 cm'^ for pumiliotoxins and at about 3521 cm"^ for allopumiliotoxins, characteristic for the 8-hydroxyl group hydrogen-bonded to the nitrogen. Pumiliotoxins have a characteristic Bohlmann band

J. W. Daly, H. M. Garraffo and T. F. Spande

14

'OH

H (-)-209F R: R: — OH 225E

237B

H (-)-225F R: R: - O H 241H

R: H (+)-251D* R: - O H (+)-267A* R: - - 0 H 7-epi-267A

R: H 281A R: - O H 297A

R: H 237A R: - O H 253A

2511

R: H 289C Tentative R: —OH 305C Tentative

Figure 3. Structures of the pumiliotoxins and allopumiliotoxins. Pumiliotoxin A is 307 A and pumiliotoxin B is 323A. *Absolute stereochemistry as shown. More than one diastereomer of 307A, 307G, 323B and 325A have been detected (see text).

Alkaloids from Amphibian Skins

R: R:

15

H 305B OH 321C R

R: H (+)-307A * (PTX A) R: - O H (+)-323B *

305D Tentative

307B

307C

(-)-307F'

307F"

307G

OH

307H Tentative Figure 3 (continued)

R: H 309A R: - O H 325A

J. W. Daly, H. M. Garraffo and T. F. Spande

16

OH

I

Ft

'OH

309D

R: H (+)-323A * (PTX B) R: —OH (+)-339A* R: - - 0 H (+)-339B* .OH

erythrO'323A 341A Figure 3 (continued) pattern with an absorption near 2798 cm"* and a shoulder at 2750-2600 cm"^ Allopumiliotoxins have a sharper Bohlmann band near 2800 cm"* with no shoulder. The fmger-print regions for pumiliotoxins and allopumiliotoxins are diagnostic. The EI-MS fragmentation pattern has been used to define an alkaloid as a member of the pumiliotoxin or allopumiliotoxin class. The pumiliotoxins are proposed to fragment as shown in Scheme 1 to yield as major ions CioHi6NO^ (m/z 166) and C4H8N^ (m/z 70), while the allopumiliotoxins would fragment in the same manner to yield as major ions CioHi6N02'^ (m/z 182) and C4H8N"^ (m/z 70). A brief summary with references to the physical (optical rotation) and spectral (IR, MS and NMR) properties of the pumilotoxins/ allopumiliotoxins has been provided [11]. Two of the alkaloids of the pumiliotoxin class appear to be structurally unique, namely 307H and 341A. Alkaloid 307H has been detected only in skin extracts from one species of Madagascan mantelline frogs. The vapor-phase FTIR of 307H [54] exhibits an enamine absorption at 1654 cm'*, a side-chain hydroxyl absorption at 3650 cm"* and an absorption at 993 cm"* typical of the 13,14-trisubstituted double bond of pumiliotoxin A and B. The hydrogen-bonded 8-hydroxyl absorption for pumiliotoxins at 3544 cm"* was replaced with an absorption at 3589 cm"*. There were no Bohlmann bands. It was proposed that 307H is the 5,6-double bond isomer of 307A [56, see Figure 3]. The other unique alkaloid, 341 A, was recently isolated in sufficient quantities from skin extracts of Epipedobates tricolor, a dendrobatid frog of southwestern Ecuador, to allow for proton NMR spectra. The NMR

17

Alkaloids from Amphibian Skins

1000-1 ^876.46

8

c € o

M

^

251D

2796.25

1460.24

.

600-^

1380.03 1312.75 1138.92 1094.79 962.646 21.7491

2800

2400 2000 Wavenumber (cm-1)

2800

2400 2000 Wavenumber (cm-1)

860

Figure 4. Vapor-phase FTIR spectra of pumiliotoxin 251D and allopumiliotoxin 267A. spectral analysis, combined with MS and vapor-phase FTIR spectra, led to a proposed 6,13cyclic ether structure [58, see Figure 3]. The ND3-CI-MS indicated three exchangeable hydrogens. Alkaloid 341B (no structure proposed) may be a diastereomer of 341 A. Alkaloid 357 is a hydroxy analog of 341 A. Certain coded alkaloids (structures not shown) of the pumiliotoxin/allopumiliotoxin class are apparently artefacts due to solvolysis or rearrangement at the C-15 ally lie hydroxyl

J. W. Daly, H. M. Garraffo and T. F. Spande

18

R"

R"

m/z166 166inPTXs, R' = H 182inalloPTXs, R' = OH 150in8-desoxyPTXs 152 in 8-desmethylPTXs -

^

m/z70 70 in PTXs 70 in alloPTXs 70 in 8-desoxyPTXs 70 in 8-desmethylPTXs Scheme 1 group of the side chain [see ref. 11]. These are 321A (15-O-methyl-307A), and 307A" and 323B" (epimers at C-15 of pumiliotoxin A (307A) and allopumiliotoxin 323B). Pumiliotoxin 307G is represented in skin extracts by more than one diastereomer, i.e., 307G* and 307G". Pumiliotoxin 307F"' appears to be the epimer at C-14 of 307F". Pumiliotoxin 267D, originally proposed as another new alkaloid, based on an apparently different GC elution time from that of pumiliotoxin 267C [59], was subsequently shown to be identical with 267C. A^-Oxides of pumiliotoxins/allopumiliotoxins have been detected on GC-MS analysis and afford longer retention times, but have EI and NH3-CI-MS virtually identical to the unoxygenated amines [60]. Deoxygenation appears to occur with NH3, but A^-oxides can be differentiated from the parent alkaloid by isobutane-CI-MS. A^-Oxides of pumiliotoxin B (323A) and allopumiliotoxin 267A were isolated and characterized [60]. Alkaloids 307D, 309C and 323F are probably A^-oxides of 307A, 309A and 323A, respectively. All of the allopumiliotoxins have the hydroxyl groups at C-7 and C-8 in the transdiaxial configuration except for 7-epi-267A [61] and 339B [52]. Such alkaloids with the hydroxyl groups at C-7 and C-8 in a cz.y-configuration form cyclic phenylboronides, while allopumiliotoxins with hydroxyl groups in the /ra«5-configuration do not [53]. Cyclic phenylboronides also form with pumiliotoxins/allopumiliotoxins such as 323A, that have vicinal hydroxyl groups in the side-chain [55]. Dimethylsilanates of certain pumiliotoxins have been detected as GC artefacts [see refs. 10,11]. All of the pumiliotoxins/allopumiliotoxins are tabulated in the Appendix.

Alkaloids from Amphibian Skins

19

Several structural congeners of the pumiliotoxins/allopumiliotoxins have been detected in frog skin extracts. These include the 8-deoxypumiliotoxins and the 8-desmethylpumiliotoxins (Section 3.1.1). The homopumiliotoxins, 9-deoxyhomopumiliotoxins, 9-desmethylhomopumiliotoxins and putative 8,9- dehydrohomopumiliotoxins contain a 7-alkylidenylquinolizidine ring (Section 3.2) rather than the 6-alkylidenylindolizidine ring of the pumiliotoxins/allopumiliotoxins. Synthesis. The early syntheses of pumiliotoxins and allopumiliotoxins were due to L. Overman and colleagues. Review^s on syntheses of pumiliotoxins and allopumiliotoxins are available [10,62,63]. Pumiliotoxin 251D was the first of such alkaloids to be synthesized [64], followed by syntheses of pumiliotoxins A (307A) and B (323A), and allopumiliotoxins 267A and 339B [65-67]. These enantioselective syntheses established the absolute configurations of these pumiliotoxins/allopumiliotoxins. The unnatural enantiomers have not been synthesized. Subsequent to the initial syntheses in the early eighties, other laboratories have reported synthetic routes to pumiliotoxin 251D [68,69] and allopumiliotoxins 267A [70], 339A [70], and 339B [71]. Overman has now provided further, more efficient routes, based not on the earlier iminium ion-vinylsilane cyclizations to pumiliotoxins [64-66] and allopumiliotoxins [72], or the lithioallene addition to a keto indolizidine derived from (-)-proline [67,73], but instead on reductive iminium ion-alkyne cyclizations, leading to pumiliotoxins A (307A) and B (323A) [74,75] and allopumiliotoxins 267A, 323B, 339A and 339B [76,77]. An unsaturated lactam was an intermediate in one synthesis of pumihotoxin 251D in which an aldol condensation introduced the side-chain [68]. This intermediate lactam was later synthesized by other routes [78-80], which thus represent formal syntheses of pumiliotoxin 251D. Recently, (-)-pumiliotoxin 209F has been synthesized, using an intramolecular palladium-catalyzed carbonylation-cyclization of an intermediate pyrrolidinyl vinyl iodide [81]. Two efficient synthetic routes to pumiliotoxins/ allopumiliotoxins are now available, utilizing either the reductive iminium ion-alkyne cyclizations developed by Overman and colleagues [74-77] or chromium-nickel-mediated cyclizations as developed by Kibayashi and colleagues [70,82]. The latter method has been utilized to synthesize allopumiliotoxins 267A and 339A [70,82]. Synthesis and biological investigation of 1 l-epi-, 1 l-desmethyl-16-e/^/- and IS-epi-lG-epi analogs of pumiliotoxin B (323A) and of pumiliotoxin analogs with 6'-hexenylidene and 7'-hydroxyheptylidene sidechains have been reported [83,84]. Iodide-promoted iminium ion-alkyne cyclizations have been used to prepare pumiliotoxin 251D and twelve analogs for biological study [85]. Occurrence. The pumiliotoxins and allopumiliotoxins have been detected in Nature only in extracts from amphibian skin. They often occur together and frequently are major alkaloids in skin extracts from frogs of the dendrobatid genera Dendrobates, Epipedobates and Minyobates. They are absent or only trace alkaloids in extracts from frogs of the dendrobatid genus Phyllobates [11]. Pumiliotoxins and/or allopumiliotoxins are major alkaloids in skin extracts of South American toads of the bufonid genus Melanophryniscus, as initially discovered for Melanophryniscus moreirae from subtropical Brazil [59]. Extracts from two

20

J. W. Daly, H. M. Garraffo and T. F. Spande

populations oi Melanophryniscus stelzneri from temperate Argentina had only minor/trace amounts of pumiliotoxin 251D, while a population from Uruguay had major amounts [86]. Pumiliotoxins are major alkaloids in skin extracts from nine species of Madagascan frogs of the mantelline genus Mantella, while an allopumiliotoxin is a major alkaloid in extracts from only three of the nine species [56,61]. Pumiliotoxins occur as major alkaloids in skin extracts from certain populations of four of seven species of Australian frogs of the myobatrachid genus Pseudophryne [25]. An allopumiliotoxin occurs as a major alkaloid in only one species. Pumiliotoxin B (323A) and allopumiliotoxin 323B are the most widely distributed of their classes, being detected in dendrobatid frogs from Central and South America, bufonid toads from subtropical/temperate South America, mantelline frogs from Madagascar and myobatrachid frogs from Australia. Captive-raised dendrobatid or mantelline frogs do not contain pumiliotoxins/allopumiliotoxins, but can accumulate such alkaloids into skin when provided in the diet [5,7,8]. A natural dietary source of pumiliotoxins/allopumiliotoxins is not known. Unlike most of the bicyclic alkaloids detected in extracts of frog skin, the pumiliotoxins/allopumiliotoxins cannot derive biosynthetically from a straight chain precursor. Instead, isoprenoid units are required. Several of the simpler bicyclic alkaloids probably are derived from dietary ants (see Sections 3.4, 3.5, 3.6.1, 3.7.1), but that appears less likely for the isoprenoid-containing pumiliotoxins/allopumiliotoxins. Whatever the putative dietary source, it must be widely distributed in tropical/subtropical regions of the world, since pumiliotoxins/allopumiliotoxins are present in dendrobatid frogs of tropical Central and South America, in bufonid toads of subtropical/temperate southeastern South America, in mantelline frogs of Madagascar and in myobatrachid frogs of Australia. Pumiliotoxins and an allopumiliotoxin are major alkaloids in a dendrobatid frog {Dendrobates auratus) that was introduced from Panama into Hawaii in 1932 [5]. Although the distribution of the putative small arthropods, serving as dietary source of pumiliotoxins/allopumiliotoxins, appears to be world-wide, it also appears that such arthropods do not occur uniformly in the tropics, since different populations of the same species, in some instances populations at different sites on a small island, can contain levels of pumiliotoxins/allopumiliotoxins ranging from none to major. Thus, only certain habitats appear suitable for the putative dietary source. The prey item, containing pumiliotoxins and allopumiliotoxins, must be very small in size, since minute frogs of the genus Minyobates contain pumiliotoxins/allopumiliotoxins as major alkaloids and such miniature frogs eat only arthropods of fruit fly-size or smaller. Arboreal dendrobatid frogs contain pumiliotoxins/allopumiliotoxins as major alkaloids and hence the putative prey item must occur not only in leaf-litter, but also high in trees of the rain forest. Finally, pumiliotoxins/allopumiliotoxins occur in skin extracts of the nocturnal myobatrachid frogs {Pseudophryne) and hence the prey items must be available both during the day when the diurnal anurans of the family Dendrobatidae and the bufonid genus Melanophryniscus and the mantelline genus Mantella are active and during the night when nocturnal frogs of the myobatrachid genus Pseudophryne are active. Captive-raised dendrobatid frogs maintained in outside enclosures in Hawaii had the 3,5-disubstituted indolizidine 195B, precoccinelline (193C), decahydroquinoline 195A and a pumiliotoxin/allopumiliotoxin pair, 251D and

Alkaloids from Amphibian Skins

21

267A, in varying amounts dependent on the species and each frog's history [5]. The indolizidine 195B occurred along with decahydroquinoline 195A and the pumiliotoxin/ allopumiliotoxin pair in Dendrobates auratus and D. leucomelus, but was the only alkaloid in Phyllobates aurotaenia. These frogs were fed mainly wild-caught termites and fruit flies, but also occasionally leaf-litter insects. In addition, small insects would have had access into the outside enclosures. Presumably 195B originates from myrmicine ants. The discovery of a prey item containing pumiliotoxins/allopumiliotoxins remains a major challenge. Activity. Pumiliotoxins A and B are relatively toxic substances with a minimum lethal dose for mice of about 2 mg/kg [50]. Simpler pumiliotoxins, such as allopumiliotoxin 267A, are relatively nontoxic to mammals [see ref. 10], but not so towards insects where pumiliotoxin 251D is quite toxic [85]. Insecticidal activities against tobacco budworms (Heliothis virescens) for pumiliotoxin 251D and thirteen synthetic analogs have been reported [85]. Both the initial convulsant activity and the toxicity were measured with the latter probably influenced by metabolic deactivation of the pumiliotoxin. Pumiliotoxin analogs with the natural 8S,11R configuration (see Figure 3) were in all cases more active than diastereomers epimeric at one or the other of these positions. An isomer of pumiliotoxin 251D with the 8R,1 IS configuration was inactive. Lengthening the side-chain of 25ID by two carbons had no effect on activity, while lengthening it by four carbons reduced activity. The analogs that lacked the 11-methyl or had an W-gem dimethyl structure were, respectively, markedly or modestly less active than 251D. An analog in which the 11-methyl of 251D was replaced with a butyl group was only somewhat less active than 251D. Remarkably, pumiliotoxins A (307A) and B (323A), which are quite toxic for mammals [50], were stated to be ca. 20-fold less active than 251D in the budworm assays [85]. The molecular pharmacology of the pumiliotoxins/allopumiliotoxins remains incompletely defined. Pumiliotoxin B was initially shown to have myotonic and cardiotonic activity [87]. In 1985, structure-activity relationships for the cardiotonic activity of natural pumiliotoxins and two synthetic analogs were reported [84]. Pumiliotoxin B (323A) was the most potent, followed by allopumiliotoxins 323B, 339A, 339B and pumiliotoxin A (307A). Pumiliotoxins without a side-chain hydroxy 1 group, such as 25ID and a synthetic analog with a 6'-hexenylidene side chain, and a synthetic analog with a 7'-hydroxyheptylidene sidechain, were cardiac depressants. Electrophysiological studies with neuromuscular preparations suggested that enhanced mobilization of calcium from internal stores was involved in the myotonic activity [88,89, see also review in 10]. Pumiliotoxin B caused repetitive neuronal firing in neuromuscular preparations [88-90], in sympathetic neurons [91], and in hippocampal preparations [92]. Remarkably, pumiliotoxin B, while causing repetitive firing in normal chick muscle, had little or no effect in muscle from dystrophic chicks [89]. Apparent inhibitory effects of pumiliotoxin B preparations on calcium-dependent ATPase of sarcoplasmic reticulum [93] were later found to be due to phenolic impurities [94]. Pumiliotoxin B was then found to stimulate influx of sodium ions through voltage-dependent sodium channels [95] and the stimulatory effects of pumiliotoxin B in neuromuscular preparations were reinterpreted as due primarily to effects on sodium channels [90]. Pumiliotoxin B, like

22

J. W. Daly, H. M. Garraffo and T. F. Spande

the sodium channel activators batrachotoxin, veratridine and aconitine, had been shown to stimulate phosphoinositide breakdown in synaptoneurosomes, an effect blocked by tetrodotoxin [96,97]. Pumiliotoxin B was later reported to alter rates of opening and closing of sodium channels in hippocampal neurons, resulting in spontaneous firing [92]. The stimulatory effects of pumiliotoxin B on sodium influx in cultured cells were blocked by tetrodotoxin and enhanced by other sodium channel modulators, namely a-scorpion toxin, pscorpion toxin and brevetoxin [95]. The effects of pumiliotoxin B on mobilization of calcium might be secondary to influx of sodium and, indeed, pumiliotoxin B and various congeners stimulate phosphoinositide breakdown in brain and heart preparations [83,96100], which would lead through inositol trisphosphate to release of calcium from intracellular stores. The stimulation of phosphoinositide breakdown by pumiliotoxin B was blocked by tetrodotoxin, but appeared in brain to involve sodium channels relatively resistant to tetrodotoxin [97]. Structure-activity relationships for pumiliotoxins/allopumiliotoxins for sodium influx, phosphoinositide breakdown and cardiotonic activity have been defined and correlated [83,84,99,100]. Three hydroxy 1 groups appeared critical for high activity in all three assays. Two of the required hydroxyl groups could be provided from the side-chain, as in pumiliotoxin B (323A), or by the ring hydroxyl groups, as in allopumiliotoxin 323B (Figure 3). The configuration at the side-chain hydroxyl groups was critical for pumiliotoxin B, with the natural 15,16-threo isomer (15R,16R) having high activity, while a natural l5,\6'erythro isomer (15S,16R) (IS-epi) and a synthetic \5,\6'threo isomer (15S,16S) (15epi-\6'epi) had much lower activity. Synthetic analogs of pumiliotoxin B lacking the 11methyl or having an 1 \-epi structure were less active and inactive, respectively [83]. Certain of the simpler pumiliotoxins, such as 251D, appeared to antagonize activation of sodium channels. Pumiliotoxin B, like several other alkaloids, can enhance binding of the L-type calcium channel blocker nitrendipine [101]. A pumiliotoxin, present in an alkaloid fraction purified from extracts of the Australian myobatrachid frog Pseudophryne coriacea, was estimated to be much more potent than pumiliotoxin B [102 and ref. therein]. However, only pumiliotoxin B was ultimately isolated from the active fraction [55]. It is possible that factors that enhance the activity of pumiliotoxin B had been present [see ref 25], or that a highly active congener had converted to pumiliotoxin B on storage. The stimulatory effects of pumiliotoxin B on sodium channels have now been proposed to be due to interaction with a subdomain of the modulatory site on such channels at which batrachotoxin, veratridine, and aconitine act [103]. Thus, pumiliotoxin B did not affect binding of batrachotoxinin A [^H]benzoate, but antagonized the ability of aconitine to inhibit the binding. The pumiliotoxins/allopumiliotoxins represent yet another class of agents that modulate function of voltage-dependent sodium channels. Further definition of sites of action and possible development of clinically useful myotonic and cardiotonic agents remain as challenges for further research.

23

Alkaloids from Amphibian Skins

3.1.1. 8-Deoxypumiliotoxins and Other Pumiliotoxin Congeners 8'Deoxypumiliotoxins. A minor alkaloid (251H) in skin extracts of the Ecuadoran poison frog, Epipedobates tricolor, was recently isolated [104]. NMR spectral analysis of the one milligram isolated, defined the structure as that of an 8-deoxypumiliotoxin (Figure 5).

193H

251H

291E 8,sOH

249G Tentative

281B

293D HO

281F Tentative

Figure 5. Structure of 8-deoxypumiliotoxin 251H and tentative structures for other deoxypumiliotoxins, an 8-desmethylpumiliotoxin 249G and a 6,10-dihydropumiliotoxin 281F. Alkaloid 251H is the 8-deoxy analog of pumiliotoxin 267C. The configuration of the 14hydroxyl group is unknown in both alkaloids. The EI-MS of 251H shows a base peak of CioHieN"*" (m/z 150) and a significant peak of C4H8N"^ (m/z 70), consonant with the structure (see Scheme 1). The vapor-phase FTIR spectrum of 251H has been reported [104] and is similar to spectra of pumiliotoxins, both in the Bohlmann band region and the finger-print region, including an absorption at 965 cm"^ for the 6,10-trisubstituted double bond. The

24

J. W. Daly, H. M. Garrafib and T. F. Spande

absorption at 3544 cm'^ for the hydrogen-bonded 8-hydroxyl group of pumiliotoxins is missing. Five other alkaloids have been tentatively assigned to the 8-deoxypumiliotoxin subclass, based mainly on characteristic EI-MS with a base peak at m/z 150 and a significant fragment ion at m/z 70 [61 and unpublished]. Structures of the putative 8-deoxypumiliotoxins 193H, 281B, 291E and 293D are proposed in Figure 5. The vapor-phase FTIR spectrum of 291E is shown in Figure 6. The structures of the side-chains are based in part on analogy to the parent pumiliotoxins 307A and 309A (see Figure 3). A structure for the fifth putative 8-deoxypumiliotoxin 281G is not proposed because of insufficient data. Properties of the 8-deoxypumiliotoxins are tabulated in the Appendix. 8-Deoxypumiliotoxins have been detected only rarely in frog skin extracts as minor or trace alkaloids. Such alkaloids have been detected in a population of the Ecuadoran Epipedohates tricolor, a population of the Panamanian Dendrobates pumilio, an undescribed Panamanian Dendrobates species, a population of Costa Rican Phyllobates vittatus, and several species of mantelline frogs of the Madagascan genus Mantella.

-

2967.37

.008-

'*^36.32

/"V^'^.-^ ^"^^V^^^ OH

N'^

7883.91 I 1

.004-

2788.65

291E

2734.14 1461.21

1 0-

>Ay/\/v^^,AA/^-^J^K

VV.VJV^'^-A''"^"*'*^^

3000

1380.1

li

970.796

1301.9

A A AA, A

^--^

11 ^V

\J\JwJ^

2600 2200 Wavenumber (cm-1)

Figure 6. Vapor-phase FTIR spectrum of 8-deoxypumiliotoxin 291E. Other Pumiliotoxin Congeners. Two other putative subclasses of pumiliotoxin congeners have been or can be proposed, based in both cases on a single alkaloid. Alkaloid 249G can be proposed to be an 8-desmethylpumiliotoxin based mainly on mass spectral fragmentation to two major ions of C9Hi4NO'' (m/z 152) and C4H8N'^ (m/z 70) (see Scheme 1). The tentative structure of 249G is shown in Figure 5. It has been detected only in a skin extract from a Peruvian dendrobatid frog, D. variabilis (unpublished results). In the homopumiliotoxin class, there are three alkaloids that appear to be 9-desmethylhomopumiliotoxins (see

Alkaloids from Amphibian Skins

25

Section 3.2). A 6,10-dihydropumiliotoxin subclass has been proposed to be represented by alkaloid 281F, whose tentative structure [56], shown in Figure 5, is based on spectral properties, in particular a comparison of the vapor-phase FTIR spectrum [56] with the spectrum of the 6,10-dihydro derivative of pumiliotoxin 267C, and an ion-trap pseudo-EIMS fragmentation yielding a base peak of C4H8N"*' (m/z 70) and a major fragment of CsHioN"^ (m/z 84). Later, EI-MS analyses of 281F afforded a more complex fragmentation; the base peak was at m/z 70, but the m/z 84 fragment ion was minor (unpublished data). Thus, the proposed structure of 281F must be considered tentative. Alkaloid 281F has been detected only in populations of the Madagascan frog Mantella baroni [56,61]. Properties of 249G and 281F are tabulated in the Appendix.

3.2.

Homopumiliotoxins

The structure of alkaloid 223G, the parent member of the homopumiliotoxin class of frog skin alkaloids, was proposed in 1987, based on EI-MS and NMR spectral analyses [105]. The structure of 223G and tentative structures for nine other homopumiliotoxins are shown in Figure 7. A structure is not proposed for a putative homopumiliotoxin 317 for which a satisfactory FTIR could not be obtained [86]. Properties of the eleven alkaloids proposed to be homopumiliotoxins are tabulated in the Appendix. The EI-MS fragmentation is diagnostic for homopumiliotoxins with major ions of CiiHigNO^ (m/z 180) and CsHioN"" (m/z 84) (see Scheme 2). The vapor-phase FTIR spectra of 223G and 319B have been reported [56,86] and the FTIR spectrum of 265N is depicted in Figure 8. The FTIR spectra of homopumiliotoxins show a characteristic Bohlmann band pattern with an absorption peak near 2750 cm"* and a shoulder near 2800 cm'^ and an absorption for a hydrogen-bonded 9-hydroxyl group near 3555 cm'^ In an earlier review [11], alkaloids 207H, 235J, 249F, 251L and 251L-0-acetate (now 293G) were included, tentatively, in the homopumiliotoxin class even though the mass spectra exhibited only the diagnostic m/z 84 fragment and not the diagnostic m/z 180 fragment. The vapor-phase FTIR spectra were stated to be consonant with a homopumiliotoxin structure [56]. In the present review, these alkaloids, for which homopumiUotoxin structures consonant with the mass spectra cannot be formulated, are reported as unclassified alkaloids in the Appendix. Alkaloid 207H always occurs in skin extracts with alkaloid 207G. The former shows m/z 178 and m/z 84 fragments, while the latter shows the m/z 180 and the m/z 84 fragments, characteristic of homopumiliotoxins. Alkaloids 207G and 207H are closely related in structure and are reported as unclassified alkaloids in the Appendix. Synthesis. Homopumiliotoxin 223G has now been synthesized [81]. The stereoselective route began with 2S-pipecolic acid in order to arrive at a (+)-homopumiliotoxin 223G with the same absolute stereochemistry in the ring as that found in pumiliotoxins/allopumilio-

J. W. Daly, H. M. Garraffo and T. F. Spande

26

HO'

223G

239M

251R

265N

'OH

319A

319B

321B

323E

335

337

Figure 7. Structures of homopumiliotoxins. toxins. The synthesis involved an intramolecular palladium-catalyzed carbonylationcyclization of an intermediate piperidinyl vinyl iodide. Synthetic 223G was identical in spectral properties with natural 223G, thereby confirming the structure and the relative

27

Alkaloids from Amphibian Skins

Stereochemistry. The optical rotation of the hydrochloride salt was [a]^^+48.1° (c = 0.51, CH3OH) (C. Kibayashi, personal communication, 1997). Unfortunately, a rotation had not been determined for the natural compound [105].

m/z180 180inhomoPTXs 164 in 9-deoxyhomoPTXs 166 in 9-desmethylhomoPTXs

m/z84 84 in homoPTXs 84 in 9-deoxyhomoPTXs 84 in 9-desmetliylhomoPTXs Scheme 2

"OH 1455.81

265N

1386.54 1322.36 1271.99 1165.85 1120.74 ^1059.46 853.4911

X 2600 2200 Wavenimtfjer (cm-1)

Figure 8. Vapor-phase FTIR spectrum of homopumiHotoxin 265N.

idoo

28

J. W. Daly, H. M. Garraffo and T. F. Spande

Occurrence. Homopumiliotoxins have been detected relatively rarely in extracts of frog/ toad skin. Homopumiliotoxin 223G v^as first detected in the dendrobatid species Dendrobates pumilio [105] and has subsequently been detected in two further neotropical dendrobatid species and three species of Madagascan frogs of the mantelline genus Mantella [11, 56,61]. Homopumiliotoxins 251R and 265N have been detected only in skin extracts from D. lehmanni and Mantella baroni [61 and unpublished results]. The remaining homopumiliotoxins have been found only in one population [86] of the bufonid toad Melanophryniscus stelzneri (319A, 319B, 321B) or in mantelline frogs [56,61] of the genus Mantella (239M, 317, 321B, 323E, 335, 337). A dietary source for homopumiliotoxins is unknown. Activity. The biological activities of homopumiliotoxins have nbt been examined. Because of close similarities in structure to the pumiliotoxins/allopumiliotoxins, it is probable that homopumiliotoxins will also prove to be myotonic and cardiotonic due to enhancement of sodium channel function.

3.2.1. Homopumiliotoxin Congeners 9'Deoxyhomopumiliotoxins and 9-Desmethylhomopumiliotoxins. Two apparent subclasses of homopumiliotoxins, which correspond to similar subclasses of pumiliotoxins, have been detected in extracts of frog skin. A tentative 9-deoxyhomopumiliotoxin structure can be proposed for alkaloids 193F and 207O (Figure 9), based on EI-MS fragmentation yielding a base peak of CnHigN"^ (m/z 164) and a significant fragment peak of CSHIQN"^ (m/z 84) (see Scheme 2). The 9-deoxyhomopumiliotoxins 193F and 207O, the latter of which would correspond to the 9-deoxy analog of homopumiliotoxin 223G, have been detected in one Costa Rican population of Dendrobates pumilio (unpublished results). 9-Deoxyhomopumiliotoxin 207O has also been detected in Mantella viridis of Madagascar. Properties of 193F and 207O are tabulated in the Appendix. Three alkaloids, all from Madagascan frogs of the genus Mantella, can be proposed to be members of a 9-desmethylhomopumiliotoxin subclass. These are 209H, 267N and 323C. Tentative structures are shown in Figure 9. Formerly such alkaloids were proposed to represent an undefined "isopumiliotoxin" class [61], but the 9-desmethylhomopumiliotoxin structure is completely consonant with the observed properties of these alkaloids. The mass spectra have a base peak of CioHi6NO'^ (m/z 166) and a significant fragment ion of CSHIQN"^ (m/z 84) (see Scheme 2). The EI-MS of 339C, also formerly termed an "isopumiliotoxin" [61], is not readily explained by a 9-desmethylhomopumiliotoxin structure and it is now reported as an unclassified alkaloid in the Appendix. The vapor-phase FTIR spectra of the putative 9-desmethylhomopumiliotoxins show a Bohlmann band pattern similar to that of homopumiliotoxins, an absorption peak near 3563 cm'^ apparently owing to a hydrogen-bonded 9-hydroxyl group, and a sharp absorption peak near 1111 cm"^ (unpublished results with N.R. Andriamaharavo). Properties of

Alkaloids from Amphibian Skins

29

alkaloids of the proposed 9-desmethylhomopumiliotoxin subclass are tabulated in the Appendix.

193F

207O 9,xOH

209H

.NOH

267N

323C

Figure 9. Tentative structures for 9-deoxyhomopumiliotoxins (193F, 207O) and 9desmethylhomopumiliotoxins (209H, 267N and 323C).

"Dehydrohomopumiliotoxins ". A set of three alkaloids, exhibiting on EI-MS analysis a large M"*"-! fragment, a base peak of CnHi^N"^ (m/z 162), and a significant fragment ion of CUHMN"^ (m/z 160), was detected in extracts of the Madagascan mantelline frogs Mantella aurantiaca and M crocea [56]. These alkaloids, 22IF, 233F and 235C, were tentatively proposed to represent an 8,9-dehydrohomopumiliotoxin subclass. The proposed tentative structures are shown in Figure 10. A fourth alkaloid, 251G, detected in an earlier study [59], also belongs to this set of alkaloids, apparently being a hydroxylated analog of 235C. The major alkaloid 235C of the putative 8,9-dehydrohomopumiliotoxin subclass was found to represent a 2:1 mixture of diastereomers [56]. The proposed tentative structure of 235C is consonant with chemical (exchange of hydrogen, hydrogenation, acetylation) and most of the spectral (MS, FTIR) properties [56]. The EI-MS of 235C exhibits major M'"-l and M""C4H9O fragments as would be expected of the proposed structure. The FTIR of 235C has been reported [56]. It exhibits absorption peaks at 3650 cm"^ typical for a non-hydrogenbonded hydroxyl group, 3020 cm"^ indicating an internal double bond, and 939 cm'\

J. W. Daly, H. M. GarrafTo and T. F. Spande

30

221F Tentative

235C

Tentative

OH

233F Tentative

251G Tentative

Figure 10. Tentative structures for alkaloids of a putative 8,9-dehydrohomopumiliotoxin subclass [56]. consonant with a trisubstituted double bond as part of a conjugated diene system. A Bohlmann band at 2792 cm"^ is w^eak. The very weak Bohlmann band of 235C would not be expected for the proposed 8,9-dehydrohomopumiliotoxin structures. A diastereomeric mixture of tetrahydro-derivatives was obtained on catalytic reduction of 235C [56]. Earlier results had indicated that only a dihydro-derivative was formed [59]. Until sufficient material is isolated for NMR spectral analysis, the putative 8,9-dehydrohomopumiliotoxin subclass must be considered as very tentative. None of these alkaloids has been isolated for UV spectral analysis, all analyses having been done by GC on mixtures. These alkaloids possibly represent analogs of alkaloid 245F, which in the Appendix is postulated to be a 6,7dehydro-5,8-disubstitutedindolizidine. Another alkaloid, 265F, was proposed to be related in structure to 235C, based on EIMS fragmentation [56]. Alkaloid 265F yielded a base peak of Ci2H2oNO"^ (m/z 194) and a significant fragment ion of CnHigNO"^ (m/z 192), corresponding to the pair of peaks at m/z 162 and 160 for 235C. However, the large M"^-l fragment of 235C was absent in 265F. The EI-MS of the 0-acetyl derivative of 265F indicated the presence of a hydroxy 1 group in the major fragment ion of 265F, since the base peak was shifted from m/z 194 to m/z 236 on acetylation. The other oxygen was present in a side-chain, but apparently not as a ketone or a hydroxyl group, based on the FTIR spectrum. That oxygen is probably present as an ether. The infrared spectrum of 265F indicated a single non-hydrogen-bonded hydroxyl group (3656 cm"^), and showed a weak Bohlmann band at 2803 cm'^ Only a dihydro derivative was obtained on hydrogenation. Alkaloid 265F is tabulated as unclassified in the Appendix,

Alkaloids from Amphibian Skins

31

but may be related to 245F, another unclassified alkaloid, which in the Appendix is postulated to be a member of a 6,7-dehydro-5,8-disubstituted indolizidine subclass. The properties of the four members of the postulated 8,9-dehydrohomopumiliotoxin subclass (Figure 10) are tabulated in the Appendix. Such alkaloids have been detected only from two swampdwelling mantelline species of Madagascar, suggesting that the dietary source is also limited to swamp habitats.

3.3.

Histrionicotoxins

Histrionicotoxin and isodihydrohistrionicotoxin were isolated in 1971 as the two major alkaloids from skin extracts from about 800 specimens of a medium-size dendrobatid frog, Dendrobates histrionicus, collected from a particularly abundant population in southwestern Colombia [106]. Such alkaloids had been previously detected in the late sixties in skin extracts from a D. histrionicus population in the Rio San Juan drainage to the north in Colombia, a population that was microsympatric with the batrachotoxin-containing poisondart frog Phyllobates aurotaenia, but not particularly abundant. The structures of histrionicotoxin and isodihydrohistrionicotoxin were determined by X-ray crystallography [106]. Further histrionicotoxins were isolated after later collections from the same site in southwestern Colombia and the structures determined by MS and NMR spectral analyses [107109]. The structures of the sixteen known histrionicotoxins are shown in Figure 11. All have the same 8-hydroxy-l-azaspiro[5.5]undecane ring system and differ only in the length and nature of unsaturation of the side-chains. The side-chain at C-2 has either three or five carbons, while the side-chain at C-7 has either two or four carbons, resulting in histrionicotoxins with fifteen, seventeen or nineteen carbons. The mass spectra of histrionicotoxins have three diagnostic fragments [107,110]. One results from a-cleavage of the side-chain at C-2. The second results from a-cleavage at the spiro-junction (C-6) and loss of all but one carbon of the cyclohexyl ring. The third diagnostic ion, often the base peak, is C6HioN^ (m/z 96), present in the EI-MS of all histrionicotoxins. Fragmentation pathways for histrionicotoxins are proposed in Scheme 3. The EI-MS of octahydrohistrionicotoxin (291 A) is somewhat different in yielding a major fragment ion at m/z 250, corresponding to loss of a propenyl group, while a fragment ion at m/z 178, rather than m/z 96, is the base peak. Fragmentation pathways for 291A are proposed in Scheme 4. The identification of individual histrionicotoxins when several are present together in an extract often presents a major challenge, which can be approached through GC-EI-MS and GC-FTIR spectral analysis. The butylboronate derivatives offer advantages for such GC analyses [110]. Certain artefacts formed from histrionicotoxins have been detected in extracts, including formaldehyde condensation products and a trans-diene isomer (tranS'2S3A) presumably formed by photoisomerization [see ref 110]. Vapor-phase FTIR spectra of such trans-isomQVS have been presented [110].

J. W. Daly, H. M. Garraffo and T. F. Spande

32

(-)-285B NeodihydroHTX (-)-285A * IsodihydroHTX

(-)-285C AllodihydroHTX

287A IsotetrahydroHTX (-)-285E DihydroHTX

287B TetrahydroHTX

291A OctahydroHTX 287D AliotetrahydroHTX

Figure 11. Structures of histrionicotoxins(HTX). * Absolute configuration as shown. The configuration is presumably the same for all histrionicotoxins, since all have been levorotatory.

33

Alkaloids from Amphibian Skins

a-cleavage at C-2 ,NH +•

a-cleavage atC-6

OH

^/xZ^-CT/

"

k^NH

a-cleavage at C-2 R

NH

a-cleavage at C-2 H

OH

(M - Rf

m/z96 (generally base peak)

(M-QHidORr

(M-CsHeOnr Scheme 3 Vapor-phase FTIR spectra of twelve of the natural histrionicotoxins have been published [110]. A vapor-phase FTIR spectrum of 239H is presented in Figure 12. The vapor-phase FTIR spectra of histrionicotoxins do not show a Bohlmann band, but do show a diagnostic absorption at 3330-3400 cm"^ for an 8-hydroxyl group that is strongly hydrogenbonded to the nitrogen, and diagnostic absorptions for terminal allenic, terminal acetylenic and terminal and internal olefmic groups. Brief summaries with references for physical (optical rotation) and spectral (UV, IR, MS and NMR) properties of histrionicotoxins have been provided [10,11]. The properties of the histrionicotoxins are tabulated in the Appendix.

J. W. Daly, H. M. Garraffo and T. F. Spande

34

a-cleavage at C-2 2^NH

OH

^/\/-J-^

^ ^^N"

m/z 222

O"

291A

a-cleavage atC-6

./\/"tir/

^

L

a-cleavage at C-2 NH

NH

OH

m/z 250

m/z 96

+

m/z 274

m/z 178 (base peak) (m/z 180, base peak, in perhydroHTX) Scheme 4

Synthesis. The early syntheses were directed towards perhydrohistrionicotoxin, which does not occur naturally. The extensive synthetic work in many laboratories towards perhydrohistrionicotoxin has been reviewed in detail [10-12,111,112]. Most of the early work focused on construction of the l-azaspiro[5.5]undecane system and then elaboration of the 8-hydroxyl group and the alkyl side-chains. Various azaspiro lactams, oximes and olefins [see ref. 10] represented key intermediates in the syntheses of racemic perhydrohistrionicotoxin and racemic 2-despentylperhydrohistrionicotoxin. The "Corey-Kishi lactam", namely 7-«-butyl-8-hydroxy-2-keto-l-azaspiro[5.5]undecane, was a key intermediate [see ref. 10]. It was resolved and one enantiomer afforded (-)-perhydrohistrionicotoxin with the same configuration as natural histrionicotoxin, while the other gave the unnatural (+)-perhydrohistrionicotoxin [113,114]. Various derivatives and analogs of perhydrohistrionicotoxin have been prepared [see ref. 10], including the 7,8-dehydro derivative [109], 8-deoxyperhy-

35

Alkaloids from Amphibian Skins

drohistrionicotoxin [107,109], A^-methylperhydrohistrionicotoxin [107], and 7-desbutyl and 2-despentyl analogs [115,116]. A^-Methylhistrionicotoxin has been prepared [117].

3396.96

1461.33 1342.72 1216.73 1131.82 1031.23

1^73.52

26'00 ^ 2200 Wavenumber(cm-I)

Figure 12. Vapor-phase FTIR spectrum of histrionicotoxin 239H. In 1963, Kishi and colleagues reported the synthesis of racemic octahydrohistrionicotoxin using an intramolecular Michael-addition of an amide to form the 7-butenyl analog of the "Corey-Kishi lactam" [118]. In 1985, utilizing the same 7-butenyl "Corey-Kishi lactam" as a starting point, Kishi and colleagues were successful in the synthesis of racemic histrionicotoxin [119]. A 7-desbutyl analog of the "Corey-Kishi lactam" has been reported [120]. An enantioselective synthesis of a functionalized l-azaspiro[5.5]undecane system suitable for elaboration of natural histrionicotoxins was based on photolysis of a dioxolenone derived from (+)-glutamic acid [121]. In 1990, utilizing an enantiospecific route involving an ally lie epoxide cyclization to intermediate lactones, followed by a second cyclization yielding the l-azaspiro[5.5] undecane system, Stork and Zhao reported the syntheses of (-)-histrionicotoxin (283 A) and (-)-235A [122]. We are not aware of any other further published synthetic work in this area. Occurrence. Histrionicotoxins are known to occur in Nature only in the skin of neotropical dendrobatid frogs. Three histrionicotoxins, namely histrionicotoxin (283A), isodihydrohistrionicotoxin (285A) and allodihydrohistrionicotoxin (285C) represented the major alkaloids in most populations oiDendrobates histrionicus from western Colombia [1,123]. In one population from northwestern Ecuador, octahydrohistrionicotoxin (291 A) replaced those alkaloids and represented the major skin alkaloid, while in certain populations in Colombia on the lower Rio San Juan, the nineteen-carbon histrionicotoxins were replaced

36

J. W. Daly, H. M. Garraffo and T. F. Spande

by the fifteen- and seventeen-carbon histrionicotoxins 235A and 259A (unpublished results). Histrionicotoxins have not been detected in skin extracts from a very closely related Colombian species D. lehmanni and the lack of histrionicotoxins and presence of alkaloid 275A (see section 3.8), now identified as a l-azabicyclo[5.3.0]decane, were two considerations in proposing D. lehmanni as a separate species, distinct from the extremely variable (with respect to skin color and pattern) populations of D. histrionicus [123]. Histrionicotoxins were present as major alkaloids in skin extracts from other dendrobatid frogs including eleven species oi Dendrabates,fivespecies of Epipedobates and one species ofPhyllobates [1 and unpublished results]. Histrionicotoxins were absent in skin extracts from the tiny dendrobatid frogs of the genus Minyabates. They were absent or only trace alkaloids in two arboreal Dendrobates species, and in two species, D. lehmanni and D. speciosus, which occur at elevations greater than 1000 meters; i.e., elevations higher than those of dendrobatid frogs that contain major amounts of histrionicotoxins. Certain populations of the small dendrobatid frog D. pumilio contained histrionicotoxins as major alkaloids, while other populations contained none or only trace amounts. Extracts from skins of a population of the dendrobatid frog D. auratusfromIsla Taboga, Panama, had histrionicotoxins as significant alkaloid components, but the descendents of 200 of these frogs introduced into Hawaii in 1932 had no histrionicotoxins in their skin, and instead had a pumiliotoxin and an allopumiliotoxin as major alkaloids [5]. Histrionicotoxins have not been detected in skin extracts of bufonid toads of the subtropical/temperate South American genus Melanophryniscus [86], nor in extracts of wild-caught Madagascan frogs of the mantelline genus Mantella [56,61]. They were not detected in skin extracts of Australian frogs of the myobatrachid genus Pseudophryne. Histrionicotoxins are presumably derived unchanged from a dietary arthropod, like the other so-called "dendrobatid" alkaloids of frog skin. Based on the occurrence in frog skins, it would appear that the prey item occurs only in tropical regions of Central and South America, and is not abundant or perhaps absent at higher elevations and in arboreal habitats. In addition, it would appear that the unknown arthropod that provides histrionicotoxins is not uniformly available even in similar lowland habitats. For example, it is likely to be absent as a significant prey item for some, but not all populations of Z). pumilio from lowlands of Costa Rica and Panama. As yet, no arthropod that contains histrionicotoxins has been discovered. It should be noted that the nineteen-carbon histrionicotoxins 283A, 285 A and 285C are often accompanied in skin extracts of dendrobatid frogs by nineteencarbon decahydroquinolines, particularly 269AB (see Section 3.4). 2,5-Disubstituted decahydroquinolines have recently been discovered in myrmicine ants [124,125]. It appears possible that myrmicine ants v^U prove to be the source not only of the decahydroquinolines, but also the histrionicotoxins. Alternate pathways from a 2,6-disubstituted piperideine could lead in ants to both alkaloid classes [see ref. 124]. Histrionicotoxins are readily accumulated in skin when dendrobatid frogs are provided with fruit flies dusted with powder containing histrionicotoxins [5,7]. The histrionicotoxin 285A, however, did not appear to readily accumulate from dusted fruit flies into a mantelline frog [8]. In experiments in Panama, nineteen-carbon histrionicotoxins 283A, 285A,

Alkaloids from Amphibian Skins

37

285C and 287A and decahydroquinoline 269AB were present as minor alkaloids in the dendrobatid frogs (D. auratus) that had been reared for seven months to adulthood in terraria and provided during that period with arthropods, obtained using Berlese funnels from leaf-litter of a mainland site where D. auratus occurs [6]. The identity of the leaf-litter arthropod providing those histrionicotoxins remains unknown. In a subsequent experiment, frogs were raised in large, screened, outdoor cages, repeatedly supplied with leaf-litter from the same mainland site, but no histrionicotoxins or decahydroquinolines were detected in their skin extracts (unpublished results with A.S. Rand and C. Jaramillo). Ants with nineteen-carbon histrionicotoxins or decahydroquinolines have not been found at either the mainland leaf-litter site or the island site of the parental population of the frogs (unpublished results with A.S. Rand and C. Jaramillo). Wild-caught frogs from the mainland collection site had the nineteen-carbon histrionicotoxins as major alkaloids, while wild-caught frogs from the parental population at the island site had none, but did have a seventeen-carbon histrionicotoxin as a minor alkaloid [6]. Activity. The name histrionicotoxin is a misnomer since these alkaloids have relatively low toxicities [see reviews on biological activities in refs. 10,11]. The biological activity of histrionicotoxins proved to be due to blockade of cation conductance through the channel formed by nicotinic receptors of the neuromuscular junction [126,127]. Histrionicotoxins also blocked potassium [127,128] and sodium channels [128]. The structure-activity relationships for histrionicotoxins at the three types of ion channels differed significantly [128]. For example, histrionicotoxin (283 A) was one of the two most active histrionicotoxins tested at the nicotinic and potassium channels, while being the least active at the sodium channel. Structure-activity relationships for histrionicotoxins also differed in sympathetic ganglia, which contain ganglionic-type nicotinic receptor-channels [129]. Histrionicotoxins acted as noncompetitive blockers at the muscle-type (aiPiyS pentamer) nicotinic receptor-channels of the neuromuscular junction [126-128] and medulloblastoma TE671 cells [130], and also at ganglion-type (a3P4(5) or a3p2 pentamers) of adrenal chromaffin cells [131,132], sympathetic ganglia [129] and pheochromocytoma PC 12 cells [130,133] and at central neuronal-type (a4P2 pentamer) of striatal and hippocampal neurons [134]. Allodihydrohistrionicotoxin (285C) had similar blocking potency at the muscle-type receptor of TE671 cells and the ganglionic-type receptor of PC 12 cells [133]. Histrionicotoxin blocked nicotinic receptor-channels of an insect [134]. Histrionicotoxins had virtually no effect on binding of an agonist, [^H]nicotine, to central neuronal-type nicotinic receptorchannels (unpublished results). A fourth channel, namely that of the glutamatergic NMDA receptor, was also blocked by histrionicotoxins [135,136]. Histrionicotoxins inhibited binding of batrachotoxinin A [^H]benzoate to sodium channels and binding of [^H]phencyclidine, probably to potassium channels, in brain membranes [137]. Histrionicotoxins enhanced binding of [^H]nitrendipine to L-type calcium channels [101,137]. The extensive early investigations of effects of histrionicotoxins on nicotinic receptorchannels have been reviewed in detail [10, see also ref 11]. Histrionicotoxins block the conductance of the open channel and can enhance the rate of inactivation/desensitization of

38

J. W. Daly, H. M. Garraffo and T. F. Spande

the channel. Histrionicotoxins appear to stabilize the desensitized state, which has a high affinity for agonists. They are now considered classic representatives of the structurally diverse, noncompetitive blockers of nicotinic receptor-channels. A [^H]perhydrohistrionicotoxin was developed as a radioligand for binding sites of noncompetitive blockers on the nicotinic receptor-channel [138]. [-^HlPerhydrohistrionicotoxin proved to be a useful probe for such sites, but only in membranes from the electroplax of the electric ray (Torpedo species), in which neuromuscular-type nicotinic receptor-channels occur at very high density. In membranes from brain, heart, muscle and ganglia, the densities of nicotinic receptor-channels are relatively low and nonspecific binding now overwhelms the specific binding of [^H]perhydrohistrionicotoxin to the binding sites on nicotinic receptor-channels [137]. The extensive use in the seventies and early eighties of [^H]perhydrohistrionicotoxin as a tool to study the affinity of a diverse array of compounds, including phencyclidine, quinacrine, phenothiazines, local anesthetics, and other frog skin alkaloids, at the so-called "high affinity" noncompetitive binding sites on the nicotinic receptor-channels of electroplax membranes was reviewed in detail in 1986 [10]. The effects of histrionicotoxin and twenty-two analogs on binding of [^H]perhydrohistrionicotoxin and [^H]phencyclidine to electroplax membranes were reported in 1985 [117]. The most potent was isotetrahydrohistrionicotoxin (287A). Both low affinity and high affinity sites for binding of noncompetitive blockers to nicotinic receptors have been proposed [139,140]. Ligands for the high affinity site include the histrionicotoxins, quinacrine, phencyclidine, meproadifen and certain other local anesthetics, chlorpromazine, and ethidium. All of these agents appear to interact with the histrionicotoxin-binding site. The relatively high affinity site has been referred to as the "high affinity histrionicotoxin-sensitive binding site" [141-143] and histrionicotoxins are among the most potent noncompetitive blockers of nicotinic receptorchannels. Fluorescent assays with quinacrine or ethidium as ligands have been developed and used to study the localization of the high affinity histrionicotoxin-sensitive sites [142, 143]; histrionicotoxins inhibited quinacrine- and ethidium-binding. Histrionicotoxin also inhibited photoaffinity-labeling of the putative noncompetitive binding site for a trimethylphenylphosphonium salt [144]. [^H]Perhydrohistrionicotoxin has continued to be used as a radioligand, in spite of the fact that it has never been commercially available. A number of studies with [^H]perhydrohistrionicotoxin have appeared since our 1986 review. Several have focused on the ability of nicotinic agonists, in particular anatoxin analogs, to enhance binding of [•^H]perhydrohistrionicotoxin to electroplax membranes [145-148]. Certain anatoxin analogs inhibited binding. The noncompetitive blocker of the glutamatergic NMD A receptor-channel dizocilpine (MK 801) was shown to inhibit binding of [^H]perhydrohistrionicotoxin to electroplax membranes, commensurate with activity as a noncompetitive blocker of nicotinic receptor-channels [149]. A wasp venom component, philanthrotoxin, inhibited binding of [^H]perhydrohistrionicotoxin to electroplax membranes, again commensurate with the activity of philanthrotoxin as a noncompetitive blocker of nicotinic receptor-channels [150]. Certain pyrethroids [151] and forskolin [152] inhibited [•^H]perhydrohistrionicotoxin binding and a variety of alkaloids, many from frog skin, inhibited [•^H]perhydrohistrionicotoxin binding to nicotinic receptor-channels of electroplax

Alkaloids from Amphibian Skins

39

membranes. Such alkaloids include decahydroquinolines [133,153,154] (Section 3.4), indolizidines [133,155,156] (Section 3.6), gephyrotoxin [133,155,157] (Section 4.1) and piperidines and pyrrolidines [133,158] (Section 5).

3.4.

Decahydroquinolines

A decahydroquinoline was isolated in 1968, along with pumiliotoxins A and B, from skin extracts from 250 specimens of a small dendrobatid frog Dendrobates pumilio found in abundance on Isla Bastimentos, Bocas Province, Panama [159]. The original name pumiliotoxin C is not only misleading, suggesting a structural relationship to pumiliotoxins A and B, but a true misnomer, since the alkaloid has very low toxicity. We now refer to the alkaloid as decahydroquinoline cz5-195A; however, the pumiliotoxin C nomenclature persists in the synthetic literature. Decahydroquinoline c/.s'-195A represents the parent member of an alkaloid class that now consists of about forty 2,5-disubstituted cis- and rmw^-decahydroquinolines. The structure of decahydroquinoline c/^-195A was revealed by X-ray crystallography [159]. The mass spectrum was dominated by a fragment of CioHigN"^ (m/z 152) owing to a-cleavage of the 2-propyl side chain. The fragment at mass 152 is also formed by loss of three carbons, C-6, C-7, C-8, from the alicyclic ring. An exchangeable hydrogen on nitrogen, detected by ND3-CI-MS, has served to distinguish frog skin decahydroquinolines from the many izidine alkaloids also found in frog skin extracts. Most of the izidine alkaloids also have mass spectra dominated by a base peak resulting from cleavage next to nitrogen (a-cleavage), but being tertiary amines have no exchangeable hydrogen. Structures for decahydroquinolines from frog skin are presented in Figure 13. The relative configuration at C-5 remains in question for many decahydroquinolines. The tranS'\9SA has been reported in only one extract, that from Peruvian Epipedobates bassleri [160]. It has also recently been detected in extracts from Peruvian populations of Dendrobates imitator and D. variabilis (unpublished results). Eleven alkaloids in addition to those depicted in Figure 13 are tentatively tabulated in the Appendix as 2,5-disubstituted decahydroquinolines. These are as follows: 209A, tentatively a ring-hydroxylated 2-allyl-5-methyldecahydroquinoline; 209J, tentatively a 5-ethyl-2-propyldecahydroquinoline; 21 IK, tentatively a ring-hydroxylated 5-methyl-2-propyldecahydroquinoline; 219C, tentatively a 5-methyl-2-pentadienyldecahydroquinoline; 219D, tentatively a 2-propargyl-5-propyldecahydroquinoline; 221C, tentatively a 5-methyl-2-pentenyldecahydroquinoline; 221D, tentatively a 2-allyl-5-propyldecahydroquinoline; 223Q, tentatively a c/^-2-methyl-5-pentyldecahydroquinoline; 223S, tentatively a 2-pentyl-5-methyldecahydroquinoline; 231E, tentatively a 2-hexenynyl-5methyldecahydroquinoline and 251A, tentatively a 2-heptyl-5-methyldecahydroquinoline. Further data are needed on these putative decahydroquinolines. X-ray crystallography and spectral (MS, FTIR, NMR) properties have led to elucidation of structures and proposal of structures [86,105,124,125,159-162]. The decahydroquinolines differ in relative stereochemistry at the chiral carbons and also differ in the length and degree of unsaturation of the side-chains at C-2 and C-5.

40

J. W. Daly, H. M. Garraffo and T. F. Spande

H-C/S-195A*

C/S-195J

frans-195 A

H-C/S-211A

C/S-223F

tranS'223F

tranS'253D

(+)-c/s-243A

(-)-frans-243A

(-)-5-ep/-frans-243A

Figure 13. Structures of decahydroquinolines. * Absolute configuration as shown.

Alkaloids from Amphibian Skins

41

C/S-245E

C/S-249D

tranS'249D

tranS'249E

r ^ ^

^ .

H H C/S-267L

5-ep/-frans-269AB

C/S-269AB

frans-269AB

frans-269A

frans-269B

C/S-271D Figure 13 (continued)

42

J. W. Daly, H. M. Garraffo and T. F. Spande

if. N H H

^^

or

^^a N H H

frans-271D

iso-S-epi-tranS'll^ D

C/S-275B

2-ep/-c/s-275B = c/s-275B'

Figure 13 (continued) Virtually all of the decahydroquinolines have either thirteen, fifteen, seventeen or nineteen carbons (Figure 13). The highly unsaturated side-chains of the seventeen- and nineteen-carbon decahydroquinolines find counterparts in the side-chains of the corresponding seventeen- and nineteen-carbon histrionicotoxins. Structures of several of the nineteencarbon decahydroquinolines have only recently been defined [124]. In addition, there are several minor isomers of the nineteen-carbon decahydroquinolines, whose structures and configurations remain unknovm [see ref 124]. The mass spectra of the decahydroquinolines are dominated by a-cleavage of the sidechain at C-2, particularly if such cleavage produces an allylic radical. If not, then a loss of a 43 amu fragment, consisting of the three ring carbons C-6, C-7 and C-8, is significant. A cleavage of the side chain at C-5 often occurs. Possible pathways are shovm in Scheme 5. Recently, NH3-CI-MS/MS spectra for collision-activated protonated parent ions have been investigated for decahydroquinolines and other bicyclic alkaloids of frog skin [163]. The fragmentation pathways for EI-MS and for CI-MS/MS are markedly different, since acleavage occurs only for the positively charged free radical molecular ion of EI-MS, and not for the coUision-activated protonated molecular ion of CI-MS/MS. The CI-MS/MS of the decahydroquinoline c/^-195A is characterized by odd-mass fragments (m/z 81,95,109) [163]. Vapor-phase FTIR spectra for ciS'219A and trans-219A are shown in Figure 14. FTIR spectra for decahydroquinolines c/5-195A, tranS'195A, c/^-195J, cis-219A, trans-219A, cis223F, trans'223¥, C/5-243A, trans-243A, 5-epi'tranS'243A, C/5-267L, C/5-269AB, trans269AB, tranS'269A, tranS'269B, C/5-271D, trans-271J), cis-215B and 2-epi-cis-275B have been presented [86,124,125,160,161 ]. Vapor-phase FTIR spectra have proven to be particularly useful in establishing the cis versus trans nature of ring junction in decahydroquinolines, and the relative configurations of hydrogens at C-2 and C-8a [86,124,160,161].

43

Alkaloids from Amphibian Skins

In the cz\y-isomers there are two significant ring conformations for the molecule, while in the tranS'isomQTS one conformation dominates. This apparently is reflected in the FTIR spectra as split peaks in the regions 1300 cm"^ and 1100 cm'^ for the c/5-isomers and single peaks in the same regions for the trans-isomeis. The configuration at C-2 relative to that at C-8a can be assigned, based on the presence (2,8aZ) or absence (2,8aQ of a significant Bohlmann band, characteristic of c/5-2,6-disubstituted piperidines (2806 cm'^ band for trans-2\9A, Figure 14). Only the relative stereochemistry at C-5 cannot be deduced from the FTIR spectrum.

(M-43f

a-cleavage

(M - Rf

cleavage at C-5 H Scheme 5 Brief summaries with references for the physical (optical rotation) and spectral (FTIR, MS and NMR) properties of decahydroquinolines have been provided [10,11]. A current report contains further NMR spectral data and assignments [124]. The properties of all the decahydroquinolines and putative decahydroquinolines are tabulated in the Appendix. Certain alkaloids (153A, 167D, 181E), tentatively proposed to be 5-monosubstituted decahydroquinolines [11], and two previously proposed 2,5-disubstituted decahydroquinolines (181D, 293A) [11] have now been tabulated as unclassified in the Appendix, since they do not give the ring cleavage and loss of three carbons expected from such decahydroquinolines. Certain alkaloids detected in frog skin extracts appear to be tetrahydro- (189) and octahydro- (193D) quinolines and apparent "dimers" (384A/384B); the "dimers" perhaps are derived as Diels-Alder adducts from hexahydro- and octahydro-quinolines [56]. These alkaloids occurred together in certain mantelline frogs of the genus Mantella, along with decahydroquinoline cz5-195A. An aromatic tetrahydroquinoline structure for 189 and an

44

J. W. Daly, H. M. Garraffo and T. F. Spande

R868.3 2805.52 2720.3 z ^-^*V._x'^^

1831.52 1639.61 1448.85

\^N^^>^^^^

2800

2400 2000 Wavenumber(cm-I)

1338.81

15?

Figure 14. Vapor phase FTIR spectra of decahydroquinolines c/5-219A and tranS'219A. enamine structure for 193D have been proposed (Figure 15), based on exchange data, EI-MS and vapor-phase FTIR spectra [56]. The "dimers" 384A/384B are perhaps derived from alkaloid 193D and a presumed, as yet undetected, hexahydroquinoline of mol. wt. 191. "Dimers" 384A and 384B occur in roughly 1:1 ratio in skin extracts that also contain 193D [56]. It is proposed [164] that these "dimers" represent Diels-Alder adducts with postulated structures, such as the "exo" structure shown in Figure 15. It is possible that they are artefacts formed during the isolation procedure. The FTIR spectra of 384A and 384B are virtua-

45

Alkaloids from Amphibian Skins

lly identical with absorptions indicating an internal double bond (3020 cm"^) and an enamine or imine moiety (1647 cm'^). The EI-MS shows one major fragment at m/z 341 corresponding to loss of a propyl group. CI-MS with NH3 or ND3 apparently is accompanied by reversal to components with mol. wts. of 191 and 193, as might be expected of such a DielsAlder adduct. Perhydrogenation yields dihydroderivatives. Acetylation fails to give a reaction. The proton NMR spectrum is consonant with the proposed structure [164]. Another "dimer", 382, was detected as a trace alkaloid. Alkaloids 189,193D, 382, and 384A/384B are tabulated in the Appendix.

189 Tentative

193D Tentative

(^Hy^^^-y^ 384A/384B Tentative Figure 15. Tentative structures for unsaturated analogs 189 and 193D of decahydroquinolines and a tentative exo-structure for one of the Diels-Alder "dimers" 384A/384B. Synthesis. Decahydroquinoline ciS'\9SA has been the target of extensive synthetic work in many laboratories, and the synthetic efforts have been reviewed in detail [10-12,165]. Diels-Alder reactions had been used in several approaches, while others used tetrahydroindanones or enamine cyclizations. There continues to be extensive synthetic work, directed primarily at decahydroquinoline cz5-195A, and the current review will merely reference such efforts. A review of the use of 2-cyano adducts of 5,6-dihydropyridinium salts and 2-cyano6-oxazolopiperidines as synthons for asymmetric syntheses of decahydroquinoline (-)-cw195A, indolizidines (-)-5Z,9Z-195B ((-)-monomorine I) and (-)-5Z,9£-223AB (Section 3.6.1), and piperidine i+ytrans-lS^i (solenopsin A) (Section 5.2) has been provided by HP. Husson [166]. A total of nearly twenty syntheses of d5-195A [167-185] have appeared since our review of 1986 [10], including some efficient and versatile routes. A variety of

46

J. W. Daly, H. M. Garraffo and T, F. Spande

approaches, some stereoselective and some enantioseiective, have been reported. The 2epimer [172,176] and the 5-epimer [185] ofcis-195A and the 2-epimer oftrans-195A [186] have been prepared. Enantioseiective syntheses have been reported for (+)-perhydro-/ra«5219A and the 2-epimer of perhydro-c/5-219A [187] and for (+)-tranS'219A [188]. Synthesis of diastereomers of C/5-195A and oftranS'195A with inverted centers at C-2 and C-5 were reported in 1983 [189], although at that time the occurrence of a tranS'195A in Nature was not known. TranS'l95A had also been obtained as a major byproduct in an early synthesis of CW-195A [190]. A diastereomer obtained as a byproduct in another early synthesis of c/5195A [191], also now appears to have been trans-195A. An intramolecular Diels-Alder approach had been used to prepare four cw-decahydroquinolines, namely 2,5-dimethyl-, 2butyl-, 2-(5-hydroxypentyl)-5-methyl- and 2,5-dipropyl-d5-decahydroquinoline [153,192]. The last corresponds to the 2-epimer of perhydro-c/5-219A. The three disubstituted decahydroquinolines had the same relative configuration. A 5-hydroxymethyl analog of cis195A has been synthesized [193]. Occurrence. 2,5-Disubstituted decahydroquinolines occur in skin extracts of neotropical dendrobatid frogs, Madagascan frogs of the mantelline genus Mantella, and toads of the bufonid genus Melanophryniscus of subtropical/temperate southeastern South America. Decahydroquinolines have not been reported in skin extracts of Australian frogs of the myobatrachid genus Pseudophryne. The distribution of the different decahydroquinolines in skin extracts of frogs/toads was recently tabulated [124]. In dendrobatid frogs only the 219A, 243A, and 269AB alkaloids occurred commonly, having been detected in each case in about twenty of fifty dendrobatid species. Decahydroquinoline ciS'\95A was less common, having been detected in only nine of the fifty dendrobatid species. The other decahydroquinolines were all relatively rare in skin, extracts of dendrobatid frogs with several having been detected in only one species or population. Decahydroquinolines can be major alkaloids in extracts from one population of a species, while being completely absent in other populations [see tabulation in ref 1]. They are uncommon in Madagascan frogs of the mantelline genus Mantella, where only ciS'\9SA has been detected, and only in three of nine species [56,61]. In one of these species ciS'\95A was accompanied by cz.s'-195J [125]. The proposed analogs 189 and 193D of c/5'-195C and the proposed Diels-Alder "dimers" 384A/384B have accompanied cw-195A in the Mantella species. The decahydroquinolines that have been detected in toads of the bufonid genus Melanophryniscus are cis- and trans223F, cis- and trans-249D, trans-249E and c/5'-275B, all of which occur together in two Argentinian populations of Melanophryniscus stelzneri [83]. No decahydroquinolines were detected in skin extracts from an Uruguayan population of Melanophryniscus stelzneri, nor in extracts from the Brazilian species Melanophryniscus moreirae. It is noteworthy that the nineteen-carbon decahydroquinolines of the 269AB group with a high degree of unsaturation in the side-chains frequently accompany the correspondingly highly unsaturated nineteen-carbon histrionicotoxins and that such nineteen-carbon decahydroquinolines, like the histrionicotoxins, occur virtually only in neotropical dendrobatid frogs. Until recently a dietary source for the decahydroquinolines of frog skin was a mystery, but now decahydroquinolines have been discovered in myrmicine ants [124,125].

47

Alkaloids from Amphibian Skins

Diastereomers of the frog skin decahydroquinolines cis-275B and trans-215B have been discovered in virgin queens of a myntiicine ant of the Solertopsis subgenus Diplorhoptrum^ from Puerto Rico [124]. These ant diastereomers have been termed S-epi-cis-llSW and 5epi'tranS'llSB with structures shown in Figure 16. In addition, the decahydroquinolines CW-195A and cw-195J have now been detected in a 1:1 ratio in extracts of a Brazilian myrmicine ant of the Solertopsis subgenus Diplorhoptrum [125]. The two alkaloids occur in about the same ratio in skin extracts of a Madagascan mantelline frog, Mantella betsileo. Recently, a series of nineteen-carbon decahydroquinolines of the 269 AB group have been detected in another Brazilian myrmicine ant extract (T. Jones, personal communication, 1997). Such alkaloids have highly unsaturated side-chains, typical of the more common decahydroquinolines found in skin of dendrobatid frogs. Thus, it would appear likely that the decahydroquinolines found in frog skin extracts are obtained from dietary myrmicine ants. Decahydroquinolines were readily accumulated into skin by dendrobatid [7] and mantelline [8] frogs fed fruit flies dusted with powder containing decahydroquinolines. Captive-raised dendrobatid frogs {Dendrobates auratus, D. leucomelus) maintained in outdoor enclosures in Hawaii had c/^-195A present in skin extracts [5].

C/S-195A

5-ep/-c/s-275B'

C/S-195J

5-ep/-frans-275B

Figure 16. Structures of decahydroquinolines detected in myrmicine ants. C/^-195A and cis-195J have also been found in frog skin extracts. *A trans-llSB has not been detected, as yet, in frog skin or ants. Activity. Decahydroquinolines have relatively low toxicity [see ref 10,11]. Synthetic decahydroquinolines, including C/5-195A, caused noncompetitive blockade of nicotinic ^ The myrmicine genus Solertopsis is divided into three subgenera Solertopsis, Diplorhoptrum and Euophthaline; the subgenus will be cited as appropriate in this review.

48

J. W. Daly, H. M. Garraffo and T. F. Spande

receptor-channels in the neuromuscular junction [153]. 2,5-Dipropyl-cw-decahydroquinoline was more potent than decahydroquinoline ciS'195A. Both cis- and rra«5-decahydroquinolines inhibited binding of [^H]perhydrohistrionicotoxin to nicotinic receptor-channels of electroplax membranes [133,154,155]. Both cis- and rm/75-decahydroquinolines blocked ganglionic-type nicotinic receptor-channels in PC 12 cells and enhanced desensi-tization [133]. Decahydroquinoline cw-195A also appeared to inhibit sodium and potassium channels [153]. Further details on biological activity of decahydroquinolines were provided in two earlier reviews [10,11].

3.5.

Pyrrolizidines

3,5-Disubstituted pyrrolizidines represent one of seven classes of izidine alkaloids that have been discovered in skin extracts of the neotropical dendrobatid, Madagascan mantelline, and South American bufonid anurans. The other izidine alkaloids are the 3,5-disubstituted, 5,8-disubstituted, and 5,6,8-trisubstituted indoHzidines (Section 3.6), the 1,4-disubstituted and 4,6-disubstituted quinolizidines (Section 3.7) and the azabicyclo[5.3.0]decanes (Section 3.8), the last being izidines with fused seven- and five-membered rings, sharing a bridge-head nitrogen. Such izidine alkaloids did not occur in skin extracts of Australian myobatrachid frogs. That certain of the unclassified dendrobatid alkaloids were 3,5-dialkylpyrrolizidines was first realized during analysis of alkaloids in extracts of the bufonid toad Melanophriniscus stelzneri, where six such alkaloids were detected [86]. The mass spectra, dominated by a-cleavage of one and/or the other of two side chains, and the vapor-phase FTIR spectra with Bohlmann bands being weak or absent led to postulation of the pyrrolizidine structures. 3,5-Disubstituted pyrrolizidines had initially been discovered, first in extracts from a Floridian myrmicine thief ant of the Solenopsis subgenus Diplorhoptrum [194], and later in various species of myrmicine ants of the genus Monomorium [195-197] and one species of myrmicine ants of the genus Megalomyrmex [198]. The myrmicine ant Chelaner antarcticus, which contained pyrrolizidines [195], has been reclassified as a Monomorium species [196]. One of the pyrrolizidines in skin extracts of a bufonid toad, Melanophryniscus stelzneri, was identified by comparison with the thief ant alkaloid 5Z,8£"-3-heptyl-5-methylpyrrolizidine [194] and termed pyrrolizidine cis-223U. [86]. In the same skin extracts, pyrrolizidines cis-223B, trans-223B, cis-251K and tram'251K were identified [86] in each case by comparison with synthetic mixtures of the four diastereomers generously supplied by T. Jones. The sixth pyrrolizidine alkaloid in the Melanophryniscus stelzneri extract was cis-237G. The structures of seventeen 3,5-disubstituted pyrrolizidines detected in frog skin extracts and characterized by EI-MS and, except for 167F and 209K, by vapor-phase FTIR spectra are depicted in Figure 17. The absolute configurations are not known. In the case of trans-251K, capillary GC did not separate the synthetic 5^,8£' trans-isomQr from the 5E,SZ trans-isomQT and thus the peak corresponding to alkaloid trans-251K could prove to represent either one rra«5-diastereomer or both ^mw^-diastereomers. There are four other trans-

Alkaloids from Amphibian Skins

49

pyrrolizidines {trans-223M, trans-239K, trans-2510 and trans-265li) for which synthetic pyrroHzidines were not available for comparison, and hence the configuration of these transpyrrolizidines remains incompletely defined. The two possible diastereomers of trans223M may not separate on GC, as was the case for trans-251K. In the case oftranS'223B with identical butyl substituents at C-3 and C-5 position only one diastereomer is possible.

167F

C/S-195F 5Z,8E

C/S-223B 5Z,8E

tranS'223B 5E,8Z

C/S-223M 5Z,8E

trans-223M 5E,8E and/or 5E,8Z^ 5E,8E shown

C7H15O C/S-239K (OH) 5Z,8E

/T„

C/S-237G 5Z,8E Tentative

C7H15O tranS'239K (OH) 5E,8Eor5E,8Z 5E,8E shown

Figure 17. Structures of 3,5-disubstituted pyrrolizidines. In the pyrrolizidines, the designations cis- and trans- refer to the ring substituents. The configurational nomenclature {Z,E) follows that proposed by Sonnet [199] for 3,5-disubstituted indolizidines. The hydrogens at C-5 and C-8 in the pyrrolizidines are designated as either on the same face (Z) or the opposite face (E) to the hydrogen at C-3.

w

50

J. W. Daly, H. M. Garraffo and T. F. Spande

frans-251K 5E,8E and/or 5E,8Z^ 5E,8E shown

frans-2510

5E,8Eor5E,8Z 5E,8E shown

C7H13O frans-265H (C=0) 5E,8Eor5E,8Z 5E,8Eshown

/

C/S-265H 5Z,8E

(C=0)

C7H15O C/S-267H 5Z,8E

(OH)

Figure 17 (continued) The cis- and /m«^-3,5-disubstituted pyrrolizidines 265J have either butyl and oxohexyl substituents or heptyl and oxopropyl substituents. High resolution mass measurements of fragment ions were not obtained. Therefore, a tentative structure cannot be presented at this time and c/5-265J and /ra«5'-265J are merely tabulated in the Appendix. A vapor-phase FTIR spectrum was not obtained for 2491, which appears based on MS to be a 3-butyl-5hexenylpyrrolizidine and is so tabulated in the Appendix. Two alkaloids were formerly tentatively proposed to be 5-propyl- and 5-hexyl-indolizidines, 167B and 209D. The EI-MS, obtained in the 1980s, of the natural alkaloids did not correspond to the spectra of synthetic indolizidines 167B and 209D, and the natural alkaloids are now designated 167F and 209K. A vapor-phase FTIR was not obtained for these trace alkaloids and their relative configuration (Figure 17) remains undefined. The EI-MS fragmentation of the 3,5-disubstituted pyrrolizidines, the 3,5-disubstituted indolizidines (Section 3.6.1) and the 4,6-disubstituted quinolizidines (Section 3.7.1) is dominated by a-cleavage of the side-chains. A collision-activated NH3-CI-MS/MS technique has now been applied to several classes of bicyclic alkaloids from frog skin extracts [163]. The EI-MS and NH3-CI-MS/MS of pyrrolizidines are quite different. For example, the EI-MS of 223H shows a major fragment ion at m/z 124 for a-cleavage of the heptyl side-chain and a minor fragment ion at m/z 208 for a-cleavage loss of methyl, while CIMS/MS shows two major even-mass fragment ions, one for the ring with a methyl substituent (m/z 84) and the second for the other ring with a heptyl substituent (m/z 168). The EIMS and CI-MS/MS fragmentations for pyrrolizidine 223H are proposed in Scheme 6. Thus, for pyrrolizidines as is the case for other izidines, the two MS techniques provide complementary data relative to the nature of the side-chains (EI-MS) and the rings (CI-MS/MS).

51

Alkaloids from Amphibian Skins

.N /

C7H1 115 223H

^N^ C7H15

m/z 223

CI-NH3

C7H15 m/z 208

m/z 124

MS/MS

m/z 84

m/z 168

Scheme 6 Vapor-phase FTIR spectra are diagnostic for the relative configuration of pyrroUzidines. Only the cw-isomer with the 5Z,8Z configuration shows significant Bohlmann bands. Such bands are weak in the c/5-isomer with the 5Z,SE configuration and are virtually absent in the /ra«5-isomers with the 5^,8^ or 5E,SZ configurations. A cw-isomer with the 5Z,8Z configuration has not been detected in Nature either in ants or frog skin extracts. This appears consistent with the prevalence of /raw^-pyrrolidines as putative precursors in ants. Vapor-phase FTIR spectra for cis-223B, trans-llZE and c/5'-223H have been presented [86]. Vapor-phase FTIR spectra for pyrrolizidines cis-25\K and trans-lSlK. are presented in Figure 18. Properties of the pyrrolizidines detected in frog/toad skin extracts are tabulated in the Appendix. Synthesis. The structures of the ant pyrrolizidines, which include the frog skin alkaloid cis223H, have been established through non-stereoselective syntheses of reference mixtures of all four possible diastereomers [194-197]. The syntheses involved preparation of the appropriate triketone followed by reductive amination and cyclization. Structures were established for each of the four diastereomers, after preparative-scale GC separation, by NMR spectral analysis. GC comparison of each set of diastereomers with the corresponding natural pyrrolizidine(s) identified the relative configuration of the natural isomer(s). Subsequently, the absolute configuration of natural 5Z,8£-3-heptyl-5-methylpyrrolizidine (cz\s'-223H), also known as xenovenine, was targeted by four laboratories, using enantioselective syntheses to obtain both the dextrorotatory (3S,5R,8S)-3-heptyl-5-methylpyrrolizidine [200-205] and the levorotatory enantiomer [201,206]. Another ant alkaloid, 3methyl-5-(non-8-enyl)pyrrolizidine, was also synthesized as the dextrorotatory 3S,5R,8S enantiomer [202]. Comparisons to ascertain the absolute configurations of the naturally occurring pyrrolizidines apparently were not conducted.

J. W. Daly, H. M. Garraffo and T. F. Spande

52

~ 12-

2959.22

i 935.27

~

v-N-y

.08-

c 2870.31

1

J

1"

.04-

0-

V^^

frans-251K V,^^

J v_

1463.51 1362.02 ft

___AA^--V_

2600 2200 Wavenumber (cm-1)

Figure 18. Vapor-phase FTIR spectra for pyrrolizidines c/^-251K and trans-lSlK. It is possible that trans-251K represents a mixture of the SE^E- and 5£,8Z-/ra«5-diastereomers. The 5E,SE diastereomer is depicted. Occurrence. 3,5-Disubstituted pyrrolizidines have been detected in skin extracts of several species of dendrobatid frogs of the genus Dendrabates and to a limited extent in dendrobatid species of the genera Minyabates and Epipedobates, They have not been detected in skin extracts of dendrobatid frogs of the genus Phyllobates. In most cases, such pyrrolizidines

Alkaloids from Amphibian Skins

53

are minor or trace alkaloid constituents in dendrobatid frogs. 3,5-Disubstituted pyrrolizidines also occur in skin extracts from various mantelline frogs of the genus Mantella [56,61] and in skin extracts from the bufonid toad Melanophryniscus stelzneri [86]. The dietary source for the 3,5-disubstituted pyrrolizidines detected in skin extracts of frogs/toads undoubtedly is myrmicine ants. Such pyrrolizidines have been identified in extracts of myrmicine ants of the Solenopsis subgenus Diplorhoptrum and the genera Monomorium and Megalomyrmex [194-198]. The following four pyrrolizidines have been reported from ants: 5Z,8^-3-heptyl-5-methylpyrrolizidine {cis-21SR\ 5£,8£'-3,5-di(hex-5enyl)pyrrolizidine, 5£:,8Z-3-(non-8-enyl)-5-(£:-prop-l-enyl)pyrrolizidine and 5£,8£-3-butyl5-hexylpyrrolizidine {trans-2S1¥). Of the twenty some pyrrolizidines detected in frog/toad skin extracts, only two, namely cw-223H and trans-lSlK, have been reported from myrmicine ants. Both cis- and trans-lSlK have recently been detected in a 3:1 ratio in myrmicine ants and a dendrobatid frog, Dendrobates auratus, which occur in microsympatry in Panama (unpublished results with A.S. Rand and C. Jaramillo). Activity. The biological activity of 3,5-disubstituted pyrrolizidines apparently has not been investigated, although such pyrrolizidines, like other ant alkaloids, probably serve as insecticidal venoms or pheromones, and, like other izidine alkaloids, will probably be noncompetitive blockers of nicotinic receptor-channels.

3.6.

Indolizidines

Three classes of indolizidines have now been detected in skin extracts from frogs/toads. The first class to be discovered was 3,5-disubstituted indolizidines, whose gross structures were proposed in 1978 [49]. Nearly ten years later the structures of a second class, the 5,8disubstituted indolizidines, were established by NMR spectral analysis [105]. And after another ten years, the structure of the first member of a third class, the 5,6,8-trisubstituted indolizidines, was demonstrated [207]. The 3,5-disubstituted indolizidines, like the histrionicotoxins, decahydroquinolines and 3,5-disubstituted pyrrolizidines, could be formed biosynthetically from a precursor with a linear carbon chain. The 5,8-disubstituted indolizidines and 5,6,8-trisubstituted indolizidines have branch points in their carbon skeleton and thus are probably formed by a different biosynthetic pathway. The 3,5-disubstituted indolizidines also occur in myrmicine ants, while the 5,8-disubstituted and 5,6,8-trisubstituted indolizidines are unknown in Nature except in skin of frogs/toads.

3.6.1. 3,5-Disubstituted Indolizidines The postulated structure of indohzidine 223AB [49] was confirmed in 1981 through GC comparison with the four synthetic diastereomers of 3-butyl-5-propylindolizidine [208]. The natural indolizidine 223 AB isolated from skin extracts of the dendrobatid frog Dendrobates

54

J. W. Daly, H. M. Garraffo and T. F. Spande

histrionicus proved to be the levorotatory 5E,9E diastereomer [209]. The configuration of the hydrogens at C-5 and C-9 are designated as either trans (£) or cis (Z) relative to the hydrogen at C-3. Two side-chain hydroxylated congeners, 239AB and 239CD, from D. histrionicus, also had the 5E,9E configuration [209]. Remarkably, indolizidine 223AB isolated from skin extracts of another dendrobatid frog, D. speciosus [60], ultimately proved to be the 5Z,9Z isomer [see ref. 11]. Three of the four possible diastereomers of 223AB, namely the 5Z,9Z, the 5E,9Z and the 5E,9E, occurred together in skin extracts of the bufonid toad Melanophryniscus stelzneri with 5Z,9Z-223AB being the major isomer [86]. A different set of three isomers, namely the 5Z,9Z, 5Z,9E and 5£,9Z, occurred together in skin extracts of a Peruvian dendrobatid frog, D. imitator (unpublished results). Another 3,5-disubstituted indolizidine, 195B, isolated from D. histrionicus was shown to be the dextrorotatory 5£,9jE-3-butyl-5methylindolizidine [162]. All four diastereomers of indolizidine 195B occurred together in skin extracts of the bufonid toad Melanophryniscus stelzneri with 5Z,9£'-195B being the major isomer [86]. The 5Z,9Z-isomer of 195B had been isolated in the early seventies from Pharaoh's ant (Monomoriumpharaonis) and named monomorine I [210]. The early structure elucidation of 223AB, 195B and related congeners has been reviewed in detail [10]. Enantioselective syntheses provided levorotatory 5^,9^-223AB [211] and dextrorotatory 5£,9E-195B [212,213], thereby allowing definition of the absolute configuration of natural levorotatory 5£,9£-223AB as 3R,5R,9R and that of natural dextrorotatory 5£,9£195B as 3S,5S,9S. It is remarkable that natural 5£,9E-223AB proved to have chirality opposite to that of natural 5£,9J^-195B; both were isolated from skin extracts of Dendrobates histrionicus [162,209]. Structures of thirteen 3,5-disubstituted indolizidines characterized from skin extracts of frogs/toads are shown in Figure 19. The structure of 5Z,9Z-167E was confirmed by GC comparison with synthetic diastereomers provided by T. Jones (unpublished results with P. Jain). The mass spectra of 3,5-disubstituted indolizidines are diagnostic showing major fragments due to loss of one or the other a-substituent and a fragment of m/z 124, particularly in pseudo-EI spectra obtained with an ion-trap spectrometer. The m/z 124 fragment could arise from a McLafferty rearrangement during cleavage of the second substituent as proposed in Scheme 7 for 223 AB. The EI-MS and NH3-CI-MS/MS fragmentations of indolizidine 223AB are quite different [163]. The dominant a-cleavages in the EI-MS of 223AB, which yield major fragment ions at m/z 180 and 166, are absent in the collision-activated NH3-CI-MS/MS, which exhibits a major ion at m/z 126 corresponding to both the sixmembered ring bearing the propyl substituent and the five-membered ring bearing the butyl substituent. Vapor-phase FTIR spectra for 5£,9£-239AB and 5Z,9Z-275C are shown in Figure 20. Vapor-phase FTIR spectra for 249A [56] and for the four diastereomers of 223AB have been presented [86]. The FTIR spectra are diagnostic for determination of the relative configuration of hydrogens at the three chiral carbons, C-3, C-5 and C-9. The 3,5-disubstituted indolizidines have a broad Bohlmann band pattern with weak fine structure when H-3, H-5 and H-9 are all cis (5Z,9Z). The Bohlmann bands decrease in intensity in the other

Alkaloids from Amphibian Sl(in$

55

diastereomers in the order 5E,9E > 5E,9Z> 5Z,9£ with the last isomer having virtually no Bohlmann bands. 9

Is

3\

5Z,9Z-167E

N~

,N-

5Z,9Z-195B

(+)-5E9E-195B *

5Z,9Z-223AB

(-)-5E,9E-223AB * 5E,9Z-223AB

N5Z,9Z-249A

5E,9Z-195B

5Z9E-195B

5Z,9E-223AB

.N f (-)-5E,9E-239AB * OH OH

Nf^5Z,9Z-275C

,N. ^^-)-5E,9E-239CD

Figure 19. Structures of 3,5-disubstituted indolizidines. * Absolute configuration as shovra.

56

J. W. Daly, H. M. Garraffo and T. F. Spande

C^7

C4H9

m/z 224

m/z124

m/z 124

Scheme 7 Seven alkaloids, in addition to those depicted in Figure 19 are tentatively tabulated in the Appendix as 3,5-disubstituted indolizidines. These alkaloids are as follows: 211E, tentatively a 5£,9^-3-(hydroxybutyl)-5-methylindolizidine based on MS and FTIR; 223R, tentatively a 3-hexyl-5-methylindolizidine; 237E, tentatively an indolizidine with 3-ethyl and 5-C5H9O substituents, the latter containing a double bond and a hydroxyl group; 239E, tentatively a 3-ethyl-5-(hydroxypentyl)indolizidine; 247C, tentatively a 3,5-disubstituted 5£,9iE'-indolizidine with butenyl and pentenyl substituents; 265M, tentatively a 3,5-disubstituted indolizidine with pentenyl and C4H9O substituents; and 271F, tentatively a 3,5-disubstituted indolizidine with HC=C(CH2)3- and HC=C(CH2)4- substituents. Further data are needed on these putative indolizidines. Two bicyclic alkaloids were tentatively proposed in previous reviews [9-11, see also ref. 161] to be monosubstituted indolizidines. These were 167B and 209D. It now appears that these frog skin alkaloids were 5-methyl-3-propylpyrrolizidine and 3-hexyl-5-methylpyrrolizidine, respectively. The proposed monosubstituted indolizidines 167B and 209B have been synthesized (see below) and hence the code numbers will be retained for these structures. However, in the tabulation in the Appendix it will be noted that the natural occurrence of these alkaloids has not been confirmed. The natural alkaloids are now proposed to be pyrrolizidines and are so tabulated in the Appendix as 167F and 209K (Figure 17). Neither show a fragment ion at m/z 96 that would form from a 5-substituted indolizidine by a retro-Diels Alder reaction after initial a-cleavage. There is one alkaloid, 195H, that is presently proposed in the Appendix to be a monosubstituted indolizidine (5pentylindolizidine) and it does exhibit a fragment ion at m/z 96. A brief summary with references on physical (optical rotation) and spectral (FTIR, NMR) properties of 3,5-disubstituted indolizidines has been provided [11, see also references to synthetic compounds in ref. 10].

57

Alkaloids from Amphibian Skins 2937.13

1600-

1

r'VA

9R>

1200

r^

800-

239AB

^OH

2874.03

1A2796.84

1

400-

L_^ ^J V 2800

2400 2000 Wavenumber(aTvl)

1053.27 1455.27 1374.86

.

1030.41 912.971

lv^A^A^/-^AAAA^-

3800

3400

2600 2200 Wavenumber (cm-1)

Figure 20. Vapor-phase FTIR spectra for indolizidines 5^,9£-239AB and 5Z,9Z-275C. Synthesis. The initial syntheses [208] were in efforts to verify a proposed structure of indolizidine 223AB [49], which at that time was referred to as gephyrotoxin 223AB. That designation is unfortunate because of possible confusion with the tricyclic gephyrotoxins that are also found in extracts of frog skin and because indolizidines, such as 223AB, have low toxicity. However, the bicyclic gephyrotoxin designation persists in the synthetic literature and has been applied not only to indolizidine 223AB, but to other mono- and disubstituted indolizidines. A detailed summary of the early synthetic efforts towards

58

J. W. Daly, H. M. Garraffo and T. F. Spande

indolizidine 223 AB was provided in 1986 [10]. All of the four possible diastereomers of 3butyl-5-propylindolizidine were defined through complementary synthetic routes to all diastereomers by Spande [208], to the 5Z,9Z and 5Z,9E isomers by Hart and Tsai [214], and to the 5E,9E isomer by MacDonald [215]. In a collaborative effort the natural indolizidine 223 AB, isolated from skins of the dendrobatid frog Dendrobates histrionicus, was shown to be the 5E,9E diastereomer [208, see review in ref 10]. An "iso" series of diastereomers of 5-butyl-3-propylindolizidine also was prepared by Spande [see ref 10]; the SE,9E diastereomer cochromatographed with natural indolizidine 223AB, but differed in its mass spectrum. Husson's laboratory was the first to prepare the 3R,5R,9R enantiomer of 5£',9£-223AB [166,211]; synthetic and natural alkaloids were both levorotatory and, thus, appeared identical in absolute configuration. An X-ray analysis of racemic 5E,9£'-223AB has been provided [216]. Later, enantioselective syntheses of levorotatory indolizidines 5E,9E239AB and 5£,9£-239CD [213,217] and dextrorotatory indolizidine 5^,9^-1956 [212] defined the absolute configurations of these alkaloids. An alkaloid found to be a trail marker component for Pharaoh's ant {Monomorium pharaonis) was determined in 1973 to be a 3-butyl-5-methylindolizidine [210], and was named monomorine I. The earlier syntheses of diastereomers of monomorine I [199,218221] preceded the syntheses of indolizidine 223AB. The absolute configuration of (+)monomorine I was later shown to be 3R,5S,9S (222,223), which would correspond to a (+)5Z,9Z-195B; the rotation and hence absolute configuration of the frog skin alkaloid 5Z,9Z195B are not known. There exist many more syntheses of monomorine I, either as the racemate [215,224-235], the (-)-enantiomer [166] or the (+)-enantiomer [236-248]. In 1994, (-)-monomorine I along with {-y5E,9E, (-)-5£,9Zand (+)-5Z,9£ isomers were obtained [249]. The (-)-5£,9£ isomer is enantiomeric to the frog (+)-5£,9£-195B. The 5Z,9Z-3-ethyl-5-methyl and 5Z,9Z-3-hexyl-5-methyl analogs of monomorine I have been identified in myrmicine ants by comparison with synthetic diastereomers [250]. The (+)enantiomers have been prepared [241]. The first of these two ant alkaloids corresponded to indolizidine 5Z,9Z-167E (unpubhshed results with P. Jain) from frog skin extracts. The second would correspond to 223R, but with uncertain configuration. An enantioselective synthesis of the ant alkaloid 5£,9Z-3-butyl-5(pent-4-enyl)indolizidine has been reported [251]. A putative 5i?,9£-3-butenyl-5-pentenylindolizidine 247C occurred in frog skin extracts. The absolute configurations (3R,5S,9S and 3R,5R,9S) of 5Z,9Z- and 5£:,9Z-3butyl-5-(l-oxopropyl)indolizidines from the myrmicine ant Myrmicaria eumenoides were recently determined by comparison on chiral GC columns with synthetic diastereomers and enantiomers [252]. The names "myrmicarin 237A" and "myrmicarin 237B" were suggested for these alkaloids. Epimerization at C-5 occurred readily. These ant alkaloids have not been detected in frog skin extracts. Further syntheses of indolizidine 5 J?,9£-223AB fromfi-ogskin, both in racemic form [227,253-259] and as the (-)-enantiomer [213,248,260-264], have appeared, following the initial synthetic efforts directed at structure elucidation. The 5Z,9Z [255,260], 5£,9Z [253] and 5Z,9£ [257,258] diastereomers of indolizidine 223 AB have also been prepared. A mixture of 5Z,9Z- and 5Z,9£-223AB was obtained in one synthesis [265].

Alkaloids from Amphibian Skins

59

Both enantiomers of the frog skin indoHzidine 5£,9£^-195B were synthesized by C. Kibayashi and colleagues [212,213], thereby estabUshing the absolute configuration of the dextrorotatory natural alkaloid. Both enantiomers of the 5Z,9£' isomer were also synthesized. Two routes to racemic 5£,9£-195B [225,232] and three enantioselective syntheses of 5Z,9Z-195B have been reported [236,249,266,267]. In the 1980's, two of the alkaloids, 167B and 209D, which had been detected in trace quantities in three frog skin extracts, were tentatively proposed to be a 5-propylindolizidine and a 5-hexylindolizidine [9,10]. Such indolizidines were subsequently synthesized in several laboratories [240,268-274]. Unfortunately, the trace alkaloids 167B and 209D could no longer be detected for GC comparison in the original three extracts that had been stored for six to thirteen years. It now appears more likely based on EI-MS data that the natural alkaloids were disubstituted pyrrolizidines, rather than monosubstituted indolizidines and they are so tabulated as 167F and 209K in the Appendix. The code names 167B and 209D are retained for the synthetic indolizidines, which have not yet been detected in Nature. Occurrence. The 3,5-disubstituted indolizidines have been detected rather infrequently in skin extracts of neotropical frogs of the dendrobatid genus Dendrohates and usually as minor or trace alkaloids [1]. Indolizidines 223AB and 195B have been the most common. Indolizidine 223AB was a major alkaloid in one population of Colombian Dendrohates histrionicus, where it was shown to be the levorotatory 5E,9E isomer. Remarkably, in Panamanian D. speciosus, where it also was a major alkaloid, 223AB proved to be the 5Z,9Z isomer, with optical rotation unknown. Indolizidine 223AB also occurred as a major alkaloid in one population of D. auratus but in that case and in several other species of Dendrohates, the stereochemistry is not known. Indolizidines 5£,9^-239AB and 5E,9£-239CD, which are side-chain terminally hydroxylated congeners of 5£',9£-223AB, have been detected almost exclusively in extracts from a few populations of D. histrionicus [1]. They were also detected in a closely related species, D, occultator. The indolizidine 195B, isolated from a Colombian population ofD. histrionicus, proved to be a dextrorotatory 5E,9E diastereomer of 5-methyl-3-propylindolizidine; the ant alkaloid monomorine I is the 5Z,9Z diastereomer. Indolizidine 195B has been detected as a minor or trace alkaloid in relatively few frog species of the genus Dendrohates and the stereochemistry in most cases is unknown. Both 5E,9E- and 5Z,9Z-195B occurred in one Panamanian population of D. pumilio (unpublished results). Three diastereomers (5Z,9Z, 5Z,9£, 5E,9Z) were detected in a Peruvian population of Z). imitator. Indolizidine 5Z,9Z-167E was present as a trace alkaloid in one population of £). pumilio. 3,5-Disubstituted indolizidines have not been detected in skin extracts of frogs of the dendrobatid genera Epipedohates and Minyohates. Indolizidine 223AB occurred in small amounts as the major volatile alkaloid in Phyllohates aurotaenia and 195B as a trace alkaloid in P. hicolor and P. terrihilis. Various 3,5-disubstituted indolizidines were present in skin extracts of the bufonid toad Melanophryniscus stelzneri, including several diastereomers of 195B and 223AB [86]. None were detected in the Brazilian species Melanophryniscus moreirae. 3,5-Disubstituted indolizidines, such as 223AB, 249A and 275C, were present in skin extracts of certain frogs of the mantelline

60

J. W. Daly, H. M. Garraffo and T. F. Spande

genus Mantella [56]. 3,5-Disubstituted indolizidines were not detected in skin extracts of frogs of the myobatrachid genus Pseudophryne [25]. It would appear that dietary myrmicine ants are the source of the 3,5-disubstituted indolizidines detected in frog/toad skin. Indeed, dendrobatid frogs of the genus Dendrobates appear to be "ant specialists" [275-278]. 3,5-Disubstituted indolizidines have been detected in myrmicine ants of the genus Monomorium [197,210,221], the Solenopsis subgenus Diplorhoptrum [250,279,280] and the genus Myrmicaria [252]. Ant indolizidines include 5Z,9Z-3-butyl-5-methylindolizidine ((-)-monomorine I) [210], 5Z,9Z-3-ethyl-5methyhndolizidine [250], 5Z,9Z-3-hexyl-5-methylindolizidine [250], 5£,9£:-3-hexyl-5methylindolizidine [280], 5Z,9£'-3-hexyl-5-methylindolizidine [280], 5£,9Z-3-butyl-5-(pent4-enyl)indolizidine [197], 5£,9Z-3-butyl-5-(l-oxopropyl)indolizidine [252] and 5Z,9£-3butyl-5-(l-oxopropyl)indolizidine [252]. Vapor-phase FTIR spectra of the four diastereomers of 3-hexyl-5-methylindolizidine have been presented [280]. Both 5E,9E- and 5^9Z-3butyl-5-propylindolizidine were detected in Puerto Rican myrmicine ants of the Solenopsis subgenus Diplorhoptrum [279]. These correspond to the major diastereomers 5£,9£-223AB and 5Z,9Z-223AB isolated from extracts of dendrobatid frog skin. Of the other ant indolizidines, monomorine I has been identified in frog skin as a minor 5Z,9Z diastereomer of 195B in extracts of the bufonid toad Melanophryniscus stelzneri, where 5Z,9i?-195B was the major isomer [86] and as a trace alkaloid in an extract of the dendrobatid frog Dendrobates imitator (unpublished results). Indolizidine 195B, isolated from skin extracts of one population of the dendrobatid frog D. histrionicus, was the 5E,9E-isomQT [162]. Indolizidine 195B was present in skin extracts from three captive-raised species (D. auratus, D. leucomelus, Phyllobates aurotaenid) maintained in outside enclosures in Hawaii [5, see Section 3.1]. The ant alkaloid 5Z,9Z-3-ethyl-5-methyl-indolizidine corresponds to 167E. The ant alkaloid 5£,9Z-3-butyl-5-(pent-4-enyl) indolizidine [197] is a diastereomer of 5Z,9Z-249A found in skin extracts of one mantelline species. An alkaloid 223R from the dendrobatid frog D. auratus probably corresponds to one diastereomer of the ant alkaloid, 3-hexyl-5-methylindolizidine. Twenty 3,5-disubstituted indolizidines have been detected in skin extracts, of which only four (5£,9£-223AB, 5Z,9Z-223AB, 5Z,9Z-195B and 5Z,9Z-167E) have been reported in ant extracts. Activity. 3,5-Disubstituted indolizidines of amphibian skin appear to have low toxicity [see ref. 10], but isomers of 223AB are all relatively potent non-competitive blockers of muscletype and ganglionic-type nicotinic receptor-channels [133,155]. A hydroxy 1 group in the side-chain, as in 239AB and 239CD, markedly reduces affinity at muscle-type nicotinic receptor channels of Torpedo electroplax, but not at ganglionic-type channels of pheochromocytoma cells [133,155]. Monomorine I acts as a trail-marker component for the ant Monomoriumpharaonis [210], but may also have "arresting" effects [221]. The four synthetic diastereomers of monomorine I differed in their behavioral effects on the ant Monomorium pharaonis [221].

Alkaloids from Amphibian Skins

61

3.6.2. 5,8-Disubstituted Indolizidines A number of unclassified "dendrobatid" alkaloids were bicyclic with no exchangeable hydrogen on nitrogen and afforded a simple EI-MS consisting of a single major fragment ion (base peak) of C9Hi6N'^ (m/z 138). Four of these alkaloids were isolated in the mid1980's in sufficient quantities for NMR spectral analysis. 5-Substituted-8-methyl-indolizidine structures for 205A, 207A, 235B" (formerly 235B) and 235B' were then proposed [60,105]. Indolizidine 205A had a 5-pent-4-ynyl and 207A, a 5-pent-4-enyl side-chain. Indolizidine 235B" and 235B' differed in stereochemistry and in the position of the double bond in the 5-heptenyl side-chain with 235B' from Dendrobates speciosus having a terminal double bond [60] and 235B from D. pumilio having a Z-4',5*-double bond [105]; indolizidine 235B is now referred to as 235B" [11] and 235B is used as a generic code for such isomers. Further 5,8-disubstituted indolizidines were later isolated and NMR spectral studies were used to define structures for 203A, 233D and 251B [161]. The ring configuration in the seven indolizidines that were subjected to NMR spectral analysis was 5,9Z and both the 5and 8-substituents were equatorial. Mass spectra, dominated by a base peak at m/z 138 and accompanied by CeHioN"^ ion (m/z 96), are diagnostic for 5-substituted 8-methylindolizidines, while the characteristic vapor-phase FTIR spectra with a sharp Bohlmann band near 2789 cm"^ allows assignment of a 5,9Z configuration to most such alkaloids. There are two 5-substituted-8-methylindolizidines (2231,259B) that have been postulated to have an atypical 5,9£ configuration based on a weak Bohlmann band. Many alkaloids from skin extracts are now classified as 5,8-disubstituted indolizidines, based on MS and FTIR characteristics. Most have an 8-methyl group, but some appear to have other alkyl, alkenyl or alkynyl groups at C-8 and, therefore, give base peaks greater than the m/z 138 peak for 8methylindolizidines. The structures for thirty-two 5,8-disubstituted indolizidines are shown in Figure 21. Alkaloids 243D and 245C have been tentatively proposed to be 5,8-disubstituted indolizidines with a trans double bond adjacent to C-5 reducing the usual facile acleavage [54]. Most of the 5,8-disubstituted indolizidines have side-chains at C-5 containing three, five, seven or nine carbons, while at C-8 most have either methyl or side-chains containing two or four carbons. A pattern of side-chains differing in length by two carbons is reminiscent of the pattern that pertains for histrionicotoxins and decahydroquinolines. In both EI and ion-trap pseudo-EI-MS, a base peak due to a-cleavage dominates the fragmentation for 5,8-disubstituted indolizidines. A diagnostic peak at m/z 96, due to a retro-Diels-Alder elimination of an alkene from the base peak, is very prominent in ion-trap pseudo-EI-MS, while being a minor ion in normal EI-MS. A fragmentation pathway is proposed in Scheme 8. The NH3-CI-MS/MS for 5,8-disubstituted indolizidines is quite complex, involving retro-Diels-Alder elimination and McLafferty cleavages from a fragment ion containing the ring bearing the 5,8-substituents [163]. As expected, a-cleavage does not occur. The FTIR spectra for 5,8-disubstituted indolizidines with the hydrogens at C-5 and C-9 on the same face show a characteristic sharp and intense Bohlmaim band near 2789 cm' . Only two of the proposed 5,8-disubstituted indolizidines, namely 2231 and 259B, show a

J. W. Daly, H. M. GarrafTo and T. F. Spande

62

weak absorbance peak in the Bohlmann band region [86,281]. Vapor-phase FTIR spectra of a typical (5,9Z) 5,8-disubstituted indolizidine, 205A, and an atypical (5,9E) indolizidine, 259B, are depicted in Figure 22. Vapor-phase FTIR spectra for typical (5,9Z) 5,8-disubstituted indolizidines 203A, 207A", 217B, and 2211 and for atypical (5,9£) indolizidines 2231 and 259B have been published [56,86,161,281]. Indolizidine 207A" has an internal double

181B

N-

N-

(-)-203A

(-)-205A

N207A

219F

209B

219L

N-

223J

231C

2091

2211

N-

233D

2231 Tentative Atypical

N~ (-)-235B'

Figure 21. Structures of 5,8-disubstituted indolizidines. * Absolute configuration as shown. *Absolute configurations of natural 235B" and 251B, which were dextrorotatory, are depicted opposite to the configuration of levorotatory 203A, 205A and 235B', and to synthetic levorotatory235B".

63

Alkaloids from Amphibian Skins

C4H9O

(+)-235B" *

237D

C5H9O (OH) 237H

C7H13O (C=0) 251U

239C

C9H17O

(OH)

259B Tentative Atypical Figure 21 (continued) bond rather than the terminal double bond of 207A (see Appendix). There is another isomer 207A' that also has an internal double bond.

J. W. Daly, H. M. Garraffo and T. F. Spande

a-cleavage

uvT"^ >

^ Q>NV (M - Rf

Diels-Alder

[|

\

%^NV m/z 96

Scheme 8

A brief summary with references on physical (optical rotation) and spectral (FTIR, NMR) properties has been provided for 5,8-disubstituted indolizidines [11]. Indolizidines 203A, 205A, 233D«HC1 and 235B' are levorotatory; however, the observed rotations were much less than those of the corresponding synthetic levorotatory 205A and 235B* [11]. Indolizidines 235B" and 251B were dextrorotatory but in the case of 235B" only weakly ([OC]D +11°)? compared to the corresponding synthetic levorotatory 235B" ([a]D -85°). The synthetic 5R,8R,9S-235B" had other spectral properties identical with natural 235B" [282, 283]. It is possible that 235B** and perhaps 205A and 235B* either contain small amounts of strongly dextrorotatory impurities or are mixtures of both enantiomers. Twenty-six alkaloids, besides the thirty-two shown in Figure 21 are tabulated as putative 5,8-disubstituted indolizidines in the Appendix. The alkaloids are as follows: 1951, tentatively a 5-butyl-8-methylindolizidine; 197C, tentatively an 8-(hydroxymethyl)-5propylindolizidine; 207Q, tentatively a 5-allyl-8-propylindolizidine; 219J, tentatively an 8butynyl-5-propylindolizidine; 221A, tentatively a 5-hexenyl-8-methylindolizidine; 221K, tentatively an 8-butyl-5-propenylindolizidine; 223D, tentatively a 5-hexyl-8-methylindolizidine; 225D, tentatively a 5-(hydroxypentyl)-8-methylindolizidine; 239A, tentatively a 5butyl-8-(hydroxypropyl)indolizidine; 239B, tentatively an 8-butyl-5-(hydroxypropyl)indolizidine; 239D, tentatively a 5-(hydroxybutyl)-8-propylindolizidine; 239F, tentatively an 8(hydroxyethyl)-5-pentylindolizidine; 239G, tentatively a 5-(hydroxyhexyl)-8-methylindolizidine; 241C, tentatively a 5-(dihydroxybutyl)-8-ethyl-indolizidine; 245D, tentatively an 8methyl-5-octenynylindolizidine; 247E, tentatively an 8-butenyl-5-pentenylindolizidine; 249L, tentatively an 8-methylindolizidine with a C7H11O side-chain at C-5; 257C, tentatively an 8-methyl-5-nonadienynylindolizidine; 261D, tentatively an 8-methyl-5-nonadienyl- or 5-nonynyl-indolizidine; 263F, tentatively an 8-methyl-5-nonenylindolizidine; 271A, tentatively an 8-butenyl-5-heptenynylindolizidine; 273B, tentatively an 8-butenyl-5-heptadienylor 8-butenyl-5-heptynylindolizidine; 275F, tentatively an 8-butenyl-5-heptenylindolizidine; 295A, tentatively a 5-(hydroxydecyl)-8-methylindolizidine; and 295B, which was proposed [56] to be a ring-hydroxylated 5-(hydroxynonenyl)-8-methylindolizidine. Alkaloid 167A, proposed as a 5,8-disubstituted indolizidine in an earlier review [11], does not show a significant retro-Diels-Alder peak at m/z 96, and is now tabulated as unclassified in the Appendix. Certain of the alkaloids listed above as putative 5,8-disubstituted indolizidines may also have to be reclassified if further data can be obtained. There may be a 6,7-dehydro subclass of 5,8-disubstituted indolizidines (see unclassified alkaloid 245F in the Appendix).

Alkaloids from Amphibian Skins

65

1600^ 2936.37

1

,r

1200-

1

3326.95

-

2882.85

il

2786.77 £

800-

1

2708.82

1

S

< 400-

u

f

, 0-

1 I1

s ^ N ^

^ 205A 1455.5 1375.75 1246.13

i

1142.07

V

1

2800

2117.6

2400 2000 Wavenumber (cm-1)

j

A ''4'/\

^ > ^ . . . , 86o

2600 2200 Wavenumber (cm-1)

Figure 22. Vapor-phase FTIR spectra for a typical (5,9Z) 5,8-disubstituted indolizidine 205A and a putative atypical (5,9£r) 5,8-disubstituted indolizidine 259B. Synthesis. Synthetic efforts in several laboratories towards some of the natural 5,8-disubstituted indolizidines have confirmed structures, which were proposed based on NMR spectral analyses [60,105,161], and have established the absolute configuration of several indolizidines. Racemic 205A [285,286], 207A [285,286], 209B [287] and 235B' and 235B" [288]

66

J. W. Daly, H. M. Garraffo and T. F. Spande

have been synthesized. Enantioselective syntheses have provided the following levorotatory alkaloids: 5R,8R,9S (-)-205A [282-284], (-)-207A [281,283,284,289,290], (-)-209B [272, 281,283-286,291,292], (-)-235B' [289], and (-)-235B" [282-284]. The dextrorotatory enantiomer of 209B has also been synthesized [293]. Both racemic 209B and the 8-epimer have been synthesized [294]. Synthesis of a chiral lactam intermediate, 8R,9R-8-methylindolizidin-5-one, for the synthesis of 5-substituted 8-methylindolizidines has been reported [295]. An (-)-8-hydroxymethyl-5-pentylindolizidine was reported as an intermediate in the synthesis of (-)-209B [272]. Levorotatory 5R,8R,9S-8-butyl-5-propylindohzidine was recently synthesized and the FTIR spectrum was consonant with the structure (5,9Z), but quite different from natural 2231 [281]. The latter was concluded, based on the FTIR spectrum, to have H-5 and H-9 in a trans arrangement (see Figure 21). The FTIR spectrum of synthetic 5,9Z-(-)-8-butyl-5-propylindolizidine was almost identical with that of natural 223J, an indolizidine with 5-butyl and 8-propyl substituents [281]. Occurrence. The 5,8-disubstituted indolizidines represent the largest class of alkaloids detected in skin extracts from frogs/toads. However, some of the near sixty such alkaloids, tabulated in the Appendix of the present review, may ultimately prove to belong to other classes. As yet, the 5,8-disubstituted indolizidines found in frog skin extracts have not been detected elsewhere in Nature. A dietary source, thus, remains a mystery. Certainly, with a branch point in the carbon skeleton, such indolizidines cannot be formed by cyclization of a single precursor with a linear carbon chain, as could occur for the histrionicotoxins, decahydroquinolines, 3,5-disubstituted pyrrolizidines, 3,5-disubstituted indolizidines, 4,6-disubstituted quinolizidines and 3,5-disubstituted azabicyclodecanes; two linear carbon chain precursors would seem to be required. 5,8-Disubstituted indolizidines were readily accumulated into skin by a dendrobatid frog [7] and by a mantelline frog [8] from fruit flies dusted with a powder containing 5,8-disubstituted indolizidines. Of the dendrobatid genera that have lipophilic skin alkaloids, the 5,8-disubstituted indolizidines were commonly detected in Dendrabates, Epipedobates, and Minyobates, while being virtually absent in Phyllobates [1 and unpublished results]. 5,8-Disubstituted indolizidines occurred in skin extracts of five of nine species of Madagascan frogs of the mantelline genus Mantella, but the majority have substituents other than methyl at C-8 [56,61], in contrast to dendrobatid frogs where an 8methyl group predominates. Only one of the 5,8-disubstituted indolizidines, namely the atypical 259B, has been detected in skin extracts of toads of the bufonid genus Melanophryniscus [86]. Such indolizidines have not been detected in frogs of the myobatrachid genus Pseudophryne [25]. Activity. The biological activity of 5,8-disubstituted indolizidines has not been investigated in detail. In common with histrionicotoxins, decahydroquinolines and other izidines, the 5,8-indolizidines, such as 205A, 207A, 209B and 235B' and 235B", were noncompetitive blockers of both neuromuscular-type {Torpedo electroplax) and ganglionic-type (PC 12 cells) nicotinic receptor-channels [156]. The 235B isomers were as potent as perhydrohistrionicotoxin, while the others were less potent. Indolizidine 205A at low concentrations

Alkaloids from Amphibian Skins

67

enhanced rather than inhibited binding of [^H]perhydrohistrionicotoxin to electroplax membranes; the inhibition is a measure of activity as a noncompetitive blocker, while stimulation is usually indicative of agonist activity [156]. The other 5,8-disubstituted indolizidines caused only inhibition. A synthetic compound with an 8-hydroxymethyl group had very low activity as an inhibitor in electroplax membranes.

3.6.3. 5,6,8-Trisubstituted Indolizidines The structure of the frog skin izidine alkaloid 223 A was finally elucidated when about one milligram was isolated from extracts of twenty skins of a Panamanian Dendrabates pumilio [207]. NMR spectral analysis established the structure as that of a 6,8-diethyl-5propylindolizidine with the relative configuration shown in Figure 23. Nine other alkaloids are at present proposed to be 5,6,8-trisubstituted indolizidines based on EI-MS and FTIR spectra and the structures are shown in Figure 23. One of these, alkaloid 249H, based on MS, FTIR and NMR spectral analysis, is unique among indolizidines in having a branched side-chain; the proposed structure is that of a 5,9i^-5-((F)-hex-3-en-3-yl)-6-methyl-8-ethylindolizidine [295a]. Five other alkaloids are tabulated as 5,6,8-trisubstituted indolizidines in the Appendix, based only on EI-MS. They are as follows: 245G, tentatively a 6-ethyl-5hexenynyl-8-methylindolizidine; 263D, tentatively a 6,8-diethyl-5-hexenyl-indolizidine; 2651, tentatively a 6-methyl-8-ethylindolizidine with a 5-C6HiiO substituent; 277C, tentatively an 8-butyl-6-ethyl-5-pentenylindolizidine; 279F, tentatively a 6,8-dimethyl-5-(hydroxyoctenyl)indolizidine. Alkaloid 223A had been tentatively proposed to be a 1,4-dipropylquinolizidine, based on an EI-MS dominated by loss of a propyl radical [5,11]. Alkaloid 273A also had been previously proposed to be a 1,4-disubstituted quinolizidine [56]. The FTIR Bohlmann bands, however, clearly distinguish 5,6,8-trisubstituted and 5,8-disubstituted indolizidines from 1,4-disubstituted quinolizidines. The indolizidines have a sharp, intense Bohlmann band [161,207], while the quinolizidines (Section 3.7.2) have a broader, somewhat less intense Bohlmann band [56,86]. Indolizidine 267J, however, exhibits a very weak Bohlmann band and an atypical 5,9E structure was proposed for this alkaloid [207], which shows on EI-MS the m/z 124 and m/z 70 fragments expected of a 5,8-disubstituted 6-ethylindolizidine. There are five other "atypical" alkaloids that can be tentatively proposed to be 5,6,8trisubstituted indolizidines based on EI-MS; all five showed a very weak Bohlmann band, suggestive of a 5,9E structure. These are as follows: 259C, tentatively a 6,8-dimethyl-5octenynylindolizidine; 263A, tentatively a 6,8-dimethyl-5-octenylindolizidine; 265L, tentatively a 6,8-dimethyl-5-(hydroxyheptenyl)indolizidine; 275E, tentatively a 6,8-dimethyl-5nonadienylindolizidine; 277E, tentatively a 6,8-dimethyl-5-nonenylindolizidine; 293C, tentatively a 6,8-dimethyl-5-(hydroxynonenyl)indolizidine (see Appendix).

J . W . Daly, H. M. Garraffo and T. F. Spande

68

8

9

6 . -

- -

_

195G

~

223A

i ~

.~/

233G

HO

231B

249H

HO

Tentative Atypical

273A Figure 23. Structures of 5,6,8-trisubstituted indolizidines. The EI-MS of 5,6,8-trisubstituted indolizidines exhibit a base peak corresponding to cleavage of the substituent at C-5 and a retro Diels-Alder daughter ion at either m/z 124 (6ethyl) or m/z 110 (6-methyl). There often is a fragment ion at m/z 70. A fragmentation pathway for 5,6,8-trisubstituted indolizidines is proposed in Scheme 9. However, assign-

Alkaloids from Amphibian Skins

69

ments of many of the izidine alkaloids to either the 5,6,8-trisubstituted indolizidine class or the 1,4-disubstituted quinolizidiiie class cannot be made based solely on EI-MS, since both 5,8-disubstituted-6-methylindolizidines and 1,4-disubstituted quinolizidines have a retroDiels-Alder fragment ion at m/z 110. The presence of a fragment ion at m/z 70 for indoHzidines or at m/z 84 for quinolizidines can be diagnostic, but often such ions are quite weak. Several alkaloids, some of which were previously tentatively assigned to the 1,4-disubstituted quinolizidine class [11], are now merely tabulated as being izidines in the Appendix. The alkaloids tabulated as izidines are 181A, 193G, 195D, 207C, 209C, 209E, 219B, 219E, 223C, 237C, 251S, 251T, 255B and 261B. All do show a major a-cleavage base peak and most show a diagnostic izidine peak at m/z 110, but FTIR spectra have not been obtained. All probably represent either a 5,6,8-trisubstituted indolizidine with a 6-methyl substituent or a 1,4-disubstituted quinolizidine.

a-cleavage

L^T

\

retro Diels-Alder ler

fTA

R' (M-R)^

R" = CH3; m/z 110 R" = C^5; m/z 124

Scheme 9 The vapor-phase FTIR spectra for 5,6,8-trisubstituted indolizidines with the hydrogens at C-5 and C-9 on the same face show the same sharp, intense Bohlmann band near 2784 cm'^ as do the 5,8-disubstituted indolizidines having the same 5,9Z configuration (Section 3.6.2). Some alkaloids, such as 267J (Figure 23), that appear to be 5,6,8-trisubstituted indolizidines do not show an intense Bohlmann band, and hence probably have a 5,9£' configuration (see above). The vapor-phase FTIR of the typical 5,9Z indolizidine 231B and of the atypical 5,9£ indolizidine 267J are depicted in Figure 24. The FTIR spectrum of 223 A has been published along with the proton NMR spectrum [207]. Synthesis. To our knowledge, no syntheses of the 5,6,8-trisubstituted indolizidines, detected in frog skin extracts, have been reported. Occurrence. The 5,6,8-trisubstituted indolizidines 223A and 231B were relatively common in skin extracts of dendrobatid frogs of the genus Dendrobates, where each had been detected in about ten of fifty species [1 and unpublished results]. Other 5,6,8-trisubstituted indolizidines have been detected only rarely in Dendrobates species. Indolizidines 223A and 231B occurred in skin extracts from many populations of Dendrobates pumilio from Panama and Costa Rica [11,207 and unpublished results]. Remarkably, indolizidine 223A was the major alkaloid in skin extracts from one population of D. pumilio, while being

70

J. W. Daly, H. M. Garraffo and T. F. Spande

absent in another population from the same small Panamanian island. Indolizidines 223 A and 231B occurred in skin extracts of three species of the dendrobatid genus Minyobates, namely M altobueyensis, M. bombetes and M minutus. The 5,6,8-trisubstituted indolizidines have been detected only rarely in species of the dendrobatid genera Phyllobates and Epipedobates. Both 223A and 231B were present in extracts of the Amazonian Epipedobates bassleri from Peru, while 231B was present in extracts of Amazonian E. pulchripectus

1459.37 1382.71 1210.52 1148.02 \ 1287^

r^ u. 3800

3400

2600 2200 Wavenumber(cm-I)

8 c

I

267J

oos^

2600 ' 22*00 Wavenumber (cm-1)

Figure 24. Vapor-phase FTIR spectra for 5,6,8-trisubstituted indolizidine 231B and a putative 5,6,8-trisubstituted indolizidine, 267J.

Alkaloids from Amphibian Skins

71

from Brasil and from E. tricolor from western Ecuador. In frogs of the genus Phyllobates, the trisubstituted indolizidine 223A has been detected only twice, both times in extracts from Panamanian populations of P. lugubris. A trace alkaloid in skin extracts from mantelline Mantella laevigata of Madagascar was identified as 223A [61]. IndoHzidine 273A was detected in extracts from Mantella haroni [56]. 5,6,8-Trisubstituted indolizidines have not been detected in South American bufonid toads {Melanophryniscus\ nor in Australian myobatrachid frogs (Pseudophryne). A dietary source is unknown. Based on the occurrence of such trisubstituted indolizidines, mainly in extracts of dendrobatid frogs, it appears probable that the dietary arthropods are found mainly in neotropical rain-forests. Activity. Nothing is known of the biological activity of 5,6,8-trisubstituted indolizidines. Presumably, like many izidine alkaloids from frog skin, they will prove to be noncompetitive blockers of nicotinic receptor-channels.

3.7.

Quinolizidines

Two classes of quinolizidines are known to occur in skin extracts of anurans, namely the 4,6-disubstituted quinolizidines and the 1,4-disubstituted quinolizidines. The former are analogous to the 3,5-disubstituted pyrrolizidines and indolizidines in having substituents a and a' to the nitrogen and being apparently derived from a linear carbon-chain precursor. All three izidine classes (pyrrolizidines, indolizidines and quinolizidines) with a,a'-disubstitution are now known to occur in myrmicine ants. The 1,4-disubstituted quinolizidines are analogous to the 5,8-disubstituted indolizidines in having only one a-substituent and having a branch point in the carbon skeleton of a putative precursor. They have been detected in Nature as yet only in extracts of frog/toad skin.

3.7.1. 4,6-Disubstituted quinolizidines The structure of alkaloid 195C, found relatively often in skin extracts from dendrobatid frogs, has now been established by MS and FTIR spectral analysis and comparison with synthetic diastereomers [125]. Alkaloid 195C proved to be 6Z,10£-4-methyl-6-propylquinolizidine as shown in Figure 25. The absolute configuration is unknown. Only one other alkaloid, 2371, is tentatively assigned to this class. Previously, alkaloid 195C had been speculated to be either a pyrrolizidine or an indolizidine [5]. The EI-MS of 195C shows a base peak at m/z 152 and a minor peak at m/z 180 corresponding to a-cleavage loss of propyl or methyl, respectively [125]. The NH3-CI-MS/MS yielded fragment ions at m/z 126, corresponding to the six-membered ring bearing a propyl group, and m/z 98, corresponding to the six-membered ring bearing a methyl. The proposed fragmentation pathways for 195C are shown in Scheme 10.

J. W. Daly, H. M. Garraffo and T. F. Spande

72

2371

195C Figure 25. Structures of 4,6-disubstituted quinolizidines.

.N

CI-NH3

EI-MS

195C

m/z180

m/z195

m/z152

MS/MS

m/z196

m/z98

m/z126

Scheme 10 The vapor-phase FTIR spectrum of 195C (Figure 26) shows a relatively weak Bohlmann band at 2813 cm'^ The NMR spectrum had an overlapping methyl doublet and methyl triplet and three downfield CHN protons [125]. Synthesis. The four diastereomers of 4-methyl-6-propylquinolizidine, were synthesized starting from 2,6-dimethylpiperidine by three pathways that allowed the unambigous assignment of the relative configurations of each of the diastereomers [125]. The 6Z,10E diastereomer proved identical to 195C. A c/5-fused quinolizidine conformation is anticipated for this structure. Occurrence. Quinolizidine 195C has been detected in eight species of dendrobatid frogs of the genus Dendrobates, in two species of the genus Epipedohates and in one species of the genus Minyobates [1]. It has not been detected in Phyllobates. It occurred in four species of

Alkaloids from Amphibian Skins

73

the mantelline genus Mantella [56,61], but has not been detected in bufonid toads {Melanophryniscus) nor myobatrachid frogs {Pseudophryne). It was a major alkaloid in an extract from a Brazilian myrmicine ant of the Solenopsis subgenus Diplorhoptrum [125]. Presumably, myrmicine ants provide the dietary source for quinoHzidine 195C of frog skin.

Figure 26. Vapor-phase FTIR spectrum of 4,6-disubstituted quinoHzidine 195C. Activity. The biological activity of quinoHzidine 195C is unknown, but presumably, like other frog skin izidines, quinoHzidine 195C will prove to be a noncompetitive blocker of nicotinic receptor-channels.

3.7.2. 1,4-Disubstituted Quinolizidines A 1,4-disubstituted quinoHzidine class of alkaloids was proposed in 1993 [11], based on MS and FTIR analysis of alkaloids present in skin extracts of frogs/toads [56,86]. Subsequently about one milligram of alkaloid 217A was isolated from skin extracts from twenty specimens of a small dendrobatid frog, Dendrobates pumilio, and proton NMR analysis established the structure as that of l-methyl-4-(Z)-(l-pent-2-en-4-ynyl)quinolizidine [296]. The structures of 217A and five other alkaloids from skin extracts of frogs/ toads, currently assigned to the 1,4-disubstituted quinoHzidine class, are shown in Figure 27. Some of the alkaloids assigned to this class in a previous review [11] now appear to be 5,6,8-trisubstituted indolizidines (Section 3.6.3) or azabicyclo[5.3.0]decanes (Section 3.8). Some are now merely classified as izidines in the Appendix.

J. W. Daly, H. M. Garraffo and T. F. Spande

74

N.

:4

2071

'^^^Z

'^=^^"

217A

231A

N>

233A

CgHn (C/S-C=C) 235E' Tentative

247D

Figure 27. Structures of 1,4-disubstituted quinolizidines Alkaloids of the 1,4-disubstituted quinolizidine class exhibit an EI-MS base peak due to a-cleavage of the C-4 substituent and a diagnostic daughter ion of m/z 110 from a retroDiels-Alder process. Some imcertainty can arise since 5,8-disubstituted-6-methylindolizidines also show a peak at m/z 110. The quinolizidines often show a small diagnostic peak at m/z 84 instead of the small peak at m/z 70 frequently shown by the indolizidines. A fragmentation pathway is proposed in Scheme 11.

retro Diels-Alder

a-cleavage

m/z 152

^^N m/z 110

Scheme 11 All of the alkaloids currently assigned to the 1,4-quinolizidine class display in FTIR spectra a Bohlmann band at about 2790 cm"^ that is broader and weaker than the one for the 5,8-disubstituted and 5,6,8-trisubstituted indolizidines. The vapor-phase FTIR spectrum of 231A is depicted in Figure 28. The vapor-phase FTIR spectra of 2071,217A and 235E' (incorrectly labeled 223E' in ref. 86) have been published [56,86,281]. The quinolizidine structure proposed for 235E* [86] and shown in Figure 27 is labeled tentative since the EIMS shows a fragment ion at m/z 70 rather than the m/z 84 fragment ion expected of a quinolizidine (unpublished results). The configuration at C-1 for 2071 is proposed to be as shown

Alkaloids from Amphibian Skins

75

in Figure 27, based on comparison of vapor-phase FTIR spectra of 2071 and synthetic (-)l,10£-4-allyl-l-ethylquinolizidine having the same relative configuration as in 217A [281] (see below). Based only on EI-MS, one further alkaloid appears to be a 1,4-disubstituted quinolizidine: 257D, tentatively a l-butynyl-4-pentynyl-quinolizidine (see Appendix).

2600 2200 Wavenumber(cnvl)

Figure 28. Vapor-phase FTIR spectrum of the 1,4-disubstituted quinolizidine 231 A. Synthesis. An enantioselective synthesis has provided (-)-lR,4S,10S-4-allyl-l-ethylquinolizidine [281] diastereomeric with 2071. The FTIR spectrum was very similar, but not identical to that of quinolizidine 2071. Both showed identical Bohlmann bands, indicating 4,10Z configurations. They had slightly different retention times on GC analysis. It was concluded that 2071 was the epimer at C-1 (Figure 27) of the synthetic quinolizidine. Occurrence, The alkaloids currently assigned to the 1,4-disubstituted quinolizidine class occurred in several species of frogs of the dendrobatid genera Dendrabates and Minyabates, only rarely in frogs of the genus Epipedobates, and not at all in frogs of the genus Phyllobates [1 and unpublished data], Quinolizidines 217A and 231A occurred as major alkaloids in skin extracts of the mantelline frog Mantella baroni [56,61]. Other 1,4-disubstituted quinolizidines also occurred in mantelline frogs. The tentative quinolizidine 235E' was detected in skin extracts of the bufonid toad Melanophryniscus stelzneri [86] (see above). An isomer of 235E*, designated 235E, had been detected in a dendrobatid frog. 1,4-Disubstituted quinolizidines were not detected in myobatrachid (Pseudophryne) frogs. A dietary source for 1,4-disubstituted quinolizidines is unknown.

76

J. W. Daly, H. M. Garraffo and T. F. Spande

Activity. The biological activity of 1,4-disubstituted quinolizidines has not been investigated. Presumably, like other izidines, they will be noncompetitive blockers of nicotinic receptor-channels.

3.8.

Azabicy clo [5.3.0] decanes

The structure of a bicyclic "izidine" alkaloid 275A, first detected in the seventies in skin extracts of a Colombian dendrobatid frog, Dendrobates lehmanni, has finally been elucidated [297]. The alkaloid was first tentatively proposed as a 1,4-disubstituted quinolizidine [11]. The lack of a fragment ion at m/z 110, diagnostic for 1,4-disubstituted quinolizidines, and NMR and vapor-phase FTIR spectra then suggested a 4-methyl-6-(8-nonynyl) quinolizidine structure, but perhydro-275A was not identical in capillary GC retention time with any of the synthetic diastereomers of 4-methyl-6-nonylquinolizidine, although the EIMS and vapor-phase FTIR spectra were very similar to those of one of the synthetic diastereomers [297]. The solution to the structure came from NH3-CI-MS/MS, which revealed that the non-8-ynyl substituent was on afive-memberedring, while the methyl was on a seven-membered ring [163,297]. Thus, alkaloid 275A and four congeners were 3,5disubstituted l-azabicyclo[5.3.0]decanes with 275A being 5-methyl-3-(non-8-ynyl)azabicyclo[5.3.0]decane. The structure was confirmed by comparison of perhydro-275A with the synthetic diastereomers of 5-methyl-3-nonylazabicyclo[5.3.0]decane [297]. Perhydro-275A was identical with one of the synthetic diastereomers, but the relative stereochemistry of that diastereomer has not yet been defined. It is not the 5Z,10Z-diastereomer. The structure of 275A and tentative structures for two congeners are depicted in Figure 29. The minor isomer designated 275G that accompanies 275A appears based on the vapor-phase FTIR spectrum to have a terminal diene moiety rather than a terminal acetylene. Alkaloid 277A is proposed to have a terminal double bond, but lacking FTIR data, the structure is only tentative. A third congener, 289A, appears to have a 3-C9H13O side-chain containing a keto group, while a fourth, 293F, appears to have a 3-C9H17O side-chain (see Appendix). The EI-MS of 275A has a base peak at m/z 152, resulting from a-cleavage of the nonynyl group, and a fragment ion at m/z 260, resulting from a-cleavage of the methyl group. In contrast, the NH3-CI-MS/MS had fragment ions at m/z 112, representing the seven-membered ring bearing the methyl, and at m/z 192, representing the five-membered ring bearing the nonynyl substituent [163,297]. Some odd-mass hydrocarbon fragments were also present. Bohlmann bands were virtually absent in the vapor-phase FTIR spectrum of 275A [297]. The properties of the azabicy clo[5.3.0]decanes are tabulated in the Appendix.

Alkaloids from Amphibian Skins

77

Figure 29. Structures of 3,5-disubstitutedazabicyclo[5.3.0]decanes. The relative 3,10E stereochemistry depicted for the alkaloids is tentative, based on the likelihood that 275A is an ant alkaloid, where a ^mw^-pyrrolidine would be the most likely precursor. Synthesis. The synthesis of all four diastereomers of 5-methyl-3-nonylazabicyclo[5.3.0] decane allowed GC comparison with the perhydro-derivative of natural 275 A [297]. The third diastereomer to emerge from a GC capillary column coeluted with perhydro-275A and exhibited identical FTIR and NH3-CI-MS/MS spectra. Isolation and NMR analysis will be necessary to establish the relative stereochemistry of this third diastereomer, which probably has the partial configuration shown in Figure 29. Occurrence. The azabicyclodecane 275A and congeners were discovered in skin extracts of Dendrobates lehmanni, a montane species of dendrobatid frog found initially near Cali, Colombia. As yet, it has only been detected in widely separated populations of Z). lehmanni and as a trace alkaloid in one population each of Panamanian D. auratus, D. speciosus and D. pumilio and one population each of Costa Rican D. granuliferus and D. pumilio [1 and unpublished results]. Such disubstituted azabicyclodecanes are unknown elsewhere in Nature. Activity. The biological activity of 275A is unknown, but presumably will include activity as a noncompetitive blocker of nicotinic receptor-channels.

4.

TRICYCLIC ALKALOIDS

A remarkable range of structural classes of tricyclic alkaloids have been characterized from amphibian skin and it appears likely that further structural classes will be discovered. The alkaloids include the gephyrotoxins, which are structurally related to the decahydroquinolines, the unique cyclopenta[6]quinolizidines, which are unprecedented in Nature, the coccinellines and related tricyclics previously known from coccinellid beetles and the spiropyrrolizidines, including an alkaloid previously isolated from a millipede. In addition, the

78

J. W. Daly, H. M. Garraffo and T. F. Spande

pyridylazabicycloheptane epibatidine (Section 6.1), the dipyridylpiperidine noranabasamine (Section 6.2) and the pseudophrynamines (Section 7.1) are tricyclic alkaloids, but are distinguished by the presence of aromatic rings.

4.1.

Gephyro toxins

One of the alkaloids isolated from extracts of 1100 skins from a particularly abundant population of the Colombian dendrobatid frog Dendrohates histrionicus was initially termed HTX-D, even though it was noted that the MS fragmentation was not that of a histrionicotoxin [107]. Histrionicotoxins were the most abundant alkaloids in that same extract. The name was later changed to gephyrotoxin when the structure was elucidated by X-ray analysis [108]. The name derives from the Greek gephyra meaning bridge and the structure does "bridge" several classes of frog skin alkaloids, since gephyrotoxin has a decahydroquinoline ring, an indolizidine ring and a enyne side-chain at that time knovm only from histrionicotoxins. The name is unfortunate in one regard since gephyrotoxin is relatively non-toxic. Structures of the gephyrotoxin 287C and dihydrogephyrotoxin 289B are shown in Figure 30.

3al..»^H

287C

L

X

X'»^H

289B

Figure 30. Structures of gephyrotoxins. The configuration shown is that obtained by X-ray analysis of a single crystal of 287C«HBr and synthesized by Y. Kishi and colleagues [298]. Whether this or the enantiomer is the major alkaloid in frog skin remains in doubt (see text). The absolute configuration of gephyrotoxin present in skin extracts of Dendrobates histrionicus remains in doubt. The structure shown is that derived from X-ray analysis of a single crystal of the hydrobromide salt of gephyrotoxin isolated from skin extracts obtained in 1971 of an abundant population of Z). histrionicus from the environs of the town Guayacana in southwestern Colombia [108]. An unambigous synthesis of this enantiomer in 1980 afforded a dextrorotatory ([a]D^ +50.0°) gephyrotoxin [298]. An optical rotation had not been obtained from the 1971 sample and the gephyrotoxin proved quite labile to oxidation.

Alkaloids from Amphibian Skins

79

Thus, the gephyrotoxin from the first collection had decomposed by 1980, but an optical rotation on gephyrotoxin isolated from skin extracts obtained in 1974 from the same population ofD. histrionicus was levorotatory {[o]^ -51.5°). Thus, the gephyrotoxin from the 1974 collection appeared to be mainly the levorotatory enantiomer opposite in absolute configuration to the gephyrotoxin hydrobromide of the single crystal subjected to X-ray analysis and derived from skin extracts of a 1971 collection. A satisfactory explanation is not evident [see discussion in ref. 10]. An error in X-ray analysis, in the enantioselective synthesis or in the sign of the optical rotation appears unlikely. It may be that the major natural enantiomer is not that of the single crystal subjected to X-ray analysis. The X-ray structure for gephyrotoxin (287C) depicted in Figure 30 corresponds in absolute configuration at C-3a with the presumably biosynthetically analogous C-2 carbon in the histrionicotoxins (see Figure 11) that are always present in extracts containing gephyrotoxins. Gephyrotoxin (287C) and dihydrogephyrotoxin (289B) represent the only members of this class of alkaloids as yet detected. The MS fragmentation of gephyrotoxins is dominated by a-cleavage resulting in loss of the CH2CH2OH substituent [107,108]. The vapor-phase FTIR spectrum of gephyrotoxin (287C) has been published [11]. The Bohlmann band near 2800 cm"^ is relatively weak as it is in the corresponding 5^,9Z-3,5-disubstituted indolizidines. Brief summaries with references for physical (optical rotation) and spectral (UV, NMR) properties have been provided [10,11]. A detailed NMR analysis of gephyrotoxin is available [299]. The properties of the two gephyrotoxins are tabulated in the Appendix. Synthesis. Synthetic routes to racemic and dextrorotatory gephyrotoxin (287C), racemic dihydrogephyrotoxin (289B) and racemic perhydrogephyrotoxin were reviewed in detail in 1986 [10]. Minor amounts of the epimer at C-1 were produced in certain synthetic routes. Another stereoselective synthesis of racemic perhydrogephyrotoxin was reported in 1986 [300]. Further synthetic approaches to gephyrotoxins have appeared [301-303]. We are unaware of any more recent synthetic efforts. Occurrence. The gephyrotoxins are known only from skin extracts of dendrobatid frogs of the neotropical genus Dendrohates. Within that genus, gephyrotoxins have been detected only rarely, primarily in skin extracts from various Colombian populations of Dendrobates histrionicus, where they are always accompanied by histrionicotoxins as the major alkaloids [1]. Gephyrotoxin 287C has now been detected in skin extracts from a population of the Panamanian D. auratus [6]. Nineteen-carbon histrionicotoxins were major alkaloids in the same skin extracts. A dietary source for gephyrotoxins is unknown. However, both gephyrotoxin and nineteen-carbon histrionicotoxins were detected in skin extracts of^D. auratus raised on leaf-litter insects from Ancon Hill in Panama [6]. Gephyrotoxins, hke 3,5-disubstituted pyrrolizidines, 3,5-disubstituted indolizidines, 4,6-disubstituted quinolizidines, 3,5-disubstituted azabicyclo[5.3.0]decanes, decahydroquinolines and histrionicotoxins, could be derived by cyclizations of a precursor with a linear carbon-chain. Except for the histrionicotoxins all such alkaloid classes have been detected in myrmicine ants.

80

J. W. Daly, H. M. Garraffo and T. F. Spande

Thus, it appears likely that myrmicine ants will prove to be the dietary source for gephyrotoxins. Activity. Gephyrotoxin is relatively nontoxic and exhibits only weak activity as a muscarinic antagonist and as a noncompetitive blocker of nicotinic receptor-channels [87,133, 155,157, 304, see ref. 10]. Gephyrotoxin appears somewhat selective as a noncompetitive blocker for ganglionic-type versus neuromuscular-type nicotinic receptor-channels [133 and unpublished results].

4.2.

Cyclopenta[^]quinolizidines

A unique tricyclic alkaloid, 251F, was detected in skin extracts from two populations of the dendrobatid frog Minyabates bomhetes found in a mountainous region west of Cali, Colombia [305]. The EI-MS of alkaloid 251F and congeners were interesting in yielding an odd mass fragment as the base peak. The amount of alkaloid 251F present in skin extracts obtained in 1983 from 100 frogs from a montane population of Minyobates bombetes at that time was deemed insufficient to warrant an isolation for NMR spectral analysis. Some seven years later, 340 |Xg of alkaloid 251F were isolated by chromatography and detailed MS and NMR spectral analysis led to the 3,7,10-trimethyl-2-hydroxymethylcyclopenta[Z?] quinolizidine structure shown in Figure 31 [306]. The absolute configuration is unknown. Tentative structures of nine congeners are also shown. A structure for a tenth congener, 253G, is not proposed (see Appendix). The EI-MS fragmentation of 251F is very complex with a base peak at m/z 111. Major fragmentation pathways are proposed in Scheme 12. The vapor-phase FTIR spectrum of 251F has been reported [306]. It has a strong Bohlmann band at 2755 cm"^ The MS and NMR spectral properties of 251F and the 0-acetyl derivative of 251F have been presented along with MS data of the nine congeners depicted in Figure 31 [306]. The properties of the cyclopenta[^]quinolizidines are tabulated in the Appendix. Synthesis. The diastereoselective synthesis of 251F was recently reported [307]. The synthesis involved the enantiospecific preparation of a substituted cyclopentyl intermediate, followed by coupling to a substituted piperidine and finally a rhodium-mediated cyclization. The synthetic alkaloid was identical with natural 251F, based on GC-MS, GC-FTIR and NMR spectral analysis. Occurrence. Cyclopentaquinolizidine 251F represents the parent member of a unique structural class of alkaloids unknown elsewhere in Nature. The structure and presence of isoprenoid units suggests that, unlike many alkaloids from frog skin, it has a terpenoid origin. Cyclopentaquinolizidine 251F has been detected only from dendrobatid frogs and seemingly is limited to the tiny dendrobatid frog Minyobates bombetes from Colombia, being a major alkaloid in a population from a montane remnant of forest and a trace alkaloid

81

Alkaloids from Amphibian Skins

in a population from a stream-side gallery forest [305]. A dietary arthropod source must be extremely small to serve as a prey item for this tiny diurnal frog (17 imn snout-vent length). Activity. The biological activity of 251F is unknown and could not be investigated with the limited amount of natural alkaloid that was isolated.

CHO

,N 235H

249B

251F

251F

245A R = CHO

251J R = H

265B R = CH3

247A R = CH20H

279C R = C2H5

279B R = C2Hfe

Figure 31. Structures of cyclopenta[6]quinolizidines. All structures except 251F are tentative, being based primarily on EI-MS and analogy.

J. W. Daly, H. M. Garraffo and T. F. Spande

82

N^ m/2l12

t

/ - ^ N

m/z 250

CH2OH

v- H

y/

H

m/z 220

m/z 251

m/z 194 m/z 112

m/z 111 (base peak) Scheme 12

m/z 150

Alkaloids from Amphibian Skins

43.

83

Coccmellines

A tricyclic alkaloid, 193C, detected as a minor alkaloid in skin extracts from an introduced Hawaiian population of the Panamanian dendrobatid frog Dendrobates auratus^ was identified as precoccinelline [5], one member of a 9Z?-azaphenalene class of alkaloids from coccinellid beetles. Precoccinelline (193C) was also detected from the Costa Rican dendrobatid frog Phyllobates vittatus (unpublished results) and from a population of the bufonid toad Melanophryniscus stelzneri [86]. The MS fragmentations of precoccinelline and other coccinellines are complex with a major M^-1 fragment along with losses of hydrocarbon fragments including methyl, ethyl, propyl, butyl and pentyl. Hydrogen is the only radical that can be lost in an a-cleavage for these tricyclic alkaloids. There are several tricyclic alkaloids that have been detected in frog skin extracts that have complex MS similar to that of preccocinelline. One of these (191B) from Epipedobates silverstonei has recently been identified by MS as the beetle alkaloid propyleine (unpublished results). Another trace tricyclic alkaloid, 205B, was isolated from skin extracts of several thousand frogs of an abundant population of Dendrobates pumilio from Isla Bastimentos, Panama, and a tentative structure of a 4,6,8-trimethyl-8^-azaacenapthylene was proposed based on NMR spectral analysis [105]. Later, the Bohlmann bands of the vapor-phase FTIR spectrum of 205B led to a reevaluation of the NMR spectral data and a stereochemical revision of the structure, in which the hydrogens at C-8a and C-6 of the previously proposed structure had to be inverted [308]. The structure of 205B depicted in a review [11] is incorrect with respect to the position of one methyl group. The structure of propyleine (191B), precoccinelline (193C) and the proposed structure [308] of the tricyclic alkaloid 205B are depicted in Figure 32.

191B

193C*

205B

Figure 32. Structures of tricyclic alkaloids propyleine (191B), precoccinelline (193C) and azaacenaphthylene 205B. * Absolute configuration shown is based on that of coccinelline, the iV-oxide of precoccinelline [309]. Unfortunately, the sample of 205B has now decomposed and restrictions on collecting large numbers of dendrobatid frogs mean that any frirther studies on structure and absolute configuration of 205B are unlikely. The optical rotation was [a]D-8.5° (c = 0.5, CHCI3) [105]. Tricyclic 205B exhibited a complex EI-MS fragmentation with a major M^-1 fragment, a base peak due to loss of methyl and a series of other fragments due to loss of

84

J. W. Daly, H. M. Garraffo and T. F. Spande

higher alkyl moieties. Such complex patterns and a major M^-1 fragment are probably diagnostic for tricyclic alkaloids related in structure to the coccinelline class. The vaporphase FTIR spectra of 193C and 205B have been reported [86,308]; both have weak Bohlmann bands. A number of other tricyclic alkaloids from frog skin extracts probably have tricyclic ring systems similar to coccinellines or 205B, based on EI-MS fragmentation. All of these alkaloids are tentatively tabulated as tricyclics in the Appendix. They include 191 A, 201B, 205E, 207J, 207P, 207R, 2191, 219K, 221G, 221M, 2351, 235K, 235M, 235P and 261C. Synthesis. The synthesis of the azaacenaphthylene 205B has not been reported. Syntheses of the beetle alkaloid precoccinelline and related tricyclic alkaloids have been accomplished; the first synthesis w^as reported in 1976 [310]. Other stereoselective syntheses have been reported [311-313 and ref. therein]. Occurrence. Precoccinelline (193C) has been detected in skin extracts from only a few dendrobatid species of the genus Dendrobates [5 and unpublished results] and in skin extracts from one population of the bufonid species Melanophryniscus stelzneri [86]. The dehydro analog propyleine (191B) has recently been detected from the Peruvian frog Epipedobates silverstonei (unpublished results). Propyleine exists in equilibrium with isopropyleine [312]. Other coccinellines that are known from beetles, such as hippodamine, myrrhine, hippocasine, and 2-dehydrococcinelline, have not been detected in skin extracts. The 8Z?-azaacenaphthylene 205B has been detected only in skin extracts from certain populations of the Central American dendrobatid species D. pumilio and D. auratus [105 and unpublished results]. Neither 193C nor 205B have been detected from mantelline {MantelId) frogs or myobatrachid (Pseudophryne) frogs, nor has 205B been detected in bufonid (Melanophryniscus) toads. Other tricyclic alkaloids, probably related in structure to the coccinelline class, have been detected in skin extracts of dendrobatid frogs (201B, 205E, 207J, 207P, 207R, 2191, 221G, 221M, 235M and 235P) [unpublished results] and mantelline (Mantella) frogs (207J, 219K, 235K, 235M and 261C) [61]. Only tricyclic 2351 has been detected in bufonid (Melanophryniscus) toads [86]. The dietary sources for the perhydro-9Z>-azaphenalene 193C and presumably other related tricyclic alkaloids detected in frog skin extracts are probably small beetles of the family Coccinellidae. The ladybug beetles of the genera Coccinella, Hippodamia, Anisosticta, Coccinula, Micraspis, Myrrha, Propylaea, Calvia, Anatis and Coleomegilla contain alkaloids of the coccinelline class [314-317]. Precoccinelline and/or related alkaloids have also been reported from other beetles, namely a soldier beetle of the genus Chauliognathus [318] and a boll weevil of the gQims Anthonomus [317]. "Dimeric" alkaloids containing an octahydro-8Z>-azaacenaphthylene moiety have been reported from beetles of the coccinellid gQUQiSi Exochomus [319] and Chilocorus [320,321]. Thus, the azaacenaphthylene ring system of 205B is present in certain beetle alkaloids. Similar dimeric and trimeric alkaloids, which were named myrmicarins, have now been reported from an African myrmicine ant of the genus Myrmicaria [322,323]. In addition, a series of

Alkaloids from Amphibian Skins

85

monomeric octahydro-8Z?-azaacenaphthylenes, which were also named myrmicarins, were present as major alkaloids in these ants [322]. Such alkaloids have not been detected in frog skin extracts, nor have other classes of beetle alkaloids been detected in frog skin extracts. Activity. The biological activity of the azaacenaphthylene 205B has not been investigated. The beetle alkaloids of the coccinelline class serve as repellants to ants and at high levels even to quail [see ref. 314]. Adult ladybug beetles (Coccinella) were rejected as distasteful by captive-raised frogs of the dendrobatid species Phyllobates hicolor and were refused when reoffered as prey at a later time (unpublished observations).

4.4.

Spiropyrrolizidines

Three novel tricyclic alkaloids were isolated from skin extracts obtained in 1983 from several thousand Dendrobatespumilio frogs from Isla Bastimentos, Panama [105]. Such alkaloids had not been present in extracts from the same population collected a decade earlier. The EI-MS and NMR spectral analyses, along with the presence of an IR absorption peak at 1660 cm'^ for the major alkaloid 236 in chloroform, led to a tentative proposal of amidine structures. Amidines typically show strong absorbtion at about 1670 cm"^ However, later vapor-phase FTIR spectra indicated that the absorption peak at 1660 cm"^ was due to an impurity. A reexamination of the NMR spectral data led to revised spiropyrrolizidine oxime structures for the three alkaloids (222,236 and 252A) [324]. The structures of spiropyrrolizidine alkaloids detected in frog skin extracts are shown in Figure 33 along with the structure of the spiropyrroline polyzonimine (151B), recently detected as a trace alkaloid in skin extracts from certain populations of Costa Rican D. pumilio and one population of Peruvian Epipedohates macero (unpublished results). The absolute configurations are probably as shown. The EI-MS fragmentation of oximes 222 and 236 yields a base peak at m/z 112 (C5H8N20'*'), and m/z 126 (C6HioN20^), respectively, while oxime 252A yields a base peak at m/z 142 (C6HioN202'^). An oxime 252B has a base peak at m/z 126 and, thus, this isomer of 252A would appear to have the hydroxyl group at either C-7 or in the dimethylcyclopentyl ring [25]. Nitropolyzonamine (238) to which the spiropyrrolizidine oximes are closely related in structure, fragments to yield a base peak at m/z 82 (CsHgN"^), as does polyzonimine (151B). An apparent hydroxynitropolyzonamine 254, which also yields a base peak at m/z 82, has been detected in skin extracts of one population of Dendrobates auratus (unpubhshed results). The fragmentation pathways for 151B, 236 and 238 leading to the base peaks are proposed in Scheme 13.

86

J. W. Daly, H. M. Garraffo and T. F. Spande

151B

^^Kjo"

222

(+)-238*

-^^O^joo"'

(+)-236

<::;q
(+)-252A

Figure 33. Structures of spiropyrrolizidines and the spiropyrroline polyzonimine (151B). * Absolute configuration for dextrorotatory nitropolyzonamine (238) from millipedes [325] is as shown [326]. Alkaloids 236 and 252A are also dextrorotatory and presumably have the same absolute configuration. Vapor-phase FTIR spectra of the spiropyrrolizidine oximes 222,236 and 252A have been presented [324]. Bohlmann bands are weak (222,236) or absent (252A). An intense absorption at about 1050 cm'^ is characteristic for the N-OCH3 of 236 and 252A, while the NO-H stretching vibration of 222 results in an intense, sharp absorption at 3641 cm"^ The vapor-phase FTIR spectrum of polyzonimine (151B) and nitropolyzonamine (238) are shown in Figure 34. The proton NMR spectra of 222,236 and 252A have been presented [105, see ref. 324 for a detailed analysis of the NMR spectral data]. Optical rotations (dextrorotatory) have been reported [324]. The five spiropyrrolizidines and the spiropyrroline polyzonimine are tabulated in the Appendix. Synthesis. Synthesis of oximes 222 and 236 confirmed the structures of the natural alkaloids [327]. The route was based on the prior syntheses of the millipede alkaloids polyzonimine and nitropolyzonamine [325,328,329]. The bicyclic polyzonimine was alkylated with 3iodonitropropane followed by thermal cyclization to nitropolyzonamine, which was then oxidized to the desired ketone precursor of oximes 222 and 236 [327]. Only one oxime stereoisomer, in each case identical with the natural alkaloid, was detected after reaction with hydroxylamine or O-methylhydroxylamine.

Alkaloids from Amphibian Skins

87

m/z151 NOCH.

m/z82 (base peak) NOCH3

m/z126 (base peak)

m/z82 (base peak) Scheme 13 Occurrence. The spiropyrrolizidine oxime 236 has been detected in only a Umited number of species of dendrobatid frogs, including populations of Dendrobates pumilio, where it was discovered [105], D. auratus, D. tinctorius, Epipedobates macero, E. pulchripectus and E. tricolor (unpublished results). It has not been detected in dendrobatid frogs of the genera Minyobates or Phyllobates. It has been detected in one species of mantelline {Mantelld) frogs [61] and in one species of bufonid (Melanophryniscus) toads [86]. The oxime 252B was detected in one population of a myobatrachid {Pseudophryne) species [25]. Nitropolyzonamine (238) has now been detected in one Panamanian population of Dendrobates auratus, and the spiropyrroline polyzonimine in three populations of D. pumilio and one population of Epipedobates macero (unpublished results). Undoubtedly, such spiropyrrolizidines and polyzonimine are sequestered into skin by the frogs from small millipedes, whose identity remains unknown. Polyzonimine and nitropolyzonamine were previously isolated from defensive secretions of a North American millipede Polyzonium rosalbum, where the volatile polyzonimine predominated [325,328]. The emergence of spiropyrroHzidines as minor alkaloids in a population ofD. pumilio, where previously they had been absent, presumably reflects environmental factors leading to a greater availability of a dietary millipede [6].

J. W. Daly, H. M. Garraffo and T. F. Spande 756^

1600-

i

1

1200-

i

800-

^ V-N

^882.32

u

400-

0-

^

—

4C)'oo

/

1622.12

1

1467.96 h

1 ^..

32'00

3600

151B

28*00

.1381.02

iUW^..^V>V^^ 24*00 2000 Wavenumber(cm-I)

16'00

'

12'00

'

860

2965.37

8 i

800-1

"

€

^ 238

36'00

^

325o

'

28!5o

' 24^0 ' 255o Wavenumber(cm-1)

^

1600

860

Figure 34. Vapor-phase FTIR spectra of polyzonimine (151B) and nitropolyzonamine (238). Activity. The spiropyrrolizidine oximes 222 and 236 and nitropolyzonamine are noncompetitive blockers of nicotinic receptor-channels with selectivity for the ganglionic-type compared with the neuromuscular-type [130]. Both nitropolyzonamine and oxime 236 inhibited radioligand binding at brain sigma receptors with submicromolar potencies. Nitropolyzonamine is a weak inhibitor of bombesin-binding and had no effect on radioligand binding to a

Alkaloids from Amphibian Skins

89

variety of other receptors. The bicycUc polyzonimine of millipedes is an effective topical irritant to ants [328].

5.

MONOCYCLIC ALKALOIDS

Monocyclic alkaloids occur only rarely and usually as trace constituents in frog skin extracts. The sole exceptions are rra«5-2-butyl-5-pentylpyrrolidine (197B), a major alkaloid from one Colombian population of the dendrobatid frog Dendrobates histrionicus [209], and c/5',c/^-4-hydroxy-2-methyl-6-nonylpiperidine (241D) from a Panamanian montane species of the dendrobatid frog, D. speciosus [60]. a,a'-Disubstituted pyrrolidines and piperidines are vs^ell-known venom constituents in myrmicine ants, v^here they have been proposed as precursors of the ant pyrrolizidines, indolizidines, and decahydroquinolines [124,197,250]. Ants are a major prey item for dendrobatid frogs, particularly for frogs of the genus Dendrobates [275-278].

5.1.

Pyrrolidines

Pyrrolidine 197B, a major alkaloid in one Colombian population of Dendrobates histrionicus, was identified as /ra«5"-2-butyl-5-pentylpyrrolidine by GC comparison with a mixture of the synthetic cis- and rra«5-pyrrolidines [209]. The EI-MS fragmentation was dominated by a-cleavages to yield ions at m/z 140 and 126. Based on MS data, a minor monocyclic alkaloid 225C from the same extract was proposed to be a 2-butyl-5-heptylpyrrolidine. Seven other monocyclic alkaloids, detected as trace components of alkaloid fractions from frog skin extracts, are proposed to be 2,5-disubstituted pyrrolidines, based primarily on MS analysis. The structures of pyrrolidines from frog skin extracts are presented in Figure 35. The absolute configuration oitrans-\91B was determined by chiral GC comparison with the synthetic enantiomers [330]. The absolute configuration at C-2 of trans'\91^ is the same as that at C-3 of the indolizidine 195B (See Section 3.6.1). Vapor-phase FTIR spectra can be diagnostic for cis- and /raw^-pyrrolidine isomers. Thus, the presence {cis) or absence (trans) of a small Bohlmann band near 2797 cm"^ in 2,5disubstituted pyrrolidines distinguish the two stereoisomers, even though the Bohlmann band in the cz5-pyrrolidine is often very weak [160]. The A^-methyl derivatives, conveniently prepared by Eschweiler-Clarke methylation, show a significantly stronger Bohlmann band for the c/5-isomer compared with the trans. The vapor-phase FTIR spectrum oi trans197B is depicted in Figure 36. The vapor-phase FTIR spectra of cis- and trans-2-h\xty\-5heptylpyrrolidine, which would correspond with 225C, have been presented along with FTIR spectra of iV-methyl derivatives [160]. The properties of the nine 2,5-disubstituted pyrrolidines detected in frog skin extracts are tabulated in the Appendix.

J. W. Daly, H. iM. Garraffo and T. F. Spande

90

N H 183B

(+)-frans-197B

9r^15

'C7H-13

223N

9"^ 17

277D

Figure 35. Structures of 2,5-disubstituted pyrrolidines. * Absolute stereochemistry as shown. Synthesis. The 2,5-disubstituted pyrrolidines of myrmicine ant venoms have provided synthetic targets for several laboratories. The present review will cite only those leading to frog skin alkaloids. Syntheses of racemic cis- and/or trans-197B have been reported [331,332]. Two enantioselective syntheses of one or both of the enantiomers oftranS'197B have been reported [330,333]. Chiral GC comparison of the two enantiomers with natural trans'1973 from frog skin established the natural alkaloid as the dextrorotatory enantiomer [330]. Racemic trans-llSC [332] and racemic cis- and trans-225C [334] have been synthesized. Enantioselective syntheses of both enantiomers of c/5-225C and both enantiomers of trans-225C have been reported [335]. Both cis- and trans'225li have been synthesized [331] as have cis- and trans'2531 [336]. Occurrence. 2,5-Disubstituted pyrrolidines occur relatively rarely in frog skin extracts and then usually as trace components. The one exception is trans-197B, which was a major alkaloid in skin extracts from one population of Dendrobates histrionicus from near El Valle, Choco, Colombia [209]. Pyrrolidine 197B was a minor or trace alkaloid in skin

91

Alkaloids from Amphibian Skins 2932.45

2962.43

frans-197B

34'00

3000

2600 2200 Wavenumber(cm-I)

Figure 36. Vapor-phase FTIR spectrum of pyrrolidine tranS'191B. extracts from populations on a small mountain. Altos de Buey, only 25 kilometers distant. Pyrrolidine 197B was a major alkaloid in skins of two specimens of D. granuliferus from western Costa Rica, while being a trace alkaloid in other specimens or extracts of this species. Pyrrolidines in trace amounts, including 197B, have been detected in skin extracts of some populations of D. pumilio (unpublished results). Pyrrolidine 223N has been detected in D. auratus and one mantelline {Mantella) species. The cis- and trans-isomexs of 225H have been detected in one population of Z). pumilio and one mantelline {Mantella) species. It is perhaps not surprising that pyrrolidines are neither common nor major alkaloids in dendrobatid frog skin extracts, since pyrrolidines are accumulated very poorly into skin when fed to a dendrobatid frog on alkaloid-dusted fruit flies [7]. 2,5-Disubstituted pyrrolidines are major alkaloids in venoms of myrmicine ants of the genus Monomorium [196,197,336-340]. They have also been detected from myrmicine ants of the genus Megalomyrmex [198,341] and the Solenopsis subgenera Diplorhoptrum [195] and Solenopsis [342]. The /ran^-isomers predominate although traces of c/^-isomers have been detected. Pyrrolines also occur in ants. The occurrence of pyrrolidines and other alkaloids in ants has been reviewed [343-345]. Of the nearly twenty 2,5-disubstituted pyrrolidines reported from myrmicine ants only five (rraw5-197B, 225C, cis- and trans225H, 2531) have been detected in frog skin extracts. The remaining four from frog skin extracts have not yet been reported from ants. Activity. The 2,5-disubstituted pyrrolidines of the myrmicine ants serve as repellants for other ants and have insecticidal activity [336,339,340]. The synthetic mixture of cis- and

92

J. W. Daly, H. M. Garraffo and T. F. Spande

trans'197B was a noncompetitive blocker of nicotinic receptor-channels with apparent selectivity towards a ganglionic subtype [133].

5.2.

Piperidines

Piperidine 241D, a major alkaloid in one population of a Panamanian montane dendrobatid frog, Dendrobates speciosus, was isolated in sufficient quantities from extracts of 258 frogs collected in 1985 for structure determination by MS and NMR spectral analysis [60]. It represents the only 2,6-disubstituted piperidine detected in frog skin extracts at more than trace levels. The EI-MS fragmentation was dominated by a-cleavage of the nonyl sidechain to yield a base peak at m/z 114. a-Cleavage of the methyl group was minor. Based on MS data, a minor congener, 255A appears to have a keto group in the nonyl side-chain. Another minor congener, trans-213, has been detected. In addition, other monocyclic alkaloids detected as trace components in alkaloid fractions from frog skin extracts can be proposed based on EI-MS to be 2,6-disubstituted piperidines. The structures of the 2,6disubstituted piperidines detected in frog skin extracts are presented in Figure 37. Many yield on EI-MS a fragment ion at m/z 98 as the base peak indicating that such alkaloids are 2-methyl-6-substituted piperidines. Most of the piperidines detected in frog skin extracts were present in trace levels too low for satisfactory vapor-phase FTIR spectra. In cases where the FTIR spectra could be obtained, the absence (trans) or presence (cis) of a small Bohlmann band at 2780 cm"^ was diagnostic of the configuration [160]. The vapor-phase FTIR spectrum of synthetic cis241D has been presented [347] as have the FTIR spectra of cis- and trans-l-mQthyl-Sundecylpiperidine [160], which would correspond with 253J. The optical rotation [a]D^ +39° (c = 0.2, CH3OH) of natural 241D was measured on about one milligram of free base [60]. The properties of the nineteen 2,6-disubstituted piperidines detected in frog skin extracts are tabulated in the Appendix. In addition, the alkaloid 211J is tentatively tabulated as an A^-methyl-2,6-disubstituted piperidine (see Appendix). Unclassified alkaloids 205C and 207M might also be A/^-methylpiperidines. Synthesis. The 2,6-disubstituted piperidines of myrmicine ant venoms have been the target for extensive synthetic efforts, which are beyond the scope of this review [see refs. 343-345 for lead references]. A stereoselective Mannich-type synthesis involving a one-stage reaction of an a,(3-unsaturated ketone, an aldehyde and ammonia followed by reduction of the 2-methyl-6-nonyl-4-piperidone provided racemic c/5,cw-4-hydroxy-2-methyl-6-nonylpiperidine identical with the natural alkaloid 241D [347]. An enantioselective synthesis of 241D utilized a lipase-mediated desymmetrization of a c/5,cz5-2,6-diacetoxy-methyl-4hydroxypiperidine with appropriate protection on the 4-hydroxyl and piperidine amine groups [346]. The resulting monoacetoxymethyl product was converted to an oxazolidinone with COCI2, followed by reduction of an 0-mesylate derivative to yield a 2-methyl group.

Alkaloids from Amphibian Skins

93

Deprotection, followed by oxidation and a Wittig reaction on the resulting aldehyde provided, after final deprotection, the dextrorotatory hydrochloride sah of 241D. Both the synthetic salt and natural free base were dextrorotatory, but an assignment of absolute configuration remains tentative, until rotations of free bases of synthetic and natural compounds are compared.

hi

N H

221L

223K

"C9H17

N H

C10H19

237J

N ' 'C9H17O H (C=0) C/S-239L

s^"

a

N- -C9H17O H (C=0)

tranS'239L

Figure 37. Structures of 2,6-disubstituted piperidines. *The absolute configuration shown is that of a dextrorotatory salt of synthetic 241D [346]. The free base of natural 241D is also dextrorotatory [60].

J. W. Daly, H. M. Garraffo and T. F. Spande

94

N'

"C8H15O2

241G

N H

C9H17O

(C=0)

255A

H

C11H21O

(C=0)

C/S-267K

.a •N'

N H

'C11H21O

trans-267K

(C=0)

„..a N H

C11H23O

(OH)

trans-269C Figure 37 (continued)

The absolute configuration of the ant piperidine solenopsin B (trans-l-rnQthyl-Sdodecylpiperidine) has been established by an enantiomeric synthesis of the 2R,6R enantiomer, followed by chiral HPLC comparison of racemic solenopsin B, natural solenopsin B and the synthetic 2R,6R-solenopsin B [348]. Another study demonstrated that the configuration of the ant trans-pipcndinQS is always 2R,6R and that the ant cz.s-piperidines are always 2R,6S [349]. The absolute configuration of the 2,6-disubstituted piperidines from frog skin have not been studied. By analogy to the ant alkaloids, the configurations of transand ci^-piperidines from frog skin are shown as 2R,6R and 2R,6S, respectively, in Figure 37. The frog skin alkaloid 253J corresponds with the common ant alkaloid solenopsin A (2methyl-6-undecylpiperidine). Both cis- and trans-isornQTS of solenopsins A and B occur in ant venoms. A number of enantioselective syntheses of solenopsins A and B have been reported [166,348-352 and references therein]. The frog skin alkaloid 2251 appears to correspond with a relatively common ant alkaloid 2-methyl-6-nonylpiperidine. Both cisand trans-isomers occur in ant venoms and both have been synthesized [250]. The frog skin alkaloid 223K appears to correspond with an ant alkaloid 2-methyl-6-(4-nonenyl)piperidine, although the position of the double bond is unknown in 223K. Both cis- and trans-isomQis occur in ant venoms and both have been synthesized [279].

Alkaloids from Amphibian Skins

95

Occurrence. 2,6-Disubstituted piperidines occur rather rarely in frog skin extracts. It appears likely that two factors contribute to the rare occurrence of such alkaloids even as trace constituents. First, piperidines, such as 2-methyl-6-nonylpiperidine, are accumulated very poorly into skin of dendrobatid and mantelline frogs compared with other alkaloids [7,8]. Second, 2,6-disubstituted piperidines occur mainly in fire ants of the Solenopsis subgenus Solenopsis [345]. Such ants, which can inflict painful stings, will undoubtedly be avoided as prey items by small frogs. However, 2,6-disubstituted piperidines also occur, but less commonly in thief ants of the Solenopsis subgenus Diplorhoptrum and in myrmicine ants of the genus Monomorium. Such ants do represent possible prey items for frogs. Only one piperidine, 241D, has been detected as a major alkaloid in a frog skin extract; it was a major alkaloid in only one population of the Panamanian montane dendrobatid frog Dendrobates speciosus [60]. It was a minor or trace alkaloid in other populations. Piperidine 241D occurs in other dendrobatid species only rarely and mainly as a trace alkaloid in various populations of Central American D. pumilio [1 and unpublished results]. It has not been detected in bufonid (Melanophryniscus) toads, mantelline (Mantelld) frogs or myobatrachid (Pseudophryne) frogs. The dietary source of 241D and other 2,6-disubstituted-4-hydroxypiperidines (213, 255A) is of some interest. With a 4-hydroxyl group, 241D has no piperidine counterpart in ant alkaloids. Of the more than twenty 2,6-disubstituted piperidines found in ants, only three (223K, 2251 and 253 J) appear to be present in frog skin extracts, where they are trace constituents. Activity. The 2,6-disubstituted piperidines solenopsins A and B from ants have a wide range of toxic effects towards other insects, bacteria, fungi, plants and vertebrates [345 and references therein]. The molecular mechanisms of action remain poorly defined. The 2methyl-6-undecylpiperidines, which correspond with solenopsin A (253J), were potent noncompetitive blockers of nicotinic receptor-channels of the neuromuscular-subtype [158,353] and the ganglionic-subtype (unpublished data). The cis- and trans-isomers had equivalent activities as blockers at the neuromuscular-subtype [353]. Piperidine 24ID also was a potent noncompetitive blocker of neuromuscular and ganglionic-subtypes of nicotinic receptor-channels in cultured cells[133].

6.

PYRIDINE ALKALOIDS

Two structural classes of pyridine alkaloids have been detected in frog skin extracts. These are the epibatidines, represented by epibatidine and a trace A^-acyl congener, and a bipyridylpiperidine, noranabasamine. Epibatidines have been detected only in three species of the dendrobatid genus Epipedobates from Ecuador and Peru and always in trace amounts. Noranabasamine has been detected only in skin extracts of the three Colombian poison dart frogs of the genus Phyllobates. Epibatidine is a potent nicotinic agonist and analgetic. Both epibatidine and noranabasamine are related in structure to the plant alkaloid nicotine. A dietary source is not known for epibatidine nor for noranabasamine.

96

6.1.

J. W. Daly, H. M. Garraffo and T. F. Spande

Epibatidine

In 1974, skin extracts were obtained from a few specimens of a lowland population of Epipedobates anthonyi and a highland population of Epipedobates tricolor on the Pacific versant of the Andes in Ecuador; these frogs are now considered to be one species, Epipedobates tricolor. Both skin extracts contained a trace alkaloid that elicited a Straubtail response in mice, a response typical of opioid-class alkaloids. The objective of a subsequent field trip in 1976 was to obtain ample skin extracts for isolation and structure elucidation of the trace alkaloid. The lowland population from a cacao grove had disappeared and skin extracts of frogs from a nearby banana plantation proved to have no alkaloids at all. Extracts from 750 skins of the highland population provided about sixty milligrams of alkaloids from which less than a milligram of the trace Straub-tail alkaloid was obtained. Initially designated alkaloid 208/210 [10], the trace alkaloid proved to be 200-fold more potent than morphine as an analgetic, but acted through a non-opioid mechanism, since neither the Straub-tail response nor the analgesia were blocked by an opioid receptor antagonist naloxone [354]. The apparent molecular ions at m/z 208 and 210 (ratio 3:1) contained a chlorine atom and had an empirical formula of CnHi3N2Cl. The alkaloid was suspected to contain a pyridyl ring and two further rings containing the second nitrogen as a secondary amine. An amidine-like structure was also considered with one nitrogen contributed by the pyridine ring [10]. The sensitivity of NMR spectroscopy was not sufficient at that time for a definitive analysis. Further extracts obtained in 1979 and 1984 from different locales in the highland area contained minimal amounts of 208/210 and in 1984 all brightly colored frogs of the family Dendrobatidae were designated by an international convention (CITES) as "threatened", thereby limiting any further collections to only a few frogs, in spite of the abundant, widespread and non-threatened status of many dendrobatid species, including Epipedobates tricolor. Thus, the remaining sample of 208/210 of about 700 ^ig represented the sole means to ever elucidate the structure of this potent non-opioid analgetic. In 1990, the remaining 208/210 from the 1976 collection was converted to the 7V-acetyl derivative, thereby allowing quantitative separation from some contaminating tertiary amines of the pumiliotoxin class [354]. NMR spectral analysis of the iV-acetyl derivative now defined the structure of alkaloid 208/210, which was given the name epibatidine in recognition of the Ecuadoran dendrobatid frog Epipedobates tricolor from which it was obtained. A more detailed history of the discovery and investigation of epibatidine has been presented [355]. Epibatidine was eA:o-2-(6-chloro-3-pyridyl)-7azabicyclo[2.2.1]heptane (Figure 38). The absolute configuration (1R,2R,4S) of natural epibatidine was established by chiral HPLC comparison with synthetic (+)- and (-)epibatidine; the absolute configuration of the latter was defined by X-ray analysis [356,357]. The free base of the synthetic epibatidine, corresponding to the natural enantiomer, is levorotatory ([a]D^ -6.7° (c = 0.82, HCCI3)), while salts are dextrorotatory (oxalate [a]D'^

Alkaloids from Amphibian Skins

97

+37.3° (c = 0.44, CH3OH), hydrochloride +34.7° (c = 0.36, CH3OH)) [357-359]. A similar reversal of rotation occurs when natural (-)-nicotine is converted to a salt.

H-Epibatidine Figure 38. Structure of epibatidine (208/210). Absolute configuration as shown. The vapor-phase FTIR spectrum of epibatidine has been presented [354]. There are strong absorption bands at 1428 and 1112 cm'^ owing to the chloropyridine ring. The UV spectrum in methanol exhibits a X^nax at 217 nm and a broad shoulder at 250-280 nm typical of a pyridine. The EI-MS of epibatidine is dominated by a base peak at m/z 69. A fragmentation pathway has been proposed (Scheme 14). The proton NMR spectrum of A^-acetylepibatidine has been presented [354]. Trace amounts of an apparent A^-acylated congener (308/ 310) of epibatidine have been detected. The two epibatidines are tabulated in the Appendix.

CKM

•o- m/z 69 (base peak)

'/

o fit

m/z 68

V% T m/z 179/181

m/z 208/210

C k ^N.

Ck ^N.

m/z 140/142 Scheme 14

98

J. W. Daly, H. M. Garraffo and T. F. Spande

Synthesis. In 1993 within a year of the report of the structure elucidation [354], five syntheses of epibatidine were reported [357-362]. Both the racemate and the enantiomers of epibatidine were prepared. These initial syntheses and some of the six further syntheses reported in 1994 [363-369] have been reviewed in detail [370-373]. A further eleven synthetic routes to epibatidine have now appeared [374-383]. Most routes have utilized cycloaddition reactions of pyrrole derivatives with activated dienophiles or an intramolecular cyclization of 4-aminocyclohexanol derivatives. In addition to epibatidine, a variety of analogs have been synthesized. These include replacement of the 6-chloro-3-pyridyl ring with 2-chloro-3-pyridyl [367], 6-methyl-3-pyridyl [359], 6-bromo-3-pyridyl [384], 6-fluoro3-pyridyl [384], 6-A^,A^,A^-trimethylammoniumyl-3-pyridyl [378], 6-iodo-3-pyridyl [359,384386], 3-pyridyl [359,367] and 6-methyl-3-isoxazolyl [387] rings. An alternate numbering of the pyridyl ring of epibatidine and analogs; i.e., 2-chloro-5-pyridyl, has also been used. Synthetic studies in which the electronegative pyridyl or isoxazolyl ring necessary for nicotinic activity of such compounds has been replaced with either an endo- or an exo-diVyX ring have been reported [388,389]. The endo-\somQv of epibatidine has been reported [358, 361] as have endo- and exo-3-pyridy 1-1',4'-dimethyl-analogs of epibatidine [390]. Analogs of epibatidine with carboethoxy and other substituents at C-3' of the azabicycloheptane ring have been reported [365]. A pyridyl analog of epibatidine with a hydroxy 1 at C-2' has been reported [367]. The azabicyclo[2.2.1]heptane system of epibatidine was synthesized for the first time about thirty years ago [391,392] and more recently in 1995 [393]. Analogs in which the azabicycloheptane ring of epibatidine has been replaced with 8-azabicyclo[3.2.1] octane, [380,394-396] or 9-azabicyclo[4.2.1]nonane rings [396] have been synthesized and were referred to as homoepibatidines and Z?w-homoepibatidines, respectively. 2-Pyridyl and 4-pyridyl analogs of epibatidine in which the nitrogen of the azabicycloheptane ring was replaced with oxygen have been reported [397]. Occurrence. The alkaloid epibatidine as yet has only been detected in Nature from a few populations of Ecuadoran frogs of the dendrobatid genus Epipedobates. The highest levels of this trace alkaloid were detected in a lowland population ofE. tricolor from a cacao plantation and in a highland population from a roadside, drainage area in the Pacific versant of the Andes in Ecuador. Highland populations from nearby streamside sites had much lower levels, while lowland populations from a banana plantation had none. Epibatidine has been detected at very low trace levels in extracts from streamside populations of another Pacific coast species, E. espinosai of Ecuador, in one Peruvian, rain-forest population of an Amazonian species, E. pictus, and in one population of a highland Amazonian species, E. silverstonei [354 and unpublished results]. It is possible that epibatidine with structural features reminiscent of both nicotine and tropane alkaloids may be accumulated from small arthropods that have obtained it from a plant source. Activity. The mechanism underlying the potent non-opioid analgetic activity of epibatidine [354] remained unknown until synthetic material became available in 1993. Epibatidine

Alkaloids from Amphibian Skins

99

was then shown to be a potent full agonist at ganglionic- and neuromuscular-type nicotinic receptors [398] and to compete with agonists [-^Hjhicotine, [^HJcytisine and [^H]ABT-418 for binding sites on brain neuronal-type (a4P2) nicotinic receptors [398-406]. Epibatidine had picomolar affinities for the nicotinic a4p2 receptor. Epibatidine also was a potent antagonist versus binding of [^H]nicotine, [^H]cytisine, [^H]acetylcholine or [^^^I]abungarotoxin to other subtypes of nicotinic receptors, including neuromuscular (ai(Jiy6), ganglionic (a3P4) and neuronal (ay) subtypes. It was potent versus binding at various nicotinic subtypes (aiPiyS, Oi^^i^ ^Z^A, OC4P2, ay, ag) expressed in Torpedo electroplax [405,407,408], cultured cells [355,358,407-413], brain membranes [405,414] ovXenopus oocytes [408]. The inhibition by epibatidine of binding of [^H]nicotine to ag nicotinic receptors was biphasic [408]. The affinity of epibatidine for nicotinic receptors appeared greatest at neuronal a^ receptors and ganglionic a3 receptors and much less at neuromuscular ai receptors and neuronal a-] receptors. Remarkably, the (+)- and (-)-enantiomers of epibatidine had similar potencies for nicotinic receptors, both in binding assays and in functional and behavioral responses. Epibatidine was many-fold more potent than (-)nicotine at all nicotine receptor subtypes. The inhibition of [^H]nicotine binding to human temporal cortical membranes by epibatidine appeared biphasic, suggestive of binding to two subtypes [414]. The major lower affinity component (a4|J2) was reduced in membranes from temporal cortex of humans with Alzheimer's disease. Epibatidine was only about 2fold more potent than (-)-nicotine in blocking binding of [^H]imidacloprid to nicotinic receptors in brain membranes of the fly Musca domestica and had only a "moderate potency" as an insecticide [415]. Epibatidine had low affinity or no effect on binding of radioligands to a variety of other receptors and ion channels [398,400-405]. Epibatidine proved not only to have high affinity for nicotinic receptors, but in functional assays with cultured cells was a full agonist [398]. Functional activity of epibatidine has been assessed with synaptosomes [407,416], cultured cells [355,398,405, 407,410,417,418], Xe«opw5 oocytes [407-409,417,419,420], retina [413], ganglia [421] and hippocampal neurons [422,423]. Such assays have involved measurement of cation flux or current through nicotinic receptor-channels or release of neurotransmitters. Epibatidine appeared functionally to be most potent at neuronal a^ receptors and ganglionic a^ receptors and much less potent at neuromuscular ai receptors and neuronal a? receptors, consonant with data from binding assays. In Xenopus oocytes, (+)- and (-)-epibatidine were very potent agonists for a3P2, CX3P4, a4P2 and ag nicotinic receptors with EC50 values of 10-70 nM, while for Torpedo and human aiPi76 nicotinic receptors the epibatidines had EC50 values from 1600 to 16,000 nM [408]. For chicken and human a-j nicotinic receptors, expressed in Xenopus oocytes, the epibatidines had EC50 values from 1100 to 2200 nM. In Xenopus oocytes with rat nicotinic receptors, racemic epibatidine was most potent for a4P2 receptors (EC50 16 nM), where it appeared to be a partial agonist, less potent for a3P2 and a3p4 receptors (EC50 100 nM), even less potent for a2p2 receptors (EC50 300 nM) and least potent for a? receptors (EC50 1000 nM) [409]. In rat hippocampal neurons, racemic epibatidine was very potent (EC50 ca. 20 nM) in eliciting an a4p2-mediated current, but was

100

J. W. Daly, H. M. Garraffo and T. F. Spande

much less potent (EC50 ca. 3500 nM) in eliciting an av-mediated current [422]. Epibatidine has potent activity as a desensitizing agonist at nicotinic receptors [410,413,416]. Epibatidine elicited depolarization of rat vagus nerve, constriction of guinea pig ileum and contraction of frog rectus abdominis muscle [404,417]. Neuronal, ganglionic and neuromuscular-type nicotinic receptors, respectively, were involved. Behaviorally, both (+)- and (-)-epibatidine were extremely potent analgetic agents that appeared to act through both central and peripheral nicotinic receptors [355,398,407,410, 424-430]. The analgetic and other effects of epibatidine were antagonized by ganglionic nicotinic blockers, such as mecamylamine, pempidine and chlorisondamine [355,398,404, 410,427] and in one study by peripherally acting nicotinic blockers, such as hexamethonium and decamethonium [410]. The analgetic effects were either reduced or unaffected by the a4P2 receptor antagonists erysodine and dihydro-p-erythrodine [398,410,428] and not by the a-i receptor antagonist methyllycaconitine [403,428]. Another nicotinic ligand, a-lobeline, effectively blocked analgesia elicited by intrathecal epibatidine [428]. The analgetic effects of epibatidine were reported to be reduced by the a-adrenergic antagonist phenoxybenzamine and by pretreatment with the noradrenergic toxin A^-(2-chloroethyl)-A/-ethyl-2-bromobenzylamine [403]. Studies have indicated either no effect or a partial blockade of epibatidine-elicited analgesia by muscarinic antagonists [402,403]. Ibogaine effectively blocked epibatidine-elicited analgesia [429]. The opioid antagonist naloxone had no effect on epibatidine-elicited analgesia under a variety of conditions. The only exception was the antagonism of the analgetic effect of (+)-epibatidine using a chemically-induced pain model [425]. The analgetic effect of (-)-epibatidine was not antagonized by naloxone in the same pain model. The activator of L-type calcium channels Bay K 8644 potentiated the analgetic effects of epibatidine, while the L-type channel blocker nifedipine antagonized the analgetic effects [410,430]. Chronic treatment with nicotine or caffeine was reported to reduce the analgetic effects of epibatidine [410, however, see ref. 426]. Chronic treatment with (-)epibatidine caused some tolerance to the analgetic effect, while no tolerance developed after chronic treatment with the (+)-enantiomer [401]. Epibatidine has other behavioral effects besides those involved in sensory pathways for pain perception. Epibatidine caused locomotor depression [401,405,426,431], reduced body temperature [401,405,426], elicited pressor effects on blood pressure [407,421,428] and caused respiratory stimulation [421]. The initial tachycardia elicited by intrathecal epibatidine correlated with an initial irritation response, while a subsequent bradycardia correlated with analgesia [428]. a-Lobeline selectively reduced the analgetic response, while methyllycaconitine and an antagonist for glutamatergic NMDA receptors selectively reduced the pressor, tachycardie and irritation reponses [428]. Tolerance to intrathecal racemic epibatidine developed [428]. The analgesia, but not the locomotor depression and hypothermia elicited by epibatidine, was effectively antagonized by chlorisondamine [427]. Epibatidine has been reported in one study to cause locomotor stimulation in rats and ipsilateral turning in rats with unilateral dopaminergic lesions [432]. Both effects were blocked by dopamine receptor antagonists. Epibatidine was highly toxic at doses only somewhat greater than those eliciting analgesia [387,428]. All the diverse effects of epibatidine are also evoked to

Alkaloids from Amphibian Skins

101

varying degrees by nicotine, and appear to be due to agonist activity of epibatidine at nicotinic receptors. In drug discrimination studies, epibatidine generalized to nicotine [401,407,433]. In a radial-arm maze, epibatidine had modest effects, both positive and negative, on performance of rats [434]. However, epibatidine has also been stated to have no anxiolytic or cognitive-enhancing properties [407]. Chronic treatment with epibatidine caused a reduction in weight gain in obese Ob/Ob mice and weight loss in normal mice [435]. Activation of nicotinic receptors in brain enhances neurotransmitter release and epibatidine, like nicotine, enhanced synaptosomal release or turnover of dopamine [405,432, 436], norepinephrine [432,436-438] and acetylcholine [439]. The epibatidine-elicited release of dopamine from striatal synaptosomes and acetylcholine from hippocampal synaptosomes was reduced by the nicotinic a4p2 selective antagonist dihydro-P-erythroidine and by the relatively nonselective antagonist mecamylamine, while release of norepinephrine from hippocampal and thalamic neurons was reduced only by mecamylamine. Intrathecal epibatidine elicited a mecamylamine-sensitive release of spinal aspartate and glutamate [440]. Epibatidine had no effect on release of serotonin from hippocampal slices [441,442]. The structures of (+)- and (-)-epibatidine have been compared with (-)-nicotine and other nicotinic agonists in efforts to delineatefiirtherinteractions of agonists with a nicotinic pharmacophore [355,399,419,443,444]. Such modeling suggests that the distance between the two nitrogens in epibatidine (5.5 A) may be optimal and is slightly longer than in nicotine (4.9 A). The lack of enantioselectivity of the epibatidines suggests that the hydrophobic azabicycloheptane ring interacts equally well with the nicotinic receptor on either side of the bicyclic moiety. The effect of an A^-methyl group, either negative, neutral, or positive, on affinity of nornicotine, epibatidine and anatoxin, respectively, may reflect different orientations of the TV-methyl group with respect to the nicotinic receptor [443]. The high affinity of epibatidine and anatoxin may reflect steric volume, which is relatively large compared with nicotine and/or lack of conformational freedom, again relative to nicotine [399,419,443]. However, such conclusions are confounded by the recent development of 3-(2-azetidinylmethoxy)pyridines, which have picomolar affinities for neuronal a4P2 receptors, but do not have steric bulk and have great conformational freedom [445]. One conformer of such pyridyl azetidines can be superimposed on epibatidine. A chloropyridyl azetidine, ABT594, has broad spectrum analgetic activity [446] and, probably because of reduced affinities for neuromuscular and ganglionic nicotinic receptors, such azetidines exhibit little of the cardiovascular and toxic effects of epibatidine [445,446]. Although there is no evidence implicating opioid receptors in the actions of epibatidine, a comparison of molecular models of epibatidine and morphine has been reported [447]. Structure-activity relationships for the analogs of epibatidine have been studied to a limited extent. The A^-methyl analogs of (+)- and (-)-epibatidine are, respectively, similar to or more potent than the parent alkaloid at neuronal, ganglionic and neuromuscular nicotinic receptors [410]. The A^-methyl analog also was equipotent with epibatidine in analgetic assays [424]. The A^-acetyl derivative of epibatidine was inactive in nicotinic assays [355, 415,424]. An A^-allyl analog was 200-fold less potent versus [^H]nicotinic binding to a4p2

102

J. W. Daly, H. M. Garraffo and T. F. Spande

receptors [444]. Replacement of the chloropyridyl moiety of epibatidine with a 2-(3methyl-5-isoxazolyl) moiety provided epiboxidine, an analog ten-fold less potent than epibatidine at neuronal a4P2 receptors, equipotent at ganglionic a3p4(5) receptors and fivefold less potent at neuromuscular receptors [387]. Epiboxidine was about ten-fold less potent as an analgetic than epibatidine, but was also much less toxic. The chloro substituent on the pyridine ring did not appear critical to the activity of epibatidine, since the deschloro analog of (+)-epibatidine was similar in potency to epibatidine at neuronal, ganglionic and neuromuscular nicotinic receptors [398]. However, deschloroepibatidine was reported to be much less potent than epibatidine in an analgetic assay [367]. The chloropyridyl analog of nicotine appeared about four-fold more potent than nicotine at neuronal nicotinic receptor [399]. Replacement of the chloro substituent of (+)-epibatidine with a methyl group reduced affinity for neuronal (a4P2) receptors by only two-fold, while replacement with iodine reduced affinity by twenty-fold [398]. The iodo analogs of (+)- and (-)-epibatidine exhibited no enantioselectivity and were about nine-fold less potent than epibatidine in binding assays for neuronal (a4P2) nicotinic receptors [386]. Replacement of the chloro substituent with fluorine had no effect on affinity in binding assays for neuronal (a4p2) receptors [385]. The epibatidine analog with the chlorine in the 2-position of the pyridyl ring appeared more than 30-fold less potent as an analgetic than epibatidine [367]. Epibatidine analogs with an A^-methylpyridonyl moiety in place of the pyridyl moiety were inactive [367]. Both [^H]epibatidine [408,428,437,448-456] and [^H]deschloroepibatidine [385,457] have been used as radioligands for nicotinic receptors. The KD values for the major rat brain neuronal (a4P2) receptors were similar at about 20 pM [385,449]. A lower affinity binding site (KD 360 pM) was also detected in brain membranes [449]. [^H]Epibatidine appears to be an effective radioligand for a3P25 o^3P20C5, a3p4, a3P4a5, and a4P2 nicotinic receptors [408,449,453,455,456]. An analog in which the chlorine of epibatidine is replaced with fluorine-18 has been prepared and utilized as a PET scanning agent for nicotine receptors [384,385,458,459]. Analogs in which the chlorine is replaced with iodine-123 or iodine-125 have been prepared [384] and the iodine-125 analog has been utilized as a high affinity nicotinic radioligand for autoradiography [386] and as a PET scanning agent [460]. The effects on activity upon substitution or alterations of the azabicycloheptane ring of epibatidine have also been investigated. The ewJo-stereoisomer has proven inactive in nicotinic assays [415,421,424]. A deschloroepibatidine analog with a tertiary hydroxy 1 group at C-2' of the azabicycloheptane ring was inactive as an analgetic [367]. An analog containing the bicyclic ring of anatoxin-a instead of the azabicycloheptane ring of epibatidine has been synthesized [461]. The enantiomer corresponding with natural anatoxin-a was about 10-fold less active than epibatidine at a4P2 nicotinic receptors, while the other enantiomer was over 200-fold less active. One homoepibatidine (6-(6-chloro-3-pyridyl)-8azabicyclo [3.2.1] octane) was about four-fold less potent as an analgesic compared with epibatidine [380,395]. The A^-methyl derivative had a similar potency, while the A^-isopropyl derivative was less active. Another homoepibatidine (2-(6-chloro-3-pyridyl)-8-azabicyclo[3.2.1]octane) was 25- to 30-fold less active than epibatidine in binding assay at Torpedo neuromuscular-type (aipiy8) receptors and in increasing blood pressure in rats

Alkaloids from Amphibian Skins

103

[394]. The endo-stQVQoisomQY was much less active. No biological activity for a bis-homoepibatidine (7-(6-chloro-3-pyridyl)-9-azabicyclo[4.2.1]noriane) was reported [396]. A recent comprehensive review on the pharmacology of epibatidine is available [462].

6.2.

Noranabasamine.

Noranabasamine was isolated as a trace alkaloid from extracts of 426 skins of the Colombian dendrobatid frog Phyllobates terribilis [32]. The structure of this bipyridyl piperidine (Figure 39) was elucidated by NMR spectroscopy. It corresponds to the desmethyl analog of the plant alkaloid anabasamine. The EI-MS shows a base peak at m/z 84 corresponding to the piperidine moiety and a major ion at m/z 157 corresponding to a pyridylpiperidine fragment [32]. The proton- and carbon-NMR spectral assignments for noranabasamine, and UV spectral data with A^max 244 and 275 nm have been provided [32]. The optical rotation was [a]^^ -14.4° (CH3OH).

H-239J Figure 39. Structure of noranabasamine (239 J). The absolute configuration (2S) shown is based on that of the plant alkaloid anabasine (2-(3-pyridyl)piperidine), which like noranabasamine, is levorotatory [see ref. 32]. Synthesis. The pyridylpiperidine anabasine and the pyridyldehydropiperidine anabaseine, both from plants, have been synthesized [463,464], but apparently anabasamine and noranabasamine have not. Occurrence. Noranabasamine was a minor or trace alkaloid in the three Colombian species oiPhyllobates [32], but apparently has not been detected elsewhere in Nature. The A^methyl analog anabasamine occurs in the Caucasian plant Anabasis aphylla (Chenopodiaceae) along with anabasine [465]. Anabaseine (2-(3-pyridyl)-l,2-dehydropiperidine) occurs in venom glands of myrmicine ants of the genus Aphaenogaster [466]. It also occurs in a marine nematode, where it is accompanied by related bipyridyl and tetrapyridyl alkaloids [467]. A dietary source for the frog skin alkaloid noranabasamine is unknown.

104

J. W. Daly, H. M. Garraffo and T. F. Spande

Activity. The biological activity of noranabasamine has not been investigated. Other bipyridyl alkaloids and anabaseine are toxic, apparently due to agonist activity at nicotinic receptors [467]. Recently, anabaseine was shown to have potent agonist activity at neuronal, ganglionic and neuromuscular subtypes of nicotinic receptors [417]. The 3pyridylpiperidine anabasine, which is more closely analogous to the frog skin alkaloid noranabasamine, also exhibited nicotinic agonist activity. The rank order of potencies at the different subtypes of nicotinic receptors was as follows: Neuromuscular (aipiyS), anabaseine > nicotine > anabasine; neuronal (a-j), anabaseine > anabasine > nicotine; ganglionic (a3P4(5), PC 12 cell), anabaseine « nicotine « anabasine; ganglionic (rat colon), anabaseine « nicotine > anabasine. For ants of the myrmicine genus Aphaenogaster that contain anabaseine, the alkaloid acts as an attractant, but also as a feeding deterrent, being apparently distasteful to the ants [466]. Anabasine has insecticidal activity.

7.

INDOLE ALKALOIDS

The pseudophrynamines represent the one major class of indole alkaloids that occur in frog skin extracts. They occur only in Australian myobatrachid frogs of the genus Pseudophryne [24,25]. Another class of indole alkaloids are represented by calycanthine/ chimonanthine, detected only in Colombian dendrobatid frogs of the genus Phyllobates [32]. Two simple indole alkaloids, dehydrobufotenine from bufonid toads and trypargine from a hyperoliid frog, which are obviously derived from the biogenic amines serotonin and tryptamine, respectively, are not treated in the present review, nor are the spinceamines, imidazole alkaloids that are obviously derived from histamine [see refs. 10,11].

7.1.

Pseudophrynamines

Three alkaloids were isolated from skin extracts from 166 frogs of the Australian myobatrachid frog Pseudophryne coriacea [24]. These represented the indolic compounds first noted by Erspamer and colleagues in 1976 [468]. The structures of the three major alkaloids were elucidated by MS and NMR spectral analysis and chemical intraconversion. The alkaloids, which were 3a-prenylpyrrolo[2,3-Z^]indoles reminiscent in structure to physostigmines, were named pseudophrynamines [24]. The structures of the three major alkaloids, pseudophrynaminol (258), pseudophrynamine A (512) and pseudophrynamine 286A are shovm in Figure 40. Pseudophrynamines 286A and 258 also were obtained by base-catalyzed methanolysis of 512, but it seems unlikely that 286A is an artefact formed by methanolysis of 512 [see ref 24]. A number of other pseudophrynamines were minor or trace alkgdoids in skin extracts oi Pseudophryne species [25]. Structures that were formulated based primarily on MS analysis [25] are shown in Figure 40. Structures for three other pseudophrynamines (272B, 286B, 524) are not proposed (see Appendix). Both enantiomers of pseudophrynaminol (258) have been synthesized [469-471].

Alkaloids from Amphibian Skins

105

The MS fragmentation of pseudophrynamines without substituents in the aromatic ring leads to major ions of m/z 173 and 130 as proposed in Scheme 15. The 2D proton NMR spectrum of 512 along with proton and carbon NMR, MS, IR and UV spectral data for 258, 286A and 512 have been presented [24]. Spectral data for other pseudophrynamines and vapor-phase FTIR spectra of 258 and 286 A have been presented [25]. The properties of the fifteen pseudophrynamines are tabulated in the Appendix. Other analogs of pseudophrynamine A (512) with molecular ions at m/z 526, 540 and 542 were detected by thermospray and direct probe CI-MS [25]. Synthesis. Syntheses of racemic pseudophrynaminol (258) were reported first in 1990 [472,473]. In one synthetic route, pseudophrynaminol was obtained in five steps from tryptamine in a sequence involving an addition-cyclization with 4-bromo-2-methyl-2-butene to yield an 3a-prenyl intermediate, which could be oxidized to an E ally lie alcohol, which on deprotection yielded pseudophrynaminol in an overall yield of 2.3% [472]. In the second synthetic route, the A^-carbomethoxyoxindole-3-ethylamine, obtained by oxidation of A^carbomethoxytryptamine with DMSO, was converted by an addition-cyclization with methyl 4-bromotiglate and then LiAlH4 reduction to racemic pseudophrynaminol [473]. In addition, after conversion to an iV-Boc derivative, a two-step oxidation provided a carboxylic acid corresponding with 286A. Esterification of this acid with A^-Boc-protected pseudophrynaminol, followed by deprotection, yielded pseudophrynamine A (512) identical with natural 512 in spectral properties [473]. A deoxypseudophrynaminol was prepared in onestep addition-cyclization from A^-methyltryptamine with 4-bromo-2-methylbutene [474]. Both enantiomers of pseudophrynaminol have now been synthesized [469-471], allowing confirmation of the proposed absolute configuration of natural levorotatory pseudophrynaminol (258). Synthesis of (-)- and (+)-pseudophrynaminol was first accomplished by protection of the oxindole nitrogen of A^-carbomethoxyoxindole-3-ethylamine through reaction with (-)-l-phenylethylimidazole-l-carboxamide to yield an intermediate urea, which on reaction with methyl 4-bromotiglate gave a mixture of diastereomeric carbomethoxyprenyl intermediates, separable by flash chromatography [469]. Cyclization, reduction of the carbomethoxy group and deprotection with LiAlH4 of each of the diastereomers provided the (+)- and (-)-enantiomers of pseudophrynaminol. An enantioselective synthesis of (+)-pseudophrynaminol involved an addition-cyclization to yield an intermediate allylated cyclic tautomer of L-tryptophan from which the allyl alcohol moiety of pseudophrynaminol was readily elaborated [470]. The absolute configuration of the levorotatory pseudophrynaminol obtained by this route from L-tryptophan corresponds with natural (-)-pseudophrynaminol. Another enantioselective synthesis involved an asymmetric nitroolefination with a chiral nitroenamine of a preny lated oxindole, followed by conversion of the resulting nitroolefin to an aldehyde, reaction with methylamine and reductive cyclization; deprotection yielded (-)-pseudophrynaminol [471]. Racemic pseudophrynaminol has recently been synthesized from TV-methyltryptamine in a one-step addition-cyclization with isoprene oxide in ca. 50% yield [475].

J. W. Daly, H. M. Garraffo and T. F. Spande

106

'^ 3^v^^^CO 2C H3

R,R' 302

R = H, R' = OH

286A R = CO2CH3

316

R = H, R' = 0CH3

256

332

R = OH, R' = 0CH3

(-)-258 * R = CH2OH

R = CHO

346A R = R' = OCH3 HgC^^CHgOH ''3^'*v^^x^^^2^''3

Figure 40. Structures of pseudophrynaminol (258), pseudophrynamine 286A, pseudophrynamine A (512) and other pseudophrynamines. *Absolute configuration as shown. Other pseudophrynamines are shown as having the same configuration.

Alkaloids from Amphibian Stains

107

m/z173

m/z130 Scheme 15 Occurrence. Pseudophrynamines have been detected in Nature only in skin extracts from Australian myobatrachid frogs of the genus Pseudophryne [24,25]. Alkaloids, including pseudophryamines, were not detected in species from six other myobatrachid genera [25]. Skin extracts from frogs of the genus Pseudophryne contained both pseudophrynamines and pumiliotoxins/allopumiliotoxins [25]. The profile and amount of pseudophrynamines differed among species and populations of Pseudophryne. Two species, P. guentheri and P. occidentalis from Western Australia had only small amounts of pseudophrynaminol (258) as the major alkaloid with little or no pumiliotoxins or other pseudophrynamines. In contrast, species and populations of Pseudophryne from eastern Australia usually had larger amounts of alkaloids. Pseudophrynamines predominated in most of the eastern Pseudophryne, but in one population of P. corrohoree and in one population of P. coriacea, pumiliotoxins/allopumiliotoxins predominated [25]. Presumably, both the pseudophrynamines and the pumiliotoxins/allopumiliotoxins found in skin extracts of Pseudophryne frogs are derived from dietary arthropods. In contrast to dendrobatid, mantelline and bufonid anurans with skin alkaloids, the frogs of the genus Pseudophryne are nocturnal and, hence, the dietary arthropod would need to be available at night in the streamside debris in which Pseudophryne frogs forage. Activity. (±)-Pseudophrynaminol has recently been shown to be a potent (IC50 0.3 M.M) noncompetitive blocker for ganglionic and neuromuscular subtype nicotinic receptors [475].

J. W. Daly, H. M. Garraffo and T. F. Spande

108

At 10 \xM pseudophrynaminol had little or no inhibitory effects on radioligand binding to a range of receptors. It had no effect on acetylcholine esterase.

7.2.

Chimonanthine/Calycanthine

The closely related plant alkaloids chimonathine and calycanthine were isolated as minor alkaloids from extracts of 426 skins of the Colombian dendrobatid frog Phyllobates terribilis [32]. The EI-MS and NMR spectral analyses suggested that the alkaloids were chimonanthine and calycanthine and comparison with reported spectral properties confirmed their identities. However, the optical rotations were equal, but opposite in sign to those of the plant alkaloids [32] and h appeared that the dextrorotatory chimonathine and levorotatory calycanthine isolated from frog skin extracts were the enantiomers of the plant alkaloids. Structures are shown in Figure 41.

9H3

HH9"3

r

N

? CH3

(+)-346B

V 1 A HN H HX H-346C

Figure 41. Structures of chimonanthine (346B) and calycanthine (346C). *The absolute configurations shown are based on observed opposite optical rotations to those of the plant alkaloids, whose absolute configurations are knovm. Note that the absolute configuration proposed for frog skin chimonanthine (346B) and that of pseudophrynaminol (258) (Figure 40) are the same. The EI-MS fragmentation of calycanthine is dominated by cleavage of the "dimeric" alkaloid to "monomeric" fragment ions at m/z 173 and 172. In contrast, the base peak in the EI-MS of calycanthine is the parent ion at m/z 346. The proton NMR spectra have been presented [32] and the alkaloids are tabulated as 346B and 346C in the Appendix. It should be noted that another plant alkaloid, morphine, has been reported in trace amounts from skin extracts of the bufonid toad Bufo marinus [476]. Synthesis. Both chimonanthine and calycanthine have been synthesized [see ref. 477]. Oxidative dimerization of A^-methyltryptamine yields in one step (±)-chimonanthine and we^o-chimonanthine. The former can be isomerized with acid to (±)calycanthine [478].

Alkaloids from Amphibian Skins

109

Occurrence, Chimonanthine and calycanthine occur in extracts of plants of the family Calycanthaceae [477]. It is thought that calycanthine may be derived biosynthetically by a rearrangement of chimonanthine. These two alkaloids have been detected in frog skin extracts only as minor or trace alkaloids from two Colombian species of dendrobatid frogs of the genus Phyllohates [32]. A dietary pathway from plant to arthropod to frog is likely. Activity. Calycanthine is highly toxic and can cause cardiac depression, paralysis and convulsions [477], while little appears to be known of the toxicity of chimonanthine. The mechanism and sites of action to our knowledge have not been delineated.

8.

SUMMARY

Amphibian skin has yielded over 500 lipophiUc alkaloids, which can be put in some twenty structural classes. The steroidal samandarines from salamandridid salamanders of the Eurasian genus Salamandra appear to be synthesized by the salamander and stored in parotoid glands. The remainder of the lipophilic alkaloids detected in skin of anurans appear to be of dietary origin. Four lineages of anurans appear to have developed or overexpressed a transport system that accumulates dietary alkaloids unchanged into epithelial secretory (poison) glands. These are dendrobatid frogs (Phyllobates, Dendrobates, Epipedobates, Minyabates) of the neotropics, bufonid toads {Melanophryniscus) of subtropical/temperate southeastern South America, mantelline frogs (Mantelld) of Madagascar and myobatrachid frogs (Pseudophryne) of Australia. All such frogs with the exception of the nocturnal Pseudophryne are diurnal; most are terrestrial. Thus, the putative arthropods that are the dietary source of the alkaloids must be moving during the day in leaflitter for dendrobatid, mantelline and bufonid anurans, but noctumally for myobatrachid frogs in Australia. The arthropod prey must be small, ranging from the size of mites, springtails, and small ants to the size of immature crickets, because of the small size of the anurans. Alkaloids of one class (spiropyrrolizidines) have been identified in millipedes, while alkaloids of another class (coccinellines) are known from beetles. But the prey that at present represents the richest source of frog skin alkaloids is myrmicine ants. Six classes of alkaloids detected in frog skin (decahydroquinolines, 3,5-disubstituted pyrrolizidines, 3,5disubstituted indolizidines, 4,6-disubstituted quinolizidines, 2,5-disubstituted pyrrolidines and 2,6-disubstituted piperidines) occur in myrmicine ants. Only a few hundred of the thousands of myrmicine species have been analyzed for alkaloids. The diversity of profiles of lipophilic alkaloids in different populations and species strongly suggests that the availability of alkaloid-containing arthropods differs markedly from site to site. A major challenge remains with respect to about a dozen structural classes of anuran skin alkaloids for which a putative prey remains a mystery. The steroidal batrachotoxins occur only in dendrobatid frogs of the genus Phyllobates. Only the three species from rain forest of western Colombia contain high levels of batracho-

110

J. W. Daly, H. M. Garraffo and T. F. Spande

toxins in skin and only these true "poison-dart" frogs are used to poison blow darts. Whatever the arthropod source, the batrachotoxins in such arthropods undoubtedly serve as a chemical defense, or as a venom. The Phyllobates frogs are able to eat such toxic prey, since their sodium channels are altered to provide insensitivity to the activating effects of batrachotoxins. One of the batrachotoxins, namely homobatrachotoxin, has now been detected in skin and feathers of Papua New Guinean birds of the genus Pitohui. Whether or not homobatrachotoxin in such birds is of dietary origin is unknown, but it appears likely that it is. Batrachotoxins are potent toxins with selective activating effects on sodium channels. The pumiliotoxins, allopumiliotoxins, homopumiliotoxins and some related subclasses are widely distributed among all the frogs/toads that accumulate alkaloids into skin glands. Therefore, the arthropod prey that contains such alkaloids must be widely distributed in tropical, subtropical and even temperate areas of the world. Pumiliotoxins, allopumiliotoxins and homopumiliotoxins are positive modulators of sodium and perhaps calcium channels and, thereby, exhibit cardiotonic and myotonic activity. The histrionicotoxins occur only in frogs of the neotropics, namely the dendrobatid genera Dendrobates and Epipedobates. Frogs of the genus Dendrobates are ant specialists and histrionicotoxins are often major alkaloids in Dendrobates species. It appears likely that myrmicine ants are the prey item that provides histrionicotoxins, but as yet the histrionicotoxins have not been detected in Nature except in skin extracts from the neotropical dendrobatid frogs. Histrionicotoxins are potent, noncompetitive blockers of nicotinic receptorchannels. 2,5-Disubstituted decahydroquinolines occur in all genera of frogs/toads that have accumulated lipophilic alkaloids in skin with the exception of the nocturnal Australian myobatrachid frogs of the genus Pseudophryne. The dietary prey that provides decahydroquinolines appears to be myrmicine ants from which several decahydroquinoline alkaloids have now been detected. Highly unsaturated nineteen-carbon decahydroquinolines are usually accompanied by highly unsaturated nineteen-carbon histrionicotoxins in frog skin extracts, suggesting a possible common dietary source. The bicyclic izidine alkaloids, consisting of pyrrolizidines, indolizidines, quinolizidines and azabicyclo[5.3.0]decanes, occur in many species of anurans that have accumulated alkaloids into skin. The 3,5-disubstituted pyrrolizidines, 3,5-disubstituted indolizidines, and 4,6-disubstituted quinolizidines, all of which could arise from cyclization of a straight chain precursor, have been detected in myrmicine ants. The 5,8-disubstituted and 5,6,8-trisubstituted indolizidines, the 1,4-disubstituted quinolizidines and the 3,5-disubstituted azabicyclo [5.3.0]decanes have as yet only been detected in skin extracts of anurans. Izidines are commonly found in dendrobatid frogs of the genus Dendrobates. Such Dendrobates species are ant specialists suggesting that ants may be the dietary source of all of the seven classes of izidines, not just the three classes of izidines that have now been detected in myrmicine ants.

Alkaloids from Amphibian Skins

11]

The tricyclic gephyrotoxins occur only in a few populations/species of dendrobatid frogs of the genus Dendrobates arid are always accompanied by histrionicotoxins. Thus, a common dietary prey containing both gephyrotoxins and histrionicotoxins appears likely. The tricyclic cyclopenta[^]quinolizidines have been detected almost uniquely in skin extracts of a tiny Colombian dendrobatid frog of the genus Minyobates. Whatever the dietary prey that provides such alkaloids, it must be very small and may have a very limited range. The tricyclic coccinellines and related compounds occur rather rarely in anuran skin. Coccinellines occur in several species of beetles and thus, small beetles are probably the dietary source of such tricyclic alkaloids detected in skin extracts. The tricyclic spiropyrrolizidines occur rather rarely in dendrobatid frogs, and very rarely in other frogs/toads that have accumulated alkaloids into skin. The major three spiropyrrolizidines of frog skin have oxime or oxime ether moieties, while a less common trace alkaloid, nitropolyzonamine, has a nitro group. Nitropolyzonamine has been detected in a small millipede and it seems likely that millipedes will prove to be the dietary source for the frog skin spiropyrrolizidine oximes. The spiropyrrolizidines are potent noncompetitive blockers of nicotinic receptor-channels with selectivity towards the ganglionic subtype. The monocyclic 2,5-disubstituted pyrrolidines and 2,6-disubstituted piperidines are well-known venom alkaloids in myrmicine ants. However, the transport systems that serve to accumulate alkaloids into anuran skin do not appear to transport effectively such pyrrolidmes and piperidines. Thus, it is perhaps not surprising that such monocyclic alkaloids occur virtually only in trace amounts in skin extracts of anurans. In one extract from a Colombian dendrobatid frog of the genus Dendrobates, a 2,5-disubstituted pyrrolidine was found as a major alkaloid and in one extract from a Panamanian dendrobatid frog of the genus Dendrobates, a 2,6-disubstituted-4-hydroxypiperidine was found as a major alkaloid. Such a 4-hydroxypiperidine has not been detected elsewhere in Nature. Epibatidine has been detected in four species of frogs of the genus Epipedobates from Ecuador and Peru. It is unknown elsewhere in Nature. Based on a structural resemblance to the plant alkaloid nicotine, it is possible that a food-chain from a plant to an arthropod to frogs provides the trace amount of epibatidine found in only a few species/populations of Epipedobates. Epibatidine is a remarkably potent agonist at nicotinic receptors. It is also quite toxic. Another pyridine alkaloid, noranabasamine, and two indole alkaloids, chimonanthine/ calycanthine may also be obtained via a food chain originating from a plant source. Such alkaloids were detected in minor amounts only in Colombian species of dendrobatid frogs of the genus Phyllobates. The pseudophrynamines occur only in skin extracts from Australian myobatrachid frogs of the genus Pseudophryne. Such frogs are noctumal and hence a dietary prey item would need to be active at night. As yet pseudophrynamines have not been detected in Nature except in Pseudophryne frogs. The pseudophrynamines are reminiscent in structure to physostigmine. One pseudophrynamine was a potent noncompetitive blocker of nicotinic receptor-channels.

112

J. W. Daly, H. M. Garraffo and T. F. Spande

The diversity of alkaloids detected in anuran skin is truly amazing and suggests that the dietary prey may provide a treasure-trove of further alkaloids. Over five hundred alkaloids have now been detected in skin extracts of anurans, the majority of which can be placed in twenty-some structural classes. Only a few of these alkaloids from amphibian skin have been detected elsewhere in Nature. Two of the batrachotoxins have been reported from the Pitohui birds (Section 2.2). At present six of the decahydroquinolines have been detected from myrmicine ants (Section 3.4). Two of the pyrrolizidines, four of the indolizidines, and one of the quinolizidines also occur in myrmicine ants (Sections 3.5, 3.6.1, 3.7.1). Two of the tricyclic coccinelline-like alkaloids occur in coccinellid beetles (Section 4.3). A spiropyrroline and one of the spiropyrrolizidines occur in a millipede (Section 4.4). Three alkaloids occur also in plants (Section 7.2). Thus, of the over five hundred alkaloids of anuran skin extracts, less than twenty have been detected in arthropods, which could serve as dietary prey. Over half of the different structural classes have not been detected elsewhere in Nature. At present there is no evidence that dietary precursors of alkaloids are involved, particularly since all alkaloids fed to frogs accumulate unchanged into secretory skin glands. Many of the minor or trace alkaloids reported in the present review require further study to define some tentative structural assignments and in many cases the relative stereochemistry at certain carbons. In some cases larger quantities and isolation of pure compounds will be required in order to conduct NMR spectral analysis. This is especially true of many alkaloids that cannot be assigned, even tentatively, to one of the present twenty structural classes of frog skin alkaloids. Such unclassified alkaloids are included in the complete tabulation of lipophilic anuran skin alkaloids presented in the Appendix. There are now severe restrictions on collection of adequate numbers of amphibians, particularly the brightly colored dendrobatid frogs, all of which were arbitrarily placed by an international convention (CITES) in a "threatened" status in spite of evidence to the contrary [see ref 479]. Thus, the challenges of structural definition for alkaloids with tentatively assigned structures and for those which are unclassified will rely on new and/or more sensitive spectral and chemical approaches and on discovery of the groups of alkaloid-containing arthropods, that provide batrachotoxins, pumiliotoxins, certain izidines, epibatidine, pseudophrynamines and the over one hundred unclassified alkaloids detected, as yet, only in amphibian skin extracts.

Acknowledgement. Our research on the chemistry, biology and pharmacology of alkaloids of anuran skin over the past thirty some years has progressed through the contributions of many students, postdoctorates, colleagues and collaborators. During almost the entire three decades, two individuals have contributed so much. They are Dr. Charles W. Myers of the American Museum of Natural History in New York and Dr. Takashi Tokuyama of the Department of Chemistry, Osaka City University. Dr. Edson X. Albuquerque pioneered the pharmacology of anuran skin alkaloids. In recent years, Drs. Fabian Gusovsky and Barbara Badio have made major contributions to the pharmacology and Drs. Tappey H. Jones,

Alkaloids from Amphibian Skins

113

Michael W. Edwards and Poonam Jain, and N. Rabe Andriamaharavo to the chemistry. To all we owe a debt of gratitude.

APPENDIX The distribution of twenty classes of lipophilic alkaloids in skin extracts of amphibians is presented in Table 1. All of the lipophilic alkaloids detected in frog/toad skin extracts are listed below with the exception of the samandarines and batrachotoxins using a code designation based on the molecular weight and an identifying letter(s) in bold face. The tabulation updates and revises prior tabulations [1,10,11]. The Rf values on TLC and emergent temperatures on GC on 1.5% OV-1 columns of prior tabulations have been omitted. The protocol for preparation of an alkaloid fraction from methanol extracts of amphibian skin has been kept relatively constant as have the conditions for a flame-ionization GC profile of alkaloids present in the equivalent of 2 mg (wet weight) of skin [see Appendix in ref. 11 for details of preparation and GC analysis of alkaloid fractions from amphibian skin]. Flame-ionization GC profiles on 1.5% OV-1 columns have been presented for many species and populations as follows: Dendrobatids - Phyllohates [11]; Dendrobates [1,6,7,10,11,49, 60,123,162,209,480,481]; Epipedobates [11,49,482]; Minyobates [11,49,305,483]; Bufonids - Melanophryniscus [59]; Mantellines - Mantella [8,56,59,61]; Myobatrachids - Pseudophryne [25,59]. Distribution of alkaloids in anuran skins has been tabulated in 1986 [10], and in 1987 [1]. Since the 1987 review, the occurrence of alkaloids in skin extracts have been tabulated for additional populations, species or groups of species of dendrobatid [5,6,60,481], bufonid [86], mantelline [56,61] and myobatrachid [25] anurans. The tabulation of alkaloids contains the following: 1) The code designation. A number of alkaloids occur in frog skin extracts as more than one diastereomer or are isomeric with respect to position of double bonds, etc. In most cases, such isomeric alkaloids are tabulated under a generic code designation, since such isomers were usually not differentiated in early studies on skin extracts. The isomers or diastereomers are then listed and properties noted under the tabulated generic code designation. Examples are the following: Pumiliotoxm 307F\ F" and F"*; Erythro- and threO'323A; Decahydroquinolines cis- and trans-195A; Indolizidines 195B and 223AB, where four diastereomers have been detected; Indolizidine 235B, where double bond isomers 235B' and B" exist; Indolizidines 207A, 207A* and 207A". In some cases, it is possible that more than one alkaloid may have been listed under the same code designation, based solely on early GC-MS data. Examples are indolizidine 223A and quinolizidine 195C. 2) The class of alkaloid. The abbreviations are as follows: Pyr, a 2,5-disubstituted pyrrolidine; Pip, a 2,6-disubstituted piperidine, PTX/aPTX/hPTX, a pumiliotoxin, an allopumiliotoxin, a homopumiliotoxin; DeoxyPTX, DesmethylPTX, DihydroPTX,

114

TABLE 1.

J. W. Daly, H. M. Garraffo and T. F. Spande

Occurrence of Various Classes of Lipophilic Alkaloids in Amphibian Skins • - D e n d r o b a t i d a e - •;•••-,"•., '':•;;•'-;'-:HVV;V^

Phyttobates Monocyclics-;^; Pvr Pip BicycOcs PTX/aPTX hPTX HTX DHO 3,5-P 3,5-1 5,8-1 5,6.8-1 1,4-0 4,6-0 AzabicvcloD Tricyclics GTX CPO Coccin. class SpiroP Epibatidine Pyridines Pseudo Indoles S t e r o i d a l .^V;:;:^^^:;:^:'; BTX

X X X X

X X

Dendrobates Epipedpbates X X X X X X X X X X X X X X X X

Minyobates 1

X X

X

X X

X

X X X

X X X

X X X X

X X X

X

Abbreviations. Pyr = pyrrolidines; Pip = piperidines; PTX/aPTX = pumiliotoxins/allopumiliotoxins; hPTX = homopumiliotoxins; HTX = histrionicotoxins; DHQ = decahydroquinolines; P = pyrrolizidines; I = indolizidines; Q = quinolizidines; AzabicycloD = azabicyclo [5,3,0]decanes; GTX = gephyrotoxins; CPQ = cyclopentaquinolizidines; Coccin = coccinellines; SpiroP = spiropyrrolizidines; Pseudo = pseudophrynamines; BTX = batrachotoxins. Anurans from the above 7 genera of the families Dendrobatidae, Rhacophoridae (subfamily Mantellinae), Bufonidae and Myobatrachidae, contain lipophilic alkaloids as shown in the Table. Anurans from 52 genera of 8 other families did not contain lipophilic alkaloids in skin extracts [59,86 and unpublished resuhs]. Anurans from the remaining 150 genera of these 8 families have not been examined for alkaloids. The 11 remaining families of anurans v^ith 29 genera have not been examined for alkaloids. Family Dendrobatidae: Frogs of the dendrobatid genera Colostethus d^nd Aromahates did not contain lipophilic alkaloids, nor did 2 species of Epipedobates.

Alkaloids from Amphibian Skins

TABLE 1.

115

Continued

~!!iiP~i{iii;i}

i

i;!!!iIII.......... i ...... ]!i i, !!: !n

!ili ,[,i .........i

~i{.......!.l::.........i! ::Ants :~ !

• ~l~l~Ii:i{~i~iiiiiiiiiiiii!ii i:~!:.:;!ii:~!!:.i;:;i:,!~' ii!il

I ~

i

i

X X

X X X X X X X

X X X X

isa

iii',ii i !ilil

~

i

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

S

iiii!':!i~.i:.:iii :t

ii!i :::::::~::!~i;!; !:i;i; :~i~!iiii:~ :.:#ili;~il iii!ii:.i ~iiiii::!i!i!~ili ~iil;.i!! I

Ants Ants Ants

X Ants

........................................ ;iiii~:ii?i'~iiii!}i :.;i~' i:)!:i!: .:!~!i::

~

X

i!ii!i!!!ii!!!i!i~ii!!:i~iii~i~i!iiii!ii!ii;

~ifli3,5~ti~:ii:i~iiiiiiiii~:;i! ,~ill

i

'Ants X X

,,

X X

X X

X

Beetles Millipedes

X Bird

Family Rhacophoridae, Subfamily Mantellinae: Frogs of the mantelline genus Mantidactylus did not contain lipophilic alkaloids. The remaining mantelline genus Laurentomantis has not been examined. Three other genera of rhacophorid frogs did not contain lipophilic alkaloids, while 7 have not been examined for alkaloids. Family Bufonidae: Toads of the bufonid genera Bufo, Atelopus and Dendrophryniscus did not contain lipophilic alkaloids. The remaining 18 bufonid genera have not been examined for alkaloids. Family Myobatrachidae: Frogs of the myobatrachid genera Adelotus, Cyclorana, Heleioperus, Notaden and Uperoleia did not contain lipophilic alkaloids. The remaining 6 myobatrachid genera have not been examined for alkaloids.

116

J. W. Daly, H. M. Garraffo and T. F. Spande

DeoxyhPTX, DesmethylhPTX, DehydroPTX; HTX, a histrionicotoxin; DHQ, a decahydroquinoline; OHQ, an octahydroquinoline; THQ a tetrahydroquinoline; DHQ-dimer, alkaloids 382,384A/384B; 3,5-P, a 3,5-disubstituted pyrrolizidine; 3,5-1, a 3,5-disubstituted indolizidine; 5,8-1, a 5,8-disubstituted indolizidine; 5,6,8-1, a 5,6,8-trisubstituted indolizidine; 4,6-Q, a 4,6-disubstituted quinolizidine; 1,4-Q, a 1,4-disubstituted quinolizidine; Azabicyclodecane; GTX, a gephyrotoxin; CPQ, a cyclopenta[^]quinolizidine; Tricyclic, a tricyclic coccinelline-like alkaloid; SpiroP, a spiropyrrolizidine; Epib, an epibatidine analog; Pseudo, a pseudophrynamine. Some alkaloids are termed "Izidine" and those of as yet undefined structures are designated as "Unclass", unclassified. 3) An empirical formula based on high resolution MS. Tentative formulae are indicated by single quotation marks. 4) Diagnostic EI-MS (EI) or ion-trap pseudo-EI-MS (IT-EI) ions with intensities, relative to a base peak set equal to 100, in parentheses. Intensities in IT-EI can vary considerably with conditions. 1) Vapor-phase FTIR references to spectra and/or data. Spectra presented in this chapter are designated by Figure number. Significant absorption peaks are reported including Bohlmann bands, OH, C=0, cis CH=CH, conjugated (conj.) CH=CH, C=CH2, C=C=C, C=CH, etc. 6) NMR references to spectra and/or data. 7) The number of hydrogens exchangeable with ND3 (OD, ID, 2D, etc). The number of hydrogens in a perhydro-derivative (HQ, H2, H4, etc). In earlier reviews [1,10,11] MS data for some perhydro-derivatives were reported. 8) Other data or comments, including occurrence of diastereomers or isomers. 9) Occurrence in dendrobatids, bufonids, mantellines and myobatrachids. 10) Citation of Figure number for the structure or tentative structure. Tabulation of Alkaloids 151A. 151B.

153A. 153B. 155.

161. 167A.

Unclass. 'CioHnN'. EI: 151(100), 150(25). OD. Dendrobatid. Polyzonimine. CioHnN. EI: 151(4), 150(3), 136(21), 108(20), 96(57), 82(100), 81(60). FTIR spectrum (Figure 34): C=N 1623 cm'^ NMR data [329]. OD. Dendrobatid. Figure 33. Unclass. 'C,oH,9N'. EI: 153(100), 152(60). ID, HQ. Previously postulated to be DHQ [11]. Dendrobatid. Unclass. 'CjoHigN'. EI: 153(45), 152(100). OD, HQ. Dendrobatid. Unclass. CpHpNO. EI: 155(40), 140(11), 126(100), 114(78), 113(88), 98(26), 84(16), 70(54). FTIR: Weak Bohlmann band 2808 cm"^; OH 3614 cm'^ ID. Dendrobatid. Unclass. C9H11N3. EI: 161(76), 160(100), 133(10, C8H9N2), 119(8, C7H7N2), 107(22). FTIR data [56]: Aromatic 3050,1594 cm"^ OD. Mantelline. Unclass. C11H21N. EI: 167(1), 138(100). OD,Ho. Previously postulated to be a 5,8-1 [11]. Dendrobatid.

Alkaloids from Amphibian Skins 167B.

117

167E.

5-1. C11H21N. Synthetic 5-propyl I [268]. EI: 167(1), 124(100), 96(23). FTIR: Synthetic 5,9Z: strong Bohlmann band 2788 cm'^; synthetic 5,9£: weak Bohlmann band 2811 cm'^ A postulated 5-propyl I (167B) [10] has not been confirmed in skin extracts. Unclass. C11H21N. EI: 167(100), 166(55). OD, HQ. Dendrobatid. Unclass. 'CnH2iN'. EI: 167(100), 166(53). lD,Ho. Previously postulated to be a DHQ[11]. Dendrobatid. 3,5-1. 'CnH2iN'. EI: 167(<1), 138(100). FTIR: Moderate Bohlmann band 2788

167F.

cm'^ Dendrobatid. Figure 19. 3,5-P. •CnH2iN'. EI: 167(8), 166(5), 124(100). OD, HQ. Tentatively, a 5-methyl-

167C. 167D.

179. 181A. 181B. 181C. 181D. 181E. 183A. 183B. 185. 189. 191A.

191B. 193A. 193B. 193C.

193D.

3-propyl P. Previously tabulated as a 5-propyl 1167B [10]. Dendrobatid. Figure 17. Unclass. 'C12H21N'. IT-EI: 180(17), 136(100), 134(30), 70(35). Dendrobatid. Izidine. 'C12H23N'. EI: 181(2), 152(100). Previously postulated to be a 1,4-Q [11]. Dendrobatid. 5,8-1. C12H23N. EI: 181(2), 138(100), 96(10). OD, HQ. Dendrobatid. Figure 21. Unclass. 'C12H23N'. EI: 181(100), 180(68), 166(10). FTIR: Strong, sharp Bohlmann band 2786 cm"^ OD, HQ. Dendrobatid. Unclass. 'C12H23N'. EI: 181(3), 152(100). ID. Previously postulated to be a DHQ[11]. Dendrobatid, mantelline. Unclass. 'C12H23N'. EI: 181(100), 180(46). lD,Ho. Previously postulated to be a DHQ[11]. Dendrobatid. Unclass. C12H25N. EI: 183(3), 154(100). lD,Ho. Previously postulated to be a 2ethylpiperidine with a pentyl at C-3, C-4 or C-5 [11]. Dendrobatid. Pyr. 'Ci2H25N'. EI: 183(10), 126(100). lD,Ho. Dendrobatid. Figure 35. Unclass. 'CiiH23NO'. EI: 185(1), 170(100). Dendrobatid. THQ. 'CnHipN'. IT-EI: 190(10), 174(26), 161(100), 146(94), 91(10). FTIR data [56]: Aromatic 3050, 1590 cm'^ Mantelline. Figure 15. Tricyclic. 'C13H21N'. IT-EI: 192(43), 191(28), 163(58), 152(20), 148(30), 134(33), 120(100), 106(33), 95(50). Tentatively a dehydro 193C [56]. Mantelline. Propyleine. 'Ci3H2iN'. EI: 191(100), 190(82), 176(43), 148(26), 134(23), 122(20). Dendrobatid. Figure 32. Unclass. C13H23N. EI: 193(100), 191(65). OD, HQ. Dendrobatid. Unclass. C13H23N. EI: 193(8), 192(7), 178(9), 150(100), 122(8), 70(7). OD, HQ. Dendrobatid. Precoccinelline. C13H23N. EI: 193(45), 192(95), 178(35), 164(55), 151(80), 150(100), 137(40), 136(50), 122(40), 108(30), 96(30), 94(30). FTIR spectrum [86]: Weak Bohlmann band. Dendrobatid, bufonid. Figure 32. OHQ. C13H23N. EI: 193(9), 178(6), 150(100), 122(6), 110(5), 96(12). FTIR data [56]: Weak Bohlmann band 2807 cm"^; enamine 3020, 1641 cm"^ ID. Mantelline. Figure 15.

118

193E. 193F.

193G. 193H. 195A.

195B.

195C.

195D. 195E. 195F. 195G. 195H. 1951. 195J. 197A. 197B. 197C.

J. W. Daly, H. M. Garraffo and T. F. Spande

5,8-1. C13H23N. EI: 193(<1), 192(3), 152(100), 96(22). FTIR: Strong, sharp Bohlmann band 2788 cm-^ C=CH2 3083 cm-^ OD. Mantelline. Figure 21. DeoxyhPTX. C13H23N. EI: 193(20), 192(23), 178(27), 164(100), 151(19), 150(31), 136(27), 122(37), 84(26). FTIR: Moderate, broad Bohlmann band 2745 cm'^ Dendrobatid. Figure 9. Izidine. 'CnHssN'. IT-EI: 194(1), 152(100), 110(31). Either a 5,6,8-1 or a 1,4-Q. Dendrobatid. DeoxyPTX. 'C13H23N'. EI: 193(14), 192(20), 178(17), 150(100), 136(21), 108(27), 70(38). Dendrobatid. Figure 5. DHQ. C13H25N. CW-195A: EI: 195(3), 194(5), 152(100), 109(8). FTIR Spectrum [160]: Weak Bohlmann band 2804 cm"^ NMR spectrum [190] and data [105,124,161,162]. ID, Ho. 7V-Acetyl derivative. rra«5-195A: IT-EI: 196(1), 152(100), 109(11). FTIR spectrum [160]: No Bohlmann band. Dendrobatid, mantelline. Figure 13. 3,5-1. C13H25N. EI: 195(2), 194(2), 180(6), 138(100). FTIR data [86]: Bohlmann bands: 5Z,9Z: moderate 2788 cm"^; 5E,9E: moderate 2793 cm"^; 5£,9Z: weak 2800 cm'^; 5Z,9£^: none. NMR data [162]. OD, HQ. Dendrobatid (all diastereomers), bufonid (all diastereomers). Figure 19. 4,6-Q. C13H25N. EI: 195(6), 180(13), 152(100). FTIR spectrum and data [125] (Figure 26): Weak Bohlmann band 2813 cm-^ NMR data [125]. OD,Ho. Dendrobatid, mantelline. Figure 25. Izidine. 'C13H25N'. EI: 195(2), 166(100), 110(12). OD, HQ. Either a 5,6,8-1 or a 1,4-Q. Dendrobatid. Unclass. 'C13H25N'. EI: 195(45), 194(100). OD, HQ. Dendrobatid. 3,5-P. C13H25N. EI: 195(<1), 124(100). FTIR: Weak Bohlmann band 2805 cm'^ Dendrobatid. Figure 17. 5,6,8-1. 'C13H25N'. EI: 195(<1), 194(<1), 152(100), 110(13), 70(11). FTIR: Strong, sharp Bohlmann band 2787 cm"^ Dendrobatid, mantelline. Figure 23. 5-1. 'C,3H25N'. EI: 195(1), 194(3), 124(100), 96(9). OD. Tentatively a 5-pentyl I. Mantelline. 5,8-1. 'Ci3H25N'. EI: 195(<1), 138(100), 96(23). OD. Tentatively a 5-butyl-8methyll. Mantelline. DHQ. 'Ci3H25N\ IT-EI: 196(5), 180(32), 152(100), 110(10). Cz5-195J: FTIR data [125]: Weak Bohlmann band 2804 cm'^ ID. Mantelline. Figure 13. Unclass. 'C12H23NO'. EI: 197(1), 180(100), 126(35). lD,Ho. Dendrobatid. Pyr. C,3H27N. EI: 197(1), 196(2), 140(78), 126(100). FTIR (Figure 36): No Bohlmann band. ID, HQ. Dendrobatid. Figure 35. 5,8-1. C12H23NO. EI: 197(1), 196(1), 154(100, C9H16NO), 96(38). FTIR: Strong, sharp Bohlmann band 2789 cm"^ OH 3485 cm'^ ID. Tentatively an 8hydroxymethyl-5-propyl I. Dendrobatid, mantelline.

Alkaloids from Amphibian Skins

197D.

197E. 201A. 201B.

203A.

203B. 205A.

205B.

205C. 205D. 205E.

207A.

207B. 207C. 207D.

119

Unclass. 'C13H27N'. IT-EI: 198(50), 197(35), 196(23), 180(47), 125(52), 108(39), 71(72), 58(100). Other alkaloids with a base peak at m/z 58 are 211G, 241B, 253E, 2671, 2670,269E. Dendrobatid. Pip. 'C13H27N'. IT-EI: 198(3), 197(1), 182(8), 98(100). ID. Dendrobatid. Figure 37. Unclass. C14H19N. EI: 201(<1), 200(2), 136(100), 134(41), 120(15). OD. See 245F. Dendrobatid. Tricyclic 'Ci4H,9N'. Three isomers. IT-EI: 201B': 202(30), 201(43), 186(100), 158(41), 144(21), 130(26); 201B": 202(27), 201(40), 186(100), 173(12), 158(40), 144(19), 130(25); 201B'": 202(41), 201(100), 186(32), 172(13), 158(21), 144(56), 130(16). Dendrobatid. 5,8-1. C14H21N. EI: 203(1), 202(2), 138(100), 96(13). FTIR data [56] and spectrum [161]: Strong, sharp Bohlmann 2789 cm"^; conj. CH=CH 3038 cm"^; C=CH3327cm-'. NMR data [161]. OD, Hg. A minor isomer, 203A\ has been detected and it is probably the 5,9E diastereomer since there is no Bohlmann band. Dendrobatid, mantelline. Figure 21. Unclass. 'C14H21N'. EI: 203(<1), 166(100). OD. The parent ion may have been misidentified, since loss of 37 is improbable. Dendrobatid. 5,8-1. C14H23N. EI: 205(1), 204(2), 138(100), 96(15). FTIR spectrum (Figure 22): Strong, sharp Bohlmann 2787 cm'^;C=CH 3327 cm"^ NMR spectrum and data [105,161]. OD, H4. Minor isomers, 205A* and 205A", have been detected. Both have strong, sharp Bohlmann bands at 2787 cm"^ and CsCH 3327 cm"^ Dendrobatid, mantelline. Figure 21. Tricyclic. C14H23N. EI: 205(31), 204(46), 190(100), 162(17), 148(10), 134(16), 96(15), 70(8). FTIR spectrum [308]: Weak Bohlmann band 2796 cm-^ NMR spectrum [105] and reinterpretation [308]. OD, H2. Dendrobatid. Figure 32. Unclass. 'C14H23N. IT-EI: 206(5), 190(5), 176(23), 140(84), 126(100). Tentatively an A^-methyl Pip. Mantelline. Unclass. 'C14H23N'. EI: 205(35), 190(33), 176(13), 162(11), 150(64), 148(73), 134(33), 82(100). Dendrobatid. Tricyclic. C13H19NO. IT-EI: 206(28), 204(13), 190(24), 177(12), 163(100), 162(52), 134(30), 122(15), 108(11), 70(29). FTIR: Moderate Bohlmann bands 2794,2739 cm"^; C=0 1728 cm"^ Dendrobatid. 5,8-1. C14H25N. EI: 207(1), 206(2), 138(100), 96(13), 70(9). FTIR: 207A: Strong, sharp Bohlmann band 2787 cm'h C=CH2 3084 cm•^ NMR data [60]. OD, H2. Two minor isomers, 207A' and 207A", have been detected. Both appear to have an internal double bond. 207A": FTIR spectrum [86]. Dendrobatid. Figure 21. PTX. C13H21NO. EI: 207(10), 190(15), 166(100), 70(80). ID, H2. A structure fitting the data is not apparent. Dendrobatid. Izidine. 'C14H25N'. EI: 207(16), 152(100), 110(13). OD, H2. Either a 5,6,8-1 or a 1,4-Q. Dendrobatid. Unclass. 'C14H25N'. EI: 207(58), 180(100). 0D,H2. Dendrobatid.

120

207E.

J. W. Daly, H. M. Garraffo and T. F. Spande

Unclass. 'C14H25N'. EI: 207(10), 206(12), 178(30), 164(100), 162(29), 120(22). FTIR: Moderate, sharp Bohlmaim band 2787 cm•^ OD. See245F. Dendrobatid, mantelline. 207F. Unclass. 'C,4H25N'. EI: 207(9), 192(100), 126(41), 110(54), 94(44). OD, H2. Dendrobatid. 207G. Unclass. C14H25N. EI: 207(38), 206(100), 192(16), 180(29), 136(18), 97(40), 96(56), 84(87). FTIR: Strong, sharp Bohlmann band 2788 cm'^ OD. Probably an isomer of 207H with which it always occurs. Previously postulated to be a hPTX [56]. Dendrobatid, mantelline. 207H. Unclass. C14H25N. EI: 207(46), 206(100), 192(16), 179(13), 178(17), 136(20), 97(37), 96(49), 84(81). FTIR: Strong, sharp Bohlmann band 2788 cm^'. OD. Probably an isomer of 207G with which it always occurs. Previously postulated to be a hPTX [11], Dendrobatid, mantelline. 2071. 1,4-Q. C14H25N. EI:207(<1),206(2), 166(100), 110(11). FTIR spectrum and data [56,281]: Moderate, broad Bohlmann band 2789 cm^^; C=CH2 3084 cm^^ Mantelline. Figure 27. 207J. Tricyclic. C14H25N. EI: 207(21), 206(11), 192(15), 178(23), 164(19), 152(8), 150(11), 136(100), 110(11), 108(14). FTIR data [56]: Strong Bohlmann band 2789 cm'^ Dendrobatid, mantelline. 207K. Unclass. C14H25N. An isomer of 207H. EI: Nearly the same. Dendrobatid. 207L. Unclass. 'C14H25N'. EI: 207(9), 206(6), 178(15), 164(14), 123(85), 109(23), 81(45), 70(100). Dendrobatid. 207M. Unclass. C14H25N. EI: 207(2), 206(1), 164(100), 140(82). FTIR: Moderate Bohlmann band 2805 cm"^; CsCH 3328 cm"^ Possibly an A^-methyl Pip. Mantelline. 207N. Unclass. C,4H25N. EI: 207(13), 206(5), 192(7), 178(6), 152(100), 150(27), 136(22), 122(11), 70(11). Dendrobatid. 207O. DeoxyhPTX. C,4H25N. EI: 207(17), 206(19), 192(31), 164(100), 150(21), 136(11), 122(23), 109(13), 84(58). FTIR: Moderate, broad Bohlmann bands 2799, 2657 cm"^ Dendrobatid, mantelline. Figure 9. 207P. Tricyclic. C14H25N. EI: 207(48), 206(19), 192(100), 164(35), 152(68), 136(20), 126(52), 110(43), 94(37), 83(60), 82(83). Dendrobatid. 207Q. 5,8-1. 'C14H25N'. IT-EI:208(<1), 166(100), 96(35). Tentatively a 5-allyl-8-propyl I. Dendrobatid. 207R. Tricyclic. 'C14H25N'. EI: 207(4), 206(3), 192(47), 178(9), 164(18), 150(38), 136(100), 70(70). FTIR: Moderate, broad Bohlmann band 2807 cm"^ OD. A lower homolog of 221M. Dendrobatid. 208/210. Epibatidine. CnHi3N2Cl. EI: 210(4), 208(12), 181(1), 179(3), 142(3), 140(9), 69(100), 68(30). FTIR spectrum [354]: No Bohlmann band; pyridine 1460, 1110 cm"^ NMRofA^-acetyI derivative [354]. ID. Dendrobatid. Figure 38. 209A. DHQ. 'C13H23NO'. EI: 209(5), 168(100). H2. Tentatively a ring-hydroxylated 2allyl-5-methyl DHQ. Dendrobatid.

Alkaloids from Amphibian Skins

209B. 209C. 209D.

209E. 209F.

209G.

209H. 2091. 209J. 209K. 211A. 211B. 211c. 211D. 211E.

211F. 211G.

211H.

121

5,8-1. C14H27N. EI: 209(5), 138(100), 96(12). FTIR: Strong, sharp Bohlmann band 2786 cm'^ OD,Ho. Dendrobatid. Figure 21. Izidine. C,4H27N. EI: 209(11), 152(100), 110(5). OD, HQ. Either a 5,6,8-1 or a 1,4-Q. Dendrobatid. 5-1. C14H27N. Synthetic 5-hexyl I [268]: EI: 209(1), 208(2), 124(100), 96(10). FTIR spectra [161]: Synthetic 5,9Z: moderate, sharp Bohlmann band 2788 cm'^; synthetic 5,9E\ weak Bohhnann band 2813 cm'^ A postulated 5-hexyl I (209D) [10] has not been confirmed in skin extracts. Izidine. 'C14H27N'. EI: 209(17), 166(100), 110(8). OD, H2. Either a 5,6,8-1 or a 1,4-Q. Dendrobatid. PTX. C13H23NO. EI: 209(6), 208(4), 166(60), 112(15), 84(24), 70(100). FTIR spectrum [57]: Moderate, sharp Bohlmann bands, 2798,2752 cm"^; OH 3542 cm"^ NMRdata[57]. ID. Dendrobatid. Figure 3. Unclass. C14H27N. EI: 209(13), 166(35), 138(100), 125(51), 110(43), 82(42). FTIR: Moderate Bohlmann band 2759 cm"^ OD. Two isomers detected. Dendrobatid. DesmethylhPTX. 'C,3H23NO'. EI: 209(25), 208(21), 194(15), 166(100), 122(17), 84(47). Mantelline. Figure 9. 5,8-1. C14H27N. EI: 209(<1), 208(<1), 166(100), 138(11), 96(17), 70(11). FTIR: Strong, sharp Bohlmann band 2786 cm"^ Dendrobatid. Figure 21. DHQ. 'C,4H27N'. EI: 205(2), 204(3), 166(100). ID. Tentatively a 5-ethy 1-2propyl DHQ, but DHQs usually have 13,15,17 or 19 carbons. Dendrobatid. 3,5-F. 'C,4H27N'. EI: 209(2), 124(100). OD,Ho. Tentatively a 3-hexyl-5-methyl P. Previously tabulated as a 5-hexyl 1209D [10]. Dendrobatid. Figure 17. DHQ. C13H25NO. EI: 211(3), 210(2), 168(100), 152(32), 150(13). NMR spectrum and data [105]. 2D, HQ. Dendrobatid. Figure 13. Unclass. •C13H25NO'. EI: 211(4), 160(100). ID. Dendrobatid. Unclass. 'C13H25NO'. EI: 211(2), 210(1), 168(100), 124(12), 110(21), 70(9). FTIR data [56]: No Bohlmann band. ID. Dendrobatid, mantelline. Unclass. •Ci4H29N'. EI: 211(2), 210(3), 168(85), 126(100). FTIR data [56]: Very weak Bohlmann band 2799 cm"^ Two isomers. Mantelline. 3,5-1. C13H25NO. EI: 211(<1), 210(1), 196(5), 138(100). FTIR: Moderate Bohlmann band 2791 cm"^; OH 3653 cm"^ ID. Tentatively a 5£,9£-3(hydroxybutyl)-5-methyl I. Dendrobatid, mantelline. Unclass. C13H25NO. EI: 211(<1), 210(<1), 140(100, CgHnNO). FTIR: Moderate Bohlmann band 2801 cm"^; OH 3556 cm'^ ID. Dendrobatid. Unclass. 'C13H25NO'. EI: 211(20), 210(18), 194(15), 178(27), 58(100). Other alkaloids with a base peak at m/z 58 are 197D, 241B, 253E, 2671,2670,269E. Dendrobatid. Unclass. C13H25NO. EI: 211(<1), 210(<1), 196(5), 154(100), 124(27). Dendrobatid.

122

2111. 211J. 211K. 213. 217A.

217B.

217C. 219A.

219B. 219C. 219D. 219E. 219F. 219G. 219H. 2191.

219J. 219K.

J. W. Daly, H. M. Garraffo and T. F. Spande

Pip. 'CMHSQN'. E I : 2 1 1 ( 2 ) , 2 1 0 ( 1 ) , 154(81), 140(100). ID. Dendrobatid. Figure 37. Pip. C14H29N. EI:211(<1), 196(6), 112(100). OD. Tentatively an A/-methyl Pip. Dendrobatid. DHQ. 'Ci3H25NO'. EI: 211(9), 210(4), 196(23), 168(100), 150(10). 2D. Tentatively a ring-hydroxylated 5-methyl-2-propyl DHQ. Dendrobatid. Pip. 'Ci3H27NO'. EI: 213(<1), 198(3), 114(100), 96(11). FTIR: No Bohlmann band; OH 3658 cm"^ 2D. Dendrobatid. Figure 37. 1,4.Q. C15H23N. EI: 217(<1), 216(<1), 152(100), 110(14). FTIR spectrum [56] and data [296]: Moderate, broad Bohlmann band 2788 cm'^ NMR data [296]. OD, Hg. Trace amounts of another isomer were detected [56]. Mantelline. Figure 27. 5,8-1. C15H23N. EI: 217(<1), 216(<1), 152(100), 96(13). FTIR spectrum [56]: Strong, sharp Bohlmann band 2789 cm"^ OD. Dendrobatid, mantelline. Figure 21. Unclass. 'C15H23N'. EI: 217(1), 216(3), 202(6), 178(28), 160(8), 150(7), 138(100), 110(23), 91(13), 70(15). OD. Dendrobatid, mantelline. DHQ. C15H25N. EI: 219(1), 218(1), 178(100). FTIR spectra [161] (Figure 14): a5-219A: No Bohlmann band. TranS'219A: Weak Bohlmann band 2800 cm"^ 5Epi-trans'219A: Weak Bohlmann band 2800 cm"^ NMR spectrum and data [161,162]. ID. A^-Acetyl derivative. Dendrobatid. Figure 13. Izidine. 'C15H25N'. EI: 219(1), 218(2), 152(100). OD, H4. Previously postulated to be a 1,4-Q [11]. Dendrobatid. DHQ. C15H25N. EI: 219(<1), 152(100). 1D,H4. Tentatively a 5-methy 1-2pentadienyl DHQ. Dendrobatid. DHQ. C15H25N. EI: 219(3), 180(100). 1D,H4. Tentatively a 2-propargy 1-5propyl DHQ. Dendrobatid. Izidine. 'Ci5H25N'. EI: 219(1), 218(3), 166(100). OD. Previously postulated to be a 1,4-Q [11]. Dendrobatid. 5,8-1. C15H25N. EI: 219(<1), 152(100), 96(12). FTIR: Strong, sharp Bohlmann band 2789 cm'^ CHCH 3329 cm"^ Dendrobatid, mantelline. Figure 21. Unclass. 'C15H25N'. IT-EI: 220(20), 164(100), 162(46), 120(30). FTIR data [86]: Strong Bohlmann band 2787 cm'^ 0D,H4. See245F. Bufonid. Unclass. 'CisHssN'. EI: 219(8), 204(5), 150(100), 135(33), 134(30), 121(59), 107(21). OD. Dendrobatid. Tricyclic. C15H25N. EI: 219(30), 218(9), 204(25), 190(18), 176(36), 164(18), 162(22), 148(100), 108(28), 93(54), 70(71). FTIR: Strong, sharp Bohlmann band 2792 cm"^;d5CH=CH 3020 cm'^ Two isomers. Dendrobatid. 5,8-1. C15H25N. EI: 219(1), 218(1), 176(100), 96(6), 70(10). OD. Tentatively an 8-butyny 1-5-propyl I. Mantelline. Tricyclic. 'C15H25N'. EI: 219(31), 218(30), 204(16), 190(14), 178(20), 152(100), 136(52), 112(23), 99(57), 91(34), 86(28), 70(40). OD. Mantelline.

Alkaloids from Amphibian Skins

219L.

123

5,8-1. C15H25N. EI: 219(<1), 152(100), 96(21). FTIR: Strong, sharp Bohlmann band 2788 cm'^;C=CH2 3090 cm'^;conj.CH=CH 3030 cm-^ OD. Mantelline. Figure 21. 219M. Unclass. •C15H25N'. EI: 219(<1), 150(100), 135(31), 134(16), 121(60), 107(14). Dendrobatid. 221A. 5,8-1. 'Ci5H27N'. EI: 221(<1), 220(2), 138(100), 96(9). 0D,H2. Tentatively a 5hexenyl-8-methyl I. Dendrobatid. 221B. Unclass. 'C14H23NO'. EI: 221(2), 192(100). 1D,H2. Dendrobatid. 221c. DHQ. C15H27N. EI: 221(2), 152(100). 1D,H2. Tentatively a 5-methy 1-2pentenyl DHQ. Dendrobatid. 221D. DHQ. C15H27N. EI: 221(3), 180(100). 1D,H2, Tentatively a 2-ally 1-5-propyl DHQ. Dendrobatid. 221E. Unclass. 'C15H27N'. IT-EI: 222(45), 220(8), 152(100), 148(28), 134(10). OD. Bufonid. 221F. DehydrohPTX. •C14H23NO'. IT-EI: 222(40), 176(10), 162(100), 160(40), 148(18), 134(42), 120(38), 91(35). Proposed to be a member of a postulated dehydrohPTX subclass [56]. Alkaloid 221J is probably an isomer. Other structures are possible, see245F. Mantelline. Figure 10. 221G. Tricyclic. C15H27N. EI: 221(33), 220(47), 206(17), 179(19), 178(16), 150(19), 136(16), 124(14), 110(27), 98(70), 97(100), 83(53). FTIR: Moderate Bohlmann bands 2800, 2758 cm"^ OD. Two isomers detected. Dendrobatid. 221H. Unclass. 'C15H27N'. EI: 221(14), 220(11), 178(57), 166(100), 136(45). FTIR: Moderate, sharp Bohlmann band 2795 cm"^. Dendrobatid. 2211. 5,8-1. C15H27N. EI: 221(<1), 152(100), 96(11). FTIR: Strong, sharp Bohlmann band 2787 cm"^;c/^CH=CH 3020 cm"^ OD. Mantelline. Figure 21. 221J. Unclass. 'C15H27N'. IT-EI: 222(1), 220(3), 164(100), 162(65), 120(28). Probably an isomer of 221F. See also 245F. Dendrobatid. 221K. 5,8-1. 'C15H27N'. IT-EI: 222(1), 180(100), 96(35). OD. Tentatively an 8-butyl-5propenyl I. Dendrobatid. 221L. Pip. 'C15H27N'. IT-EI: 222(<1), 98(100). ID. Dendrobatid. Figure 37. 221M. Tricyclic. 'C15H27N'. EI: 221(3), 220(5), 206(8), 192(50), 178(17), 164(34), 150(100), 136(22), 126(21), 124(29), 110(22), 70(100). FTIR: Moderate Bohlmann band 2800 cm"^ OD. Ahigherhomologof 207R. Dendrobatid. 222. SpiroP. C13H22N2O. EI: 222(1), 221(2), 112(100), 82(16). FTIR spectrum [324]: Weak Bohlmann band 2827 cm"^; =NOH 3641 cm"^ NMR spectrum [105] and data [324]. ID. Dendrobatid. Figure 33. 223AB. 3,5-1. C15H29N. EI: 5£,9£: 223(<1), 180(88), 166(100); 5^92: 223(<1), 180(27), 166(100); 5E,9Z: 223(<1), 180(100), 166(75); 5Z,9£: 223(<1), 180(100), 166(86). IT-EI: Minor ion at m/z 124 for all diastereomers. FTIR spectra for all diastereomers [86]: Bohlmann bands moderate to none 5E,9E 2795 cm"^>5Z,9Z 2792 cm-^>5E,9Z 2803 cm-^»5Z,9£ none. NMR data for 5EM [162,209]. OD,

124

J. W. Daly, H. M. Garraffo and T. F. Spande

Ho. Dendrobatid (all diastereomers), bufonid (5Z,9Z>5£,9Z>5E,9F), mantelline (5Z,9Z). Figure 19. 223A. 5,6,8-1. C15H29N. EI: 223(3), 222(2), 180(100), 124(10). FTIR spectrum [207]: Strong, sharp Bohlmann band 2784 cm"^ NMR spectrum and data [207]. OD, HQ. Dendrobatid, mantelline. Figure 23. 223B. 3,5-P. C15H29N. EI: 223(1), 222(2), 166(100). FTIR spectra [86]: Cw-223B: Weak Bohlmann band 2802 cm"^ Trans-223B: No Bohlmann band. OD, HQ. Dendrobatid, bufonid. Figure 17. 223C. Izidine. 'C15H29N'. EI: 223(1), 222(2), 152(100), 110(16). OD, H2. Either a 5,6,8I or a 1,4-Q. Dendrobatid. 223D. 5,8-1. 'C15H29N'. EI: 223(2), 222(1), 138(100), 96(28). OD. Tentatively a 5hexyl-8-methyl I. Dendrobatid. 223E. Unclass. •C14H25NO'. EI: 223(2), 222(3), 168(100). 1D,H2. Dendrobatid. 223F. DHQ. C15H29N. EI: 223(2), 222(1), 180(100). FTIR spectra [86]: Cz5-223F: Weak Bohlmann band 2801 cm"^ rrfl«i:-223F: Virtually no Bohlmann band. ID, HQ. A S-epi isomer of trans-223¥ has been detected. Dendrobatid, bufonid. Figure 13. 223G. hPTX. C14H25NO. EI: 223(10), 208(6), 180(37), 98(23), 84(100). FTIR spectrum [56]: Moderate, broad Bohlmann band 2753 cm"^; OH 3555 cm"^ NMR spectrum [105]. 1D,H2. Dendrobatid, mantelline. Figure 7. 223H. 3,5-P. C15H29N. EI: 223(1), 208(4), 124(100). FTIR spectrum [86]: Cw-223H: Very weak Bohlmann band 2803 cm'^ OD, HQ. A minor isomer has also been detected. Dendrobatid, mantelline, bufonid. Figure 17. 2231. 5,8-1. C15H29N. EI: 223(3), 222(1), 194(7), 180(100), 166(8), 152(12), 138(7), 96(10). FTIR spectrum [281]: Weak Bohlmann band 2781 cm'^ Dendrobatid. Figure 21. 223J. 5,8-1. C15H29N. EI: 223(<1), 222(1), 166(100), 96(11). FTIR spectrum [281]: Strong, sharp Bohlmann band 2786 cm"^ OD. Some early reports of 223B [1] may instead correspond to 223 J. Dendrobatid, mantelline. Figure 21. 223K. Pip. 'C15H29N'. EI: 223(1), 222(1), 208(3), 124(31), 111(23), 98(100). ID. Dendrobatid. Figure 37. 223L. Unclass. 'C15H29N'. EI: 223(1), 222(2), 208(2), 194(23), 180(40), 167(53), 166(24), 152(18), 137(20), 138(28), 124(15), 96(42), 82(100). OD. Dendrobatid. 223M. 3,5-P. C,5H29N. Cis- and trans-223M: EI: 223(3), 222(2), 180(55), 152(100). FTIR: Virtually no Bohlmann band. OD. Mantelline. Figure 17. 223N. Pyr. C15H29N. EI: 223(12), 222(6), 152(85), 140(100). ID. Dendrobatid, mantelline. Figure 35. 2230. Unclass. C13H21NO2. EI: 223(<1), 222(2), 206(4), 184(100), 166(11), 148(7), 134(9), 120(13), 70(87). FTIR: Strong, sharp Bohlmann band 2810 cm'^; C=C=C 1955 cm"^; OH 3658, 3511 cm'^ Mantelline.

Alkaloids from Amphibian Skins

223P.

125

Unclass. 'CMHSSNO'. EI: 223(22), 208(11), 206(15), 180(9), 168(14), 164(50), 152(35), 150(100), 138(35), 136(39), 122(28), 110(58), 96(58), 81(42), 70(88). ID. Dendrobatid. 223Q. DHQ. 'Ci5H29N'. EI: 223(11), 222(9), 208(45), 180(67), 152(100). FTIR: Weak Bohlmann band 2801 cm'^. ID. Tentatively a c/5-2-methyl-5-pentyl DHQ. Dendrobatid. 223R. 3,5-1. 'C15H29N'. EI: 223(<1), 222(1), 138(100). OD. Tentatively a 3-hexyl-5methyll. Dendrobatid. 223S. DHQ. 'C15H29N'. EI: 223(2), 180(100), 152(38). FTIR: Moderate Bohlmann band 2802 cm'^ ID. Tentatively a 2-pentyl-5-methyl DHQ. Dendrobatid. 225A. Unclass. 'CnHaTNO'. EI: 225(3), 224(6), 208(2), 168(100), 152(25). ID, HQ. Dendrobatid. 225B. Pip. C15H31N. EI: 225(1), 224(1), 154(100). lD,Ho. Dendrobatid. Figure 37. 225C. Pyr. C15H31N. EI: 225(1), 224(2), 168(70), 126(100). lD,Ho. Dendrobatid. Figure 35. 225D. 5,8-1. C14H27NO. EI: 225(<1), 138(100), 96(12). lD,Ho. Tentatively a 5(hydroxypentyl)-8-methyl I. Dendrobatid. 225E. aPTX. C13H23NO2. EI: 225(10), 208(28), 182(16), 114(27), 112(38), 70(100). NMR [cited inref. 57]. 2D. Dendrobatid. Figure 3. 225F. PTX. C13H23NO2. EI: 225(21), 194(28), 166(79), 112(19), 84(26), 70(100). FTIR spectrum [57]: Strong, sharp Bohlmann band 2794 cm"^ shoulder 2750 cm'^; homoallylic OH 3650, 3600 cm"^; OH 3542 cm'^ NMR data [57]. 2D. 0-Acetyl derivative. Dendrobatid. Figure 3. 225G. Unclass. C14H27NO. EI: 225(8), 138(100), 125(63), 110(33), 96(9), 82(10), 70(23). FTIR: Weak Bohlmann band 2791 cm"^; OH 3638 cm'^ A hydroxy congener of 209G. Dendrobatid. 225H. Pyr. C15H31N. EI: 225(<1), 154(100), 140(91). FTIR: Cis-llSU: Weak Bohlmann band 2798 cm"^ rrfl«5-225H: No Bohlmann band. ID. Dendrobatid, mantelline. Figure 35. 2251. Pip. 'Ci5H3iN'. EI: 225(<1), 224(1), 98(100). ID. Dendrobatid. Figure 37. 231A. 1,4-Q. C16H25N. EI: 231(2), 230(1), 166(100), 110(12). FTIR spectrum and data [56] (Figure 28): Moderate, broad Bohlmann band, 2789 cm^^ conj CH=CH 3029 cm"^;C=CH3328cm'^ OD, H6. A minor isomer has been detected. Dendrobatid, mantelline. Figure 27. 231B. 5,6,8-1. C16H25N. EI: 231(2), 230(1), 152(100), 110(13). FTIR (Figure 24): Strong, sharp Bohlmann band 2788 cm"*; conj CH=CH 3031 cm"*; C=CH 3327 cm"^ OD, He. Previously proposed to be a 1,4-Q [11]. A minor isomer has been detected. Dendrobatid. Figure 23. 231C. 5,8-1. C,6H25N. EI: 231(3), 230(4), 138(100), 96(19). FTIR: Strong, sharp Bohlmann band 2787 cm"*; conj CH=CH 3034 cm"*; C=CH 3327 cm'^ OD, Hg. Dendrobatid. Figure 21. 231D. Unclass. 'C15H21NO'. EI: 231(1), 154(100). ID, He- Dendrobatid.

126

231E. 231F. 231G. 231H.

2311. 233A.

233B. 233C. 233D.

233E. 233F.

233G. 233H.

2331. 233J. 235A.

235B.

235C.

J. W. Daly, H. M. Garraffo and T. F. Spande

DHQ. 'CifiHssN'. EI: 231(<1), 230(3), 152(100). ID, He- Tentatively, a 2hexenynyl-5-methyl DHQ. Dendrobatid. Unclass. 'C16H25N'. EI: 231(1), 180(100). 0D,H6. Dendrobatid. Unclass. 'Ci6H25N'. EI: 231(4), 232(2), 202(10), 152(42), 138(100), 110(8), 96(21). Dendrobatid. Unclass. C16H25N. EI: 231(<1), 216(16), 162(17), 160(20), 150(100), 148(28), 134(17), 122(10), 107(12). FTIR: Weak Bohlmann band 2801 cm'^; C=CH 3329 cm•^ Two isomers detected. Dendrobatid. Unclass. C16H25N. EI: 231(18), 190(100), 178(14), 148(11). Dendrobatid. 1,4-Q. C,6H27N. EI: 233(2), 232(1), 166(100), 110(4). FTIR data [56]: Moderate, broad Bohlmann band 2789 cm"^; C=CH 3329 cm'^ OD, H4 Mantelline. Figure 27. Unclass. 'C15H23NO'. EI: 233(<1), 168(100). ID. Dendrobatid. Unclass. 'C,6H27N'. EI: 233(1), 216(1), 192(100). OD. Dendrobatid. 5,8-1. C16H27N. EI: 233(3), 232(2), 164(17), 151(17), 138(100), 96(20). FTIR: Strong, sharp Bohlmann band 2787 cm^^; cis CH=CH 3017 cm"^; C=CH2 3083 cm-^ NMR data [161]. OD, H4. Dendrobatid. Figure 21. Unclass. C,6H27N. EI: 233(4), 232(9), 152(36), 136(100), 134(32). Dendrobatid. DehydrohPTX. C,5H23NO. EI: 233(12), 232(27), 218(14), 162(100), 160(31), 134(16), 120(11). FTIR data [56]: Weak Bohlmann band 2790 cm"*; cis CH=CH 3020 cm'^; C=0 1731 cm"^ Proposed to be a member of a postulated dehydrohPTX subclass [56]. Other structures are possible, see 245F. Mantelline. Figure 10. 5,6,8-1. C16H27N. EI: 233(2), 231(1), 152(100), 110(9). FTIR: Strong, sharp Bohlmann band 2785 cm"^;C=CH 3329 cm"^ Dendrobatid. Figure 23. Unclass. 'CisHssNO'. EI: 233(66), 218(23), 202(100), 159(17), 131(14), 115(33), 82(50). FTIR: Weak Bohlmann band 2784 cm"^; C=CH2 3067 cm"^; strong bands 1487, 1275, 1050 cm-'. Dendrobatid. Unclass. •C,6H27N'. EI: 233(1), 232(<1), 190(100), 138(8), 110(9), 84(19). Mantelline. Unclass. 'Ci6H27N\ IT-EI: 234(32), 233(37), 232(12), 218(24), 204(15), 190(25), 178(100), 176(30), 148(19), 108(17). Dendrobatid. HTX. C15H25NO. EI: 235(5), 234(2), 218(15), 194(76), 176(25), 150(28), 96(100). FTIR spectrum [110]: No Bohlmann band; OH broad 3345 cm"'; C=CH2 3084 cm-^ 2D,H4. Dendrobatid. Figure 11. 5,8-1. C16H29N. 235B': EI: 235(2), 234(1), 138(100), 96(10). FTIR: Strong, sharp Bohlmann band 2787 cm"'; C=CH2 3083 cm"^ NMR data [60]. OD, H2. 235B" (formerly 235B): EI: 235(2), 234(1), 138(100), 96(10). FTIR data [56]: Strong, sharp Bohlmann band 2787 cm"'; cis CH=CH 3010 cm"^ NMR spectrum and data [105,161]. 0D,H2. Dendrobatid, mantelline. Figure 21. DehydrohPTX. C15H25NO. EI: 235(28), 234(53), 220(20), 176(11), 162(100), 160(25), 134(23), 120(15). FTIR spectrum [56]: Weak Bohlmann band 2792 cm"';

Alkaloids from Amphibian Skins

127

conj CH=CH 3029 cm'^; OH 3654 cm"^ ID, H2 and H4. O-Acetyl derivative. Two diastereomers. Proposed to be a member of a postulated dehydrohPTX subclass [56]. Other structures are possible, see 245F. Mantelline. Figure 10. 235D. Unclass. 'Ci5H25NO'. EI: 235(<1), 196(20), 170(100). 1D,H6. Dendrobatid. 235E. 1,4-Q. 'C16H29N'. EI: 235(5), 152(100), 110(18), 70(13). FTIR spectrum [86, mislabeled 223E']: 235E': Strong Bohlmann band 2787 cm"^; cis CH=CH 3020 cm"^ OD, H2. Two isomers have been detected. There is a strong possibility that this alkaloid is instead a 6,8-dimethyl-5-hexenyl indolizidine, based on the shape of the relatively sharp Bohlmann band and the presence of an m/z 70 fragment typical of such indolizidines. Dendrobatid, bufonid. Figure 27 (the previously proposed quinolizidine structure). 235F. Unclass. 'C16H29N'. EI: 235(5), 234(3), 166(36), 138(100). OD. Dendrobatid. 235G. Unclass. 'C16H29N'. EI: 235(2), 206(100), 194(65). Dendrobatid. 235H. CPQ. 'C,6H29N'. EI: 235(60), 234(68), 220(28), 178(48), 164(20), 152(28), 150(32), 112(50), 111(100), 98(52), 96(40). OD. Dendrobatid. Figure 31. 2351. Tricyclic. C16H29N. EI: 235(63), 234(100), 220(61), 208(12), 206(21), 192(26), 178(11), 150(20), 138(18), 136(50), 122(21), 110(20), 96(20), 84(27), 70(30). FTIR spectrum [86]: Strong, sharp Bohlmann band 2791 cm"^ Bufonid. 235J. Unclass. 'C15H25NO'. IT-EI: 235(12), 220(2), 208(2), 192(2), 138(8), 109(18), 84(100). FTIR data [56]: Moderate Bohlmann band 2755 cm-^ ID. Previously postulated to be an hPTX [56]. Mantelline. 235K. Tricyclic. C16H29N. EI: 235(6), 234(21), 220(4), 206(7), 192(100), 178(33), 164(14), 150(15), 136(14), 122(9). FTIR: Weak Bohlmann band 2792 cm"^; C=CH2 3084 cm-^ OD. Mantelline. 235L. Unclass. Ci4H2,N02. EI: 235(<1), 192(11), 170(100,C9Hi6NO2), 152 (34,C9H,4NO), 136(6), 134(14,C9Hi2N), 112(8), 96(19), 70(30). FTIR: Strong, sharp Bohlmann band 2806 cm"^; conj CH=CH 3036 cm"^; C=CH 3328 cm"^; OH 3680, 3550 cm"^ Mantelline. 235M. Tricyclic. C16H29N. EI: 235(24), 234(100), 220(13), 206(24), 192(38), 166(29), 164(32), 151(25), 150(28), 136(24), 122(8). FTIR: No Bohlmann band. Dendrobatid, mantelline. 235N. Unclass. C15H25NO. EI: 235(2), 234(1), 194(5), 168(100,CioHi8NO), 112(4). Mantelline. 2350. Unclass. C16H29N. EI: 235(<1), 234(2), 192(100). Mantelline. 235P. Tricyclic. C16H29N. EI: 235(6), 234(5), 220(10), 206(82), 192(13), 178(31), 164(22), 150(46), 140(52), 138(58), 124(28), 122(42), 98(63), 70(100). OD. Dendrobatid. 236. SpiroP. C,4H24N20. EI: 236(8), 126(100). FTIR spectrum [324]: No Bohlmann band, =N-0CH31055, 864 cm^^ NMR spectrum [105] and data [324]. OD. Dendrobatid, bufonid, mantelline. Figure 33.

128

237A.

J. W. Daly, H. M. Garraffo and T. F. Spande

PTX. C,5H27NO. EI: 237(6), 236(4), 194(20), 166(54), 84(19), 70(100). FTIR: Moderate Bohlmann bands 2797, 2749 cm-^; OH 3545 cm-^ ID, H2. Dendrobatid, mantelline. Figure 3. 237B. aPTX. C14H23NO2. EI: 237(11), 182(60), 114(30), 112(25), 70(100). 2D. Dendrobatid. Figure 3. 237C. Izidine. 'C16H31N'. EI: 237(1), 236(2), 180(100), 110(7), 96(8), 70(13). OD, HQ. Probably a 5,6,8-1. Previously proposed to be a 1,4-Q [11]. Dendrobatid. 237D. 5,8-1. 'C16H31N'. EI: 237(1), 236(<1), 138(100), 96(10). OD, HQ. Dendrobatid. Figure 21. 237E. 3,5-1. 'C15H27NO'. EI: 237(1), 236(3), 208(70), 152(100). H2. Tentatively a 3ethyl-5-(hydroxypentenyl) I. Dendrobatid. 237F. HTX. C15H27NO. EI: 237(13), 220(11), 194(39), 176(12), 166(27), 152(58), 139(24), 110(10), 96(100). 2D,H2. Dendrobatid. Figure 11. 237G. 3,5-P. 'C15H27NO'. IT-EI: 238(2), 124(100). FTIR data [86]: as'237G: Very weak Bohlmann band 2804 cm'^ Dendrobatid, bufonid. Figure 17. 237H. 5,8-1. 'C15H27NO'. EI: 237(<1), 236(1), 152(100), 96(11). FTIR: Strong, sharp Bohlmann band 2788 cm"^; OH 3646 cm"^; cis CH=CH 3010 cm"^ Previously proposed to be an 8-ethyl-5-hexyl I [56]. Mantelline. Figure 21. 2371. 4,6-Q. C16H31N. EI: 237(2), 236(1), 194(45), 180(100), 138(12), 110(8), 97(15). Dendrobatid. Figure 25. 237J. Pip. 'Ci6H3,N'. EI: 237(<1), 98(100). ID. Dendrobatid. Figure 37. 237K. Unclass. C15H27NO. EI: 237(23), 236(11), 222(13), 178(100, C12H20N), 164(53, CiiHigN). Mantelline. 237L. 5,6,8-1. C16H31N. IT-EI: 238(2), 180(100), 124(15). Dendrobatid. Figure 23. 237M. Unclass. 'C15H27NO'. IT-EI: 238(26), 237(8), 210(4), 168(100), 150(14), 110(68), 70(25). FTIR: No Bohlmann band; OH 3656 cm"^ Dendrobatid. 237N. Unclass. 'C15H27NO'. IT-EI: 238(25), 206(18), 192(100), 124(15). Dendrobatid. 238. Nitropolyzonamine. C13H22N2O2. EI: 238(2), 122(20), 108(23), 82(100). FTIR spectrum (Figure 34): Moderate Bohlmann band 2810 cm'^; NO21562, 1367 cm"^ Dendrobatid. Figure 33. 239AB. 3,5-1. C15H29NO. EI: 239(2), 238(3), 182(100), 180(90). FTIR (Figure 20): Weak Bohlmann band 2798 cm-^; OH 3670 cm-^ NMR data [162,209]. lD,Ho. OAcetyl derivative. Dendrobatid. Figure 19. 239A. 5,8-1. 'C15H29NO'. EI: 239(2), 238(3), 182(100). ID, HQ. Tentatively a 5-butyl-8(hydroxypropyl) I. Dendrobatid. 239B. 5,8-1. 'C15H29NO'. EI: 239(2), 238(3), 180(100). ID, HQ. Tentatively an 8-butyl5-(hydroxypropyl) I. Dendrobatid. 239CD. 3,5-1. C15H29NO. EI:239(4),238(3), 196(100), 166(60). FTIR: Weak Bohlmann band 2796 cm'^; OH 3669 cm'^ NMR data [162,209]. ID, HQ. 0-Acetyl derivative. Dendrobatid. Figure 19.

Alkaloids from Amphibian Skins

239C.

129

5,8-1. C15H29NO. EI: 239(<1), 238(1), 196(100), 124(4), 96(4), 70(4). FTIR: Strong, sharp Bohlmann band 2788 cm"^; OH 3660 cm"^ ID, HQ. A minor isomer has been detected. Dendrobatid, mantelline. Figure 21. 239D. 5,8-1. 'C15H29NO'. EI: 239(2), 238(3), 166(100). HQ. Tentatively a 5(hydroxybutyl)-8-propyl I. Dendrobatid. 239E. 3,5-1. 'C15H29NO'. EI: 239(2), 238(3), 210(40), 152(100). lD,Ho. Tentatively a 3-ethyl-5-(hydroxypentyl) I. Dendrobatid. 239F. 5,8-1. 'Ci5H29NO'. EI: 239(1), 168(100). lD,Ho. Tentatively an 8(hydroxyethyl)-5-pentyl I. Dendrobatid. 239G. 5,8-1. C15H29NO. EI: 239(1), 238(3), 138(100), 96(8). lD,Ho. Tentatively a 5(hydroxyhexyl)-8-methyl I. Dendrobatid. 239H. HTX. C15H29NO. EI: 239(11), 222(16), 196(51), 168(39), 152(100), 139(33), 110(11), 96(75). FTIR (Figure 12): No Bohlmann band; OH broad 3328 cm"^ 2D, HQ. Dendrobatid. Figure 11. 2391. Pip, C16H33N. EI:239(3), 182(40), 140(100). lD,Ho. Dendrobatid. Figure 37. 239J. Noranabasamine. C15H17N3. EI: 239(75), 238(30), 210(25), 183(20), 182(35), 157(80), 84(100). ID, Ho. Dendrobatid. Figure 39. 239K. 3,5-P. 'C15H29NO'. EI: 239(<1), 224(2), 124(100). FTIR: C/5-239K: Very weak Bohlmann band 2800 cm"^; OH 3560 cm"^ Trans-239K: No Bohlmann band; OH 3560 cm'^ Mantelline. Figure 17. 239L. Pip. C15H29NO. EI: 239(<1), 238(4), 224(6), 98(100). FTIR: C/5-239L: Very weak Bohlmann band 2810 cm"^; C=0 1733 cm"^ Trans-239L: No Bohlmann band; C=0 1733 cm"^ Dendrobatid. Figure 37. 239M. hPTX. C14H25NO2. EI: 239(10), 238(4), 208(14), 180(56), 96(23), 84(100). FTIR: Moderate, broad Bohlmann band 2760 cm"^; homoallyHc OH 3610, 3580 cm-^;OH3550cm"^ 2D. Mantelline. Figure 7. 239N. Unclass. C17H21N. EI: 239(11), 200(100). Mantelline. 2390. Pip. 'C16H33N'. EI: 239(2), 168(85), 154(100). ID. Dendrobatid. Figure 37. 241A. Unclass. 'Ci4H27N02'. EI: 241(2), 240(3), 166(100), 126(48). Dendrobatid. 241B. Unclass. C16H35N. EI: 241(15), 125(45), 58(100). OD,Ho. Possibly an aliphatic dimethylamine. Other alkaloids with a base peak at m/z 58 are 197D, 21IG, 253E, 2671,2670, 269E. Mantelline (pet trade). 241C. 5,8-1. •C14H27NO2'. EI: 241(1), 152(100). Tentatively a 5-(dihydroxybutyl)-8ethyll. Dendrobatid. 241D. Pip. C15H31NO. EI: 241(<1), 240(2), 226(6), 114(100), 96(8), 70(28). FTIR spectrum synthetic [347]: Weak Bohlmann band 2808 cm"^ OH 3649 cm'^ NMR data [60]. 2D,Ho. Dendrobatid. Figure 37. 241E. Unclass. 'Ci4H27N02'. EI: 241(3), 222(62), 154(100). 2D. Dendrobatid. 241F. 5,8-1. C17H23N. EI: 241(<1), 176(100), 96(12). FTIR: Strong, sharp Bohlmann band 2790 cm"^; conj CH=CH 3039 cm"^; two C^CH 3328 cm'^ Mantelline. Figure 21. 241G. Pip. C14H27NO2. EI: 241(<1), 226(6), 98(100). Dendrobatid. Figure 37.

130

241H. 243A.

243B. 243C.

243D.

243E. 243F. 245A. 245B.

245C.

245D. 245E. 245F.

J. W. Daly, H. M. Garraffo and T. F. Spande

aPTX. 'Ci3H23N03'. IT-EI: 242(5), 240(3), 182(15), 114(100), 96(14), 84(15), 70(78). 3D. Dendrobatid. Figure 3. DHQ. C17H25N. EI: 243(2), 242(1), 202(100). FTIR spectra [161]: Cis-243A: No Bohlmann band. Trans-243A and 5-epi-tranS'243A: Weak Bohlmann band 2805 cm"^ NMR spectra and data [161,162]. In ref. 162, the NMR data for 243A' appears to be that reported for tranS'243A in ref. 161 and the data for 243A in ref. 162 appears to be that reported in ref 161 for 5-epi-trans-243A. ID, Hg. A^Acetyl derivative. Dendrobatid. Figure 13. 5,8-1. C17H25N. EI: 243(<1), 176(100), 96(12). OD, Hg. Dendrobatid, mantelline. Figure 21. 5,8-1. C17H25N. EI: 243(<1), 178(100), 96(15). FTIR data [56]: Strong, sharp Bohlmann band 2789 cm"^; C=CH2 3085 cm"^; cis CH=CH 3020 cm'^; C=CH 3328 cm'^ OD. Mantelline. Figure 21. 5,8-1. 'C17H25N'. IT-EI: 243(<1), 242(17), 176(18), 164(12), 152(32), 122(44), 96(45), 91(50), 70(100). FTIR data [56]: Strong, sharp Bohlmann band 2789 cm'^ conj CH=CH 3035 cm"^; trans CH=CH 970 cm"^; C^CH 3328 cm'^ OD. Mantelline. Figure 21. Unclass. 'C^HssN'. EI: 243(3), 242(35), 214(27), 200(30), 176(40), 164(29), 146(48), 124(100), 122(55), 96(42). Dendrobatid. Unclass. C17H25N. IT-EI: 244(<1), 242(12), 150(100), 148(93), 120(34). See 245F. Dendrobatid. CPQ. 'C16H23NO'. EI: 245(20), 109(100), 108(55), 107(60), 94(30). Dendrobatid. Figure 31. 5,8-1. C17H27N. EI: 245(2), 244(1), 178(100), 96(9). FTIR data [56]: Strong, sharp Bohlmann band 2788 cm"^; C=CH2 3085 cm'^; CsCH 3328 cm"^ OD. Mantelline. Figure 21. 5,8-1. 'C17H27N'. IT-EI: 245(<1), 244(15), 216(20), 206(10), 204(10), 202(10), 188(15), 175(15), 174(17), 164(15), 152(32), 134(15), 132(15), 122(35), 96(45), 91(48), 79(60), 70(100). FTIR data [56]: Strong, sharp Bohlmann 2788 cm"^; /ra«5CH=CH 970 cm-^;CHCH 3329 cm"^ OD. Mantelline. Figure 21. 5,8-1. 'C,7H27N'. EI: 245(<1), 138(100), 96(10). OD. Tentatively an 8-methyl-5octenynyl I. Dendrobatid. DHQ. 'C17H27N'. EI: 245(5), 244(3), 202(100), 180(43), 178(19). FTIR data [124]: CW-245E: Weak Bohlmann band 2807 cm"^ Dendrobatid. Figure 13. Unclass. C17H27N. EI: 245(<1), 244(2), 216(6), 150(100), 148(21), 120(12). FTIR: Moderate, sharp Bohlmann band 2787 cm"^;CsCH 3328 cm"^ OD, He. The perhydro derivative affords a base peak at m/z 152 and a fragment ion at m/z 96 diagnostic for a 5,8-1. Tentatively, 245F is a member of a putative 6,7-dehydro5,8-1 class of alkaloids; see also 201 A, 207E, 219G, 221J, 243F, 265F, 269D, 275D. All have a significant fragment ion at m/z 120 (CgHioN"*") as do putative dehydrohPTXs 221F, 233F, 235C, 251G. Mantelline.

Alkaloids from Amphibian Skins

245G. 247A. 247B. 247C.

247D. 247E. 249A.

249B.

249C.

249D.

249E. 249F.

249G. 249H.

2491. 249J. 249K. 249L.

131

5,6,8-1. 'C17H27N'. IT-EI: 246(4), 245(2), 192(12), 166(100), 124(18). OD. Tentatively a 6-ethyl-5-hexenynyl-8-methyl I. Dendrobatid. CPQ. C16H25NO. EI: 247(15), 110(37), 109(100). FTIRdata [306]: Enamine 1692 cm-^ 1D,H4. Dendrobatid. Figure 31. Unclass. 'C17H29N'. EI: 247(1), 192(4), 178(100). OD. Dendrobatid. 3,5-1. C17H29N. EI: 247(2), 246(1), 192(100), 178(37), 138(6), 124(5). FTIR: Moderate Bohlmann band 2787 cm'^;C=CH2 3089 cm"^ OD. Tentatively a 5^,9£-3-butenyl-5-pentenyl I. Mantelline. 1,4-Q. C17H29N. EI: 247(<1), 166(100), 110(11). FTIR: Moderate, broad Bohlmann band 2790 cm"^; C=CH 3328 cm"^ Mantelline. Figure 27. 5,8-1. 'C,7H29N'. IT-EI: 248(<1), 204(12), 178(100), 96(27), 70(25). Tentatively a 8-butenyl-5-pentenyl I. Dendrobatid. 3,5-1. C17H31N. EI: 249(3), 248(2), 192(100), 180(25). FTIR spectrum [56]: Weak Bohlmann band 2791 cm'^; C=CH2 3085 cm'^ Dendrobatid, mantelline. Figure 19. CPQ. •C16H27NO'. EI: 249(18), 234(48), 222(26), 221(98), 220(100), 206(30), 192(28), 186(13), 178(20), 172(63), 168(30), 166(30), 164(48), 152(100), 136(18), 124(15), 114(53), i l 1(66), 98(30). OD. Dendrobatid. Figure 31. Unclass. C17H31N. EI: 249(4), 248(1), 220(1), 206(2), 192(4), 178(6), 152(100), 110(12), 95(5), 70(9). FTIR: Weak Bohlmann band 2811 cm'^ OD, HQ. Probably a tricyclic. Previously proposed to be a 1,4-Q [56]. Dendrobatid, mantelline. DHQ. 'CnHsiN'. EI: 249(2), 248(3), 206(100), 180(15). FTIRdata [86]: Cis249D: Moderate Bohlmann band 2804 cm"^; C=CH2 3085 cm•^ TranS'249D: No Bohlmann band; C=CH2 3085 cm"^ ID, H2. Bufonid. Figure 13. DHQ. 'C17H31N'. IT-ET: 250(15), 206(29), 180(100). FTIRdata [86]: Trans249E: No Bohlmann band; C=CH2 3084 cm-^ 1D,H2. Bufonid. Figure 13. Unclass. 'C16H27NO'. IT-EI: 249(12), 220(22), 123(16), 84(100). FTIR data [56]: Moderate, broad Bohlmann band 2755 cm"^; OH 3540 cm"^ Previously proposed to be a hPTX [56]. Mantelline. DesmethylPTX. 'C16H27NO'. IT-EI: 249(14), 152(93), 124(25), 96(34), 70(100). Dendrobatid. Figure 5. 5,6,8-1. CnHsiN. EI: 249(7), 220(6), 166(100), 110(8), 95(10), 70(16). FTIR: Very weak Bohlmann band. OD, H2. NMR data [295a]. A minor isomer has been detected. Dendrobatid, mantelline. Figure 23. 3,5-P. 'C17H31N'. EI:249(5), 192(75), 166(100). Tentatively a 3-buty 1-5-hexenyl P. Dendrobatid. Unclass. C16H27NO. EI: 249(2), 176(100), 124(12), 96(15). FTIR: Moderate Bohlmann band 2800 cm'^; conj CH=CH 3030 cm"^; OH 3525 cm"^ Mantelline. Unclass. C16H27NO. EI: 249(2), 248(3), 192(11), 136(100). FTIR: Moderate Bohlmann band 2787 cm^^; cis CH=CH 3015 cm"^; C=0 1731 cm^^ Mantelline. 5,8-1. C16H27NO. IT-EI: 250(<1), 138(100), 96(23). Tentatively an 8-methyl I with a 5-C7H11O substituent. Dendrobatid.

132

251A.

J. W. Daly, H. M. Garraffo and T. F. Spande

DHQ. 'CnHssN'. EI: 251(2), 208(6), 152(100). ID, HQ. Tentatively a 2-hepty 1-5methylDHQ. Dendrobatid. 251B. 5,8-1. C16H29NO. EI: 251(2), 250(1), 164(8), 151(8), 138(100), 96(9), 70(7). FTIR: Strong, sharp Bohlmann band 2787 cm-^; cis CH=CH 3012 cm"^; OH 3647 cm'^ NMR data [161]. ID, H2. Two isomers have been detected. Dendrobatid, mantelline. Figure 21. 251C. Unclass. •Ci6H29NO'. EI: 251(2), 234(4), 154(100). 1D,H2. Dendrobatid. 251D. PTX. C,6H29NO. EI: 251(6), 250(4), 194(16), 166(68), 84(12), 70(100). FTIR spectrum and data [86] (Figure 4): Moderate, broad Bohlmann band 2797 cm"^; OH 3544 cm"^ NMR spectrum [51] and data [52,57]. ID, H2. An isomer may be present in mantelline extracts. Dendrobatid, mantelline, bufonid. Figure 3. 251E. Unclass. •C16H29NO'. EI: 251(3), 250(1), 168(30), 84(18), 70(100). Dendrobatid. 251F. CPQ. C16H29NO. EI: 251(54), 250(65), 236(27), 222(28), 221(30), 220(68), 194(62), 164(19), 152(35), 150(17), 112(43), 111(100), 98(35). FTIR spectrum [306]: Strong Bohlmann band 2755 cm"^; OH 3666 cm"^ NMR spectrum and data [306]. ID, HQ. 0-Acetyl derivative. A minor diastereomer, 251F', has been detected [306]. Dendrobatid. Figure 31. 251G. DehydrohPTX. C15H25NO2. EI: 251(26), 250(45), 162(100), 160(40), 134(13), 120(10). 2D, H2. Proposed to be a member ofa postulated dehydrohPTX subclass [56]. Other structures are possible, see 245F. Mantelline. Figure 10. 251H. DeoxyPTX. 'Ci6H29NO'. EI: 251(3), 250(4), 178(10), 150(100), 70(20). FTIR spectrum and data [104]: Moderate Bohlmann bands 2790, 2740 cm"^; OH 3652 cm'^ NMR spectrum and data [104]. ID. A minor diastereomer has been detected. Dendrobatid. Figure 5. 2511. aPTX. •C15H25NO2'. EI: 251(7), 236(4), 210(15), 209(20), 182(11), 70(100). 2D, H4. Dendrobatid. Figure 3. 251J. CPQ. 'C16H29NO'. EI: 251(92), 250(86), 236(24), 234(63), 222(23), 195(23), 194(23), 178(27), 164(28), 152(100), 150(85), 112(38), 111(82), 98(42). ID. Dendrobatid. Figure 31. 251K. 3,5-P. C17H33N. EI: 251(4), 194(76), 166(100). FTIR spectrum and data [86] (Figure 18): Cis-251K: Very v^eak Bohlmann band 2805 cm'^ Trans-ISIK: No Bohlmann band. OD, HQ. Dendrobatid, bufonid. Figure 17. 251L. Unclass. C15H25NO2. EI: 251,176, 84 (cochromatographs with another alkaloid). Occurs mainly as an 0-acetate, which is tabulated as 293G. Previously proposed to be a hPTX [56]. Mantelline. 251M. 5,6,8-1. 'CnHsaN'. EI: 251(<1), 250(1), 180(100), 124(7). OD, Dendrobatid, mantelline. Figure 23. 251N. 5,8-1. 'C17H33N'. EI: 251(<1), 250(2), 180(100), 96(13), 70(10). FTIR: Major isomer: Strong, sharp Bohlmann band 2787 cm'^ Minor isomer: Weak Bohlmann band 2804 cm'^ Dendrobatid. Figure 21. 2510. 3,5-P. C17H33N. EI: 251(2), 250(2), 208(35), 152(100). FTIR: Trans-lSlO: No Bohlmann band. OD. Mantelline. Figure 17.

Alkaloids from Amphibian Skins

251P.

133

Unclass. C16H29NO. EI: 251(5), 250(11), 236(4), 190(6, C13H20N), 136(100, C9H14N), 122(34), 70(17). FTIR: Moderate Bohlmann bands 2791, 2734 cm-^; OH 3660 cm"^ ID. Mantelline. 251Q. Unclass. C16H29NO. EI: 251(2), 250(1), 236(5), 122(100, CgH^N), 120(17). FTIR: Moderate Bohlmann band 2787 cm-^; conj CH=CH 3025 cm"^; OH 3654 cm"^ Two isomers detected. Mantelline. 251R. hPTX. C,6H29NO. EI: 251(1), 250(2), 208(10), 180(35), 84(100). ID. Dendrobatid, mantelline. Figure 7. 251S. Izidine, 'C16H29NO'. EI: 251(1), 250(1), 238(6), 152(100), 110(16). ID. Either a 5,6,8-1 or a 1,4-Q. Dendrobatid. 251T. Izidine. 'CnHgsN'. EI: 251(<1), 152(100), 110(31). OD. Either a 5,6,8-1 or a 1,4Q. Dendrobatid. 251U. 5,8-1. C16H29NO. EI: 251(<1), 250(<1), 164(6), 151(7), 138(100), 96(9), 70(5). FTIR: Strong, sharp Bohlmann band 2787 cm"^; C=0 1731 cm"^ Dendrobatid. Figure 21. 252A. SpiroP. C14H24N2O2. EI: 252(4), 251(3), 142(100). FTIR spectrum [324]: No Bohlmann band; OH 3611 cm'^; =N-0CH3 strong band 1048 cm"^ ID. Dendrobatid, mantelline. Figure 33. 252B. SpiroP. 'C14H24N2O2'. EI: 252(1), 221(5), 126(100). ID. Myobatrachid. 253A. aPTX. C15H27NO2. EI: 253(4), 236(22), 182(16), 114(27), 112(26), 98(15), 70(100). FTIR: Strong, sharp Bohhnann band 2803 cm^^; OH 3648, 3520 cm^^ Dendrobatid. Figure 3. 253B. 5,8-1. C16H31NO. EI: 253(1), 252(2), 238(4), 138(100), 96(9). FTIR data [56]: Strong, sharp Bohlmann band 2787 cm'^; OH 3650 cm'^ ID. Mantelline. Figure 21. 253C. Unclass. 'C16H31NO'. EI: 253(3), 192(100). ID. Dendrobatid. 253D. DHQ. C15H27NO2. EI: 253(<1), 222(6), 212(100). NMR spectrum and data [162]. 3D. Dendrobatid. Figure 13. 253E. Unclass. •Ci6H3,NO'. EI: 253(7), 180(23), 110(59), 58(100). Other alkaloids with base peak at m/z 58 are 197D, 211G, 241B, 2671, 2670, 269E. Dendrobatid. 253F. PTX. 'C15H27NO2'. EI: 253(<1), 235(6), 166(63), 98(23), 70(100). 2D. Dendrobatid. Figure 3. 253G. CPQ. 'C,5H27N02'. EI 253(26), 238(39), 224(14), 212(35), 210(56), 196(20), 182(44), 168(61), 154(50), 140(42), 126(100), 112(33), 111(36), 98(95), 84(23). Dendrobatid. 253H. 5,6,8-1. 'Ci6H3iNO'. EI: 253(<1), 196(100), 180(5), 124(32), 70(18). ID. Dendrobatid. Figure 23. 2531. Pyr. 'C17H35N'. IT-EI: 254(45), 168(45), 154(100). ID. Dendrobatid. Figure 35. 253J. Pip. 'C17H35N'. EI: 253(<1), 98(100). ID. Dendrobatid. Figure 37. 254. SpiroP. •C13H22N2O3'. EI: 254(<1), 122(8), 108(6), 82(100). An apparent hydroxynitropolyzonamine. Dendrobatid. 255A. Pip. C15H29NO2. EI: 255(3), 114(100). 2D,Ho. Dendrobatid. Figure 37.

134

255B. 256.

257A.

257B. 257C. 257D. 258.

259A.

259B.

259C.

261A.

261B. 261C.

261D.

263A.

263B.

J. W. Daly, H. M. Garraffo and T. F. Spande

Izidine. 'C18H25N'. IT-EI: 256(1), 255(<1), 190(100), 110(28). Either a 5,6,8-1 or a 1,4-Q. Dendrobatid. Pseudo. 'C16H20N2O'. EI: 256(22), 228(10), 227(10), 199(10), 185(28), 173(100), 171(20), 144(28), 130(95), 110(25), 109(30). FTIR data [25]. ID. Myobatrachid. Figure 40. Unclass. 'C18H27N'. EI: 257(1), 256(2), 216(100). FTIR: Moderate Bohlmann band 2781 cm"^; C=CH 3327 cm'^; C=CH2 3080 cm'^ cis CH=CH 3020 cm-^ OD, Hg. Dendrobatid. Unclass. 'C18H27N'. EI: 257(60), 256(100), 152(20). ID. Dendrobatid. 5,8-1. 'Ci8H27N'. EI: 257(<1), 138(100), 96(8). 0D,H8. Tentatively an 8-methyl5-nonadienynyl I. Dendrobatid. 1,4-Q. C18H27N. EI: 257(5), 256(3), 190(100), 110(9), 84(13). OD, Hg. Tentatively a l-butynyl-4-pentynyl Q. Dendrobatid. Pseudophrynaminol. C16H22N2O. EI: 258(25), 185(18), 173(100), 130(90). FTIR spectrum and data [24,25]: Moderate broad Bohlmann band 2802 cm"^; OH 3657 cm"^; NH 3429 cm"^ aryl H 3061 cm"^; strong, sharp bands 1606,1476, 1244, 1015 cm"^ NMRdata[24]. 2D. Myobatrachid. Figure 40. HTX. C17H25NO. EI: 259(4), 218(28), 200(14), 164(9), 150(18), 96(100). FTIR spectrum [110]: No Bohlmann band, C=CH2 3083 cm'^; conj CH=CH 3038 cm"^; CsCH 3328 cm'^; OH broad band 3330 cm'^ NMR spectrum and data [109]. 2D, Hg. Dendrobatid. Figure 11. 5,8-1. 'CigH29N'. EI: 259(3), 138(100), 96(20). FTIR spectrum [86] (Figure 22): Weak Bohlmann band 2812 cm'^; conj CH=CH 3025 cm"^; CsCH 3327 cm'^ Bufonid. Figure 21. 5,6,8-1. CigH29N. EI: 259(<1), 258(2), 152(100), 110(12), 70(8). FTIR: Weak Bohlmann band 2810 cm-^C=CH 3328 cm"^ OD. Tentatively a 5,9£-6,8dimethyl-5-octenynyl I. Dendrobatid. HTX. C17H27NO. EI: 261(8), 220(100), 204(10), 96(68). FTIR spectrum [110]: No Bohlmann band; C=CH2 3089 cm"^; conj CH=CH 3035 cm^^; OH broad band 3361 cm'^ 2D,H6. Dendrobatid. Figure 11. Izidine. 'CigHsiN'. IT-EI: 262(<1), 152(100), 110(23). Either a 5,6,8-1 or a 1,4-Q. Dendrobatid. Tricyclic. CigHsiN. EI: 261(19), 260(29), 246(8), 232(77), 220(64), 218(100), 204(12), 192(9), 190(12), 178(9), 164(14), 162(5), 150(24). FTIR: Weak Bohlmann band 2811 cm^^ C=CH2 3083 cm"^ OD. Mantellme. 5,8-1. C18H31N. IT-ET: 262(<1), 220(8), 164(13), 151(18), 138(100), 96(38). Tentatively an 8-methyl I with a 5-nonadienyl or a 5-nonynyl substituent. Dendrobatid. 5,6,8-1. C18H33N. EI: 263(2), 192(4), 152(100), 110(10), 70(6). FTIR: Weak Bohlmann band 2790 cm'^ Tentatively a 5,9£-6,8-dimethyl-5-octenyl I. Dendrobatid. Unclass. CigHssN. EI: 263(3), 198(100). OD. Dendrobatid

Alkaloids from Amphibian Skins

263C.

135

HTX. C17H29NO. EI: 263(1), 222(100), 204(10), 96(48). 2D, H4. Dendrobatid. Figure 11. 263D. 5,6,8-1. 'CigHasN'. EI: 263(3), 234(5), 180(100), 124(16), 70(15). Tentatively a 6,8-diethyl-5-hexenyl I. Dendrobatid. 263E. Unclass. 'C18H33N'. EI: 263(100), 246(72), 232(20), 204(15), 190(58), 176(84), 174(45), 158(28), 134(37), 126(25), 120(31), 91(52), 70(48). OD. Dendrobatid. 263F. 5,8-1. 'CigHssN'. EI: 263(<1), 262(<1), 164(13), 151(14), 138(100), 96(10), 70(8). Tentatively an 8-methyl-5-nonenyl I analogous to 249H. Dendrobatid. "265A". Non-alkaloidal. In early GC studies, the apparent molecular ion at m/z 265 led to assignment of an alkaloid code number to an unknown compound, even though the MS appeared atypical [49]. Later "265A" proved to be octadecenoic acid methyl ester with a true parent ion at m/z 296 and a major loss of 31 amu [1]. Fatty acid methyl esters do not afford a major protonated parent ion on NH3-CI-MS and often are trace contaminants of alkaloid fractions. 265B. CPQ. CyHsiNO. EI: 265(18), 264(22), 250(12), 236(17), 234(20), 194(16), 166(25), 126(30), 125(100), 112(28). lD,Ho. Dendrobatid. Figure 31. 265C. Unclass. C17H31NO. EI: 265(14), 236(10), 210(100, C,3H24NO), 192(10), 138(21), 84(45). ID, H2. Dendrobatid. 265D. Unclass. 'Ci7H3iNO'. EI: 265(5), 222(14), 166(100), 138(18), 124(13), 70(25). 2D. Dendrobatid. 265E. HTX. C17H31NO. EI: 265(5), 264(3), 248(10), 224(48), 222(20), 168(24), 152(95), 139(63), 96(100). FTIR spectrum [110]: No Bohlmann band; C=CH2 3080 cm"^; OH broad band 3329 cm"^ Dendrobatid. Figure 11. 265F. Unclass. C16H27NO2. EI: 265(5), 206(8), 194(100, C12H20NO), 192(30), 148(11), 134(15), 120(19). FTIR: Weak Bohlmann band 2803 cm"^; OH 3660 cm'^ H2. O-Acetyl derivative. See 245F. Mantelline. 265G. PTX. C,6H27N02. EI: 265(8), 222(6), 194(6), 166(100), 84(11), 70(45). FTIR: Strong Bohhnann band 2799 cm"^; OH 3541 cm^^; C=0 1731 cm'^ Mantelline. Figure 3. 265H. 3,5-P. C17H31NO. Cis- and tranS'265n: EI: 265(2), 264(1), 250(1), 152(100). FTIR data [56]: Weak or no Bohlmann bands; C=0 1730 cm"^ Mantelline. Figure 17. 2651. 5,6,8-1. 'C17H31NO'. IT-EI: 266(13), 265(4), 248(10), 166(100), 110(22), 70(14). Tentatively a 6-methyl-8-ethyl I with a 5-C6HnO substituent. Dendrobatid. 265J. 3,5-P. •Ci7H3iNO'. EI: 265(2), 264(1), 208(76), 166(100), 126(18). FTIR: Cis265J: Weak Bohlmann band; C=0 1731 cm"^ Trans-265J: No Bohlmann band, C=0 1731 cm'^ OD. The nature of the two substituents in cis- and trans-265J remains uncertain. Either a butyl and oxohexyl or an oxopropyl and heptyl are present. Dendrobatid. 265K. Unclass. C,6H27N02. EI: 265(14), 248(18), 236(11), 222(7), 204(7), 84(100). FTIR: Moderate, broad Bohlmann band 2805 cm"^; OH 3650, 3523 cm"^ Mantelline.

136

265L.

J. W. Daly, H. M. Garraffo and T. F. Spande

5,6,8-1. C17H31NO. EI: 265(7), 152(100), 110(9), 70(13). FTIR: Weak, broad Bohlmann band, 2790 cm"^; OH 3650 cm"^ ID. Tentatively a 5,9i&-6,8-dimethyl5-(hydroxyheptenyl) I. Mantelline. 265M. 3,5-1. CnHsiNO. EI: 265(2), 264(2), 250(4), 196(3l,Ci2H22NO), 192(100). Tentatively a 5-pentenyl I with a C4H9O substituent at C-3. Mantelline. 265N. hPTX. C17H31NO. EI: 265(8), 222(9), 220(6), 180(64), 98(23), 84(100). FTIR spectrum (Figure 8): Moderate, broad Bohlmann 2752 cm'^ OH 3553 cm'^ Dendrobatid, mantelline. Figure 7. 2650. Unclass. 'C17H31NO'. IT-EI: 264(<1), 248(3), 208(15), 192(12), 181(63), 168(7^), 166(11), 160(25), 150(25), 136(9), 122(22), 110(100), 96(23), 83(20), 75(35). Dendrobatid. 267A. aPTX. C16H29NO2. EI: 267(8), 250(23), 182(21), 114(24), 112(28), 70(100). FTIR spectrum (Figure 4): Strong, sharp Bohlmann band 2803 cm"^; OH 3649, 3522 cm-^ 1-Epi-261A.\ Strong Bohlmann band 2802 cm"^; OH 3581, 3515 cm'^ NMR data [52,60]. 2D, H2. A^-Oxide: NMR data [60]. 0-Acetyl derivative. Dendrobatid, mantelline. Figure 3. 267B. Unclass. 'C,6H29N02. EI: 267(7), 266(4), 250(1), 170(100), 152(4), 112(13). 2D. Dendrobatid. 267C. PTX. C16H29NO2. EI: 267(11), 266(7), 224(9), 222(7), 194(12), 166(100), 112(8), 84(18), 70(75). FTIR spectrum [25]: Strong, broad Bohlmann band 2798 cm'^; OH 3655, 3545 cm"^ NMR data [59]. 2D,H2. Two isomers have been detected. Dendrobatid, mantelline, bufonid, myobatrachid. Figure 3. 267D. PTX. C16H29NO2. All properties, including GC retention time, now indicate that the alkaloid reported from a mantelline frog as 267D [59] is identical to 267C. 267E. Unclass. 'CnHssNO'. EI: 267(18), 266(11), 196(100), 96(58). OD, HQ. Dendrobatid. 267F. Unclass. C16H29NO2. EI: 267(12), 266(18), 250(10), 178(100,Ci2H2oN), 126(40), 70(56). Dendrobatid. 267G. Unclass. 'Ci6H29N02'. EI: 267(4), 152(100). 2D,H2. Dendrobatid. 267H. 3,5-P. C17H33NO. EI: 267(2), 266(1), 224(48), 152(100). FTIR data [56]: Cis267H: Weak Bohlmann band; OH 3653 cm'^ ID. A minor isomer with the same ring configuration was also detected. Mantelline. Figure 17. 2671. Unclass. •C18H37N'. IT-EI: 268(17), 267(15), 198(6), 171(19), 152(13), 138(9), 111(10), 96(7), 84(14), 69(17), 58(100). FTIR: Moderate Bohlmann band 2775 cm"^. Three isomers detected. Other alkaloids with a base peak at m/z 58 are 197D, 211G, 241B, 253E, 2670 and 269E. Dendrobatid. 5,6,8-1. C,7H33NO. EI: 267(<1), 266(<1), 196(100), 124(23), 70(14). FTIR 267J. spectrum (Figure 24): Weak Bohlmann band 2805 cm"^; OH 3650 cm'^ Dendrobatid. Figure 23. 267K. Pip. C17H33NO. EI: 267(<1), 266(<1), 250(4), 210(6), 98(100). FTIR: Trans267K: No Bohlmann band; C=0 1732 cm"^ ID. The trans-\soxnQi is preceded on <JC by a minor isomer, presumably ci5-267K. Dendrobatid. Figure 37.

Alkaloids from Amphibian Skins

267L.

137

DHQ. 'C19H25N'. EI: 267(<1), 202(100). FTIR spectrum [124]: Cis-ieiU Weak Bohlmann band 2800 cm-^conjCH=CH 3035 cm-^;CsCH 3328 cm-^ ID. Two trace isomers were also detected. Dendrobatid. Figure 13. 267M. Unclass. C16H29NO2. EI: 267(9), 250(49), 224(8), 222(10), 206(21), 196(13), 178(9), 128(13), 126(19), 84(100). FTIR: Moderate, broad Bohlmami band 2805 cm-^; OH 3652, 3525 cm^^ Mantelline. 267N. DesmethylhPTX. C16H29NO2. EI: 267(3), 194(27), 166(100), 84(31). FTIR: Moderate, broad Bohlmann band 2755 cm"^; OH 3660, 3563 cm'^; strong band 1111 cm"^. Mantelline. Figure 9. 2670. Unclass. 'C17H33NO'. EI: 267(8), 238(4), 194(36), 124(33), 58(100). Other alkaloids with a base peak at m/z 58 are 197D, 211G, 241B, 253E, 2671 and 269E. Dendrobatid. 269AB. DHQ. C19H27N. EI: 269(4), 268(12), 226(6), 204(100), 202(80), 148(20). FTIR spectra [124]: Trans-269AB: Weak Bohlmann band 2803 cm"^; conj CH=CH 3036 cm"^; C ^ H 3328 cm"^; C=C=C 1952 cm'^ Cw-269AB: Weak Bohlmann band 2800 cm-^; conj CH=CH 3035 cm'h C=CH 3328 cm"^; C=C=C 1952 cm'^ NMR data [124]. ID, HIQ. A minor diastereomer, 5-ep/-rraw5-269AB, did not separate from trans-269AB on GC. The A^-acetyl derivatives separate on GC. Variable amounts of the two diastereomers of tranS'269AB occur in different extracts. At least five isomeric DHQs are included under the generic designation 269AB [124]. Dendrobatid. Figure 13. 269A. DHQ. 'C19H27N'. EI: 269(4), 226(10), 204(100), 202(10). FTIR spectrum [124]: Trans-269A: Weak Bohlmann band 2801 cm"^; conj CH=CH 3034 cm"^; CsCH 3327 cm"^;C=C=C 1952 cm"^ ID, HIQ. Dendrobatid. Figure 13. 269B. DHQ. 'C19H27N'. EI: 269(4), 226(5), 204(11), 202(100). FTIR spectrum [124]: Trans-269B: Weak Bohlmann band 2803 cm"^; conj CH=CH 3035 cm"^; CsCH 3328 cm"^ ID, HIQ. Dendrobatid. Figure 13. 269C. Pip. 'C17H35NO', EI: 269(<1), 254(3), 98(100). FTIR: Trans-269C: No Bohlmann band; OH 3653 cm"^ 2D. Dendrobatid. Figure 37. 269D. Unclass. •C19H27N'. EI: 269(4), 268(7), 228(29), 176(100), 174(17), 134(25), 120(16). FTIR: Moderate BoWmann band 2788 cm"^; C=CH2 3086 cm"^ conj CH=CH 3031 cm-^;C<:H 3328 cm-^ See245F. Dendrobatid. 269E. Unclass. 'C17H35NO'. EI: 269(4), 142(18), 100(9), 58(100). FTIR: Moderate, broad Bohlmann band 2776 cm'^; C=0 1715 cm"^ Other alkaloids with a base peak at m/z 58 are 197D, 211G, 241B, 253E, 2671 and 2670. Dendrobatid, mantelline. 271A. 5,8-1. C19H29N. EI: 271(<1), 270(2), 230(7), 228(6), 204(4), 178(100), 96(13), 70(17). FTIR: Strong, sharp Bohlmann band 2788 cm"*; C=CH2 3086 cm"*; conj CH=CH 3031 cm"^ C=CH 3328 cm'^ Tentatively a 5,8-1 with 5-C7H9 and 8-C4H7 substituents, but losses of 41 and 43 amu are puzzling. See 273B, 275F. Dendrobatid.

138

271B. 271C. 271D.

271E. 271F.

272A.

272B.

273A.

273B.

275A.

275B.

275C.

275D.

J. W. Daly, H. M. Garraffo and T. F. Spande

Unclass. C19H29N. EI: 271(7), 228(100). FTIRdata [56]:NoBohlmannband; enamineorimine 1656cm'^ OD. Dendrobatid, mantelline. Unclass. C,9H29N. EI: 271(3), 256(44), 206(100), 136(16). Dendrobatid. DHQ. 'C,9H29N'. EI: 271(6), 270(17), 228(13), 204(100). FTIR spectra [124]: Cis-lllD: Weak Bohlmann band 2804 cm"^; C=CH 3328 cm'^; C=C=C 1953 cm'^ Trans-llVD: Weak Bohlmann band 2801 cm'^ conj. CH=CH 3036 cm"^; C=CH 3327 cm''; C=C=C 1954 cm'^ Iso-S-epi-trans-llW: Weak Bohlmann band 2798 cm"'; two C=C=C 1952 cm"'. ID. Five isomeric DHQs are included under the generic designation 271D. In addition to the three of Figure 13, there is an iso-cis271D with two terminal allene side-chains and an iso-trans-llVD with a terminal allene and a terminal diene side-chain [124]. Dendrobatid. Figure 13. Unclass. 'Ci9H29N'. EI: 271(9), 162(100), 106(15), 94(8), 93(11). Dendrobatid. 3,5-1. C19H29N. EI: 271(<1), 204(100), 190(67), 126(33). FTIR: Moderate, sharp Bohlmann band 2790 cm"'; two C=CH 3328 cm"'. Tentatively a 3,5-1 with HC=C(CH2)3- and HC=C(CH2)4- substituents, but the Bohlmann band appears too strong. Mantelline. Pseudo. 'C17H24N2O'. EI: 272(50), 255(12), 199(27), 187(80), 144(100), 143(23), 130(18). ID. Tentatively an A^(8)-methyl analog of 258. Myobatrachid. Figure 40. Pseudo. 'C,7H24N20'. EI: 272(50), 255(15), 199(33), 187(54), 144(100), 143(25). 2D. A tentative structure was previously proposed [25], but the EI-MS, which is very similar to that of 272A, is not consonant with that structure. A structure fitting the data is not apparent. Myobatrachid. 5,6,8-1. C19H31N. EI: 273(2), 272(1), 234(7), 152(100), 110(8), 70(4). FTIR data [56]: Strong, sharp Bohlmann band 2788 cm"'; conj CH=CH 3030 cm"'; C=CH 3328 cm"'. A minor isomer has an unconjugated cis CH=CH 3016 cm"'. OD. Previously postulated to be a 1,4-Q [56]. Mantelline. Figure 23. 5,8-1. C,9H3iN. EI: 273(3), 272(2), 232(2), 230(5), 218(6), 204(11), 178(100), 96(15), 70(22). OD. Tentatively a 5,8-1 with 5-C7Hn and 8-C4H7 substituents, but the loss of 41 and 43 amu is puzzling. See 271 A, 275F. Dendrobatid. Azabicyclodecane. C19H33N. EI: 275(<1),274(<1), 260(3), 152(100). FTIR spectrum [297]: Very weak Bohlmann band 2805 cm"'; C=CH 3328 cm"'. OD, H4. Previously postulated to be a 1,4-Q [11]. Dendrobatid. Figure 29. DHQ. C19H33N. EI: 275(4), 274(3), 232(21), 206(100). FTIR data [86] and spectrum [124]: Cw-275B: Weak Bohlmann band 2803 cm"'; C=CH2 3086 cm"'. 2'Epi-cis-215B: No Bohlmann band, C=CH2 3086 cm"'. NMR data [124]. ID, H4. Dendrobatid, bufonid. Figure 13. 3,5-1. C19H33N. EI: 275(<1), 274(<1), 206(28), 192(100). FTIR spectrum and data [56] (Figure 20): 5Z,9Z-275C: Weak Bohlmann band 2790 cm"'; C=CH2 3086 cm"'. OD. Dendrobatid, mantelline. Figure 19. Unclass. 'C19H33N'. EI: 275(<1), 274(1), 192(47), 150(100), 148(18), 120(10). See245F. Mantelline.

Alkaloids from Amphibian Skins

275E.

275F.

275G. 277A. 277B.

277C. 277D. 277E.

279A. 279B.

279C.

279D.

279E. 279F. 279G. 281A. 281B. 281C.

139

5,6,8-L C19H33N. EI: 275(<1), 152(100), 110(6), 70(4). FTIR: Weak, broad Bohlmann band 2790 cm-^; C=CH2 3079 cm"^ A minor isomer has a C=CH 3328 cm"^ Tentatively a 5,9£-6,8-dimethyl-5-nonadienyl I and the minor isomer a 5,9£-6,8-dimethyl-5-nonynyl I. Dendrobatid. 5,8-1. 'C19H33N', EI: 275(2), 234(2), 232(6), 178(100), 96(10), 70(18). OD. Tentatively a 5,8-1 v^ith 5-C7H13 and 8-C4H7 substituents, but the loss of 41 and 43 amu is puzzling. See 271A, 273B. Dendrobatid. Azabicyclodecane. C19H33N. EI: 275(2), 274(1), 260(4), 152(100). FTIR: No Bohlmann band; C=CH2 3085 cm'^ Dendrobatid. Figure 29. Azabicyclodecane. C19H35N. EI: 277(5), 152(100). Dendrobatid. Figure 29. PTX. C17H27NO2. EI: 277(2), 206(15), 194(34), 193(48), 176(14), 166(25), 153(55), 98(28), 84(26), 70(100). FTIR: Strong, sharp Bohhnann band 2802 cm-\ shoulder 2750 cm"^; OH 3544 cm"^; C=0 1710 cm"^ ID. Dendrobatid, mantelline, myobatrachid. Figure 3. 5,6,8-1. 'C19H35N'. IT-EI: 278(5), 208(100), 124(22). Tentatively an 8-butyl-6ethyl-5-pentenyl I. Dendrobatid. Pyr. 'C19H35N'. IT-EI: 278(13), 194(68), 152(100). Dendrobatid. Figure 35. 5,6,8-1. C19H35N. EI: 277(2), 152(100), 110(11), 70(5). FTIR: Weak, broad Bohlmann band 2805 cm'^ A minor isomer has a C=CH2 3085 cm"^ Tentatively a 5,9£'-6,8-dimethyl-5-nonenyl I. Dendrobatid. Unclass. •C18H33NO'. EI: 279(35), 210(90), 190(75), 84(100). lD,Ho. Dendrobatid. CPQ. 'C18H33NO'. EI: 279(85), 278(100), 264(40), 250(45), 249(45), 248(100), 236(28), 233(40), 222(20), 220(50), 194(58), 180(35), 139(40), 117(42). ID. Dendrobatid. Figure 31. CPQ. •C18H33NO'. EI: 279(100), 278(95), 264(43), 262(64), 250(21), 243(23), 236(23), 222(22), 195(22), 180(64), 140(23), 139(33), 126(28). ID. Dendrobatid. Figure 31. 5,8-1. 'C18H33NO'. EI: 279(<1), 278(1), 164(12), 151(17), 138(100), 96(11). FTIR data [56]: Strong, sharp Bohlmann band 2786 cm"^; cis CH=CH 3013 cm"^; OH 3655 cm-^ ID. Mantelline. Figure 21. Unclass. 'C18H33NO'. EI: 279(4), 210(100), 164(21), 154(11), 150(14), 138(18), 84(45). ID. Dendrobatid. 5,6,8-1. C18H33NO. EI: 279(2), 220(4,Ci5H26N), 152(100), 110(9), 70(7). ID. Tentatively a 6,8-dimethyl-5-(hydroxyoctenyl) I. Dendrobatid. Pyr. 'C19H37N'. IT-EI: 280(28), 194(53), 154(100). Dendrobatid. Figure 35. PTX. C17H31NO2. EI: 281(4), 280(2), 264(2), 194(12), 166(72), 70(100). 2D, H2. Dendrobatid. Figure 3. DeoxyPTX. 'C17H31NO2'. EI: 281(4), 264(12), 208(25), 206(20), 150(65), 98(5), 96(20), 70(100). 2D. Dendrobatid, mantelline. Figure 5. Unclass. 'C17H31NO2'. EI: 281(25), 208(100). 2D, HQ. Dendrobatid.

140

281D.

281E.

281F.

283A.

283B. 283C. 285A.

285B.

285C.

285D. 285E.

286A.

J. W. Daly, H. M. Garraffo and T. F. Spande

Unclass. 'C17H31NO2'. EI: 281(16), 210(100). A somewhat different spectrum was obtained years later after repurification of the extract. EI: 281(<1), 210(33), 154(9), 152(7), 110(100), 70(34). 2D. Dendrobatid. Unclass. 'C17H31NO2'. EI: 281(8), 196(100). A somewhat different spectrum was obtained years later after repurification of the extract. EI: 281(6), 196(62), 178(22), 123(25), 100(100), 82(78). 2D. Dendrobatid. DihydroPTX. C17H31NO2. EI: 281(3), 264(11), 234(5), 224(7), 222(5), 196(8), 128(16), 126(23), 110(10), 84(7), 70(100). FTIR spectrum [56]: Strong, sharp Bohlmann band 2803 cm'\ shoulder 2750 cm"^; OH 3580, 3512 cm"^ Mantelline. Figure 5. Histrionicotoxin. C19H25NO. EI: 283(9), 282(5), 266(5), 218(48), 200(21), 174(14), 160(22), 132(12), 124(14), 96(100). FTIR spectrum and discussion [110]. NMR spectrum [109] and data [107,109]. 2D, H12. 0-Acetyl derivative: FTIR spectrum [110]. A 17,18-/raw5-isomer has also been detected: FTIR spectrum [110]. NMR spectrum and data [109]. PerhydroHTX: EI: 295(12), 278(13), 252(18), 224(73), 196(27), 180(100), 168(39), 96(68). FTIR spectrum [110]. NMR spectrum [107] and data [109]. Dendrobatid. Figure 11. Unclass. C17H33NO2. EI: 283(<1), 282(1), 254(2), 212(40), 152(23,CioHi8N), 140(100,C9Hi8N). ID, Ho. O-Acetyl derivative. Dendrobatid. Unclass. C17H33NO2. EI: 283(<1), 282(1), 240(5), 226(28), 224(10), 166(60,CiiH2oN), 126(100,C8Hi6N). ID, HQ. O-Acetyl derivative. Dendrobatid. Isodihydrohistrionicotoxin. C19H27NO. EI: 285(11), 284(5), 268(15), 218(13), 190(23), 176(37), 162(35), 148(18), 134(27), 122(25), 120(21), 109(40), 108(31), 96(100). FTIR spectrum and discussion [110]. NMR data [107,109]. 2D,Hio. Dendrobatid. Figure 11. Neodihydrohistrionicotoxin. C,9H27NO. EI: 285(8), 284(3), 268(12), 220(39), 202(19), 190(11), 176(14), 160(28), 132(18), 96(100). FTIR spectrum and discussion [110]. NMR data [107,109]. 2D, HIQ. Dendrobatid. Figure 11. Allodihydrohistrionicotoxin. C,9H27NO. EI: 285(14), 284(10), 268(9), 218(18), 190(13), 176(25), 162(19), 144(23), 122(25), 96(100). FTIR spectrum and discussion [110]. NMR spectrum and data [108,109]. 2D, HIQ. Dendrobatid. Figure 11. Unclass. •C19H27NO'. EI: 285(3), 270(2), 256(2), 180(35), 140(100). Dendrobatid. Dihydrohistrionicotoxin. C19H27NO. EI: 285(18), 284(8), 268(25), 218(100), 200(84), 176(13), 145(43), 96(78). FTIR spectrum and discussion [110]. NMR data [107,109]. 2D,Hio. Dendrobatid. Figure 11. Pseudo. C17H22N2O2. EI: 286(38), 199(20), 185(32), 173(100), 157(62), 156(52), 130(97). FTIR and IR data [24,25]: Moderate Bohlmann band 2800 cm"^; NH 3427 cm-^; aryl H broad 3050 cm"^; ester 1738 cm'^ NMR data [24]. ID. Myobatrachid. Figure 40.

Alkaloids from Amphibian Skins

286B.

287A.

287B.

287C.

287D.

289A.

289B. 289C.

291A.

291B. 291C. 291D. 291E.

293A.

141

Pseudo. 'CnH22N202'. EI: 286(<1), 199(100), 173(7), 156(10), 130(7), 70(50). FTIR data [25]. OD. Proposed to be a double bond isomer of 286A [25] but that is not consistent with OD. Myobatrachid. Isotetrahydrohistrionicotoxin. C19H29NO. EI: 287(12), 286(6), 270(7), 220(18), 202(23), 176(42), 162(51), 148(29), 122(30), 120(33), 109(58), 96(100). FTIR spectrum and discussion [110]. NMR data [107,109]. 2D, Hg. Dendrobatid. Figure 11. Tetrahydrohistrionicotoxin. C19H29NO. EI: 287(13), 286(5), 270(5), 220(63), 202(25), 176(15), 96(100). FTIR spectrum and discussion [110]. NMR data [107,109]. 2D,H8. Dendrobatid. Figure 11. Gephyrotoxin. C19H29NO. EI: 287(5), 286(3), 242(100), 222(45), 122(14). FTIR spectrum [11]: Very weak Bohlmann band 2800 cm'^; conj CH=CH 3032 cm"^; C=CH 3326 cm-'; OH 3666 cm"^ NMR spectrum and data [108]. 1D,H6. OAcetyl derivative. Dendrobatid. Figure 30. Allotetrahydrohistrionicotoxin. C19H29NO. EI: 287(14), 286(6), 270(16), 220(24), 202(36), 176(38), 162(49), 148(21), 134(70), 122(24), 120(38), 106(40), 96(100). NMR data [108]. 2D,H8. Dendrobatid. Figure 11. Azabicyclodecane. 'C19H31NO'. EI: 289(2), 287(2), 274(3), 152(100). OD, H4. Tentatively a keto analog of 275A with 5-methyl and 3-C9H13O substituents. Dendrobatid. Dihydrogephyrotoxin. C19H31NO. EI: 289(4), 288(3), 245(21), 244(100), 222(49), 122(20). 1D,H4. Dendrobatid. Figure 30. PTX. 'C19H31NO'. EI: 289(9), 206(42), 193(34), 166(100), 70(30). FTIR: Strong, sharp Bohlmann band 2798 cm"\ shoulder 2750 cm"^; conj CH=CH 3040 cm"^; OH 3541 cm"^ Mantelline. Figure 3. Octahydrohistrionicotoxin. C19H33NO. EI: 291(6), 290(2), 274(11), 250(54), 222(26), 194(24), 192(20), 178(100), 165(22), 136(23), 122(17), 96(65). FTIR spectrum and discussion [110]. NMR data [107]. 2D, H4. Dendrobatid. Figure 11. Unclass. 'C,9H33NO'. EI: 291(2), 290(3), 276(6), 209(4), 168(100), 114(8), 70(38). 1D,H4. Dendrobatid. Unclass. 'C,9H33NO'. EI: 291(1), 290(2), 276(4), 210(10), 152(100). 1D,H4. Dendrobatid. Unclass. C19H33NO. EI: 291 (<1), 276(5), 168(100), 114(6), 70(28). ID, H4. Probably the iV-oxide of 291B with a longer GC retention time. Dendrobatid. DeoxyPTX. C19H33NO. EI: 291(5), 246(8), 190(18), 178(20), 150(100), 136(8), 70(21). FTIR spectrum (Figure 6): Moderate Bohlmann bands 2789, 2734 cm"^; OH 3650 cm"^ ID, H4. Mantelline. Figure 5. Unclass. 'C20H39N'. EI: 293(2), 150(100). lD,Ho. Previously postulated to be a DHQ[11]. Dendrobatid.

142

293B.

293C.

293D.

293E. 293F. 293G.

295A. 295B.

297A. 297B. 300.

301. 302. 305A.

305B.

305C.

J. W. Daly, H. M. Garraffo and T. F. Spande

Unclass. 'CigHasNO'. EI: 293(<1), 292(1), 150(33), 95(35), 81(70), 67(100). FTIR data [56]: Moderate Bohlmann band 2790 cm-^; OH 3655 cm"^ ID. Mantelline. 5,6,8-1. C19H35NO. EI: 293(2), 278(3), 192(5), 152(100), 110(10), 70(9). FTIR: Weak Bohlmann band 2737 cm'^ OH 3596 cm-^ ID. Tentatively a 5,9£-6,8dimethyl-5-(hydroxynonenyl I). Dendrobatid. DeoxyPTX. C19H35NO. EI: 293(3), 292(5), 278(2), 264(4), 178(13), 150(100), 70(25). FTIR: Moderate Bohlmann bands 2788, 2750 cm'^; OH 3665 cm^^ ID. Mantelline. Figure 5. PTX. •C,8H3iN02'. EI: 293(17), 276(9), 206(27), 194(29), 193(19), 166(100), 70(38). 2D. Dendrobatid, mantelline. Azabicyclodecane. 'CIQHSSNO'. EI: 293(<1), 278(2), 152(100). OD. Probably an analog of 275A with a keto group in a saturated side-chain. Dendrobatid. Unclass. C17H27NO3. EI: 293(<1), 250(20,Ci5H24NO2), 222(35,Ci3H2oN02), 176(100,Ci2Hi8N), 148(15), 134(8), 84(44). FTIR data [56]: Moderate, broad Bohlmann band 2750 cm'^; OH 3550 cm"^; O-Acetyl 1749,1184 cm"^ Previously tabulated as 251L-(9-Ac and postulated to be an hPTX [56]. Mantelline. 5,8-1. 'C19H37NO'. EI: 295(3), 278(4), 138(100). Tentatively a 5-(hydroxydecyl)8-methylI. Dendrobatid. 5,8-1. C18H33NO2. EI: 295(<1), 278(2), 236(4,Ci5H26NO), 180(17,CHHI8NO), 154(100,C9Hi6NO), 112(20), 94(13). FTIR data [56]: Moderate, sharp Bohlmann band 2784 cm^^; cis CH=CH 3018 cm'^ OH 3655 cm•^ 0,0-Diacetyl derivative. Tentatively a ring-hydroxylated 5-(hydroxynonenyl)-8-methyl I [56]. Mantelline. aPTX. C17H31NO3. EI: 297(3), 296(4), 280(9), 182(21), 114(27), 112(16), 70(100). 3D,H2. Dendrobatid. Figure 3. PTX. 'C,7H3iN03'. EI: 297(10), 166(92), 70(100). 3D. Dendrobatid. Pseudo. 'Ci7H2oN203*. EI: 300(100), 269(23), 241(40), 240(43), 225(10), 213(20), 198(16), 187(47), 185(30), 170(19), 159(25), 106(34). OD. A trace isomer was also detected [25]. The structure of 300 was shown incorrectly in ref. 25. Myobatrachid. Figure 40. Unclass. 'C21H35N'. EI: 301(<1), 260(100). Dendrobatid. Pseudo. C17H22N2O3. EI: 302(66), 300(24), 190(35), 189(100), 173(20), 146(30). 2D. A trace isomer with ID was also detected [25]. Myobatrachid. Figure 40. aPTX. C19H31NO2. EI: 305(6), 288(2), 222(29), 209(23), 182(100), 114(33), 70(55). FTIR: Strong, sharp Bohlmann band 2801 cm"^; conj CH=CH 3031 cm"^; OH 3645, 3520 cm-^ 2D, He. Dendrobatid. PTX. C,9H3iN02. EI: 305(2), 304(3), 206(25), 193(100), 166(55), 150(15), 70(43). FTIR spectrum [56]: Strong, sharp Bohlmann band 2799 cm"^ shoulder 2750 cm"^ conj C=0 1689 cm"*; OH 3550 cm'^ Mantelline. Figure 3. aPTX. C19H31NO2. EI: 305(1), 304(2), 209(100,Ci2Hi9NO2), 182(5), 114(10), 70(30). FTIR: Strong, sharp Bohhnann band 2802 cm"*; conj CH=CH 3030 cm'*; OH 3650, 3519 cm-^ Mantelline. Figure 3.

Alkaloids from Amphibian Skins

305D.

307A.

307B.

307C. 307D.

307E. 307F.

307G.

307H.

143

PTX. C,9H3iN02. EI: 305(<1), 206(5), 193(100,C,2Hi9NO), 166(18), 150(12), 70(22). FTIR: Moderate Bohlmann bands 2798,2749 cm"*; conj CH=CH 3025 cm"*; OH 3640, 3541 cm"*. Mantelline. Figure 3. Pumiliotoxin A. C19H33NO2. EI: 307(7), 290(5), 278(4), 206(10), 194(18), 193(13), 176(10), 166(85), 84(14), 70(100). FTIR spectrum [56,57]: Strong, sharp Bohlmann band 2798 cm"*, shoulder 2750 cm"*; OH 3647, 3544 cm"*. NMR spectrum and data [51,52,105]. 2D, H4. Both C-15 epimers have been detected. The 15R-epimer (307A*), corresponding in configuration to 323A, is assumed to be the natural alkaloid, while the 15S-epimer (307A") results from facile epimerization. A 15-0-methyl derivative of 307A has been detected and given the code number 321 A. Dendrobatid, mantelline. Figure 3. PTX. C19H33NO2. EI: 307(12), 306(4), 290(2), 194(24), 193(45), 166(100), 70(56). FTIR spectrum [56]: Strong, sharp Bohlmann band 2798 cm'*, shoulder 2750 cm"*; OH 3647, 3544 cm"*. 2D, H4. Two diastereomers have been detected. Pumiliotoxins 307B and 307F cochromatograph on packed GC columns, and early studies [1] may not have always correctly identified such alkaloids. Possibly an artefact formed on allylic rearrangement of 307A [57]. Dendrobatid, mantelline. Figure 3. aPTX. C19H33NO2. EI: 307(9), 306(8), 290(4), 222(8), 210(9), 182(62), 114(28), 70(100). Dendrobatid. Figure 3. PTX. 'C19H33NO2'. EI: 307(8), 290(4), 278(3), 206(14), 194(23), 193(31), 176(10), 166(100), 84(11), 70(63). 2D, H4. Probably the TV-oxide of 307A with a longer GC retention time. Dendrobatid. PTX. 'C19H33NO2'. EI: 307(8), 166(100), 70(80). 2D. Probably identical with or an isomer of 307A. PTX. •C19H33NO2'. EI: 307(12), 194(24), 193(46), 166(100), 70(68). The m/z 194 fragment predominates in 307F*, while m/z 193 predominates in 307F". Early GC did not separate the isomers. FTIR spectra [57]: 307F*: Strong Bohlmann band 2799 cm"*, shoulder 2750 cm"*; C=0 1719 cm"*; OH 3541 cm"*. 307F": Strong Bohlmann band 2798 cm"*, shoulder 2750 cm"*; C=0 1724 cm"*; OH 3541 cm"*. NMR spectrum and data [57,105]. ID. PTX 307F' and 307F" differ with respect to the position of the keto group. PTX 307F'* (formerly 307F), the major isomer, is accompanied by a diastereomer 307F*", which probably is the C-14 epimer [57]. Dendrobatid, mantelline. Figure 3. PTX. 'C19H33NO2'. EI: 307(15), 262(23), 206(27), 194(23), 176(11), 166(100), 70(43). FTIR spectrum [56]: 307G: Strong, sharp Bohlmann band 2799 cm"*, shoulder 2750 cm"*; homoallylic OH 3650, 3610 cm"*; OH 3545 cm"*. Two diastereomers, 307G* and 307G" have been detected. Mantelline. Figure 3. PTX. 'C19H33NO2'. IT:EI: 306(13), 206(22), 193(100), 166(45), 150(25), 84(20), 70(78). FTIR spectrum [56]: No Bohlmann band; OH 3650, 3580 cm"*; enamine 1653 cm"*. Proposed to be an isomer of 307A with a 5,6-double bond [56]. Mantelline. Figure 3.

144

3071.

J. W. Daly, H. M. Garraffo and T. F. Spande

Unclass. C18H29NO3. EI: 307(<1), 252(3), 212(21,Ci2H22N02), 168(53,CHH22N), 152(100,CioHi8N). Dendrobatid. 308/310. Epib. •Ci6H2iN202Cr. EI: 310(<1), 308(<1), 207(25), 169(40), 143(5), 141(10), 140(20), 69(100). ID. Tentatively an A^-(hydroxyacyl) derivative of epibatidine. Dendrobatid. 309A. PTX. C19H35NO2. EI: 309(9), 308(3), 292(2), 194(15), 166(100), 110(10), 84(20), 70(51). FTIR spectrum [56]: Strong Bohlmann band 2798 cm•^ shoulder 2750 cm"^; OH 3655, 3544 cm"^ 2D,H2. Dendrobatid, mantelline. Figure 3. 309B. Unclass. •C20H39NO'. EI: 309(1), 152(100). Dendrobatid. 309C. PTX. 'C19H35NO2'. EI: 309(3), 308(2), 292(1), 194(15), 166(100), 70(90). 2D, H2. Probably the A^-oxide of 309A with a longer GC retention time. Dendrobatid. 309D. aPTX. C19H35NO2. EI: 309(13), 292(34), 210(14), 182(37), 114(26), 112(35), 70(100). FTIR: Moderate Bohlmann band 2802 cm"^; OH 3646, 3521 cm'^ NMR data [57]. 2D,H2. Dendrobatid. Figure 3. 309E. Unclass. 'C18H31NO3'. EI: 309(32), 266(13), 240(100), 205(22), 124(35), 114(25). 3D,H2. Dendrobatid. 309F. Unclass. •C19H35NO2'. EI: 309(2), 152(100). 2D, HQ. Dendrobatid. 316. Pseudo. 'C18H24N2O3'. EI: 316(90), 215(22), 203(100), 188(37), 174(16), 160(85), 146(18). ID. Myobatrachid. Figure 40. 317. hPTX. 'C20H31NO2'. IT-EI: 318(15), 220(10), 208(15), 207(20), 190(12), 180(100), 164(12), 148(10), 98(18), 84(72). 2D. Mantelline. 319A. hPTX. 'C20H33NO2'. IT-EI: 320(15), 276(35), 261(37), 220(34), 208(30), 190(27), 180(90), 98(20), 84(100). FTIR data [86]: Moderate, broad Bohhnann band 2757 cm"^; C=0 1728 cm'^ OH 3557 cm"^ Bufonid. Figure 7. 319B. hPTX. 'C20H33NO2'. IT-EI: 320(7), 276(12), 261(12), 180(100), 84(83). FTIR spectrum [86]: Moderate, broad Bohlmann band 2756 cm'^; conj C=0 1702,1622 cm"^; OH 3554 cm'^ ID. Bufonid. Figure 7. 321A. PTX. C20H35NO2. EI: 321(3), 304(8), 166(65), 70(100). FTIR data [56]. NMR data [105]. ID. Proposed to be a 15-O-methyl artefact formed from 307A. Dendrobatid, mantelline. 321B. hPTX. C20H35NO2. EI: 321(18), 276(13), 220(11), 208(17), 180(100), 84(47). FTIR data [86]: Moderate, broad Bohlmann band 2757 cm"^; homoallylic OH 3640, 3600 cm'^; OH 3556 cm"^ 2D. O-Acetyl derivative. Bufonid. Figure 7. 321C. aPTX. C19H31NO3. EI: 321(1), 304(10), 209(100), 192(12), 182(9), 114(32), 70(83). FTIR: Strong, sharp Bohlmann band 2803 cm"^; conj C=0 1688 cm"^; OH 3647, 3520 cm"^ Mantelline. Figure 3. 323A. Pumiliotoxin B. C,9H33N03. EI: 323(10), 306(5), 290(2), 278(12), 206(15), 194(26), 193(22), 176(15), 166(75), 84(15), 70(100). FTIR spectrum [56,57]: Strong, sharp Bohlmann band 2799 cm"^ shoulder 2750 cm"^; OH 3655, 3614, 3545 cm"^ NMR spectrum and data [49,52,105]. 3D, H4. A^-Oxide [60]. Threo323A is the only isomer detected in dendrobatid skin extracts [25]. Trace amounts of erythro-'^l^K have been detected in certain extracts of myobatrachid and

Alkaloids from Amphibian Skins

145

mantelline frog skin [25,55]. £ryr/2ro-323A: FTIR spectrum [57]. Dendrobatid, bufonid, mantelline, myobatrachid. Figure 3. 323B. aPTX. C19H33NO3. EI: 323(5), 306(10), 210(4), 209(3), 182(50), 114(20), 70(100). FTIR spectrum [57]: Strong, sharp Bohlmann band 2803 cm"^; OH 3647, 3522 cm•^ NMR spectrum and data [52,57]. 3D, H4. Both C-15 epimers have been detected. The 15R-epimer (323B*) is assumed to be the natural alkaloid, while the 15S-epimer (323B") results from facile epimerization. A 15-0-methyl artefact formed from 323B has been detected: NMR data [57]. Dendrobatid, bufonid, mantelline, myobatrachid. Figure 3. 323C. DesmethylhPTX. C19H33NO3. EI: 323(10), 278(23), 206(15), 194(42), 193(30), 166(100), 84(68). FTIR: Moderate, broad Bohlmann band 2751 cm"^; OH 3650, 3600, 3569 cm"^; strong, sharp band 1112 cm'*. Mantelline. Figure 9. 323D. Unclass. C19H33NO3. EI: 323(8), 306(11), 278(19), 266(16), 238(100,Ci3H2oN03), 222(30), 218(16), 70(60). FTIR: Moderate, sharp Bohhnann bands 2803, 2769 cm"*; OH 3552, 3516 cm"*; strong band 1094 cm"*. Mantelline. 323E. hPTX. C20H37NO2. EI: 323(7), 294(6), 280(4), 278(6), 180(100), 84(73). FTIR: Moderate, broad Bohlmann band 2753 cm"*; OH 3650, 3556 cm"*. Mantelline. Figure 7. 323F. PTX. C19H33NO3. EI: 323(13), 278(21), 260(7), 206(19), 194(27), 193(20), 166(100), 84(11), 70(57). Probably the iV-oxide of 323A with a longer GC retention time. Mantelline. 325A. aPTX. C19H35NO3. EI: 325(12), 308(22), 182(100), 114(25), 112(21), 70(73). FTIR spectrum [57]: Strong, sharp Bohlmann band 2803 cm"*; OH 3649, 3520 cm"*. NMR data [57]. 3D,H2. Both C-15 epimers have been detected. The 15Repimer (325A*) is presumed to be the natural alkaloid, while the 15S-epimer (325A") results by facile epimerization. Dendrobatid, mantelline, myobatrachid. Figure 3. 325B. PTX. C19H35NO3. EI: 325(6), 309(8), 166(85), 70(100). 3D, H2. Dendrobatid. Tentatively a 13,14-dihydro analog of 323A. 330. Pseudo. C18H22N2O4. EI (direct probe): 330(100), 302(45), 296(90), 282(32), 243(32),217(30),215(30), 199(25), 189(87). OD. Red color. Myobatrachid. Figure 40. 332. Pseudo. C18H24N2O4. EI: 332(40), 282(10), 219(100), 217(30), 189(25), 176(35), 161(15). FTIR data for two isomers [25]. 2D. Myobatrachid. Figure 40. 335. hPTX. C21H37NO2. EI: 335(23), 318(11), 292(14), 220(28), 207(34), 206(39), 180(100), 84(72). Mantelline. Figure 7. 337. hPTX. C21H39NO2. EI: 337(9), 294(11), 208(6), 180(100), 84(63). Mantelline. Figure 7. 339A. aPTX. C19H33NO4. EI: 339(3), 322(3), 192(14), 182(75), 114(25), 70(100). FTIR: Strong, sharp Bohlmann band 2803 cm"*; OH 3600 (broad), 3524 cm"*. NMR spectrum and data [52]. 4D,H4. Dendrobatid. Figure 3.

146

339B.

J. W. Daly, H. M. Garraffo and T. F. Spande

aPTX. C19H33NO4. EI: 339(3), 322(3), 192(10), 182(70), 114(25), 70(100). NMR spectrum and data [52]. 4D, H4. An atypical aPTX with cw-hydroxyl groups in the ring. Dendrobatid. Figure 3. 339C. Unclass. 'C19H33NO4'. EI: 339(8), 282(25), 264(5), 212(4), 210(12), 166(35), 110(43), 84(100), 70(31). Mantelline. 341A. aPTX. C19H35NO4. EI: 341(4), 324(3), 323(2), 298(12), 266(11), 254(13), 182(10), 126(15), 114(20), 112(75), 84(38), 70(100). FTIR spectrum [58]: Strong, Bohlmann band 2817 cm"^; OH 3535 (broad) cm"^; strong ether band 1070 cm'^ NMR spectrum and data [58]. 3D. Dendrobatid. Figure 3. 341B. aPTX. 'C,9H35N04'. EI: 341(1), 324(4), 182(60), 114(20), 112(20), 70(100). Tentatively an isomer of 341 A. Dendrobatid. 341C. Unclass. •C20H39NO3'. EI: 341(<1), 268(100), 222(14), 164(16), 110(12), 96(9), 70(11). Mantelline. 346A. Pseudo. 'C19H26N2O4'. EI: 346(60), 233(100), 218(22), 190(60), 175(25). ID. Myobatrachid. Figure 40. 346B. Chimonanthine. C22H26N4. EI: 346(<1), 173(70), 172(100). Dendrobatid. Figure 41. 346C. Calycanthine. C22H26N4. EI: 346(100). Dendrobatid. Figure 41. 351. Unclass. 'C21H37NO3'. EI: 351(6), 350(2), 336(4), 152(38), 138(65), 70(100). 4D. Dendrobatid. 353A. PTX. •C2iH39N03'. EI: 353(4), 338(10), 336(5), 194(20), 166(80), 70(100). 3D. Dendrobatid. 353B. Unclass. •C21H39NO3'. EI: 353(<1), 266(100), 178(13), 148(4), 124(8), 98(3), 96(5), 70(11). Dendrobatid. 357. aPTX. C19H35NO5. EI: 357(3), 339(15), 282(11), 254(16), 182(20), 112(83), 84(35), 70(100). FTIR: Strong, sharp Bohlmann band 2817 cm"^; OH broad 3574 cm"^; strong ether band 1070 cm'^ 4D. Hydroxy-congener of 341 A. Dendrobatid. 382. DHQ-dimer. 'C26H42N2'. IT-EI: 383(38), 339(100), 194(8), 192(7), 136(19), 108(7). Mantelline. 384A/B. DHQ-dimer. C26H44N2. EI: 384(5), 341(100), 190(13), 150(15), 136(10). FTIR spectrum [164]: No Bohlmann band; cis CH=CH 3020 cm"^; enamine 1647 cm'\ NMR spectrum and data [164]. 2D. Two isomers, 384A and 384B, occur together and have virtually identical MS and FTIR spectra [56]. Reversion to "monomers" appears to occur to some extent on GC-MS analysis. Mantelline. Figure 15. 392. Unclass. C22H36N2O4. EI: 392(2), 310(20), 294(14), 278(23), 252(100), 234(13), 222(43), 220(31), 178(13), 164(15), 162(11), 136(28), 126(14), 110(19), 98(39), 84(27), 82(21), 68(32). FTIR: No Bohlmann band; OH 3585 cm"^ enamine or imine 1643 cm'^; strong band 1063 cm"^ Mantelline. 434. Unclass. C24H38N2O5. EI: 434(<1), 352(18), 320(20), 294(100), 276(23), 234(53), 222(25), 220(25), 136(39), 110(33), 98(51), 84(65), 83(51), 82(47), 68(85). FTIR:

Alkaloids from Amphibian Skins

512.

524. 528.

147

No Bohlmann band; 0-acetyl 1761, 1229 cm'^; enamine or imine 1643 cm"^ Mantelline. Pseudophrynamine A. C32H40N4O2. EI (direct probe): 512(56), 456(23), 455(13), 340(55), 338(100), 273(20), 241(40), 211(17), 199-197(20-23), 185-182(22-25), 173(80), 172(60), 144(20), 138(38). IR data [24]. NMRdata[24]. 2D. Myobatrachid. Figure 40. Pseudo. C33H40N4O2. EI (direct probe): Present in a mixture; fragments not assigned. OD. Tentatively an A^(8)-methyldehydro analog of 512. Myobatrachid. Pseudo. C32H40N4O3. EI (direct probe): 528(10), 472(5), 356(15), 354(25), 173(100), 130(95). 3D. Myobatrachid. Figure 40.

REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22.

JW Daly, CW Myers, and N Whittaker, Toxicon 25:1023 (1987). CL Bevins and M Zasloff, Ann Rev Biochem 59:395 (1990). T Yasumoto, D Yasumura, M Yotsu, T Michishita, A Endo, and Y Kotaki, Agric Biol Chem 50:793 (1986). JW Daly, WL Padgett, RL Saunders, and JF Cover Jr, Toxicon 35:705 (1997). JW Daly, SI Secunda, HM Garraffo, TF Spande, A Wisnieski, C Nishihira, and JF Cover Jr, Toxicon 30:887 (1992). JW Daly, HM Garraffo, TF Spande, C Jaramillo, and AS Rand, J Chem Ecol 20:943 (1994). JW Daly, SI Secunda, HM Garraffo, TF Spande, A Wisnieski, and JF Cover Jr, Toxicon 32:657 (1994). JW Daly, HM Garraffo, GSE Hall and JF Cover Jr, Toxicon 35:1131 (1997). JW Daly, in: Progress in the Chemistry of Organic Natural Products, Vol 41, W Herz, H Grisebach and GW Kirby, Eds., Springer-Verlag, Vienna, 1982, pp. 205340. JW Daly and TF Spande, in: Alkaloids: Chemical and Biological Perspectives, Vol 4, SW Pelletier, Ed., John Wiley and Sons, New York, 1986, pp. 1-274. JW Daly, HM Garraffo, and TF Spande, in: The Alkaloids, Vol. 43, GA Cordell, Ed., Academic Press, San Diego, 1993, pp. 185-288. B Witkop and E Gossinger, in: The Alkaloids, Vol. 21, A Brossi, Ed., Academic Press, New York, 1983, pp. 139-253. CI Schopf, Experientia 17:285 (1961). G Habermehl, in: The Alkaloids, Vol. 9, RHF Manske, Ed., Academic Press, New York, 1967, pp. 427-439. G Habermehl, in: Venomous Animals and their Venoms, Vol. 2, W Bucherl and EE Buckley, Eds., Academic Press, New York, 1971, pp. 569-584. S Hara and K Oka, J Am Chem Soc 89:1041 (1967). K Oka and S Hara, Tetrahedron Lett 1193 (1969). G Habermehl and A Haaf, Chem Ber 98:3001 (1965). K Oka and S Hara, J Am Chem Soc 99:3859 (1977); see also K Oka, Y Ike, and S Hara, Tetrahedron Lett 4543 (1968). K Oka and S Hara Tetrahedron Lett 1189 (1969). G Habermehl, Tetrahedron Lett 3 815 (1969). K Oka and S Hara, Tetrahedron Lett 1987 (1969).

148

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. 48a. 49. 50. 51. 52. 53. 54. 55. 56. 57. 58.

J. W. Daly, H. M. Garraffo and T. F. Spande

G Haberaiehl, Zeit Naturforsch 20B:1129 (1965). TF Spande, MW Edwards, LK Pannell, and JW Daly, J Org Chem 53:1222 (1988). JW Daly, HM Garraffo, LK Pannell, and TF Spande, J Nat Prod 53:407 (1990). F Marki and B Witkop, Experientia 19:329 (1963). CW Myers, JW Daly, and B Malkin, Bull Am Mus Nat Hist 161:307 (1978). T Tokuyama, JW Daly, B Witkop, IL Karle, and J Karle, J Am Chem Soc 90:1917 (1968). IL Karle, Proc Natl Acad Sci USA 69:2932 (1972). JW Daly, B Witkop, P Bommer, and K Biemann, J Am Chem Soc 87:124 (1965). T Tokuyama, JW Daly, and B Witkop, J Am Chem Soc 91:3931 (1969). T Tokuyama and JW Daly, Tetrahedron 39:41 (1983). R Imhoff, E Gossinger, W Graf, L Bemer-Fenz, H Bemer, R Schaufelberger, and H Wehrli, Helv Chim Acta 56:139 (1973). BI Khodorov, LD Zaborvskaya, EA Elin, MZ Maksudov, OB Tikhomirova, and VN Leonov, Biol Membranes 5:475 (1988). P Hudson, G Pairaudeau, PJ Parsons, AW Jahans, and MGB Drew, Tetrahedron Lett 34:7295 (1993). TJ Grinsteiner and Y Kishi, Tetrahedron Lett 35:8333 (1994). TJ Grinsteiner and Y Kishi, Tetrahedron Lett 35:8337 (1994). JW Daly, CW Myers, JE Wamick, and EX Albuquerque, Science 208:1383 (1980). M Neuwirth, JW Daly, CW Myers, and LW Tice, Tissue Cell 11:755 (1979). EX Albuquerque, JE Wamick, FM Sansone, and JW Daly, J Pharmacol Exp Therap 184:315(1973). JP Dumbacher, BM Beehler, TF Spande, HM Garraffo, and JW Daly, Science 258:799(1992). J Dumbacher and S Pruett-Jones, Current Omithol 13:137 (1996). EX Albuquerque, JW Daly, and B Witkop, Science 172:955 (1971). GB Brown, SC Tieszen, JW Daly, JE Wamick, and EX Albuquerque, Cell Mol Neurobiol 1:19(1981). S Kongsamut, SB Freedman, BE Simon, and RJ Miller, Life Sci 36:1493 (1985). CR Creveling and JW Daly, in: Methods in Neuroscience, Vol. 8, PM Conn, Ed., 1992, pp. 25-37. TL Casebolt and GB Brown, Toxicon 31:1113 (1993). VL Trainer, GB Brown, and WA Catterall, J Biol Chem 271:11261 (1996). S-Y Wang and GW Wang, Proc Natl Acad Sci USA 95:2653 (1998). JW Daly, GB Brown, M Mensah-Dwumah, and CW Myers, Toxicon 16:163 (1978). JW Daly and CW Myers, Science 156:970 (1967). JW Daly, T Tokuyama, T Fujiwara, RJ Highet and IL Karle, J Am Chem Soc 102:830 (1980). T Tokuyama, JW Daly, and RJ Highet, Tetrahedron 40:1183 (1984). T Tokuyama, K Shimada, M Uemura, and JW Daly, Tetrahedron Lett 23:2121 (1982). M Uemura, K Shimada, T Tokuyama, and JW Daly, Tetrahedron Lett 23:4369 (1982). HM Garraffo, MW Edwards, TF Spande, JW Daly, LE Overman, C Severini, and V Erspamer, Tetrahedron 44:6795 (1988). HM Garraffo, J Caceres, JW Daly, TF Spande, NR Andriamaharavo, and M Andriantsiferana, JNat Prod 56:1016 (1993). T Tokuyama, T Tsujita, HM Garraffo, TF Spande, and JW Daly, Tetrahedron 47:5415(1991). P Jain, TF Spande, HM Garraffo, and JW Daly, Heterocycles, submitted (1998).

Alkaloids from Amphibian Skins

59. 60. 61. 62. 63. 64. 65. 66. 67. 68. 69. 70. 71. 72. 73. 74. 75. 76. 77. 78. 79. 80. 81. 82. 83. 84. 85. 86. 87. 88. 89. 90. 91. 92. 93

149

JW Daly, RJ Highet, and CW Myers, Toxicon 22:905 (1984). MW Edwards, JW Daly, and CW Myers, J Nat Prod 51:1188(1988). JW Daly, NR Andriamaharavo, M Andriantsiferana, and CW Myers, Am Mus Novitates No. 3177, pp. 1-34 (1996). H Takahata and T Momose, in: The Alkaloids, Vol. 43, GA Cordell, Ed., Academic Press, San Diego, 1994, pp. 189-256. AS Franklin and LE Overman, Chem Rev 96:505 (1996). LE Overman and KL Bell, J Am Chem Soc 103:1851 (1981). LE Overman and N-H Lin, J Org Chem 50:3669 (1985). LE Overman, KL Bell, and F Ito, J Am Chem Soc 106:4192 (1984). LE Overman and SW Goldstein, J Am Chem Soc 106:5360 (1984). DNA Fox, D Lathbury, MF Mahon, KC Molloy, and T Gallagher, J Am Chem Soc 113:2652(1991). T Honda, M Hoshi, K Kanai, and M Tsubuki, J Chem Soc Perkin Trans I 2091 (1994). S Aoyagi, T-C Wang, and C Kibayashi J Am Chem Soc 115:11393 (1993). BM Trost and TS Scanlan, J Am Chem Soc 111:4988 (1989). RM Lett, LE Overman, and J Zablocki, Tetrahedron Lett 29:6541 (1988). SW Goldstein, LE Overman, and MH Rabinovs^itz, J Org Chem 57:1179 (1992). LE Overman and MJ Sharp, Tetrahedron Lett 29:901 (1988). N-H Lin, LE Overman, MH Rabinowitz, LA Robinson, MJ Sharp, and J Zablocki, J Am Chem Soc 118:9062 (1996). LE Overman, LA Robinson, and J Zablocki, J Am Chem Soc 114:368 (1992). C Caderas, R Lett, LE Overman, MH Rabinowitz, LA Robinson, MJ Sharp, and J Zablocki, J Am Chem Soc 118:9073 (1996). T Honda, M Hoshi, and M Tsubuki, Heterocycles 34:1515 (1992). J Cossy, M Cases, and D Gomez Pardo, Synlett 909 (1996). J Cossy, M Cases, and D Gomez-Pardo, Bull Soc Chim Fr 134:141 (1997). S Aoyagi, Y Hasegawa, S. Hirashima, and C Kibayashi, Tetrahedron Lett 39:2149 (1998). S Aoyagi, T-C Wang, and C Kibayashi, J Am Chem Soc 114:10653 (1992). JW Daly, F Gusovsky, ET McNeal, S Secunda, M Bell, CR Creveling, Y Nishizawa, LE Overman, MJ Sharp, and DP Rossignol, Biochem Pharmacol 40:315 (1990). JW Daly, ET McNeal, LE Overman, and DH Ellison, J Med Chem 28:482 (1985). TM Barger, RM Lett, PL Johnson, JE Hunter, CP Chang, DJ Pemich, MR Sabol, and MR Dick, J Agric Food Chem 43:1044 (1995). HM Garraffo, TF Spande, JW Daly, A Baldessari, and EG Gros, J Nat Prod 56:357 (1993). M Mensah-Dwumah and JW Daly, Toxicon 16:189 (1978). EX Albuquerque, JE Warnick, MA Maleque, FC Kauffman, F Tamburini, Y Nimit, and JW Daly, Mol Pharmacol 19:411 (1981). EX Albuquerque, JE Warnick, FC Kauffman, and JW Daly, in: Membranes and Transport, A Martonosi, Ed., Vol. 2, Plenum Publishing Corp, New York, 1982, pp. 335-365. KS Rao, JE Warnick, JW Daly, and EX Albuquerque, J Pharmacol Exp Therap 243:775 (1987). GG Schofield, FF Weight, and SR Ikeda, Eur J Pharmacol 147:39 (1988). RE Sheridan, SS Deshpande, FJ Lebeda and M Adler, Brain Res 556:53 (1991). R Tamburini, EX Albuquerque, JW Daly, and FC Kauffman, J Neurochem 37:775 (1981).

150

94.

J. W. Daly, H. M. Garraffo and T. F. Spande

PM Sokolove, EX Albuquerque, FC Kauffman, TF Spande, and JW Daly, FEES Lett 203:121 (1986). 95. F Gusovsky, DP Rossignol, ET McNeal, and JW Daly, Proc Natl Acad Sci USA 85:1272(1988). 96. F Gusovsky, EB Hollingsworth, and JW Daly, Proc Natl Acad Sci USA 83:3003 (1986). 97. F Gusovsky, ET McNeal, and JW Daly, Mol Pharmacol 32:479 (1987). 98. F Gusovsky and JW Daly, Neuropharmacology 27:95 (1988). 99. JW Daly, ET McNeal, and F Gusovsky, Biochim Biophys Acta 930:470 (1987). 100. JW Daly, ET McNeal, F Gusovsky, F Ito, and LE Overman, J Med Chem 31:477 (1988). 101. GT Bolger, MF Rafferty and P Skolnick, Pharmacol Biochem Behav 24:417 (1986). 102. GF Erspamer, C Severini, V Erspamer, and P Melchiorri, Neuropharmacology 28:319(1989). 103. F Gusovsky, WL Padgett, CR Creveling, and JW Daly, Mol Pharmacol 42:1104 (1992). 104. P Jain, HM Garraffo, TF Spande, HJC Yeh, and JW Daly, J Nat Prod 58:100 (1995). 105. T Tokuyama, N Nishimori, A Shimada, MW Edwards, and JW Daly, Tetrahedron 43:643 (1987). 106. JW Daly, I Karle, CW Myers, T Tokuyama, JA Waters, and B Witkop, Proc Natl Acad Sci USA 68:1870 (1971). 107. T Tokuyama, K Uenoyama, G Brown, JW Daly, and B Witkop, Helv Chim Acta 57:2597 (1974). 108. JW Daly, B Witkop, T Tokuyama, T Nishikawa, and IL Karle, Helv Chim Acta 60:1128(1977). 109. T Tokuyama, J Yamamoto, JW Daly, and RJ Highet, Tetrahedron 39:49 (1983). 110. TF Spande, HM Garraffo, JW Daly, T Tokuyama, and A Shimada, Tetrahedron 48:1823(1992). 111. EJ Corey and RD Balanson, Heterocycles 5:445 (1976). 112. Y Inubushi and T Ibuka, Heterocycles 17:507 (1982). 113. K Takahashi, B Witkop, A Brossi, MA Maleque, and EX Albuquerque, Helv Chim Acta 65:252 (1982). 114. K Takahashi, AE Jacobson, C-P Mak, B Witkop, A Brossi, EX Albuquerque, JE Wamick, MA Maleque, A Bavoso, and JW Silverton, J Med Chem 25:919 (1982). 115. W Gessner, K Takahashi, B Witkop, and A Brossi, Helv Chim Acta 68:49 (1985). 116. D Tanner and P Somfai, Tetrahedron 42:5657 (1986). 117. RS Aronstam, CT King Jr, EX Albuquerque, JW Daly, and DM Feigl, Biochem Pharmacol 34:3037 (1985). 118. T Fukuyama, LV Dunkerton, M Aratani, and Y Kishi, J Org Chem 40:2011 (1975). 119. SC Carey, M Aratani, and Y Kishi, Tetrahedron Lett 26:5887 (1985). 120. JJ Venit, M Dipierro, and P Magnus, J Org Chem 54:4298 (1989). 121. JD Winkler, PM Hershberger and JP Sprmger, Tetrahedron Lett 27:5177 (1986). 122. G Stork and K Zhao, J Am Chem Soc 112:5875 (1990). 123. CW Myers and JW Daly, Bull Am Mus Nat Hist 157:173 (1976). 124. TF Spande, P Jain, HM Garraffo, LK Pannell, HJC Yeh, JW Daly, S Fukumoto, K Imamura, T Tokuyama, JA Torres, RR Snelling, and TH Jones, J Nat Prod submitted (1998). 125. TH Jones, JST Gorman, RR Snelling, JHC Delabie, MS Blum, HM Garraffo, P Jain, JW Daly, and TF Spande, J Chem Ecol submitted (1998). 126. EX Albuquerque, EA Bamard, TM Chiu, AJ Lapa, JO Dolly, S-E Jansson, JW Daly, and B Witkop, Proc Natl Acad Sci USA 70:949 (1973).

Alkaloids from Amphibian Skins

127. 128. 129. 130. 131. 132. 133. 134. 135. 136. 137. 138. 139. 140. 141. 142. 143. 144. 145. 146. 147. 148. 149. 150. 151. 152. 153. 154. 155.

151

AJ Lapa, EX Albuquerque, JM Sarvey, JW Daly, and B Witkop, Exp Neurol 47:558 (1975). CE Spivak, MA Maleque, AC Oliveira, LM Masukawa, T Tokuyama, JW Daly, and EX Albuquerque, Mol Pharmacol 21:351 (1982). T Ogura and A Warashima, Comp Biochem Physiol 88C:249 (1987). B Badio, D Shi, Y Shin, KD Hutchinson, WL Padgett, and JW Daly, Biochem Pharmacol 52:933 (1996). M Klepper, M Hans, and K Takeda, J Physiol (London) 428:545 (1990). A Wada, Y Uezono, M Arita, K Tsuji, N Yanagihara, K Kobayashi, and F Izumi, Neuroscience 33:333 (1989). JW Daly, Y Nishizawa, MW Edwards, JA Waters, RS Aronstam, Neurochem Res 16:489(1991). DB Sattelle and JA David, Neurosci Lett 43:37 (1983). C Rapier, S Wonnacott, GG Lunt, and EX Albuquerque, FEBS Lett 212:292 (1987). G Kato and J-P Changeux, Mol Pharmacol 12:92 (1976). T Lovenberg and JW Daly, Neurochem Res 11:1609 (1986). AT Eldefrawi, ME Eldefrawi, EX Albuquerque, AC Oliveira, N Mansour, M Adler, JW Daly, GB Brown, W Burgermeister, and B Witkop, Proc Natl Acad Sci USA 74:2172(1977). T Heideman, RE Oswald, and J-P Changeux, Biochemistry 22:3112 (1983). ND Boyd and JB Cohen, Biochemistry 23:4023 (1984). HR Arias, CF Valenzuela, and DA Johnson, J Biol Chem 268:6348 (1993). DA Johnson and JM Nuss, Biochemistty 33:9070 (1994). HR Arias and DA Johnson, Biochemistry 34:1589 (1995). W Oberthiir, P Muhn, H Baumann, F Lottspeich, B Wittmann-Liebold, and F Hucho, EMBO J 5:1815 (1986). KL Swanson, Y Aracava, FJ Sardina, H Rapoport, RS Aronstam, and EX Albuquerque, Mol Pharmacol 35:223 (1989). ACS Costa, KL Swanson, Y Aracava, RS Aronstam, and EX Albuquerque, J Pharmacol Exp Therap 252:507 (1990). P Kofiigi, Y Aracava, KL Swanson, RS Aronstam, H Rapoport, and EX Albuquerque, J Pharmacol Exp Therap 252:517 (1990). KL Swanson, RS Aronstam, S Wonnacott, H Rapoport, and EX Albuquerque, J Pharmacol Exp Therap 259:377 (1991). AS Ramoa, M Alkondon, Y Aracava, J Irons, GG Lunt, SS Deshpande, S Wonnacott, RS Aronstam, and EX Albuquerque, J Pharmacol Exp Therap 254:71 (1990). R Rozental, GT Scoble, EX Albuquerque, M Idriss, S Sherby, DB Sattelle, K Nakanishi, K Konno, AT Eldefrawi, and ME Eldefrawi, J Pharmacol Exp Therap 249:123 (1989). SM Sherby, AT Eldefrawi, SS Deshpande, EX Albuquerque, and ME Eldefrawi, Pesticide Biochem Physiol 26:107 (1986). Y Nishizawa, KB Seamon, JW Daly, and RS Aronstam, Cell Mol Neurobiol 10:351 (1990). JE Wamick, PJ Jessup, LE Overman, ME Eldefrawi, Y Nimit, JW Daly, and EX Albuquerque, Mol Pharmacol 22:565 (1982). JW Daly, Y Nishizawa, WL Padgett, T Tokuyama, PJ McCloskey, L Waykole, AG Schultz, and RS Aronstam, Neurochem Res 16:1207 (1991). RS Aronstam, JW Daly, TF Spande, TK Narayanan, and EX Albuquerque, Neurochem Res 11:1227 (1986).

152

156. 157. 158. 159. 160. 161. 162. 163. 164. 165. 166. 167. 168. 169. 170. 171. 172. 173. 174. 175. 176. 177. 178. 179. 180. 181. 182. 183. 184. 185. 186. 187. 188. 189. 190. 191. 192. 193. 194.

J. W. Daly, H. M. Garraffo and T. F. Spande

JW Daly, Y Nishizawa, WL Padgett, T Tokuyama, AL Smith, AB Holmes, C Kibayashi, and RS Aronstam, Neurochem Res 16:1213 (1991). C Souccar, WA Varanda, RS Aronstam, JW Daly, and EX Albuquerque, Mol Pharmacol 25:395 (1984). RS Aronstam, MW Edwards, JW Daly, and EX Albuquerque, Neurochem Res 13:171(1988). JW Daly, T Tokuyama, G Habermehl, IL Karle, and B Witkop, Justus Liebigs Ann Chem 729:198 (1969). HM Garraffo, LD Simon, JW Daly, TF Spande, and TH Jones, Tetrahedron 50:11329(1994), T Tokuyama, T Tsujita, A Shimada, HM Garraffo, TF Spande, and JW Daly, Tetrahedron 47:5401 (1991). T Tokuyama, N Nishimori, IL Karle, MW Edwards, and JW Daly, Tetrahedron 42:3453 (1986). HM Garraffo, TF Spande, TH Jones, and JW Daly, Tetrahedron in preparation (1998). TF Spande, HM Garraffo, P Jain, LK Pannell, HJC Yeh, and JW Daly, J Nat Prod in preparation (1998). Y Inubushi and T Ibuka, Heterocycles 8:633 (1977). H-P Husson, J Nat Prod 48:894 (1985). M Bonin, J Royer, DS Grierson, and HP Husson, Tetrahedron Lett 27:1569 (1986). AG Schultz, J Patrick, and JJ Court, J Am Chem Soc 109:6493 (1987). NA LeBel and N Balasubramanian, J Am Chem Soc 111:3363 (1989). DL Comins and A Dehghani, Tetrahedron Lett 32:5697 (1991). A Brandi, FM Cordero, A Goti, and A Guema, Tetrahedron Lett 33:6697 (1992). RP Polniaszek and LW Dillard, J Org Chem 57:4103 (1992). S Murahashi, S Sasao, E Saito, and T Naota, J Org Chem 57:2521 (1992). DL Comins and A Dehghani, J Chem Soc Chem Commun 1838 (1993). S Murahashi, S Sasao, E Saito, and T Naota, Tetrahedron 49:8805 (1993). AI Meyers and G Milot, J Am Chem Soc 115:6652 (1993). M Naruse, S Aoyagi, and C Kibayashi, Tetrahedron Lett 35:9213 (1994). G Mehta and M Praveen, J Org Chem 60:279 (1995). M Toyota, T Asoh, M Matsuura, and K Fukumoto, J Org Chem 61:8687 (1996). SG Davies and G Bhalay, Tetrahedron: Asymmetry 7:1595 (1996). M Toyota, T Asoh, and K Fukumoto, Tetrahedron Lett 37:4401 (1996). M Naruse, S Aoyagi, K Sakae, and C Kibayashi, J Chem Soc Perkin Trans 11113 (1996). M Toyota, T Asoh, and K Fukumoto, Heterocycles 45:147 (1997). JT Kuethe and A Padwan, Tetrahedron Lett 38:1505 (1997). K Paulvannan and JR Stille, Tetrahedron Lett 34:6673 (1993). PA Grieco and DT Parker, J Org Chem 53:3658 (1988). PJ McCloskey and AG Schultz, J Org Chem 53:1380 (1988). DL Comins and A Dehghani, J Org Chem 60:794 (1995). M Bonin, R Bellelievre, DS Grierson, and H-P Husson, Tetrahedron Lett 24:1493 (1983). G Habermehl, H Andres, K Miyahara, B Witkop, and JW Daly, Justus Liebigs Ann Chem 1577 (1976). K Abe, T Tsugoshi, and N Nakamura, Bull Chem Soc Japan 57:3351 (1984). LE Overman and PJ Jessup, J Am Chem Soc 100:5179 (1978). LE Overman and T Yokomatsu, J Org Chem 45:5229 (1980). TH Jones, MS Blum, HM Tales, and CR Thompson, J Org Chem 45:4778 (1980).

Alkaloids from Amphibian Skins

195. 196. 197. 198.

TH Jones, RJ Highet, AW Don, and MS Blum, J Org Chem 51:2712(1986). TH Jones, SM Stahly, AW Don, and MS Blum, J Chem Ecol 14:2197 (1988). TH Jones, A Laddago, AW Don, and MS Blum, J Nat Prod 53:375 (1990). TH Jones, MS Blum, HM Fales, CRF Bandao, and J Lattke, J Chem Ecol 17:1897 (1991). 199. PE Sonnet, DA Netzel, and R Mendoza, J Heterocyclic Chem 16:1041 (1979). 200. S Arseniyadis, PQ Huang, and HP Husson, Tetrahedron Lett 29:1301 (1988). 201. H Takahata, H Bandoh, and T Momose, Tetrahedron: Asymmetry 2:351 (1991). 202. H Takahata, H Bandoh, and T Momose, J Org Chem 57:4401 (1992). 203. O Provot, JP Celerier, H Petit, and G Lhommet, J Org Chem 57:2163 (1992). 204. C Grandjean, S Rosset, JP Celerier, and G Lhommet, Tetrahedron Lett 34:4517 (1993). 205. GD Cuny and SL Buchwald, Synlett 519 (1975). 206. W Oppolzer, CG Bochet, and E Merifield, Tetrahedron Lett 35:7015 (1994). 207. HM Garraffo, P Jain, TF Spande, and JW Daly, J Nat Prod 60:2 (1997). 208. TF Spande, JW Daly, DJ Hart, Y-M Tsai, and TL MacDonald, Experientia 37:1242 (1981). 209. JW Daly, TF Spande, N Whittaker, RJ Highet, D Feigl, N Nishimori, T Tokuyama, and CW Myers, J Nat Prod 49:265 (1986). 210. FJ Ritter, lEM Rogans, E Talman, PEJ Verwiel, and F Stein, Experientia 29:530 (1973). 211. J Royer, and H-P Husson, Tetrahedron Lett 26:1515 (1985). 212. N Yamazaki and C Kibayashi, J Am Chem Soc 111:1396 (1989). 213. N Machinaga and C Kibayashi, J Org Chem 57:5178(1992). 214. DJ Hart and Y-M Tsai, J Org Chem 47:4403 (1982). 215. TL MacDonald, J Org Chem 45:193 (1980). 216. M Przybylska and RF Ahmed, Acta Crystallogr 042:832 (1986). 217. N Machinaga and C Kibayashi, J Chem Soc Chem Commun 405 (1991). 218. JE Oliver and PE Sonnet, J Org Chem 39:2662 (1974). 219. PE Sonnet and JE Oliver, J Heterocyclic Chem 12:289 (1975). 220. FJ Ritter and CJ Persons, Netherland J Zool 25:261 (1975). 221. JP Edwards and DB Pinniger, Ann Appl Biol 89:395 (1978). 222. J Royer and HP Husson, J Org Chem 50:670 (1985). 223. N Yamazaki and C Kibayashi, Tetrahedron Lett 29:5767 (1988). 224. RV Stevens and AWM Lee, J Chem Soc Chem Commun 102 (1982). 225. H lida, Y Watanabe, and C Kibayashi, Tetrahedron Lett 27:5513 (1986). 226. R Yamaguchi, E Hata, T Matsuki, and M Kawanishi, J Org Chem 52:2094 (1987). 227. Y Watanabe, H lida, and C Kibayashi, J Org Chem 54:4088 (1989). 228. CW Jefford, Q Tang, and A Zaslona, Helv Chim Acta 72:1749 (1989). 229. T Nagasaka, H Kato, H Hayashi, M Shioda, H Hikasa, and F Hamaguchi, Heterocycles 30:561 (1990). 230. E Zeller and DS Grierson, Synlett 878 (1991). 231. M Vavrecka and M Hesse, Helv Chim Acta 74:438 (1991). 232. PL McGrane and T Livinghouse, J Org Chem 57:1323 (1992). 233. AM Castafio, JM Cuerva, and AM Echavarren, Tetrahedron Lett 35:7435 (1994). 234. TT Shawe, CJ Sheils, SM Gray, and JL Conard, J Org Chem 59:5841 (1994). 235. A Fleurant, C Saliou, JP Celerier, N Platzer, Thuy Vu Moc, and G Lhommet, J Heterocyclic Chem 32:255 (1995). 236. M Ito and C Kibayashi, Tetrahedron Lett 31:5065 (1990). 237. M Ito and C Kibayashi, Tetrahedron 47:9329 (1991). 238 T Momose, N Toyooka, S Seki, and Y Hirai, Chem Pharm Bull 38:2072 (1990).

153

154

239. 240. 241. 242. 243. 244. 245. 246. 247. 248. 249. 250. 251. 252. 253. 254. 255. 256. 257. 258. 259. 260. 261. 262. 263. 264. 265. 266. 267. 268. 269. 270. 271. 272. 273. 273a. 274. 275. 276. 277. 278. 279.

J. W. Daly, H. M. Garraffo and T. F. Spande

C Saliou, A Fleurant, JP Celerier, and G Lhommet, Tetrahedron Lett 32:3365 (1991). CW Jefford, Q Tang, and A Zaslona, J Am Chem Soc 113:3513 (1991). H Takahata, H Bandoh, and T Momose, Tetrahedron 49:11205 (1993). SR Angle and JG Breitenbucher, Tetrahedron Lett 34:3985 (1993). K Higashiyama, KNakahata, and H Takashi, J Chem Soc Perkin Trans 1 351 (1994). CW Jefford, K Sienkiewicz and SR Thorton, Tetrahedron Lett 35:4759 (1994). MJ Munchhof and AI Meyers, J Am Chem Soc 117:5399 (1995). G Solladie and G-H Chu, Tetrahedron Lett 37:111 (1996). O Muraoka, B-Z Zheng, K Okumura, E Tabata, G Tanabe, and M Kubo, J Chem Soc Perkin Trans I 113(1997). T Momose, M Toshima, S Seki, Y Koike, N Toyooka, and Y Hirai, J Chem Soc Perkin Trans 11315 (1997). DR Artis, I-S Cho, S Jaime-Figueroa, and JM Muchowski, J Org Chem 59:2456 (1994). TH Jones, RJ Highet, MS Blum, and HM Fales, J Chem Ecol 10:1233 (1984). H Takahata, H Bandoh, and T Momose, Heterocycles 42:39 (1996). W Francke, F Schroder, F Walter, V Sinnwell, H Baumann, and M Kaib, Justus Liebigs Ann Chem 965 (1995). RV Stevens and AWM Lee, J Chem Soc Chem Commun 103 (1982). H lida, Y Watanabe, and C Kibayashi, J Am Chem Soc 107:5534 (1985). CA Broka and KK Eng, J Org Chem 51:5045 (1986). OE Edwards, AM Greaves, and WW Sy, Can J Chem 66:1163 (1988). A Brandi, F Cordero, and C Querci, J Org Chem 54:1748 (1989). FM Cordero, A Brandi, C Querci, A Goti, F DeSarlo, and A Guama, J Org Chem 55:1762(1990). TK Yang, ST Yeh, and YY Lay, Heterocycles 38:1711 (1994). Y Nakagawa and RV Stevens, J Org Chem 53:1871 (1988). DF Taber, PB Deker, and LJ Silverberg, J Org Chem 57:5990 (1992). A Fleurant, JP Celerier, and G Lhommet, Tetrahedron: Asymmetry 4:1429 (1993). O Muraoka, K Okumura, T Maeda, G Tanabe, and T Momose, Tetrahedron: Asymmetry 5:317 (1994). RA Pilli, LC Dias, and AO Maldaner, J Org Chem 60:717 (1995). E Zeller and DS Grierson, Heterocycles 27:1575 (1988). H Takahata, H Bandoh, and T Momose, Heterocycles 36:2777 (1993). R Bloch, C Brillet-Femandez, P Kuhn, and G Mandville, Heterocycles 30:1589 (1994). RP Polniaszek and SE Belmont, J Org Chem 55:4688 (1990). E Zeller, H Sajus, and DS Grierson, Synlett 44 (1991). A Fleurant, JP Celerier, and G Lhommet, Tetrahedron: Asymmetry 3:695 (1992). CW Jefford and JB Wang, Tetrahedron Lett 34:3119 (1993). CW Jefford, K Sienkiewicz, and SR Thornton, Helv Chim Acta 78:1511 (1995). S Seiji, M Sodeoka, H Sasai, and M Shibasaki, J Org Chem 60:398 (1995). SR Angle and RM Henry, J Org Chem 62:8549 (1997). M Weymann, W Pfrengle, D Schollmeyer, and H Kunz, Synthesis 1151(1997). CA Toft, Oecologia (Berlin) 45:131 (1980). MA Donnelly, Copeia 723 (1991). CA Toft, Herpetologia 51:202 (1995). JP Caldwell, J Zool (London) 240:75 (1996). TH Jones, JA Torres, TF Spande, HM Garraffo, MS Blum, and RR Snelling, J Chem Ecol 22:1221 (1996).

Alkaloids from Amphibian Skins

280.

155

JST Gorman, TH Jones, TF Spande, RR Snelling, JA Torres, and HM Garraffo, J ChemEcol 24:933 (1998). 281. N Toyooka, K Tanaka, T Momose, JW Daly, and HM Garraffo, Tetrahedron 53:9553 (1997). 282. RP Polniaszek and SE Belmont, J Org Chem 56:4868 (1991). 283. Y Shishido and C Kibayashi, J Org Chem 57:2876 (1992). 284. DL Comins, DH LaMunyon, and XH Chen, J Org Chem 62:8182 (1997). 285. AL Smith, SF Williams, AB Hoknes, LR Hughes, Z Lidert, and C Swithenbank, J Am Chem Soc 110:8696 (1988). 286. AB Holmes, AL Smith, SF WiUiams, LR Hughes, Z Lidert, and C Swithenbank, J Org Chem 56:1393 (1991). 287. DL Comins and E Zeller, Tetrahedron Lett 32:5889 (1991). 288. I Collins, ME Fox, AB Holmes, SF Williams, R Baker, IJ Forbes, and M Thompson, J Chem Soc Perkin Trans 1175 (1991). 289. T Momose and N Toyooka, J Org Chem 59:943 (1994). 290. DF Taber, M Rahaminizadeh, and KK You, J Org Chem 60:529 (1995). 291. D Gnecco, C Marazano, and BC Das, J Chem Soc Chem Commun 625 (1991). 292. A Satake and I Shimizu, Tetrahedron: Asymmetry 4:1405 (1993). 293. J Aube, PS Rafferty, and GL Milligan, Heterocycles 35:1141 (1993). 294. J Aehman and P Somfai, Tetrahedron 51:9747 (1995). 295. JP Michael and D Gravestock, Synlett 981 (1996). 295a. T Tokuyama, A Shimada, HM Garraffo, TF Spande, and JW Daly, Heterocycles, submitted (1998). 296. P Jain, HM Garraffo, HJC Yeh, TF Spande, JW Daly, NR Andriamaharavo, and M Andriantsiferana, J Nat Prod 59:1174 (1996). 297. HM Garraffo, TF Spande, JW Daly, JST Gorman, and TH Jones, Proc Nat Acad Sci USA in preparation (1998). 298. R Fujimoto and Y Kishi, Tetrahedron Lett 22:4197 (1981). 299. MW Edwards and A Bax, J Am Chem Soc 108:918 (1986). 300. T Ibuka and GN Chu, Chem Pharm Bull 34:2380 (1986). 301. WH Pearson and Y-F Poon, Tetrahedron Lett 30:6661 (1989). 302. AB Holmes, AB Hughes, AL Smith, and SF Williams, J Chem Soc Perkin Trans I 1089(1992). 303. AB Holmes, AB Hughes and AL Smith, J Chem Soc Perkin Trans I 633 (1992). 304. C Souccar, WA Varanda, JW Daly, and EX Albuquerque, Mol Pharmacol 25:384 (1984). 305. CW Myers and JW Daly, Am Mus Novitates No. 2692, pp.1-23 (1980). 306. TF Spande, HM Garraffo, HJC Yeh, Q-L Pu, LK Pannell, and JW Daly, J Nat Prod 55:707(1992). 307. DF Taber and KK You, J Am Chem Soc 117:5757 (1995). 308. T Tokuyama, HM Garraffo, TF Spande, and JW Daly, Anal Asoc Quim Arg, in press (1998). 309. R Karlsson and D Losman, J Chem Soc Chem Commun 626 (1972). 310. WA Ayer and K Furuichi, Can J Chem 54:1494 (1976). 311. RV Stevens and AWM Lee, J Am Chem Soc 101:7032 (1979). 312. RH Mueller, ME Thompson, and RM DiPardo, J Org Chem 49:2217 (1984). 313. C Yue, J-F Nicolay, J Royer, and H-P Husson, Tetrahedron 50:3139 (1994). 314. JM Pasteels, C Deroe, B Tursch, JC Braekman, D Daloze and C Hootele, J Insect Physiol 19:1771 (1973). 315. B Tursch, D Daloze, JC Braekman, C Hootele, and JM Pasteels, Tetrahedron 31:1541(1975).

156

316. 317.

J. W. Daly, H. M. Garraffo and T. F. Spande

B Lebrun, JC Braekman, D Daloze, and JM Pasteels, J Nat Prod 60:1148 (1987). PA Hedin, RC Gueldner, RD Henson, and AC Thompson, J Insect Physiol 20:2135 (1974). 318. BP Moore and WV Brown, Insect Biochem 8:393 (1978). 319. M Timmerman, J-C Braekman, D Daloze, JM Pasteels, J Merlin, and J-P Declercq, Tetrahedron Lett 33:1281 (1992). 320. KD McCormick, AB Attygalle, S-C Cheng, A Svatos, J Meinwald, MA Houck, and T Eisner, Tetrahedron 50:2365 (1994). 321. X Shi, AB Attygalle, J Meinwald, MA Houck, and T Eisner, Tetrahedron 51:8711 (1995). 322. F Schroder, S Francke, W Francke, H Baumann, M Kaib, JM Pasteels, and D Daloze, Tetrahedron 52:13539 (1996). 323. F. Schroder, V Sinnwell, H Baumann, M Kaib, and W Francke, Angew Chem Int Ed 36:77(1997). 324. T Tokuyama, JW Daly, HM Garraffo, and TF Spande, Tetrahedron 48:4247 (1992). 325. J Meinwald, J Smolanoff, AT McPhail, RW Miller, T Eisner, and K Hicks, Tetrahedron Lett, 2367 (1975). 326. RW Miller and AT McPhail, J Chem Res 2:76 (1978). 327. KD Hutchinson, JV Silverton, and JW Daly, Tetrahedron 50:6129(1994). 328. J Smolanoff, AF Kluge, J Meinwald, A McPhail, RW Miller, K Hicks, and T Eisner, Science 188:734 (1975). 329. T Sugahara, Y Komatsu, and S Takano, J Chem Soc Chem Commun 214: (1984). 330. N Machinaga and C Kibayashi, J Org Chem 56:1386 (1991). 331. TH Jones, MS Blum, and HM Fales, Tetrahedron Lett 1031 (1979). 332. W Gessner, K Takahashi, A Brossi, M Kowalski, and MA Kaliner, Helv Chim Acta 70:2003 (1987). 333. H Takahata, H Takehara, N Ohkubo, and T Momose, Tetrahedron: Asymmetry 1:561 (1990). 334. J-E Backvall, HE Schink, and ZD Renko, J Org Chem 55:826 (1990). 335. K Shiosaki and H Rapoport, J Org Chem 50:1229 (1985). 336. TH Jones, MS Blum, P Escoubas, and TMM Ali, J Nat Prod 52:779 (1989). 337. TH Jones, MS Blum, RW Howard, CA McDaniel, HM Fales, MB Dubois, and J Torres, J Chem Ecol 8:285 (1982). 338. TH Jones, MS Blum, and HM Fales, Tetrahedron 38:1949 (1982). 339. TH Jones, MS Blum, AN Anderson, HM Fales, and P Escoubas, J Chem Ecol. 14:35 (1988). 340. D Bacos, JJ Basselier, JP Celerier, C Lange, E Marx, G Lhommet, P Escoubas, M Lemaire, and JL Clement, Tetrahedron Lett 29:3061 (1988). 341. TH Jones, PJ DeVries, and P Escoubas, J Chem Ecol 17:2507 (1991). 342. DJ Pedder, HM Fales, T Jaouni, M Blum, and J MacConnell, Tetrahedron 32:2275 (1976). 343. TH Jones and MS Blum in: Alkaloids: Chemical and Biological Perspectives, Vol. 1, SW Pelletier, Ed., John Wiley and Sons, New York, 1983, pp. 33-84. 344. A Numata and T Ibuka in: The Alkaloids, Vol. 31, A Brossi, Ed., Academic Press, New York, 1987, pp. 193-315. 345. MS Blum, J Toxicol Toxin Rev 11:115 (1992). 346. R Chenevert and M Dickman, J Org Chem 61:3332(1995). 347. MW Edwards, HM Garraffo, and JW Daly, Synthesis 1167 (1994). 348. DF Taber, PB Deker, HM Fales, TH Jones, and HA Lloyd, J Org Chem 53:2968 (1988).

Alkaloids from Amphibian Skins

349. 350. 351. 352. 353. 354. 355. 356. 357. 358. 359. 360. 361. 362. 363. 364. 365. 366. 367. 368. 369. 370. 371. 372. 373. 374. 375. 376. 377. 378. 379. 380. 381. 382. 382a. 383. 384.

157

S Leclercq, I Thirionet, F Breeders, D Daloze, R Vandermeer, and JC Braekman, Tetrahedron 50:8465 (1994). DS Grierson, J Royer, L Guerrier, and HP Husson, J Org Chem 51:4475 (1986). H Kotsuki, T Kusumi, M Inoue, Y Ushio, and M Ochi, Tetrahedron Lett 32:4159 (1991). CW Jefford and JB Wang, Tetrahedron Lett 34:2911 (1993). JZ Yeh, T Narahashi, and RR Almon, J Pharmacol Exp Therap 194:373 (1975). TF Spande, HM Garraffo, MW Edwards, HJC Yeh, L Pannell, and JW Daly, J Am Chem Soc 114:3475(1992). B Badio, HM Garraffo, TF Spande, and JW Daly, Med Chem Res 4:440 (1994). AP Watt, HM Verrier, and D O'Connor, J Liquid Chromat 17:1257 (1994). SR Fletcher, R Baker, MS Chambers, RH Herbert, SC Hobbs, SR Thomas, HM Verrier, AP Watt, and RG Ball; J Org Chem 59:1771 (1994). SR Fletcher, R Baker, MS Chambers, SC Hobbs, and PJ Mitchell, J Chem Soc Chem Commun 1216 (1993). EJ Corey, T-K Loh, S AchyuthaRao, DC Daley, and S Sarshar, J Org Chem 58:5600 (1993). CA Broka, Tetrahedron Lett 34:3251 (1993). DF Huang and TY Shen, Tetrahedron Lett 34:4477 (1993). SC Clayton and AC Regan, Tetrahedron Lett 34:7493 (1993). C Szantay, Z Kardos-Balogh, I Moldvai, C Szantay Jr, E Temesvari-Major and G Blasko, Tetrahedron Lett 35:3171 (1994). K Sestanj, E Melenski, and I Jirkovsky, Tetrahedron Lett 35:5417 (1994). G Pandey, TD Bagul, and G Lakshmaiah, Tetrahedron Lett 35:7439 (1994). E Albertini, A Barco, S Benetti, C De Risi, GP Pollini, R Romagnoli, and V Zanirato, Tetrahedron Lett 35:9297 (1994). K Senokuchi, H Nakai, M Kawamura, N Katsube, S Nonaka, H Sawaragi, and N Hamanaka, Synlett 343 (1994). K Okabe and M Natsume, Chem Pharm Bull 42:1432 (1994). SY Ko, J Lerpinieve, ID Linney, and R Wrigglesworth, J Chem Soc Chem Commun 1775(1994). CA Broka, Med Chem Res 4:449 (1994). C Szantay, Z Kardos-Balogh and C Szantay Jr in: The Alkaloids, vol. 46, GA Cordell, Ed., Academic Press, San Diego 1995, pp. 95-125. CE Miiller, Pharm Unserer Zeit 25:85 (1996). EV Dehmlow, J Prakt Chem 337:167 (1995). PL Kotian and FI Carroll, Synth Commun 25:63 (1995). A Hernandez, M Marcos, and H Rapoport, J Org Chem 60:2683 (1995). C Szantay, Z Kardos-Balogh, I Moldvai, C Szantay Jr, E Temesvari-Major, and G Blasko, Tetrahedron 52:11053 (1996). BM Trost and GR Cook, Tetrahedron Lett 37:7485 (1996). C Zhang and ML Trudell, J Org Chem 61:7189 (1996). R Xu, G Chu and D Bai, Tetrahedron Lett 37:1463 (1996). D Bai, R Xu, G Chu, and X Zhu, J Org Chem, 61:4600 (1996). JA Campbell and H Rapoport, J Org Chem 61:6313 (1996). NP Pavri and ML Trudell, Tetrahedron Lett 38:7993 (1997). M Ikeda, Y Kugo, Y Kondo, T Yamasaki, and T Sato, J Chem Soc Perkin Trans I 3339 (1997). HF Olivo, MS Hemenway, and MH Gezginei, Tetrahedron Lett 39:1309 (1998). A Horti, HT Ravert, ED London, and RF Dannals, J Labelled Compounds Radiopharm 38:355 (1996).

158

385. 386. 387. 388. 389. 390. 391. 392. 393. 394. 395. 396. 397. 398. 399. 400. 401. 402. 403. 404. 405. 406. 407. 408. 409. 410. 411. 412. 413.

J. W. Daly, H. M. Garraffo and T. F. Spande

F Liang, HA Navarro, P Abraham, P Kotian, Y-S Ding, J Fowler, N Volkow, MJ Kuhar, and FI Carroll, J Med Chem 40:2293 (1997). MI Davila-Garcia, JL Musachio, DC Perry, Y Xiao, A Horti, ED London, RF Dannals, and KJ Kellar, J Pharmacol Exp Therap 282:445 (1997). B Radio, HM Garraffo, CV Plummer, WL Padgett, and JW Daly, Eur J Pharmacol 321:189(1997). RWM Aben, J Keijsers, B Hams, CO Kruse, and HW Sheeren, Tetrahedron Lett 35:1299(1994). K Hiroya, K Uwai, and K Ogasawara, Chem Pharm Bull 43:901 (1995). J Gonzalez, JI Koontz, LM Hodges, KR Nilsson, LK Neeley, WH Myers, H Sabat, and WD Harman, J Am Chem Soc 117:3405 (1995). A Shafi'ee and G Hite, J Org Chem 33:3435 (1968). RR Fraser and RB Swingle, Can J Chem 48:2065 (1970). A Hassner and AM Belostotskii, Tetrahedron Lett 36:1709 (1995). C Zhang, L Gyermek, and ML Trudell, Tetrahedron Lett 38:5619 (1997). R Xu, G Chu, J Tao, and X Zhu, Bioorg Med Chem Lett 6:279 (1996). JR Malpass, DA Hemmings, and AL Walhs, Tetrahedron Lett 37:3911 (1996). B Sundermann and HD Scharf, Synlett 703 (1996). B Badio and JW Daly, Mol Pharmacol 45:563 (1994). MD Dukat, MI Damaj, W Glassco, D Dumas, EL May, BR Martin, and RA Glennon, Med Chem Res 4:131(1994). C Qian, T Li, TY Shen, L Libertine-Garahan, J Eckman, T Biftu, and S Ip, Eur J Pharmacol 250:R13 (1993). MI Damaj, KR Creasy, AD Grove, JA Rosecrans, and BR Martin, Brain Res 664:34 (1994). NMJ Rupniak, S Patel, R Marwood, J Webb, JR Traynor, J Elliott, SB Freedman, SR Fletcher, and RG Hill, Brit J Pharmacol 113:1487 (1994). TS Rao, LD Correa, RT Reid, and GK Lloyd, Neuropharmacology 35:393 (1996). DW Bouhaus, KR Bley, CA Broka, DJ Fontana, E Leung, R Lewis, A Shieh, and EHF Wong, J Pharmacol Exp Therap 272:1199 (1995). JP Sullivan, MW Decker, JD Brioni, D Donnelly-Roberts, DJ Anderson, AW Bannon, C-H Kang, P Adams, M Piattoni-Kaplan, MJ Buckley, M Gopalakrishnan, M. Williams, and SP Americ, J Pharmacol Exp Therap 271:624 (1994). DJ Anderson, M Williams, JR Pauly, JL Raszkiewicz, JE Campbell, G Rotert, B Surbor, SB Thomas, J Wasicak, SP Americ, and JP Sullivan, J Pharmacol Exp Therap 273:1434 (1995). JP Sullivan, CA Briggs, D Donnelly-Roberts, JD Brioni, RJ Radek, DG McKenna, JE Campbell, SP Americ, MW Decker, and AW Bannon, Med Chem Res 4:502 (1994). V Gerzanich, X Peng, F Wang, G Wells, R Anand, S Fletcher, and J Lindstrom, Mol Pharmacol 48:774 (1995). RL Papke, JS Thinschmidt, BA Moulton, EM Meyer, and A Poirier, Brit J Pharmacol 120:429(1997). B Badio, D Shi, HM Garraffo, and JW Daly, Dmg Dev Res 36:46 (1995). M. Gopalakrishnan, LM Monteggia, DJ Anderson, EJ Molinari, M Piattoni-Kaplan, D Donnelly-Roberts, SP Americ, and JP Sullivan, J Pharmacol Exp Therap 276:289 (1996). M Gopalakrishnan, B Buisson, E Tuoma, T Giordano, JE Campbell, IC Hu, D Donnelly-Roberts, SP Americ, D Bertrand, and JP Sullivan, Eur J Pharmacol 290:237(1995). RH Loring, T McHugh, J McKay, and XG Zhang, Med Chem Res 4:517 (1994).

Alkaloids from Amphibian Skins

414. 415. 416. 417.

159

U Warpman and A Norberg, Neuroreport 6:2419 (1995). M-Y Liu, B Latli, and JE Casida, Pesticide Biochem Physiol 52:170 (1995). MJ Marks, SF Robinson, and AC Collins, J Pharmacol Exp Therap 277:1383 (1996). WR Kern, VM Mahnir, RL Papke, and CJ Lingle, J Pharmacol Exp Therap 283:979 (1997). 418. O Delbono, M Gopalakrishnan, M Renganathan, LM Monteggia, ML Messi, and JP Sullivan, J Pharmacol Exp Therap 280:428 (1997). 419. JC Cooper, O Gutbrod, V Witzemann, and C Methfessel, Eur J Pharmacol 309:287 (1996). 420. CA Briggs, DG McKenna, and M Piattoni-Kaplan, Neuropharmacology 34:583 (1995). 421. M Fisher, D Huangfu, TY Shen, and PG Guyonet, J Pharmacol Exp Therap 270:702 (1994). 422. M Alkondon and EX Albuquerque, J Pharmacol Exp Therap 274:771 (1995). 423. EX Albuquerque, EFR Pereira, NG Castro, M Alkondon, S Reinhardt, H Schroder, and A Maelicke, Ann New York Acad Sci 757:48 (1995). 424. T Li, C Qian, J Eckman, DF Huang, and TY Shen, Bioorg Med Chem Lett 3:2759 (1993). 425. MI Damaj, SM Tucker, MD Aceto, and BR Martin, Med Chem Res 4:483 (1994). 426. MI Damaj and BR Martin, Eur J Pharmacol 300:51 (1996). 427. AW Bannon, KL Gunther, and MW Decker, Pharmacol Biochem Behav 51:693 (1995). 428. IM Khan, TL Yaksh, and P Taylor, Brain Res 753:269 (1997). 429. B Badio, WL Padgett, and JW Daly, Mol Pharmacol 51:1-5 (1997). 430. AW Bannon, KL Gunther, MW Decker, and SP Americ, Brain Res 678:244 (1995). 431. F Menzaghi, KT Whelan, VB Risbrough, TS Rao, and GK Lloyd, J Pharmacol Exp Res 280:384 (1997). 432. AI Sacaan, F Menzaghi, JL Dunlop, LD Correa, KT Whelan, and GK Lloyd, J Pharmacol Exp Therap 276:509 (1996). 433. JL Wiley, JR James, and JA Rosecrans, Drug Dev Res 38:222 (1996). 434. ED Levin, K Toll, G Chang, NC Christopher, SJ Briggs, and W Fiedler, Med Chem Res 6:543 (1996). 435. C Qian, S Chen, L Libertine-Garahan, DM Wypij, P Yao, F-M Zuo, and J Eckman, Med Chem Res 4:493 (1994). 436. AI Sacaan, RT Reid, EM Santori, P Adams, LD Correa, LS Mahaffy, L Bleicher, NDP Cosford, KA Stauderman, lA McDonald, TS Rao, and GK Lloyd, J Pharmacol Exp Therap 280:373 (1997). 437. K Sershen, A Balla, A Lajtha, and ES Vizi, Neuroscience 77:121 (1997). 438. TD White and K Semba, Neurosci Lett 235:125 (1997). 439. GI Wilkie, P Hutson, JP Sullivan, and S Wonnacott, Neurochem Res 21:1141 (1996). 440. IM Khan, M Marsala, MP Printz, P Taylor, and TL Yaksh, J Pharmacol Exp Therap 278:97 (1996). 441. B. Lendvai, H Sershen, A Lajtha, E Santha, M Baranyi, and ES Vizi, Neuropharmacology 35:769 (1996). 442. TS Rao, LD Correa, and GK Lloyd, Neuropharmacology 36:39 (1997). 443. RA Glennon, JL Hemdon, and M Dukat, Med Chem Res 4:461 (1994). 444. MI Damaj, W Glassco, M Dukat, EL May, RA Glennon, and BR Martin, Drug Dev Res 38:177 (1996). 445. MA Abreo, N-H Lin, DS Garvey, DE Gunn, A-M Hettinger, JT Wasicak, PA Pavlik, YC Martin, DL Donnelly-Roberts, DJ Anderson, JP Sullivan, M Williams, SP Americ, and MW Holladay, J Med Chem 39:817 (1996).

160

446. 447. 448. 449. 450. 451. 452. 453. 454. 455. 456. 457. 458. 459. 460. 461. 462. 463. 464. 465. 466. 467. 468. 469. 470. 471. 472. 473. 474. 475. 476. 477. 478. 479.

J. W. Daly, H. M. Garraffo and T. F. Spande

AW Bannon, MW Decker, MW Holladay, P Curzon, D Donnelly-Roberts, PS Puttfarken, RS Bitner, A Diaz, AH Dickenson, RD Porselt, M Williams, and SP Americ, Science 279:77 (1998). W Brandt and A Barth, SAR QSAR Environ Res 1:345 (1993). RA Houghtling, MI Davila-Garcia, SD Hurt, and KJ Kellar, Med Chem Res 4:538 (1994). RA Houghtling, MI Davila-Garcia, and KJ Kellar, Mol Pharmacol 48:280 (1995). DC Perry and KJ Kellar, J Pharmacol Exp Therap 275:1030 (1995). ED London, U Scheffel, AS Kimes, and KJ Kellar, Eur J Pharmacol 278: Rl (1995). CM Flores, MI Davila-Garcia, YM Ulrich, and KJ Kellar, J Neurochem 69:2216 (1997). J McKay, J Lindstrom, and RH Loring, Med Chem Res 4:528 (1994). IM Khan, KL Youngblood, MP Printz, TL Yaksh, and P Taylor, Hypertension 28:1093(1996). CM Flores, RM DeCamp, S Kilo, SW Rogers, and KM Hargreaves, J Neurosci 16:7892(1996). F Wang, V Gerzanich, GB Wells, R Anand, X Peng, K Keyser, and J Lindstrom, J Biol Chem271:17656(1996). U Scheffel, GT Taylor, JA Kepler, FI Carroll, and MJ Kuhar, Neuroreport 6:2483 (1995). Y-S Dmg, SJ Gatley, JS Fowler, ND Volkov^, D Aggarwal, J Logan, SL Dewey, F Liang, FI Carroll, and MJ Kuhar, Synapse 24:403 (1996). VL Villemagne, A Horti, U Scheffel, HT Ravert, P Finley, DJ Clough, ED London, HN Wagner Jr, and RF Dannals, J Nucl Med 38:1737 (1997). X Liu, JL Musachio, HN Wagner Jr, T Mochizuki, RF Dannals, and ED London, Synapse 27:378 (1997). E Wright, T Gallagher, CGV Sharpies, and S Wonnacott, Bioorg Med Chem Lett 7:2867(1997). JP Sullivan and AW Bannon, CNS Drug Rev 2:21 (1996). E Spath and L Mamoli, Berichte 69:1082 (1936). AM Duffield, H Budzikiewicz, and C Djerassi, J Am Chem Soc 87:2926 (1965). AS Sadykov, SZ Mukhamedzhanov, and KHA Aslanov, Dokl Akad Nauk Uzb SSR 24:34 (1967) (Chem Abst 68:78473e, 1988, see also Chem Abst 89:126093p, 1978). JW Wheeler, O Olubajo, CB Storm, and RM Duffield, Science 211:1051 (1981). WR Kem, KM Scott, and JH Duncan, Experientia 32:684 (1976). M Roseghini, V Erspamer, and R Endean, Comp Biochem Physiol C: Comp Pharmacol 54C:31 (1976). WY Sun, Y Sun, YC Tang, and JQ Hu, Synlett 337 (1993). D Crich, AB Pavlovic, and R Samy, Tetrahedron Lett 51:6379 (1995). K Fuji, T Kawabata, T Ohmuri, and M Node, Synlett 367 (1995). MO Mitchell and PW LeQuesne, Tetrahedron Lett 31:2681 (1990). PG Cozzi, C Palazzi, D Potenza, C Scolastico, and WY Sun, Tetrahedron Lett 31:5661(1990). MO Mitchell and P Dorroh, Tetrahedron Lett 32:7641 (1991). B Badio, HM Garraffo, WL Padgett, NH Greig, and JW Daly, Biochem Pharmacol 53:671 (1997). K Oka, JD Kantrowitz, and S Spector, Proc Natl Acad Sci USA 82:1852 (1985). GA Cordell, in: The Alkaloids, vol. 20, RHF Manske and RGA Rodrigo, Eds., Academic Press, New York, 1981, pp. 1-296. ES Hall, F McCapra, and AI Scott, Tetrahedron 23:4131 (1967). CW Myers and JW Daly, Science 262:1193 (1993).

Alkaloids from Amphibian Skins

480. 481. 482. 483.

CW Myers, JW Daly, and V Martinez, Am Mus Novit No. 2783 pp.1-20 (1984). CW Myers, JW Daly, HM Garraffo, A Wisnieski, and JF Cover Jr, Am Mus Novit No. 3144 pp.1-21 (1995). CW Myers and JW Daly, Am Mus Novit No. 2674 pp. 1-24 (1979). CW Myers and JW Daly, Occ Papers Mus Nat Hist, Univ Kansas No. 59 pp. 1 -12 (1976).

161

This Page Intentionally Left Blank

Chapter Two

Naturally Occurring Cyclotryptophans and Cyclotryptamines Uffe Anthoni, Carsten Christophersen and Per Halfdan Nielsen Marine Chemistry Section Department of Chemistry, University of Copenhagen Universitetsparken 5, DK-2100, Copenhagen Denmark

CONTENTS 1. INTRODUCTION

165

2. BACTERIA Streptomyces

166 168

3. CYANOBACTERIA Oscillatoria Microcystis

168 168 168

4. FUNGI Penicillium Amauroascus Chaetomium Acrostalagmus Gliocladium Verticillium Pithomyces Corollospora Nannizzia Aspergillus Leptosphaeria

170 170 173 174 176 177 111 111 179 179 180 18 5 163

164

U. Anthoni, C. Christophersen and P. H. Nielsen

5. HIGHER PLANTS 186 188 5.1 LilmcQSiQ Allium 5.2 Leguminosae, Physostigma 188 5.3 Calycanthaceae, Calycanthus, Chimonanthus 189 5.4 Idiospermaceae, Idiospermum 191 5.5 Rutaceae, Flindersia 191 5.6 Rubiaceae, Hedyotis, Borreria, Hodgkinsonia, Calycodendron and Psychotria 192 5.7 Apocynaceae 195 Hunteria 195 Rauwolfia 199 Vinca (Catharanthus) 200 Alstonia 201 Pe/c/z/a 204 Cabucala 204 Tabernaemontana 206 Rhazya, Aspidosperma, Gonioma 206 5.8 Loganiaceae 207 Strychnos 207 6. ANIMALS 6.1 Porifera Phakellia 6.2 Bryozoa Flustra Securiflustra 6.3 Ascidiacea Ciona, Botrylloides 6.4 Amphibia Pseudophryne 6.5 Mammalia

207 207 208 208 209 212 212 212 213 213 215

M^AZ ««<^ Rat

215

7. BIOSYNTHESIS

215

8. CYCLIZATIONS IN PEPTIDES AND PROTEINS? 8.1 Potential role of cyclotryptophans in senescence and aging 8.2 Acetylcholine esterase

220 221 222

8.3 Conceivable regulatory functions

223

9. CONCLUSION

225

REFERENCES

226

Naturally Occurring Cyclotryptophans and Cyclotryptamines

165

1. INTRODUCTION The term cyclotryptophans is proposed in this review to describe alkaloids containing the l,2,3,3a,8,8a-hexahydropyrrolo[2,3-Z?]indolering system. The cyclotryptophans are formally derived from tryptophan by cyclisation through addition of the A^b-amino group to the double bond of the pyrrole ring, i.e. are cyclic tautomers of tryptophan:

Cxj^ H

COOH

'^'^N'^N^COOH

H2

Tryptophan

Cyclotryptophan

When discussing the occurrence of alkaloids it is practical to distinguish cyclotryptamines as those cyclotryptophans which are formally derived from tryptamine:

Tryptamine

Cyclotryptamine

Although relatively rare among natural products these alkaloids exhibit wide systematic occurrence, being known from such diverse sources as bacteria, fungi, higher plants, marine bryozoans, frogs, and human metabolism. Many derivatives are formed by simple substitution in the pyrroloindole ring system by alkyl, aryl, acyl, OR, NR2, halogen, etc. or they contain an additional ring condensed across the 3a,8a-junction. A considerable number of alkaloids are derived from dimeric, trimeric and polymers of cyclotryptamine or cyclotryptophan. The carboxylic group in cyclotryptophans allows for formation of cyclic peptides ranging from simple 6-rings to large depsipeptides. Several treatises deal with limited groups of compounds encompassing the 1,2,3,3a,8,8ahexahydropyrrolo[2,3-^]indole ring system. Thus, the occurrence of bisindole alkaloids have been reviewed by Cordell and Saxton in 1981 [1] and the monoterpenoid derivatives by Sapi and Massiot in 1994 [2]. A list of fungal metabolites were given by Turner in 1971 [3] and updated by Turner and Aldridge in 1983 [4]. In spite of the wide systematic distribution a full treatment of these alkaloids have only been given once before by Hino and Nakagawa in 1989

166

U. Anthoni, C. Christophersen and P. H. Nielsen

[5]. The latter review is entitled "Chemistry and Reactions of Cyclic Tautomers of Tryptamines and Tryptophans" and as indicated in the title the emphasis is on the chemistry and less on occurrence. Hence, while the chemistry and synthetic aspects are excellently treated in depth, the occurrence, biological/biochemical implications and structural details of these alkaloids are less exhaustively addressed. Therefore the total synthesis of the natural products will not be treated in any detail in this context, but will only be invoked in the discussion as a further proof of the correctness of the structural assignments. The intention of the present review is to list the available structures as exhaustively as possible. Moreover, the chemical and biological information has been evaluated critically in order to pinpoint existing errors and expose any regularities in appearance or biological function. However, first of all we want to draw attention to the possible implications of the accumulated knowledge concerning the synthesis, occurrence, and biochemistry of this class of compounds. As thoroughly documented in the previous review [5], tryptophan is the only naturally occurring essential amino acid where derivatives can be demonstrated experimentally to be in equilibrium with the isomeric cyclic form (cyclotryptophan). Several cyclotryptophan alkaloids discussed in the following section have also been shown to exist in equilibrium with the open form. Some cyclotryptophan alkaloids are polypeptides, i.e. chemically related to proteins. Finally, the biogenesis of cyclotryptophans have in some instances been proved to proceed from the open forms. In our opinion these facts strongly suggest the existence of enzymes which may convert protein-bound tryptophan to cyclotryptophan and vice versa. In the final chapter it will be shown that such an equilibrium might act as a "molecular switch" necessary for a hitherto unexplained function of acetylcholinesterase.

2. BACTERIA The only bacterial genus, where derivatives of cyclotryptophan have so far been encountered is Streptomyces. Except for the physostigmine alkaloids the only known compound at present is a dimeric cyclohexapeptide. Other comparable bacterial cyclopeptides are non-ribosomatically produced by peptide synthetases. Prominent among the intensively studied members of this group of enzymes is the multifunctional L-5-(a-aminoadipoyl)-L-cysteinyl-D-valine(ACV) synthetase producing the precursor L-5-(Q:-aminoadipoyl)-L-cysteinyl-D-valine in penicillin biosynthesis from the L-amino acid primary metabolites [6]. This enzyme is structurally and functionally related to other large modular multifunctional enzymes [7, 8] involved in nonribosomal peptide bond formation. Examples of such products are antibiotics like penicillin, gramicidin S, bacitracin, the immunosuppressive cyclosporin A and the surfactant surfactin [7]. The size of these enzymes range from approximately 126 kDa for bacterial one modul enzymes to more than 1600 kDa for the 11 modules comprizing the fungal cyclosporin A synthetase. The multienzymes have all modules aligned on a single polypeptide chain ranking them among the largest functional proteins known. Since a frequent occurring module concerns the epimerization domain, the presence of D-amino acid units in a peptide is indicative of the

167

Naturally Occurring Cyclotryptophans and Cyclotryptamines

Me

MeNHCOO

I H I Ri R, >i

1 2 3 4

H Me H Me

Me Me COMe COMe

A/' -Norphysostigmine Physostigmine

HO HhL^O

OH 5 Himastatin

-J2

operation of this class of enzymes. If a comparable biogenetic origin holds true in the Streptomyces case, it means that a whole series of such compounds await isolation because the motif responsible for the incorporation of cyclotryptophan is but one feature in the combined enzyme system necessary for the production of this compound. Furthermore, unless the hydroxylated cyclotryptophan is formed in an independent step, an enzyme module transforming tryptophan to the 3ahydroxylated cyclotryptophan exists. Such a module could form part of the enzymatic machinery responsible for the formation of this characteristic unit in several natural products.

168

U. Anthoni, C. Christophersen and P. H. Nielsen

Therefore, if the information for the production of such an enzyme is once expressed, available comparisons would predict this motif to be part of a family of genes present in the genome of other microorganisms as well. Streptomyces A^-Norphysostigmine (1) and physostigmine (2), known since 1864 from Calabar beans, were found to be responsible for the insecticidal activity of a Streptomyces species from soil [9]. Physostigmine has also been isolated from cultures ofS, pseudogriseolus [10]. The acetyl derivatives 3 and 4 produced by another soil living Streptomyces species are inhibitors for acetylcholine esterase and patented as a remedy for Alzheimer's dementia [11]. The unusual antitumor antibiotic himastatin (5) has been obtained from the culture broth of S. hygroscopicus [12-15]. The structure of this dimeric cyclohexadepsipeptide joined through a biphenyl linkage incorporates valine, leucine, threonine, a-hydroxyisovaleric acid, and 5-hydroxypiperazic acid, in addition to the dimeric l,2/?,3,3a/?,8,8a/?-hexahydro-2carboxy-3a-hydroxypyrrolo[2,3-b]indole-5-yl moiety.

3. CYANOBACTERIA The only secondary cyclotryptophan metabolites described from the procaryotic cyanobacteria (former blue-green algae) are a cyclodeca- and a cycloundecapeptide. The comments regarding peptide synthetases for bacteria are equally valid here. Cyanobacteria are efficient producers of cyclopeptide toxins with unusual amino acids [16] attesting to their genetic competence within this area. It would be of considerable interest to learn more about such enzymes capable of introducing the 3a-alkylated (prenylated) cyclotryptophan unit, part of many of the alkaloids considered in this review. Oscillatoria The toxic freshwater cyanobacterium O. agardhii has given rise to the isolation [17] of the chymotrypsin inhibitor (at 1.3 x 10'^ M) oscillatorin (6). The cyclic decapeptide is derived entirely fromL-amino acids but encompass the unusual amino acid unit (3a-cis)-l,2,3,3a,8,8ahexahydro-3a-(3-methyl-2-butenyl)-pyrrolo[2,3-^]indole-2-carboxylic acid called oscillatoric acid. The absolute configuration of oscillatoric acid has not been determined. The similarity between this amino acid and the alkaloids isolated from the marine bryozoan Flustra foliacea (see later) prompts the authors to suggests that the latter alkaloids may in fact be produced by the symbiotic cyanobacteria in the marine bryozoans. Microcystis M. aeruginosa is responsible for water blooms producing potent hepatotoxic cyclic peptides as e.g. microtoxins. The cyclic undecapeptide kawaguchipeptin A (7) isolated from one of these strains [18] is derived from L-amino acids as 6 from Oscillatoria ^ but deviates by incorporating D-leucine and two prenylated cyclotryptophan groups. The absolute stereochem-

Naturally Occurring Cyclotryptophans and Cyclotryptamines

169

J NH,

°Y 0

HOOC--~Y^°

6 Oscillatorin

T

H

HO

OH 6 " V I 1

o^x

0 CONH,

0<^N^0 H COOH

^ ' N^f 0

^O'^NHz

7 Kuwaguchipeptin A istry of the amino acids was determined from an acidic hydrolysis experiment followed by chiral gas chromatographic analysis and from HPLC analysis of the Marfey (l-fluoro-2,4dinitrophenyl-5-L-alanine amide) derivatives. The absolute configuration of the three chiral centers of the cyclotryptophan unit were all found to be S. The latter determination was performed on an acidic hydrolysis (1 % phenol in 6 N HCl

170

U. Anthoni, C. Christophersen and P. H. Nielsen

at 110°C for 4 hours) mixture derivatized with Marfey's reagent. The result showed L-tryptophan as part of the mixture. Loss of the isoprene unit during the hydrolysis reaction is hard to explain. Intuitively the result of the acidic hydrolysis would appear to be the 3a-prenylated cyclotryptophan. This compound would be expected to tolerate acidic conditions quite well. When treated with Marfey's reagent it could form either a N-1 monosubstituted or a iV-1, A^-8 disubstituted derivative. It would be unfortunate if any of these derivatives accidentally had the same retention time as that of the Marfey derivative of L-tryptophan. This question ought to be clarified, not only because of the extensive use of these reactions in modern biological chemistry, but especially in order to clarify the stability of the cyclotryptophan natural products. Interestingly, the same extract also contained kawaguchipeptin B [19] having mstead of the prenylated cyclotryptophan moieties the normal L-tryptophan residues. Except for this feature kawaguchipeptin B deviates from A only in the exchange of the D-leucine of A with a L-leucine in B. These facts indicate that these peptides do not represent post-translationally modified ribosomal peptides. Also, apparently kawaguchipeptin B is not a precursor for A. The most likely explanation, in our view, is that different peptide synthetases are responsible for the synthesis of the two compounds. Both peptides inhibit growth of the Gram-positive Staphylococcus aureus (MIC 1 /ig/ml).

4. FUNGI The kingdom of fungi encompasses the filamentous microfungi (moulds) treated in this section. The classification of the microfungi is difficult and still in a state of flux [20]. Accordingly, at least some of the species treated here are presumably not identified with the currently correct nomenclature. Some species produce mycotoxins and are responsible for rendering food toxic. Apart from higher plants the fungi has given rise to the identification of the largest number of cyclotryptophans. Of the 61 structures present in the following section all but three are diketopiperazines. The remaining three (41, 42 and 48) are built over the same scheme where an amino acid unit is anthranilic acid expanding the piperazine ring with one C-atom. This means that all are in fact cyclodipeptides. The comments given above concerning the existence of peptide synthetases holds equally well for the fungal dipeptides treated below. A note of warning: Access to the chemical literature on these dipeptides is hampered by the presence of a relative abundance of misprinted formulas, especially with regard to absolute stereochemistry. Penicillium Roquefortine, or roquefortine C (8), is an important neurotoxic metabolite of P. roquefortii, the mold of many varieties of blue cheese [21-23]. The absolute configuration of 8 has been established by chemical and spectroscopic techniques [24]. The occurrence, structural characterization, biological activity, conditions for production, and analytical

171

Naturally Occurring Cyclotryptophans and Cyclotryptamines

H

8 (R=H)

Roquefortine (C)

10 Roquefortine D=

9 (R=CHO)

Formylroquefortine

3,12-Dihydroroquefortine

determination of 8 was reviewed in 1984 [25]. Almost 50 papers have appeared since then and only the most important will be mentioned here. Thus, 8 has recently been detected in strains of P. roquefortii from feed grain and maize silage [26, 27], P. verrucosum var. cyclopium from sausage and other strains [28, 29], P. terrestre [29], two strains of P. chrysogenum [30], and from P. crustosum in samples of reject kidney beans and contaminated beer [31-33]. Roquefortine and the dihydroderivative roquefortine D (10) have been isolated from cultures of P. roquefortii [34, 35], P. purpurrescens [29], P. corymbiferum [36], four strains of P. expansum [37], and from P. farinosum [38-43]. In the latter fungus four additional metabolites were shown to be ringopened isomers of the cyclotryptophans 8 and 10. By feeding the mycelium with ^'^Croquefortine one of the metabolites was shown to be formed by degradation of 8 with initial ring-opening, i.e. reminiscent of the tryptophan-cyclotryptophan conversion mentioned in the introduction. The mechanisms for excretion and uptake of 8 by P. crustosum have recently been thoroughly characterized [44]. In P. oxalicum, P. glandicola, and P. atramentosum 8 is a precursor of other alkaloids [45, 46]. Although 8 is neurotoxic the concentrations usually found in blue cheese (0.2-2.3mg/kg) are too small to present any health problems [47, 48]. The antibiotic properties have been shown in cultures of Corynehacterium fluccumfaciens to arise from inhibition of the RNA synthesis [49]. The production of 8 in different isolates of P. roquefortii from Cabrales blue cheese has been studied in order to select a non-toxic strain for food manufacture [50]. A screening method for the detection of thirteen mycotoxins, including roquefortine, has

172

U. Anthoni, C. Christophersen and P. H. Nielsen

appeared [51]. A total of 16 mycotoxins, including again 8, can be detected by their action on selected human or porcine cell lines [52], but the use of eucaryotes and bacteria as biosensors is also feasible [53]. TLC and HPLC have been standardized for detection of mycotoxins, including 8, especially for use in chemotaxonomy of Penicillium strains [54-57].

a

^

MeCO ^

11 Fructigenine A

12 Fructigenine B=Verrucofortine

During screening for bioactive metabolites, fructigenine A (11) and B (12) were isolated from P. fructigenum [58]. Only 11 inhibited growth of Avena coleoptiles and L-5178Y cells. The other metabolite 12 is identical with verrucofortine isolated the preceding year from P, verrucosum var. cyclopium [59]. A strain of this species (RV 67718) isolated from ground cassava collected in Burundi [60] in addition to 8 furnished formylroquefortine (9). Other diketopiperazine metabolites, the symmetrical dimer nigrifortine (13) and brevianamide E (14), were isolated from P. nigricans [61], and P. brevi-compactum [62], respectively. The asynmietric total synthesis of 14 has been performed and has established that

Naturally Occurring Cyclotryptophans and Cyclotryptamines

' 13 Nigrifortine=Amauromine

kXi^^Ao 14 Brevianamide E

OH ^"

OH ^"

_LHi

173

X

-4^\

OMe

1^

16 Okaramine B

it is derived from L-tryptophan and L-proline. Since the tryptophan derivative deoxybrevianamide E is slowly converted to 14 by atmospheric oxidation (cf section 7), reported syntheses of the former constitutes the synthesis also of 14 [63-66]. Okaramine A (15) and B (16) are insecticidal alkaloids from P. simplicissimum [67-70]. Amauroascus The symmetrical alkaloid 13 (1.35 g pure compound) was also isolated from a liquid culture (75 1) of an Amauroascus sp. and given the name amauromine [71-73]. The compound has a low toxicity in mice (LD50 > 200 mg/kg), but exhibit potent hypotensive and vasodilating activity. The vasodilating activity is believed to originate in a calcium antagonism. The partial and total synthesis from L-tryptophan has been reported [74-78].

U. Anthoni, C. Christophersen and P. H. Nielsen

174

CH2OH

m 17

H

2

2

Chaetocin

18

H

2

3

Chaetocin B

19

H

3

3

Chaetocin C

20

OH

2

2

Dihydroxychaetocin=Melinacidin E

21

OH

4

4

Chetracin A

Chaetomium All cyclotryptophan metabolites produced by this and many of the following fungal species contain sulfur. Sulfur containing alkaloids have been reviewed in 1985 [79] and 1992 [80]. Chaetocin (17) has been isolated from C. minutum [81], four soil strains of Chaetomium [82], and from C. thielavioideum [83]; the absolute configuration has been determined by Xray techniques [84]. It is most unfortunate that the stereostructure was depicted incorrectly since this has given rise to a fair amount of confusion regarding the absolute configuration of this series of compounds. As correctly stated in the paper, the stereochemistry of the chiral centers of the diketopiperazine rings are all of S configuration in contrast to the findings for the sporidesmins (32 - 40 except 38). Chaetocin is antibacterial and cytostatic but is devoid of activity against herpes, vaccinia and Newcastle Disease vira [81, 85]. The antibacterial activity is especially potent against Staphylococcus aureus where the minimum inhibitory concentration (MIC) was determined as 0.01 Mg/ml against a penicillin sensitive strain and, most interestingly, 0.001 /xg/ml against a resistant strain [81]. The acute toxicity expressed as LD50

Naturally Occurring Cyclotryptophans and Cyclotryptamines

175

in mice was determined as 1.7 mg/kg intraperitoneally and > 1000 mg/kg orally. Some studies toward the total synthesis have been described [86].

1

a

Me

N " Me'^^T ^CH2 OH 0

N4Y° I

CH2OH 22 Chetomin

CO

MeS

Me i^SMe 0

23 DethiO"-tetra(methylthio)chetomin The corresponding epitetrasulfide, chetracin A (21), was obtained from C. abuense, C. tenuissimum, C. nigricolor, and C. retardatum [87, 88]. Two strongly cytotoxic congeners of chaetocin with trisulfide bridges, chaetocin B (18) and C (19), originate from C. virescens var. thielavioideum [87]. C. retardatum also produces the cytotoxic dihydroxychaetocin, melinacidin IV [88].

176

U. Anthoni, C. Christophersen and P. H. Nielsen

The antibiotic chetomin (22) is a metabolite of C. cochlioides, C. globosum [89, 90], C. funicola, C. umbonatum [91], and C. subglobosum [83]. The effect on growth and fermentation of rumen bacteria is strongest for Gram-positive strains [92]. Treatment of 22 with NaBH4 and CH3I furnished 23 (structure and absolute stereochemistry confirmed by X-ray crystallography) which has also been isolated from C globosum [93] and proved to be active against several bacteria. The role of antibiotics produced by C. globosum in biocontrol of Pythium altimum, a causal agent of damping-off, has been discussed [94].

9H2-R 'N'

H^^N I

CH2 —R2

24 25 2 26 2 27 2 28 2 (or 3

m

R1

2 (or 2 2 2 3 2)

H H

OH

H

OH

Melcnicidin 1

OH

OH

OH

OH

OH

H) H

Melanicidin H

H

H

OH

OH

Verticillin A

H

OH

OH

OH

Verticillin B

H

OH

OH

OH

Verticillin C

Acrostalagmus The epidithiodiketopiperazines, melinacidins II (24), III (25), and IV (20) from Acrostalagmus cinnabarinus var. melinacidinus inhibit a variety of Gram-positive bacteria in vitro but are toxic and ineffective in treatment of bacterial infections in vivo [95, 96]. The structure of several other related metabolites from this fungus have never been clarified.

Naturally Occurring Cyclotryptophans and Cyclotryptamines

177

Gliocladium Two inhibitors of a human proto-oncogene induction, Sch 52900 (30) and Sch 52901 (29), have been isolated from the fermentation broth of the fungal culture (SCF-1168) Gliocladium sp. [137]. Verticillin A (26) was isolated from the same fermentation broth and displays the same activity as 30 and 31.

29

R=H

Sch 52901

30

R=OH

Sch 52900

Verticillium A strain of Verticillium isolated from a basidiocarp of Coltricia cinnamomea produced three antibiotic and cytotoxic epidithiodiketopiperazines, verticillins A (26), B (27), and C (28) [97, 98]. The verticillins are active against Gram-positive bacteria and mycobacteria but inactive against Gram-negative bacteria and fungi. The most toxic component in cultures of V. dahliae also is 26 [99]. From the mycelium and culture of V. tenerum 20 has been reported [100]. Pithomyces Other epithiadiketopiperazine indole alkaloids are the sporidesmins of most strains [101] of the fungus P. chartarum and treated in several reviews [102, 103], most recently by White et al. [104]. Several sporidesmins have been subjected to X-ray crystallographic structure determination [105-109]. Sporidesmin (or Sporidesmin A) (32), B (33), C (36), D (37), E (34), F (38), G (35), H (39, tentative structure) and J (40) have all been described from P, chartarum [110-114]. The infrared spectra of 32, 33, 34 and 37 have been discussed [115],

U. Anthoni, C. Christophersen and P. H. Nielsen

178

31

Gypsetin

Me n 32 33

2 2

OH H

Sporidesmin (A) Sporidesmin B

34

3

OH

Sporidesmin

E

35

4

OH

Sporidesmin

G

36

Sporidesmin C

as have the ^^C NMR spectra of 32 and 37 [116], the electrospray mass spectra of zinc and cadmium complexes of 32 [117], and the circular dichroism of 32 [118]. Sporidesmin derivatives have been prepared synthetically [119, 120] as has 32 [121] and (±)-33 [122]. Sporidesmin (32) is responsible for the "facial eczema" in ruminants, reviewed in 1985 [123]. From a mechanistic point of view sporidesmin is the most intensively investigated compound. The disulfide group is essential to the activity and was early shown to be involved in the action on swelling and respiration of liver mitochondria [124]. The reduced dithiol form readily undergoes autoxidation in vitro generating the superoxide radical (O2"). The superoxide radical is also generated in a cyclic reduction/autoxidation reaction of the disulfide with glutathione [125]. In addition to superoxide radical and hydrogen peroxide the extremely reactive hydroxyl radical is formed in the reaction [126] and it is interesting that sporidesmin itself generates exactly the agents (superoxide ion and hydroxyl radicals) capable of affecting the ring closure reaction of the substituted tryptophans to sporidesmins. The mutagenicity [127]

Naturally Occurring Cyclotryptophans and Cyclotryptamines

^N-^Y"

179

MeO

Me "SMe 37

Sporidesmin D

38

MeO

Sporidesmin F

MeO

MeO M'eo^N^ Me 39

Sporidesmin H

Me

Me 40

Sporidesmin J

and effect of sporidesmin on vitamin B12 levels [130], liver [129], kidney [130], and lipid bilayers [131] have been reported. The radical forming reaction is inhibited in vitro by mercaptide-forming metals and iron [132], zinc [133], molybdenum and sulfur [134] can be used for protection of sheep against sporidesmin intoxication. Copper and to a lesser extent iron ions catalyze the autoxidation reaction. Copper-chelating agents were found to inhibit the reactions [135]. Corollospora The marine Ascomycete, C. pulchella was isolated from the surfaces of sand grains. From a culture broth of this fungus cultivated in sea water three diketopiperazines, gancidin W, 20 and 25 were isolated by bioassay-guided fractionation using the activity against Staphylococcus aureus. The identification was performed by comparison of physico-chemical and spectroscopical data with the ones reported [136]. Nannizzia Gypsetin (31) is an inhibitor of acyl-CoA: cholesterol acyltransferase produced by N, gypsea var. incurvata, the structure of which was clarified by X-ray analysis [138, 139]. Since it is an important drug discovery lead, a highly concise total synthesis has been developed

U. Anthoni, C. Christophersen and P. H. Nielsen

180

[140]. Aspergillus In this section the focus is placed on updating the review of the Aspergillus alkaloids which appeared in 1986 [141].

a

,<^

N . I H

?4°

41

R=H

Aszonalenin

42

R=MeCO

LL-S490p

Ardeemin 43 R^ sRj =H 44 Ri =H, R2 =MeCO AZ-Acetyladeemin 45 Ri =0H. R2 =MeCO A. zonatus has given rise to the isolation and structure elucidation [142] of a cyclotryptophan anthranilic acid diketopiperazine aszonalenin (41) and the acetyl derivative LLS490iS (42) previously found in an uidentified Aspergillus [143]. The relative and absolute configurations of aszonalenin have been determined by the synthesis of (-)-dihydroaszolenalenin from L-tryptophan [144, 145]. Since 41 furnish (+)-dihydroaszolenalenin on hydrogenation it is, according to this investigation, derived from /^-tryptophan. A. fisheri var. brasiliensis produces three cyclotryptophans, ardeemin (43) and two congeners, A/-acetylardeemin (44) and the hydroxy derivative 45. The metabolites were secured

Naturally Occurring Cyclotryptophans and Cyclotryptamines

181

^

HH 1

^ H R

46

R=H

Epiamauromine

47

R=Me

A/-Methylepiamauromine

a 48

Asperlicin E

by a bioassay-guided isolation monitoring the attenuation of multiple drug resistance in tumor cells. The structure of 44 was confirmed by single crystal X-ray diffraction analysis [146, 147] and by total synthesis [78]. The sclerotia of A. ochraceus has yielded three diketopiperazine metabolites which exhibit moderate activity towards the corn earworm Helicoverpa zea, resulting in a reduction of weight gain when present in the feed of the lepidopteran insects. They have been patented as insecticides. Two of the metabolites are the cyclotryptophans epiamauromine (46) and A^methylepiamauromine (47) [148]. These cyclotryptophans are closely related to amauromine (13, nigrifortine) except for the absolute configuration of the carbon atoms forming the junction between the two five-membered rings. All three compounds are derived from Ltryptophan. Asperlicin E (48) is a cyclotryptophan fromy4. alliaceus [149]. 48 originate biosynthetically from asperlicin C, which is also the precursor of asperlin. By the use of ^'^C-labelled precursors, 48 was shown to originate from tryptophan and anthranilate [150]. The resting

182

U. Anthoni, C. Christophersen and P. H. Nielsen

RH

N"IN H0

( . N"

RH 0

49 50

0 NxM e

R=H Ditryptophenaline R=Ph~--~_S

cells of this organism, which were used for the biosynthesis experiments, interestingly enough, were able to incorporate amino acid analogues into the alkaloids. In this way several new compounds were produced among which were three fluoro-substituted asperlicin E's. A total synthesis of 48 from L-tryptophan has been published [151]. The synthesis also gave the diastereomeric compound with the inverse absolute configuration in the ring junction. The latter structure was secured by an X-ray structural determination and thus also proving the stereochemical assignment of 48. Asperlicin E is an antagonist of the neuropeptide cholecystokinin receptor. The dimeric alkaloid ditryptophenaline (49 with S, S, S, S-configuration) originates from A. flavus var columnaris and several related strains and the structure was clarified by X-ray methods [152]. The absolute configuration and conformation in solution of 49 has been established by total synthesis [153] and from NMR studies [154]. From soil treated for long periods with pesticides, a strain of A. flavus was isolated which in addition to 49 produced cyclo-(L-tryptophyl-L-phenylalanyl) [155]. Since the latter metabolite on photooxidation gave 49, it is a possible precursor. In another study I'-(E-2phenyl-ethylene)-ditryptophenaline (5tl) was isolated from A. flavus; the stereochemistry at the point of dimerisation may be an important determinant of biological potency for these

Naturally Occurring Cyclotryptophans and Cyclotryptamines

51

R=Benzyl

WIN 64821

52

R=lsobutyl

WIN 64745

53

Asperazine

183

U. Anthoni, C. Christophersen and P. H. Nielsen

184

H

00 Oc; ^ O^^^Me 54

n=2,m=2

Leptosin A

61

n=2

Leptosin D

55

n=3,m=2

Leptosin B

62

n=3

Leptosin E

56

n=4,m=2

Leptosin C

63

n=4

Leptosin F

57

n=4,m=3

Leptosin G

58

n=3,m=3

Leptosin Gi

59

n=2,m=3

Leptosin G2

60

n=2,m=4

Leptosin H

cyclotryptophans [156]. An Aspergillus sp., SC319 isolated from soil yielded two related diketopiperazine dimers, WIN 64821 (51) and WIN 64745 (52), both with /?,/?,/?,/?-configuration, in addition to cydc>(L-tryptophyl-L-phenylalanyl) [157]. The latter peptide could also be isolated after treatment of 51 with hot acetic acid. As correctly pointed out by Barrow et al. [157] ditryptophenaline has opposite chirality at the indoline junction to the WIN alkaloids; it is therefore unfortunate that the latter authors depict 49 incorrectly in their paper. In ensuing studies the fermentation, isolation, biological activity and preparation of 36 biosynthetic analogs from this strain useful as ligands for the neurokinin-1 receptor were studied [158-161]. The authors concluded that both the mdoline and phenyl moieties were involved in binding to the receptor. A culture of A. niger derived from a salt water fermentation of an isolate from a Caribbean marine sponge, Hyrtios proteus, yielded the selective cytotoxic asperazine (53) [162]. This dimeric alkaloid is composed of L-tryptophan and D-phenylalanine.

185

Naturally Occurring Cyclotryptophans and Cyclotryptamines

R2^ >Rl

H HI

H HT i ^

N-"

S^N \

a

,0H

Me

^Me

64

Ri=CH20H. R2=H

Leptosin I

66

n=2 Leptosin K

65

Ri=H. R2=CH20H

Leptosin J

67

n=3 Leptosin Ki

68

n=4 Leptosin K2

Leptosphaeria The leptosins (54 - 68) comprises a series of epipolythiodioxopiperazines isolated from the mycelium of a strain of Leptosphaerica sp., OUPS-4, attached to the marine alga Sargassum tortile collected in the Tanabe Bay of Japan [163-166]. The metabolites, secured from a culture of the fungus in artificial sea water, were isolated ensuing a bioassay-guided isolation procedure. Ten leptosins (54 - 60 and 66 - 68) are dimeric cyclotryptophans only differing in the number of sulfur atoms in the polythio bridge. The leptosins D-F (61 - 63) incorporate the 3-indolyl moiety while 64 and 65 are isomeric epitetrathioderivatives. It is noteworthy that all leptosins, except D, E and F, have one cyclotryptophan unit (upper one in the formulas depicted) with epimeric 3a,8a configuration to that observed in the chaetocin (17) - chetracin A (21) series. Furthermore, the leptosins K - Kj (66 - 68) in addition have opposite absolute configuration around one of the sulfur bridges. All compounds showed significant cytotoxic activity (ED50 around 4 ng/ml), exceeding that of mitomycin C (ED50 around 20 ng/ml), in the P388 lymphocytic leukemia test system in cell culture. For those possessing an epipolythiobriged piperazinedione moiety the activity is essentially independent of the number of sulfur atoms. Two compounds, 54 and 56, moreover exhibited antitumor activity against Sarcoma 180 ascites. The structure of 66 was established by X-ray analysis [166]. This compound exist in a mixture of four different conformers incorporated in an equal ratio in a single crystal, while only one or two conformers are present in solution.

186

U. Anthoni, C. Christophersen and P. H. Nielsen

5. HIGHER PLANTS Higher plants have yielded the largest number of compounds of the cyclotryptophan series. However, of the about 80 structures the majority originate with Rubiaceae and Apocynaceae and only a few with six other families. It is thought provoking that a family as large as Leguminosae (the Pea Family) with around 700 genera and 17,000 species has only yielded a single cyclotryptamine-containing species and the Lily Family (Liliaceae, 250 genera and 3,500 species) only one cyclotryptamine. Likewise, inLoganiaceae (30 genera, around 600 species) only one cyclotryptamine is known. In this context it should be borne in mind that the classical procedure for isolating alkaloids almost always invoked treatment with rather strong acid in the purification steps. Since acid catalyzed formation of cyclotryptophans may be reversed in a ring opening process and/or other rearrangement processes the native natural products must in many cases have been more or less transformed in this way [172]:

H

^

H

H

" IX. OMe

I

H

" IX. OMe

Especially the Leguminosae are well known to harbour symbiotic microorganisms in nodular rhizomal aggregates. The physostigmines have also been identified from a bacterial source. Perhaps the physostigmines from the Calabar bean originate with associated bacteria. Furthermore, most higher plants associate with symbiotic fungi. An investigation of the nutrient transfer between trees in a temperate forrest, where at least 90% of the roots are colonized by fungi in an ectomycorrhizal relationship, has shown that a nutrient flux exists between species [167, 168]. It is to be expected that other plant species also experience an analogous situation allowing secondary metabolites to be transferred through the hyphae of the mycorrhizal fungi to materialize in other as well as the host species. Alternatively, the symbiontic fungi could be responsible for the biogenesis of the alkaloids. More than half of the known compounds from plants are dimers or polymers. Except for one example (from Idiospermaceae), all 14 trimeric or higher polymeric cyclotryptophans originate with the Rubiaceae. With the exception of two rearranged compounds all polymers

Naturally Occurring Cyclotryptophans and Cyclotryptamines

187

represent variations of conjugation of A^b-methylcyclotryptamine. The structures indicate that the mechanism of polymerization could well proceed via free radical mechanisms. In this connection it is interesting that the tryptophan radical cation has shown evidence of anchimeric spin delocalization [169]. This spin delocalization results in spin density being transferred through space from the TT system of the indole nucleus into the alanyl side chain. Furthermore, under free radical conditions cyclotryptophans suffers oxidation at the 3a-position [170] which is often the position joined to another cyclotryptophan molecule in the polymeric derivatives. Comparable reactions are observed by electrooxidation of methylindoles [171]. Other mechanisms are possible as well. Tryptophan derivatives are known to cyclize by cationic mechanisms on treatment with acid [172, 173] forming the diastereomeric cyclotryptophan derivatives from the intermediate 3-H indolinium ion. The latter species may in a competing reaction give rise to formation of dimers [174, 175] and mimic the biogenesis of many alkaloids. Furthermore, the unique complex formed between trifluoroacetic anhydride and pyridine [176] react with cyclization and 3a-arylation [177, 178] comparable, e.g. to the structures of the leptosins 61 - 63).

® H

OMe OMe OMe By combinations of reactions of this type it is possible to rationalize the biogenesis of most of these alkaloids. Investigations concerning the structural restraints in this type of

188

U. Anthoni, C. Christophersen and P. H. Nielsen

reaction is in progress [179] and may cast new light on these problems. Most of the remaining alkaloids are of the monoterpenoid or iridoid class, often with loss of one carbon atom. The enzymology of the genesis of the indole alkaloids has been reviewed [180], however, the bulk of information relate to the important Catharanthus alkaloids and nothing is known of the enzymes responsible for the generation of cyclotryptophans.

OH

HH I Me 69

Alline

5.1 Liliaceae In the Liliaceae only one cyclotryptamine is known. Alline, 3a-hydroxy-l-methyll,2,3,3a,8,8a-hexahydropyrrolo[2,3-^]indole(69), was'vs>o\2XtdftomAllium odorum [181]. The structure was established by an X-ray crystallographic investigation. Later the same compound was obtained from A. senescens and A. anisopodium [182] and detected by TLC in seven {A. victorialis, A. altaicum, A. anisopodium, A. senescens, A. stelleranum, A. ramosum syn. A. odorum, A. splendens) but not in A. sibiricum, A. bidentatum and/l. leucocephalum out often species [183]. The relative stereochemistry is revealed from the X-ray structure as cw-annelation of the pyrrolidine rings. Alline is optically active ([a:]D +136.3° in CHCI3) but the absolute configuration seems to be undetermined. Accordingly formula 69 is not intended to depict absolute configuration. Being the simplest member identified of the 3a-hydroxycyclotryptamines, representing the N-\ methylated skeleton, the structure is of considerable interest not at least for reference purposes. Since alline has never been synthesized, reisolation and study of the structure, reactions and activity is warranted.

5.2 Leguminosae. From this family originate the first, best known, and from a conmiercial point of view, the most important compound, physostigmine (2). The seeds of the Calabar Bean, Physostigma venenosum, contain apart from 2 also 1, eseramine (70) and a series of chemically related alkaloids. The isolation, structure elucidation, synthesis and pharmacology of the Calabar Bean alkaloids have been reviewed covering the literature until the end of 1988 [184-188]. The absolute configuration of naturally occurring 2 and the other alkaloids was established by a

Naturally Occurring Cyclotryptophans and Cyclotryptamines

189

Me

MeNHCOO.

f

•P

N

Me 70

CONHMe

Eseramine

RO. I H i ^-OH Me Me

I H "^ Me

R=MeNHCO Physostigmine /V-oxide

Geneserine

R=H Geneseroline AZ-oxide

Geneseroline

•Me

combination of chemical and CD studies [189, 190]. Geneserine, isolated from the basic extracts of the P. venenosum seeds, isomerizes in acid to physostigmine A'-oxide [191]. Identical behavior was found for geneseroline, where the structure of the N-oxide was determined by X-ray crystallography. The solution and solid state structures of the (-)-A/-heptylcarbamate of geneseroline and its hydrochloride salt, where an analogous equilibrium exist, have been studied by NMR and X-ray crystallography [192]. Since the A/-oxides are in equilibrium with the corresponding hexahydro-l,2-oxazino-[5,6^]indoles, the presence of one or the other in the biological material becomes a question of the local pH in the plant. There are indications that equilibria of this type could be of quite common occurrence since several alkaloids contain the structural elements required for the isomerization to proceed. Another example involving the bryozoan alkaloid flustrarine B is discussed later. Phenylcarbamates of 2 and related compounds are inhibitors of cholinesterase and may have potential for treatment of Alzheimers's desease [193, 194]. A considerable amount of papers related to the synthesis of 2 have appeared since 1989 [195-208].

5.3 Calycanthaceae. This family is represented only by two genera, Calycanthus and Chimonanthus. Chimonanthine (71), folicanthine (72) and calycanthidine (73) are bisindole alkaloids identified from plants in this family. The alkaloids have been reviewed several times, most recently in 1988 [1, 5]. (±)-Folicanthine and (±)-chimonanthine have been synthesized [209]. The

U. Anthoni, C. Christophersen and P. H. Nielsen

190

Ri

a

Me

\ H I. N. .N

:]

Co? R2

H

R3

71

R,=R2=H, R3=CH3

Chimonanthine

72

Ri=R3=CH3, R2=H

Calycanthidine

73

Ri=R2=R3=CH3

Folicanthine

Me Me J. H .1 .N

n

^

a a 74

H Me I H N. ,N

;]

J

N I H I Me Me

Idiospermuline

structure of a bis-borane complex of synthetic (±)-folicanthine has been subjected to X-ray stractural determination [210].

Naturally Occurring Cyclotryptophans and Cyclotryptamines

191

5.4 Idiospermaceae From the seeds of the tree Idiospermum australiense, the sole member of the primitive angiosperm family Idiospermaceae closely related to the Calycanthaceae, the trimeric pyrroloindole alkaloid idiospermuline (74) has been isolated [211]. The absolute stereochemistry was established by an X-ray crystallographic study of the trimethiodide. Chimonanthine (71), isolated as well, shows suppression of the cholinergic transmission as does idiospermoline (74). The alkaloids were isolated as a result of a bioassay-guided fractionation usmg this depressing activity. The seeds have been implicated in cattle poisoning.

H

Me "

75 76

R=NHMe R=N(Me)2

Borreverine Methylborreverine=Auricularine

77

R=OH

Spermacoceine

H 78

Me H

Hydroxydihydroborreverine

5.5 Rutaceae. The alkaloids isolated from Flindersia foumieri from New Caledonia include the cyclotryptamines borreverine (75), methylborreverine (76), and hydroxydihydroborreverine (78) and the related isoborreverines [212-215]. Several reviews of 75 [1, 216], the X-ray structure [217], antibacterial activity [218], biomimetic synthesis [219] and analysis of the "C NMR spectra have appeared [220, 221]. Apart from these alkaloids Rutaceae is mainly characterized by the occurrence of polymeric hexahydropyrroloindoles.

U. Anthoni, C. Christophersen and P. H. Nielsen

192

H H Me N. N

H H Me N. N R

Cx

:] H H Me N ,N

a

R S

Me H

H+

:i

'} ^N

H

N' HH Me 79

a a 81

80

Hodgkinsine

H

Me

H

a H

H

i

Me

a

H

Calycosidine

Me

H

cx

Me

H

cx

1

Me

Me

J

J

H

Me Quadrigemine-A

Me

82

Me

Quadrigemine~B

5.6 Rubiaceae Only a few bis-indole alkaloids have been reported. Thus, 4-methylborreverine (76) is identical with auricularine from extracts of dried and powdered Hedyotis auricularia [111]. Borreverine (75) and the new spermacoceine (77) were both isolated from the dried aerial parts

Naturally Occurring Cyclotryptophans and Cyclotryptamines

HH

H H Me 83

Quadrigemlne-C

193

Me

O:

H H Me

J

H H Me

84

Psycholeine

of Borreria verticillata [223]. The tri-indole hodgkinsine (79) was isolated from the leaves of a shrub growing in tropical Queensland, Hodgkinsoniafrutescens, and the structure was solved by X-ray crystallography [224, 225]. The H. frutescens alkaloids have been thoroughly reviewed [1]. Hodgkinsine (79), also isolated from Psychotria rostrata [226] and Psychotria oleoides [227], under mild acidic conditions is transformed into calycosidine (80) [228]. Since both compounds are found together in Calycodendwn milnei the latter alkaloid may be an artifact [229]. The two tetra-indolic alkaloids quadrigemine-A (81) and -B (82) were first described from Hodgkinsoniafrutescens [230] but have also been found in Psychotria forsteriana [231233]. The absolute stereochemistry has not been unambiguously determined; however, 81 was considered to be a 1:1 mixture of diastereomers. Clearly, this problem calls for reinvestigations relying on X-ray methods. The compounds possess potent bioactivity. Cytotoxicity against cultured rat hepatoma cells greatly exceed that of the antitumor chemotherapeuticum vincristine. Two further tetra-indolic alkaloids, quadrigemine C (83) and psycholeine (84), both with undetermined absolute stereochemistry, were co-occurring in Psychotria oleoides [111]. Since they bear a similar relationship to each other as 79 and 80, 84 may be an sutifact formed from 83 during extraction and purification. Interestingly, psycholeine (84) was isolated in a bioassay-guided fractionation owing to its somatostatin antagonistic activity. A total of four penta-indolic alkaloids have been found in the Rubiacea. These include psychotridine (85) from Psychotria beccaroides [234] and Calycodendron milnei [229], 85 with isopsychotrodine C (92) in Psychotria forsteriana [231-233], and 85 with the isopsychotridines A (90) and B (91) in Calycodendron milnei [229]. The chemistry of 85 has been reviewed [1].

U. Anthoni, C. Christophersen and P. H. Nielsen

194

Me H

Me H

C

c

^

H

la

Me H

Me

C

;]- I n H

30

^

H

Me

Me

;]

a

H H

Me

CK- J H

Me

•N Me

H

J

Me

85

n=1

Psychotridine

90

Isopsychotridine A

86

n=2

Valine

91

Isopsychotridine B

87

n=2

Vatine A

92

Isopsychotridine C

88

n=3

Vatamine

89

n=4

Vatamidine

The stereochemistry of the isopsychotridines is not known. The cyclotryptamines 81, 82, 85 and 92 are potent inhibitors of aggregation of human platelets [232]. Polymers containing six, seven, and eight cyclotryptamine units have been isolated from Calycodendron milnei [235]. They include vatine (86) with the stereoisomer vatine A (87), vatamine (88), and vatamidine (89). The in vitro cytotoxicity of the trimers, tetramers, pentamers, and hexamers have been investigated on human leukemic and rat hepatoma cell lines [236, 237]. They show strong dose-dependent activity in the micromolar range, increasing with the number of, but varying with the position of the bond connecting the cyclotryptamine units.

Naturally Occurring Cyclotryptophans and Cyclotryptamines

93

Ri =H, R2 =Me

Corymine

94

Ri=COMe,

Acetylcorymine

95

Ri =R2 =H

96 97

98 99

R2=Me

195

Demethylcorymine

R=Me

Isocorymine

R=H

Norisocoiymine

R=Me

Erinine

R=CH2Me

Erinicine

5.7 Apocynaceae.

Hunteria H. umbellata is a glabrous tropical tree with leaves, bark and roots used in local Western Nigerian medicine. The major cyclotryptamine alkaloids in the seeds are the lipophilic corymine (93), acetylcorymine (94) and isocorymine (96) [238], while the leaves contain 93, erinine (98), erinicine (99) and eripine (100) [239]. The aqueous extracts of the seeds, having

U. Anthoni, C. Christophersen and P. H. Nielsen

196

Ri COOMe

100

R^^CHjOH, R2=Me

Eripine

101

Ri=CHO, R2=Me

Eripind

102

Ri=CHO, R2=H

Noreripinal

103

Ri=R2=0H, R3=R4=H

104

Ri=R2=H, R3=R4=0H

105

Ri=R3=H, R2=R4=0H

106

Ri=R4=H. R2=R3=0H Abereamines

ethnomedical use as a cure for, e.g. diabetes and stomach ulcer, yielded four isomeric abereamines (103 - 106) [240]. H. ebumea in addition to 93, 94, 98 and 99 and desformocorymine (107) [241] has given rise to several quaternary alkaloids including the cyclotryptamine hunteracine chloride (109) [242], The latter alkaloid has been subjected to an X-ray crystallographic structure determination clarifying also the absolute stereochemistry [243]. H. elliotii has yielded a variety of alkaloids [244, 245] but the cyclotryptamines 93, 94, 107, and ep/-3-dihydrocorymine (110) were only present in the leaves. The seeds of H. congolana contain five cyclotryptamines, 93, 94, 107, 110, with the 3-acetate (111) and 3,17diacetate (113). Norisocorymine (97), 96, and 98 -102 were characteristic of the leaves [245, 246]. H. zeylanica var. africana, collected in Kenya, has yielded twenty alkaloids in one investigation [247] and the same species, collected in Sri Lanka, fourteen indole alkaloids in another [248] including 93, 96, 97, 110, 111, and 3-^pz-dihydrocorymine 17-acetate (112).

197

Naturally Occurring Cyclotryptophans and Cyclotryptamines

COOMe

107

R=Me

Desformocorymine=Deformylcorymine

108

R=H

Demethyldeformylcorymine

109

Hunteracine

RiOCH2^ XOOMe

110

R^ =R2=H

3 - £ p / -dihydrocorymine

111

Ri=H, R2=MeC0

3-fp/-dihydrocorymine 3-acetate

112

R^=MeCO, R2=H

3-fp/-dihydrocorymine

113

R-|=R2=MeC0

3-£p/-dihydrocorymine 3,17-diacetate

17-acetate

Coryzeylamine (114) and deformylcoryzeylamine (115), two dimeric indole alkaloids composed of sarpagine and echitamine-type monoterpenoid indole alkaloids, were identified from leaves from this species collected in Thailand [249], which species also gave rise to the isolation and structure elucidation of A^g-demethylcorymine (95) and A'a-demethyldeformylcorymine (108) [250]. The ^H NMR spectra of the hemiacetalic 114 was considerably simplified in DMSO-^e or pyridine-cfj as compared with CDCI3 solution owing to the predominance of the hemiacetal shown in structure 114. In acidic solution, 107 and 110 easily rearrange, and in the former case with conversion to the akuammiline skeleton [251]. In the case of 110 a series of rather complex events follow

U. Anthoni, C. Christophersen and P. H. Nielsen

198

COOMe CH2 OH Me

COOMe

114

Coryzeylamine

COOMe CH2 OH Me

115

Def ormylcoryzeylamine

protonization. However, the different transformations are all in accord with the acid-catalyzed opening of the cyclotryptamine ring by cleavage of the C-8a - N-1 bond succeeded by various rearrangement reactions resulting in reestablishment of the indole nucleus. These observations are of interest for two reasons, one of which being that the acid-catalyzed reactions mimic the reverse transformations of those usually believed to take place during the biogenesis of these compounds, thus lending some support to the notion that many of these reactions are in

Naturally Occurring Cyclotryptophans and Cyclotryptamines

199

principle reversible transformations. The second reason is that many of the cyclotryptamines described from higher plants have actually been exposed to quite drastic conditions during isolation and purification, including elevated temperatures for extended periods of time, and strong basic reactions followed by acidification with strong mineral acids. Considering these conditions it is likely that some of the structures reported are actually artifacts. According to the evidence presented [251] it could be argued that the cyclotryptamines are more numerous than anticipated at present. The stereochemistry of 96, 97, 98 and 100 have been studied by ^H NMR spectroscopy [252]. Behavioral studies of alkaloids extracted from the leaves of H. zeylanica have been reported [253].

COOMe

y^^Q

COOMe

COOMe MeO

MeO

116

Flexicorine

117

Cabufiline

Rauwolfia The dark-red bis-indole alkaloid flexicorine (116) originatesfromthe leaves oiR, reflexa and is unusual by being most stable in air in the oxidized iminoquinone form rather than as the reduced hydroxyindoline [254]. Originally the stereochemistry of the methoxycarbonylbearing carbon atom was believed to be the opposite to that found in the other members of this class of compounds. However, this assignment rests on an erroneous assignment of the ^^C NMR signals. From extracts of the leaves of R. sumatrana collected in Thailand 116 was isolated together with three other bis-indole alkaloids, cabufiline (117), rausutrine (118) and (probably the biogenetically derived) rausutranine (119) [255]. The two latter also are iminoquinones composed of an akuammilane unit and a vincorane unit. In the latter study the stereochemistry of these alkaloids was firmly established.

U. Anthoni, C. Christophersen and P. H. Nielsen

200

H-.^COOMe

r^ 0.

H

COOMe

COOMe MeO

MeO

118

Rausutrine

COOMe

120

Vincoridine

119

COOMe

MeO

121 122

Rausutramine

R=H R=Me

Norvincorine Vincorine=Vincovine

Vina (Catharanthus) The Vinca or, systematically more correct, Catharanthus alkaloids have been reviewed [256]. All cyclotryptamines were isolated from the leaves of the lesser periwinkle V. minor, Vincoridine (120, stereochemistry assigned in analogy with that of related compounds), norvincorine (121) and vincorine (122) are monomeric alkaloids with only little cytotoxic activity [257-260,258 features a misprinted formula]. Vincarubine (123) from the dried leaves is a dark red bis-indole iminoquinone alkaloid chemically related to 116, 118, and 119 [261, 262]. Probably because of the quinoneimine structure it has a significant antitumor activity but at the same time without any mutagenic effect [262]. It is isolated from V. minor and the content can be determined easily by HPLC [263]. During the structural elucidation of vincarubine (123) reduction of the quinoneimine to the corresponding aminophenol followed by O-acetylation resulted in isolation of two very closely related isomers [262, two structural formulas misprinted]. The same type of isomers were isolated in the case of norvincorme

201

Naturally Occurring Cyclotryptophans and Cyclotryptamines

H-^COOMe MeCH

124 125

Ri =R2 =OMe Ri =OMe, R2=H

Alstonamide Demethoxyalstonamide

(121). The isomers were believed to differ in the conformation of the seven-membered azepine ring encompassing the two methylene groups connected to the cyclotryptamine 8a position (E ring), the carbocyclic six-membered ring (D ring) and the eight-membered ring containing the cyclotryptamine N-l. Based on NMR experiments it was argued that one isomer has the D, E and eight-membered rings in the boat conformation while these conformations are changed to the twisted boat, twisted chair and chair forms, respectively. Isolation of conformers as stable molecules is an extremely rare phenomenon. Since there are other possibilities for isomerism in these structures, a reinvestigation is called for. Alstonia Since extracts of Alstonia species have been used for many years for medical purposes, the alkaloids have been repeatedly reviewed covering the literature until 1973 and only recent reports concerning cyclotryptophans are mentioned here [1, 264-266]. Alstonamide (124) and demethoxyalstonamide (125) co-occur with 122 ini4. macrophylla from Sri Lanka [267, 268]. Pleiocorine (126) and 122 are known from A. deplanchei [269]. A. odontophora, in addition to 126 contains N(r)-demethylpleiocorine (127) [270]. From A. plumosa was isolated 126, the dimeric desoxycabufiline (128), and nordesoxycabufiline (129) [271]. The two latter alkaloids are also found with 120 in A. sphaerocapitata [111]. From A. ondulata 128 and 122 are known [273]. In A. vitiensis var. novo ebudica monachino, a large tree from the New Hebrides, 122

202

U. Anthoni, C. Christophersen and P. H. Nielsen

COOMe COOMe

^-^

--

126

R=Me

Pleicorine

127

R=H

yV-Demethylpleicorine

COOMe MeOOC.

128

R=R'=Me

129

R=Me , R'=H or R=H, R'=Me

Desoxycabufiline Nordesoxycabufiline

was the only cyclotryptamine present [274]. The dried, ground leaves of A. pittieri were investigated in order to clarify the botanical classification [275]. Among the nine alkaloids identified were 117, vincorine (122) and three dimeric vincorine derivatives 130 - 132. Compounds 130 and 131 are isomers differing only in the epimeric hydroxy groups, and 132 is the acetate of 131. The alkaloids all belonged to the first type of the Le Men-Taylor chemotaxonomic classification expected for Alstonia. The authors concluded that the separation of the genera Tonduzia and Alstonia could not be justified from composition of the alkaloid content. A large group of Alstonia species, including A, angustifolia from Indonesia [276] and A. undulifolia from the Malay Peninsula [277], contains the cyclotryptamines echitamine (133) and norechitamine or Nb-demethylechitamine (135). These two alkaloids are also typical of ^4. scholaris collected in Thailand [278, 279]. The corresponding Phillippine species have yielded 133, 135, and 17-acetoxynorechitamme (136) [280] as has the African A. congensis closely related to A, scholaris [281]. From an ethanolic extract of the roots of the Chinese Winchia

Naturally Occurring Cyclotryptophans and Cyclotryptamines

MeOOC

203

Ri

130

Ri=H, R2=0H

131

Ri=OH. R2=H

132

Ri=OOCMe, R2=H

HOCH

R0CH2^_^C00Me

133 R=COOMe Echitamine

135 R=H

134 R=COCr

136 R=MeCO 17-0-Acetylnorechitamine

Echitaminic acid

Norechitamine

H0CH2,.._^C00Me

137

Norechitamine A/^oxide

calophylla (syn. Alstonia pachycharra) 133, 135, and norechitamine A^oxide (137) were isolated [282]. A recent investigation of the stem bark of ^4. glaucescens showed the presence of all the five known echitamine derivatives, 133, echitaminic acid (134), 135, 136 and 137 [283]. Some of these alkaloids are closely related to the Hunteria alkaloids, e.g. norechitamine (135) is Ar-demethyl-3-^/?i-dihydrocorymine (demethyl 110).

204

U. Anthoni, C. Christophersen and P. H. Nielsen

The metabolism [284] and pharmacology [285, 286] of 133 have been studied. Moreover, the possibility of using 133 as antitumor agent [287, 288], for inhibition of HIV reverse transcriptases [289], or in malaria treatment [290] have been investigated.

H^>^COOMe

"JXXX

N^

MeOOC 138

R=Me

Peceyline

139

R=H

Demethylpeceyline

J ''^

"-^axn

H^.^COOMe

MeOOC

l4

140

Pelankine

Petchia P. ceylanica is an endemic species found in the lowlands of Sri Lanka. In addition to vincorine (122) the leaves has yielded five dimeric indole alkaloids, peceyline (138), demethylpeceyline (139), pelankine (140), peceylanine (141), and ceylanine (142) [291-294]. From an ethanolic extract of the air dried stem ceylanicine (143) was isolated [294]. In the case of 138, 140 and 141 the position of the ethylidene and epoxy groups were not unambiguously determined and they could be interchanged. However, since the structure of 139 was determined as depicted, the remaining alkaloids presumably are epoxydated in the positions indicated. Cabucala Several indole alkaloids have been isolated from C. erythrocarpa var. erythrocarpa) including the cyclotryptamines cabuamine (144). Gentle heating of an alkaline solution of 144

205

Naturally Occurring Cyclotryptophans and Cyclotryptamines

COOMe MeOOC

H...^^COOMe

gives rise to the retro-aldol product desformocabuamine identical with vincovine and vincorine (122) [295, 297 (wrong structures)] also present in the plant material. Interestingly, in this series of compounds the formation of the pyrrolidine ring seems to occur under gentle chemical conditions from the corresponding jS-carbolines with an indoline C=N. Thus treatment of 10-methoxy-akuammilin (from Vinca minor L.) with Zn/HCl results in a fair yield of the 0-acetyl derivative of A/-demethylcabuamine [297] and analogously the same treatment of strictamine gave demethoxy-A'-demethylvincovine [298]. Admittedly the two latter cyclotryptamine structures rest exclusively on mass spectrometry and the occurrence of hypsochromic shifts in the UV spectra on acidification. Since the plants containing cyclotryptamines usually have jS-carbolines it is tempting to ponder the potential connection between the biogenetic pathways leading to the two types of compounds. From the leaves of C. caudata, also known from Rauwolfia sumatrana and Alstonia pittieri. 111 has been identified [299].

206

U. Anthoni, C. Christophersen and P. H. Nielsen

143

Ceylanicine

ROCH2

cOOMe

1 Me ^

144 Cabuamine

145 R=MeCO 146 R=H

Tacraline Desacetyltacraline

Tabemaemontana Among forty-five alkaloids isolated from the root bark of the small tree Tabemaemontana chippii (syn. Conopharyngia chippii)fromthe Ivory Coast, the trace components tacraline and desacetyltacraline tentatively formulated as 145 and 146 were characterized by chromogenic reactions, MS, and ^H NMR spectra. However, due to the small amount of material available they were not conclusively identified [300]. Curiously enough, the published ^H NMR data, except for the signals from the NH protons, do not show signals in the 6 4-5 range expected for the methine proton of the doubly substituted nitrogen position. Moreover, it is possible that the alkaloids are artifacts arising during the treatment of the crude mixture of alkaloids with ammonia. A reinvestigation is warranted. Rhazya, Aspidosperma, Gonioma From the alkaloid fractions of Rhazya stricta, Aspidosperma quebracho bianco or Gonioma kamassi either the quaternary cyclotryptamine rhazidine (147) or the precursor rhazidigenine was isolated. It seems clear that 147 is not an artifact formed during isolation, and since the two compounds are in tautomeric equilibrium it becomes irrelevant to discuss

207

Naturally Occurring Cyclotryptophans and Cyclotryptamines

147

Rhazidigenine

Rhazidine

which represents the native alkaloid [301]. The tautomerization process can conveniently be followed by optical rotation. In neutral or acidic ethanol a low rotation is observed ([aJo -37°), approaching -612° on addition of strong base and again returning to the low negative value on re-acidification. The relative configuration around the newly formed ring junction is presumably cis, however the absolute stereochemistry is still undecided [302].

148

Minfiensine

5.8 Loganiaceae. Although alkaloids are very common in the Loganiaceae, minfiensine (148) from the African Strychnos minfiensis, is the only example of a cyclotryptamine from this source [303].

6. ANIMALS 6.1 Porifera Phylum Porifera constitute the most primitive of the multicellular animals. Most chemical investigations deal with members of the class Demospongiae constituting 95% of all sponge species. Cyclopeptides isolated from lithistid sponges have been suggested as originating with filamentous microorganisms (presumably bacteria or cyanobacteria) since such organisms were consistently found in large quantities in the sponge tissues [304]. It is remarkable that in spite of the enormous diversity of secondary metabolites isolated from sponges the two cyclopeptides

U. Anthoni, C. Christophersen and P. H. Nielsen

208

described below are, so far, the only examples of cyclotryptophans known from these animals while a wealth of other cyclic, highly bioactive, cyclopeptides are known [305]. This indicates that these two compounds might be procaryotic metabolites since the cyanobacteria have the genetic setup necessary for transcription of the multienzyme peptide synthetase complexes. However, sponges are known to produce cyclopeptides of the same general type as those originating from the synthetases. Cyclocinamide-A is an unusual halogenated cyclic hexapeptide isolated from the marine sponge Psammocinia sp. [306]. There is at present no available information to decide the origin of the peptides.

149 150

R=(R)-OH R=(S)-OH

Phakellistatin 3 Isophakellistatin 3

Phakellia So far the only examples of cyclotryptophans from a marine sponge are phakellistatin 3 (149, yield 2 10"^% of wet weight) and isophakellistatin 3 (150, 1.3 10-^%) from Phakellia carted (order Axenellida, class Demospongia) [307]. The two cycto-heptapeptide alkaloids are diastereomeric and both have the dj-annelated cyclotryptophan structural unit. Phakellistatin 3 exhibits significant activity against the murine P388 lymphocytic leukemia cell line (ED50 0.33 /ig/mL) while 150, in spite of the only minor stereochemical difference, was inactive in this assay. The structure of 150 was established by an X-ray crystal structure determination. The cyclotryptophan moiety was referred to as photo-Trp indirectly implying that the two peptide alkaloids might be formed in a photochemical reaction. However, it would be more likely, if they are artifacts, that they were formed from the tryptophan precursor by reaction with singlet oxygen or peroxide.

209

Naturally Occurring Cyclotryptophans and Cyclotryptamines

151 X=H2 Flustramine A

153 R=Br, X=H2 Flustramine B

152 X=0

154 R=H, X=H2

Debromoflustramine 8

155 R=Br, X=0

Flustramide B

Flustramide A

J

r^

Br-V^?V Me 156

Flustramine C

sX^

N ^

158

B r - ^ * * ^HH

I Me

157

Flustramine E

159

Flustraminol B

OH )

'N Me

Flustraminol A

6.2 Bryozoa Phylum Bryozoa (Polyzoa, Ectoprocta or moss animals) is one of the major (around 4000 extant species) animal phyla. They are colonial sessile animals. The chemistry of this phylum is but little investigated. In the genus Flustra all cyclotryptamines are simple N^methylated mono- or diprenylated derivatives characterized with one exception by the bromosubstitution in the tryptophan 6-position. The enzymatic requirements of these transformations are not known but prenyltransferases are well known in other connections [308, 309]. In the case of the closely related genus Securiflustra the alkaloids are characterized

210

U. Anthoni, C. Christophersen and P. H. Nielsen

by being monoprenylated dipeptides derived from cyclotryptamin and histidine. Flustra The series of indole alkaloids including many bromocyclotryptamines known from the marine bryozoan F, foliacea have been reviewed several times [310-316]. Colonies collected in the North Sea have given rise to the isolation and structure elucidation of the flustramines A (151) [317, 318], B (153) [317, 318], C (156) [319], and E (157) [320], the debromoflustramine B (154) [320], the flustramides A (152) [321] and B (155) [322], and the flustraminols A (158) and B (159) [319]. The same species collected from Nova Scotia, Canada, gave instead dihydroflustramine C (160), dihydrpflustramine C A/-oxide (163), flustramine D (161), flustramine D A/-oxide (164), and isoflustramine D (162) [323, 324]. Since so far not a single cyclotryptamine has been found to co-occur in the two populations it is tempting to speculate that the metabolites, in fact, originate from different populations of associated microorganisms. This speculation is supported by circumstantial evidence since many bryozoan species have been found to be obligately associated with, what seems to be, species specific bacteria [325]. In the case of F. foliaceae and Securiflustra securifrons a small green alga, Epicladiaflustrae, has been found only in these bryozoans [326]. Furthermore, in accordance with the theory of adaptive variance the metabolite profile should be optimized to give maximal protection against different local predators [327, 328]. Available information on the antibacterial activity of these cyclotryptamines lends some support to this point of view [320]. In addition the Flustra alkaloids have been implicated in the chemical defence of this organism, mainly because of their larvotoxic activity [329-331]. Flustramine A (151) and B (153) exhibit muscle relaxant activity against skeletal as well as smooth muscles both in vitro and in vivo [332].

- Cri Br

Flustramine B /V-oxide

Flustrarine B

By analogy with the findings reported for the acid-base catalyzed equilibrium between geneserine - physostigmine A^-I-oxide and geneseroline - geneseroline A^-1-oxide [191], flustrarine B (hexahydro-l,2-oxazino[6,5-&]indole) participates in an equilibrium with the corresponding hexahydropyrrolo[2,3-ft]indole A^-1-oxide [333]. Also in this case the transformation is acid - base dependent, the A^-oxide being the structure present in acid and the oxazine the isomer present in basic solution. The two isomers have the 15, 3a5, 8aiS

211

Naturally Occurring Cyclotryptophans and Cyclotryptamines

R, ^2

"^M".L^oe

160 Ri=R2=H Dihydroflustramine C 163 A/-oxide of 160 161 Ri=H, R2 =CH2 CH=CMe2 Flustramine D 164 /V-oxide of 161 162 R2=H, Ri=CH2CH=CMe2 Isoflustramine D

165 166

R=H Securamine A R=Br Securamine B

R=H Securine A R=Br Securine B

BrA^N-f-K|J

167 168 169 170

R,=Br, R2=R3=H, R4=CI Ri=R2=R3=H, R4=CI Ri=R2=Br, R3=H, R4=CI Ri=R2=Br, R3=CI. R4=H

Securonnine C Securamine D Securamine E Securamine F

171

Securamine G

configurations. It would be interesting to know if a comparable tautomeric equilibrium exist with the two flustramine A/-oxides, 163 and 164. Flustramine A and B exhibit muscle-relaxant activity in vitro as well as in vivo and both

212

U. Anthoni, C. Christophersen and P. H. Nielsen

skeletal and smooth muscle are affected [334]. The alkaloids are of quite low acute toxicity and show a minimum lethal dose of 500 mg/kg in mice. The preparation of racemic 153 [335], racemic 154 [336, 337], racemic flustramine E [337] , racemic debromoflustramide B and E [337], optically active 154 [338], and racemic 156 [339] have all been reported. Debromo-8,8a-dihydroflustramme C has been synthesized [340, 341]. Securiflustra From another marine bryozoan, S. securifrons, originate the imidazole alkaloids securamines A (165) and B (166) [342]. When dissolved in DMSO a reversible ring opening occurs to give the corresponding securines A and B. Furthermore, the securamines C (167), D (168), E (169), F (170), and G (171) were isolated, derived from the former group of metabolites by formation of an additional C-N bond [342, 343]. Interestingly, a closely related genus, Chartella, of marine bryozoans elaborate indole alkaloids built from tryptophan, histidine and an isoprene unit as well [344]. The latter genus seems not to be capable of closing the cyclotryptamine skeleton and consequently may not possess the modular enzymatic unit needed for affecting this transformation.

Me

Y "Me 0

~)

172 R=(SVMeCH2

Urochordamine A

173 R=(R)-MeCH2

Urochordamine B

6.3 Ascidiacea The ascidians or tunicates (sea squirts) belong in class Ascidiaceae in subphyllum Urochordata, one of the two subphyla of phylum Chordata. They only possess a notochord in the larval state. Several interestmg biological active cyclopeptides {e.g, the didemnins) have emerged from members of this class [345, 346] attesting to the prescence of peptide synthetases in the subphylum (or associated symbionts). Urochordamines A (172) and B (173) result from a bioassay-guided isolation from the ascidians Ciona savignyi (the tunic part, yield 3.7 lO""^ and 1.2 10"*% of wet weight.

213

Naturally Occurring Cyclotryptophans and Cyclotryptamines

respectively) and a colonial Botrylloides sp. (whole body, yield 6.8 10"* and 6.0 W^% of wet weight, respectively) [347]. The names were coined after the subphylum, Urochordata, to which the tunicates belong. The compounds rearrange in the pteridine moiety in protic solvents to urochordamine A' and B' where the =NH group at the 2-position is exchanged with the 3 NMe- group [348]. Formulas 172 and 173 are only intended to depict relative stereochemistry since the absolute configurations are unknown. All alkaloids promoted ascidian larval settlement and metamorphosis and the order of activity with larvae of the ascidian Halocynthia roretzi was A' > A > B > B' and induced metamorphosis of the pediveliger larvae of the mussel Mytilus edulis galloprovencialis. These and other results imply that urochordamines do not act via a chemoreceptor but via an internal mechanism. The natural products, urochordamines A and B, exhibited activity against the Gram-positive bacterium Bacillus marinus but not against the fungus Mortierella ramaniana or the Gram-negative Pseudomonas nautica or Flavobacterium marinotipycum.

Me

c

r

N ^

I Me

Hf4 I Me 174 c/-Chimonanthine

175 R=CH2 0H

Pseudophrynaminol

176 R=COOMe 177 R=CHO Since the bromocyclotryptamines 151 - 171 are characteristic of bryozoans while C6substituted pteridines have been reported from different marine organisms (sponges, anthozoans and polychaetes), it is proposed [347] that microorganisms play a role in the biosynthesis of the urochordamines. 6.4 Amphibia Pseudophryne Members of the Australian myobatrachid frog genus Pseudophryne have yielded a series of indole alkaloids with the cyclotryptamine skeleton. The chemistry, pharmacology, and biological role of the amphibian alkaloids have been thoroughly reviewed and discussed recently [349, 350].

214

U. Anthoni, C. Christophersen and P. H. Nielsen

I I R2 Me

N' I Me

I I Me Rj 181

178 Ri =R2 =R3 =H Pseudophrynomine A 179 R,=OH. I^=R3=H 180 Ri=R2=H, R3=Me or Ri=R3=H, R2=Me

y

0^.^. RO^^^N^

COOMe

0

.-^^^.

R2-

H

1

182 R=HMe 183 R=Me

y

COOMe

184 185 186 187

1 Me

Ri=H, R2=0H R,=H, R2=0Me R,=OH, R2=0Me Ri=R2 = OMe

The dendrobatid Colombian poison-dart frog Phyllobates terribilis have given rise to the isolation of t/-chimonanthine (174) and the acidic rearrangement product /-calycanthine, both enantiomeric to the compounds found in plants (e.g. 71 obtained from Calycanthus) [351]. This observation seems to exclude the possibility that the alkaloids are of dietary origin and accumulated in the skin of P. terribilis. The skin of the burrowing frog Pseudophryne coriaceae contains three indole alkaloids, namely pseudophrynaminol (175), pseudophrynamine A (178), and the methyl ester (176) corresponding to the acid present in 178 [350]. The latter may be an artifact. In an examination of seven species of Pseudophryne the indole alkaloids were detected in all members [353] and the new cyclotryptamines 177 and 179 - 187 (all structures are tentative) were identified. The total synthesis of (±)-175 [354], (-)-175 [355], (+)-(ent)-175 [356] and (±)-178 [357] have been reported. The primary role for these alkaloids is considered defense against predation and against skin infections [349].

Naturally Occurring Cyclotryptophans and Cyclotryptamines

MeO

215

H

XX

^

"*"~^COM, 188

Cyclic 2-hydroxymelatonin

6.5 Mammalia Man and rat The cyclic isomer of 2-hydroxymelatonin (188) was isolated from urine following administration of synthetic melatonin to rat and man [358]. It was present in unconjugated form as a minor (5%) excretion product together with the main urinary metabolite 6hydroxymelatonin present as the sulfate and the glucoronide conjugates. The structure was inferred based on ^H NMR (400 Mhz) and MS (high resolution) studies. The structure is of considerable interest since it is highly unusual and the only example of a l,2,3,3a,8,8ahexahydro-8a-hydroxypyrrolo[2,3-^]indole (8a-hydroxycyclotryptamin). There are several equivocal points in the published ^H NMR data of the compound. A peculiar trait is the presence of an intense molecular ion at m/z 248 in the EI (70 eV) mass spectrum even though the sample was introduced around 200°C, obviously without any appreciable thermal or mass spectrometric loss of water. Unfortunately nothing is known about the stereochemistry of the compound. If H-3a and the 8a-hydroxy group are cw-oriented, the groups may not be able to adopt the anti-periplanar transition state necessary for water elimination. In view of these peculiarities an alternative structure cannot at present be completely ruled out. If, for example, the initial metabolic reaction was an AT-oxidation of N-l then the A/-oxide might rearrange to the corresponding six-membered hexahydro-l,2-oxazino[5,6-&]indole skeleton like in the cases of flustrarine B [333], geneserine and geneseroline [191]. Flustrarine B shows a loss of C2H5NO occurring with 100% relative intensity from the molecular ion which is also observed (50%) in the cyclic 2-hydroxymelatonin. Clearly the whole matter calls for a reinvestigation including a synthetic approach to this kind of cyclotryptamines. The biological activity and physiological function, if any, is unknown.

7. BIOSYNTHESIS

Based upon the available evidence and in accordance with biomimetic syntheses [5], biogenesis of cyclotryptophans/cyclotryptamines may proceed from tryptophan/tryptamine via the scheme outlined below. Following eventual derivatization of the side chain, the 3-position

216

U. Anthoni, C. Christophersen and P. H. Nielsen

W

NH2 COOH

Ocji H' NH. Tryptamine

Tryptophan

1 NH V

H

COs, l+RX ^ ^

H

CO\

+RX

I I CO.,

I

NH

i-

R H

H

I

Cyclotryptophan

CIS? H

I

H

I C0>

t

Cyclotryptamine

in the indole ring is substituted with RX (R = hydroxyl, alkyl or hydrogen) to give the indolenine in equilibrium with the corresponding salt. In the final step cyclization occurs by electrophilic attack of the side chain nitrogen. Regulation of biosynthesis and metabolism has only been studied in some detail for the production of roquefortine (8) from tryptophan in Penicillium species [359-363]. The available evidence has recently been discussed [364] and it is concluded that the alkaloids in this species participate actively in metabolism dependent on the physiological state of the fungus. For example, it is known that 8 is an intermediate in the biosynthesis of oxaline in cultures of P.

217

Naturally Occurring Cyclotryptophans and Cyclotryptamines

oxalicum [365]. The amount of 8 present is not only influenced by transport, cultivation, and excretion but also controlled by the concentrations of precursors and end products. The biogenesis of 8 involves the building blocks of tryptophan, histidine, and mevalonate. This has been demonstrated by feeding P. roqueforti [l-^'^Clmevalonic acid lactone, [methylene-^'^Cltryptophan, and [l-^'^Qhistidine resulting in 0.08, 0.15, and 1.12% incorporation, respectively [366]. The specific activity of the resulting radioactive 8 was lower than the simultaneously formed roquefortine D (10) suggesting, that 8 is formed via 10. However, this has never been corroborated and other evidence although not unambiguous indicates that the formation of 8 proceeds according to the general scheme. This means that tryptophan is initially derivatized in the side chain with formation of the diketopiperazine. Prenylation in the 3 position forms the indolenine intermediate which easily cyclizes by addition of amide nitrogen to the CN double bond. Ring closure of the open indolenine form undoubtedly has a very low activation energy. Reversible equilibria have been found in several cases, cf. the securamines (165 and 166) and rhazidine (147) mentioned above. The open indolenine form has been reported on feeding 8 to Penicillium. Using ^^C-labelled precursors the prenyl group was shown to arises from acetate and mevalonate [367, 368]. Several mechanisms are possible for the introduction of the inverted isoprene moiety at position 3 in the indole nucleus. The pathway may encompass an aza-

V^

GOGH

Tryptophan-ds

r^

r^

V-N^^MffY^

V,

N

Roquefortine-d 5

U. Anthoni, C. Christophersen and P. H. Nielsen

218

Claisen-type rearrangement of an initially formed l-(7,7-dimethylallyl) substituent to an inverted isoprene at position 2, direct alkylation at this position followed by rearrangement of the 2 substituent to the 3 position, or direct alkylation of the 3 position. Bhat et al. found that L-[2,4,5,6,7-4]tryptophan was incorporated into 8 (as well as into 41 from Aspergillus zonatus) with retention of all five deuterium atoms as shown on the scheme [369, 370] in contrast to an earlier report [371]. This result shows that the 2 position cannot be involved in the alkylation reaction. N-prenylation is improbable since the tritiated precursor was not incorporated in related fungal metabolites [341] leaving direct prenylation of the 3 position involving a prenyltransferase as the most attractive mechanism [373]. 8 has the E configuration around the dehydrohistidine unit. By incorporation of (25,35)- and (25,3/?)-[3-^H]histidine into the alkaloids in each case ihtpro-S hydrogen atom was shown to be eliminated in the dehydrogenation reaction [374]. The biogenesis of several 3-hydroxycyclotryptophanes (R = OH in the general scheme) has been investigated. Tritiated tryptophan was incorporated in Sporidesmin A (32) from Pithomyces chartamm confirming that 32 is derived biosynthetically from tryptophan, alanine and methionine [375]. The 3/?- and 35-forms of [3-^H]-tryptophan were used to show that side chain hydroxylation takes place with retention of configuration at the side of attack. However, the individual steps in the biosynthetic pathway were not investigated.

Oo B

(fH, COOH

Tryptophan

Asperlicin C

Asperlicin E

Naturally Occurring Cyclotryptophans and Cyclotryptamines

219

Later, work incorporating ^"^C-labelled amino acids in asperlicin E (48) from Aspergillus alliaceus established that this is biosynthesized from tryptophan and two anthranilate moieties [376]. Initially the side chain is derivatized to the benzodiazepine asperlicin C, which can be isolated and characterized. Following the metabolism of tritiated tryptophan it was shown that asperlicin C was further hydroxylated and cyclized to 48. Using 5-fluorotryptophan a series of fluorinated analogs of 48 were produced showing that the existing biosynthetic machinery in A. alliaceus is not specific for tryptophan. Such directed biosyntheses of cyclotryptophane analogs can be very useful in studies of receptor binding. Recently, 36 biosynthetic analogs of the dimeric WIN 64821 (51) were produced by feeding analogs of phenylalanine, tryptophan and other amino acids to intact cells of an Aspergillus sp. [377]. It is tempting to speculate whether the same or closely related enzymes are used for all cyclisations of cyclotryptophans and whether cyclization always is the final step in the biosyntheses. This would be in accordance with the result, that cyclisation is performed with an enzyme with little specificity. In the case of the aspercilins it was not possible to establish the succession of the final steps, hydroxylation and cyclisation. However, by using ^H and ^"^C-labelled precursors and analyzing the possible biosynthetic pathways for the closely related brevianamides from Penicillium brevicompactum, it was possible to conclude that hydroxylation precedes cyclisation in production of brevianamide E (14) [378]. This result validates the proposed general biogenetic pathway for hydroxy lated cyclotryptophans. Since biogenesis of both prenylated and hydroxylated cyclotryptophans fit the same general scheme it would be expected to apply also in other cases including unsubstituted cyclotryptophans, i.e. R = H. Such a mechanism is well known from related cases, e.g. from reactions of the indole ring in tryptophan synthetase [379]. The enzymatic proton transfer to the 3 position occurs from a COOH group located appropriately and the ring closure may occur concerted or in a separate step. In biomimetic syntheses proton transfer always occur in a stepwise process [5]. Very little information is available regarding the related formation of cyclotryptamines but as far as known the general scheme is followed. In Chimonanthus fragrans 2-tritiated (and ^'^C labelled) tryptophan and tryptamine are both converted into the appropriately labelled chimonanthine (71) establishing the sequence tryptophan - > tryptamine - > 71. However, it could not be confirmed with certainty that tryptamine was methylated prior to cyclisation [380]. Interestingly, the producer of the protein kinase C inhibitor staurosporine 5. staurosporeus is capable of transforming added tryptamine to (3a/?,8a5)-l-acetyl-3a-hydroxyl,2,3,3a,8,8a-hexahydropyrrolo[2,3-Z?]indole in 8% yield [381, 382]. The first reaction can be concluded to be a selective N^acetylation triggered by an acyl transferase while it was not possible to decide whether hydroxylation preceded cyclisation or vice versa. Analogous results were obtained for 5- and 6-fluorotryptamine indicating that the cyclisation step is not entirely specific. Tryptophan was not metabolized to the cyclized derivative but yielded anthranilic acid (o-aminobenzoic acid) as the major product (40%) in addition to 3-(hydroxyacetyl)indole (4%) and indole-3-carbaldehyde (1 %). In this connection it should be borne in mind that anthranilic acid is not only a precursor for tryptophan in the chorismate pathway but also a product in

220

U. Anthoni, C. Christophersen and P. H. Nielsen

OH

9

NHa

^^

N

NH

^

COMe Tryptamine

/V-Acetyltryptamine

^H^ COMe

Hydroxyacetylcyclotryptamine

tryptophan catabolism formed by the action of kynureninase on kynurenine produced enzymatically from A^-formylkynurenine, the initial product from the action of tryptophan oxygenase on tryptophan.

8. CYCLIZATION IN PEPTIDES AND PROTEINS? Tryptophan is an exceptional protein amino acid in almost every aspect. Together with methionine tryptophan is the only protein amino acid without degenerate codon. Together with methionine and threonine it is coded with a different codon in mitochondria. Tryptophan is the protein amino acid with: The highest molecular mass (186 g/mol), the largest volume (227.8 A^), the largest free surface area (255A^) defined as the part of the surface area that is available for contact with a spheric probe such as a water molecule, the lowest procentual occurrence (1.1%) in proteins, the highest index of hydrophobicity (7r=2.14) for the indole group (7r=log(P/Po), where P is the partition coefficient between n-octanol and water and Po is the same figure for the compound where the indole group is replaced with -H), the lowest relative mutability meaning that tryptophan is the amino acid most often conserved in evolution of proteins. The physico-chemical and biochemical singular character of tryptophan is matched with the seemingly inexhaustive plethora of chemical reactions in which it participates. In spite of the uniqueness the function of tryptophan is most often unaccounted for; it is known only that it is frequently essential for proper function of the proteins of which it is part. Taking into account the myriad of chemical transformations the tryptophan molecule and especially the indole nucleus engage in under gentle (physiological) conditions, it is almost inevitable that tryptophan in peptides and proteins undergoes purely chemical transformations in the living organism. It is, for example, well known that tryptophan and derivatives react smoothly with singlet oxygen to form the 3a-hydroperoxylated cyclotryptophans which are easily reduced to the corresponding 3a-hydroxycyclotryptophan [5, 383]. The same compound was identified on 7-radiolysis [384], on reaction with superoxide ion [385], peroxyacetic acid [386] and on electrochemical oxidation [387]. Also dipeptides have been shown to react analogously [388].

Naturally Occurring Cyclotryptophans and Cyclotryptamines

221

8.1 Potential role of cyclotryptophans in senescence and aging

OH

UCT^yO

Nu',e

0 ^

0^/JV The 3a-hydroxycyclotryptophans react readily with nucleophiles such as cysteine with the sulfur attacking the 8a-position with ring opening and elimination of water to form the 2thiosubstituted tryptophan [386, 389]. Analogous reactions can be carried out using A'^bromosuccinimide or tert-b\iXy\ hypochlorite as oxidation reagent [390]. Wieland and coworkers have utilized this reaction ingeniously to prepare several synthetic phallotoxins [391-394]. Other examples are known as well (e.g. 395). Since all organisms endure free radicals [396], singlet oxygen, superoxide ion and hydrogenperoxide this reaction must occur in the living organism as well. If a cysteine residue in the same polypeptide reacts a cyclic artefact is formed. Cysteine from another peptide (or another molecule containing nucleophilic groups) will give rise to a junction of the two molecules. A transformation of this kind would undoubtedly mark the artefact for hnmediate removal by the proteolytic machinery of the cell. However, owing to the covalent bond formed to the tryptophan 2-position the complex would presumably not be degradable by the ordinary proteolytic enzymes. It is now a question to what degree the molecular shape would have to be changed in order to evoke a responce from the immune system. If the complex is detected and deemed foreign by the immune system a response would be initiated and an autoimmune reaction ensue. This is exactly what seems to happen when an organism ages. The number of autoimmune diseases accelerate [397]. In addition oxygen free radicals have been implicated in a long list of deseases ranging from cancer over various tissue injuries to inflammatory deseases [398]. It is interesting to note that the age pigment, lipofuscin is generated by peroxidation [399] and thus could very well encompass structures discussed above. In addition it has been shown [400] that this pigment is accumulated through the life span of an organism and thus can serve as a measure of the chronological age. This result signifies that the fuscin complexes are not easily removed by the enzymatic machinary of the intact cell. There is an overwhelming literature concerned with the biochemistry of senescence

222

U. Anthoni, C. Christophersen and P. H. Nielsen

(including many international journals). It is outside the scope of the present review to analyse the multitude of studies where the above presented ideas may explain the often extensive experimental results. In the case of the 3a-hydroxy derivatives the reaction with nucleophiles became irreversible by the ensuing elimination of water. The 3a-hydrogenated or alkylated cyclotryptamines participate in a comparable reaction with nucleophiles only without the elimination step. In these cases the reaction consequently becomes reversible. When physostigmine, 2, was treated with sodium bisulfite in aqueous solution at pH 5.5 the reversible ring opening was monitored by spectroscopic and optical activity measurements [401]. Changing the pH to 9.5 and extraction with heptane served to recover physostigmine with the original configuration. This is but one documented example on the existence of the easily established ring-chain equilibrium between certain cyclotryptamines and the corresponding tryptamines. In this case the transformation is brought around by an exterior nucleophile (bisulfite) whereas often the nucleophile is inherent in the alkaloid structure and the equilibrium then becomes an example of tautomerism. Examples of the latter type are the equilibria between rhazidigenine/rhazidine (147) and securamines A and B/securines A and B (165, 166).

H MeN

Y' 0

Me HSOa^^

k^N'i-N'J I HI

Me Me

8.2 Acetylcholine esterase This enzyme which functions to remove, by hydrolysis, the neurotransmitter acetylcholine in the synapses has been intensively studied and the structure solved to 2.8 A resolution [402]. Macroscopically it consists of a large protein forming a deep narrow tunnel leading to the active site. The tunnel is lined with 14 conserved aromatic residues. The highly

Naturally Occurring Cyclotryptophans and Cyclotryptamines

223

conserved tryptophan-84 near the active site functions to bind by x-cation interaction the substrate [403]. Since the rate of hydrolysis of the substrate is near the diffusion controlled limit the products cannot be transported backwards through the gorge but instead are released through the "bottom" of the enzyme. At the crucial area of the end of the tunnel tryptophan-84 is situated functioning like a cork. Investigations utilizing the Cambridge protein structural database revealed that this tryptophan residue had the peptide nitrogen of the tryptophan CONH group placed within the Van der Walls distance from the indole 2-position. In other words it is ideally situated for the ring closure reaction forming cyclotryptophan at this position. Molecular modelling experiments (T Fatum, F Jorgensen, U Anthoni, PH Nielsen and C Christophersen, unpublished results) revealed that the transformation of this tryptophan into the corresponding cyclotryptophan resulted in an opening of the tunnel bottom allowing the products free passage out of the enzyme. We submit that even if there has been no formal documentation of this mechanism it should be investigated further. Cursory investigations of protein structures exposed many structures where the position of tryptophan residues had a spatial arrangement like the one described for the tryptophan-84 above. It is thus plausible that analogous entropy facilitated reactions are of common occurrence in proteins. 8.3 Conceivable regulatory functions

NH

NH " 0 Kapakahine B A priori, any tryptophan unit in a peptide chain has the possibility of forming a macrocyclic cyclopeptide ring, a five-membered condensed pyrrolidine ring or, except for a

224

tl- Anthoni, C. Christophersen and P. H. Nielsen

C-terminal tryptophan unit, a six-membered piperidone ring (a-carboline derivative) on oxonium ion catalyzed cyclization. The latter prospect is realized in kapakahine B, a cyclic hexapeptide, from the marine sponge Cribrochalina olemda, isolated in a yield of 2.0 mg from 4 kg wet weight of sponge [404]. The a-carboline moiety is a highly unusual feature previously encountered only in marine metabolites in grossularine-1 and -2fromthe tunicate Dendrodoa grossularia [405] and a few terrestrial species, e.g. Streptomyces griseoflavus [406]. However, it does demonstrate the existence, although presumably rare, in Nature of enzymes (presumably modular peptide synthetases) capable of performing such transformations. If, on the other hand, a cyclotryptophan moiety is formed, the consequences from a structural point of view would amount to the reversible transformation of a normal peptide unit to what amounts to a proline residue. Since proline is a very special amino acid (e.g. imparts a kink in an a-helix) the transformation undoubtedly would impress considerable restraints on the tertiary structure of the peptide or protein involved. A brief, but informative, discussion of the structural and dynamic role of proline in transmembrane helices has been published [407]. To our knowledge no example of a natural protein containing the cyclotryptophan unit has so far been identified. This could of course mean that this structural element does not exist in natural proteins. It could, however, also mean that proteins potentially capable of displaying the cyclo-element have not been investigated in sufficient details to reveal the presence of the element. Since the ring closure as well as the ring opening is acid catalyzed, isolation of the native molecule could easily result in opening of the potentially existing cyclotryptophans. Also in determination of the primary structure any cyclotryptophan would under the conditions usually employed count in the determination as tryptophan. The fact that reactions of this type can occur in proteins was demonstrated early in the case of lysozyme. When lysozyme was treated with iodine a product was formed where a carboxylic group (glutamic acid 35) formed an ester bond to the 2-position of tryptophan 108 [408-411], thereby creating a macrocyclic ring in the now inactive enzyme. The mechanism presumably consists of electrophilic attack of the iodine at the tryptophan 3-position followed by attack of the glutamic acid anion at the 2-position and elimination of hydrogen iodide to regenerate the tryptophan ring. This mechanism is analogous to the one encountered in the generation of 2-substituted tryptophans from 3a-hydroxycyclotryptophans discussed above. The structure was confirmed by X-ray analysis [411] and is now regarded as a classical example. In the acetylcholine esterase example treated above the enzyme itself dispensed the catalytic activity required for the cyclization. If this reaction has relevance in the regulation of intermediary metabolism, enzymes capable of effecting the reactions must exist. Until recently no example of posttranslational modification of tryptophan (and glycine, alanine, leucine, isoleucine and valine) was known. The "bromosleeper peptide", a 33-amino acid, L-6bromotryptophan containing peptide of the venom of the carnivorous fish hunting marine cone snail, Conus radiatus, has been investigated and shown to offer an example of posttranslational bromination of the peptide tryptophan residues [412]. The descriptive name originates from the sleep-like state induced on administration to mice. Analogous findings were reported for

Naturally Occurring Cyciotryptophans and Cyclotryptamines

225

a heptapeptide from the venom of a worm hunting cone, Conus imperialis. A few other examples of posttranslational modifications of tryptophan are now known. As has been touched upon earlier in this review, there is compelling evidence that nonribosomal peptide formation requires enzymes capable of effecting the cyclization equilibria. There is thus reason to believe that multienzymes contain modules possessing this capacity. We submit that the tryptophan cyclotryptophan equilibria has regulatory significance in the physiology of the cell and/or organism and that a whole class of hitherto unrecognized regulatory enzymes await discovery.

9. CONCLUSION The biological function and genetic origm of the tricyclic derivatives is a matter of considerable uncertainty. However, it is time to evoke, in natural products chemistry, the contemporary results emerging rapidly from biochemistry, molecular biology and genetics. Recently the complete genome sequence has been determined for microorganisms, e.g. Bacillus subtilis [413]. This impressive event allows the inspection of the total genetic capability of the organism. It appears that many of the genes are involved in the synthesis of secondary metabolites including antibiotics. These are of special interest in view of the discussion presented in this review of the genes coding for peptide synthetases (almost four percent of the genome codes for large multifunctional enzymes) like surfactin synthetase. At present the discussions and conclusions can often only be tentative. Obviously there exists a lack of cross fertilization within the different disciplines as evidenced by the lack of cross references in the papers of each area. In order to plan and carry out meaningful investigations a working hypothesis is indispensable. It is our hope that the discussions presented in the present review concerning the existence or nonexistence of metabolic processes governed by the principle outlined will prompt multidisciplinary investigations into this realm. Regardless of the actual outcome of such endeavors the results are bound to generate much needed information affecting future directions of research.

226

li. Anthoni, C. Christophersen and P. H. Nielsen

REFERENCES 1. 2. 3. 4. 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. 34. 35. 36. 37.

GA Cordell and JE Saxton, The Alkaloids: Chemistry and Physiology, Vol 20, RGA Rodrigo, Ed, Academic Press, New York 1981, Ch 1: Bisindole Alkaloids, p 3. J Sapi and G Massiot, Monoterpenoid Indole Alkaloids, Vol 25 Suppl. Part 4, JE Saxton, Ed, Chemistry of Heterocyclic Compounds, John Wiley and Sons, Chichester 1994, Ch 11: Bisindole Alkaloids, p 523. WB Turner, Fungal Metabolites, Academic Press, London 1971. WB Turner and DC Aldridge, Fungal Metabolites 2, Academic Press, London 1983. T Hino and M Nakagawa, The Alkaloids: Chemistry and Pharmacology, Vol 34, A Brossi, Ed, New York. 1989, Ch 1: Chemistry and Reactions of Cyclic Tautomers of Tryptamines and Tryptophans, p 1. MF Byford, JE Baldwin, C-J Shiau and CJ Schofield, Chem Rev 97:2631 (1997). MA Marahiel, T Stachelhaus and HD Mootz, Chem Rev 97:2651 (1997). H von Dohren, U Keller, J Vater and R Zocher, Chem Rev 97:2675 (1997). S Murao and H Hayashi, Agric Biol Chem 50:523 (1986). T Iwasa, S Harada and Y Sato, J Takeda, Biol Lab 40:12 (1981). H Shirafuji, S Tsuhoya and S Harada, JP 2-270875 (5.11.90), Chem Abs 114:120278r (1990) JE Leet, DR Schroeder, BS Krishnan and JA Matson, J Antibiot 43:961 (1990). KS Lam, GA Hesler, JM Mattel, SW Mamber, S Forenza and K Tomita, J Antibiot 43:956 (1990). SW Mamber, KW Brookshire, BJ Dean, RA Firestone, JE Leet, J A Matson and S Forenza, Antimicrob Agents Chemother 38:2633 (1994) JE Leet, DR Schroeder, J Golik, JA Matson, TW Doyle, KS Lam, SE Hill, MS Lee, JL Whitoey and BS Krishnan, J Antibiot 49:299 (1996). WW Carmichael, NA Mahmood and EG Hyde, Natural Toxins from Cyanobacteria (Blue-Green Algae) in Marine Toxins, Origin, Structure, and Molecular Pharmacology, S Hall and G Strichartz, Eds. ACS Symposium Series 418, American Chemical Society, Washington, DC 1990, Ch 6, p. 87. T Sano and K Kaya, Tetrahedron Lett 37:6873 (1996). K Ishida, H Matsuda, M Murakami and K Yamaguchi, Tetrahedron 52:9025 (1996). K Ishida, H Matsuda, M Murakami and K Yamaguchi, J Nat Prod 60:724 (1997) S Gravesen, JC Frisvad and RA Samson, Microfungi, 1994, Munksgaard, Copenhagen. S Ohmomo, T Sato, T Utagawa and M Abe, Agric Biol Chem 39:1333 (1975). S Ohmomo, T Sato, T Utagawa and M Abe, J Agric Chem Soc Jap 49:615 (1975). PM Scott, M-A Merrien and J Polonsky, Experientia 32:140 (1976). T Yamiguchi, K Nozawa, S Nakajima, K Kawai and S. Udagawa, Maikotokishin 34:29 (1991). PM Scott, Mycotoxins: Production, Isolation, Separation and Purification, V Betina, Ed, Elsevier, Amsterdam 1984, Ch 22: Roquefortine, p 463. P Haggblom, Appl Environ Microbiol 56:2924 (1990). S Ohmomo, HK Kitamoto and T Nakajima, J Sci Food Agric 64:211 (1994). RF Vesonder, L Tjarks, W Rohwedder and DO Kieswetter, Experientia 36:1308 (1980). TF Solovyeva and BP Baskunov, Prikl Biokhim Mikrobiol 28:880 (1992). TA Reshetilova, VI Shevchenko, VM Adanin and AG Kozlovsky, Prikl Biokhim Mikrobiol 29:418 (1993). V Sanchis, PM Scott and JM Farber, Mycopathologia 104:157 (1988). RJ Cole, JW Domer, RH Cox and LW Raymond, J Agric Food Chem 31:655 (1983). N Kyriakidis, ES Waight, JB Day and PG Mantle, Appl Environ Microbiol 42:61 (1981). S Ohmomo, K Oguma, T Ohashi and M Abe, Agric Biol Chem 42:2387 (1978). AG Kozlovskii, TA Reshetilova, TN Medvedeva, MU Arinbasarov, VG Sakharovskii and VM Adanin, Biokhimiya 44:1691 (1979). S Ohmomo, T Ohashi and M Abe, Agric Biol Chem 44:1929 (1980). TA Reshetilova, TF Solov'eva, LM Fadeeva and AG Kuzlovskii, Mikrobiologiya 61:391 (1992).

Naturally Occurring Cyclotryptophans and Cyclotryptamines

227

38.

AG Kozlovskii, TA Reshetilova, VG Sakharovskii, VM Adanin and AM Zyakun, Prikl Biokhim Mikrobiol 24:642 (1988).

39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50. 51. 52. 53.

72. 73. 74. 75. 76. 77. 78.

TA Reshetilova, OV Kuleshova and AG Kozlovskii, Mikrobiologiya 55:435 (1986). AG Kozlovsky, TF Solovieva, TA Reshetilova and GK Skryabin, Experientia 37:472 (1981). TA Reshetilova, TV Kulakovskaya, TN Kuvichkina and AG Kozlovsky, Mikrobiologiya 63:411 (1994). LP Dudina, TA Reshetilova, AG Kozlovsky and VK Eroshin, Prikl Biokhim Mikrobiol 29:700 (1993). TA Reshetilova and AG Kozlovskii, J Basic Microbiol 30:109 (1990). TV Kulakovskaya, TA Reshetilova, TN Kuvichkina and NG Vinokurova, Process Biochem 32:29 (1996). TA Reshetilova, NG Vinokurova, VN Khmelenina and AG Kozlovskii, Mikrobiologiya 64:36 (1995). PS Steyn and R Vleggaar, J Chem Soc Chem Conunun 1983:560. U Schoch, J Liithy and C Schlatter, Mitt Gebiete Lebensm Hyg 74:50 (1983). U Schoch, J Liithy and C Schlatter, Milchwissenschaft 39:76 (1984). B Kopp and HJ Rehm, J Appl Microbiol Biotechnol 13:232 (1981). M Medina, P Gaya and M Nunez, J Food Protect 48:118 (1985). CP Gorst-Allman and PS Steyn, J Chromatogr 175:325 (1979). A Lompe and K-E v Milczewski, Z Lebensm Unters Forsch 169:249 (1979). L Benitez, A Martin-Gonzalez, P Gilardi, T Soto, J Rodriguez de Lecea and JC Gutierrez, Lett Appl Microbiol 19:489 (1994). JC Frisvad, Prikl Biokhim Mikrobiol 29:19 (1993). J Frisvad and U Thrane, J Chromatogr 404:195 (1987). JC Frisvad, J Chromatogr 392:333 (1987). RRM Paterson, J Chromatogr 368:249 (1986). K Arai, K Kimura, T Mushiroda and Y Yamamoto, Chem Pharm Bull 37:2937 (1989). RP Hodge, CM Harris and TM Harris, J Nat Prod 51:66 (1988). A Musuku, MI Selala, T de Bruyne, M Claeys, PJC Schepens, A Tsatsakis and MI Shtilman, J Nat Prod 57:983 (1994). I Laws and PG Mantle, Phytochem 24:1395 (1985). AJ Birch and JJ Wright, J Chem Soc Chem Commun 1969:644. R Ritchie and JE Saxton, J Chem Soc Chem Commun 1975:611. T Kametani, N Kanaya and M Ihara, J Am Chem Soc 102:3974 (1980). T Kametani, N Kanaya and M Ihara, J Chem Soc Perkin I 1981:959. R Ritchie and JE Saxton, Tetrahedron 37:4295 (1981). S Murao, H Hayashi, K Takiuchi and M Arai, Agric Biol Chem 52:885 (1988). H Hayashi, K Takiuchi, S Murao and M Arai, Agric Biol Chem 52:2131 (1988). H Hayashi, K Takiuchi, S Murao and M Arai, Agric Biol Chem 53:461 (1989). H Hayashi, K Takiuchi, S Murao and M Arai, Agric Biol Chem 55:2177 (1991). S Takase, M Iwami, T Ando, M Okamoto, K Yoshida, H Horiai, M Kohsaka, H Aoki and H Imanaka J Antibiot 37:1320 (1984). S Takase, Y Kawai, I Uchida, H Tanaka and H Aoki, Tetrahedron Lett 25:4673 (1984). S Takase, Y Kawai, I Uchida, H Tanaka and H Aoki, Tetrahedron 41:3037 (1985). S Takase, Y Itoh, I Uchida, H Tanaka and H Aoki, Tetrahedron 42:5887 (1986). S Takase, Y Itoh, I Uchida, H Tanaka and H Aoki, Tetrahedron Lett 26:847 (1985). S Takase, I Uchida, H Tanaka and H Aoki, Tetrahedron 42:5879 (1986). S Takase, I Uchida, H Tanaka and H Aoki, Heterocycles 22:2491 (1984). SP Marsden, KM Depew and SJ Danishefsky, J Am Chem Soc 116:11143 (1994).

79. 80. 81. 82.

The Alkaloids, Vol 26, 1985, p 53. The Alkaloids, Vol 42, 1992, p 249. D Hauser. HP Weber and HP Sigg, Helv chim Acta 53:1061 (1970). S Udagawa, T Muroi, H Kurata, S Sekita, K Yoshihira, S Natori and M Umeda, Can J Microbiol 25:17i

54. 55. 56. 57. 58. 59. 60. 61. 62. 63. 64. 65. 66. 67. 68. 69. 70. 71.

228

83. 84. 85. 86. 87. 88. 89. 90. 91. 92. 93. 94. 95. 96. 97. 98. 99. 100. 101. 102. 103. 104.

105. 106. 107. 108. 109. 110. 111. 112. 113. 114. 115. 116. 117. 118. 119. 120. 121.

U. Anthoni, C. Christophersen and P. H. Nielsen (1979). S Sekita, K Yoshihira, S Natori, S Udagawa, T Muroi, Y Sugiyama, H Kurata and M Umeda, Can J Microbiol 27:766 (1981). HP Weber, Acta Cryst B28:2945 (1972). B Alarcon, JC Lacal, JM Femandez-Sousa and L Carrasco, Antiviral Research 4:231 (1984). JG Hamiff, Diss Abstr Int B39:750 (1978). T Saito, Y Suzuki, K Koyama, S Natori, Y litaka and T Kinoshita, Chem Pharm Bull 36:1942 (1988). T Saito, K Koyama, S Natori and Y litaka, Tetrahedron Lett 26:4731 (1985). D Brewer, JM Duncan, WA Jerram, CK Leach, S Safe, A Taylor, LC Vining, RMcG Archibald, RG Stevenson, CJ Mirocha and CM Christensen, Can J Microbiol 18:1129 (1972). S Safe and A Taylor, J Chem Soc Perkin I 1972:472. D Brewer and A Taylor, Can J Microbiol 24:1082 (1978). W-C Jen and GA Jones, Can J Microbiol 29:1399 (1983). T Kikuchi, S Kadota, K Nakamura, A Nishi, T Taga, T Kaji, K Osaki and K Tubaki, Chem Pharm Bull 30:3846 (1982). A Di Pietro, M Gut-Rella, JP Pachlatko and FJ Schwinn, Phytopathology 82:131 (1992). AD Argoudelis, J Antiobiot 25:171 (1972). AD Argoudelis and SA Mizsak, J Antiobiot 30:468 (1977). H Minato, M Matsumoto and T Katayama, J Chem Soc Chem Comm 1971:44. H Minato, M Matsumoto and T Katayama, J Chem Soc Perkin I 1973:1819. LI Chepenko, GI Borodin, VI Runov and BS Salikhova, Immunitet Pokoi Rast 1972:151, Chem Abs 78:156654. D Hauser, HR Loosli and P Niklaus, Helv chim Acta 55:2182 (1972). K Ueno, CS Giam and WA Taber, Mycologia 66:360 (1974). A Taylor, Biochemistry of some Foodbome Microbial Toxins, RI Mateles and GN Wogan, Eds,MIT Press, Cambridge, Massachusetts, 1967, p 69-107. A Taylor, Microbial Toxins, S Kadis, A Ciegler and SJ Ajl, Eds, Vol 7, Academic Press, New York, p 337-276. EP White, PH Mortimer and ME diMenna, Mycotoxic Fungi, Mycotoxins, Mycotoxicoses, An Encyclopedic Handbook, Vol 1, TD Wyllie and LG Morehouse, Eds, Marcel Dekker Inc, New York and Basel, 1975, p 427-447. J Fridrichsons and A McL Mathieson, Tetrahedron Lett 26:1265 (1962). J Fridrichsons and A McL Mathieson, Acta Cryst 18:1043 (1965). M Przybylska, EM Gopalakrishna, A Taylor and S Safe, J Chem Soc Chem Comm 1973:554. M Przybylska and EM Gopalakrishna, Acta Cryst B30:597 (1974). RT Gallagher, SJ Eichler, PT Holland, AL Wilkins and BK Nicholson, J Agric Food Chem 40:701 (1992). JW Ronaldson, A Taylor, EP White and RJ Abraham, J Chem Soc 1963:3172. WD Jamieson, R Rahman and A Taylor, J Chem Soc C 1969:1564. R Rahman, S Safe and A Taylor, J Chem Soc C 1969:1665. E Francis, R Rahman, S Safe and A Taylor, J Chem Soc Perkin I 1972:470. R Rahman, S Safe and A Taylor, J Chem Soc Perkin I 1978:1476. J W Ronaldson, Aust J Chem 34:1215 (1981). J W Ronaldson, Aust J Chem 29:2307 (1976). W Henderson, CO Miles and BK Nicholson, J Chem Soc Chem Comm 1995:889. R Nagarajan and RW Woody, J Am Chem Soc 95:7212 (1973). M Ohno, TF Spande and B Witkop, J Am Chem Soc 92:343 (1970). R Plate, MAJ Akkerman and HCJ Ottenheijm, J Chem Soc Perkin I 1987:2481. Y Kishi, S Nakatsuka, T Fukuyama and M Havel, J Am Chem Soc 95:6493 (1973).

Naturally Occurring Cyclotryptophans and Cyclotryptamines 122. 123. 124. 125. 126. 127. 128. 129. 130. 131. 132. 133. 134. 135. 136. 137.

229

S Nakatsuka, T Fukuyama and Y Kishi, Tetrahedron Lett 1974:1549. BL Smith, Trichothecenes Other Mycotoxins, Proc. Int. Mycotoxin Symp 1985:325. MC Middleton, Biochem Pharmacol 23:811 (1974). R Munday, Chem Biol Interactions 41:361 (1982). R Munday, J Appl Toxicol 7:17 (1987). LR Ferguson, J Berriman, A Pearson, R Munday, EA Fowke and NR Towers, Mutat res 268:199 (1992). RG Clark, DM Duganzich, L Mortleman and AJ Fraser, N Z Vet J 36:51 (1989). M Bonnefoi, M Hasim, P Sauvagnac, P De Saqui-Sannes, A Rico and V Burgat, C R Acad Sci Ser 3 311:169(1990). BL Smith and E Payne, N Z Vet J 39:46 (1991). GC Upreti and MK Jain, Biosci Rep 13:233 (1993). R Munday and E Manns, N Z Vet J 37:65 (1989). R Munday, J Appl Toxicol 4:182 (1984). SR Gobnerame, BL Smith and NR Towers, Plant-Assoc Toxins, SM Colegate and PR Dorling, Eds, CAB International, Wallingford, UK, 1994, 507. R Munday, J Appl Toxicol 5:69 (1985). K Furuya, M Okudaira, T Shindo and A Sato, Annu Rep Sankyo Res Lab 37:140 (1985). M Chu, T Truumees, ML Rotrofsky, MG Patel, F Gentile, PR Das, MS Puar and SL Lin, J Antibiot 48:1440(1995).

138. 139. 140. 141. 142. 143. 144. 145. 146.

C Shinohara, K Hasumi, Y Takei, A Endo, B Nuber, F Hansske and S Miura, J Antibiot 47:163 (1994). B Nuber, F Hansske, C Shinohara, S Miura, K Hasumi and A Endo, J Antibiot 47:168 (1994). JM Schkeryantz, JCG Woo and SJ Danishefsky, J Am Chem Soc 117:7025 (1995). The Alkaloids, Vol 29, 1986, pl85. Y Kimura, T Hamasaki, H Nakajima and A Isogai, Tetrahedron Lett 23:225 (1982). GA EUestad, P Mirando and MP Kunstmann, J Org Chem 38:4202 (1973). B Bhat and DM Harrison, Tetrahedron Lett 27:5873 (1986). B Bhat and DM Harrison, Tetrahedron 49:10655 (1993). JP Karwowski, M Jackson, RR Rasmussen, PE Humphrey, JB Poddig, WL Kohl, WH Scherr, S Kadam and JB McAlpine, J Antibiot 46:374 (1993). 147. JE Hochlowski, MM MuUally, SG Spanton, DN Whittem, P Hill and JB McAlpine, J Antibiot 46:380 (1993). 148. FS De Guzman, JB Gloer, DT Wicklow and PF Dowd, J Nat Prod 55:931 (1992). 149. JM Liesch, OD Hensens, DL Zink and MA Goetz, J Antibiot 41:878 (1988). 150. DR Houck, J Ondeyka, DL Zmk, E Inamine, MA Goetz and OD Hensens, J Antibiot 41:882 (1988). 151. MG Bock, RM DiPardo, SM Pitzenberger, CF Homnick, JP Springer and RM Freidinger, J Org Chem 52:1646(1987). 152. JP Springer, G Biichi, B Kobbe, AL Demain and J Clardy, Tetrahedron Lett 1977:2403. 153. M Nakagawa, H Sugumi, S Kodato and T Hino, Tetrahedron Lett 22:5323 (1981). 154. CM Maes. M Potgieter and PS Steyn, J Chem Soc Perkin I 1986:861. 155. AG Kozlowskii, TF Solov'eva, YE Bukhtiyarov, YV Shurukhin, VG Sakharovskii, VM Adanin, MY Nefedova, RN Pertsova, VG Tokarev and LA Golovleva, Mikrobiologiya 59:409 (1990). 156. CJ Barrow and DM Sedlock, J Nat Prod 57:1239 (1994). 157. CJ Barrow, P Cai, JK Snyder, DM Sedlock, HH Sun and R Cooper, J Org Chem 58:6016 (1993). 158. DM Sedlock, CJ Barrow, JE Brownell, A Hong, AM Gillum and DR Houck, J Antibiot 47:391 (1994). 159.

JJ Oleynek, DM Sedlock, CJ Barrow, KC Appell, F Casiano, D Haycock, SJ Ward, P Kaplita and AM

160. 161. 162.

Gillum, J Antibiot 47:399 (1994). JL Popp, LL Musza, CJ Barrow, PJ Rudewicz and DR Houck, J Antibiot 47:411 (1994). CJ Barrow, LL Musza and R Copper, Biiorg Med Chem Lett 5:377 (1995). M Varoglu, TH Corbett, FA Valeriote and P Crews, J Org Chem, m press.

230 163. 164. 165. 166. 167. 168. 169. 170. 171. 172. 173. 174. 175. 176. 177. 178. 179. 180. 181. 182. 183. 184. 185. 186. 187. 188. 189. 190. 191. 192. 193. 194. 195. 196. 197. 198.

U. Anthoni, C. Christophersen and P. H. Nielsen C Takahashi, A Numata, Y Ito, E Matsumura, H Araki, H Iwaki and K Kushida, J Chem Soc Perkin I 1994:1859. C Takahashi, Y Takai, Y Kimura, A Numata, N Shigematsu and H Tanaka, Phytochem 38:155 (1995). C Takahashi, A Numata, E Matsumura, K Minoura, H Eto, T Shingu, T Ito and T Hasegawa, J Antibiot 47:1242(1994). C Takahashi, K Minoura, T Yamada, A Numata, K Kushida, T Shingu, S Hagishita, H Nakai, T Sato and H Harada, Tetrahedron 51:3483 (1995). D Read, Nature 388:517 (1997). SW Simak et al.. Nature 388:579 (1997). SE Walden and RA Wheeler, J Am Chem Soc 119:3175 (1997). M Bnincko, D Crich and R Sami, Heterocycles 36:1735 (1993). A Berlin, A Canavesi, G Schiavon, S Zecchin and G Zotti, Tetrahedron 52:7947 (1996). U Anthoni, C Christophersen, PH Nielsen and EJ Pedersen, Acta Chem Scand 48:91 (1994). U Anthoni, L Chortsen, C Christophersen and PH Nielsen, Acta Chem Scand 49:441 (1995). PB Hoist, U Anthoni, C Christophersen, S Larsen, PH Nielsen and A Puschl, Acta Chem Scand 52:In press (1998). T Fatum, U Anthoni, C Christophersen and PH Nielsen, Acta Chem Scand 52:In press (1998). U Anthoni, D Christensen, C Christophersen and PH Nielsen, Acta Chem Scand 49:203 (1995). U Anthoni, C Christophersen, PH Nielsen and EJ Pedersen, 51:407 (1997). U. Anthoni, C Christophersen, A Obel and PH Nielsen, Acta Chem Scand 48:334 (1994). U Anthoni, C Christophersen, C Flensburg, MH Jakobsen, J Jensen and PH Nielsen, Structural Chemistry 7:103(1996). V De Luca, Methods in Plant Biochemistry, PJ Lea, Ed, Vol 9, 1993, Academic Press, London, Ch 13: Enzymology of Indole Alkaloid Biosynthesis, p. 345. B Tashkhodzhaev, K Samikov, MR Yagudaev, TP Antsupova, R Shakirov and SY Yunusov, Khim Prim Soedin 5:687 (1985). K Samikov, R Shakirov, TP Antsupova and SY Yunusov, Khim Prir Soedin 3:383 (1986). TP Antsupova and AV Polozhii, Rastit Resur 23:436 (1987). L Marion, The Alkaloids: Chemistry and Physiology, Vol 2, RHF Manske and HL Holmes, Eds, Academic Press, New York 1952, Ch 13: The Indole Alkaloids, p 438. E Coxworth, The Alkaloids: Chemistry and Physiology, Vol 8, RHF Manske, Ed, Academic Press, New York 1965, Ch 2: Alkaloids of the Calabar Bean, p 27. B Robinson, The Alkaloids: Chemistry and Physiology, Vol 10, RHF Manske, Ed, Academic Press, New York 1968, Ch 5: Alkaloids of the Calabar Bean, p 383. B Robinson, The Alkaloids: Chemistry and Physiology, Vol 13, RHF Manske, Ed, Academic Press, New York 1971, Ch 4: Alkaloids of the Calabar Bean, p 213. S Takano and K Ogasawara, The Alkaloids: Chemistry and Pharmacology, Vol 36, A Brossi, Ed, Academic Press, New York, 1989, Ch 5: Alkaloids of the Calabar Bean, p 225. RB Longmore and B Robinson, Chem and Ind 1969:622. RB Longmore and B Robinson, J Pharm Pharmac 21:118S (1969). Q-S Yu, HJC Yeh, A Brossi and JL Flippen-Anderson, J Nat Prod 52:332 (1989). E Redenti, M Delcanale, G Amari, P Ventura, A Bacchi and G Pelizzi, J Pharm Sci 84:1126 (1995). M Brzostowska, X He, NH Greig, SI Rapoport and A Brossi, Med Chem Res 2:238 (1992). A Brossi, X-F Pei and NH Greig, 1994 International Congress on Natural Products Research, Halifax, Nova Scotia, Canada. P Rosenmund, S Gektidis, H Brill and R Kalbe, Tetrahedron Lett 30:61 (1989). JP Marino, MW Kim and R Lawrence, J Org Chem 54:1782 (1989). M Node, X Hao and K Fuji, Chem Lett 1991:57. QS Yu, WM Luo, YQ Li and A Brossi, Heterocycles 36:1279 (1993).

Naturally Occurring Cyclotryptophans and Cyclotryptamines 199. 200. 201. 202. 203. 204. 205. 206. 207. 208. 209. 210. 211. 212. 213. 214. 215. 216. 217. 218. 219. 220. 221. 222. 223. 224. 225. 226. 227.

231

S Bonnotlours, C Crouzel, C Prenant and F Hitmen, J Label Compound Radiopharm 33:277 (1993). M Pallavicini, E Valoti, L Villa and I Resta, Tetrahedron-Asymmetry 5:963 (1994). MS Moralesrios, MA Bucio, C Garciamartinez and P Josephnathan, Tetrahedron Lett 35:6087 (1994). QS Yu, BY Lu and XF Pei, Heterocycles 39:519 (1994). XF Pei, NH Greig, JL Flippen-Anderson, S Bi and A Brossi, Helv Chim Acta 77:1412 (1994). N Pallavicini, E Valoti, L Villa and F Lianza, Tetrahedron-Asymmetry 5:111 (1994). PF Santos, AM Lobo, S Prabhakar, Tetrahedron Lett 36:8099 (1995). MS Moralesrios, MA Bucio and P Josephnathan, Tetrahedron 52:5339 (1996). M Node, X Hao, K Nishide and K Fuji, Chem Pharm Bull 44:715 (1996). XF Pei, QS Yu, BY Lu, NH Greig and A Brossi, Heterocycles 42:229 (1996). T Hino, S Kodato, K Takahashi, H Yamaguchi and M Nakagawa, Tetrahedron Lett 1978:4913. CL Fang, S Home, N Taylor and R Rodrigo, J Am Chem Soc 116:9480 (1994). RK Duke, RD Allan, GAR Johnston, KN Mewett, AD Mitrovic, CC Duke and TW Hambley, J Nat Prod 58:1200(1995). F Tillequin, M Koch, M Bert and T Sevenet, J Nat Prod 42:92 (1979). F Tillequin and M Koch, Phytochem 18:1559 (1979). F Tillequin and M Koch, Phytochem 18:2066 (1979). F Tillequin, R Rousselet, M Koch, M Bert and T Sevenet, Ann Pharm Fr 37:543 (1979). PG Waterman, Alkaloids: Chemical and Biological Perspectives, Vol 4, SW Pelletier, Ed, Wiley, New York, 1986, Ch. 3, p 331.

234. 235. 236.

J-L Pousset, A Cave, A Chiaroni and C Riche, J Chem Soc Chem Comm 1977:261. G Maynart, J-L Pousset, S Mboup and F Denis, C R Soc Biol 174:925 (1980). F Tillequin, M Koch, J-L Pousset and A Cave, J Chem Soc Chem Comm 1978:826. F Tillequin, M Koch and A Rabaron, J Nat Prod 48:120 (1985). MS Morales.Rios, J Espineira and P Joseph-Nathan, Magn Reson Chem 25:377 (1987). KK Purushothaman and A Sarada, Phytochem 20:351 (1981). AM Balde, LA Pieters, A Gergely. V Wrav, M Claeys and AJ Vlietinck, Phytochem 30:997 (1991). J Fridrichsons, MF Mackay and AM Mathieson, Tetrahedron Lett 1967:3521. J Fridrichsons, MF Mackay and AM Mathieson, Tetrahedron 30:85 (1974). NH Ujis, Z Mahmud and RF Toia, Planta Med 59:383 (1993). F Gueritte-Voegelein, T Sevenet, J Pusset, M-T Adeline, B Gillet, J-C Beloeil, D Guenard, P Potier, R Rasolonjanahary and C Kordon, J Nat Prod 55:923 (1992). F Libot, N Kunesch, J Poisson, M Kaiser and H Duddeck, Heterocycles 27:2381 (1988). F Libot, C Miet, N Kunesch, JE Poisson, J Pusset and T Sevenet, J Nat Prod 50:468 (1987). KF Parry and GF Smith, J Chem Soc Perkin 1 1978:1671. A Roth, B Kuballa, P Cabalion and R Anton, Planta Med 51:289 (1985). A Beretz, A Roth-Georger, G Corre, B Kuballa, R Anton and J-P Cazenave, Planta Med 51:300 (1985). A Roth, B Kuballa, C Bounthanh, P Cabalion, T Sevenet, JP Beck and R Anton, Planta Med 52:450 (1986). NK Hart, SR Johns, JA Lamberton and RE Summons, Aust J Chem 27:639 (1974). Y Adjibade, H Saad, T Sevenet, B Kuballa, JC Quirion and R Anton, Planta Med 56:212 (1990). Y Adjibade, B Kuballa, P Cabalion, ML Jung, JP Beck and R Anton, Planta Med 55:567 (1989).

237.

Y Adjibade, H Saad, B Kuballa, JP Beck, T Sevenet, P Cabalion and R Anton, J Ethnopharmac 29:127

228. 229. 230. 231. 232. 233.

(1990). 238.

CWL Bevan, MB Patel, AH Rees and AG Loudon, Tetrahedron 23:3809 (1967).

239.

Y Morita, M Hesse and H Schmid, Helv chim Acta 51:1438 (1968).

240. 241. 242.

EA Adegoke and B Alo, Phytochem 25:1461 (1986). A-M Morfaux, L Olivier. J Levy and J Le Men, Ann pharm franc 27:679 (1969). RH Bumell, A Chapelle and MF Khalil, Can J Chem 52:2327 (1974).

232 243. 244. 245. 246. 247. 248. 249. 250. 251. 252. 253. 254. 255. 256. 257. 258. 259. 260. 261. 262. 263. 264. 265. 266. 267. 268. 269. 270. 271. 272. 273. 274. 275. 276. 277. 278.

U. Anthoni, C. Christophersen and P. H. Nielsen RH Bumell, A Chapelle, MF Khalil and PH Bird, J Chem Soc Chem Comm 1970:772. A-M Morfaux, J Vercauteren, J Kerharo, L Le Men-Olivier and J Le Men, Phytochem 17:167 (1978). L Le Men-Olivier, Plant medicin phytotherap 12:173 (1978). J Vercauteren, G Massiot, L Me Men-Olivier, J Levy and C Delaude, Bull soc chim France 1982-11:291. C Lavaud, G Massiot, J Vercauteren and L Le Men-Olivier, Phytochem 21:445 (1982). LSR Arambewela and F Khuong-Huu, Phytochem 20:349 (1981). H Takayama, S Subhadhirasakul, J Mizuki, M Kitajima, N Aimi, D Ponglux and S Sakai, Chem Pharm Bull 42:1957 (1994). S Subhadhirasakul, H Takayama, Y Miyabe, N Aimi, D Ponglux and S Sakai, Chem Pharm Bull 42:2645 (1994). G Massiot, C Lavaud, J Vercauteren, L Le Men-Olivier, J Levy, J Guilhem and C Pascard, Helv chim Acta 66:2414 (1983). F Heatley, DI Bishop and JA Joule, J Chem Soc Perkin II 1981:725. P Leewanich, M Tohda, K Matsumoto, S Subhadhirasakul, H Takayama and H Watanabe, Biol Pharm Bull 19:394 (1996). A Chatterjee, AK Ghosh and EW Hagaman, J Org Chem 47:1732 (1982). S Subhadhirasakul, H Takayama, N Aimi, D Ponglux and S Sakai, Chem Pharm Bull 42:1427 (1994). The Akaloids, 20:297 (1981). I Kompis and J Mokry, Coll Czech Chem Comm 33:4328 (1968) H Meisel and W Dopke, Tetrahedron Lett 1971:1285. B Proksa, D Uhrin, E Grossmann and Z Voticky, Planta medica 1987:120. M Sturdikova, J Fuska, E Grossmann and Z Voticky, Pharmazie 41:270 (1986). B Proksa, D Uhrin, E Grossmann and Z Voticky, Tetrahedron Lett 27:5413 (1986) B Proksa, D Uhrin, E Grossmann, Z Voticky and J Fuska, Planta medica 1988:214. B Proksa and E Grossmann, Phytochem Anal 2:74 (1991). JE Saxton, The Alkaloids: Chemistry and Physiology, Vol 8, RHF Manske, Ed, Academic Press, New York 1965. Ch 8, p 159. JE Saxton, The Alkaloids: Chemistry and Physiology, Vol 12, RHF Manske, Ed, Academic Press, New York 1970, Ch 3, p 207. JE Saxton, The Alkaloids: Chemistry and Physiology, Vol 14, RHF Manske, Ed, Academic Press, New York 1973, p 168. CK Ratnayake, LSR Arambewela, KTD De Silva, Atta-Ur-Rahman and KA Alvi, Phytochem 26:868 (1987). Atta-ur-Rahman, SA Abbas, F Nighat, G Ahmed, MI Choudhary, KA Alvi and Habib-Ur-Rehman, J Nat Prod 54:750 (1991). BC Das, JP Cosson, G Lukacs and P Potier, Tetrahedron Lett 1974:4299. J Vercauteren, G Massiot, T Sevenet, J Levy, L Le Men-Olivier and J Le Men, Phytochem 18:1729 (1979). MJ Jacquier, J Vercauteren, G Massiot, L Le Men-Olivier, J Pusset and T Sevenet, Phytochem 21:2973 (1982). C Caron, Y Yachaoui, G Massiot, L Le Men-Olivier, J Pusset and T Sevenet, Phytochem23:2355 (1984). D Guillaume, AM Morfaux, B Richard, G Massiot, L Le Men-Olivier, J Pusset and T Sevenet, Phytochem 23:2407 (1984). S Mamatas-Kalamaras, T Sevenet, C Thai and P Potier, Phytochem 14:1637 (1975). A-M Morfaux, P Mouton, G Massiot and L Le Men-Olivier, Phytochem 31:1079 (1992). W Hu, J Zhu and M Hesse, Planta Medica 55:463 (1989). G Massiot, A Boumendjel, J-M Nuzillard, B Richard, L Le Men-Olivier, B David and HA Hadi, Phytochem 31:1078 (1992). W Boonchuay and WE Court, Phytochem 15:821 (1976).

Naturally Occurring Cyclotryptophans and Cyclotryptamines 279. 280. 281. 282. 283. 284. 285. 286. 287. 288. 289. 290. 291. 292. 293. 294. 295. 296. 297. 298. 299. 300. 301. 302. 303. 304. 305. 306. 307. 308. 309. 310. 311. 312. 313.

314. 315.

233

W Boonchuay and WE Court, Planta Medica 29:380 (1976). T Yamauchi, F Abe, WG Padolina and FM Dayrit, Phytochem 29:3321 (1990). C Caron, A Graftieaux, G Massiot, L Le Men-Olivier and C Delaude, Phytochem 28:1241 (1989). CW Ming, ZP Ling and G Riicker, Planta Medica 1988:480. N Keawpradub, H Takayama, N Aimi and S-I Sakai, Phytochem 37:1745 (1994). B Chandrasekaran and B Nagarajan, J Biosci 3:395 (1981). JAO Ojewole, Fitoterapia 54:99 (1983). JAO Ojewole, Int J Crude Drug Res 22:121 (1984). B Chandrasekaran, R Vijayendran, KK Purushothaman and B Nagarajan, Indian J Biochem Biophys 19:148 (1982). B Chandrasekaran and B Nagarajan, Arogya 9:60 (1983). GT Tan, JF Miller, AD Kinghom, A Douglas, SH Hughes and JM Pezzuto, Biochem Biophys Res Commun 185:370(1992). CW Wright, D Allen, JD Phillipson, GC Kirby, DC Warhurst, G Massiot and L Le Men-Olivier, J Ethnopharmac 40:41 (1993). N Kunesch, A Cave, EW Hagaman and E Wenkert, Tetrahedron Lett 21:1727 (1980). A Cave, N Kunesch, J Bruneton, R Goutarel and GP Wannigama, J Nat Prod 50:1178 (1987). Atta-ur-Rahman, A Pervin, I Ali, A Muzaffar, KTD DeSilva and WSJ Silva, Planta Medica 1988:37. Atta-ur-Rahman, A Pervin, A Muzaffar, KTD DeSilva and WSJ Silva, Heterocycles 27:2051 (1988). L Douzoua, M Mansour, M-M Debray, L Le Men-Olivier and J Le Men, Phytochem 13:1994 (1974). M Mansour, L Le Men-Olivier, J L^vy and J Le Men, Phytochem 13:2861 (1974). S Savaskan, I Kompis, M Hesse and H Schmid, Helv. Chim. Acta 55:2861 (1972). HK Schnoes, K Biemann, J Mokry, I Kompis, A Chatterjee and G Ganguli, J. Org. Chem. 31:1641 (1966). G Massiot, J Vercauteren, B Richard, M-J Jacquier and L Le Men-Olivier, C R Acad Sc Paris II 294:579 (1982). TA Van Beek, R Verpoorte, AB Svendsen and R Fokkens, J Nat Prod 48:400 (1985). B Witkop, Heterocycles 20:2059 (1983). S Markey, K Biemann and B Witkop, Tetrahedron Lett 1967:157. G Massiot, P Thepenier, M-J Jacquier, L Le Men-Olivier and C Delaude, Heterocycles 29:1435 (1989). DJ Faulkner, H-Y He, MD Unson, CA Bewly and MJ Garson, Gazz Chim Ital 123:301 (1993). N Fusetani and S Matsunaga, Chem Rev 93:1793 (1993). WD Clark, T Corbett, F Valeriote and P Crews, J Amer Chem Soc 119:9285 (1997). GR Pettit, R Tan, DL Herald, RL Cemy and MD Williams, J Org Chem 59:1593 (1994). PM Dewick, Nat Prod Rep 14:111 (1997). WR Alonso and R Croteau, Methods in Plant Biochemistry, Ed PJ Lea, Vol 9, 1993, Academic Press, London, Ch 9: Prenyltransferases and Cyclases, p 239. C Christophersen, Acta Chem Scand B39:517 (1985). C Christophersen, Marine Natural Products, Chemical and Biological Perspectives, PJ Scheuer, Ed, Academic Press, New York, Vol 5, 1983, Ch 5: Marine Indoles, p 259. C Christophersen, The Alkaloids, Chemistry and Physiology, A Brossi, Ed, Academic Press, New York, Vol 24, 1985, Ch 2: Marine Alkaloids, p 25. GW Gribble, Progress in the Chemistry of Organic Natural Products, Vol 68, W Herz, GW Kirby, RE Moore, W Steglich and C Tamm, Eds, Springer, New York, 1996. Namrally Occurring Organohalogen Compounds - A Comprehensive Survey, p 1. J Kobayashi and M Ishibashi, The Alkaloids, Chemistry and Physiology, A Brossi and GA Cordell, Eds, Academic Press, New York, Vol 41, 1992, Ch 2: Marine Alkaloids II, p 41. W Fenical, Alkaloids: Chemical and Biological Perspectives, SW Pelletier, Ed, Wiley and Sons, New York 1986, Vol 4, Ch 2: Marine Alkaloids and Related Compounds, p 275.

234

U. Anthoni, C. Christophersen and P. H. Nielsen

316. 317. 318. 319. 320. 321. 322. 323. 324. 325. 326. 327. 328.

M Alvarez, M Salas and JA Joule, Heterocycles 32:1391 (1991). JS Carle and C Christophersen, J Amer Chem Soc 101:4012 (1979). JS Carle and C Christophersen, J Org Chem 45:1586 (1980). JS Carle and C Christophersen, J Org Chem 46:3440 (1981). PB Hoist, U Anthoni, C Christophersen and PH Nielsen, J Nat Prod 57:997 (1994). P Wulff, JS Carle and C Christophersen, Comp Biochem Physiol 718:523 (1982). P Keil, EG Nielsen, U Anthoni and C Christophersen, Acta Chem Scand B40:555 (1986). JLC Wright, J Nat Prod 47:893 (1984). MV Laycock, JLC Wright, JA Findlay and AD Patil, Can J Chem 64:1312 (1986). U Anthoni, PH Nielsen, M Pereira and C Christophersen, Comp Biochem Physiol 96B:431 (1990). R Nielsen, Br Phycol J 19:371 (1984). C Christophersen, Comp Biochem Physiol 98:427 (1991). C Christophersen, Studies in Natural Products Chemistry, Atta-ur-Rahman, Ed., Elsevier Science BV, Amsterdam, Theory of the Origin, Function and Evolution of Secondary Metabolites, 18:667 (1996). 329. PEJ Dyrynda, Developmental and Comparative Immunology 7:621 (1983). 330. PEJ Dyrynda, Proc 19^^ European Marine Biology Symposium, Cambridge Univ Press 1985:411. 331. PEJ Dyrynda, Bryozoa: Ordovician to Recent, C Nielsen and GP Larwood, Eds. Olsen and Olsen, Fredensborg, Functional AUelochemistry in Temperate Waters: Chemical Defence of bryozoans, 1985:95. 332. T Sjoblom, L Bohlin and C Christophersen, Acta Pharm Suec 20:415 (1983). 333. PB Hoist, U Anthoni, C. Christophersen and PH Nielsen, J Nat Prod 57:1310 (1994). 334. T Sjoblom, L Bohlin and C Christophersen, Acta Pharm Suec 20:415 (1983). 335. T Hino, T Tanaka, K Matsuki and M Nakagawa, Chem Pharm Bull 31:1806 (1983). 336. P Muthusubramanian, JS Carle and C Christophersen, Acta Chem Scand B37:803 (1983). 337. J Jensen, U Anthoni, C. Christophersen and PH Nielsen, Acta Chem Scand 49:68 (1995). 338. M Bruncko, D Crich and R Samy, J Org Chem 59:5543 (1994). 339. T Kawasaki, R Terashima, K Sakaguchi, H Sekiguchi and M Sakamoto, Tetrahedron Lett 37:7525 (1996). 340. S Takase, I Uchida, H Tanaka and H Aoki, Heterocycles 22:2491 (1984). 341. S Takase, I Uchida, H Tanaka and H Aoki, Tetrahedron 42:5879 (1986). 342. L Rahbaek, U Anthoni, C Christophersen, PH Nielsen and BO Petersen, J Org Chem 61:887 (1996). 343. L Rahbaek and C Christophersen, J Nat Prod 60:175 (1997). 344. U Anthoni, L Chevolot, C Larsen, PH Nielsen and C Christophersen, J Org Chem 52:4709 (1987). 345. BS Davidson, Chem Rev 93:1771 (1993). 346. C Christophersen and U Anthoni, Sulfur Reports 4:365 (1986). 347. S Tsukamoto, H Hirota, H Kato and N Fusetani, Tetrahedron Lett 34:4819 (1993). 348. S Tsukamoto, H Hirota, H Kato and N Fusetani, Experientia 50:680 (1994). 349. JW Daly and TF Spande, Alkaloids: Chemical and Biological Percpectives, SW Pelletier, Ed, Wiley and Sons, New York, 1986, vol 4, Ch 1: Amphibian Alkaloids: Chemistry, Pharmacology, and Biology, p 1. 350. 351. 352. 353. 354. 355. 356. 357. 358.

JW Daly, HM Garraffo and TF Spande, The Alkaloids: Chemistry and Pharmacology, GA Cordell, Ed, Academic Press, San Diego, California, 1993, Vol 43, p 186. T Tokuyama and JW Daly, Tetrahedron 39:41 (1983). TF Spande, MW Edwards, IK Pannell, JW Daly, V Erspamer and P Melchiorri, J Org Chem 53:1222 (1988). JW Daly, HM Garaffo, LK Pannell, TF Spande, C Severini and V Erspamer, J Nat Prod 53:407 (1990). MO Mitchell and PW Le Quesne, Tetrahedron Lett 31:2681 (1990). WY Sun, Y Sun, YC Tang and JQ Hu, Synlett 1993:337. D Crich, AB Pavlovic and R Samy, Tetrahedron 51:6379 (1995). PG Cozzi, C Palazzi, D Potenza, C Scolastico and WY Sun, Tetrahedron Lett 31:5661 (1990). 0 Vakkuri, J Tervo, R Luttinen, H Ruotsalainen, E Rahkamaa and J Leppaluoto, Endocrinology 120:2453

Naturally Occurring Cyclotryptophans and Cyclotryptamines

235

(1987). 359. 360. 361. 362. 363. 364. 365. 366. 367. 368. 369. 370. 371. 372.

AG Kozlovskii, TA Reshetilova and TN Medvedeva, Mikrobiologiya 51:48 (1982). AG Kozlovskii and TA Reshetilova, Mikrobiologiya 53:81 (1984). TA Reshetilova and AG Kozlovskii, Mikrobiologiya 54:699 (1985). J Kusch and HJ Rehm, Appl Microbiol Biotechnol 23:394 (1986). I Laws and PG Mantle, J Gen Microbiol 135:2679 (1989). TA Reshetilova and AG Kozlovsky, J Basic Microniol 30:109 (1990). PS Steyn and R Vieggaar, J Chem Soc Chem Comm 1983:560. S Ohmono, T Ohashi and M Abe, Agric Biol Chem 43:2035 (1979). CP Gorst-Allman, PS Steyn and R Vieggaar, J Chem Soc Chem Comm 1982:652. PG Mantle, KPWC Perera, NJ Maishman and GR Mundy, Appl Environm Microbiol 45:1486 (1983). B Bhat, DM Harrison and HM Lamont, J Chem Soc Chem Comm 1990:1548. B Bhat, DM Harrison and HM Lamont, Tetrahedron, 46:10663 (1993). KD Barrow, PW CoUey and DE Tribe, J Chem Soc Chem Comm 1979:225. MF Grundon, MR Hemblin, DM Harrison, JND Logue, M Maguire and J A McGrath, J Chem Soc Perkin I 1980:1294. 373. JC Gebler, AB Woodside and CD Poulter, J Am Chem Soc 114:7354 (1992). 374. R Vieggaar and PL Wessels, J Chem Soc Chem Comm 1980:160. 375. GW Kirby and MJ Varley, J Chem Soc Chem Comm 1974:833. 376. DR Houck, J Ondeyka, DL Zink, E Inamine, MA Goetz and OD Hensens, J Antibiot 41:882 (1988). 377. JL Popp, LL Musza, CJ Barrow, PJ Rudewicz and DR Houck, J Antibiot 47:411 (1994). 378. JF Sanz-Cervera, T Glinka and RM Williams, J Am Chem Soc 115:347 (1993). 379. S Rhee, KD Parris, CC Hyde, SA Ahmed, EW Miles and DR Davies, Biochemistry 36:7664 (1997). 380. GW Kirby, SW Shah and EJ Herbert, J Chem Soc C 1969:1916. 381. S-W Yang and GA Cordell, J Nat Prod 60:44 (1997). 382. S-W Yang and GA Cordell, J Nat Prod 60:230 (1997). 383. M Nakagawa, Y Yokoyama, S Kato and T Hino, Tetrahedron 41:2125 (1985). 384. A Singh, SA Antonsen, GW Koroll, W Kremers and H Singh, Oxygen Radicals in Chemistry and Biology, 1984, Walter de Gruyter & Co. Berlin, p. 491. 385. K Itakura, K Uchida and S Kawakishi, Tetrahedron Lett 33:2567 (1992). 386. WE Savige and A Fontana, J C S Chem Comm 1976:600. 387. NT Nguyen, MZ Wrona and G Dryhurst, J Electroanal Chem 199:101 (1986. 388. U Anthoni, C Christophersen, PH Nielsen, MW Christoffersen and D Sorensen, Acta Chem Scand 52:In press (1998). 389. WE Savige and A Fontana, Int J Peptide Protein Res 15:102 (1980). 390. M Ohno, S Tanaka, T-C Shieh and TF Spande, J Org Chem 49:5069 (1984). 391. G Zanotti, C Birr and T Wieland, Int J Peptide Protein Res 12:204. 392. T Wieland, B Beijer, A Seeliger, J Dabrowski, G Zanotti, AE Tonelli, A. Gieren, B Dederer, V. Lamm and E Hadicke, Liebigs Ann Chem 1981:2318. 393. G Zanotti, B Beijer and T Wieland, Int J Peptide protein Res 30:323 (1987). 394. G Zanotti. T Wieland, G D'Auria, L Paolillo and E Trivellone, Int J Peptide Protein Res 35:263 (1990). 395. EV Kudryavtseva, MV Sidorova, MV Ovchinnikov, ZD Bespalova and VN Bushuev, J Pept Res 49:52 (1997). 396.

G Poli, E Albano and MU Dianzani, Eds, Free Radicals: From Basic Science to Medicine, Birkhauser

397. 398.

Verlag, Basel, 1993. I Emerit and B Chance, Eds, Free Radicals and Aging, Birkhauser Verlag, Basel, 1992. M Tarr and F Samson, Eds, Oxygen Free Radicals in Tissue Damage, Birkhauser Verlag, Boston, 1993.

399.

MR Marzabadi, D Yin and UT Brunk, Lipofuscinogenesis in a model system of cultured cardiac myocytes, p 78. Reference 396.

236 400. 401. 402. 403. 404. 405. 406. 407. 408. 409. 410. 411. 412. 413.

U. Anthoni, C. Christophersen and P. H. Nielsen KT Hill and C Womersley, Marine Biology 109:1 (1991). A Hussain, H Wahner and J Triplet!, J Pharai Sci 67:742 (1978). JL Sussman, M Harel, F Frolov, C Oefher, A Goldman, L Toker and I Silman, Science 253:872 (1991). JC Ma and DA Dougherty, Chem Rev 97:1303 (1997). Y Nakao, BKS Yeung, WY Yoshida, PJ Scheuer and M Kelly-Borges, J Am Chem Soc 117:8271 (1995). C Moquin-Pattey and M Guyot, Tetrahedron 45:3445 (1989). JS Kim, K Shinya, K Furihata, Y Hayakawa and H Seto, Tetrahedron Lett 38:3431 (1997). KA Williams and CM Deber, Biochemistry 30:8919 (1991). T Imoto, FJ Hartdegen and JA Rupley, J Mol Biol 80:637 (1973). FJ Hartdegen and JA Rupley, J Mol Biol 80:649 (1973). T Imoto and JA Rupley, J Mol Biol 80:657 (1973). CR Beddell, CCF Blake and SJ Oatley, J Mol Biol 97:643 (1975). AG Craig, EC Jimenez, J Dykert, DB Nielsen, J Gulyas, FC Abogadie, J Porter, JE Rivier, U Cruz, BM Olivera and JM Mcintosh, J Biol Chem 272:4689 (1997). F Kunst et al. (with 150 other authors), Nature 390:249 (1997).

Chapter Three

Recent Research on Pyrrole Alkaloids Philip W. Le Quesne and Ying Dong Department of Chemistry and Barnett Institute Northeastern University, 360 Huntington Avenue Boston, MA 02115-5096, U.S.A. Todd A. Blythe Vertex Pharmaceuticals, 130 Waverly Street Cambridge, MA 02139-4242, U.S.A. CONTENTS 1. Introduction

238

2. Simple Pyrroles

238

3. Pyrrolnitrin

240

4. Prodigiosin

242

5. Alkaloids from Quararibea funebris

242

6. Pyrrole Alkaloids from Sponges 6.1. Pyrrololactams 6.2. Phakellins and Related Compounds 6.3. Pyrroloquinolines 6.4. The Phorbazoles 6.5. Miscellaneous Pyrroles from Sponges 7. Pyrrole Alkaloids from Ascidians 7.1. Eudistomins 8. Pyrrole Alkaloids from Molluscs 8.1. Lamellarins 9. Miscellaneous Marine Pyrrole Alkaloids 9.1. Triketramine 9.2. Rhodophycan Pyrrole Alkaloids 9.3. Tambjamines and Related Compounds 9.4. Pyrrolomycins and Pyrroindomycins 9.5. Pyrrolethers 9.6. Keronopsins A and B, Protozoan Defense Agents 10. Miscellaneous Pyrroles from Microorganisms

245 245 248 256 262 262 263 263 265 265 267 267 267 268 269 272 272 273

11. Minor Piper Alkaloids Containing a Pyrrole Skeleton

276

12. Miscellaneous Pyrroles from Higher Plants

278

237

238

P- W. LeQuesne, Y. Dong and T. A. Blythe

13. Roseophilin

279

14. Conclusion

282

15. Acknowledgments

282

References

282

INTRODUCTION

Alkaloids have long been defined, in an approximate way, as natural nitrogen-containing compounds, found particularly in plants. The name is derived from the characteristic basic properties (alkali-like) of many alkaloids, which are induced by the lone electron pair of nitrogen [1]. Investigation of alkaloids during nearly two hundred years has revealed that many contain different heterocyclic skeletons, some of which occur much more frequently than others [2]. One class which is relatively little known is that containing the comparatively non-basic pyrrole (1) ring system. Only a few examples [3-5] appeared in the literature between 1940 and 1980.

O N I

R 1 However, over the past fifteen years there has been a dramatic increase in identification of pyrrole alkaloids. The purpose of this chapter is to review the literature during this time to report on members of the small but expanding class of alkaloids containing the pyrrole moiety. This review will encompass where these compounds are found and isolated, their structures as well as their biological activities, and selected chemical syntheses of some pyrrole alkaloids. The literature has also been surveyed to investigate biosynthetic pathways, either known or postulated, to pyrrole alkaloids.

SIMPLE PYRROLES

Metabolites incorporating a halogenated pyrrole ring are well known in marine organisms (see below). Even the simplest of these may show antibacterial activity and possibly have a

Recent Research on Pyrrole Alkaloids

239

physiological role, such as 2,3,4-tribromopyiTole (2) isolated from the polychaete Polyphysia crassa [6]. 2-Acetylpyrrole (3), isolated from a culture broth of Streptomyces sp. A-5071, Br-s.^

^Br

// N^ H

Br

W

.CH,

N H

was found to protect primary cultured rat hepatocytes against D-galactosamine-induced cytotoxicity [7]. Glycerinopyrin (4) from a Sri Lankan strain of Streptomyces violaceus possesses the intriguing A^-hydroxypyrrole nucleus and was reported to be 85% S, 15% R at the side-chain stereogenic center [8]. Pyrrole-2-carboxylic acid is frequently found esterified to other secondary metabolite nuclei; an example is its ester with 9, 21-didehydroxyryanodine, a constituent of insecticidal ryania powder [9]. Two A^, 2-substituted pyrroles (5) and (6) (see section 9.2.) may be related biogenetically to compound 4.

o N R

II O

5 R=CH2CH(OH)CH3 OH ' - ' " 6 R=CH2CH20H

The Chinese herbal medicine tai-zi-shen (Pseudostellaria heterophylla ), used as a pediatric or geriatric tonic, contains 3-furfuryl pyrrole-2-carboxylate (7) [11]. Oxidative metabolism of the plant cytokinin zeatin riboside gives the pyrrole derivative 8 which was synthesized by a biomimetic oxidation reaction [12]. 4-Geranylpyrrole-2-carboxylic acid (9), isolated from Streptomyces chrestomyceticus [13], is an inhibitor of lipid peroxidation in rat brain homogenate, and was named pyrrolostatin. From another Streptomyces species (sp. MI 424-38 Fl) [14], the interesting polychlorinated antibiotic neopyrrolomycin (10) was obtained. This compound, which as isolated is optically active by virtue of restricted intemuclear rotation, has a broad range of antibacterial and antifungal activity.

240

P. W. LeQuesne, Y. Dong and T. A. BIythe

.Me

^OHrbH

HOOC

"^N H

PYRROLNITRIN

Pyrrolnitrin (11), 3-(2-nitro-3-chlorophenyl)-4-chloropyrrole, is one of the earliest-known pyrrole alkaloids. It is an antifungal antibiotic produced by the bacterium Pseudomonas pyrrocinia as well as other organisms. It was first reported by Arima and co-workers [15] in 1964 and its biosynthesis from tryptophan was first investigated by Lively et al. in 1966 [16]. In the most recent work on pyrrolnitrin biosynthesis, Van Pee and coworkers [17] propose the biosynthetic

241

Recent Research on Pyrrole Alkaloids

Scheme 1

Scheme 2 pathway shown in Scheme 1. They have shown [17] that both chlorines of pyrrolnitrin are introduced via NADPH-dependent enzymes (see Scheme 2), rather than conventional, unspecific haloperoxidases. They propose the intermediacy of arene oxides such as 16 which can be attacked by CI ® before undergoing aromatization by dehydration of the resulting chlorohydrin.

242

P. W. LeQuesne, Y. Dong and T. A. Blythe

PRODIGIOSIN

Microorganisms producing the long-known metabolite prodigiosin (18) also give rise to a variety of cyclic alkylated derivatives, of which cycloprodigiosin (19) [18-19] is the simplest example. Cycloprodigiosins having larger alicyclic rings are also known, such as the recently described butyl-m-cycloheptylprodigiosin (20) from actinomycete strain B 4358 [20].

OMe

// \v// ^ \

0

(CH2)4 Me

OMe

^

18 OMe

5.

ALKALOIDS FROM Quararibea funebris

Funebrine (21) was the first alkaloid isolated from a plant of the family Bombacaceae. It was obtained from the flowers of the Mexican tree Quararibea funebris [21]. The biosynthetic pathway has been proposed as shown in Scheme 3 [21-23].

H Me

21

243

Recent Research on Pyrrole Alkaloids

Lactone 22 was postulated to combine with a six carbon, sugar-derived fragment such as 2,5diketogluconic acid. What follows is a series of nucleophilic additions, dehydrations, reductions, and aromatizations forming the pyrrole moiety and thus the funebrine congener and presumed precursor funebral (23). Another molecule of 22 condenses with 23 to afford the Schiff base, funebrine (21). Subsequently, Zennie et al. isolated and characterized funebral (23) [22] and funebradiol (24) [23] to lead credence to this biogenetic hypothesis.

H Me

O + HOH2C'

H 22

NH,

HO

HH

O CO2H OH

-^23

+22

21

2,5-Diketogluconic acid Scheme 3.

\JI 24 Several synthetic studies have been performed. The two characteristic amino lactone units of funebrine (21) are derived from a rare amino acid, (2S, 3S, 4R)-Y-hydroxyisoleucine. LeQuesne et al. [24] have described two approaches to this compound. In the first (see Scheme 4), the rran^-crotyl ester of ?-Boc-glycine was formed by DCC coupling. Then the ester/enolate Claisen rearrangement was realized in a >20:1 favorable stereoselectivity. Aminolactone 22 was obtained following iodolactonization, reduction, and hydrolysis. The second approach (see Scheme 5) employed a highly stereoselective addition of methyl, then amino equivalents to p-angelicalactone.

With access to the aminolactone 22, the total

synthesis of (±)-funebral (23) v^as accomplished by Yu and LeQuesne [25] (see Scheme 6).

244

P. W. LeQuesne, Y. Dong and T. A. BIythe

O O ^j^ B o c - H N ^ ^ Q ,;; LDA Boc-HN„ J L Q H Boc-NHCHoCOoH^^^-^^OH J '^^TMSCI , Me/ —rrr ^ ^ ^ - 7 8 - 2 0 ° C then DCC ^ 55-60°C H DMAP " i) CH2N2 ^-^ /) ^ O // ii) Phthalic // Y ^ o riH NL /w; I2, CH3CN

Y

r j b Me^

/() 6N HCI

Me-'^"

i

Me

1^

22 Hydrochloride

Scheme 4.

O a-Angelicalactone

O p-Angelicalactone Me.

-^

Me^

>NH2

Ml

Raney nickel

,^/^ > = 0 Me*^ O

Scheme 5. Me Me'

22

w //

Ti(0/Pr)4, toluene, heat Me

HIO4

! OHC^N

\J

CHO

HCOOH

Scheme 6.

Q^

245

Recent Research on Pyrrole Alkaloids

Using titanium (IV) isopropoxide as the cyclizing reagent, the Paal-Knorr condensation of 22 with 2,9-dimethyldeca-2,8-dien-4,7-dione occurred without polymerization. Alkene oxidation followed by reduction of one of the aldehydic functions yielded 23. Several other examples were given to show that this is a general synthetic procedure to produce 1,2,5-substituted pyrroles.

6.

PYRROLE ALKALOIDS FROM SPONGES

The chemical literature over the past thirty years has shown that sponges are a rich source of alkaloids of pharmacological, physiological, and biological importance. The pyrrole nucleus is well represented in sponge alkaloids, especially in combination with other functional groups as in pyrrololactams and pyrroloquinolines.

6.1.

Pyrrololactams

In 1980, Sharma et al.[26] isolated the yellow pyrrololactam 25 from the Great Barrier Reef sponge Phakellia flabellata. They suggested that 25 has a biosynthetic origin in conmion with a class of compounds isolated from this sponge called phakellins (see below). These compounds either possess or are derived from compounds possessing a "C^Ns" skeleton. Albizati and Faulkner [27] isolated and spectrally characterized a new metabolite, stevensine (26),

26 from an unidentified Micronesian sponge. This compound has the same 6,7-dihydropyrrolo[2,3c]azepin-8-one ring system. One can envision that the biosynthetic progenitors of these two alkaloids and this class of natural products might be oroidin (27), hymenidin (33) or similar compounds as depicted in Scheme 7 [28].

246

P. W. LeQuesne, Y. Dong and T. A. Blythe

Schmitz and co-workers [29] isolated two related pyrrololactams from several different sponges. Aldisin (28) and bromoaldisin (29) were isolated from Hymeniacidon aldis de Laubenfels of Guam and an unidentified sponge from Fiji. Specimens of a Lissodendoryx sp. sponge of Sri Lanka yielded bromoaldisin (29) and the pyrrole-2-carboxylic acid methyl esters (30) and (31). The aldisins are thought to be formed by oxidative degradation of the appropriately substituted guanidine 25. However, a second pathway could be the addition of H2O to the intemuclear double bond of 25 followed by reverse aldol loss of the guanidine function. Another pyrrole alkaloid which appears to be closely related to 25, 26, 27, 28, and 29 biogenetically has been described. Kobayashi et al [30] isolated hymenin (32) from a Hymeniacidon sp. collected at Ishigaki Island, Okinawa. In a succeeding communication they describe the biological activity of 32 as an alpha-adrenoceptor blocker [31]. The same group isolated and structurally characterized hymenidin (33) [32]. This compound should be a biosynthetic precursor to the cyclized class of compounds and an oroidin analogue. They demonstrated the antiserotonergic action of 33 on rabbit aorta. Van der Helm et «/.[33] isolated three pyrrololactams from the marine sponge P. fusca collected in the South China Sea. Their structures were determined by X-ray analysis, as well as spectroscopic means. One of them was named fuscin (34) [34].

HoN.

27

Xi=Br X2=Br

33

Xi=Br X2=H Scheme 7.

r ' N H

COgMe

o

Aldisin 28 X=H Bromoaldisin 29 X=Br

30 X=H 31 X=Br

247

Recent Research on Pyrrole Alkaloids

HN- '

H

O

Fuscin/Axinohydantoin

34 R=Br

Hymenialdisine

36 R=Br

Debromoaxinohydantoin

35 R=H

Debromohymenialdisine

37 R=H

Another group isolated the same compound from a different sponge, Monanchora from Papua New Guinea, and called it axinohydantoin (34) [35]. They also characterized three other related analogues, 35, 36, and 37. These alkaloids demonstrated moderate biological activity against lymphocytic leukemia in vitro.

248

6.2.

P. W. LeQuesne, Y. Dong and T. A. BIythe

Phakellins and Related Compounds

In 1988 and 1989, two groups isolated four different bromopyrroles, 38, 39, 40, and 41 from the deep water marine sponge, Agelas flabelliformis. Tada and Tozyo [36] characterized 2,3-dibromopyrrole (38) and 2,3-dibromo-5-methoxymethylpyrrole (39) by NMR and found Br \ ^ /r~\ Br-^,.>-Rp N 2

38 39 40 ^^

Ri=H R2=H Ri=H R2=CH20Me Ri=H R2=C02H f\^=:Me R2=C02Me

them to be very unstable. Gunasekera et ai [37] described 4,5-dibromo-2-pyrrollic acid (40) and the A^-methyl, methyl ester derivative (41). Their work shows that 40 suppresses the two-way murine mixed lymphocyte reaction in a dose responsive manner. Compound 40 has been previously isolated from Agelas oroides. All four of the compounds are analogues to the previously described bromopyrrolic acid esters 30 and 31. These simple pyrroles are mentioned here because they may arise from the more complex phakellins by degradation. The pyrrole-containing alkaloid, dibromoisophakellin (42), was isolated from the marine sponge Acanthella carteri collected off the Madagascar coast [38]. The structure was established by X-ray crystallography. The same group isolated and characterized dibromoagelaspongin hydrochloride (43) from the marine sponge Agelas sp. found off the coast of Tanzania [39]. These compounds contain the commonly found guanidine group and are structurally similar to all the previously shown pyrrololactams. However, the skeletal structures are unique and are thought to be related to phakellin (44) and hymenialdisine (36) [40-42]. The biosynthesis is postulated as shown in Scheme 8.

42

Recent Research on Pyrrole Alkaloids

249

k /

OH HN—[ H N ^ H2N-<\ \

Lr° -H20 i[0]

Scheme 8 The highly cytotoxic dibromophakellstatin (45) was isolated by Pettit and coworkers [43] from the Indian ocean sponge Phakella mauritiana and is clearly related to the guanidinium derivative dibromoisophakellin (42). During their study of Caribbean sponges, Faulkner and coworkers structurally elucidated the bromopyrrole alkaloid 46 from the sponge Agelas sceptum, collected at Glover Reef, Belize [44]. This compound demonstrated antimicrobial activity much greater than that of oroidin (14). Compound 46 could be biosynthetically derived from a photochemical [2+2] addition reaction between two molecules of hymenidin (33).

P. W. LeQuesne, Y. Dong and T. A. BIythe

250

Brs

// \^

NHp

P

HoN'^^NH

HN Br>

// V NH2 46 An alternative combination might give rise to the cyclohexane-containing ageliferins (such as 47) from Astroclera willeyana [45]. H2N Br>

Three novel styloguanidines, 48-50, were isolated from the sponge Stylotella aurantium collected in the Yap sea [46]. They are chitinase inhibitors that arrest the moulting process of the cyprid larvae of barnacles. Accompanying these compounds, palau'amine (50a), which had been isolated shortly before [46] by Scheuer and coworkers from S. agminata, was also obtained.

HN HgN-

^

N CI

CH2NH2

48; Ri=R2=H 49; Ri=Br, R2=H 50 ; Ri=R2=Br

CI 50 a

CH2NH2

Recent Research on Pyrrole Alkaloids

251

Recently, Japanese researchers isolated mauritiamine (51) from the marine sponge Agelas mauritiana collected off Hachijojima Island, Japan [47]. Both 51 and oroidin (27) were shown to inhibit metamorphosis of the barnacle Balanus amphitrite. Interestingly, dibromopyrrole metabolite 52 promoted larval metamorphosis of the ascidian Ciona savignyi. These workers postulate a possible biogenetic pathway of 51 from two molecules of oroidin (27) as shown in Scheme 9. H

0V-NH2 N H

•

^

' 52

From the related sponge A. clathrodes, Fattorusso and coworkers [48] have isolated the clathramides, such as 53. They have mild antifungal activity. These workers have also isolated the novel pyrrolopiperazine 54 from A. longissima [49].

Br>

NH

// V O

CO2 Me Me

e

53

CH3 54

A number of other oroidin-related metabolites have been discovered. Among them are the dispacamides 55 and 56, obtained by Fattorusso and coworkers from Agelas conifera, A. longissima, A. clathrodes and A. dispar [50], the taurine-containing compound mauritamide A (57) [51], from A. mauritiana , the non-brominated clathrodin 58, from A.clathrodes [52], keramadine 59 from an Okinawan Agelas species, and the manzacidins 60-62 [53] obtained from an Okinawan Hymeniacidon species and incorporating a tetrahydropyrimidine ring. Also related are the novel imides milpacamide (63) and its debromoderivative (64) [54].

P. W. LeQuesne, Y. Dong and T. A. BIythe

252

XO*

H r

>-NH2 N U

u oxidative^ ^N addition I >-NH2

oxidative^ dehydration

51

/ ^ 91 VNH2 rTTN

0OHH Scheme 9.

INF

Br-^

.AV H

55; R=Br 56; R=H

.N

O

H

BK^

^V Me

> NH

O

57

MeOsS

NH

Recent Research on Pyrrole Alkaloids

4

// w ^

253

NHo

HN'^S^

N

"•

H

O 58

O

Br> IV

BK^

1,

II

Mee -•-^ M Ri

COOR2

^

II

0 63; R=Br 64; R=H

60; Ri=R2=H 61 ; Ri=OH, R2=H 62 ; Ri=H, R2=Me

Agelas species are also noted for their production of "mixed biosynthesis" natural products incorporating pyrrole, terpenoid, and nucleic acid base moieties. Recent examples of these are ageline B (65) and agelasine G (66) [55]. NH2

65 ; R=H 66 ; R=Br In recent years Home and coworkers [56] have been developing chemical syntheses of the Cj 1N5 alkaloids. Stevensine (26) was synthesized from (±)-hymenin (32) (see Scheme 10). The key transformation in the sequence is a thermal transbromination of 4-bromohymenin (67) to create the olefin 26. In a full paper [57], the syntheses of (±)-hymenin (32), stevensine (26), hymenialdisine (36), and debromohymenialdisine (37) were reported. The synthesis of (±)-

254

P. W. LeQuesne, Y. Dong and T. A. Blythe

hymenin (32) is shown in Scheme 11. The synthetic imperative for this sequence was to suppress the self-dimerization commonly seen for pyrroles. Therefore, the bromines had to be introduced early in the synthesis. Reaction of 2,3-dibromo(trichloroacetyl)pyrrole with aminoethyldioxolane easily afforded 68. Removal of the acetal gave aldehyde 69. Cyclization under acidic conditions gave 70 without any dimer formation. Subsequent treatment with 2aminoimidazole proceeded smoothly to give (±)-hymenin (32). The penultimate step of the synthesis of stevensine (26), as expressed in Scheme 10, was the generation of 74, believed to involve the bridged bromonium ion 72 and/or the azafulvenium ion 73 (see Scheme 12). In accord with this mechanism, addition of bromine in methanol to 70 yielded 71. Subsequent coupHng of 75 with 2-aminoimidazole afforded 74 as the major product. When 74 was heated under acidic conditions, stevensine (26) resulted. For the syntheses of hymenialdisine (36) and debromohymenialdisine (37), Home and coworkers attempted to couple glycocyamine with 70 or 71 but failed. They then showed that 36 and 37 could be derived from 4-bromohymenin (67) by hydrolysis, protodebromination, and oxidation. Also for hymenialdisine (36), selective removal of the bromine atom at the P-position of the pyrrole ring was necessary. The experimental results are displayed in Scheme 13.

HpN 32

Bfp

^-^ CF3CO2H (95%)

//

HpN

H

6

Stevensine (26)

Scheme 10.

Recent Research on Pyrrole Alkaloids

255

Bfv

TsOH H20/acetone reflux 91%

B\r^K, H

CHO

N—, CHaSOgH^ rt. 7 days 80%

N' H CH3SO3H ft, 7 days 65%

// V Br^^ H

Hymenin (32)

Scheme 11. H,CQ

Br

f

Brj

NH

MeOH^ 20 min, rt B r - ^ ^ ^ ^ ^ H

95%

70

H

o 71

HgN-^ CH3SO3H Q^ * 90 °C ,, unsealed J/ 61% '

Scheme 12.

CH3SO3H

'^' ^^°'^°

g- and/or

Stevensine (26)

.N«

H,N^/1

P. W. LeQuesne, Y. Dong and T. A. Blythe

256 HoN-

+ 37 40%

aq. HBr 90 °C sealed

HN- '

21% .NH pj

M

67

72%^ reflux \

HPN-^^N^O

^

HNNH

Scheme

6.3.

CH3SO3H HBr (cat.) 90;C>. 36 + 37 sealed 12 h 33% 27%

13

Py rroloquinolines

In 1989, Sakemi and Sun isolated three novel alkaloids containing the 1,3,4,5tetrahydropyrrolo[4,3,2-J^]quinoline ring system [58]. These nitrogenous pigments were found in the deep water sponge, Batzella sp., from the Bahamas and were named batzellines A (76), B (77), and C (78). The structure of A (76) was determined by X-ray crystallography. The

R"! R2 Xi 76 77 78

Me H H H Me H

X2

SMeCI SMeCI H CI

structures of B (77) and C (78) were determined by NMR spectroscopy and chemical transformations. The authors compared these structures with dehydrobufotenine (79), isolated from the South American toad Bufo marinus, and discorhabdin C (80), produced by the sponges

257

Recent Research on Pyrrole Alkaloids

Latnmcula and Prianos. They suggested that batzelhnes and the tyrosine-derived sponge metabolite 81 are biosynthetic precursors of discorhabdins (such as compound 80, below).

The following year, Sun et al [59] isolated a new group of analogous alkaloids, isobatzellines A (82), B (83), C (84), and D (85) from the same Caribbean sponge. The isobatzellines have the same pyrrolo[4,3,2-c(e]quinoline ring as the batzellines. However, they also have an aminoiminoquinoline functionality different from the aminoquinoline of the batzellines. These compounds were cytotoxic against P388 leukemia cells and moderately antifungal against Candida albicans. However, the batzellines were neither antifungal nor cytotoxic.

SMe

HoN

X^ X2 82 SMe CI 83 SMe H 84 H CI

HpN-

Copp, Ireland, and Barrows reported the isolation of wakayin (86) from the ascidian Clavelina sp.[60]. The structure was proved by detailed 1 and 2D NMR studies. Wakayin (86) showed antimicrobial activity against Bac///w subtilus as well as cytotoxicity. It is claimed this is the first reported isolation of a pyrroloiminoquinoline from an ascidian, this class of compound usually being found in sponges. Therefore, the authors raised the question whether these natural products come from the ascidians and sponges or from microbes within these organisms. They

P. W. LeQuesne, Y. Dong and T. A. Blythe

258

suggested that isobatzellines are biosynthetically derived from tryptamine and that wakayin (86) incorporates a further tryptamine unit. Related to wakayin are compounds 88-91 from an unidentified South African latruncuhd sponge [61].

86

88 ; R=H 89; R=Me

90; Ri=Br, R2=R3=0 91 ; Ri=Br, R2=0H, R3=H

Epinardins A-D, (92-95) obtained from an unidentified southern Indian Ocean sponge, [62] are recently described members of this group of compounds. Epinardin C (94) is strongly cytotoxic towards doxorubicin-resistant L1210/DX tumor cells in vitro. Ireland and coworkers have described a series of iminoquinonoid compounds, the makaluvamines, from Zyzzya species of the sponge order Poecilosclerida [63-65]. These compounds, represented by structures 96-101, owe their cytotoxicity to topoisomerase 11inhibition.

Recent Research on Pyrrole Alkaloids

259

O

DMe

95

94

R. NHR4

96 97 98 99 100 101

Me H H Me H H

H H H H H H

H H H H H Br

H Me a b H H

a= ,11

OH

b= OH

Recently there has been synthetic effort in this area. Joule et al reported the formal total synthesis of batzelline C (78), isobatzelline C (84), discorhabdin C (80), and makaluvamine D (87) [66]. Their approach began with a quinoline rather than an indole. Scheme 14 shows the conversion of 6,7-dimethoxy-4-methyl-5-nitroquinoline to compound 102. This was then transformed to 103. Compound 103 has been converted to batzelline C (78) or isobatzelline C (84) in two steps by Yamamura et al. [67]. Transformation of 102 to 104 was simply carried out. The Yamamura group also converted 104 into discorhabdin C (80) in two steps [67], and into makaluvamine D (87) in one step [68].

P. W. LeQuesne, Y. Dong and T. A. BIythe

260 NO2 CH3

NO2 CH3

^

N

CI

NO2 CHO

DMSO 80 °C ^«°/°

NO2 CH(0Me)2

*

CI

NH2 CH(0Me)2

NiCl2 6Hr20

JdCt

,, _ MeO

if

1

I

MeOH, 0 °C M e O - ' \ f ^ K i ^ 90% I H

MeO

aq. 1NHCI THF

HCOOH, AC2O, rt^ 50% OHCN-

aa. NaOH MeOH, CH2CI2

MeOv

CHO

CHO

rt 82%

CHO

THF, rt 56%

aq. 2N NaOH '^®° reflux 950/0 MeO

Scheme 14.

In the full paper [69], Joule and coworkers described the formal total synthesis of damirones A (105) and B (106), batzelline C (78), isobatzelline C (84), discorhabdin C (80), and makaluvamines A-D (107, 108, 109, and 87 ). Scheme 15 demonstrates the retrosynthetic strategy. A substituted 4-methylquinoline A can be built into 5-amino-l,2,3,4-tetrahydroquinoline-4-aldehyde B. Cyclization to form the pyrrole ring system leads to the desired heterocycle.

261

Recent Research on Pyrrole Alkaloids

RiN-

RiN-

RoHN Ri

R2 R3

105 Me H Me 106 H H Me 78 Me CI H

84 107 108 109 87

Me H H H H

H H H H 4-HO-C6H4(CH2)2

CI H H 3,4-clehydroH N+5-Me H

RiM

NHR1CHO

I

?

^

110 H 111 Ci 112 H 113 H

CI CI CI H

Ci CI CI CI

CI H H H

P. W. LeQuesne, Y. Dong and T. A. BIythe

262

6.4.

The Phorbazoles

Kashman and his coworkers [70] have obtained a unique series of chlorinated phenylpyrrolyloxazoles from the Indo-Pacific sponge Phorhas aff. dathrata, collected in Sodwana Bay, South Africa. These compounds, named phorbazoles A-D, have been assigned structures 110-113, respectively.

6.5.

Miscellaneous Pyrroles from Sponges

Pseudoceratine (114), obviously derived from spermidine and the well-known 4,5dibromopyrrole-2-carboxylic acid, is a constituent of Pseudoceratina purpurea that inhibits larval settlement and metamorphosis of the barnacle Balanus amphitrite [71].

114 In a search for naturally-occurring aldose reductase inhibitors potentially useful in the treatment of diabetes, Sato and coworkers isolated three novel compounds, 115-117, from a Dictyodendrilla species [72]. COgMe

115;R=S03Na 116;R=S03H

117

Recent Research on Pyrrole Alkaloids

263

7.

PYRROLE ALKALOIDS FROM ASCIDIANS

7.1.

Eudistomines

Eudistomines are a class of natural products containing either the substituted p-carboline ring system as shown in structure 118 or the p-carboline/oxathiazepine condensed ring system of 119. This class of natural products is active against Herpes simplex virus, type 1 (HSV-1). Compounds having the skeleton of 119 are considered to be biosynthetically derived from tryptophan and cysteine [73].

118

This class of compounds has been recently reviewed [74]. We mention eudistomin A (120) and M (121) here because of the 2-pyrrolyl substituents on the P-carboline ring system of their structures. Kobayashi, Rinehart et al [75] reported the isolation and structural elucidation of eudistomin A (120) and M (121) from the colonial Caribbean tunicate Eudistoma olivaceum in 1984.

R R., R2 R3 R4 120 H OH Br H 2-pyrrolyl 121 H OH H H 2-pyrrolyl

R5 H H

Several years later, the Rinehart group published a full paper describing the synthesis of some of the eudistomins, one of them being eudistomin M (121) [76]. As shown in Scheme 16, the starting material was 5-methoxytryptamine (122). Esterification, conversion to the

264

P. W. LeQuesne, Y. Dong and T. A. BIythe

carboxamide, catalytic dehydrogenation, then dehydration gave the key l-cyano-6-methoxy-Pcarboline intermediate 123. Grignard reaction and cyclization yielded O-methyleudistomin M (125). Demethylation was carried out by boron tribromide to afford eudistomin M (121). MeO,

MeO,

%

MeOH

ylTV-/ N H

122

^

NH4OH

CO2CH3

MeQ i j \ ^ ^ ^ N ^

H

NH

xylene,

( i J \ , , N

reflux

^^'^N'^

CONH2 70%

H

65%

CONH2

MeO,

MeQ \ s = < >-...^'^ H NH4OAC HOAc reflux 67%

THF. 0 °C

CN

ii) H"- 75%

123 MeQ

124 BBrg N M'

H

125

\

VNH

^1^2^12 reflux

72%

^ ^

Eudistomin M (121)

Scheme 16.

Molina et al [77] have developed an aza-Wittig strategy to synthesize 1-substituted pcarboline derivatives. Iminophosphorane (126), prepared from 3-formyl-l-methylindole by treatment with ethyl azidoacetate then triphenylphosphine, reacted with 2-formylpyrrole in toluene at 160 °C in a sealed tube to yield compound 127 having the carbon skeleton of eudistomins A and M. From a newly discovered ascidian, Polycitor sp., found in Sodwana Bay, South Africa, Kashman and coworkers [78] have isolated a series of heavily brominated aryl-substituted pyrroles, 128-130. These compounds may be biogenetically related to the lamellarins (q. v.), and questions of their biogenesis and biological roles are still unanswered.

Recent Research on Pyrrole Alkaloids

_ / /

/COgEt

265

.COgEt

O^

CHO N H toluene 160 °C, sealed

• ^

70%

127

Scheme 17.

BrBr.

BrBi

OH 129; R=H 130; R=Me

8.

PYRROLE ALKALOIDS FROM MOLLUSCS

8.1.

Lamellarins

Lamellarins are metabolites of the marine prosobranch moUusk Lamellaria sp. Faulkner et al. [79] isolated four of these pyrrole-containing aromatic alkaloids, lamellarins A-D (131-134), from six specimens of this sponge collected near Koror, Palau. The structure of lamellarin A (131) was determined by X-ray crystallography. Characterization of B-D (132-134) was accomplished by NMR spectral studies. The authors claimed that a majority of metabolites from mollusks can be traced to dietary sources. Nothing was known of the diet of this species, but

266

P. W. LeQuesne, Y. Dong and T. A. BIythe

Other species are believed to feed on colonial tunicates. However, the lamellarins do not resemble any known metabolites or marine natural products, and their biosynthetic pathway is unknown.

R 131 H 133 H

X OH H

R"i R2 R3 R4 R5

R Ri X 132 H Me OMe 134 H H H

Re X

138

135 H Me Me H Me Me OH 136 H Me Me Me Me Me OH 137 MeH Me H Me H H Several years later, Fenical and Lindquist in collaboration with Clardy and VanDuyne [80] isolated lamellarins E-H (135-138). These compounds were found in the ascidian Didemnum chartaceum which was collected near Aldabra Atoll, Republic of the Seychelles in the Indian Ocean. Lamellarin E (135) was assigned its structure based on X-ray data. The rest were studied by 2D NMR. Prosobranch moUusks are vulnerable because they have reduced shells covered by a mantle of flesh. They are believed to feed on chemically rich ascidians and to use these dietderived compounds as chemical defense against predators.

Recent Research on Pyrrole Alkaloids

9.

267

MISCELLANEOUS MARINE PYRROLE ALKALOIDS

As well as the marine pyrrole alkaloids outlined above, a few novel pyrrole alkaloids have been obtained that do not fit into previously delineated structural categories.

9.1.

Triketramine

In 1990, Aknin et al [81] discovered an unusual pyrrole alkaloid, triketramine (139), in the sponge Trikentrion loeve found at Thiouri-ba, Dakar, Senegal. The structure was elucidated by spectroscopic and X-ray methods. This skeleton is believed to be completely new for either marine or terrestrial organisms; no biosynthetic pathways have been postulated.

CHq

9.2.

CH3

Rhodophycan Pyrrole Alkaloids

Gerwick et al [10] isolated two relatively simple pyrroles, 5 and 6, from the red alga Gracilariopsis lemaneiformis collected in intertidal pools at Cape Perpetua, Oregon. Since the extractive process involved acetic anhydride, they were not certain whether the natural products were the acetates (see structures 140-141) or the alcohols. Several years later, Jefford et al [82] synthesized 140 and 141 as shown in Scheme 18. Aminoalcohols 142 or 143 were condensed with 2,5-dimethoxytetrahydrofuran to give the A^-substituted pyrrole 144 or 145 which was then acylated to afford 146 or 147. Intramolecular acylation promoted by boron tribromide gave acetoxyacetylpyrrole 148a or b. The final acylation provided pyrroles 140 and 141. NMR data were consistent with those of the natural products. However, the optical rotation of 141 was low. This is claimed to be the result of partial racemization in the methanolic solution during measurement or in the aqueous methanol eluent of the final purification stage.

p. W. LeQuesne, Y. Dong and T. A. Blythe

268

N

COCHgOR

^N

s

OR 6 R=H 141 R=COMe

OR 5 R=H 140 R=COMe

NH OH

MeO

XX

R 142R=H 143R=Me

o

COCH2OR

HOAc 120 °C^ OMe ^^2 h 65-69%

Q

CICOCH2OAC EtN(/-Pr)2 DMAP 87-95% OH

R 144 R=H 145 R=Me BBrg L/n2Vi/l2

^N

COCH2OAC

rk^oH

k^^OCOCHgOAc"^ to -5 °C 65-70% R 146 R=H 147 R=Me

R 148a R=H 148b R=Me

AcCI EtN(/-Pr)^ DMAP 70-94%

140 or 141

Scheme 18.

9.3.

Tambjamines and Related Compounds

Tambjamines are substituted bipyrrole alkaloids occurring in bryozoans, nudibranchs, and ascidians [83-85]. They have the generalized stracture 149, where R^ represents an alkyl or arylalkyl side chain. They are yellow to yellowish-brown pigments located in the granular amebocyte blood cells of tambjamine-producing ascidians [84], and they become concentrated in the nudibranches which are the ascidians' predators. Associated with the tambjamines both in some of the ascidians and their associated nudibranches is a tetrapyrrole pigment also known as the Bugula pyrrole (see below). In recent years the new tambjamines 150-153 have been obtained from the Tasmanian bryozoan Bugula dentata [85], together with the blue "Bugula pyrrole". Related to the tambjamines is the bipyrrole 154, derived from a terrestrial Streptomyces sp. BE 18591 [86]. The variety of organisms in which tambjamines are found raises questions of possible biosynthesis by

Recent Research on Pyrrole Alkaloids

269

symbiotic microorganisms, but Lindquist and Fenical [84] believe that at least in the ascidian Atapozoa this animal produces them itself.

PMe

NHR2 149; Ri, R3=HorBr

150 151 152 153 154

Ri

Ra

R;

Br Br Br Br H

0H20rlQ CH2CH2Cng Crl2Cn(CH3)2 OH2CH(Cn3)CH2CH3 C12H25

H H H H H

Japanese authors discovered a brilliant blue pigment 155 from the bryozoan Bugula dentata Ramouroux found in the Gulf of Sagami, Japan [87]. This alkaloid is antimicrobial against Grampositive and -negative bacteria. The structure was solved spectroscopically and is thought to be identical with a tetrapyrrole isolated from a mutant strain of Serratia marcescens. Therefore, it is possible that either the bryozoan or microorganisms associated with the bryozoan may biosynthesize 155. It could also be derived from prodigiosin-producing bacteria in the diet.

OMe QMe M H

9.4.

N H^Cr 155

N H

N H

Pyrrolomycins and Pyrroindomycins

Over the years, bacterial strains have been a valuable source of alkaloids when fermented in broths. One such class of alkaloids that contains the pyrrole moiety is the pyrrolomycins. In 1981, the isolation, structure identification and biological activity of the new antibiotics pyrrolomycins A (156) and B (157) were reported [88-90]. In 1983, the same group isolated three new members C (158), D (159), and E (160) [91]. All five of these alkaloids were isolated from Actinosporangium vitaminophilum SF-2080. Pyrrolomycin C (158) and E (160) are active against Gram-positive bacteria and D (159) has a broad spectrum including Gram-positive and negative, bacteria, and fungi. Structures of these compounds were based on spectroscopic data. X-ray analysis, and synthetic correlations.

270

p. W. LeQuesne, Y. Dong and T. A. BIythe

CI Ck CI

.NO,

'I \

Pyrrolomycin A(156)

Pyrrolomycin B(157) CI

CI

O

O

OH

Pyrrolomycin C (158)

CI- - . . /

OH

Pyrrolomycin D (159)

-^

-CI

OH Pyrrolomycin E (160) Several years later a similar compound, pyrroxamycin (161), was isolated from the culture broth of Streptomyces sp. S46506 [92]. Its structure was also assigned based on NMR analysis and synthetic correlation. Pyrroxamycin (161) was active against Gram-positive bacteria and dermatophytes. However, it was not active against Gram-negative bacteria and yeasts. One can postulate that this compound arises from acetal formation from the pyrroxamycin diol (derived from 158) and formaldehyde. In 1990, Masuda et a/. [93] published the isolation of an actinomycete metabolite which they designated HS3. This compound was found to be identical with 161. Their work was focused on a search for microbial modulators of the activity of substance P (SP), a neuropeptide involved in inflammatory reaction. They established a screen based on SP-induced myeloperoxidase (MPO) release from human polymorphonuclear leukocytes (PMN). As a result of the screening, they found that pyrroxamycin (161) and all the related pyrrolomycin antibiotics had an effect on SP-induced MPO release.

Recent Research on Pyrrole Alkaloids

271

Pyrroxamycin (161) These researchers speculate whether pyrrolomycins act as antagonists of SP in the nervous system. Because baclofen [p-(4-chlorophenyl)-GABA] is a known SP antagonist and GABA is not, the authors suggested that the 4-chlorophenyl moiety, which is present in pyrrolomycins, is critical for the antagonistic activity to SP. The pyrroindomycins are analogous to the pyrrolomycins where the 2- and 3- positions of the pyrrole ring are substituted with an indole group. In 1994 two sequential articles were published about the isolation, structure determination, and biological activities of the first natural products containing the highly unsaturated pyrrolindole moiety [94-95]. Pyrrolindomycins A (162) and B (163) were found to be two principal components of the antibiotic complex isolated

OH

HoN /

'

Pyrroindomycins A (162) R=H B (163) R=CI

from fermentations of culture LL-42D005, which is a strain of Streptomyces rugisporus. Their structures were determined by using 1- and 2-D NMR, mass spectroscopy, and chemical degradations. These two compounds demonstrated excellent in vivo activity against Gram-positive bacteria. It is believed that the mode of action is interference with the integrity of the bacterial membrane.

P. W. LeQuesne, Y. Dong and T. A. Blythe

272 9.5.

Pyrrolethers

Another class of antibiotics containing the pyrrole ring system is the pyrrolethers. Westley et al [96] isolated antibiotic X-14885A (164) from Streptomyces antihioticus. X-14885A (164) was active against Gram-positive bacteria and the spirochete, Treponema hydrodysenteriae. Its structure was elucidated by X-ray crystallographic analysis.

X-14885A

9.6.

(164)

Keronopsins A and B, Protozoan Chemical Defense Agents

The ciliate Pseudokeronopsis rubra is colored deep red and exerts a powerful combination of antibiotic, deterrent, and antifeedant effects. Hofle and coworkers have recently shown that these activities arise from conjugated pyrrole derivatives 165-168 (keronopsins Al, A2, B, and B2) [97].

165 166 167 168

SOgNa SOgNa H H

H Br H Br

Compounds 165 and 166 are unstable in the crude extract, being converted into 167 and 168, respectively, and are very prone to polymerization under acidic conditions or in the solid state.

273

Recent Research on Pyrrole Alkaloids

10.

MISCELLANEOUS PYRROLES FROM MICROORGANISMS

The biosynthetic fecundity of microorganisms is seen in the varied structures of pyrrole derivatives in this section. The pyrrole tetraene rumbrin (169), somewhat resembling the keronopsins (see above), was isolated [98] from Auxarthron umbrinum. It has cytoprotective

169 properties, as do the thiazohalostatins 170 and 171 isolated from Actinomadura sp. HQ24 by the same Kirin Brewery group [99]. The chlorine-containing 170 was replaced in the cultures by 171 when the medium was enriched with KBr.

Me OH

O

Me

Me COOH

170;R=CI 171 ; R=Br

Another thiazole-containing pyrrole metabolite is BE10988 (172), a topoisomerase II inhibitor, which was isolated from a culture broth [100].

274

P. W. LeQuesne, Y. Dong and T. A. Blythe

,COOH

HoN.

MegN

172 NMeg

173

From Streptomyces rubropurpureus No. 6362 Seto and coworkers [101] isolated the antitumor antibiotic chromoxymycin (173). This structure is unusual in having an anthraquinonetype nucleus substitited by a novel A^-hydroxypyrrolyl fragment at one of the central potential carbonyl positions. A series of fused pyrrole metabolites, the pyralomycins, 174-180, was obtained [102] from Actinomadara spiralis. They show antibiotic activity against Micrococcus luteus.

H H H CI

CI Me CI CI

Me CI Me Me

Me Me H H

178 H 179 H 180 H

CI Me CI

Me CI Me

Me Me H

174 175 176 177

275

Recent Research on Pyrrole Alkaloids

A novel macrolide antibiotic, pyrrolosporin A (181), having the lactone function incorporated into a spiro-fused tetronic acid fragment within the macrocycle, was isolated [103] from a Micromonosporium species (strain C39217-R109-7) obtained from a Puerto Rican soil sample. Pyrrolosporin prolongs the life-span of mice inoculated with P388 leukemia cells.

COOH HQG

Pyrrolosporin A 181

From the myxomycete (slime mould ) Lycogala epidendrum, Asakawa and coworkers [104] have identified three bisindolic pyrroles, 182-184, named lycogarubins A-C, respectively. Compound 184 showed anti-HSV-I virus activity in vitro.

Ri

182 H 183 H 184 H

OH OH H

OH H H

276

P. W. LeQuesne, Y. Dong and T. A. Blythe

Two interesting pyrrole derivatives have been obtained recently from mushrooms. From the poisonous mushroom Clitocybe acromalalga, the amino acid L-3-(2-carboxy-4-pyrrolyl)alanine (185) was isolated [105]. Its structure was supported by synthesis and a biogenetic origin from DOPA was proposed.

HOOC

^2^

COOH 185

HaCO' 186

As part of an investigation of the pungent principle of the mushroom Tricholoma sciodes, the novel "bisindole" sciodole (186) was isolated by Sterner [106]. This intriguing structure is of an apparently unique type.

11.

MINOR PIPER ALKALOIDS CONTAINING A PYRROLE SKELETON

The species Piper sarmentosum Roxb. has found much medicinal use. This plant is found in Malaya, the Indonesian archipelago, and Thailand. Likhitwitayawuid et al [107] reported the isolation of six components from the dried fruit of P. sarmentosum.

(CH2)iiCH3

o

187

188

277

Recent Research on Pyrrole Alkaloids

Pellitorine (189)

Sarmentine (190)

Sarmentosine (191)

These are, as well as [i-sitosterol, the aromatic alkene 187, the pyrrole amide 188, pellitorine (189), and two unsaturated pyrrolidine amides, sarmentine (190) and sarmentosine (191). Relevant to this review is compound 188. The structure of this alkaloid was confirmed by comparison with a synthetically derived sample prepared as shown in Scheme 19. 3Phenylpropanoic acid was converted to the acid chloride which was then treated with anion of pyrrole to yield 188 in 77% yield. To the authors' knowledge, 188 had never been synthesized or isolated from natural sources, but they stated that 3-phenylpropanoic acid has been previously found in the leaves of P. sarmentoswn.

CO2H /; (C0CI)2

.0

N Li Scheme 19.

188

278

12.

P. W. LeQuesne, Y. Dong and T. A. Blythe

MICELLANEOUS PYRROLES FROM HIGHER PLANTS

Three oligopeptide pyrrolamides, asterinins A-C (192-194), respectively, were isolated from Aster tataricus roots [108].

The inclusion of p-phenylalanine in these structures is

interesting. The epilupinine esters 195 and 196 were isolated in a detailed study [109] of the alkaloids of Virgilia divaricata and V. oroboides from South Africa.

OH H N^^COOR C2H5

192; R=H 193; R=Me

N^ ^COOMe H,C-

OH

194

CH2OR1 .0R2 195 H 196 a-C4H4N-C0

a-C4H4N-CO H

The fascinating "dimeric" alkaloid exochomine (197) affords the European ladybird Exochomus quadripustulatus chemical protection against predators [110]. Two other fused-ring pyrroles from higher plants are the unique monomargine (198), an orange pigment from the Malaysian annonaceous plant Monocarpia marginalis [111], and the rhazinilam relative 199,

Recent Research on Pyrrole Alkaloids

279

obtained from cell cultures of Aspidosperma quebracho bianco [112]. Compounds 198 and 199 are likely formed in the metabolic degradation (catabolism) of other alkaloids.

199

197

13.

ROSEOPHILIN

Roseophilin (200) is a metabolite isolated from a culture broth of Streptomyces griseoviridis [113]. This pyrrole-containing compound exhibited cytotoxicity against several human epidermoid and leukemia cell lines in the submicromolar range. This compound has been the subject of much synthetic effort recendy [114-118]. We have selected the synthetic approach of Furstner and Weintritt in a very recent publication [117] for review here. The retrosynthetic strategy, as demonstrated in Scheme 20, was based on the synthesis of compound 201 and its subsequent condensation to the previously known pyrrole 202. To this effect, compound 203 (where R=benzyl) was synthesized in ten efficient steps (see Scheme 21). The structure of 203 was assigned based on extensive 2D NMR studies. The synthesis of 203 was based on a palladium-catalyzed macrocyclization reaction as depicted in Scheme 22. This sequence relies on the differences in reactivity of varied allyhc precursors in palladium catalyzed substitution reactions. The completion of the total synthesis is in progress.

280

P. W. LeQuesne, Y. Dong and T. A. BIythe

MeO. MeO

Roseophilin (200)

201 203

202

R=H R=benzyl

Scheme 20.

0^0-^ 216 214

215

kyd-'& O^

OH

217

O'-" "OH

218 Scheme 22. (see next page for Scheme 21).

219

OH

Recent Research on Pyrrole Alkaloids

HO

281

TBDMSCI

CI

ilci^TrT" CH2V-'i2

II 205

90%

204

BFa TBS<

ii) tetrahydrothiophene, AgBF4, acetone, rt 73%

/•) f-BuLi ii) 9-bromononanal, THF, -78 °C - rt 84%

s*206

^^Y^

SOgPh TBSO

'"cO.Me ,-^^^Q' KH, DMF, rt 68% 207

PhOgS

[^

208

Pd(PPh3)4(10mol%) dppe(20mol%) ' " ^ T ' ^ THF, reflux PhOgSM. \ HO 85% MeO O OTBS 209

TBAF, NH4J= THF, rt PhOgS 63%

210

Pd(PPh3)4(15mol%) BnNHg PhOpS THF, 35 °C 70%

Dess-Martin^ CH2CI2 PhOaS rt, 83% 211

H

NMeo

C! . ii) SnCU, CHaCIa rt, 71%

/-PrMeaZnMgCI excess f-BuOK THF, rt 47%

PhOgS

203

213

Scheme 21.

282

14.

P. W. LeQuesne, Y. Dong and T. A. Blythe

CONCLUSION

The pyrrole alkaloids still represent a relatively small group of compounds. The increase in their numbers during the last fifteen years owes much to the increase in investigations of bacterial metabolites and marine organisms. We can expect many further discoveries of novel compounds from these sources, as well as from land plants. The Quararibea alkaloids are still, to our knowledge, the only alkaloids so far from the entire family Bombacaceae.

15.

ACKNOWLEDGMENTS We are grateful to a referee for helpful conmients.

REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

KPC VoUhardt, Organic Chemistry, W. H. Freeman and Company, New York, 1987. IRC Bick, Alkaloids from Australian Flora. In Alkaloids: Chemical & Biological Perspectives', Vol.10, SW Pelletier, Ed., Pergamon, New York, 1996, pp 1-154. PR Burkholder, RM Pfister, and FH Leitz, Applied Microbiology 14, 649 (1966). K Wiesner, Z Valenta, and JA Findlay, Tetrahedron Lett. 8, 221 (1967). T Tokuyama, J Daly, and B Witkop, 7. Am. Chem. Soc. 91, 3931 (1969). R Emrich, H Weyland, and K Weber, J. Nat. Prod. 53, 703 (1990). M Ito, H Shimura, N Watanabe, M Tamai, A Takahashi, Y Tanaka, I Arai, and K Hanada, Agric. Biol. Chem. 55, 2117 (1991). M Schonewolf, S Grably, K Hiitter, R Machinek, J Wink, A Zeeck, and J Rohr, Liebigs Ann. Chem.ll (1991). AL Waterhouse, I Holden, and JE Casida, /. Chem. Soc. Chem. Commun. 1265 (1984). ZD Jiang, and WH Gerwick, J. Nat. Prod 54, 403 (1991). MG Reinecke, and YY Zhao, J. Nat. Prod. 51, 1236 (1988). M Haidoune, R Momet, and M Laloue, Tetrahedron Lett. 31,1419 (1990). S Kato, K Shindo, H Kawai, A Odagawa, M Matsuoka, and J Mochizuki, J. Antibiotics 46, 892 (1993).

Recent Research on Pyrrole Alkaloids

14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40.

283

T Nogami, Y Shigihara, N Matsuda, Y Takahashi, H Naganawa, H Nakamura, M Hamada, Y Muraoka, T Kakita, Y litaka, and t Takeuchi, J, Antibiotics 43,1192 (1990). K Arima, H Imanaka, M Kousaka, A Fukuda, and G Tamura, Agr. Biol Chem. 575 (1964). DH Lively, M Gorman, ME Haney, and JA Mabe, Antimicrobial Agents and Chemotherapy 462 (1966). K Hohaus, A Altmann, W Burd, I Fischer, PE Hammer, DS Hill, JM Ligon, and KH Van Pee, Angew. Chem. Int. Ed. Engl. 36, 2012 (1997). NN Gerber, Tetrahedron Lett. 24, 2797 (1983). H Laatsch, and RH Thomson, Tetrahedron Lett. 24, 2701 (1983). H Laatsch, M Kellner, and H Weyland, J. Antibiotics 44, 187 (1990). RF Raffauf, TM Zennie, KD Onan, and PW LeQuesne, /. Org. Chem. 49, 2714 (1984). TM Zennie, JM Cassady, and RF Raffauf, J. Nat. Prod. 49, 695 (1986). TM Zennie, JM Cassady, J. Nat. Prod. 53, 1611 (1990). SL Ablaza, NN Pai, and LeQuesne, Nat. Prod. Lett. 6, 77 (1995). S-X Yu, and PW LeQuesne, Tetrahedron Lett. 36, 6205 (1995). GM Sharma, JS Buyer, and MW Pomerantz, J. Chem. Soc, Chem. Comm. 435 (1980). KF Albizati, and DJ Faulkner, J. Org. Chem. 50,4163 (1985). G DeNanteuil, A Ahond, J Guilhem, C Poupat, E Tran Huu Dau, P Potier, M Pusset, J Pusset, and P Laboute, Tetrahedron 41, 6019 (1985). FJ Schmitz, SP Gunasekera, V Lakshmi, LMV Tillekeratne, /. Nat. Prod. 48,47 (1985). J Kobayashi, Y Ohizumi, H Nakamura, Y Hirata, K Wakamatsu, and T Miyasawa, Experientia 42, 1064 (1986). J Kobayashi, H Nakamura, and Y Ohizumi, Experientia 44, 86 (1988). J Kobayashi, Y Ohizumi, H Nakamura, and Y Hirata, Experientia 42, 1176 (1986). L Zeng, X Fu, J Su, F De Guzman, FJ Schmitz, BM Hossain, and D Van der Helm,CA 115: 203706a (1991). X Fu, L Zeng, J Su, FJ Schmitz, BM Hossain, and D Van der Helm, CA 118: 36148y (1993). G Groszek, D Kantoci, and GR Pettit, Liebigs Ann. 715 (1995). H Tada, and T Tozyo, Chem. Lett. 803 (1988). SP Gunasekera, S Cranick, and RE Longley, J. Nat. Prod. 52, 757 (1989). SA Fedoreyev, NK Urkina, SG Ilyin, MV Reshetnyak, and OB Maximov, Tetrahedron Lerr. 27, 3177(1986). SA Fedoreyev, SG Ilyin, NK Urkina, OB Maximov, and MV Reshetnyak, Tetrahedron 45, 3487 (1989). GM Sharma, and PR Burkholder, J. Chem. Soc. Chem. Comm. 151 (1971).

284

41. 42. 43. 44. 45. 46.

47. 48. 49. 50. 51. 52. 53. 54. 55. 56. 57. 58. 59. 60. 61. 62. 63. 64.

P. W. LeQuesne, Y. Dong and T. A. BIythe

GM Sharma, and B Magdoff-Fairchild, /. Org. Chem. 42, 418 (1977). I Kitagawa, M Kobayashi, K Kitanaka, M Kido, and Y Kyogoku, Chem. Pharm Bull. 31, 2321 (1983). GR Pettit, J McNulty, DL Herald, DL Doubek, JC Chapuis, JM Schmidt, LP Tackett, and MR Boyd, J. Nat. Prod. 60, 180 (1997). RP Walker, DJ Faulkner, D Van Engen, and J Clardy, J. Am. Chem. Soc. 103, 6772 (1981). DH Williams, and DJ Faulkner, Tetrahedron 52, 5381 (1996). T Kato, Y Shizuri, H Izumida, A Yokoyama, and M Endo, Tetrahedron Lett. 36, 2133 (1995); RB Kinnel, H-P Gehrken, and PJ Scheuer, J. Am. ^Chem. Soc. 115, 3376 (1993). S Tsukamoto, H Kato, H Hirota, and N Fusetani, J. Nat. Prod. 59, 501 (1996). F Cafieri, E Fattorusso, A Mangoni, and O Taglialatela-Scatati, Tetrahedron 52, 13713 (1996). F Cafieri, E Fattorusso, A Mangoni, and O Taglialatela-Scatati, Tetrahedron Lett. 36, 7893 (1995). F Cafieri, E Fattorusso, A Mangoni, and O Taglialatela-Scafati, Tetrahedron Lett. 37, 3587 (1996). C Jimenez, and P Crews, Tetrahedron Lett. 35, 1325 (1994). JJ Morales, and AD Rodriguez, J. Nat. Prod. 34, 629 (1991). H Nakamura, Y Ohizumi, J Kobayashi, and Y Hirata, Tetrahedron Lett. 25, 2475 (1984); J Kobayashi, F Kanda, M Ishibashi, and H Shigemori, J. Org. Chem. 56, 4574 (1991). R Fathi-Afshar, and TM Allen, Canad. J. Chem. 66, 45 (1988). K Ishida, M Ishibashi, H Shigemori, T Sasaki, and J Kobayashi, Chem. Pharm. Bull., 40, 766 (1992). Y-Z Xu, K Yakushijin, and DA Home, Tetrahedron Lett. 37, 8121 (1996). Y-Z Xu, K Yakushijin, and DA Home, J. Org. Chem. 62, 456 (1997). S Sakemi, and HH Sun, Tetrahedron Lett. 30, 2517 (1989). HH Sun, S Sakemi, N Burres, and P McCarthy, J. Org. Chem. 55, 4964 (1990). BR Copp, CM Ireland, and LR Barrows, J. Org. Chem. 56, 4596 (1991). GJ Hooper, MT Davies-Coleman, M Kelly-Borges, and PS Coetzee, Tetrahedron Lett. 37, 7135 (1996). M D'Ambrosio, A Guerriero, G Chiasera, and F Pietra, Tetrahedron 52, 8899 (1996). DC Radisky, ES Radisky, LR Barrows, BR Copp, RA Kramer, and CM Ireland, J. Am. Chem. Soc. 115, 1632 (1993). LR Barrows, DC Radisky, BR Copp, DS Swaffar, RA Kramer, RL Warters, and CM Ireland, Anticancer Drug Design 8, 333 (1993).

Recent Research on Pyrrole Alkaloids

65. 66. 67. 68. 69. 70. 71. 72. 73. 74.

285

DA Venables, GP Concepcion, SS Matsumoto, LR Barrows, and CM Ireland, J. Nat. Prod. 60, 408 (1997). D Roberts, M Alvarez, and JA Joule, Tetrahedron Lett 37,1509 (1996). S Nishiyama, J-F Cheng, S Yamamura, and XL Tao, Tetrahedron Lett. 32, 4151 (1991) T Izawa, S Nishiyama, and S Yamamura, Tetrahedron 50, 13593 (1994). D Roberts, JA Joule, MA Bros, and M Alvarez, J. Org. Chem. 62, 568 (1997). A Rudi, Z Stein, S Green, I Goldberg, Y Kashman, Y Benayahu, and M Schleyer, Tetrahedron Lett. 35, 2589 (1994). S Tsukamoto, H Kato, H Hirota, and N Fusetani, Tetrahedron Lett. 37, 1439 (1996). A Sato, T Morishita, T Shiraki, S Yoshioka, H Horikoshi, H Kuwano, H Hanzawa, and T Hata, J. Org. Chem. 58, 7632 (1993). KL Rinehart, Jr., J Kobayashi, GC Harbour, RG Hughes, Jr., SA Mizsak, and TA Scahill, J. Am. Chem. Soc. 106, 1524 (1984). BJ Baker, P-Carboline and isoquinoline alkaloids from marine organisms. In Alkaloids: Chemical & Biological Perspectives', Vol. 10, SW Pelletier, Ed., Pergamon, New York, 1996, pp 357-407.

75. 76. 77. 78. 79. 80. 81. 82. 83. 84. 85. 86. 87. 88. 89.

J Kobayashi, GC Harbour, J Gilmore, and KL Rinehart, Jr., J. Am. Chem. Soc. 106, 1526 (1984). KL Rinehart, Jr., J Kobayashi, GC Harbour, J Gilmore, M Mascal, TG Holt, LS Shield, and F Lafargue, J. Am. Chem. Soc. 109, 3378 (1987). P Molina, PM Fresneda, and M Canovas, Tetrahedron Lett. 33, 2891 (1992). A Rudi, I Goldberg, Z Stein, F Frolow,Y Benayahu, M Schleyer, and Y Kashman, J. Org. Chem. 59, 999 (1994). RJ Andersen, DJ Faulkner, C-H He, GD Van Duyne, and J Clardy, J. Am. Chem. Soc. 107, 5492 (1985). N Lindquist, W Fenical, GD Van Duyne, and J Clardy, /. Org. Chem. 53, 4570 (1988). M Aknin, J Miralles, J-M Komprobst, R Faure, E-M Gaydou, N Boury-Esnault, Y Kato, and J Clardy, Tetrahedron Lett. 31, 2979 (1990). CW Jefford, K Sienkiewicz, and SR Thornton, Tetrahedron Lett. 35, 6271 (1994). B Carte, and DJ Faulkner, /. Org. Chem. 48, 2314 (1983). N Lindquist, and WF Fenical, Experientia 47, 504 (1991). AJ Blackman, and C Li, Aust. J. Chem. 47, 1625 (1994). K Kojiri, S Nakajima, H Suzuki, A Okura and H Suda, J. Antibiotics 46, 1799 (1993). S Matsunaga, N Fusetani, and K Hashimoto, Experientia 42, 84 (1986). N Ezaki, T Shomura, M Koyama, T Niwa, M Kojima, S Inouye, T Ito, and T Niida, J. Antibiotics 34, 1363 (1981). M Kaneda, S Nakamura, N Ezaki, and Y litaka, J. Antibiotics 34,1366 (1981).

286

90. 91. 92. 93. 94. 95. 96. 97. 98. 99. 100. 101. 102. 103. 104. 105. 106. 107. 108. 109. 110. 111. 112.

P. W. LeQuesne, Y. Dong and T. A. Blythe

M Koyama, Y Kodama, T Tsuruoka, N Ezaki, T Niwa, and S Inouye, J. Antibiotics 34, 1569 (1981). N Ezaki, M Koyama, T Shomura, T Tsuruoka, and S Inouye, / Antibiotics 36, 1263 (1983). K Yano, J Oono, K Mogi, T Asaoka, and T Nakashima, 7. Antibiotics 40, 961 (1986). K Masuda, K Suzuki, A Ishida-Okawara, S Mizuno, and K Hotta, J. Antibiotics 44, 533 (1990). W Ding, DR Williams, P Northcote, MM Siegel, R Tsao, J Ashcroft, GO Morton, M Alluri, D Abbanat, WM Maiese, and GA Ellestad, J. Antibiotics 47, 1250 (1994). MP Singh, PJ Petersen, NV Jacobus, MJ Mroczenski-Wildey, WM Maiese, M Greenstein, and DA Steinberg, /. Antibiotics 47, 1258 (1994). JW Westley, C-M Liu, JF Blount, LH Sello, N Troupe, and PA Miller, J. Antibiotics 36, 1275 (1983). G Hofle, S Pohlan, G Uhlig, K Kabbe, and D Schumacher, Angew. Chem. Int. Ed. Engl 33, 1495 (1994). Y Yamagishi, K Shindo, and H Kawai, J. Antibiotics 46, 888 (1993). K Shindo, Y Yamagishi, and H Kawai, J. Antibiotics 46, 1638 (1993). H Suda, K Matsunaga, S Yamamura, and Y Shizuri, Tetrahedron Lett. 32,2791 (1991). Y Kawai, K Furihata, H Seto and N Otake, Tetrahedron Lett. 26, 3273 (1985). N Kawamura, R Sawa, Y Takahashi, K Issiki, T Sawa, N Kinoshita, H Naganawa, M Hamada, and T Takeuchi, J. Antibiotics 48, 435 (1995). KS Lam, GA Hesler, DR Gustavson, RL Berry, K Tomita, JL MacBeth, J Ross, D Miller, and S Forenza, J. Antibiotics 49, 860 (1996). T Hashimoto, A Yasuda, K Akazawa, S Takaoka, M Tori, and Y Asakawa, Tetrahedron Lett. 35, 2559 (1994). K Yamano, K Konno, and H Shirahama, Chem. Lett. 1541 (1991). O Sterner, Nat. Prod. Lett. 4, 9 (1994). K Likhitwitayawuid, N Ruangrungsi, GL Lange, and CP Decicco, Tetrahedron 43, 3689 (1987). D Cheng, Y Shao, R Hartman, E Roder, and K Zhao, Phytochem. 36, 945 (1994). G Veen, R Greinwald, L Witte, V Wray, and F-C Czygan, Phytochem. 30, 1891 (1991). M Timmermans, J-C Braeckman, D Daloze, JM Pasteels, J Merlin, and J-P Declercq, Tetrahedron Lett. 33, 1281 (1992). K Mahmood, M Pais, C Fontaine, HM Ali, A Hamid, A Hadi and E Guittet, Tetrahedron Lett. 34, 1795 (1993). N Aimi, N Uchida, N Ohya, H Hosokawa, H Takayama, S Sakai, LA Mendonza, L Polz, and J Stockigt, Tetrahedron Lett. 32, 4949 (1991).

Recent Research on Pyrrole Alkaloids

113. 114. 115. 116. 117. 118.

287

Y Hayakawa, K Kawakami, and H Seto, Tetrahedron Lett 33, 2701 (1992). S Nakatani, M Kirihara, K Yamada, and S Terashima, Tetrahedron Lett. 36, 8461 (1995). SH Kim, and PL Fuchs, Tetrahedron Lett. 37, 2545 (1996). SH Kim, I Figueroa, and PL Fuchs, Tetrahedron Lett. 38, 2601 (1997). Fiirstner, A.; Weintritt, H. J. Am Chem. Soc. 119, 2944-2945 (1997). T Luker, W-J Koot, H Hiemstra, and WN Speckamp, /. Org. Chem. 63, 220 (1998).

This Page Intentionally Left Blank

Chapter Four

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids Balawant S. Joshi and S. William Pelletier Institute for Natural Products Research and the Department of Chemistry, The University of Georgia Athens, Georgia 30602-2556, USA. CONTENTS 1. INTRODUCTION

292

2. ISOLATION TECHNIQUES

293

3. PHYTOCHEMICAL INVESTIGATIONS

293

4. NMR SPECTRAL DATA IN STRUCTURE DETERMINATION 5. NORDITERPENOID ALKALOIDS: ACONITINE-TYPE 5.1. 8-0-Methyl-14-0-veratroylpseudaconitine, 14-0-Veratroylbikhaconine and Balfourine 5.2. Columbidine 5.3. Falconerine, Falconerine-8-O-acetate, Falconericine and Falconeridine 5.4. Forestine, Foresticine, Acoforine, Acoforesticine, Acoforestine and Acoforestinine 5.5. Acoseptrigine, Acoseptriginine, 4-Anthranoyllapaconidine, Acoseptridinine and 14-0-Methylforesticine 5.6. Delstaphisine, Delstaphisagrine, l4-0-Acetylneoline (Delstaphisagnine), Delstaphidine, Neolinine, Delstasphisinine l-O-Acetyldelphisine, Delstaphinine, 1-Dehydrodelphisine, Delstaphigine, 14-0-Benzoyldelphonine, Staphisadrine and Staphisadrinine 5.7. Crassicaudine, Crassicausine and Crassicautine

293 294 294 295 296 297 299

300 302

6. NORDITERPENOID ALKALOIDS: LYCOCTONINE-TYPE 303 6.L 8-0-Methyllycaconitine, 6-O-Acetylacosepticine, Acosepticine, Acoseptridine, Acoseptrine, Acoseptrinine and 6-0-Demethyldelphatine 303 6.2. Ajadelphine, Ajadelphinine, 14-0-Deacetylajadine, Delajacine, Delajacirine, Delajadine, Ajanine, Ajadinine, 19-Oxoanthranoyllycoctonine and 19-Oxodelphatine 304 6.3. Barbeline, 6-0-Acetyldelpheline, Occidentaline (6-Deoxydelpheline), Occidentalidine, 14-0-Acetyldictyocarpine, Barbinine and Barbinidine 306 6.4. Delvestine, Delvestidine and Isodelectine 308 6.5. Delavaine A and Delavaine B 309 289

290

B. S. Joshi and S. W. Pelletier

6.6. 14-O-Deacetylambiguine, Tatsiensine, Tatsinine, Deltatsine, Delelatine and Tatsidine 309 6.7. Andersonine, 14-0-Deacetylnudicauline, Andersonidine and 14-0-Acetylnudicaulidine 311 6.8. Elasine, Isodelpheline, Eladine, Elanine, Blacknine and Blacknidine 312 7. METHYLENATION, DEMETHYLATION, DEOXYGENATION OF NORDITERPENOID ALKALOIDS AND HETEROLYTIC FRAGMENTATION OF DELTALINE 7.1. Methylenation of Lycoctonine-type Norditerpenoid Alkaloids 7.2. Demethylation of Lycoctonine-type Norditerpenoid Alkaloids 7.3. Demethylation of Aconitine-type Norditerpenoid Alkaloids 7.4. Deoxygenation Reactions of Norditerpenoid Alkaloids 7.5. Heterolytic Fragmentation of Deltaline

314 314 315 315 318 320

8. NORDITERPENOID ALKALOIDS: HETERATISINE-TYPE 8.1. 6-0-Acetylheteratisine 8.2. ^^C-NMR Spectral Assignments 8.3. Tangirine

373 373 373 373

9. DITERPENOID ALKALOIDS: HETISINE-TYPE 9.1. Heterophylloidine 9.2. 15-0-Deacetylvacognavine, Palmadine, Palmasine, Vakhmatine, and Vakhmadine 9.3. Septentriosine, 2-0-Acetylseptentriosine, and Septatsine 9.4. Tangutisine 9.5. 13-O-Acetylvakhmatine and Ajabicine 9.6. Chellespontine and Azitine 9.7. Andersobine 9.8. Delatisine 9.9. Tatsirine 9.10. Barbisine 9.11. Davisinol, 18-0-Benzoyldavisinol, and Davisine

326 326

10. REARRANGEMENT REACTIONS OF DITERPENOID AND NORDITERPENOID ALKALOIDS 10.1. Acid Catalyzed Rearrangement of Hetisine 10.2. Acid Catalyzed Rearrangement of 11-Dehydrohetisine and 2,11 -Didehydrohetisine 10.3. Base Catalyzed Rearrangement of ll-Acetyl-2,13-didehydrohetisine 10.4. Base Catalyzed Rearrangement of 13-Dehydro-2,ll-diacetylhetisine 10.5. Acid Catalyzed Rearrangement of Isoatisine 10.6. Acid Catalyzed Isomerization of Dihydroveatchine

327 330 332 334 336 337 339 341 343 344 347 347 348 350 352 353 355

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

10.7. Epimerization of the C(l)-Q;-Hydroxyl group of Delphisine and 8-O-Acetylneoline

291

356

11. BIOLOGICAL ACTIVITY OF NORDITERPENOID ALKALOIDS 359 11.1. Toxicity of Larkspur 359 11.2. Effect of Norditerpenoid Alkaloids on Cardiac Sympathetic Efferent and Vagal Afferent Nerve Activity 361 11.3. 6-Benzoylheteratisine 362 REFERENCES

362

B. S. Joshi and S. W. Pelletier

292

1.

INTRODUCTION Norditerpenoid alkaloids (also called Ci9-diterpenoid alkaloids) are highly oxygenated bases composed of one seven-, three six-, and two five- membered rings. They are mainly subdivided into four types: a) aconitine-type, having the skeleton (1), without substitution at C(7) by any group other than hydrogen. Some members of this class bear hydroxyl groups at C(3), C(13) and C(15); b) lycoctonine-type having the skeleton (1) with an oxygen substituent at C(7); c) heteratisine-type, a very small group of alkaloids with a 8-lactone moiety in ring C of skeleton 1, and d) pyroaconitine-type having a double bond between C(8) and C(15) in skelton 1. It is very likely that the pyroacontine-type do not occur naturally but are artifacts obtained during alkaloid isolation. The diterpenoid alkaloids (also called C20-diterpenoid alkaloids) have the basic skeleton (2), differing in the attachment of the C(15)-C(16) bridge at either C(l 1), C(12) or C(13); e.g. a) atisane-type alkaloids contain a [2.2.2] bicyclic ring system with the C(15)-C(16) bridge attached at C(12). Such a ring system incorporates an eAzr-ausane skeleton, but does not obey the isoprene rule; the atisane-type can be further divided in eight sub-skeletons: b) veatchine-type possessing a [3.2.1] bicyclic ring system with a C(15)-C(16) bridge connected to C(13), forming the five membered ring D. These compounds are modeled on an e/if-kaurane nucleus and obey the isoprene rule; c) delnudine-type where the five membered ring C in the skeleton differs from the veatchine type in that the C(15)-C(16) bridge is linked to C(ll). Biogenetic pathways for the formation of norditerpenoid and diterpenoid alkaloids have been postulated; however, supporting biosynthetic experimental evidence is lacking [1].

16

15

The diterpenoid and norditerpenoid alkaloids occur in the plant species of Compositae, Garryaceae, Ranunculaceae, Rosaceae, and Saxifragaceae. By far the largest number of diterpenoid alkaloids have been reported from Aconitum, Delphinium^ and Consolida genera of the Ranunculaceae family. Extracts of the Aconitum and Delphinium species have been used as arrow poisons since antiquity. In contrast, some of these plants have also been used for centuries in traditional Indian and Chinese medicine as cardiotonics, febrifuges, sedatives, and anti-rheumatics. Recent studies have shown that diterpenoid alkaloids are the active constituents responsible for the medicinal propenies of these plants [2].

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

293

Although the isolation of the first diterpenoid alkaloid was reported in 1819, the structures of these complex natural products were not determined for well over one-hundred years. The structure of lycoctonine was established in 1956, by X-ray crystallographic studies of a degradation product [3], the results of which marked a milestone in the development of the chemistry of diterpenoid alkaloids. In the 1970's ^H and ^^C NMR spectroscopic methods superseded chemical and degradation studies to establish the structures of these alkaloids. 2.

ISOLATION

TECHNIQUES

Since the isolation of the first alkaloid morphine by Sertiimer in 1805, a variety of methods have been used for the isolation of pure alkaloids from crude plant extracts. Many of the newer techniques include pH gradient extraction, preparative thin layer chromatography (PTLC), high performance thin layer chromatography (HPTLC), high performance liquid chromatography (HPLC), vacuum liquid chromatography (VLC), centrifugally accelerated radial thin layer chromatography ("Chromatotron"), counter current distribution (CC), droplet counter current chromatography (DCCC), and high performance centrifugal partition chromatography (HPCPC) [4]. In our work, we have made extensive use of VLC [5], "Chromatotron" [6,7] and occasionally HPLC [8], DCCC [9] and HPCPC [10] for purification of the diterpenoid alkaloids. 3.

PHYTOCHEMICAL

INVESTIGATIONS

This review embodies some of our work of the last fifteen years on the isolation, determination of structures, partial synthesis, rearrangement reactions and spectroscopic studies of diterpenoid alkaloids isolated from Aconitum, Delphinium, and Consolida species. The chemical structures given in the review are predominantly those of new alkaloids; however, structures of known alkaloids are also given where necessary. It is not uncommon to find the presence of 2030 alkaloids in a single plant species. Also, it is noteworthy that hundreds of discrete alkaloids with the norditerpenoid alkaloid skeleton (1) have been found in species of Aconitum and Delphinium species in the Ranunculaceae family. The question of how the wide variety of alkaloids in a single plant species is biosynthesized and what enzymes are responsible for each of the oxygenation and alkylation steps is unresolved. This can only be answered by intensive biosynthetic studies of these plant species.

4.

NMR SPECTRAL DATA IN STRUCTURE DETERMINATION

The only systematic study of the ^H-NMR spectra of norditerpenoid alkaloids was published by Tsuda and Marion who established the presence of acetate and benzoate groups in aconitine, delphinine, and related alkaloids [11]. The ^H NMR spectra are not as useful as the

294

B- S. Joshi and S. W. Pelletier

13c NMR spectra in the structure elucidation of these alkaloids. Jones and Benn made the initial contribution to l^C NMR spectroscopy of diterpenoid alkaloids [12,13]. This technique was further developed by Pelletier and co-workers [14,15] which greatly accelerated progress in structure elucidation of these complex natural products. Today the availability of NMR experimental techniques, such as homonuclear iH-decoupled l^C, DEPT, 2D iR COSY, hetero COSY (HETCOR), TOCSY, ROESY, lH-13c HMQC, HMQC-clean TOCSY, nOe, COLOC, HMBC and selective INEPT allow structure determination of small amounts of isolated alkaloids. The structures of more than 500 norditerpenoid and diterpenoid alkaloids have been determined by NMR studies. The structure determination of norditerpenoid alkaloids has now become straight forward, as most of these alkaloids fall in the four major structure types discussed earlier and have well-defined substituent and configurational patterns. In addition to these, a l^C NMR "data bank" is readily available for comparison of data of alkaloids which have similar structures [14,15]. However, the structure determination of diterpenoid alkaloids (C20) is more challenging because of their diverse substructure types and also the fewer number of alkaloids for which the l^C NMR data are available. The general procedure for l^C data acquisition and assignments of the resonances for each of the alkaloids, involves determination of the iR, and l^C DEPT (Distortionless Enhancement by Polarization Transfer) NMR spectra. The signals can be assigned by means of the single frequency proton off resonance decoupling technique, direct analysis of non-protonated carbon centers, application of known chemical shift rules for substituent shifts, steric effects and comparison of spectra of closely related compounds. Most of the accurate chemical shift assignments can be made by a study of HMQC (Homonuclear Multiple Quantum Coherence) and HMBC (Homonuclear Multiple Bond Coherence) NMR spectral data. The first detailed studies involving COSY, and C/H correlations were carried out on 3a-hydroxybikhaconine, obtained by hydrolysis of yunaconitine, (3a,13-dihydroxyforesaconitine) [16]. We have carried out detailed NMR spectral investigations of aconitine [17], tatsidine [18], delpheline [19], dictyzine [19], ajabicine [20], andersobine [9],tatsirine [21], and many other alkaloids. This study has enabled not only the structure derivation but also the accurate chemical shift assignments in these alkaloids. Our work indicated that accurate data collection of l^C NMR spectra of norditerpenoid and diterpenoid alkaloids is very useful in the structure determination of newly isolated compounds. Also, in the case of amorphous alkaloids, where an X-ray structure determination is not possible, l^C NMR spectroscopy can be a very useful tool for derivation of structures. 5.

NORDITERPENOID ALKALOIDS: ACONITINE-TYPE

5.1 8-0-Methyl-14-0-veratroylpseudaconine (4), 14-0-Veratroylbikhaconine (5) and Balfourine (6)

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

295

Aconitum balfourii Stapf. is found in the sub-alpine and alpine Himalayas from Garhwal to Nepal between 7,500 and 14,000 ft and its roots are reported to be highly poisonous [22]. Henry and Sharp [23] reported the isolation of pseudaconitine from A. balfourii and this alkaloid was first isolated in 1877 from the roots oiA.ferox Wall [24], a species which is used in Ayurvedic medicines [22]. After a series of investigations, the structure of pseudaconitine was established as 3 in 1963 by Tsuda and Marion [25]. lOMe

OMe

Et—

3 4

OMe OMe R^=Ac; R2=Vr(Veratroyl) R^ =Me; R2 = Vr

OMe 5

R =Vr

Three new norditerpenoid alkaloids, 8-(9-methyl-14-0-veratroylpseudaconine (4), 14-0veratroylbikhaconine (5), and balfourine (6) were isolated from the roots, together with eight known alkaloids: pseudaconitine (3), veratroylpseudaconine, indaconitine, ludaconitine, 8deacetylyunaconitine, bikhaconitine, neoline, and chasmanine [14]. Structures of the new alkaloids were determined by spectral data and chemical correlation with alkaloids of established structures [26]. Nine known norditerpenoid alkaloids were isolated from the aerial parts of A. balfourii: condelphine, bullatine C, neoline, isotalatizidine, l-O-methyldelphisine, pseudaconitine (3), yunaconitine, bikhaconitine and indaconitine [14]. Detailed NMR studies (^H, l^C, ^H-^H COSY, HETCOR, and selective INEPT) were carried out on condelphine, neoline, isotalatizidine, and indaconitine to provide accurate chemical shift assignments for these alkaloids [27]. 5.2

Columbidine (7) A. columbianum Nutt. ssp. columbianum is a perennial shrub growing in the western parts of the United States. The presence of aconine and aconitine in this plant has been recorded although no reference has been cited for their original isolation [28]. Some preliminary studies on the isolation of amorphous alkaloids from the roots and assays of the toxicity to sheep and cattle have been recorded [29, 30]. A chemical investigation of the Canadian variety reported as major alkaloids: talatizamine and cammaconine and as minor alkaloids: sachaconitine, talatizidine, isotalatizidine, 14-(9-acetyltalatizamine, and columbianine [31].

B. S. Joshi and S. W. Pelletier

296

In addition to the known alkaloids cammaconine, deltaline, dictyocarpine, talatizamine and 8-(9-methyltalatizamine, we isolated a new base, columbidine (7). The structure of 7 was derived on the basis of spectral studies and chemical correlation with talatizamine. The synthesis of 8-0-methyltalatizamine has been described [32]. Chemical shifts for the alkaloids talatizamine, 8-0-methyltalatizamine, 8,14-di-O-acetyltalatizamine, 14-acetyl-8-0-methyltalatizamine, 14-dehydrotalatizamine, cammaconine, and columbidine (7) have been assigned [32]. The 8acetoxyl group in norditerpenoid alkaloids can be replaced by an alkoxyl group by treatment with the corresponding alcohol under reflux or in a sealed tube at 110-130° C. 8-0-Acetyltalatizamine has not been isolated from a natural source and we failed to isolate it from A. columbianum. Talatizamine which has an OH group at C(8), did not fumish 7 when heated with ethanol at 50® C for a prolonged period.

.OMe 9Me 1 H > - O V r

QMe r ^ H / — O H

OEt OMe OMe

5.3 Falconerine (10), FaIconerine-8-0-acetate (11), Falconericine (12) and Falconeridine (13) A.falconeri Stapf. is found in the sub-alpine and alpine regions of the Himalayas of Garhwal and its roots are known to be poisonous [22]. Singh et al reported the isolation of the alkaloids bishatisine and bishaconitine without structure determination [33]. Pelletier et al isolated two pyroacontine-type norditerpenoid alkaloids, falaconitine (8) and mithaconitine (9)[34] from A. falconed in addition to the known compounds pseudaconitine (3), indaconitine, and veratroylpseudaconine [35]. The alkaloids 8 and 9 are most probably artifacts formed during the isolation procedure by the pyrolysis of pseudaconitine (3) and indaconitine. A reinvestigation of the alkaloids from the roots by isolation under mild conditions did not show the presence of these pyro-type alkaloids 8 and 9. The new alkaloids falconerine (10), falconerine-8-O-acetate (11) [36], falconericine (12) and falconeridine (13) were isolated from A.falconeri [37]. Another alkaloid, falconeridinine isolated from this plant, is probably an artifact resulting from the replacement of the 8-0-acetate in 11 with an ethoxy group. This was confirmed by refluxing 11 with ethanol to afford falconeridinine [37].

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

P"

OMe

OMe 8 R = Vr 9 R = Bz (Benzoyl) OMe

OMe

Et-

OMe 12 R = Ac 13 R = H

14 As (AnisoyI)

5.4 Forestine (14), Foresticine (15), Acoforine (16), Acoforesticine (17), Acoforestine (18), and Acoforestinine (19) The roots of A. forrestii Stapf. are used in the Chinese traditional medicine for the treatment of rheumatism. Chen and Breitmaier have reported the isolation of a foresaconitine (vilmorrianine C) (37; Ri = Ac, R^ = As) from A. forrestii Stapf. var. albo-villosum (Chen et Liu) W. T. Wang [38]. Our investigation of the roots resulted in the isolation of two novel norditerpenoid alkaloids, forestine (14) and foresticine (15), together with the known alkaloids, chasmanine, talatizamine, and yunaconitine [39]. In another study, we isolated four alkaloids, acoforine (16), acoforesticine (17), acoforestine (18), acoforestinine (19), in additon to crassicauline A. The structures of these alkaloids were determined on the basis of NMR spectral data and correlation with alkaloids of established structures. The structure and stereochemistry of acoforestine (18) was confirmed by an X-ray crystal structure analysis [40].

B. S. Joshi and S. W. Pelletier

298

OH

OMe

OMe

Et"

E t - -N

OMe 15 16

R ^ O H ; R2, R2 = H R^ = H; R2 = Et;R2 = Ac

17

R ^ O M e ; R 2 = H ; R 3 = AS

18 R=H

A few naturally occurring norditerpenoid alkaloids containing a methoxyl group at C(8) have been reported in the literature, e.g. alkaloid A (bicoloridine), ambiguine, puberaconitidine.

MeO

MeO

OAc-

H^ MeO

MeO

OAc-

R = Alkyl Scheme 1 sepentriosine, and hokbusine A [14,15]. Columbidine (7) is one of the alkaloids containing an ethoxyl group at C(8) [32]. The question as to whether the alkaloids containing a methoxyl or an ethoxyl group at C(8) are artifacts or are present in the plant cannot be answered unequivocally without careful investigation. The precursor of acoforine (16) should be 8,14-diacetyltalatizamine, but this compound has not been isolated from this plant. The facile conversion of the 8-

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

299

acetoxy group in some norditerpenoid alkaloids to the 8-alkoxy compounds can be considered as a synchronous fragmentation process such as described by Grob [41]. The free electron pair of the nitrogen atom is oriented anti and parallel (anti-periplanar) with respect to the C-C bond which undergoes cleavage as shown in Scheme 1. 5.5 Acoseptrigine (20), Acoseptriginine (21), 4-Anthranoyllapaconidine (22), Acoseptridinine (23), and 14-0-MethyIforesticine (24) The presence of alkaloids in A. septentrionale Koelle, a plant native to Sweden and Norway, has been recorded more than a century ago [42] and the roots are rich in alkaloids. Two novel aconitine-type norditerpenoid alkaloids, designated as acoseptrigine (20), and acoseptriginine (21), were isolated from the roots besides five known alkaloids: A/-acetylsepaconitine,N-deacetyllappaconitine, lapaconidine, lappaconine, and lappaconitine [14,15,43]. The structures of these alkaloids were determined by spectral data and chemical correlation with alkaloids of established structures. Alkaline hydrolysis of acoseptrigine (20) gave 14-(9-methylforesticine identified by comparison of the ^H and ^^C NMR spectra with those of an authentic sample (vide infra). The structure of acoseptrigine (20) could be 6-0-acetyl-14-0-methylforesticine or 8-(9-acetyl-14-0-methylforesticine. In the aconitine-type alkaloids, the ^^C chemical shifts for C(8) bearing an OH is from 72.5-74.5 ppm whereas the chemical shifts for C(8) bearing an acetoxyl group is from 85.5-86.0 ppm [14]. Thus the quaternary carbon of acoseptrigine at 74.1 ppm is assigned to C(8) (J-OH as in 20. From the same plant, by the ion OH OMe

OMe

>OMe

QMe

OMe 19

20

exchange method, three interesting norditerpenoid alkaloids of the aconitine-type, 4-anthranoyllapaconidine (22), acoseptridinine (23), and 14-0-methylforesticine (24), along with three known aconitine-type alkaloids, N-deacetyllappaconitine, sepaconitine, and lapaconidine were isolated. Their structures were derived on the basis of NMR spectral data [44]. Alkaline hydrolysis of alkaloid 22 gave lapaconidine [14]. In order to confirm that the strongly acidic ion exchange resin hydrolyses the amide group of lappaconitine, this alkaloid was processed by the ion exchange method under conditions used earlier [43] for the isolation of the crude base.

300

B. S. Joshi and S. W. Pelletier

Lappaconitine was deacetylated to afford A^-deacetyllappaconitine in quantitative yield, leaving the anthranoyl ester group intact. DMe QMe \

r

^ >---OMe

OMe

OMe

T^

OMe 24

5 . 6 . Delstaphisine (25), Delstaphisagrine (26), 14-0 - A c e t y l n e o l i n e (Delstaphisagnine), Delstaphidine (27), Neolinine (28), Delstaphisinine (29), 10-Acetyldelphisine (30), Delstaphinine (31), 1-Dehydrodelphisine (32), Delstaphigine (33), 14-0-Benzoyldelphonine (34), Staphisadrine (35) and Staphisadrinine (36) Delphinine, the major alkaloid of the seeds of D. staphisagria L. was isolated in 1819 [45]. The seeds of this plant have been a rich source of aconitine-type norditerpenoid alkaloids and eight interesting bisditerpenoid alkaloids [46]. Careful fractionation of mother liquors after separation of delphinine from an extract of the seeds led to the isolation of a number of new alkaloids. The new norditerpenoid alkaloids delstaphisine (25), delstaphisagrine (26) and 14-0acetylneoline (delstaphisagnine) were isolated by a combination of gradient pH extraction, VLC, and "Chromatotron" separation and the structures of the alkaloids were determined by 'H and ^^C NMR spectroscopy. Delstaphisine (25) is the first reported example of an aconitine-type alkaloid bearing a C(16) hydroxyl group [47]. In addition, delstaphidine (27) and neolinine (28) were also isolated from the seeds. Their structures were determined by spectroscopic studies

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

301

and established by correlation with alkaloids of known structures [48]. Delstaphisinine (29), and l-6>-acetyldelphisine (30), were also obtained and the structures were derived from spectral and ^OMe

Etv1-|

OMe 25 26

27

R U H ; R2=Me R^ = Me; R^ = H

correlation studies [49]. The structures of the alkaloids delstaphinine (31) and 1-dehydrodelphisine (32) isolated from the seeds were determined on the basis of ^H and ^^C NMR spectral data. Oxidation of neoline gave N-deethyldelstaphinine which when alkylated with ethylbromide afforded 31. Oxidation of delphisine with Cornforth reagent gave 32 [50]. Two alkaloids, delstaphigine (33) and 14-0-benzoyldelphonine (34), were isolated from the same

r

OMe

-N

OAc

H OMe 28

29 30

OMe

0R2

R\R2=H R^=AC;R2=

Me

plant. Methylation of 33 gave delphinine and deacetylation and benzoylation of delphinine afforded 34 [51]. Staphisadrine (35) and staphisadrinine (36) were also isolated from the seeds ofD. staphisagria and were assigned structures based on detailed NMR studies. The alkaloid 35 is a rare example of an alkaloid bearing a formyl group at C(4) and the carbonyl function at C(16) of alkaloid 36 is unusual. Two known alkaloids, pyrodelphinine and 14-0-acetyl-l-epJneoline, were also isolated [14,15]. An attempt to prepare 36 from 25 by selective oxidation of the C(16) hydroxyl group with pyridinium dichromate, afforded 1-dehydrodelstaphisine by oxidation of the C(l) hydroxyl group [52].

B. S. Joshi and S. W. Pelletier

302

»OMe

OMe

OMe

OMe 32

31

9^_

Me—hN

^OMe

^OMe

-K .H

0R2

^^^

33 R^=Ac; R 2 = H 34 R^ = H; R2 = Me

H CHOi OMe

OAc

35

5.7 Crassicaudine (37; Rl = Ac, R^ = Bz), Crassicausine (38; R = Me) and Crassicautine (39; R = Me) From the roots of A. crassicaule W. T. Wang, three alkaloids, crassicaudine (37; R^ = Ac, R2 = Bz), crassicausine (38; R = Me) and crassicautine (39; R = Me), were isolated and their structures determined on the basis of spectral data. The alkaloid crassicaudine was synthesized by stepwise esterification of chasmanine (37; Rl, R2 = H), and the structure of forestine (14) was confirmed by correlation with crassicauline A (38; R = Ac). Five known alkaloids, chasmanine (37; Rl, R^ = H), crassicauline A (38; R = Ac), foresaconitine (vilmorrianine C) (37; Rl = Ac, R^ = As), forestine (14) and yunaconitine (39; R = Ac), were also obtained from this plant [53]. In order to prove the structure (39; R = Me) assigned to crassicautine, the alkaloid yunaconitine (39; R = Ac) of well established structure [14,16], was heated with MeOH at 140-145° C to afford an amorphous compound identical with crassicautine.

Recent Developments in the Chemistr}' of Norditerpenoid and Diterpenoid Alkaloids

303

Me

^•^t L H / " O R ^ Et-H".N

El—

H OMe OMe

OMe

37

36 OH 1

Et—

•.N

OMe

OMe

38 6.

^OMe

JDMe

NORDITERPENOID ALKALOIDS:

OMe

OMe

39 LYCOCTONINE-TYPE

6.1 8-0-Methyllycaconitine (40), 6-0-Acetylacosepticine (41), Acosepticine (42), Acoseptridine (43; Rl, R2 = Me, R 3 = NH2), Acoseptrine (44), Acoseptrinine (45) and 6-0-Demethyldelphatine (46) Two lycoctonine-type norditeq^enoid alkaloids, 8-0-methyllycaconitine (40), 6-(9-acetylacosepticine (41), and the two known alkaloids lycoctonine and puberaconitine [14] were isolated from the roots of A. septentrionale [43]. The structures of these alkaloids were determined on the basis of their spectral data and chemical correlation studies. Thus, 8-0-methyllycaconitine (40) was transformed to septentrionine [14] by opening of the succinylamide ring with methanolic ammonia. 6-0-Acetylacosepticine (41) gave acosepticine (42) by treatment with 5% ethanolic KOH. By the ion exchange method for the isolation of alkaloids [4] from the roots of A. septentrionaley five norditerpenoid alkaloids: acosepticine (42), acoseptridine (43; Rl, R^ = Me, R3 = NH2), acoseptrine (44), acoseptrinine (45), and 6-0-demethyldelphatine (46), along with four known alkaloids, anthranoyllycoctonine, delvestidine, lycoctonine and septentrionine [14], were isolated. Their structures were derived on the basis of NMR spectral data [44]. A comparison of the l^C NMR spectrum of 42 with 6-(9-demethyldelphatine (46) indicated

304

B. S. Joshi and S. W. Pelletier

identity of chemical shifts for all carbons except the quaternary C(4) carbon in 46, vs. the tertiary carbon in 42. The '^C NMR chemical shifts of acoseptrine (44) indicated that this .OMe

_^OMe OMe

41 R^ = Ac; 42

40 45

R \ R2

= Me; R^ =-N.^

44

R2

=H

R\R2=H R^ = H ; R 2 = 0 H

1 ^ ^CO-

R \ R 2 = H ; R 3 = NH2

•OMe

^OMe

OMe OCO\^jsi*^

OMe 43

,

3

46

^

alkaloid bears an OH group at C(10) [14]. The structure 46 for 6-(9-demethyldelphatine was assigned by comparison of the ^^C NMR cheniical shifts with delphatine [14]. The structure of acoseptrinine (45) was derived by comparison of its ^^C NMR spectrum with delectine and isodelectine [54]. Acoseptridine (43; R^ R^ = Me, R^ = NH2), and anthranoyllycoctonine [14] had similar l^C NMR spectra, except for the A^-C(19) azomethine portion. 6.2 Ajadelphine (47), Ajadelphinine (48), 14-0-Deacetylajadine (49; R^ = H, R 2 = NHAc), Delajacine (50), Delajacirine (51), Delajadine (52), Ajanine (53), Ajadinine (54), 19-Oxoanthranoyllycoctonine [55; C(19) C=0] and 19Oxodelphatine [46; C(6) OMe, C(19) C=0] The aerial parts of C. ambigua (L.) P. W. Ball and V. H. Heywood (Syn. Delphinium ajacis L.) are rich in alkaloids, and most of these belong to the norditerpenoid-type. Twenty-nine alkaloids of the lycoctonine-type have been isolated from the seeds and various parts of this

305

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

plant, including: 14-0-acetylbrowniine, 14-(9-acetyldelcosine, 14-0-acetyldelectine, ajacine, ajacusine, ajadine, ambiguine, anthranoyllycoctonine, browniine, 14-0-deacetylambiguine, delcosine, delsoline, deltaline, deltatsine, gigactonine, lycoctonine, 18-methoxygadesine, methyllycaconittne, 19-oxoanthranoyllycoctonine, 19-oxodelphatine, and takaosamine [14,15]. Two new norditerpenoid alkaloids ajadelphine (47) and ajadelphinine (48) were isolated from the roots of C. ambigua and their structures established on the basis of NMR spectral data including DEPT, COSY, and HMQC experiments [55]. Ajadelphine (47) and ajadelphinine (48) are examples of lycoctonine-type alkaloids having a methoxyl at C(8), or a methylenedioxy at C(7)-C(8) positions and a hydroxymethyl group at C(4). 14-0-Deacetylajadine (49; R^ = H, R2 = NHAc), a lycoctonine-type alkaloid was isolated from the seeds of C. ambigua and its _^OMe

S—-OH Et-

Et-

-f-N'' 1 P\H OH

\ >

\

oJ

0

48

structure deducedfromspectroscopic data and chemical correlation by acetylation of (49; R^ = H, R2 = NHAc), to give ajadine [14,55], Four norditerpenoid alkaloids, delajacine (50), delajacirine (51), delajadine (52) and ajanine (53) were isolated from the leaves of C. ambigua ip. ajacis). The structures of these new alkaloids were determined on the basis of the mass, ^H and ^^C NMR spectral data. The structure of delajacirine (51) was confirmed by acylation of anthranoyllycoctonine (55) with isobutyryl chloride [56]. Ajadinine (54) has been isolated from the seeds of C. ambigua (D. ajacis) and its structure established from spectroscopic data. Detailed ^H and ^^C NMR spectral data were collected for the alkaloids ajacine, ambiguine, 14(9-deacetylajadine, and delcosine. Assignments for the aromatic protons of the C(18) Nacetylanthranilic acid ester group were made by selective INEPT NMR experiments [57]. Thirteen norditerpenoid alkaloids were isolated and identified from the stems and leaves of C, ambigua (D. ajacis), cultivated in Assiut, Egypt. The alkaloids 19-oxoanthranoyllycoctonine (55; C(19) C=0), and 19-oxodelphatine (46; C(6) OMe, C(19) C=0) were assigned structures on the basis of ^H and ^^C NMR spectral data and synthesis. Oxidation of ajacine (45; R^ = Me, R^ = H, R^ = NHAc) with osmium tetroxide gave 19-oxoajacine which when hydrolyzed afforded 19-oxoanthranoyllycoctonine and oxidation of 55 with osmium tetroxide gave the same compound. 19-Oxodelphatine could be obtained by potassium permanganate oxidation of delphatine [58].

B. S. Joshi and S. W. PeUetier

306

»OMe

OMe

Et--

Et" )Me OCO—1*«*^

«XJ

49

52 R^ = Ac; R^ = NHCOCH{Me)-Et

R^=H;R2=NHAC

50 RUMe;R2='NHC0CH(Me)-Et

53 RUCOC{Me)(OH)-Et; R^ = NHAc

51 R^ = Me; R2 = NHC0CH(Me)2

r

OMe

»OMe

9Me r^V-OAc I

9^! Et19*

54 6.3 Barbeline (56), 6-0-AcetyideIpheIine (57, R = OAc), Occidentaline (6Deoxydelpheline) (57, R = H), Occidentalidine (58, R = C0CH(Me)2), 14-0Acetyldictyocarpine (59, R = Ac), Barbinine (60), and Barbinidine (61) The occurrence of anthranoyllycoctonine, deltaline, dictyocarpine, and lycoctonine in D. barbeyi Huth. was reported in earlier literature [59]. From the ethanolic extract of D. barbeyi Huth, a novel norditerpenoid alkaloid barbeline {S6) was isolated and its structure determined by spectroscopic data and an X-ray crystal diffraction study. Barbeline was the first example of an alkaloid containing a C(19) N-azomethine group among the naturally occurring norditerpenoid alkaloids. Two new alkaloids, 6-0-acetyldelpheline (57, R = OAc) and occidentaline (6deoxydelpheline) (57; R = H), and the known alkaloids browniine, delphatine, and glaucenine were also isolated from this plant [60]. Three lycoctonine-type alkaloids, occidentaline (57; R = H), occidentalidine (58; R = C0CH(Me)2), and 14-0-acetyldictyocarpine (59, R = Ac), wvre isolated from D. occidentale S. Wats, and their structures deduced by spectroscopic methods jnd

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

307

chemical correlations. To confirm the structure assigned to occidentaline, delpheline (57; R = OH) was converted to the S-methyl dithiocarbonate (57; R = OC(=S)-SMe) and reduced with OMe

^OMe

9Me

LoK^ *\

N

\

OMe

—OMe

i

\\«H

\\

Et-

i

\ 1

.H Me OAc

0

^oJ

56 tri-rt-butyltin hydride to give compound 57, (R = H) [61]. The structure of occidentalidine was confirmed by acyladon of browniine with isobutyryl chloride to afford 58, (R = COCH(Me)2). Twelve known norditerpenoid alkaloids were isolated by gradient pH extraction, VLC and "Chromatotron" separations. These were: browniine, 14-dehydrobrowniine, 14dehydrodelcosine, delcosine, delpheline, deltaline (59; R = Me), deltamine, dictyocarpine (59; R = H), dictyocarpinine, glaucedine, glaucenine, and glaucerine [14, 61]. Chemical investigation of the aerial parts of/), barbeyi resulted in the isolation of 14-(9-acetyldictyocarpine (59; R = Ac), barbinine (60) and barbinidine (61). Their structures and stereochemistries were determined by spectral methods. Thirteen known alkaloids isolated were: barbeline (56), browniine ^OMe

OMe OMe 1

Et-

OMe 58

59

(58; R = H), 14-deacetylnudicauline, 14-dehydrobrowniine, 6-dehydrodeltamine, delcosine, delpheline (57; R = OH), delelatine (72), deltaline (59; R = Me), dictyocarpine (59; R = H), glaucenine, glaucerine and methyllycaconitine (67) [14,15]. The structure of 14-0-acetyldictyocarpine (59; R = Ac) was established by acetylation of dictyocarpine and that of barbinidine on the basis of NMR spectral data. The structure of barbinidine was confirmed as follows: alkaline hydrolysis of dictyocarpine (59; R = H) gave dictyocarpinine and oxidation of this with

B. S. Joshi and S. W. Pelletier

308

pyridinium chlorochromate furnished 6,14-didehydrodictyocarpinine, 14-dehydrodictycarpinine and 6-dehydrodictyocarpinine. Acetylation of the last compound gave barbinidine (61) [62]. 6.4

Delvestine (62), Delvestidine (63), and Isodelectine (65) D. vestitwn Wall, grows in the Western Himalayas and inner Tibetan valleys at elevations of 10,000-12,000 ft and is reported to be poisonous to goats [22]. An earlier investigation reported the isolation of two weak bases one of which when saponified gave lycoctonine [63].

9Me rrs=o r

9^^! foi^-oAc

OMe Me

L,

60

61

CO' /

Our investigation of the aerial parts of the D. vestitum gave two new norditerpenoid allcaloids, delvestine (62) and delvestidine (63) [64]. The structures were based on NMR spectral evidence and chemical conversion to known alkaloids. In order to confirm the structure, delvestine was hydrolyzed with methanolic potassium hydroxide to give an amino alcohol which on heating »OMe

OMe

Et-

HgN'

62 R U H , R 2 = Me

64 R = H

63 R^ = R2 = Me 65

RUR2 = H

with aqueous sulfuric acid afforded 64, (R = H) shown to be identical with gigactonine [65, 66]. The structure of gigactonine was established by its conversion to delsoline which was in

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

309

turn correlated with delcosine (X-ray structure) [67]. Demethylation of delvestidine with 3M sulfuric acid gave anthranoyllycoctonine (55) which on saponification gave lycoctonine. An Xray crystallographic study of delvestine confirmed the structure and stereochemistry of the alkaloid as 62 [68]. Isodelectine (65) was isolated from D. vestitum and its structure deduced from spectroscopic data and correlation with delvestine (62). The known alkaloid anthranoyllycoctonine (55) was isolated from this plant [54]. 6.5 Delavaine A (66, R = NHCOCH(Me)CH2C02Me) and Delavaine B (66, R = NHCOCH2CH(Me)C02Me) Investigation of the alkaloids of the Chinese plant D. delavayi Franch var. pogonanthum (H.-M) Wang led to the isolation of a pair of regioisomeric alkaloids delavaine A (66; R = NHCOCH(Me)CH2C02Me) and delavaine B {66\ R = NHCOCH2CH(Me)C02Me); their structures were determined from spectral evidence and two syntheses starting from methyllycaconitine (67). Delavaine A and delavaine B and the corresponding amides delsemine A (66; R = NHCOCH(Me)CH2CONH2) and delsemine B {66\ R = NHCOCH2CH(Me)CONH2) [14] were synthesized from methyllycaconitine (67) by refluxing with methanol and treatment with ammoOMe

OMe

66 nia. The regioisomers were separated on alumina rotors of a Chromatotron [69]. Six known lycoctonine-type alkaloids isolated from this plant are: anthranoyllycoctonine (55), delsemine, deltaline, deltamine, lycoctonine, and methyllycaconitine (67) [14]. 6.6 14-O.Deacetylambiguine (68; R = H), Tatsiensine (69), Tatsinine (70), Deltatsine (71, R l , R 3 = Me, R2 = H), Delelatine (72), and Tatsidine (73, Rl = Ac, R2 = H) No work was reported earlier on the chemical constituents of the Chinese plant D. tatsienense Franch. Two novel alkaloids, 14-(9-deacetylambiguine (68; R = H) and tatsiensine (69) were isolated from this plant. The alkaloid 68, (R = H) was prepared earlier by the hydrolysis of ambiguine (68; R = Ac) which was isolated from Consolida amhigua [70]. The structure

310

B. S. Joshi and S. W. Pelletier

of tatsiensine was established on the basis of spectroscopic data and correlation with delpheline (57; R = OH) [71]. Tatsiensine is a rare example of a lycoctonine-type diterpenoid alkaloid having unsaturation between C(2) and C(3). Since lycoctonine-type alkaloids do not possess an oxygen function at C(2) or C(3) positions, the formation of tatsiensine or takaosamine [14] appears to be biogenetically anomolous. The known lycoctonine-type alkaloids browniine. OMe

^OMe

OMe >^' • \ /

E t " h.Ni

1

LH V—OMe

UH < *»-'' Ar''

%

1

% i

V

Me OAc OMe

68

0

\)J

69

delcosine, and lycoctonine were also isolated [14,71]. A highly polar norditerpenoid alkaloid designated as tatsinine was isolated from the roots and its structure (70) was derived from ^H and ^^C NMR spectroscopic evidence [72]. As this structure depended purely on spectral evidence, and because of the unusual substitution pattern of the hydroxyl groups, an X-ray crystal diffraction study was undertaken. The structure and relative stereochemistry of tatsinine was confirmed by a single crystal X-ray analysis of tatsinine perchlorate [73]. Another alkaloid deltatsine was isolatedfromthe roots of D. tatsienense and its structure (71; R^ R^ = Me, R^ = H) was established on the basis of ^H, l^C NMR studies, and a chemical correlation with delcosine. Deltatsine was heated with 3M sulfuric acid to afford the crystalline alkaloid delcosine [74], the structure of which was established by an X-ray structure determination [67]. This reaction which probably proceeds by the formation of a tertiary carbonium ion is useful for dealkylation at the C(8) position [75]. The norditerpenoid alkaloid delelatine (72) was also isolated from D. tatsienense Franch andD. elatum L. and its structure was elucidated by spectroanalytical methods. Delelatine was synthetically correlated with 14-0-acetyl-lO-deoxydictyocarpine. Dictyocarpine was converted to 14-0-acetyldictyocarpine (59; R = Ac), which when treated with thionylchloride afforded the corresponding 10-chloro-lO-deoxy derivative. Reduction of this compound with tributyltin hydride gave 14-0-acetyl-lO-deoxydictyocarpine, identical with 6,14-diacetyldelelatine, thus confirming its structure and stereochemistry [76]. By chromatographic separation on silica gel rotors, another norditerpenoid alkaloid, tatsidine (73), was isolated from the pH 8 alkaloidal fractions [77]. Homonuclear ^H COSY, long-range COSY, and relay coherence transfer correlation NMR spectroscopy experiments were employed together with fixed evolution HETCOR [78] and selective INEPT spectra for complete ^H and ^^C spectral peak assignments

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

311

for tatsidine (73). Sterecx:hemical and conformational assignments were made from the two dimensional nOe spectrum. The results of these studies confirmed the location of the methylenedioxy group at C(7)-C(8) [18]. ^OMe

70 OMe

kOMe

Et" -

73

6.7 Andersonine (74; Ri = Ac, R2 = NHCOCH2CH(Me)C02Me + NHCOCH(Me)CH2C02Me), 14-O-DeacetyInudicauline (74; Rl = H, R2 = methylsuccinimido), Andersonidine (74; Rl = Ac, R^ = NH2), and 14-0-Acetylnudicaulidine (75) Two lycoctonine-type compounds andersonine (74; Rl = Ac, R^ = NHCOCH2CH(Me)C02Me + NHCOCH(Me)CH2C02Me) and 14-0-deacetylnudicauline (74; Rl = H, R2 = methylsuccinimido) were obtained from the pH 8 alkaloidal fraction of D. andersonii Gray by extensive purification by VLC and on a "Chromatotron". Their structures were deduced by spectral methods. Andersonine might be an artifact obtained by opening of the methylsuccinamide moiety of nudicauline (74; Rl = Ac, R^ = methylsuccinimido-) [15] during the isolation procedure. The structure of 14-(9-deacetylnudicauline was confirmed by acetylation to its monoacetate, which is identical with nudicauline. Six known alkaloids, delavaine, delectinine, lycoctonine, methyllycaconitine (67), nudicauline (74; Rl=Ac, R^ = methylsuccinamido), and takaosamine were also isolated [14,15,79]. In another investigation, the aerial parts afforded andersonidine (74; Rl = Ac, R^ = NH2) and 14-0-acetylnudicaulidine

312

B. S. Joshi and S. W. Pelletier

(75). In addition, VLC and "Chromatotron" separation gave the following lycoctonine-type alkaloids: 14-0-acetylbrowniine, 14-(9-acetyldelcosine, browniine (58; R = H), 14-0deacetylnudicauline (74, R^ = H, R^ = methylsuccinimido), delcosine, deltaline (59; R = Me), dictyocarpine (59; R = H), methyllycaconitine (67), and nudicauline [14,15,80]. OMe

Et"

DMe

Et"

OMe 75

6.8 Elasine (76), Isodelpheline, (77), Eladine (78), Elanine (79; R = COCH(Me)Et), Blacknine (80), and Blacknidine (81) Delphinium elatum L. has proven to be a rich source of norditerpenoid and diterpenoid alkaloids. Investigations by earlier workers had shown the presence of delpheline, deltaline, deltamine, elatine (82), and methyllycaconitine (67) [14] in the whole plant. The seeds of D. elatum cult. var. Pacific Giants Mix, afforded delelatine (72) [76]. Another study of the seeds of D. elatum cult. var. Pacific Giants Mix led to the isolation of elasine (76), isodelpheline (77), and eladine (78), besides seven known lycoctonine-type alkaloids: 14-0-deacetylnudiculine, (74, Rl = H, R2 = methylsuccinimido), delpheline (57; R = OH), deltaline, (59; R = Me), elatine (82), lycoctonine, methyllycaconitine (67), and nudicauline (74; R^ = Ac, R^ = methylsuccinamido) [14, 15]. The structures of the newly isolated bases were determined on OMe

Et

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

313

the basis of NMR spectral data and correlation with alkaloids with established structures. Thus methylation of elasine (76), and eladine (78) using proton sponge [1,8-bis(dimethylamino)naphthalene] and trimethyltetrafluoroborate gave the 16-methylethers identical with deltaline (59; R = Me), and delpheline (57; R = OH), respectively [81]. Elanine [79; R = COCH(Me)Et] was isolated from the neutral fraction of the seed extract of D. elatum cult. var. Pacific Giants Mix. The structure determination of the alkaloid was based on NMR spectral data and hydrolysis of both the ester functions at C(14) and C(18) of elanine to afford the known alkaloid delectinine [14]. Pacinine, obtained by oxidation of the C(6) hydroxyl group of delpheline (57; R = OH), and delectinine were also isolated from the pH 10 fraction of the seed extract [82]. From the whole plant ofD. elatum var. "black nighf, two alkaloids designated as blacknine (80) and blacknidine (81) in addition to five known alkaloids: 14-0-deacetylnudicauline, delectinine, delelatine (72), delpheline (57; R = OH), and methyllycaconitine (67) were isolated [14,15,83]. Wada etal have isolated from the seeds of D. elatum cult. var. Pacific Giants, the alkaloids pacifidine (43; Rl = H, R2 = Me, R^ = NH2), pacifiline [49; Rl = Me, R2 = NH2, C(19) C=0], pacifinine (N-desethylpacifiline), pacidine (l4-0-methylblacknine), paciline (14-(9-methylisodelpheUne) [15] and pacinine (6-oxoeladine) [15,84,85].

r

,OMe

OH

OMe I

QMe

OMe 78

OMe

Et" Me OH 80

kOMe

B. S. Joshi and S. W. Pelletier

314

7. METHYLENATION, DEMETHYLATION, AND DEOXYGENATION OF NORDITERPENOID ALKALOIDS AND HETEROLYTIC FRAGMENTATION OF DELTALINE 7.1

Methylenation of Lycoctonine-type Norditerpenoid Alkaloids A number of naturally occurring norditerpenoid alkaloids, e.g. delcorine, delpheline (57; R = OH), deltaline (59; R = Me), dictyocarpine (59; R = H), ajadelphinine (48), and occidentaline (57; R = H) contain a methylenedioxy group at the C(7)-C(8) position [14,55,60]. The structures of some of these alkaloids were established by acidic hydrolysis of the methylenedioxy group and comparison of the resulting products with alkaloids of known structures. The disadvantage of this method is that some of the functional groups, such as esters could also be cleaved under the experimental conditions. We anticipate that synthesis of alkaloids having a methylene- dioxy group from suitable substrates will be a useful method for structure determination. Elatine (82) isolated from D. elatum L. was assigned the structure based on chemical correlation with lycoctonine (64; R = Me) [86]. Treatment of methyllycaconitine (67) with aqueous formaldehyde andp-toluenesulfonic acid in refluxing benzene (Dean-Stark apparatus) gave elatine (82) [87]. Delbruline (83; Ri = H, R2 = Me) and delbrusine (83; RK R^ = Me), isolated from D. brunonianum Royle [88], were synthesized from known substrates [87]. Thus,

r

,OMe

OMe

OMe I Et"

9"^! Et--|-

Me 82

83

methylenation of 14-0-acetylbrowniine (58; R = Ac) with diethoxymethane and p-toluenesulfonic acid gave compound 83, (R^ = Ac, R^ = Me) which after hydrolysis furnished delbruline. Reaction of delphatine (58; R = Me) under similar conditions gave the compound 83, (R^ R^ = Me). Since the synthetic material differed in its physical constants from the reported values for natural delbrusine, compound 83, (R^ R^ = Me) was prepared by two other routes from alkaloids of established structures. The methylenedioxy derivative prepared from delsoline (71; R^ = H, R2,R3 = Me) was methylated to give 83, (R^ R2 = Me). Synthetic delbruline (83; R^ = H, R2 = Me) was methylated to afford the same compound, indicating that three different syn-

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

315

theses gave an identical product (83; R^ R^ = Me). Delsoline, ajacine, 1,14-di-O-acetyldelcosine, and 14-dehydrobrowniine [14] were converted to the corresponding methylenedioxy derivatives [87]. 7.2

Demethylation of Lycoctonine-type Norditerpenoid Alkaloids Most of the aconitine- and lycoctonine-type norditerpenoid alkaloids possess methoxyl groups at C(l), C(6), C(8), C(14), C(16) or C(18) positions, as opposed to the diterpenoid allcaloids which lack these functional groups. Although in recent years, the structure determination of these alkaloids has depended largely on ^H and ^^C NMR spectral evidence, selective demethylation studies are also useful in facilitating the structure determination of newly isolated alkaloids. Demethylation of the lycoctonine-type alkaloids delsoline (71; R^ = H, R2,R3 = Me) with HBr-AcOH gave compound 71, (R^ = H, R^ = Me, R^ = Ac) by demethylation of only the C(16) methoxyl group [89]. Similarly, ajacine (66; R = NHAc), and deltaline (59; R = Me) afforded 84 and 85, respectively. In all the cases, the C(16) methoxyl is demethylated and acetylated. In the alkaloids studied, the C(6P)-methoxyl and the C(18)-methoxyl groups were not demethylated under the experimental conditions [89]. lOAc

OAc

Et- •

AcHN 7.3

Demethylation of Aconitine-type Norditerpenoid Alkaloids Treatment of delphisine (86; R^ R^ = Ac) with 48% aqueous HBr gave the compounds delphidine (86; Rl = Ac, R2 = H) and neoline (86; RK R2 = H). When delphisine (86; Rl, R2 = Ac) was treated with 30% HBr-AcOH in glacial acetic acid, the demethylated product was identified as 87, (R = Ac) on the basis of ^H and ^^c NMR data. Alkaline hydrolysis of 87, (R = Ac) gave 87, (R = H) by deacetylation of C(8), C(14), C(16) and C(18)-acetoxyl groups. A similar reaction of delphinine (88) with HBr-AcOH gave the compounds 89 and 90 the structures of which were based on NMR spectral data. In the few cases of aconitine-type alkaloids studied, the C(18) methoxyl group is preferentially demethylated and the next methoxyl group to be demethylated is the one at C(16). In these alkaloids, the conformation of the methoxyl group at C(6) is a. The methoxyls at C(l) and C(6) are not demethylated under these conditions [89].

B. S. Joshi and S. W. Pelletier

316

^^OMe

OMe

87

86

Demethylation of delphinine (88) with trimethylsilyl iodide (TMS-I) in CH2CI2 [90] selectively demethylated first the 18-methoxyl group to afford 33 and 16,18-di-Odesmethyldelphinine. The alkaloid 33 was isolated earlier from D. staphisagria L. [51] and was designated as delstaphigine. The demethylation of aconitine (91) with HBr-AcOH gave 92 and 93 [90]. This was an unexpected result as aconitine is closely related to delphinine, except for the hydroxy] groups at the C(3) and C(15) positions and an N-ethyl instead of an N-methyl group. The probable explanation for the difference is the C(3) hydroxy] group present in aconitine and not in delphinine which readily undergoes acetylation with HBr-AcOH. The C(3)^0R2

Me-- -

88

89 R \ R3 = Ac;R2 = Me 90

R^ = H ; R 2 R 3 = AC

acetate appears to be a part of the displacement process probably by the formation of a six membered acetal; we believe that the C(18)-methoxyl is initially demethylated and is then converted to the C(18)-0-acetate (as observed in other demethylations) and is a better leaving group. The oxygen atom of C-6-O-methylether forms the furan ring with retention of configuration at C-6. Demethylation of aconitine (91) with TMS-I gave a mixture of 18-des-(9-methyl and 16,18-di-O-desmethyl aconiunes (94) and (95), respectively, as also reported by earlier investigators [91].

317

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

Demethyiation of falconerine (10) [36] which possesses an OH group at C(3), with HBrAcOH takes a similar course as in aconitine to afford the tetrahydrofuians 96,97, and 98. The structure of the demethyiation product (96) was based on detailed ^H and ^^C NMR spectral data. The demethyiation of other aconitine-type alkaloids, lappaoooitine (99) cammaconine

OMe

•-OBz •H

Et"j|--K 1 H O ' ' / > < ^ 3 V - ^ ^OAc

H'

X)H OAc

oV ^^« 91 R\R2 = Me 94 R ^ M e . R2 = 95 R \ R 2 = H

92 R\R2 = Me 93 R ^ M e . R2

H

= AC

.0R2

I '

Et-fy

Et-f

r\\! ^ f > OH

Ac0' y

V

^^0 96 R^=Me. R2 = AC 97 R \ R 2 = AC 98 Ri = H. R2 = Ac

102 R = Ac

(100) and talatizamine (101) with HBr-AcOH has also been invesdgaied. On the basis of NMR spectral data, the demethyiation product of lappaconitine (99) has been assigned the structure 14,16-di-0-desmethyl-14,16-diacetyllappaconitine (102). The demethyiation product of cammaconine (100) and one of those from talatizamine (101) have been sbo^-n to be identical with structure 103. A second product isolated in the demethyiation of talatizamine (101) has been assigned structure 104. The structure assignment of 103 was consistent with the detailed NMR spectral studies. Lappaconitine (99) when treated with TMS-I gave the polar compound 14,16di-0-desmethyllappaconitine which was difficult to purify by chromatography and it was therefore acetylated to afford 102 [90].

B. S. Joshi and S. W. Pelletier

318

Deoxygenation Reactions of Norditerpenoid Alkaloids Deoxygenation is a useful synthetic method in the structure determination of natural products. As the norditerpenoid alkaloids are highly oxygenated at various positions of the molecule, methods for selective conversion of secondary and tertiary alcohols to the corresponding deoxy7.4

^ R 2

Et"

100 R\R^ = H;R2=Me 103 R \ R 2 , R3 = AC

101 104

R^ = H;

R2 = Me

R \ R 2 = AC

genated products would be helpful for structure correlations among these alkaloids. Carmack et al had converted deltaline (59; R = Me) to delpheline (57; R = OH) via the 10-chloro derivative (105; R = OAc) by reduction with lithium aluminum hydride [92]. The disadvantage of this method is the non-selectivity which resulted in hydrogenolysis of the C(6) acetate group. Reaction of deltamine (106) with carbon disulfide and methyl iodide in alkaline solution afforded the .S-methyldithiocarbonate (107). Reduction of 107 with tri-«-butyltin hydride gave 6-deoxydeltamine (108) in good yield. Delpheline (57; R = OH) when treated with sodium hydride and a catalytic amount of imidazolide followed by addition of carbon disulfide and methyl iodide gave 57, (R = OC(=S)-SMe), reduction of which with tri-«-butyltin hydride furnished 6deoxydelpheline 57, (R = H), identical as reported earlier with occidentaline, a constituent of D. occidentalis S. Wats. [60,93]. In another study, deoxygenation reactions were carried out on delphisine, 14-0acetyldelcosine, aconitine, yunaconitine and 14-0-acetyldictyocarpine [93]. Thus the lycoctonine-type alkaloid 14-(9-acetyldelcosine (109) when heated with N/y/'-thiocarbonyldiimidazole (TCDI) in 1,2-dichloroethane gave the imidazolide (110) which on reduction with tri-n-butyltin hydride gave 14-0-acetyl-l-deoxydelcosoine (111). Alkaline hydrolysis of 111 gave 1-deoxydelcosine (112) in excellent yield. Treatment of the aconiune-type alkaloid delphisine (113) with thionyl chloride gave 1,2-dehydrodelphisine (114) as the major product and a minor chloro derivative (115). Catalytic hydrogenation of 114 in presence of Pd/C afforded 1-deoxydelphisine (116). As an alternative method to obtain 1-deoxydelphisine, 113 was treated with phenyl chlorothionocarbonate in the presence of 4-dimethylaminopyridine to afford the thionocarbonate (117) in good yield. Reduction of 117 with tri-«-butyltin hydride gave 113, identical with the compound prepared by catalytic reduction of 114.

Recent Developments in the Chemistry of Norditerpenbid and Diterpenoid Alkaloids

kOMe

Et—

E f - -N

r

319

Me

MeR 105

106 R = OH 107 R«OC(=S)-SMe 108 R = H

Treatment of aconitine (118) with TCDI gave 3-0-(imida2olylthiocarbonyl) aconitine (119) in 96% yield. Even though 118 has secondary hydroxyl groups at C(3) and C(15), only the C(3) hydroxyl is esterified. Reduction of 119 with tri-n-butyMn hydride afforded 3-desoxyaconitine (120).

r

•N H OMe

OMe

OH OMe

OMe 109

110

RU0H;R^AC

=N I ;R2=AC

RUOC-N

S

^ = '

111

RUH;R2=AC

112

R\

113 115 116 117

R = 0H R = CI R = H

R = 0C0-Ph It S

R2=H

A similar sequence of reactions on yunaconitine (39; R = Ac) gave 3-0-(imidazolylthiocarbonyl)- yunaconitine and the 3-desoxy derivative identical with crassicauline A (38; R=Ac) [93]. Marion et al had convened aconitine (118) into hypaconitine (121) in 7.7% yield [94]. N-Desethylaconitine (122) obtained by alkaline potassium pennanganate oxidation of aconitine (118) was methylated with methyl iodide to afford mesaconitine (123) in excellent

320

B. S. Joshi and S. W. Pelletier

yield. Treatment of 123 with TCDI gave 124 which on reduction with tri-n-butyltin hydride afforded hypaconitine (121) in an overall yield of 49% [93]. QH

OMe

)Me

Et-

OMe

OMe 114

118

R U O H ; R2=Et

119 R^=OC-N

I ;R^=Et

120 R^ = H;R2=Et 121 R^ = H;R2=Me 122

R^=0H;R2=H

123

RU0H;R2=Me

124 R^=OC-N

7.5

I ;R2=Me

Heterolytic Fragmentation of Deltaline Our plan to prepare 10-chlorodelpheline (105; R = OH) by the deacetylation of 10chloro-10-deoxydeltaline (105; R = OAc) was designed to obtain suitable derivatives of norditerpenoid alkaloids to evaluate their hypotensive activity. Compound 105, (R = OAc) was prepared by treatment of deltaline (59; R = Me) with thionyl chloride in dry benzene, and chromatographic purification according to literature procedure [92,93]. The treatment of deltaline with thionyl chloride in undistilled benzene (containing traces of moisture) however afforded a crystalline product C27H41NO8 in ~ 30% yield. The ^H NMR spectrum showed signals for three methoxyls, a tertiary methyl, an A^-ethyl, an acetate, a methylenedioxy and an aldehyde group. The presence of a tetra-substituted double bond was evident from the ^^C NMR spectrum. The ^^C NMR spectrum indicated the presence of six methyls, seven methylenes, eight methines and six quaternary carbon signals. On the basis of these data, structure 125 was assigned. Treatment of 105, (R = OAc) with aqueous MeOH at 45°C also gave 125 in 88% yield. Reduction of pure 105, (R = OAc) with «-tributyltin hydride afforded 10-deoxydeltaline (6-0-acetyldelpheline) (57; R = OAc) in excellent yield. In contrast, reduction

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

321

of crude 105, (R = OAc) with LiAlH4 is reported to give the elimination product 126 in 60% yield along with 16% of 10-chlorodelpheline (105; R = OH) [93]. Treatment of 105, (R = OAc) or 125 with methanolic KOH afforded a crystalline compound, mp 222-4^0 [94]. Its molecular formula C25H37NO6 was confirmed by elemental analysis and HRMS. The ^H and the l^c NMR spectral data of this compound differed from that of 125 through the absence of OMe

Deltaline: R^ = Ac; R2 = 0 H Delpheline : R^ = R^ = H 10-Chlorodelpheline: R^ = H; R^ = CI 10-Chloro-10-deoxydeltaline: R^ = Ac; R^ = CI 10-deoxydeltaline: R^ = Ac; R^ = H

Figure 1: ORTEP drawing of the rearranged product (127)

OMe

322

B. S. Joshi and S. W. Pelletier

signals for an acetate group. On the basis of detailed NMR spectral data (COSY, COLOC, SINEPT, HMBC), the rearrangement product was assigned the structure 127 which was confirmed by an X-ray crystal structure determination (Figure 1). OMe

PMe

126 A plausible mechanism for the formation of 125 and 127 is shown in Scheme 2. The formation of 128 from 10-chloro-lO-deoxydeltaline (105; R = OAc) is reminiscent of the synchronous fragmentation of 3-bromoadamentylamine investigated by Grob and Schwarz [95]. In this synchronous mechanism, the Ca-Cl bond and the orbital of the lone pair of electrons on the nitrogen are anti and parallel (anti-periplanar) [96] to the Cp-Cy bond and the staggered conformation and stereoelectronic condition present in (105, R = OAc) facilitates a fragmentation to OMer

OMe

MeO Me Deltaline socig \ r / 59 R = Me •* OAc 10-Chloro-10-deoxydeltaline ^OMe + KOH - AcOH

,^^

125——rr?-127

+ HOH

Scheme 2

Recent Developments in the Chemistry of Norditerpetloid and Diterpenoid Alkaloids

323

give 128. The azomethine (128) adds a water molecule to form 129 which undergoes further rearrangement to afford 125 [97]. The tertiary aldehyde at C(7) in the rearrangement products 125 and 127 arises from the methine carbon C(17) of the precursor. The ethylamine derivative 125 undergoes base catalyzed elimination of the C(6p)-acetate as the leaving group to give the pyrrolidine (127) [98]. 8.

NORDITERPENOID ALKALOIDS: HETERATISINE-TYPE

8.1

6-0-Acetylheteratisine (130) Very few heteratisine-type of norditerpenoid alkaloids have been isolated. These alkaloids contain a 5-lactone grouping in ring C in place of the C(14) carbon atom in 1. Heteratisine, the major alkaloid of this class occurs as a weak base in A. heterophyllum WalL[99] along with 6-benzoylheteratisine, and minor alkaloids of this type, heterophyllidine, heterophylline, and heterophyllisine [14]. The structure of heteratisine was established by an Xray crystal structure determination [100] and chemical degradation studies [101]. Heteratisine (130; R = H), and 6-0-acetylheteratisine (130; R = Ac), were isolated from the roots oi A, palmatwn Don. [102]. The structure of 6-0-acetylheteratisine was established by the ^H and ^^C NMR spectral evidence and acetylation of heteratisine to give (130; R = Ac). 8.2

13C-NMR Spectral Assignments 13c NMR chemical shift assignments made earlier for the heteratisine-type alkaloids heteratisine, 6-0-acetylheteratisine (130; R = Ac), 6-(9-benzoylheteratisine (134), have been revised from a study of ID, 2D and selective INEPT NMR spectral data. For the related alkaloids heterophyllidine (131), heterophylline (132) and heterophyllisine (133), definitive l^C NMR shift assignments have been made [103]. 8.3

Tangirine (135) A dimeric diterpenoid alkaloid, designated as tangirine, was isolated from Aconitum tanguticum (Maxim.) Staff, W.T.Wang, a plant native to China [104]. The molecular formula of tangirine (135) was established to be (^9H62N207 by HRFAB-MS. All the 62 protons and 49 carbons were visible in the ^H and ^^C NMR spectra of 135 in CDCI3 and all of the one-bond ^H-l^C connectivities were established by a HETCOR experiment. The DEPT spectrum indicated that tangirine possessed 11 quaternary carbons, 19 methines, 15 methylenes and 4 methyls. Of these, the six carbon singlets at 5 173.3, 166.6, 130.8, 78.7, 48.8, 34.7, the eleven doublets at 5 132.3, 130.2 (2C), 128.2 (2C), 82.3, 75.1, 74.0, 62.9, 55.9, 48.2, 44.7, 43.0, the seven triplets at 5 57.4, 48.8, 36.3, 31.2, 29.7, 29.2, 26.7 and the three quartets at 5 55.0, 25.9 and 13.5 are consistent with 6-benzoylheteratisine, C29H37NO6 (134) [14]. However, a ~ 4-5 ppm upfield shift for C(7) and C(15) was observed when compared with the values for these carbons in 6-benzoylheteratisine. The ^H-^H COSY, the NOESY, and the

324

B. S. Joshi and S. W. Pelletier

HMBC spectral results are consistent with a partial structure for tangirine where a bulky substituent group (T) is attached at the C(8) position in 6-benzoylheteratisine (134). The mass spectral fragmentations also support the presence of a methoxyl group at C(l) [105] and a benzoate ester group in tangirine.

Et—+-J

Et-

131 R^ = H. 130 134 R = COPh

R2 = 0 H

132

R\R2=:H

133

R^ = Me,

R2 = H

This group (T), C20H26NO, in tangirine is attached at C(8) through an oxygen function forming an ether and appears as a fragment ion in the MS (m/z 296, T^; m/z 494 M"*'-T). The group T showed in the ^^c NMR spectrum, one sp^ singlet at 5 146.0, four sp^ singlets at 5 72.5, 45.0, 44.9, 43.1, two sp2 doublets at 5 169.2, 127.8, four sp3 doublets at 5 80.3, 46.4, 44.3, 31.5, eight methylenes at 5 60.4, 42.8, 31.5, 30.6, 30.5, 28.4, 27.4, 20.6 and one methyl at 8 19.0. The ^H NMR signal at 5 3.82 (2H, dd, AB, 7=12.0 Hz; 13C, 60.4 ppm), showing a connection (COSY, nOe) with 5 4.98 (IH, br s; ^^C, 127.8 ppm) suggested the presence of the grouping, -CH=C-CH2-0- in T. Also, the signal at 5 7.32 (IH, d, 7=2.5 Hz; 13c, 169.2 ppm) was coupled (COSY) to 5 3.30 (IH, br s ; l^c, 80.3 ppm) indicative of the system -C-CH=N-CH- in T. The three proton singlet at 8 1.00 (I3c, 19.0 ppm) suggested the presence of a tertiary methyl group. The molecular formula, C20H26NO, biogenetic considerations, and the functional groups described, suggested one of the three possible structures A, B or C for T. The quaternary carbon singlet at 8 72.5 can be assigned to C(14') in A or to C(7') in B or C. The other singlets at 8 45.0, 44.9 and 43.1 were assigned to C(4'), C(IO') and C(8"), respectively. Of the two closely appearing signals, 45.0 ppm was assigned to C(4') as this showed a two-bond correlation to H(19') (8 7.32; 8 l^c, 169.2) in the HMBC spectrum. Also, C(IO') showed a three-bond correlation with the proton at 8 1.32 (13c at 8 27.4, HMBC) which may be assigned to C(2') or C(6'). The methine proton at 8 1.18 (13C, 8 44.3) was assigned to C(5') as this correlated with H(20') (8 3.30; 13c, 8 80.3) in the COSY spectrum and showed a threebond correlation with the tertiary methyl CH3(18') (8 iH 1.00; 13C, 8 19.0, HMBC). H(5') exhibited an nOe to H(3') (8, iH 1.22; 13c, 8 30.6). The most downfield quaternary carbon at 8 146.0 is clearly assigned to C(16'). The 2D COSY and the long-range COSY (LRCOSY) spectra showed a correlation of H(15') (8 4.98) with H(12') (8 2.00) and the methylene at C(17) (8

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

325

3.82, l^C, 5 60.4). An important clue to the attachment of the two halves of this dimeric alkaloid is an nOe observed between the H(7) proton of 6-benzoylheteratisine and the H(17') methylene of the remaining part of the molecule. In the HMBC spectrum, H(15') (5 4.98) showed a two-

1Q'

CH3

^^

18'

bond correlation with C(8'), and a three-bond correlation with C(17') which is in agreement with all the three structures A , B , orC. H(15') also showed a correlation with 31.5 ppm (overlapping signals for a CH2 and a CH). In the case of A, these carbons are assigned to C(7') and C(12') or C(9') or C(13'), whereas in B these have to be assigned to C(14'), and C(9') or C(12') and in C to C(14'), C(9') or C(13'). The crucial evidence in support of structure A was the observed correlations of H(13') in the HMBC spectrum. One of the methylene protons at 8 1.52 assigned to H(13'a) (i^C, 5 42.8) is related to C(8') (5 43.1, 3 bonds separated), C(12') (5 31.5, 2 bonds), C(16') (5 146.0, 3 bonds) and C(20') (6 80.3, 3 bonds). A correlation of any methylene protons, two or three bonds separated from C(16') (6 146.0) to C(20') (5 80.3) is possible only in the case of structure A. This evidence excludes the alternative structures B and

I

' "?

21

135

Tangirine

B. S. Joshi and S. W. Pelletier

326

C for T. After assigning five methine carbons as discussed, the remaining signal at 8 46.4 was assigned to H(9'). In the HMBC spectrum, the methylene protons at 5 1.42 O^C, 5 30.5) showed a correlation to C(5') (54.3) three-bonds separated and were assigned to H(r). The methylene carbons appearing at 27.4 and 20.6 ppm were assigned to C(2') and C(6'), respectively, by comparison with the values assigned to dihydroatisine [C(2), 23.2; C(6), 17.6 ppm] and dihydroajaconine [C(2,) 23.1; C(6), 17.6 ppm] [106]. The ^^c assignments, the proton COSY and NOESY results, and the HMBC data for T corroborate the structure 135 for tangirine. Tangirine is the only example of a dimeric alkaloid in which a heteratisine-type norditerpenoid and a diterpenoid alkaloid are joined together. Six other dimeric diterpenoid alkaloids of this class: e.g staphidine, staphisine etc., all isolated from D. staphisagria, are C20-diterpenoid alkaloids of the atisane-type, dimerized at the C(17), C(17'), C(16'), O, C(15'), C(16) positions to form spiro ethers [107]. 9.

DITERPENOID ALKALOIDS: HETISINE-TYPE

9.1

Heterophylloidine (137) An amorphous alkaloid heterophylloidine was isolated by Pelletier et al, [108] from the Indian plant A. heterophylloides Stapf. On the basis of NMR data and an X-ray analysis of the bromo derivative (136), the structure and absolute configuration (137) were assigned to heterophylloidine. In a later publication, Katz and Staehlin unaware of the work on heterophylloidine, reported the isolation of an alkaloid panicutine from the European species A. paniculatum Lam. They assigned the structure (138) for panicutine from its UV, Mass , ^H and ^^c NMR spectral analysis [109]. The structure of panicutine differed from heterophylloidine in location of one of the keto groups at C(l 1) instead of C(13) and left the stereochemistry at C(2) undecided. As the l^C NMR spectra of heterophylloidine and panicutine appeared to be identical, we undertook reinvestigation of the structure [110]. The alkaline hydrolysis products of heterophylloidine and panicutine were shown to be identical in their TLC, IR, mp and mmp. A choice between the two

19

igMe 6 137 141

R = Ac R =H

327

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

Structures was settled by application of CD measurements of 11-ketohetisine (139) and spiradine A acetate (140) [111], both having a keto group at C(ll), as expected show a positive Cotton effect. In contrast, deacetylheterophylloidine (141) exhibits a strong negative Cotton effect establishing the location of the keto groups at C(6) and C(13) in the hetisane skeleton. On addition of HCl, the CD of 141 showed a negative Cotton effect with slightly lower am-plitude due to loss of contribution by the 6-keto group. The structure assigned to panicutine was therefore revised to 137 [111].

N"'"T" L H

Me

H

i

R2

139

R^ = 0H , R2 = H

140

Ri = H,

R2 =

OAc

9.2 15-0-Deacetylvacognavine (143), Palmadine (144), Palmasine (145), Vakhmatine (151), and Vakhmadine (152) A. palmatum Don. occurs in the alpine Himalayas of Nepal, Sikkim at elevations of 10,000-16,000 ft and the roots are reported to be non-poisonous. The roots ("Vakhma") are used in the Ayurvedic medicine as a tonic, and in the treatment of diarrhea and rheumatism [22]. Singh and Singh [112] reported the isolation of five diterpenoid alkaloids: vakognavine, palmatisine, vakatisine, vakatisinine, and vakatidine without assigning structures. The structure of vakognavine (142) was established by an X-ray crystal structure determination of the hydriodide [113]. Later publications focused on the chemical and spectral confirmation of the structure of vacognavine (142) [114,115] and that of vakatisine [116]. Vacognavine is the first example of an N-C(19)-seco diterpenoid alkaloid and is of interest for biogenetic speculation. The C(19) aldehyde may be a plausible alternate to the pseudokobusine structure [117] and may be an intermediate in the biosynthesis of hetisane-type alkaloids [113]. A reinvestigation of the constituents A. palmatum in our laboratory led to the isolation of three diterpenoid alkaloids: 15-(9-deacetylvacognavine (143), palmadine (144), and palmasine (145) [102]. Other known alkaloids isolated from this plant were: vacognavine (142), isoatisine and hetidine [117]. The ^H NMR spectrum of 15-0-Deacetylvacognavine (143) showed the presence of an aldehydic proton at 5 9.24 as a singlet and an aldyhydic carbon as a doublet at 5 195.0 in the ^^c NMR spectrum. Acetylation of 143 furnished a compound identical with

B. S. Joshi and S. W. Pelletier

328

vacognavine (142). The position of the hydroxyl at C(15) was established by homonuclear decoupling experiments. Irradiation of the C(17) methylene protons at 5 5.20 and 5.23 (each IH, s), showed decoupling of the broad singlet at 5 4.08 attributed to the C(15) proton, and vice versa. The upfield signal at 5 4.08 attributed to H(15a) in 143 compared with the signal at 5 5.45 in 142 for the C(15a)-proton confirmed that the hydroxyl group in the new alkaloid is located at C(15). Palmadine (144) showed ^H NMR signals characteristic of hetisane-type diterpenoid alkaloids [118]. Selective homonuclear irradiation of the C(9) and C(14) protons showed decoupling of the C(liP) and C(13p)-protons, indicating that the two acyl groups in palmadine, an acetyl group at 5 2.02 and a cinnamoyl group at 8 6.61, are located at C(l 1) and C(13), although their respective precise positions are uncertain. In order to relate palmadine to hetisine (146), the latter alkaloid was acetylated to afford a mixture of 11-0-acetylhetisine (147), 13-(9acetylhetisine (148) and 11,13-di-O-acetylhetisine (149) [119]. Compound 148 has a ^^C NMR spectrum identical with that of naturally occurring 13-(9-acetylhetisine, whose structure has been established by an X-ray crystal structure determination [118]. Acylation of 11-0-

AcO. OAc

^^^> Jk'':^*' J OR Me-I-- rt'--f-.^^ T 1

Ji

y\»Me

\

1

OHC'

142 143

R = Ac R=H

144 R^ = OH; R2 = OAc; R^ = OCOCH=CH-Ph 145 R^ = R2 = OH; R3 = OCOCH=CH-Ph 146 R1 = R2 = R3 = OH 147 R^ = R3 = OH; R2 = OAc 148 R^ = R2 = OH; R3 = OAc 149 R^ = 0 H ; R 2 = R3 = 0 A C 156 fO= R2 = R3 = OAc

acetyl-hetisine (147) with cinnamoylchloride gave a compound identical with palmadine (144), confinning its stracture assignment. There has been some confusion and inconsistency in the literature in denoting the stereochemistry of the C(13) hydroxyl group in hetisine (indicated both a as a dotted line, and P as a thick line) [117,119,120,121,122,123,124,125,126]. For the sake of consistency, the convention adopted in our work is defined as follows: the boat conformation containing both C(l 1) and C(13) and formed by carbon atoms 8, 9, 11, 12, 13 and 14 in 150 is selected as the reference

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

329

ring [21,127,128]. The hydroxyls at C(ll) and C(13) in hetisine are both a. This representation of configuration of the \j/-axial C(13) hydroxy 1 as a is opposite to that shown in the series of earlier papers. Palmasine (145) is a minor alkaloid whose spectral data suggests structural features similar to those of palmadine (144). Palmasine is 42 mass units less than palmadine (144) and its proton NMR spectrum showed the presence of only a cinnamoyl group, compared to the presence of an acetyl and cinnamoyl group in (144). To locate the cinnamoyl group in palmasine, hetisine (146) was treated with cinnamoyl chloride in pyridine followed by sep-

, a• ^ Ha

^

•Hb

I

\ --I--N'

Hp ^ N

T "^ T

1 '

1

'

Me

150 Hetisine

J

153 Atisine

aration, to give monocinnamoyl hetisine, identical with palmasine. When this synthetic product was acetylated with acetic anhydride-pyridine, 11-acetyl-13-cinnamoylhetisine, identical with palmadine (144) was obtained, thus confirming the structure assignment 145 for palmasine.

H0-. MeHO''

n

H

Me OH

\c2'Me 'R'' 151

R\R2R3=OH

154

R \ R 2 , R 3 = OAC

155

R\ R^ = OAc; R2= OH

165

R\

R2

"K •

= OH; R3= OAc

152

B. S. Joshi and S. W. Pelletier

330

In another study, two diterpenoid alkaloids, vakhmatine (151) and vakhmadine (152), and the known alkaloids atisine (153) and hetisine (146) were isolated [128]. Comparison of the 13c NMR spectrum of vakhmatine (C20H27NO4) with that of hetisine (146) showed strong similarity in the 5c values for C(7) to C(17). The existence of a doublet at 5c 95.5 and the absence of a triplet normally at - 5c 65-70 characteristic of the C(19) signal of diterpenoid alkaloids, strongly suggested that one of the hydroxyl groups of 151 was situated at the C(19) position. Acetylation of vakhmatine (151) gave the tetra and tri O-acetyl derivatives 154 and 155, respectively. Examination of the 2D COSY NMR spectrum located three of the four acetoxyl groups at the C(2a), C(l la) and C(13a) positions. The site of the remaining acetoxyl group at C(19) was established by the observation of a downfield singlet at 5H 5.73 and a doublet at 6c 92.6. Comparison of the 5c values of 154 and tri-(9-acetylhetisine (156) showed an a effect of the 19-OAc on C(19), causing a downfield shift of 29.0 ppm, a P effect on C(4) of 6.6 ppm, a y effect on C(18) of 8.0 ppm, and also a steric effect shifting C(6) and C(20) upfield. The C(19) acetoxyl group was assigned a p-configuration on the basis of an nOe experiment. Based on detailed NMR studies, vakhmatine was assigned structure 151. Vakhmadine (152), C2iH3oN04"*' OH', was isolated as a quaternary base. Acetylation of 152 gave a triacetyl derivative (157) in which a masked 6-keto group was restored. This compound showed structural features of hetidine-like alkaloids [117] indicating an A/-methyl, an exocyclic methylene, hydroxyl groups at C(2), C(3) and a keto group at C(6). A detailed NMR study and comparison of the ^^C NMR spectra of 157 and 2,3-diacetylhetidine (158), led to the structure 152 for vakhmadine [128]. AcO^,

>.

Q

V

.OH.

^ ---1 "

AcO-., Me—

-N' T

^

AcO>.

T

Me-" -•N' T f

1

^

AcO'' Me 0 157 9.3 Septentriosine (159), 2-0-AcetyIseptentriosine (162)

158 (160), and Septatisine

The occurrence of alkaloids in A. septentrionale Koelle was recorded more than one hundred years ago [42] and the roots of this plant are found to be rich in alkaloids. In 1967, Marion et al reported the isolation of seven alkaloids of which two were the known compounds lappaconitine and deacetyllappaconitine. Structures of the remaining alkaloids were not determined [129]. More than twenty-four norditerpenoid alkaloids have now been isolated from

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

331

A. septentrionale by Marion et al [129], Yunusov and co-workers [130-132], and from our own laboratories [43,44,133,134]. Our investigation of the roots of this plant has led to the isolation and characterization of the diterpenoid alkaloids septentriosine (159) [135], 2acetylseptentriosine (160) [136], and septatisine (162) [137]. Septentriosine, C20H27NO4, formed a colorless crystalline hydrochloride salt. In the l^C NMR, one of the hydroxyls appearing as a doublet at 5c 95.3 ppm was readily associated with the C(19) methylene of the hetisane skeleton and the tertiary hydroxyl group at 5c 79.8 appearing as a singlet was assigned to C(6), C(9) or C(14). A number of possibilities remained for the placement of the remaining two secondary hydroxyl groups. Because of the uncertainties of positioning of the hydroxyl groups, the structure elucidation was completed by X-ray diffraction studies on the hydrochloride of septentriosine. The absolute configuration of 159 is defined as : C(l) 5, C(2) S, C(9) S and C(19) R. The hydroxyl groups at C(l) and C(2) are trans [C(lp), C(2a)], and appear diaxially disposed relative to the chair conformation of the C(l), C(5), C(10) of ring A. The C(9) hydroxyl is p (axial) and the a hydroxyl group at C(19) is du-ected endo to the A ring [135]. A crystalline alkaloid, C22H29NO5,2-acetylseptentriosine (2-a-acetoxyhetisaneip,9p,19a triol), was also isolated from the roots of A. septentrionale in 0.04% yield [136]. The ^H and l^C NMR spectral data indicated that this alkaloid is a monoacetyl derivative of the previously isolated alkaloid septentriosine (159). Of the three hydroxyl groups, one is tertiary (5c 79.6) and on the basis of ^^C NMR spectral analysis, it was located at C(9), after considering the positions C(5), C(6), C(12), C(14) and C(20). Of the remaining three secondary hydroxyl groups, an OH or acetoxyl group should be located at C(19); the ^^C spectral comparison with septentriosine indicated that the other hydroxyl or acetoxyl groups

rTr'

CHn

R^O

159

160

R^=H;R2=AC

161

R\

R2 = AC

should be placed at C(l) and C(2). Mild alkaline hydrolysis of the new alkaloid gave septentriosine (159) and acetylation with AC2O/ pyridine gave 1,2,19-triacetylseptentriosine (161). On the basis of NMR spectral data and correlation studies, it was not possible to decide which of the hydroxyl groups in 159 is acetylated. The complete structure and the relative

B. S. Joshi and S. W. Pelletier

332

Stereochemistry of the alkaloid were solved by a single crystal X-ray analysis. This established the structure as 2-acetylseptentriosine (160) [136]. Septatisine, C22H31NO3, (162), a crystalline atisane-type diterpenoid alkaloid, was isolated from the roots of A. septentrionale [137]. ^H and ^^c NMR spectral studies showed the presence of a normal diterpenoid alkaloid skeleton having two "additional" carbons attached to the nitrogen forming an ethanolamine grouping. Of the three diterpenoid alkaloidal skeleta, the hetisane and the veatchine-types were discounted in preference to the atisane structure with one additional bond between C(14)-C(20). The presence of two secondary hydroxyl groups and the third oxygen to form a cyclic ether were shown by ^H and ^^C NMR spectral studies. The sole methyl group was assigned to C(18). The atisane skeleton with a C(14)-C(20) bridge and the location of the carbinolamine-anchored cyclic ether, was established through selective INEPT studies. The complete structure of septatisine (162) was established on the basis of ^H COSY, LRCOSY, NOESY, DNOE, fixed-evolution HETCOR, and selective INEPT NMR techniques. It is noteworthy that septatisine (162) is the only example of an atisane-type diterpenoid alkaloid in which the C(20) carbon of the oxazolidine ring is bridged with C(14). In the case of atisine (153), the oxazolidine ring gives rise to a mixture of inseparable epimers differing in configuration at C(20) [138].

162 9.4

Stereostructure of septatisine 162

Tangutisine (163)

A. tanguticum (Maxim.) Stapf W. T. Wang has been reported to contain the known alkaloids benzoylheteratisine (135), atisine (153), heteratisine, and tanwusine; the structure of the last alkaloid was not determined [139]. We have described earlier (Section 8.3), the isolation and structure of the novel dimeric alkaloid tangirine (134), which is formed by joining of a heteratisine-type and a hetisine-type of alkaloids. From the same plant species, a diterpenoid alkaloid designated as tangutisine, C20H27NO4, possessing the hetisane skeleton was isolated; it has been assigned structure 163. Tangutisine forms a crystalline hydrochloride salt and a tetraacetate (164) [140]. On the basis of homonuclear ^H COSY, HETCOR, two dimensional

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

333

nOe, iH-l^c long range correlations (FLOCK) and selective INEPT [141] NMR techniques, the structure 163 was established for tangutisine. One of the four hydroxyl groups in 163 occurs as a tertiary alcohol associated with the quaternary carbon appearing at 5c 81.4 in the ^^C NMR. This OH group was assigned to C(14) on the basis of ^^C NMR spectral evidence after eliminating the possible substitution at C(5), C(6), C(9), C(12), and C(20). The remaining three secondary hydroxyl groups were shown to be situated on C(2), C(l 1) and C(13) from a detailed examination of the ^H and ^^C NMR data. A broad signal at 5H 4.21 in the ^H NMR was shown to be coupled to four upfield protons between 5H 1.62-2.90 in the COSY spectrum. The HETCOR experiment revealed that signals at 8 1.76 and 2.97 were for nonequivalent geminal methylene protons, as were signals for protons at 6 1.62 and 1.92. The carbinol proton signal at 5 4.21 was assigned to H(2) since this was the only position at which a proton could have scalar coupling with two adjacent methylenes. The appearance of H(2) as a broad singlet (W 1/2 = 10.4 Hz) required that H(2) must be |3-oriented. The coupling constants for H(2) and H(1|J), and H(2) and H(3p) were 4.1 and 4.3 Hz, respectively, typical of axial-equatorial coupling constants, confirming the a-orientation of the C(2) hydroxyl group. The stereochemistry of hydroxyl groups at C(ll) and C(13) was easily determined by the analysis of observed nOe's and vicinal coupling constants. The H(9) signal appeared as a doublet with a coupling constant of 8.8 Hz. This large scalar coupling must result from a near 0° dihedral angle between H(9) and H(ll), which is only possible if the OH(ll) group has an a-orientation (a dihedral angle of 180° between H(9) and H(l 1) is impossible in the hetisane skeleton). The (J-orientation of H(l 1) was confirmed by the observation of a dipolar interaction (nOe) between H(l 1) and H(15p). The observation of W-coupling between H(l 1) and H(13) in the COSY and LRCOSY spectra provided strong evidence to establish the stereochemistry of 0H(13). The H(13) proton should be p-oriented to form the W-shape with H(ll). The dipolar interaction in the COSY spectrum between H(13) and H(17a) confirmed this stereochemical assignment. Thus the stereo-structure of tangutisine was established as 163 A.

1

""r** H ] 1

J

H Me 163 R == H 164 R = Ac

163 A

334

B. S. Joshi and S. W. Pelletier

9.5

13-O-Acetylvakhniatine (165) and Ajabicine (166) We have listed earlier (Section 6.2), a large number of norditerpenoid alkaloids isolated from the aerial parts of C. ambigua L. P. Ball and V. H. Heywood (Syn. Delphinium ajacis L). Two diterpenoid alkaloids (C20) isolated from the seeds are ajaconine and dihydroajaconine [117]. Vakhmatine (151) [128] and 13-(9-acetylvakhmatine (165) were isolated from the seeds of C ambigua and the structure of the new alkaloid established on the basis of spectroscopic data and chemical correlation with 151 [10]. A novel amorphous diterpenoid alkaloid designated as ajabicine (166) was isolated in 0.0012% yield by chromatographic separation from the leaves of C. ambigua, cultivated in Assiyut, Egypt. Ajabicine, C22H33NO2, showed 22 resonances in the ^^C NMR spectrum comprised of one sp2 singlet, three sp^ singlets, seven sp^ doublets, nine methylenes, and two methyl carbons. The ^H NMR spectrum featured signals at 5 4.89, 5.00 (each IH, j , =C//2), 3.96 [IH, f, C//(OH)], 1.03 (3H, t, CH2C//5), 0.90 (3H, s, tert-CHs) [20]. The molecular formula and chemotaxonomic considerations together with the presence of an iV-ethyl group indicated that ajabicine must be a diterpenoid and not a norditerpenoid alkaloid. This conclusion

13

OH

Me^

22 \ 2 r

21 "1

r

>C|>ttlZ

-f%-

3 ^^ N r S S ^

i-S. 6 19 18 Me

166

-H ^H

14)

^

9 \^ ' ' ' % /

\/

7

16

15

OH

7

OH

15

22 M

Stereostructure of ajabicine

was supported by the absence of signals for methoxyl groups (^H and ^^C NMR), which are normally present in norditerpenoid alkaloids [14,15,55,58,117]. All the diterpenoid alkaloids containing 20 carbon atoms belong to two broad types: the atisine-type (incorporates an entatisane nucleus), and the veatchine-type (modeled on an ^«r-kaurane nucleus) [117]. These alkaloids contain an exocyclic methylene group at C(16) and possess three other quaternary carbons at C(4), C(8) and C(10). These ring systems are not consistent with a quaternary carbon signal bearing a hydroxyl group at 5 80,0. To account for this carbon signal, the C(8)-C(9) or the C(8)-C(14) bond in the bicyclic [2.2.2] octane system must be cleaved, to form a rearranged skeleton. The results of COSY, TOCSY, HETCOR, and COLOC experiments and an inspection of literature values previously reported for norditerpenoid and diterpenoid alkaloids [14,15,56,58,

335

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids I OH ^-CH2-CH-C=CH2 ^—CH-CH2-9H I H3C-CH2-N-CH-I -"^ | - C H ~ C H 2 - C H 2 - ^ ^_^^

a

b

c

117] clearly established the presence of partial structures a, b, and c in ajabicine. Besides these partial structures, a total of six degrees of unsaturation remained to be accounted for and these must consist of six rings. Connectivities of the remaining three quaternary centers, one methine, two methylenes and one methyl group were established by COSY, selective INEPT and FLOCK experiments and these data can be satisfactorily explained with structure 166 for ajabicine. Many of the assignments of the carbon atoms have been confirmed by selective INEPT experiments. The overall connectivity assignments were confirmed by the nOe studies (ID and NOESY). The biogenesis of ajabicine (166) may be speculated to proceed through a rearrangement of the fused bi-

Me

166

B

(Numbers in brackets are those as in A) Scheme 3

cyclo[2.2.2]octane A to give the bicyclo[3.2.1]octane rearrangement product (166) via the homo allylic carbocation B, which on hydration gives 166 as shown in Scheme 3. There are

B. S. Joshi and S. W. PeUetier

336

many precedents in the literature for the rearrangement of a bicyclo[2.2.2]octane system to the bicyclo[3.2.1]octane [142]. 9.6

Chellespontine (167) and Azitine (169) Consolida hellespontica (Boiss.) Charter. [Syn. Delphinium hellespontica (Boiss.) D. tomentosum (Boiss.)] grows in the Zonguldak and Kastmonu regions of Turkey at an altitude of 640 m. We investigated the alkaloids of the aerial parts of C. hellespontica and isolated seven alkaloids (A-G) by conventional chromatographic procedures [143]. The new crystalline diterpenoid alkaloid (A) designated as chellespontine (167) showed in the ^H NMR spectrum signals at 8: 0.84 (3H, 5, tert -Me), 4.00 (IH, br s, C//-OH), 5.10, 5.37 (each IH, br s, C=CH2 ), 9.40 (IH, s, CHO). The molecular formula, C22H33NO2, for the alkaloid was derived from its mass and ^^C NMR spectrum. Chellespontine showed 21 lines for 22 carbon atoms of the molecule and a DEPT spectrum revealed four non-protonated carbons at 5 156.4, 46.4, 38.1 33.4, five methines at 5 183.5, 75.0, 44.9, 40.0 36.3, twelve methylenes at 5 109.5, 64.5, 59.5, 58.3, 41.0, 35.0, 31.0, 28.1, 25.9, 25.9, 19.8, 19.4, and one methyl group at 5 24.7. Consolida and Delphinium alkaloids usually conform to two main group of diterpenoid alkaloids: the aconitine- or lycoctonine-type norditerpenoid alkaloids and the diterpenoid alkaloids having an atisane or veatchine-type ring skeleton [144]. The former class of compounds are usually methoxylated but not the latter. The 22 carbon atoms of chellespontine confirmed by its ^^C NMR spectrum together with the absence of methoxyl groups, excluded the norditerpenoid skeleton. The molecular formula of chellespontine indicated seven degrees of unsaturation, of which two are accounted for by the methylene and the aldehydic functions. The characteristic carbon resonances of an atisane- or a veatchine-type exocyclic methylene group were readily located at 5 156.4 (s) and 109,5 (t) for C(16) and C(17) and the protons at 8 5.10 and 5.37 for the exocyclic methylene group. Of the only two methine doublets downfield of 8 44.9 the signal at 8 183.5 (d) indicated an aldehydic carbon resonance and the signal at 8 75.0 (d), a carbon attached to an oxygen function. The remaining three methines are attached to carbons which are not attached to oxygen or nitrogen atoms. Chellespontine contains an exocyclic

RHgC

167 R = CHO 168 R = CH20H

169

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

337

methylene, an N-CH2CHO, and a secondary hydroxyl group. We noticed that in the ^H NMR spectrum the aldehydic proton attached to a methylene group appeared as a singlet at 8 9.40. A literature search showed that the aldehydic proton in N-CH2CHO has been reported to appear as a singlet [145]. The hydroxyl group of chellospontine should be located next to the exocyclic methylene group to account for the downfield shift of C(16) (8 155.6), On the basis of these and additional spectral data, we arrived at structure 167 for chellospontine [143]. The structure was confirmed by the reduction of chellospontine with sodium borohydride to afford a compound identical with dihydroatisine (168) which has been isolated from A. heterophyllum [46] and has also been prepared by the reduction of atisine with sodium borohydride [146]. The crystalline compound (B), C20H29NO, designated as azitine (169) was isolated from C. hellespontica during the chromatographic separation of the pH 10 alkaloidal fraction. The l^C NMR spectrum showed 20 signals for 20 carbon atoms of azitine. Resonances at 8c 156.6 (s) and 109.2 (t) and the two broad signals at 8 5.04 and 5.10 in the ^H NMR spectrum suggested that the alkaloid was of the atisane-type with an exocyclic methylene group. The signal at 8 3.69 (8c 75.8) is clearly assigned to the proton attached to a hydroxyl group and the two broad proton signals at 8 3.41 and 3.42 (8c 60.4) can be assigned to the N-CW2 g^oup. A lowfield methine proton at 8 7.87 indicated the presence of an N=CH group and is supported by the l^C signal at 8 166.2. The structure for azitine (169)was deduced from its NMR spectral data. The azomethine group is located on N-C(20) and not N=C(19) as established from the selective INEPT NMR experiments [143]. Azitine was shown to be identical with the azomethine prepared by the potassium permanganate oxidation of atisine (153) [147]. Compounds (C), (D), (E) and (F) were identified as the known norditerpenoid alkaloids 1-0-methyldelphisine [148], delphinine (88) [149], delphisine (86;R^ R2 = Ac) [150], and bullatine C [151], respectively. ^H and ^^C NMR spectra of the crystalline alkaloid (G) showed it to be a hydroxy-N-methyltrimethoxyaporphine, and comparison with an authentic sample proved its identity with (+)-corydine [152,153,154]. 9.7

Andersobine (170) D. andersonii Gray is a low larkspur growing in the 'Wildcat Hills' of Utah, at an altitude of about 4800 ft. Cattle deaths from grazing of D. andersonii have been observed. From the aerial parts of this plant, the isolation and structure determination of sixteen norditerpenoid alkaloids : 14-0-acetylbrowniine, 14-acetyldelcosine, 14-0-acetylnudicaulidine, andersonidine, andersonine, browniine, 14-deacetylnudiculine, delavaine, delcosine, delectinine, deltaline, dictyocarpine, lycoctonine, methyllycaconitine, nudicauline, and takaosamine has been described in section 6.7. In the course of our studies on the minor constituents of D. andersonii, we isolated a new polar diterpenoid alkaloid by droplet counter current chromatographic separation [4] designated as andersobine and determined its structure as 170 [9].

B. S. Joshi and S. W. Pelletier

338

The molecular formula, C22H29NO4, derived for andersobine from the low resolution mass spectrum and elemental analysis was confirmed by HRMS. The ^H and ^^C NMR spectrum in CD3SOCD3 indicated the presence of a tertiary methyl at 5 0.95 (5c 19.1), and an acetate methyl at 5 2.02 (5c 170.5, 20.8). The downfield part of the spectrum showed the presence of an exocyclic methylene group at 5 4.92,4.93 (5c 151.7,109.9). Biogenetic considerations and the absence of methoxyl, N-methyl or N-ethyl groups and the presence of an acetate, suggested that andersobine is a diterpenoid and not a norditerpenoid alkaloid. Of the various skeleta known for the diterpenoid alkaloids [117,144], andersobine possesses the hetisane skeleton as all the other skeleta have to be /V-alkyl derivatives. The partial structure of andersobine is thus expressed having one acetoxyl and two hydroxyl groups attached to the hetisane skeleton. The ^^C NMR spectrum showed 22 signals for all the carbon atoms of the molecule. An APT/DEPT spectrum revealed the expected five non-protonated carbon singlets at 5 170.5, 151.7, 48.5 (2C), 44.0, nine methine doublets at 5 87.6, 73.0, 71.8, 69.9, 61.7, 60.6, 43.5, 42.9, 33.0, six methylene triplets at 5 109.9, 32.5, 31.8, 28.0, 26.2, 25.6 and two methyl quartets at 5 20.8 and 19.1 ppm. A literature search for alkaloids (C20II27NO3), having the hetisane skeleton and possessing three hydroxyl groups, indicated six known diterpenoid alkaloids. However, the physical and spectral properties of andersobine were not in accord with the monoacetyl derivatives of these known alkaloids [155-159]. The three oxygen functions of andersobine may be located at C(l), C(2), C(3), C(7), C(ll), C(13), C(15) or C(19) positions. One of the hydroxyl or acetoxyl groups should be located at C(15), adjacent to the exocyclic methylene, as the quaternary carbon signal for C(16) appears at 5 151.7 because of the p-effect. By comparison with alkaloids bearing a C(15) hydroxyl group, the resonance at 6 71.8 in andersobine has been assigned to C(15). There are six methines in andersobine appearing downfield of 5 60.0. The methine signal at 5 87.6 indicated a carbinolamine carbon resonance suggesting the location of a hydroxyl or acetoxyl group at

H

170

Stereostructure of andersobine

'H

170

C(19). Such a carbinolamine carbon can be expected to appear around 90.0 ppm. As location of a hydroxyl group at C(20) will make this a quaternary carbon, this position is excluded. Of the remaining five methine carbon signals, two should be attached to the nitrogen atom C(6) and

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

339

C(20), two should be oxygenated and one will be the C(5) methine carbon, as in hetisine [119]. If the ring (A) is not oxygenated, a triplet for a methylene group (C2) near 5 19.8 would be expected [140,160]. The lack of this resonance locates the third oxygen function at C(l), C(2) or C(3) in ring A. The quaternary carbon signal of C(10) appears in the normal range at 48.5 ppm, and is not subjected to the p-effect of an oxygen function located at C(l). Of the five nonprotonated carbons, the signals at 8 170.5, 151.7 and 48.5 have been assigned to C(21), C(16), and C(10), respectively. The signal at 8 44.0 should be assigned to C(8) as in hetisine and the remaining quaternary carbon resonance at 8 48.5 is assigned to C(4). On the basis of these data, andersobine was assumed to bear hydroxyl groups at C(3), C(5) and C(19), with one of these being an acetate group. Two separate attempts to solve the structure of andersobine by X-ray crystallography were unsuccessful in spite of the availability of suitable crystals. In an effort to make other derivatives suitable for X-ray as well as chiroptical studies, the 4dimethylaminobenzoate ester of andersobine was prepared. X-ray crystallographic analysis of these crystals also failed. Finally, the structure of andersobine as 170 was confirmed by detailed ^H COSY, one dimensional (ID; in C5D5N), two dimensional (2D; in C5D5N and CD3SOCD3) nOe NMR studies, and selective INEPT experiments. Stereochemical assignments of andersobine (170) were confirmed by measurement of vicinal coupling constants as well as the observation of ID nOe's in C5D5N. The equatorial orientation of the C(3) OH group enables the H(3) axial proton to be seen with a large vicinal coupling constant for the axial-axial (11.4 Hz) and a smaller axial-equatorial (5.5 Hz) relation resulting in a double doublet at 8 3.83. This result can be explained by H(3) being in a p configuration, coupling with its neighboring H(2) axial proton for an A ring in a chair conformation. The H(3) proton also showed a NOESY relation with the H(5) proton thus, establishing their 1,3-diaxial relationship. A strong nOe between H(19a) and H(20) both in ID and 2D nOe spectra indicated that the OH group at C(19) is in a (J-position. H(19a) also showed an nOe in ID and 2D nOe spectra with H(6) and not with H(5p) as expected. The stereochemistry of the acetoxyl group at C(15) in an equatorial position [boat conformation of the D ring formed by C(8), C(14), C(13), C(12), C(16), and C(15)] was established by the NOESY observed between H(15a) and H(14) protons and the absence of an nOe or NOESY relationship between H(15), and H(9)[9]. 9.8

Delatisine (171) D. elatum L. has been a rich source of norditerpenoid and diterpenoid alkaloids. Delpheline, deltaline, deltamine, elatine, and methyllycaconitine were isolated from the whole plant. However, the following norditerpenoid (i-xiv) and diterpenoid (xv) alkaloids were found to be present in the seeds: i) 14-acetylnudicauline, ii) delectinine, iii) delelatine, iv) delpheline, v) deltaline, vi) eladine, vii) elanine, viii) elsasine, ix) elatine, x) isodelpheline, xi) lycoctonine, xii) methyllycaconitine, xiii) ivudicauline, xiv) pacinine [14,15] and xv) ajaconine [117,143]. A novel furanohetisine-type crystalline diterpenoid alkaloid designated as delatisine was isolated from the seeds of D. elatum by chromatographic separation [161]. The molecular for-

340

B. S. Joshi and S. W. Pelletier

mula, C20H25NO3, was derived from the HRMS. The ^^C NMR and DEPT spectrum indicated four non-protonated carbons at S 145.7, 52.7, 50.5, 45.7 ten methines at 5 100.2, 79.6, 75.7, 72.2, 66.3, 64.4, 62.0, 55.4, 50.2, 50.0, five methylenes at 5 108.2, 41.6, 37.3, 34.3, 33.9, and one methyl group at 5 21.9. Carbon and proton resonances characteristic of an exocyclic methylene and a tertiary methyl group were evident. There are six methines downfield of 5 62.0 and the signal at 5 100.2 indicated a carbinolamine carbon. Of the remaining five signals, two should be attached to the nitrogen and three should be oxygenated. The molecular formula of delatisine raised the question of attaching the three oxygen functionalities to the hetisane skeleton to give four oxygenated methine carbons. This can only be achieved when there is one ether functionality and two hydroxyl groups in the molecule. At least one oxygen function should be located in the A-ring, because of the lack of a triplet carbon signal about 8 19.8 where C(2) resonance would appear if no oxygen functionality exists from C(l) to C(3). A methine singlet at 5 4.67, correlated with the methine carbon at 5 100.2 and this proton showed interactions with H(18) methyl, H(20), and H(6) in the 2D-nOe spectrum, indicating that this signal belonged to H(19). Thus one oxygenated carbon was located at C(19), and the remaining three must be at C(l), C(2), C(3), C(l) or C(13). The location of a hydroxyl group at C(15) and C(7) was dis-

HOaH

Stereostructure of delatisine

H

171

counted on the basis of the chemical shifts of C(16) and C(8). Long range (LR) COSY spectral analysis indicated that only C(2) is oxygenated and C(l), C(3) are aliphatic methylene groups. The triplet at 6 4.50 (5c 79.6) was assigned to H(2) since no other location is possible for a CH2-CH(OH)-CH2-system. The coupling constant of H(2) with its neighboring protons suggested that H(2) was P-oriented with only axial-equatorial and equatorial-equatorial vicinal relationships with neighboring protons. A selective INEPT experiment confirmed the ether linkage between C(2) and C(19). With the C(2)/C(19) ether linkage established, only one possibility remained: location of the two hydroxyl groups at C(ll) and C(13). The proton at 5 4.11 was assigned to H(ll) and the large vicinal coupling (8.6 Hz) between H(9) and H(ll) suggested that H(ll) is p-oriented having a 10^ dihedral angle. The H(13) at 5 4.25 is also P-oriented due to the large vicinal coupling constant (9.1 Hz) between H(13) and H(14). The stereochemical assignments for the hydroxyl groups at C(ll) and C(13) were confirmed in the COSY spectrum

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

341

and structure 171 was thus assigned to delatisine. Since the structural evidence for delatisine was entirely based on NMR spectral data, a single crystal X-ray diffraction was performed and the structure confirmed. The alkaloid contains six, six-membered rings and four, five-membered rings. The rings A and C are mostly chair-like conformations with a moderate degree of puckering. Ring B in contrast, is best described as a highly puckered boat conformation. Rings D, E, and F are all part of the bicyclo[2.2.2]octane system and highly puckered, nearly boat conformations. Rings G and J are twist conformations and ring I is nearest to an envelope conformation; ring H is a nearly equal mix of twist and envelope conformations. A number of alkaloids possessing an oxide ring between C(l) and C(19) have been isolated [157,162-164]. The structure of delatisine is unusual in that there is no other diterpenoid alkaloid which contains an oxygen bridge between C(2) and C(19) to form a furan ring [161]. 9.9

Tatsirine (172) Apart from the norditerpenoid alkaloids described earlier (section 6.6), D. tatsienense Franch contains the known diterpenoid alkaloids ajaconine (173), hetisine (146), hetisinone (174), and dictyzine (175) [117,143]. Dictyzine was first isolated from D. dictyocarpwn DC in 1978 and its structure was based on an X-ray crystal structure determination [165]. This alkaloid was also isolated from D. brunonianum Royle [166] and D. tatsienense Franch [17,21]. The ^^C NMR chemical shifts were based on chemical shift rationales [17]. Unambiguous proton and ^^C NMR chemical shift assignments for dictyzine (175) were accomplished in an independent study of the DEPT, COSY, fixed evolution HETCOR, NOESY and selective INEPT NMR techniques [19]. A crystalline alkaloid designated as tatsirine, C20H27NO3, was isolated from the roots of D. tatsienense Franch by chromatographic separation. It was assigned structure 172 on the basis of ^H COSY, fixed evolution HETCOR, two-dimensional nOe and selective INEPT studies on 176 [21]. The l^C NMR spectrum of tatsirine showed 19 signals for 20 carbons at 5 149.1,106.6, 97.9, 70.6, 67.4, 66.7, 60.9 (2C), 51.8, 49.4, 48.5, 44.8, 42.9,42.3, 41.6, 36.8, 33.9, 32.4,

HCh

172

Stereostnjcture of tatsirine

342

B. S. Joshi and S. W. Pelletier

31.2, and 22.4. The exocyclic methylene signals at 6c 149.1 and 106.6 (5H 4.73, 4.85) suggested a hetisane-type skeleton and the singlet signal at 5 97.9 is clearly due to the carbinolamine carbon. These results indicated the location of one of the three hydroxyls at C(6). On the basis of the chemical shifts of C(8) and C(10), the location of hydroxyls at C(7) or C(15) positions was excluded. Since there is no resonance for a methylene carbon in the region 5 19.8, C(2) probably bears a hydroxyl group. This evidence leads to the location of the third hydroxyl group at C(ll) or C(13) and it is difficult to make an assignment on simple NMR experiments. We therefore decided to carry out high field NMR studies. The sample of tatsirine in methylene chloride was recovered after preliminary NMR studies by evaporation and purified on an alumina column. Spectral data of the recovered sample showed that the exocycHc methylene group had migrated into the ring as in 176. This was seen bytiiedisappearance of the exocyclic methylene signals and the appearance of a single vinyl resonance (8H 5.54, s) and a vinylic methyl group

HO-

174

173

(8H 1.83). Isomerization of the double bond must have taken place during the exposure to traces of acid in CDCI3 followed by contact with alumina. The key question of locating the third hydroxyl group with the carbinol signal at 5 3.53 brs (5c 75.4) was decided by examination of the COSY spectrum. From the narrow half width of the carbinol singlet [167], the carbinol proton was assigned to a pseudo-axial position (a) in the boat conformation of the ring defined by C(8), C(9), C(ll), C(12), C(13), C(14), with the hydroxyl group pseudo-equatorial (p) pointing to the exterior face of the molecule. This result suggested that the hydroxyl occupies the C(13P) position and all these results were confirmed by 2D nOe experiments leading to structure H .OH 1

Me-

-N 1

> 1

?>CH

Me 175

"""OH ^

OH

HOv

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

343

(172) for tatsirine. There are very few hetisane-type diterpenoid alkaloids in which a hydroxyl group at C(13) is located in a P position (equatorial hydroxyl in the boat conformation of the ring formed by carbons 8, 9, 11, 12, 13, and 14). Two examples are spirasine Xlll and spirasine XV, isolated from Spirea japonica L. \wr,fortunei (PL) Rehd. (Rosaceae) [167]. The facile isomerization of the exocyclic double bond of tatsirine (172) to afford 176 can be explained by the formation of a carbonium ion (with traces of HCl from CDCI3), which is stabilized with the suitably placed |3-hydroxyl group to form an oxitane. On alumina, the more stable isomer is readily formed by loss of a proton from C(15). Similar isomerization of the exocyclic methylene group takes place under acidic conditions, e. g. kaurene to isokaurene or hetisine to a mixture of isomeric products [168]. 9.10 Barbisine (177) Vacognavine (142) and 15-0-deacetylvacognavine (143) isolated from A. palmatum are the only examples known oiN-C{\9)-seco diterpenoid alkaloids [102,112,113]. The structure and stereochemistry of barbisine (177), a novel diterpenoid alkaloid from D, barbeyU were deduced by a combination of NMR spectra and a single crystal X-ray diffraction analysis [169]. Chromatography of the polar alkaloidal fraction of D. barbeyi gave barbisine.

177 178

R^ = H; R2 = OAc; R^ = p-OH R^ = OAc; R^ = OH; R^ = a-OAc

179 180

"="°-0

The presence of a hydroxyl group, three ester functions, two acetates, and one benzoate group was evident from the NMR data. A spin decoupling experiment established that two of the three ester functions were located on adjacent carbons of ring A. The 3.1 Hz coupling constant excluded an axial-axial relationship of the vicinal protons, but either axial-equatorial or equatorial-equatorial orientation remain. In the ^H NMR spectrum of barbisine monoacetate, a signal at 8 4.78 (d, / = 4.5 Hz) coupled to another at 8 3.43, corresponding to a carbinyl resonance seen at 8 3.78 in 177, establishes the presence of a secondary alcohol in 177. These protons are likely to be cis to each other. A small coupling constant (4.5 Hz) between H(9) and H(ll) and nearly zero between H(ll) and H(12) can be rationalized by locating the secondary

344

B. S. Joshi and S. W. Pelletier

hydroxyl group of 177 at C(ll) and oriented p. The aldehydic group (5H 9.23, s, H-19; 8c 196.6, C-19) was located at C(4) by analogy with the known vacognavine derivatives and comparison of their ^^c NMR cheniical shifts. The negative Cotton effect exhibited by 177 (also by vacognavine) near 300 nm, indicated that the keto group is at C(13) (111,119). The remaining problem was to establish the location of the third ester group which could be at C(7) or C(15). In view of the uncertainties, an X-ray structure analysis of barbisine was carried out which established its structure as 177 [169]. An alkaloid designated as barbaline (178), closely related to barbisine (177) has been isolated from the same plant [170]. Its structure was determined by NMR spectroscopy and a single crystal X-ray diffraction analysis. The absolute stereochemistry of barbaline has been determined to be: 15, 2/?, 3S, 45, 55, 6^, IR, %R, 9R, 10/?, 115, 12R, US, 20R. 9.11 Davisinol (179), 18-0-BenzoyIdavisinol (180), and Davisine (181) Three new hetisane-type diterpenoid alkaloids, davisinol (179), 18-(9-benzoyldavisinol (180) and davisine (181) were isolated from Delphinium davisii Munz., a plant native to Turkey [171]. The structures of these alkaloids were estabhshed by detailed NMR spectroscopic studies. The known diterpenoid alkaloids hetisine (146), hetisinone (174) and the norditerpenoid alkaloids 14-0-acetylperegrine, 6-deacetylperegrine [172, 173] and karakoline [14] were also isolated from this plant. Accurate ^H and ^^c NMR assignments were made for kobusine (182) [117] and karakoline. Chromatographic separation gave an amorphous alkaloid, C20H28NO2, designated as davisinol (179). Biogenetic considerations and the absence of a methoxyl or N-methyl group indicated that, davisinol is a hetisane-type alkaloid [9,137]. The ^^C NMR spectrum showed 20

181 183

R =H R = Ac

182

Kobusine

signals and the DEPT spectrum revealed four non-protonated carbon singlets at 5 145.8, 49.6, 43.5, 40.5, seven methines at 5 75.7, 67.4, 64.8, 59.5, 56.0, 44.0, 41.9, and nine methylene triplets at 5 110.1, 69.2, 58.2, 35.8, 33.6, 29.6, 28.4, 26.5, 18.9. Characteristic carbon and proton signals for an exocyclic methylene group were observed at 5 4.83 (2H, d, / = 1.8 Hz), 145.8 (s), 110.1 (t). Of the two low-field methylene triplets at 5 58.2 and 69.2, the former signal was assigned to C(19) (5 2.55, 2.23 AB, J = 12.5 Hz). This assignment was supported by

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

345

the NOESY correlation between H(19) (S 2.23) and H(20) (5 2.40) and the HMBC correlation of H(20) with C(19), C(6) (64.8), C(8) (43.5) and C(13) (29.6). As the normally observed signal for a C(4) methyl is absent in the ^H and ^^C NMR spectrum, the methylene signal at 5 69.2 (5 3.28, 3.43 AB, J = 10.8 Hz) should be assigned to C(18) bearing a hydroxyl group. This assignment was confirmed by the HMBC spectrum which showed a three-bond correlation of H(19a) at 6 2.55 with C(18), and H(18b) at 5 3.28 with C(5) (8 56.0; 5 1.72). In this partial structure of davisinol with an OH at C(18), the location of the second oxygen function which can only be a hydroxyl group, remained to be decided. When no hydroxyl group is present at C(l), C(2) or C(3) in ring A, the signal for C(2) appears around 5 19.8 [140,166]. Davisinol therefore, does not bear an OH in ring A. The remaining methylenes on which a hydroxyl may be located are C(7), C(ll), C(13) or C(15). The C(6) resonance at 5 64.8 showed correlation with 5 3.14 in the HETCOR spectrum, and this proton showed COSY correlation with H(7a), H(7P) (5 1.65, 1.57); C(7) does not therefore, bear a hydroxyl group. The C(16) quaternary carbon resonance at 6 145.8 indicated that C(15) does not bear an OH group [9]. This leaves the location of the OH group at C(l 1) or C(13) Positioning of the hydroxyl group at C(l 1) and not at C(13) was based on COSY and HMBC results. It is necessary to first identify either the C(9) or the H(9) signal in the NMR spectra. In the HMBC spectrum, the methylene for H(19) showed correlations with C(3) (5 28.3), C(18) and C(20), all three bonds away. In addition, correlation with the quaternary carbon at C(4)(8 43.5) was observed. The H(20) proton at 8 2.40 shows correlation with H(14) at 8 1.78, H(6) at 8 (3.14, brs), and H(18b) at 8 3.28. It also showed a W-type coupling with H(5) at 8 1.72 and a long-range coupling with H(la) at 8 1.80. The H(14) proton (8 1.78) showed a W-type coupling with H(9) (8 1.38) and H(20) (8 2.40). The H(9) proton was correlated with 8 59.5 (d) in the HETCOR and appeared as a sharp singlet. The H(9) proton showed in the COSY spectrum, correlation with H(ll) (8 4.01, d, 7 = 4.8 Hz) and a W-type coupling with H(12) (8 2.28, d, / = 4.8 Hz) and H(14), as stated earlier. These correlations are possible only by location of the hydroxyl group at C(l 1). This conclusion was supported by the HMBC spectrum in which the H(9) proton (8 1.38) was correlated with C(14) (8 44.0), C(5) (8 56.0), C(20) (8 75.7), and C(ll) (8 67.4). These correlations would not be possible if the hydroxyl group was located at C(13). Firstly, H(9) would not be a singlet, if the adjacent group at C(ll) was a methylene and secondly, this proton would not show a COSY correlation with a proton at C(13) bearing a hydroxyl group. The evidence for locating an OH at C(ll) was obtained by a few other sets of correlations. The C(17) methylene protons at 8 4.83 showed a long-range COSY correlation with H(15) (8 2.10, m). In the HMBC spectrum, a slice at 8 2.10, H(15), showed response of signals for C(9) (8 59.5), C(17) (8 110.3) both three bonds removed and C(8) (8, 40.5), and C(16) (8, 145.8) two bonds away. In the HMBC spectrum, a slice at 8 4.83 for H(17) showed responding carbon signals for C(15) (8 33.6) and C(12) (8 41.9). In the COSY spectrum H(12) was correlated with H(l 1), H(13a), H(13p) and H(9). These results indicated

346

B- S. Joshi and S. W. Pelletier

the location of an OH group at C(l 1). Some of the ^H NMR assignments were confirmed by selective irradiation experiments. Irradiation of H(7a) at 5 1.65 collapsed the H(7p) signal at 6 1.57, and irradiation of H(15a) at 5 2.10 brought a change of the signals for H(15|J) at 5 L95. Similarly, irradiation of H(19P) at 8 2.23 showed a change of the signals for H(19a) at 5 2.55. The hydroxyl group at C(l 1) can be either ^-axial (shown as a dotted line) or W- equatorial (shown as a thick line) in the twist boat ring conformation formed by C(8), C(9), C(ll), C(12), C(13) and C(14). When the OH group is ^'-axial, H(ll) should show a large coupling (-- 9-10 Hz) with H(9) (almost 0-2^) and - 4-5 Hz coupling with H(12) (-- 30^). In davisinol, the coupling with H(9) is almost nil (~ 90^) and the coupling with H(12) is ~ 5 Hz (almost 45°) as expected for a ^ equatorial OH group, confirming the structure 179. In order to enable structure derivation by spectral data, we undertook detailed NMR studies of kobusine [174], the absolute stereochemistry of which was established as 182. Another fraction gave a new alkaloid, C27H31NO3. The ^H NMR 8 (8.02, 2H, d, 7 = 7.5 Hz, H-2', 6'), 7.46 (2H, dd, J = 7.6 Hz, H-3', 5'), 7.58 (IH, dd, J = 7.4 Hz, H-4') and the 13c NMR spectral data 8 (129.6, 2C, C-2', C-6'), (128.5, 2C, C-3', C-5'), 133.1 C(4'), indicated that this is a benzoate ester of a hetisane-type alkaloid. The resemblance of the ^^C NMR chemical shifts with davisinol suggested that this alkaloid might be 18-benzyoldavisinol (180). Alkaline hydrolysis of this alkaloid gave davisinol (179) confirming the structure assignment. The ^H NMR spectral data established the location of the benzoate group at C(18) and not the C(ll) position. The signal for the H(ll) proton in davisinol and the benzoate is unchanged and appears at ~ 8 4.00. However, the H(18) protons of 179 at 8 3.28 and 3.43 are shifted downfield to 8 4.08 and 4.24 in the benzoate ester establishing structure 180 for 18-0benzoyldavisinol. Supporting evidence was obtained from the HMBC spectrum. Separation on an AI2O3 rotor of a Chromatotron gave another crystalline alkaloid, C20H27NO2, designated asdavisine (181). The ^H NMR spectrum exhibited signals for the presence of an exocyclic methylene (8 4.94, 4.96, br d), a tertiary methyl (8 1.01, 3H, s) and indicated the absence of methoxyl groups. The ^^C NMR spectrum and DEPT experiments indicated the presence of one methyl at 8 28.4, seven methylenes at 8 108.8, 62.8, 33.1, 32.5, 26.8, 27.8 and 27.1, eight methines at 8 75.6, 71.3, 66.1, 65.7, 56.5, 43.4, 41.3 and 33.7, and four quaternary carbons at 8 156.2, 55.0, 45.8 and 37.5. The molecular formula indicated eight degrees of unsaturation of which one is accounted for by the presence of an exocyclic methylene. The remaining seven degrees of unsaturation indicate the presence of a heptacyclic skeleton as in hetisane-type diterpenoid alkaloids. As there are no other unsaturations in the molecule, both the oxygens of davisine must be present as OH groups. This conclusion is also supported by the preparation of diacetyldavisine (183). The C(16) carbon signal at 8 156.2 indicates that there is an OH group located at C(15), the absence of which would have caused C(16) to appear around 8 141-148. The remaining hydroxyl should be present in ring A, because of the absence of a methylene in the ^^C NMR --8 18-19. In hetisane-type alkaloids, C(10) normally appears at ~ 8 49-51 and in davisine, the downfield singlet at 8 55.0 is assigned

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

347

to C(10), this shift being ascribed to the presence of an adjacent hydroxyl group. There are eight methine carbons in davisine of which four signals at 5 75.6, 71.3, 66.1 and 65.9 must be due to carbons attached to either nitrogen or oxygen. These are assigned to C(20), C(15), C(l) and C(6), respectively, on the basis ^H NMR and HMBC experiments. The configuration of C(l) OH (8 66.1) in 181 is considered as P, by comparison of the chemical shifts with those reported for lassiocarpine (5 70.4) [175], napelline (8 70.5) [176], lusidusculine (8 69.9) [177], and songorine (8 70.1) [178], which all bear an a OH at C(l). As a general rule, in norditerpenoid alkaloids, when C(l) has an a OH group it appears at - 8 72-73 whereas, when C(l) has a p OH group it appears at -- 4 ppm downfield (8 68-69) [14]. When the A ring in diterpenoid alkaloids has a C(l) a OH group it has been shown to take a boat conformation because of an intramolecular N - H....0 hydrogen bond. In the case of a C(l) P OH group, an intramolecular hydrogen bond is no longer possible and the A ring will assume a chair conformation. The H(l) proton in 181 at 8 4.19 and in 183 at 8 5,25 appears as broad peaks having Wl/2.'-5.5 Hz, indicating that the C(l) OH group should have a P configuration [171]. 10.

REARRANGEMENT REACTIONS OF DITERPENOID AND NORDITERPENOID ALKALOIDS

10.1 Acid-Catalyzed Rearrangement of Hetisine (146) When treated with 5% aqueous sulfuric acid or 10% hydrochloric acid, hetisine (146) gives two unusual rearrangement products 184 and 185 in a ratio of 19:1 [168,179]. The structure of the rearrangement product 184, which was also isolated from A. heterophyllum [179] was determined by an X-ray crystallographic determination [168]. Structure 185 was deduced from the ^H and ^^C NMR spectra. The mechanism for the formation of 184 from hetisine (Scheme 4) is shown by path A. Protonation of the double bond and a retroaldol reaction involving the OH(ll) group would give the aldehyde (186) which through the formation of a

HO^

184

185

hemiacetal with 0H(13) would cyclize to give the acetal (184). The rearrangement of the minor product (Scheme 4) takes path B, and involves the cleavage of the C(12), C(13) bond to form an

348

B. S. Joshi and S. W. Pelletier

intermediate aldehyde (187) which on dehydration gives 185. Under similar acidic conditions, 1 l-epZ-hetisine gave 184 and 185 in a ratio of 1:3, suggesting that changing the configuration of the OH (11) group makes the cleavage of the C(l 1), C(12) bond more difficult. Prolonging the heating of 146 with acid, gave not only 184 and 185, but also two isomeric products 188 and 189 in a ratio of 1:1. The structures of the new compounds were determined by X-ray crystal analysis. The rearrangement products 188 and 189 are probably formed through the intermediate 186. The configuration of the hydroxyl at C(l 1) has undergone a change and is p in both the compounds. However, the configuration of the hydroxyl group at C(13) is a or |J, as a result of the addition of water on either the a or P face of the stabilized allyl carbocation. Refluxing a mixture of 188 and 189 in 5% sulfuric acid gave small amounts of 184 and 185 indicating that the reactions are reversible. 10.2 Acid-Catalyzed Rearrangement of 11-Dehydrohetisine (190) and 2,11Didehydrohetisine (191) 11-Dehydrohetisine (190), and 2,11-didehydrohetisine (191) were obtained by Sarett oxi-

• - 184

185

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

HQ

349

Ha

189 dation of hetisine (146) [119]. Compound 191 was also obtained by the alkaline hydrolysis of 13-acetyl-2,l 1-didehydrohetisine, prepared by the oxidation of 13-acetylhetisine. 1 l-Dehydrohetisine (190) when refluxed with 45% sulfuric acid gave an interesting rearrangement product 192 [125]. The mass spectrum showed that the rearrangement product is isomeric having the molecular formula C20H25NO3. The structure of this compound was solved by single crystal Xray diffraction analysis of its perchlorate salt. ^H and ^^C NMR spectra showed the disappearance of the exocyclic methylene group. An interesting feature of this rearrangement product is an additional carbonyl group and the formation of a new ring incorporating the exocycHc methylene group. The rearrangement product of 190 contains five six-membered rings and two five-memHQ..

190

191

bered rings fused together. Ring A [C(l), C(2), C(3), C(4), C(5), C(10)] is a flattened distorted chair; ring B [C(5), C(6), C(7), C(8), C(9), C(10)] is also a flattened chair; ring C [C(9), C(ll), C(12), C(13), C(14), C(8)], ring D [C(8), C(14), C(13), C(17), C(16), C(15)] and the piperidine ring E [C(6), C(7), C(8), C(14), C(20), N] are also in chair conformations. The two five-membered rings F [C(4), C(5), C(6), N, C(19)] and G [C(8),C(9), C(10), C(20), C(14)] both have distorted half-chair conformations. The carbonyls at C(ll) and C(16) of the eightmembered ring [C(8), C(9), C(ll), C(12). C(13), C(17), C(16), C(15)] are almost parallel. Similarly, refluxing 191 with 45% sulfuric acid gave 193, The structure of this compound was deduced from NMR spectral data and confirmed by oxidation of 190 (the structure of which was

350

B. S. Joshi and S. W. Pelletier

192 established by an X-ray analysis) with pyridinium chlorochromate to afford 193. A plausible mechanism for the acid-catalyzed rearrangements of 190 and 191 is shown in Scheme 5 involving hydration of the exocyclic methylene group followed by dehydration and enolization to give the intermediate 194. An internal Michael addition would afford the compounds 192 and 193 [125].

190 R = a-OH 190 R = a-OH 191 R = =0

192

or 193 Scheme 5 10.3 Base-Catalyzed Rearrangement of ll-AcetyN2,13-didehydrohetisine (197) Acetylation of hetisine (146) gave a mixture of several acetates which were separated by chromatographic purification to afford 2-acetyl, 13-acetyl, 11-acetyl (195), 2,11-diacetyl (196),

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

351

11,13-diacetyl, and 2,11,13-triacetylhetisines. Oxidation of 11-acetylhetisine (195) with Sarett's reagent gave ll-acetyl-2,13-didehydrohetisine (197) [119]. Heating 197 with aqueous potassium carbonate in methanol under reflux gave a crystalline compound, C20H25NO4 [180], which formed crystalline prisms of the perchlorate salt. The infrared spectrum showed the presence of a hydroxyl, a carbonyl and a 5-lactone group. The ^H NMR spectrum indicated two tertiary methyls, one at C(4) and the other attached to an oxygen. The singlet at 8 2.78 is assigned to C(9)-H and the signals at 5 3.10-3.50 (3H, m) to the overlap of C(6), C(13), and C(20) protons. The absence of the exocyclic methylene group in the ^H and ^^C spectrum suggested that this is a rearrangement product. The structure of compound 198 was determined by an X-ray analysis of the perchlorate salt (Figure 2) An interesting feature of this structure is the skeletal rearrangement which resulted in the formation of a 5-lactone under alkaline hydrolytic conditions [180]. 0 122)

C112)

0123)

CUB)

0121)

Figure 2. ORTEP plot of (198) (HCIO4 not shown).

Ra.

195 R = H 196 R = Ac

352

B. S. Joshi and S. W. Pelletier

Ha

AcOv

198

199

10.4 Base-Catalyzed Rearrangement of 13-Dehydro-2,ll-cliacetylhetisine (199) 13-Dehydro-2,l 1-diacetylhetisine (199) was prepared by Sarett oxidation of 2,11-diacetylhetisine (196) [119]. Treatment of 199 with methanolic aqueous potassium carbonate at 25® C gave a mixture of products from which the normal deacetylation product 13-dehydrohetisine

OvJ> AcO..

Me

HO..

201 could not be isolated. However, an isomeric rearrangement product was obtained which was shown to be 2-acetyl-13-dehydro-ll-^p/-hetisine (200) [180]. The structure proof for 200 depended on the ^H NMR spectral evidence. The 11-a proton showed a doublet at 5 4.14 with a coupling constant of 5.1 Hz. This is in agreement with a dihedral angle of -- 12(P between the C(lla) and C(9P) protons. The reaction of 199 with aqueous methanolic potassium hydroxide under refluxing conditions gave the rearrangement product (201). The structure assignment of this compound was made by oxidation of 201 with chromium trioxide-pyridine to give the ketolactone (198), described earlier [180].

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

353

0

4,-.-

.#

197 or

OH-

R

#1 i

• ,4" I

"

I

#

"*

"

OH-

~

"

199

~

' 197 199

|

..._ "--

"Mr.

R=(x-OH R = OAc

/

i

_.._

/'%"e

-

197

R=(x-OH R = OAc

~

199

l

200

q. K2CO 3 MeOH

O,H

HO • 198

or

201

-.

m o ~'

/"

Scheme 6 A plausible mechanism for the formation of the rearrangement products 198 and 201 is shown in Scheme 6. The first step appears to be the hydrolysis of the C(11) acetyl group under mild hydrolysis conditions. The C(11) acetyl group is more susceptible to hydrolysis than the C(2) acetyl group. The hydrolysis is accompanied by a reverse aldol condensation as is evidenced by the formation of the C(11) epimer 200 on work up of the reaction. Under refluxing conditions, a Cannizaro-like process involving attack of a hydroxide on the formyl moiety followed by intramolecular hydride transfer, takes place [ 181 ]. On work up under mildly alkaline conditions, the rearranged lactone 198 or 201 is obtained. 10.5

Acid-Catalyzed Rearrangement of Isoatisine (204)

An acid-catalyzed allylic rearrangement of garryfoline (202) having the [3.2.1]bicyclooctane ring occurs at room temperature to afford cuauchichicine (203) [182]. However, when

B. S. Joshi and S. W. Pelletier

354

202 212

203

R = P-OH R = a-OH

isoatisine (204) containing a [2.2.2]bicyclooctane ring system, was kept in 7% aqueous hydrochloric acid for 7 days it gave 205 in 50% yield, and a mixture of two epimeric methyl ketones 206 and 207. Only the compound 206 could be isolated in pure form. The structures of these compounds were determined by ^H, l^c NMR spectra, including nOe difference, and 2D COSY [183]. Compound 206 was identified by oxidation of tetrahydroatisine (208), the structure of which was determined by an X-ray crystal analysis [184]. J"*

Ri

r1

R2

4

/I < OH $*•

-'«AA/*

206 207

R^ = Me; R^ = H R^ = H; R2 = Me

^Me

1 \ 1 \

T*'"H

H 1

"^DH

1

le 208

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

204

HCI

355

*-

209 + H2O -H +

-H2O

HCI 20% NaOH 206 + 207

20% NaOH 205 p-TsOH,
'

>M, Scheme 7

A plausible mechanism for the isomerization is given in Scheme 7. The carbocation (209) is generated by protonation of the exocyclic methylene group. At room temperature, by the addition of a molecule of water 209 gives 210, which on basic work up affords the diol (205). However, under refluxing conditions, 209 undergoes a pinacol-type hydride shift from the a face to 211, which upon epimerization gives the methyl ketones (206) and (207) [183]. 10.6 Acid-Catalyzed Isomerization of Dihydroveatchine (213) In contrast to garryfoline (202), the 15-epimer veatchine (212) is stable even to boiling hydrochloric acid. The acid-catalyzed isomerization of dihydroveatchine (213) resulted in the isolation of the aldehyde (214). The structure of the product was derived from NMR studies including ID, 2D and selective INEPT experiments [185]. The reaction was carried out in DCl and D2O to study the mechanism of the reacdon. On the basis of the results, a pionacol-type mechanism involving dehydration, rehydration and an allylic rearrangement as shown in Scheme 8 has been suggested. Two deuterium atoms were incorporated in the molecule 215.

B. S. Joshi and S. W. Pelletier

356

H

OH

214

213

215 Scheme 8 10.7 Epimerization of the C(l)-a-Hydroxyl group of Delphisine (86, R1,R2 = Ac) and 8-0-Acetylneoline (219) Some norditerpenoid alkaloids, e.g. aconitine (91), mesaconitine (N-methyl instead of N-ethyl in 91), 3-deoxyaconine, delphinine (88), 8,14-diacetyltalatizamine (101) and yunaconitine (39, R = Ac) containing an acetoxyl group at the C(8) position, upon heating with methanol, ethanol or propanol form the corresponding C(8)-alkyloxy derivatives [32,39,186]. A plausible mechanism for this conversion has been suggested earlier (Section 5.4).

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

^OMe OH

[ ..\—OAcI

Et —

R^ Et —

OMe

OMe

216 R = Me 217 R=: Et 218 R= H

OMe

OMe 219 R^ = a-OH, R2 = Ac 220 R^ = p-OH. R2=Me

Solvolysis of delphisine (86, R1,R2 = Ac) in refluxing methanol and ethanol gave 8-0deacetyl-8-6)-methyl-l-ep/-delphisine (8-0-methyl-14-acetyl-l-^p/-neoline) (216) and 8deacetyl-8-6)-ethyl-l-ep/-delphisine (217), respectively [187]. Similarly, heating delphisine in boiling water gave 14-0-acetyl -l-ep/-neoline (218) and solvolysis of 8-0-acetylneoline (219) in methanol gave 8-0-methyl-l-ep/-neoline (220). Thus, during solvolysis of these alkaloids bearing a C(l) a hydroxyl group, epimerization of the C(l) hydroxyl to form C(l)-|J-hydroxyl, accompanies replacement of the C(8) acetoxyl group by an alkoxyl function. These epimers with a P-hydroxyl show the C(l) chemical shift -- 69.0 ppm compared with the C(l) bearing an a-hydroxyl which exhibits - 72.0 ppm.[14,15]. For epimerization to occur in these alkaloids, both a C(l) a hydroxyl group and a C(8)-acetoxyl group are necessary. Two plausible mechanisms for the epmerization have been suggested. In the formation of 216 from delphisine (86, R ^ R 2 = Ac), a common intermediate to both the paths A and B is initiated by a synchronous fragmentation of the type described by Grob (Section, 5.4, Scheme 1) [41]. Pathway A proceeds by formation of an oxetane ring 222, which is opened by addition of water from the P-face [188] to give 223. Loss of hydroxyl from C(17) leads to 216 or 217. In path B, two epimeric intermediates 221 and 224 are equilibrated through an aldehyde (225). In refluxing methanol, the epimerization proceeds mainly by path B, with formation of 216 whereas in boiling water an increasing amount of 218 is formed by path A, as indicated by ^^O labeling studies [187].

B. S. Joshi and S. W. Pelletier

358

Path A .OMe

^OMe

% < - ^ ^ K ^ J v VOAc OMe

ROH, A

OMe

Delphisine 86 ( R \ R 2 = AC)

(&H^^^

OMe

^OMe H20

Et— - N ;

E t — -N

OMe

OMe

223

222

.G

I

-OH

^OMe

1 -fROH 2 -H"^ OMe

OMe

OMe 224

216 or 217

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

359

Path B OMe

Delphisine 86 (R^R2 = Ac)

Et —

224

-*- 221

216 or 217 1 1 . BIOLOGICAL ACTIVITY OF NORDITERPENOID ALKALOIDS 11.1 Toxicity of Larkspur The toxicology and pharmacology of diterpenoid alkaloids was reviewed in 1983 by Benn and Jacyno [2]. In that review the ethnopharmacology of Aconitum, Delphinium and Garraya species and the pharmacology of individual alkaloids has been discussed. The plants commonly known as larkspurs {Delphinium species) constitute the single greatest source of livestock losses attributable to poisonous plants [189]. A study conducted by Olsen et al estimates the toxicity of individual alkaloids in larkspur responsible for acute cattle deaths on mountain and high plains range lands of the western U.S.A. [190]. In the highly toxic tall larkspur of D, barbeyi the total alkaloid content is 0.23% of dry weight of the plant and the major toxic alkaloids identified are methyllycaconitine (67) and 14-deacetylnudicauline (74, Rl = H, R2 = methylsuccinimido) [15,61,79]. They estimated that the LD50 of (67) for catde is 5.0-6.3 mg/kg body wt. or less when given intraluminally as a single dose of plant. A systematic investigation was carried out to determine the relative toxicity of leaf petiole extracts of nine plants belonging to Delphinium and Consolida species at the flowering stage of growth. D. barbeyi was found to be the most toxic plant compared with: D. glaucescens, D, geyri, D, tricorne, D. occidentale, and Aconitum columbianum showing decreasing toxicity in a mouse assay (Table 1). Table 1. Relative Toxicitv Of Leaf-Petiole Extract Among Delphinium, Consolida. And Aconitum Species At The Rowering Stage Of Growth Compared Bv Mouse Assav Sprigs

JL^sO*

Confidence Interval

D. barbeyi D.glaucescens (leaf petiole)

2.0 7.8

(1.8 - 2.3) (5.0 - 6.7)

360

B. S. Joshi and S. W. Pelletier

D. glaucescens (stem) (10.0 •• 13.2) 11.5 D. geyeri (12.2 •• 17.4) 14.6 D. hybridum cv (18.1 •• 28.9) 22.9 D. tricorne (25.2 -- 30.0) 27.6 Consolida sp, cv 35.4 (28.8 •- 43.5) D. occidentale (33.7 •- 37.8) 35.7 A. columbianum 39.2 (35.8 - 42.9) *(|il of saline extract per g body wt.) They also studied the relative toxicities of the different alkaloids present in larkspur in mice (Table 2). All the larkspur examined contained a mixture of alkaloids and the degree of poisoning in an animal is determined by the type and content of a specific alkaloid. Table 2. Relative Toxicitv Of Larkspur Alkaloids In Mice (LD^n. mg/kg i.v. [1911 Methyllycaconitine (67) Delsemine (66) Anthranoyllycoctonine (55) Condelphine[14] Delcosine (109, Rl = R2 = OH) Delsoline (71; Rl = H; R2 = Me) Lycoctonine [14] Deltaline (59; R = Me)

3 6 20 35 109 175 350 >300

Manners et al examined the structure-activity relationship of norditerpenoid alkaloids occurring in toxic larkspur (Delphinium species) in mouse bioassay [192]. This toxicological study is of interest for the development of methodology to overcome the toxicity of these compounds and thereby alleviate the poisoning threat to cattle on grazing land in the Western USA. Fourteen alkaloids of the lycoctonine-type present in toxic larkspurs were examined. Of these, nudicauline (74; R^ = Ac, R^ = methylsuccinimido) [15], 14-deacetylnudicauline (74; R^ = H, R2 = methylsuccinimido)[15], methyllycaconitine 67 [14] and elatine 82 [14] were found to be highly toxic (Table 3). Table 3. Toxicitv Of Norditerpenoid Alkaloids In Mice Alk^QJ^

No. of mice LD50 (mg/kg) (Calculated)

Nudicauline (74; R^ = Ac,

LD50 (mg/kg) (Estimated)

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

R 2 = methylsuccinimido)

23

2.7

18

4.0

361

14-Deacetylnudicauline (74; R1 = H,

R 2 = methylsuccinimido) Methyllycaconitine (67)

15

7.5

Delavaine A + B (66)

23

3.3

Elatine (82)

15

9.2

Anthranoyllycoctonine (55)

19

20.8

Barbinine (60)

6

57

N-Desethylmethyllycaconitine (67, NH)

6

100

Deltaline (59, R = Me)

23

Anhweidelphinine [ 14]

1

N-Desethyldeltaline [ 192]

12

200.5 >177 210 >230

Zaliline [ 193]

1

Dictyocarpine (59, R = H)

20

282.9

Lycoctonine [ 14]

17

443.5

Correlation of structural character with observed toxicity indicated that three structural features are necessary to enhance the toxicity of the lycoctonine-type norditerpenoid alkaloids to mammals: a) an N-ethylbicyclo substituted tertiary nitrogen atom b) an ortho-imide substituted C(18) benzoate ester, and c) a C (14) hydroxyl or derivatized C(14) hydroxyl function. The 10400 fold higher toxicity of the N-(methylsuccinimido)-anthranoyllycoctonine-type alkaloids compared with desalkyl and non-anthtranilic ester alkaloids supports this structure-activity relationship. Clinical observations of larkspur toxicosis for cattle grazing the larkspur rich in methyllycaconitine (67), indicate a neuromuscular site of action. This finding was confirmed in a neuromuscular in vitro test system which showed that 67 was highly toxic having a curare-like action [194]. The alkaloid 67 is a potent competitor of ct-bungarotoxin binding to acetycholine receptors [195]. It is considered to be the most potent competitive antagonist of these receptor sites [196]. 11.2

Effect of Norditerpenoid Alkaloids on Cardiac Sympathetic Efferent and

Vagal Afferent Nerve Activity

Digitalis and some other alkaloids activate or sensitize both arterial baroreceptors and cardiac mechanoreceptors causing a) an increase in vagal efferent nervous activity and b) a withdrawal of sympathetic efferent nervous activity resulting in a decrease in heart rate and blood pressure. Non-polar compounds like digoxin, act upon both arterial and cardiac reflex receptors [197]. Some norditerpenoid alkaloids from Aconitum and Delphinium species exhibit hypotensive and bradycardiac activity which may be due to activation of autonomic reflexes. Lappaconitine reduces blood pressure and heart rate and is comparatively less toxic [2]. Caldwell and coworkers carried out a study on lappaconitine (99) and N-deacetyllappaconitine to

362

B. S. Joshi and S. W. Pelletier

determine the activation of autonomic reflex receptors [198]. Lappaconitine (99) at a dose of 150 |ig/kg (i.v.) increased cardiac vagal afferent nerve activity (16.2%) and reduced cardiac sympathetic efferent nerve activity (12.5%). At the same dose, N-deacetyllappaconitine increased cardiac vagal afferent nerve activity (40.0%) and reduced cardiac sympathetic efferent nerve activity (23.5%). Both the alkaloids reduced heart rate and blood pressure in the dog. The data show that the polar alkaloid A^-deacetyllappaconitine has a much stronger effect on cardiac vagal afferent nerve activity than the less polar lappaconitine (99). 11.3 6-Benzoylheteratisine (134) The antiarrhythmic activity of 6-benzoylheteratisine (134), an alkaloid from Aconitum tanguticum (Maxim.) Stapf was investigated in left and right guinea pig isolated atria [199]. At concentrations of more than 6x10 "^ mol/1, preincubation with the alkaloid (134) suppressed arrhythmia induced by aconitine (91), veratridine and auabain. Bradycardia of the right atria as a sign of toxicity occurred at 1x10"^ mol/1. The alkaloid significantly reduced the maximum rate of rise of the action potential as well as the action potential amplitude, indicating inhibition of voltage-dependent sodium channels as a functional principle. In addition, a use-dependent mode of action could be demonstrated. One could conclude that 6-benzoylheteratisine (134), is a naturally occurring class-1 antiarrhythmic substance. This is the major alkaloid of Aconitum tanguticum, a plant used in the preparation of a poison antidote in Tibetan and Chinese folk medicine. REFERENCES 1. G Cordell, Introduction to Alkaloids: A Biogenetic Approach, Wiley, New York, (1981). 2. MH Benn and JC Jacyno, The Toxicology and Pharmacology of Diterpenoid Alkaloids,in: "Alkaloids: Chemical and Biological Perspectives", Vol. 1. Ed. SW Pelletier, Chapter 4, Wiley-Interscience, New York, pp 153-210 (1983); JM Jacyno, The Chemistry and Toxicology of the Diterpenoid Alkaloids, in "Chemistry and Toxicology of Diverse Classes of Alkaloids" Ed. MS Blum, Chapter 5, Alaken, Inc., Fort Colhns, Co, pp 301-336 (1996). 3. M Przybylska and L Marion, Can. J. Chem., 34, 185 (1956). 4. SW Pelletier, BS Joshi, and HK Desai, Techniques for Isolation of Alkaloids, in: ''Advances in Medicinal Plant Research" Ed. AJ Vlietinick, and RA Dommisse, Wissenschaftliche Veriagsgesellschaft mbH, Stuttgart, pp. 153-195 (1985). 5. SW Pelletier, HP Chokshi, and HK Desai, /. Nat. Prod,, 49, 892 (1986). 6. HK Desai, BS Joshi, AM Panu, and SW Pelletier, 7. Chromatogr,, 322, 223 (1985). 7. HK Desai, ER Trumbull, and SW Pelletier, /. Chromatogr., 366, 439 (1986). 8. P Kulanthaivel and SW Pelletier, J. Chromatogr,, 402, 366 (1987). 9. BS Joshi, MS Puar, Y Bai, AM Panu, and SW Pelletier, Tetrahedron, 50, 12283 (1994).

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

363

10. V Venkateswarlu, SK Srivastava, BS Joshi, HK Desai, and SW Pelletier, J. Nat. Prod., 58, 1527 (1995). 11. Y Tsuda and L Marion, Can. J. Chem., 41, 1634 (1963). 12. AJ Jones and MH Benn, Tetrahedron Lett., 42, 4351 (1972). 13. AJJonesandMHBenn, Can./. C/z^m., 51, 486(1973). 14. SW Pelletier, NV Mody, BS Joshi, and LC Schramm, ^^c and Proton NMR Spectral Assignments and Physical Constants of Ci9-Diterpenoid Alkaloids, in: "Alkaloids: Chemical and Biological Perspectives", Ed. SW Pelletier, Vol. 2, John Wiley & Sons, New York, Chapter 5, pp. 206-247 (1984). 15. SW Pelletier and BS Joshi, Carbon-13 and Proton NMR Shift Assignments and Physical Constants in: "Alkaloids: Chemical and Biological Perspectives", Ed. SW. Pelletier, Vol. 7, Springer Verlag, New York, Chapter 3, pp. 298-565 (1984). 16. WS Chen, S Sepulveda-Boza, M Mortter, and E Breitmaier, Liehigs Ann. Chem., 1297 (1985). 17. BS Joshi, JK Wunderlich, and SW Pelletier, Can. J. Chem., 65, 99 (1987). 18. JK Snyder, X Zhang, BS Joshi, and SW Pelletier, Magn. Reson. Chem., 27, 1057 (1989). 19. BS Joshi, SW Pelletier, X Zhang, and JK Snyder, Tetrahedron, 47, 4299 (1991). 20. BS Joshi, MS Puar, HK Desai, SA Ross, J Lu, and SW Pelletier, Tetrahedron Lett., 34, 1441 (1993). 21. X Zhang, JK Snyder, BS Joshi, JA. Glinski, and SW Pelletier, Heterocycles, 31,1879 (1990). 22. RN Chopra, SL Nayar, and IC Chopra, "Glossary of Indian Medicinal Plants," CSIR, New Delhi, (1956). 23. TA Henry and TM Sharp, / . Chem. Soc, 1105 (1928). 24. CRA Wright and AP Luff, /. Chem. Soc, 31, 143 (1877); 33, 151 (1878). 25. Y Tsuda and L Marion, Can. J. Chem., 41, 1485 (1963). 26. KS Khetwal, BS Joshi, HK Desai, and SW Pelletier, Heterocycles, 34, 441 (1992). 27. KS Khetwal, HK Desai, BS Joshi, and SW Pelletier, Heterocycles, 38, 833 (1994). 28. WC Muenscher, "Poisonous Plants of the United States", Revised Edition, Macmillan Co., New York, p. 79 (1951). 29. PT Millar, Wyoming Agr. Exptl. Stn. Rep., 131 (1920), [Chem. Abs., 16, 3500 (1922)]. 30. OA Beath J. Am. Pharm. Assn., 15, 265 (1926); [Chem. Abs, 20, 3778 (1926)]. 31. V Boido, OE Edwards, KL Handa, RJ. Kolt, and KK Purushothaman, Can. J. Chem., 62, 778 (1984). 32. SW Pelletier, S K Srivastava, BS Joshi, and JD Olsen, Heterocycles, 23, 331 (1985). 33. GS Singh, GS Bajwa, and MG Singh, Indian J. Chem., 4, 39 (1966). 34. SW Pelletier, NV Mody, and HS Puri, /. Chem. Soc, (Chem. Comm.), 12 (1977). 35. SW Pelletier, NV Mody, and HS Puri, Phytochemistry, 16, 623 (1977).

364

36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46.

47. 48. 49. 50. 51. 52. 53. 54. 55. 56. 57. 58. 59.

60. 61. 62.

B- S. Joshi and S. W. Pelletier

HK Desai, BS Joshi, and SW Pelletier, Heterocycles, 24, 1061 (1986). HK Desai and SW Pelletier, Heterocycles, 29, 225 (1989). WS Chen and E Breitmaier, Chem, Ber., 114, 394 (1981). SW Pelletier, SY Chen, BS Joshi, and HK Desai, J. Nat. Prod., 47, 474 (1984). SW Pelletier, BS Joshi, JA Glinski, HP Chokshi, SY Chen, K Bhandary, and KT Go, Heterocycles, 25, 365 (1987). CA Grob, HR Kiefer, H Lutz, and H Wilkens, Tetrahedron Lett., 2901 (1964). HV Rosendahl, Arch. Pharmakol. Inst. Dorpat, 11, 1 (1895). SA Ross, SW Pelletier, and AJ Aasen, Tetrahedron, 48, 1183 (1992). HM Sayed, HK Desai, SA Ross, SW Pelletier, and AJ Aasen, J. Nat. Prod., SS, 1595 (1992). R Brandes, Schweigger's J. Chem. Phys., 25, 369 (1819); Lassaigne and Feneulle, Ann. Chim. Phys., 12, 358 (1819). SW Pelletier and NV Mody, The Chemistry of C20-Diterpenoid Alkaloids, in: "The Alkaloids, Chemistry and Physiology" Ed. RGA Rodrigo, Vol. XVlll, The Academic Press Inc., New York, pp. 144 (1981). SW Pelletier and MM Badawi, Heterocycles, 23, 2873 (1985). SA Ross, HK Desai, and SW Pelletier, Heterocycles, 26, 2895 (1987). SA Ross and SW Pelletier, J. Nat. Prod., 51, 572 (1988). SW Pelletier and MM Badawi, J. Nat. Prod., 50, 381 (1987). SW Pelletier, SA Ross, and JT Etse, Heterocycles, 27, 2467 (1988). X Liang, HK Desai, and SW Pelletier, /. Nat. Prod., 53, 1307 (1990). FP Wang and SW Pelletier, J. Nat. Prod., 50, 55 (1987). HK Desai, RH El Sofany, and SW Pelletier, J. Nat. Prod., 53, 1606 (1990). SW Pelletier, S Bhandaru, HK Desai, SA Ross, and HM Sayed, J. Nat. Prod., 55, 736 (1992); P Kulanthaivel, HK Desai, and SW Pelletier, J. Nat. Prod., 52, 143 (1989). J Lu, HK Desai, SA Ross, HM Sayed, and SW Pelletier, J. Nat. Prod., 56, 2098 (1993). HK Desai, BT Cartwright, and SW Pelletier, J. Nat. Prod., 57, 677 (1994). X Liang, SA Ross, YR Sohni, HM Sayed, HK Desai, BS Joshi, and SW Pelletier, J. Nat. Prod., 54, 1283 (1991). WB Cook and OA Beath, /. Amer. Chem. Soc, 74, 1411 (1952); M Carmack, JP Ferris, J Harvey, Jr., PL Magat, E Martin, and DW Mayo, J. Amer. Chem. Soc, 80, 497 (1958). BS Joshi, EA El-Kashoury, HK Desai, EM Holt, JD Olsen, and SW Pelletier, Tetrahedron Lett., 29, 2397 (1988). P Kulanthaivel, SW Pelletier, and JD Olsen, Heterocycles, 27, 339 (1988). SW Pelletier, P Kulanthaivel, and JD Olsen, Phytochemistry, 28,1521 (1989).

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

365

63. GA Miana, MI Khan, F Sultana, and M Ikram, Pak. J. Sci. and Ind. Res,, 23, 2483 (1985). 64. HK Desai, BS Joshi, and SW Pelletier, Heterocycles, 23, 2483 (1985). 65. S Sakai, N Shinma, H Hasegawa, and T Okamoto, / . Pharm. Soc, Japan, 98,1376 (1978). 66. S Sakai, N Shinma, and T Okamoto, Heterocycles, 8, 207 (1977) 67. GA Mair and M Przybylska, unpublished results quoted by L Marion, in: "Pure and applied Chemistry", 6, 621 (1963) 68. KK Bhandary, N Ramasubbu, BS Joshi, HK Desai, and SW Pelletier, Acta CrySt., C46, 1704 (1990). 69. SW Pelletier, HM Harraz, MM Badawi, S Tantiraksachai, F P Wang, and SY Chen, Heterocycles, 24, 1853 (1986). 70. SW Pelletier, RS Sawahney, HK Desai, and NV Mody, / . Nat. Prod., 43, 395 (1980). 71. SW Pelletier, JA Glinski, BS Joshi, and SY Chen, Heterocycles, 20, 347 (1983). 72. JA Glinski, BS Joshi, SY Chen, and SW Pelletier, Tetrahedron Lett., 25, 1211 (1984). 73. JA Maddry, MG Newton, and SW Pelletier, J. Nat. Prod., 49, 674 (1986). 74. BS Joshi, JA Glinski, HP Chokshi, SY Chen, SK Srivastava, and SW Pelletier, Heterocycles, 22, 2037 (1984). 75. PW Codding, KA Kerr, MH Benn, AJ Jones, SW Pelletier, and NV Mody, Tetrahedron Lett., 21, 127 (1980). 76. SA Ross, HK Desai, BS Joshi, SK Srivastava, JA Glinski, SY Chen, and SW Pelletier, Phytochemistry, 27, 3719 (1988). 77. BS Joshi, HK Desai, SW Pelletier, JK Snyder, X Zhang, and SY Chen, Phytochemistry, 29, 357 (1990). 78. WF Reynolds, DW Hughes, M Perpick-Dumont, and RG Enriguez, J. Magn. Reson., 64, 304 (1985); M Perpick-Dumont, WF Reynolds, and RG Enriguez, Magn. Reson. Chem., 26, 358 (1988). 79. SW Pelletier, AM Panu, P Kulanthaivel, and JD Olsen, Heterocycles, 11, 2387 (1988). 80. SW Pelletier, P Kulanthaivel, and JD Olsen, Heterocycles, 28,107 (1989). 81. SW Pelletier, SA Ross, and P Kulanthaivel, Tetrahedron, 45,1892 (1989). 82. SW Pelletier, SA Ross, and HK Desai, Phytochemistry, 29, 2381 (1990). 83. JC Park, HK Desai, and SW Pelletier, / . Nat. Prod., 58, 291 (1995). 84. K Wada, T Yamamoto, H Bando, and N Kawahara, Phytochemistry, 31, 2135 (1992). 85. H. Bando, K. Wada, J. Tanaka, S. Kimura, E Hasegawa, and T Amiya, Heterocycles, 29, 1293 (1989). 86. MS Rabinovich, /. Gen. Chem. USSR (English Translation) 24, 2211, 2242 (1954); AD Kuzovkov ibid, 399 (1955); AD Kuzokov and JF Platonova, ibid, 29, 2746 (1959). 87. SW Pelletier, HK Desai, P Kulanthaivel, and BS Joshi, Heterocycles, 26,2835 (1987). 88. W Deng and WL Sung, Heterocycles, 24, 873 (1986).

366

B- S. Joshi and S. W. Pelletier

89. X Liang, HK Desai, BS Joshi, and SW Pelletier, Heterocycles, 31, 1889 (1990). 90. BS Joshi, SK Srivastava, AD Barber, HK Desai, and SW Pelletier, /. Nat. Prod., 60, 439 (1997). 91. IS Blagbrough, DJ Hardick, S Wonnacott, and BVL Potter, Tetrahedron Lett., 35,3367 (1994). 92. M Caraiack, JP Ferris, J Harvey Jr., PL Magat, EW Martin, and DW Mayo, /. Amer. Chem. Soc, 80, 947 (1958). 93. P Kulanthaivel and SW Pelletier, Tetrahedron Utt., 28,3883 (1987); P Kulanthaivel and SW Pelletier, Tetrahedron , 44,4313 (1988). 94. SK Srivastava, BS Joshi, MG Newton, D Lee, and SW Pelletier, Tetrahedron Lett., 36, 519 (1995). 95. CA Grob and W Scwarz, Helv. Chem. Acta, 47, 1870 (1964); CA Grob, AngeM;. Chem., International Edn., 6 (1967), 8, 535 (1969). 96. W Klyne and V Prelog, Experientia, 16, 521.(1960) 97. G Wittig and HD Frommeld, Chem. Ber., 97, 3548 (1964); MF Bartlett, DF Dickel, WI Taylor, J. Amer. Chem. Soc, 80, 126 (1958). 98. ME Wolff, Chem. Rev., 63, 55 (1963). 99. R Aneja and SW Pelletier, Tetrahedron Lett., 669 (1964). 100. M Przybylska, Can. J. Chem., 41, 2911 (1963). 101. R Aneja, M Locke, and SW Pelletier, Tetrahedron , 29, 3297 (1973). 102. QP Jiang and SW Pelletier, Tetrahedron Lett., 29,1875 (1988). 103. HK Desai and SW Pelletier, /. Nat. Prod., 56, 2193 (1993). 104. BS Joshi, Y Bai, DH Chen, and SW Pelletier, Tetrahedron Utt., 34, 7525 (1993). 105. SW Pelletier and R Aneja, Tetrahedron Lett., 557 (1967); MS Yunusov, YV Rashkes, VA Telnov, and SY Yunusov, Khim. Prir. Soedin., 5, 575 (1969). 106. SW Pelletier and NV Mody, Tetrahedron, 34,2421 (1978); SW Pelletier, RS Sawahney, and NV Mody, Heterocycles, 9,1241 (1978). 107. SW Pelletier, NV Mody, Z Djarmati, and SD Lajsic, /. Org. Chem., 41, 3042 (1978); SW Pelletier, Z Djarmati, and NV Mody, Tetrahedron Lett., 1749 (1976). 108. SW Pelletier, NV Mody, J Finer-Moore, HK Desai, and HS Puri, Tetrahedron Lett., 22, 313 (1981). 109. A Katz and E Staehelin, Helv. Chim. Acta, 65, 286 (1982). 110. SW Pelletier, BS Joshi, HK Desai, A Panu, and A Katz, Heterocycles, 24, 1275 (1986). 111. G Goto, K Sakai, N Sakabe, and Y Hirata, Tetrahedron Utt., 1369 (1968); K Sakai, N Sakabe, and Y Hirata, J. Chem. Soc, Perkin I, 354, (1971). 112. N Singh and A Singh, J. Indian Chem. Soc, 42, 49 (1965). 113. SW Pelletier, KN Iyer, JH Wright, MG Newton, and N Singh, /. Amer. Chem. Soc, 93, 5942 (1971). 114. N. Singh and SS Jaswal Tetrahedron Lett., 2219 (1968).

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

367

115. N Singh SS Jaswal, and A Singh, Indian J, Chem , 12, 1219 (1974). 116. N Singh and A Singh, Indian J. Chem., 10, 953 (1962). 117. SW Pelletier and NV Mody, The Chemistry of C20-Diterpenoid Alkaloids, in: "The Alkaloids, Chemistry and Physiology" Vol. XVIII. Ed. RHF Manske and RGA Rodrigo, Chapter 2, Academic Press Inc., New York, pp 125,131 (1981). 118. MH Benn, JF Richardson, and W Majak. Heterocycles, 24, 1605 (1986). 119. JA Glinski, BS Joshi, QP Jiang, and SW Pelletier, Heterocycles, 27,185 (1988). 120. RT Alpin MH Benn, SW Pelletier, J Solo, SA Telang, and H Wright, Can, J. Chem,, 46, 2635 (1968). 121. YL Chu and JH Chu, Heterocycles, 17, 6007 (1982). 122. KI Varughese and SW Pelletier, / . Nat. Prod., 47,470 (1984). 123. MG Reinecke, WH Watson D Chen, and W Van, Heterocycles, 24,49 (1986). 124. F Sun, XT Liang, and DQ Yu, / . Nat. Prod., 50, 923 (1987). 125 BS Joshi, JA Glinski, KI Varughese, and SW Pelletier, Heterocycles, 27,195 (1988). 126. Y Bai, F Sun, M Benn, and W Majak, Phytochemistry, 37, 1717 (1994). 127. M Reina, A Madinaveitia, G de la Fuente, ML Rodriguez, and I Brito Tetrahedron Lett., 33, 1661 (1992). 128. QP Jiang and SW Pelletier, J. Nat. Prod., 54, 525 (1991). 129. L Marion, L Fonzes, CK Wilkins Jr., JP Boca, F Sandberg, R Thorsen, and E Linden, Can. J. Chem., 45, 969 (1967). 130. S Usmanova, V Telnov, MS Yunusov, N AbduUaev, A Shreter, and G Filippova, Khim. Prir. Soedin., 879 (1987). 131. V Telnov, MS Yunusov, N AbduUaev, and M Zhamierashvili, Khim. Prir. Soedin., 556 (1988). 132. E Sirotenko and Y. Rashkes, Khim. Prir. Soedin., 532 (1989). 133. SW Pelletier, R Sawhney, and A Aasen, Heterocycles, 12, 377 (1979). 134. SW Pelletier, NV Mody, and R Sawhney, Can. J. Chem., SI, 1652 (1992). 135. BS Joshi, HK Desai, SW Pelletier, EM Holt, and AJ Aasen, J. Nat. Prod., 51, 265 (1988). 136. SA Ross, BS Joshi, SW Pelletier, MG Newton, and AJ Aasen, J. Nat. Prod., 56,424 (1993). 137. BS Joshi, HM Sayed, SA Ross, HK Desai, SW Pelletier, P Cai, JK Snyder, and A Aasen, Can. J. Chem., 11, 100 (1994). 138. SW Pelletier, NV Mody, HK Desai, J Finer-Moore, J Nowacki, and BS Joshi, J. Org. Chem., 4S, 1787(1983). 139. DH Chen and WL Sung, Zhongcaoyao , 16, 388 (1985); [Chem. Abstr., 104, 95320g, (1986)}. 140. BS Joshi, DH Chen, X. Zhang, JK Snyder, and SW Pelletier, Heterocycles, 32, 1793 (1991).

368

B. S. Joshi and S. W. Pelletier

141. A Bax, /. Magn. Reson,, 57, 314 (1984); WF Reynolds, S McLean, M Perpick-Dumont and E Enriquez, Magn. Reson. Chem., 27, 162 (1989). 142. WE Doering, and M Farber, J. Amer, Chem., Soc, 71, 1514 (1949); HL Goering, and MF Sloan, J. Amer. Chem. Soc, 83, 1397 (1961). 143. HK Desai, BS Joshi, SW Pelletier, B Sener, F Bingol, and T Baykal, Heterocycles, 36, 1081, (1993). 144. SW Pelletier and LH Keith, Diterpenoid Alkaloids from Aconitum Delphinium and Garraya Species: The Ci9-Diterpene Alkaloids, in :"The Alkaloids, Chemistry and Physiology", Chapter 1, Vol XII, Ed. RHF Manske, Academic Press, New York, (1970). 145. RJ Sundberg and KG Gadamsetti, Tetrahedron, 47,5673 (1991); M Rubiralta, A Diez, A Balet, and J Bosch, Tetrahedron, 43, 3021, (1987); P Magnus, NL Sear, CS Kim and N Vicker, / . Org. Chem., 57, 70 (1992). 146. OE Edwards and T Singh, Can. J. Chem., 32, 465 (1954). 147. SW Pelletier and WA Jacobs, /. Amer. Chem. Soc, 78, 4139 (1956). 148. SW Pelletier and JT Etse, J. Nat. Prod., 52, 145 (1989). 149. KB Bimbaum, K Wiesner, EWK Jay, and L Jay, Tetrahedron Lett., 867 (1971); SW Pelletier and Z Djarmati, /. Amer. Chem. Soc, 98, 2626 (1976). 150. SW Pelletier, WH De Camp, S Lajsic, Z Djarmati, and AH Kapadi, J. Amer. Chem. Soc, 96,7815 (1974); SW Pelletier, Z Djarmati, S Lajsic, and W De Camp, J. Amer. Chem. Soc, 98, 2617 (1976). 151. HC Wang, DZ Zhu, Y Zhao, and RH Zhu, Acta Chimica Sinica, 38, 475 (1980); H Takayama, A Tokita, M Ito, S Sakai, F Kurosaki, and T Okamoto, /. Pharm. Soc. Japan, 102, 245 (1982); G de la Fuente, RD Acosta, and T Orribo, Heterocycles, 29, 205 (1989). 152. AH Jackson, and JA Martin, /. Chem. Soc (C), 2222 (1966). 153. AJ Marsaioli, FAM Reis, AF Magalehaes, EA Ruveda, and AM Kuck, Phytochemistry, 18, 165 (1979). 154. E. Wenkert, BZ Buckwalter, IR Burfitt, MJ Gasic, HG Gottiieb, EW Hagaman, FM Schell and PM Wovkulich, 'Topics in Carbon-13 NMR Spectroscopy" Ed. GC Levy, Vol 2, Wiley-Interscience, New York (1976). 155. H Takayama, M Ito, S Sakai, and T Okamoto, Heterocycles, 65,403 (1981). 156. S Sakai, K Yamaguchi, I Yamamoto, K Hotoda, T Okazaki, N Aimi, J Haginiwa, and T Okamoto, Chem. Pharm. Bull. Japan, 31, 3338 (1983). 157. H Bando, K Wada, T Amiya, K Kobayashi, Y Fujimoto, and T Sakurai, Heterocycles, 26, 2623 (1987). 158. Z Karimov and MG Zhamierashvili, Khim. Prior. Soedin, 335 (1981). 159. Y Bai, Ph. D. Thesis, University of Calgary, (1993). 160. A Ulubelen, AH Meri9li, F Merigli, R Ilarsan, and W Voelter, Phytochemistry, 34, 1165 (1993).

Recent Developments in the Chemistry of Norditerpenoid and Diterpenoid Alkaloids

369

161. SA Ross, BS Joshi, HK Desai, SW Pelletier, MG Newton, X Zhang, and JK Snyder, Tetrahedron, 47, 9585 (1991). 162. K Wada, H Bando, and T Amiya, Heterocycles, 23, 2623 (1985); H Bando, K Wada, T Amiya, Y Fujimoto, and K Kobayashi, Chem. Pharm. Bull. Japan, 36, 1604 (1988). 163. G de la Fuente, M Meina, E Valencia, and A Rodriguez-Ojeda, Heterocycles, 27,1109 (1988). 164. SJ Chen, SH Li, and XJ Xao, Acta Botanica Sinica, 28, 86 (1986). 165. BT Salimov, ND AbduUaev, MS Yunusov, and SYu Yunusov, Khim, Prir. Soedin, 235 (1978); B Tashkhodzhaev, Khim. Prir. Soedin, 230 (1982). 166. W Deng and WL Sung, Heterocycles, 24, 869 (1986). 167. F Sun, XT Liang, and DQ Yu, / . Nat. Prod., 51, 50 (1988). 168. LH Briggs and RW Cawley, / . Chem. Soc, 1888 (1948); SW Pelletier, JA Glinski, KI Varughese, J Maddry, and NV Mody, Heterocycles, 20, 413 (1983). 169. P Kulanthaivel, EM Holt, JD Olsen, and SW Pelletier, Phytochemistry, 29,293 (1990). 170. GD Manners, RY Young, M Benson, MH Ralphs, and JA Pfister, Phytochemistry, 42, 875 (1996). 171. A Ulubelen, HK Desai, SK Srivastava, BP Hart, JC Park, BS Joshi, SW Pelletier, AH Merigli, and F Merigli, J. Nat. Prod., 59, 360 (1996). 172. G de la Fuente, G Gavin, R Diaz-Acosta, and JA Morales, Heterocycles, 27,1 (1994). 173. G de la Fuente, L Ruiz-Mesia, and ML Rodriguez, Helv. Chim. Acta, 11, 1768 (1994). 174. SI Sakai, Y. Yamamoto, K Yamaguchi, H Takayama, M Ito, and T Okamoto, Chem. Pharm. Bull. Japan, 30, 4579 (1982). 175. H Takayama, JJ Sun, N Aimi, and S Sakai, Tetrahedron Lett., 30, 3441 (1989). 176. ZG Chen, AN Lao, HC Wang, and SH Hong, Heterocycles, 26, 1455 (1987). 177. K Wada, H Bando, T Amiya, and N Kawahara, Heterocycles, 29, 2141 (1989). 178. H Takayama, A Tokita, M Ito, SI Sakai, F Kurosaki, and T Okamoto, Yakugaku Zasshi, 102, 245 (1982). 179. SW Pelletier, NV Mody, J Finer-Moore, AMM Ateya, and LC Schramm, J. Chem. Soc, Chem. Comm., 327 (1981). 180. QP Jiang, JA Glinski, BS Joshi, JA Maddry, MG Newton, and SW Pelletier, Heterocycles, 27, 925 (1988). 181. MC Wildman, and DT Bailey, / . Org. Chem., 33, 37849 (1968); CF Murphy and WC Wildman, Tetrahedron Lett., 3863 (1964). 182. SW Pelletier, HK Desai, and NV Mody, Heterocycles, 13, 277 (1979). 183. HK Desai, QP Jiang, and SW Pelletier, J. Nat. Prod., 53, 1374 (1990). 184. SW Pelletier, NV Mody, HK Desai, J Finer-Moore, J Novaki, and BS Joshi, J. Org. Chem., 48, 1787 (1983). 185. HK Desai, Y Bai, and SW Pelletier, / . Nat. Prod., 60, 684 (1997). 186. HK Desai, BS Joshi, SA Ross, and SW Pelletier, / . Nat. Prod., 52, 720 (1985).

370

B. S. Joshi and S. W. Pelletier

187. SW Pelletier, HK Desai, QP Jiang, and SA Ross, Phytochemistry, 29, 3649 (1990). 188. S Searles in "Heterocyclic Compounds with three- and four-memberedRings" Part 2, Ed., A. Weissberger, pp 994, 998. 10054, John Wiley (1964). 189. JD Olsen, Larkspur toxicosis: A review of current research in "Effects of Poisonous Plants on Live stock" Ed. RF Keeler, KR Van Kamper, and LF James, Academic Press Inc., New York, pp 535-543 (1978); JD Olsen, Larkspur poisoning: Potential for management, in "Proceedings of the Second Meadow Symposium" Ed., EG Seimer, and RH Delaney, Colorado State University Experimental Station, Special series, 34, 6772 (1984). 190. JD Olsen, GD Manners, and SW Pelletier, Collectanea Botanica (Barcelona), 19,142 (1990). 191. JD Olsen and GD Manners, Toxicology of diterpenoid alkaloids in rangeland larkspurs (Delphinium Species), in "Toxicants of Plant Origin", Vol. I, Ed. P Cheeke, CRC Press, Inc., Boca Raton, Florida, pp. 291-326 (1989). 192. GD Manners, KE Panter, and SW Pelletier, /. Nat. prod., 58, 863 (1995). 193. F Sun and M Benn, Phytochemistry, 31, 3247 (1992). 194. VN Aiyar, MH Benn, T Hanna, J Jacyno, S Roth, and JL Wilkes, Experientia, 35, 1367 (1979). 195. KR Jennings, DG Brown, and DP Wright Jr., Experientia, 35, 611 (1986). 196. S Wonnacott, EX Albuquerque, and D Bertrand, Methods in Neurosciences, 12, 263 (1993). 197. RW Caldwell, and WR Rebagay, J. Pharmacol. Exp. Ther., 262, 1022 (1992). 198. H Chiao, SW Pelletier, HK Desai, WR Rebagay and RW Caldwell, European J. Pharmacol., 283, 103 (1995). 199. JF Heubach, F Heer, D Wilhelm, and D Peters, Phytomedicine, 4, 109 (1997).

Chapter Five

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids by Means of Transition Metal Catalyzed Carbonylations Iwao Ojima and Donna M. lula Department of Chemistry State University of New York at Stony Brook Stony Brook. NY 11794

CONTENTS 1. INTRODUCTION

372

2. SYNTHESES OF PIPERIDINE ALKALOIDS VIA TRANSITION METAL CATALYZED HYDROCARBONYLATIONS

373

2.1 Piperidine Alkaloids 2.2 Methodology for the Formation of Piperidine Ring System by Means of Catalytic Carbonylations and its Functionalization 2.3 Syntheses of 2,6-Disubstituted Piperidines via Cyclohydrocarbonylation 2.3.1 Cobalt-Catalyzed Processes 2.3.2 Rhodium-Catalyzed Processes 2.4 Syntheses of Tri- and Tetrasubstituted Piperidine Alkaloids via Cyclohydrocarbonylation 2.5 Syntheses of Piperidine Alkaloid Skeletons via Cyclohydrocarbonylation of Aminodienes 3. HYDROCARBONYLATION ROUTES TO IZIDINE ALKALOIDS 3.1 Izidine Alkaloids 3.2 General Strategies for the Construction of Izidine Alkaloid Ring Systems via Carbonylations 3.3 Syntheses of Pyrrolizidine Alkaloids 3.3.1 (±)-Isoretronecanol and (±)-Trachelanthamidine

373 373 384 385 386 390 392 395 395 395 400 400 371

I. Ojima and D. M. lula

372

3.3.2 Ant Venom (3S,5R,8S)-3-Heptyl-5-Methylpyrrolizidine 3.4 Syntheses of Indolizidine Alkaloids (±)-167B and (+)-209D 3.5 Synthesis of Functionalized Quinolizidine Alkaloids

400 403 404

4. SYNTHESES OF Q U I N A Z O L I N E A L K A L O I D SKELETONS T H R O U G H CARBONYLATIONS

406

5. C O N C L U S I O N

409

6. A C K N O W L E D G E M E N T S

410

7. R E F E R E N C E S

410

1.

INTRODUCTION Piperidine, pyrrolizidine, indolizidine, quinolizidine, and quinazoline alkaloids are found

in great numbers in nature. Because of their diverse biological activities, these alkaloids have attracted the attention of synthetic, medicinal, pharmaceutical, and organic chemists. Numerous methodologies for constructing piperidine and izidine alkaloids have been developed over the years. In this regard, it can be said that these alkaloids, as synthetic targets, have contributed to the growth and development of modem organic syntheses. Although there are many synthetic strategies in the literature, this chapter will focus on transition metal-catalyzed carbonylations as efficient and novel approaches toward the construction of alkaloid skeletons. The development of synthetic methods by means of transition metal-catalyzed reactions over the last decade or two has been explosive [1,2]. Transition metal catalysts facilitate complex organic transformations with high degrees of chemo-, regio-, and stereoselectivity. In general, carbonylations are transition metal-catalyzed reactions that use carbon monoxide, involving the incorporation of a carbonyl group into a substrate [3].

For example, the

hydroformylation reaction is one of the most important industrial processes for the production of aldehydes from alkenes. Carbonylations are also used for the homologation of a variety of organic substrates, a process which is very useful in organic synthesis. Carbonylation reactions are very "tunable" in that changes in transition metal catalyst, ligands, and reaction conditions can exert substantial effects upon the chemo-, regio-, and stereoselectivity of the process. This chapter describes the recent advances in the synthesis of piperidine, izidine, and quinazoline alkaloids by means of transition metal-catalyzed carbonylations.

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

373

2. SYNTHESES OF PIPERIDINE ALKALOIDS VIA TRANSITION METAL CATALYZED HYDROCARBONYLATIONS

2.1

Piperidine Alkaloids

Piperidine alkaloids constitute one of the major classes of alkaloids and have been the subject of numerous reviews [4-7]. Piperidine itself is a naturally ocurring compound found in plants such as Piper nigrum L., Piperaceae and piperidine alkaloids are classified according to their natural source. Piperidine alkaloids can also be categorized on the basis of their structure, for example, 2,6-disubstituted piperidines, fused-ring piperidines, A^-acylpiperidines, steroidal piperidines, piperidine alcohols, etc. Examples of piperidine alkaloids are shown in Figure 1. Prosopinine (1) was isolated from the leaves, stems, and roots of Prosopis africana Taub [4], and has a wide variety of biological activities such as acting as a sedative, hypotensive agent, spasmolytic, local anestetic, antiseptic agent, etc. Piperidine 24ID (2) was isolated from the skin of poison-dart dendrobate frogs Dendrobates speciosus [8] and blocks the action of acetylcholine by a non competative blockade of the nicotinic receptor-channel complex [9]. Piperine (3), isolated from black pepper {Piper nigrum), has numerous biological activities such as the stimulation of the pituitary adrenal axis, increase in the permeability of intestinal epitheleal cells, and inhibition of dopamine p-hydroxylase [10]. Sedamine (4) was isolated from Sedum acre [11] and has been shown to competitively inhibit pea diamine oxidase [12]. Adaline (5) is a defensive alkaloid isolated from the European two-spot ladybird beetle Adalia bipunctata [13]. Histrionicotoxin (6) is one of the components of the defensive skin secretions of Dendrobates frogs, which acts as a venom as well as a mucosal tissue irritant toward mammals and reptiles [14]. This alkaloid is believed to block the nicotinic acetylcholine receptor-channel complex as well as inhibit the binding sites associated with sodium, potassium, and calcium channels in brain membranes [15].

2.2 Methodology for the Formation of Piperidine Ring System by Means of Catalytic Carbonylations and its Functionalization An efficient methodology for the construction of the piperidine alkaloid skeleton by means of catalytic carbonylations is not only useful for the synthesis of piperidine alkaloids, but also provides routes to izidine alkaloids. The use of transition metal-catalyzed carbonylation as an approach to the syntheses of functionalized piperidine skeletons has shown that N-acyl amino alkanals (7) can be converted into cyclic AT-acylamino acids 9 under cobalt-catalyzed hydroformylation conditions (equation 1) [16-18]. The reaction is believed to proceed via

1. Ojima and D. M. lula

374

hemiamidal 8. In fact, control experiments using (9-alkylhemiamidals prepared by anodic oxidation strongly support this hypothesis (equation 2) [16-18].

'//. r\

OH CH3(CH2)8

n

CH3(CH2)ri N' ._ — n -

V

Figure 1

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

375

C02(CO)8

T

CO/H2 ^CHO (150atm, 1:1)

o

75%

"N" J

"COoH

(1)

d ^ P h

C02(CO)8 OMe

N /

-

Ph 10

CO/H2 (100 atm, 1:1)

N'

A,

O

C02Me

(2)

Ph

11

This reaction is an intramolecular version of the Wakamatsu reaction which gives an Nacyl-a-amino acid from an aldehyde and amide in the presence of a catalytic amount of Co2(CO)8 at high temperature [19]. There are several possible pathways leading to the Wakamatsu reaction product as illustrated by in Scheme 1. Studies of the mechanism of the intramolecular Wakamatsu reaction indicates that coordination of the amide carbonyl to the cobalt metal center is essential for this reaction to occur [20]. This coordination likely prevents the cobalt metal from oxidative addition of H2 so that facile hydroylsis of the acyl-cobalt bond of E takes place to give A^-acyl-a-amino acid F (Pathway B). Rhodium complexes have been shown not to catalyze the Wakamatsu reaction, but promote the intramolecular amidocarbonylation of alkenylamides. For example, 3-butenamide (12) is easily converted to dihydro-2-pyridone 13 using a catalytic amount of Rh4(CO)i2 and CO/H2 (1/1, 82 atm), in 92% yield (equation 3) [21-23].

376

Ojima and D. M. lula

.OH N^^R

T

B

/Co(CO)4

HCo(CO)4

Y

- H2O

o

o

CO

ro

X

^ ' • ^ C ' / Pathway A

Co(CO)4 Pathway B

R

0

( 9 co(co)4

( el

O

D

Co(CO)3

C

I

N

o

R

HoO

^COgH HoO

R

[Co(CO)4]-

H Scheme 1

o

377

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

o NH,

H2'CO(1:1,82atm) Rh4(CO)i2 ^ THF, 100°C 92%

12

(3)

I

H 13

The first step in the transformation of 12 to dihydropyridone 13 is the extremely regioselective rhodium-catalyzed hydroformylation of the alkene moiety as shown in Scheme 2. The observed extremely high regioselectivity is acribed to the amide-directed "chelation control", which clearly favors a six-membered ring formation as compared to the formation of a sevenmembered ring.[23] Intramolecular reaction of the aldehyde moiety with the amide nitrogen gives the 6-membered ring hemiamidal 15. Dehydration of hemiamidal 15 followed by double bond isomerization results in the formation of dihydropyridone 13.

[Rh] NHg

12

CO/H2

CHO . / ^ HgN 14

HO'

'N-^0 I H 15

N

^O

Scheme 2 3-Butenamide (17) has been shown when catalyzed by RhCl(PPh3)3 and 20 equivalents of PPhs to give 3,4-dihydro-2-pyridone (18) in 92% yield accompanied by a small amount of 4methyl-3-pyiTolin-2-one (8%) (Scheme 3) [23]. A large excess of PPhs is cracial to achieve the

378

I. Ojima and D. M. lula

observed high regioselectivity. When insufficient amount of PPhs was used, a nearly 1:1 mixture of dihydropyridone (18) and methylpyrrolinone was formed, which yielded a unique heterodimer (19) as the major product. This unique heterodimer (19) was obtained in 94 % yield when the reaction was carried out in the presence of P(0Ph)3. O NH2

RhCI(PPh3)3 PPh3 / CO/H2 (3:1, 82 atmy

\

80°C 92 %

RhCI(PPh3)3 P(0Ph)3 \ C 0 / H 2 (3:1, 82 atm)

THF, 94 %

Scheme 3 Functionalization at the a position of the piperidine ring is useful because it provides a point from which further elaboration is possible. The cyclization reactions mentioned above are particularly useful because they generate the functionalized piperidine ring in one step. Another common way in which a-substituents can be introduced is the a-lithiation of A^-r-Boc-piperidine. a-Lithiation of A^-r-Boc-pyrrolidines and piperidines followed by electrophilic substitution provides a useful method for the synthesis of alkaloids [24]. Lithiation of 20, for example, followed by the addition of dimethylsulfate results in the formation of the 2,6-dialkylpiperidine 21 which was then deprotected by trifluoroacetic acid (TFA) to give (±)-solenopsin A (22) with excellent diastereoselectivity (equation 4).

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

1)s-BuLi/ TMEDA •rjl' \--;^Q 2)Me2S04 C02*Bu 83 %

379

(4) HgC^^N^t'^o cOg'Bu

20

21

HX'

Anodic oxidation followed by nucleophilic substitution also provides an efficient method for introducing an a-substituent on a piperidine ring. Carbamates can be methoxylated a to the nitrogen by anodic oxidation in methanol [25-29]. Anodic oxidation of a-ro-diamino acid derivatives such as 23 results in methoxylation a to the co-amide moiety, giving intermediate 24 which then cyclizes to the 2,6-disubstituted pipecolate 25 when treated with acid (equation 5).

NHCOgMe 1)-2e-, MeOH MeOaCHN"

^COgMe 2) H2SO4 -MeOH MeOgC^ 47%

23

N I COaMe

(5) OMe

25 NHCOgMe

OMe

COgMe

MeOaCH 24

2-Alkoxypiperidine 25 serves as a versatile intermediate in the synthesis of (+)-JVmethylconiine [10] (30) (Scheme 4) and (+)-A^-methylpseudoconhydrine [30] (33) isolated from South African Conium species (Scheme 5). The reaction of 2-alkoxypipecolate 25 with allyltrimethylsilane in the presence of TiCU results in the exclusive formation of cis-2,6-

380

I- Ojima and D. M. lula

disubstituted pipecolate 26 after catalytic hydrogenation [27]. Hydrolysis of 26 gave 6propylpipecolic acid 27 which was then subjected to anodic decarboxylation to give Omethylhemiamidal 28. Reaction of 28 with sodium borohydride (NaBH4) followed by reduction with lithium aluminum hydride (LAH) gave (+)-A^-methylconiine (30) with 96% ee. In a similar manner, 2-alkoxy-piperidine 25 has been used in the synthesis of (+)-Nmethylpseudoconhydrine (33) (Scheme 5) [28]. 2-Alkoxypiperidine 25 was converted to 6propylpipecolic acid 27 following the same steps shown in Scheme 4. Anodic oxidation was then used to stereoselectively introduce an acetoxy group at C(5) and the hydroxyl group at C(6) was reduced using NaBH4 to give 32 in 62% yield as the predominant product with the desired stereochemistry. The diastereomers of 32 were separated and (35',65)-32 was treated with LAH followed by HCl to give (+)-A^-methylpseudoconhydrine (33) with 95% ee.

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

381

^SiMeg

1) MeOgC^

I COgMe

TiCL

2) Hg/ Pd-C

COgMe

68 %

25

26 KOH

HgCd''

'•'I, N I C02Me

-2e NaOMe, -MeOH HOoC^'

I COgMe

28

27

NaBH.

LAH ''/.

N I Me

•'I. N I COgMe

30

29 Scheme 4

I. Ojima and D. M. lula

382

^SiMeq

1) MeOgC"^'

N

TiCL

OMe

COgMe 25

2) Hg/ Pd-C

C02Me 26 1)K0H 2) H3O*

AcQ

1)-4e/AcOH 2) H2O-ACOH HO^C^^' ^ N ' I COgMe

66%

31

COaMe 27

NaBH4 HCO2H

AcQ 1)LAH I COgMe

2)HCI

32

- "TD, N I CH3 33

(3S, 6S) 62 % (3fl, 6 ^ 7 % Scheme 5

""

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

383

(±) Sedamine (4) and (-) allosedamine (37) have been synthesized in two steps from a methoxy piperidine 35 (equation 6) [25]. a-Methoxypiperidine 35 was prepared via anodic oxidation of A/'-methylcarbamate 36. Treatment of 35 with TiCU followed by a(trimethylsiloxy)styrene afforded the phenyl ketone 36. Reduction of 36 with LAH gave a mixture of 4 and 37.

(6)

OCH

34

35

36

37

2-Alkoxypiperidines such as 25 also serve as useful intermediates for the stereoselective synthesis of fran^-2,6-dialkylpiperidines. For example, the reaction of an 0-alkylamidal with a Lewis acid such as BF30Et2, generates acyliminium ion 38 which, followed by addition of a nucleophile, gives rran5-2,6-disubstituted piperidines 39 through a highly stereoselective nucleophilic substitution (Scheme 6) [31]. It is worthy of note that highly stereoselective syn- or anti-alkylation of 2-alkoxy-6-substituted piperidines can be readily achieved by the proper choice of organometallic alkylating agents and/or Lewis acids used.

384

I. Ojima and D. M. lula

BFgOEta MeO

AN

N ^COoMe I COaMe

(±)|

^COoMe

COaMe

25

38 CuBr • MeaS RLi

1V>'' R'

N ^COoMe I COgMe 39

R = allyl, /7-C3H7, />C4H9, n-CjH^s, n-CuHag Scheme 6

2.3 Syntheses of 2,6-Disubstituted Piperidines via Cyclohydrocarbonylation

(CH2)ioMe

40

22

Cis- and rran5-2,6-disubstituted piperidines such as a powerful teratogen pinidine (40) [32] and solenopsin A (22) [33] represent a common piperidine alkaloid type that possesses a wide range of interesting biological activities. Pinidine was isolated from several species of the dendrobate frogs, Pinaceae family. Solenopsin A is one of five structurally analagous piperidines isolated from the venom of fire ants Solenopsis saevissima. Transition-metal

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

385

catalyzed carbonylations offer efficient methods for the synthesis of these 2,6-disubstituted piperidines of biological significance.

2.3.1

Cobalt-Catalyzed Processes

The combination of anodic oxidation and cobalt-catalyzed carbonylation provides an efficient method for preparing N-acyl-a-amino esters and has been successfully applied to the synthesis of trans- and cw-A^-acyl-6-methylpipecolate (equation 7) [16]. Tran^-pipecolate 43 is easily epimerized to c/^-pipecolate 44 using a base. The driving force for this facile epimerization is ascribed to the release of A^'^ strain.

2e Me^^^^N^

A

41

f

"1

C02(C0)8

(7)

M^nu M e ' ^ ^ N ^ ^ ' ^ O M e ^^'^^ Me^ ^^^^ I (100 atm)

A,

ioo°c

^N^ I

'''COgMe

/^^

42

43 NaOMe

44 The highly linear selective hydroformylation and the subsequent intramolecular Wakamatsu reaction in situ of (iS^-A/'-benzoylallylglycinate 45 afforded A^-benzoylteneraic acid (48) stereoselectively in one step, which is readily hydrolyzed to teneraic acid (49) (Scheme 7) [17]. The reaction very likely proceeds via hemiamidal 47, but this intermediate cannot be isolated under the reaction conditions. These cobalt-catalyzed processes require high temperature and high pressure, i.e., lOCC and 150 atm of CO/H2 (1:1)- Thus, this process serves as an efficient industrial method, but it is not easy for academic laboratories to carry out these reactions.

I. Ojima and D. M. lula

386 Co(CO)8

^ ^ ^ HOaC^'

CO/H2 * 150atm(1:1) 100°C

NH COPh

CHO HOgC^'

HO2C COPh

'N

OH

COPh

62%

45

47

46

CO/H2 Co{CO)8

HCI HOaC""'

N

^COgH

(quant.)

HOoC'"'

CO2H

H

49 Scheme 7

2.3.2

Rhodium-Catalyzed Processes

Rhodium catalyzed cyclohydrocarbonylations may be carried out under relatively mild conditions. For example, the cyclohydrocarbonylation of JV-acylglycinate 50 catalyzed by a rhodium-diphosphite complex afforded either 6-alkoxypipecolate 51 or 5,6-didehydropipecolate 52 exclusively depending on the solvent used (equation 8) [34]. When the reaction was run in an aprotic solvent such as THF, CH2CI2, hexane, ethyl acetate or toluene, the enamide 52 was the sole product, whereas the reaction in an alcoholic solvent gave amidal 51 exclusively. It is noteworthy that no racemization takes place during the reaction when enantiomerically pure Nacylglycinate 50 is employed. The reaction is carried out under very mild conditions, i.e., 65 °C and 4 atm of CO/H2 (1/1) and versatile synthetic intermediates, l-acyl-6-alkoxypipecolates 51, are obtained in excellent yields, which is in stark contrast to the cobalt-catalyzed reactions {vide supra). The proposed mechanism for the formation of 51 and 52 is shown in Scheme 8 [34].

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

387

or |l^ R^OCHN'""^COpR2 CO/H2 (1/1, 4 atm) EtO^

65°C

50

I

^COgR^

, 1 / ^ ^

R'^

a: RU*Bu;R2 = Me b: R^ = Me ; R2 = *Bu

^N^ "O

51 (in alcohol)

1

(8)

N-^^COgR^

I

, 1 ^ ^ , R^" "O 52 (in non-protic solvent)

MeO

BIPHEPHOS =

MeO

As Scheme 8 illustrates, the first step of this process is the extremely regioselective hydroformylation catalyzed by the BIPHEPHOS-Rh complex, giving the linear aldehyde 53. Cyclization of aldehyde 53 to hemiamidal 54 followed by loss of a hydroxyl group generates acyliminium ion 55. The addition of an alcohol to 55 gives 6-alkoxypipecolate 51, while the deprotonation and 1,3-double bond migration yields enamide 52. The 6-alkoxy group of pipecolate 51 serves as a handle to introduce a substituent at C(6) stereoselectively as exemplified in Scheme 9 [34]. The nucleophilic substitution of the ethoxy group at C(6) of (5')-51a with n-BuCu'BF3 complex proceeded with excellent diastereoselectivity (>92% de), giving rr<2n5-l-r-Boc-6-n-butylpipecolate (trans-56a) in 80% isolated yield (100% de); the nucleophilic attack on the acyliminium ion 55 generated in situ takes place almost exclusively anti to the ester group at C(2). Because of A^'^ strain, trans-56 undergoes facile epimerization at C(2), giving the thermodynamically more favorable cis-57 in 72% isolated yield (100% de) upon reaction with LiHMDS in THF. The enantiomeric integrity of the starting enantiopure (5)-51a is maintained during the catalytic reaction (vide infra) as well as during the alkylation at C(6).

I. Ojima and D. M. lula

388

[Rh] OHC CO/H2

^

50

53

R^-'''^0 54

N 0 ^COgR^

55 -H*

R^OH

N'

R^-^O

R^-^0

51

52

Scheme 8

^C0,R2

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

389

i) BuCu " BF3 /~

EtO"

/J~

," ~ nBu"'

iii) NH4Cl/NH3

-N" "C02Me

tBuO"~O

C02Me

tBuO--'~O

51

56

~iHMDS

nBu~,,.~N.~ ,,,tCO2Me tBuO"~O 57

Scheme 9 Not only are the 2,6-disubstituted piperidines discussed above useful in the synthesis of piperidine alkaloids, but they also serve as key intermediates in the synthesis of izidine alkaloids. For example, short routes to indolizidine alkaloids, from ant venom, (+)-monomorine (61a), 5ethyl-3-methylindolizidine (61b), and 5-hexyl-3-methylindolizidine (61c) have been reported (Scheme 10) [35]. (+)-Monororine was isolated from Monomorium pharanosis [36] while 5ethyl-3-methylindolizidine and 5-hexyl-3-methylindolizidine was isolated from Solenopsis

(Diplorhoptrum) conjurata and Solenopsis (Diplorhoptrum) species AA respectively [37]. 2-Hydroxymethylpiperidine 58, which was synthesized in several steps from L-aniline, was oxidized to aldehyde 59 followed by the Homer-Emmons reaction using an appropriate phosphonate to give t~,[~-unsaturated ketones 60a-c. Hydrogenation of the double bond as well as hydrogenolysis of the Cbz group afforded an amino ketone which spontaneously cyclized to form the corresponding indolizidine alkaloids 61a-c in good overall yield.

. Ojima and D. M. lula

390

Me'

OH N I COgCHgPh

CHO

58

59

Me^ Me

R

61a : R = (CH)2CH3 61b I R = CH2CH3 61c : R = (CH2)5CH3

N COaCHaPh

Q

60a : R = (CH)2CH3 60b : R = CH2CH3 60c : R = (CH2)5CH3

i) (COCI)2, DMSO, EtgN; ii) (H3CO)2P(0)CH2COR, NaH, THF; iii) H2, Pd(0H)2 MeOH Scheme 10

2.4 Syntheses of Tri- and Tetrasubstituted Piperidine Alkaloids via Cyclohydrocarbonylation 2,3,6-Trisubstituted piperidine alcohol prosopinine (1), was isolated from the African mimosa Prosopis africana Taub and possesses antibiotic and anesthetic properties [4]. Dexoynojirimycin, 62, a 2,3,4,5-tetrasubstituted piperidine is a potent glucosidase inhibitor [38]. These polyfunctionalized piperidines can be prepared using the rhodium-catalyzed cyclohydrocarbonylation as the key step [39].

62

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

391

An efficient synthesis of prosopinine (1) using the rhodium-catalyzed cyclohydrocarbonylation as the key step is shown in Scheme 11. Fully protected enantiopure Dserine (64) was subject to DIBAL reduction to give aldehyde 65. Aldehyde 65 was then treated with vinylmagnesium bromide to give the corresponding alkenyl amino alcohol as the major product (trans Zeis = 4:1). The hydroxy! group of this product was protected as a silyl ether 66. The trans-isomer of 66 was isolated by column chromatography. COOH HoN^ ^ ^

i, ii, Hi

COOMe HN ^ ^ ^ HN' I COo'Bu

63

CHO I

iv

HN' 1 COa'Bu

OTBS

OTBS

65

64 V, VI

OTBS OTBS

.xV,0H

OH

OTBS - ^ ^ ' Y ^ ' ) 8 \

/

1 H

COg^Bu

68 i) 2,2-dimethoxypropane, HCI; ii) (f-Boc)20, KgCOg; iii) f-BuMegSiCI, imidazole; iv) DIBAL; v) vinyl-MgBr, -5°C (trans); vi) t-BuMegSiCI, imidazole; vii) Rh(acac)(CO)2, BIPHEPHOS, Hg/CO (1:1, 4 atm), EtOH, 65°C ; viii) f-BuLi, CuBrMegS, 69; Ix) BU4NF; x)TFA. Scheme 11

392

1. Ojima and D. M. luia

Cyclohydrocarbonylation of 66 catalyzed by Rh(acac)(CO)2-BIPHEPHOS complex in ethanol at 65°C and 4 atm of C0/H2(l:l) gave amidal 67 in excellent yield. The side chain precursor (69) was prepared in 5 steps from commercially available 1,10-dihydroxydecane and then coupled to 67. Deprotection of 68 afforded enantiopure prosopinine (1) in good overall yield [39]. Cyclohydrocarbonylation of 66 in THF instead of ethanol under the same conditions gave didehydropiperidine 70 (equation 9). Didehydropiperidine 70 can serve as the key intermediate for a variety of piperidine alkaloids including deoxynojirimycin (62).

,vOTBS

Rh(acac)(C0)2

^

BIPHEPHOS

h

^

\ ^.NOTBS

I

OTBS H2/CO (4 atm, 1:1) ^ N ' THF, 7 0 % I COg^Bu

Bu^COg-H

66

(9) .OTBS

70

J^ /

62

2.5 Syntheses of Piperidine Alkaloid Skeletons via Cyclohydrocarbonylation of Aminodienes Another approach toward the piperidine alkaloid skeleton is the regioselective cyclohydrocarbonylation of aminodienes such as 71 which gives access to 5,6dihydropiperidines or 2,6-disubstituted piperidines (equation 10) [40,41].

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

393

Rh(acac)(C0)2 BIPHEPHOS /K '\ H PG

CO/H2(4atm, 1:1) HO 2v , ;

N I

71 PG = Ts, f-Boc, Cbz

OHO

74 a: R = H CHO 74 b: R = Et Depending upon the reaction conditions, different functionalized piperidine skeletons can be selectively synthesized (Table 1). All reactions were run using 1 mol % Rh(acac)(C0)2 and 2 mol % BIPHEPHOS. As entries 1-3 illustrate, when the carbonylation reaction is run for 16 h at 65°C in THF, the enamide 73 is the only observed product. When the reaction is stopped at 2 h the formation of hemiamidal 74a is observed (entry 4). Hemiamidals 72 and 74a (72/74a = 3/2) were obtained when the reaction was run at low temperature (entry 5), while the Oethylamidal 74b was the only product isolated when the catalytic reaction was run in ethanol (entry 6).

Table 1. Cyclohydrocarbonylation of 4-amino-l,6-heptadienes (71). Entry

PG

Solvent

1 2 3 4 5 6

Cbz r-Boc Ts Ts Ts t-Boc

THF THF THF THF THF EtOH

Time (h) 18 18 18 2 3.5 18

Temp (°C) 65 65 65 65 45 65

Conversion (%) 100 100 100 100 70 100

Product 73 73 73 74a

72,74a 74b

394

m NH

I

I. Ojima and D. M. lula

Rh(acac)(C0)2 Ligand CO/H2 0HC

PG 71

[Rh] Hg/CO

Cf\ -

CCi ^ r ^ ^PG OH

^ ^ 0 ^ P G ^CHO 76

PG

CHO

74a

H+

CHO

PG

CHO

OEt 74b

73

Scheme 12 The proposed mechanism for the formation of these products is shown in Scheme 12. The first step is the linear-selective hydroformylation of one of the two double bonds, which is followed by cyclization to give alkenyl-hemiamidal 72. The hydroformylation of the remaining double bond takes place to afford hemiamidal-aldehyde 74a. When the catalytic reaction is run for a longer period of time or when 74a is subjected to column chromatography on silical gel, 74a undergoes dehydration via an iminium ion intermediate to yield the enamide 73. When the reaction is run in ethanol, ethanol adds to iminium ion 76 to give 6>-ethylhemiamidal 74b. All of the piperidines discussed above contain functionalities at critical positions which potentially allow for construction of the second ring system to yield the izidine alkaloid skeleton.

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

395

3. HYDROCARBONYLATION ROUTES TO IZIDINE ALKALOIDS

3.1

Izidine Alkaloids

The common term "izidine" alkaloid refers to a class of fused bicyclic alkaloids which bears a nitrogen at the bridgehead position. The alkaloids consisting of the 1~ azabicyclo[3.3.0]octane ring system are called pyrrolizidine alkaloids, the 1azabicyclo[4.3.0]nonane alkaloids are referred to indolizidines, and the alkaloids possessing the l-azabicyclo[4.4.0]decane skeleton are termed quinolizidines, which are often referred to as lupine alkaloids. These ring systems are found widely in nature ranging from a simple monofunctionalized to highly complex systems with various functionalities and additional ring systems. Pyrrolizidine [42-44], indolizidine [15,45-47], and quinolizidine [42,45,48] alkaloids have been the subject of numerous reviews. Examples of izidine alkaloids are shown in Figure 2. Two examples of pyrrolizidine alkaloids are (35,5J?,85)-3-heptyl-5-methylpyrrolizidine (77) [49] which is found in the venom of cryptic thief ants from the genera Monomonium and Solenopsis, and retrorsine (78) isolated from from Senecio isatideus [50]. Neotropical frog alkaloid (-)-gephyrotoxin 223AB from the Dendrobatidae family (79) [7] is an example of indolizidine alkaloids that has attracted considerable attention due to its strong biological activity as a noncompetitive blocker of neuromuscular transmission [15]. Two additional indolizidine alkaloids that are well known for their potent biological activities are swainsonine (80), isolated from the fungus Rhizoctonia leguminicola [51], which has recently been selected for clinical testing as an anti-cancer drug [52] and pumilotoxin B (81), isolated from Dendrobatid Amazonian frogs, which is a potent cardiovascular agent [53]. Examples of quinolizidine alkaloids are lupinine (82) isolated from leguminous plants [54] and porantherilidine (83) [55] from the Australian shrub Poranthera corymbrosa A. Brongn., family Euphorbiaceae. Tetracyclic quinolizidine (-)-spartein (84) is often used as a chiral ligand in asymmetric synthesis as well as a reagent for optical resolutions [56].

3.2 Strategies for the Construction of Izidine Alkaloid Ring Systems via Catalytic Carbonylations The use of transition metal catalyzed carbonylation for the construction of izidine alkaloid skeletal systems has proven to be a viable synthetic methodology. There are two different carbonylation approaches as illustrated in Scheme 13. As the retrosynthetic routes A and B indicate, izidine alkaloid skeletons can be synthesized through the cyclohydrocarbonylation of

I. Ojima and D. M. lula

396

.0 H

o-^ o

N-

H Me ^o

H

N<

Me

C7H15 C3H7 77 78

•lOH

80

/ H

PH

=

CO 82

Figure 2

79

C ^HQ '^4'-'9

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

397

alkenylpiperidines or aminoalkadienes. The carbons coming from the carbon monoxide are highlighted.

R,

nj

m c=>

nI

m PG

or

I

>

CHO n I

n I mi N H PG

m PG

OHO

Scheme 13

Functionalized indolizidine and quinolizidine skeletons can be readily prepared via route A. Cyclohydrocarbonylation of lactam 85 bearing an a-methallyl tether catalyzed by a rhodium complex gave functionalized izidine alkaloid skeleton 86 or 87 (Scheme 14), while hydroformylation followed by an intramolecular Wakamatsu reaction catalyzed by Co2(CO)8 afforded izidine alkaloid skeleton 88 bearing a carboxylic acid (Scheme 14) [20,21,57].

1. Ojima and D. M. lula

398

Rh(PPh3)3CI C0/H2 HC(0Et)3^

Rh4(CO)i2 CO/H2

-N OEt

C02(CO)8 C0/H2

o 87

86

rv

CO2H

88

Scheme 14 The quinolizidine skeleton 91, has been prepared via route B starting from aminodiene 71 as shown in Scheme 15 [41]. Cyclohydrocarbonylation of 71 catalyzed by the RhBIPHEPHOS complex in THF gave didehydropiperidine-aldehyde 73, which was reduced and mesylated to afford 90. The hydrogenolysis and hydrogenation of 90 on Pd/C gave 91 in good overall yield. Another approach to the synthesis of pyrrolizidine and indolizidine alkaloid skeletons involves the use of rhodium-catalyzed silylcyclocarbonylation of alkynylamines [58]. For example, (/?)-2-(3-butynyl)pyrrolidine 92 was converted into indolizindine 93 in one step using Rh(acac)(CO)2 as catalyst (equation 11). In a similar manner the reaction of l-(3butynyl)tetrahydroisoquinoline 94 gave tricyclic indolizidine 95 (equation 12) [58]. In both cases the E isomer was obtained exclusively. This apparant "trans" addition is due to the ZtoE isomerization of a p-silylvinyl-[Rh] or P-silylacryloyl-[Rh] intermeidate via a zwitterionic Rhcarbene intermediate. [58]

m

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

H

/\.

399

Rh(acac)(C0)2 BIPHEPHOS CO/H2{1:1;60psi)

COaCHaPh

THF

CHO ^ ^ ^

COgCHgPh

71 73 NaBH.

OMs

MsCI DMAP, py COaCHaPh

89 Pd/C

91 Scheme 15

H

NH 92

Rh(acac)(C0)2 HSiMePha CO (1 atm) toluene, 28°C

(11) SiMePh,

400

1. Ojima and D. M. lula

Rh(acac)(CO)2 HSiMegPh CO (50 atm) H

(12)

toluene, 60°C

R^Si The products of these particular carbonylation reactions have not been elaborated further, but serve as proof that functionalized izidine alkaloid skeletons can easily be obtained with these methods. Sections 3.3 to 3.5 describe how these methods can be used in the synthesis of izidine alkaloids.

3.3

3.3.1

Syntheses of Pyrrolizidines Alkaloids

(±)-Isoretronecanol and (±)-Trachelanthamidine

The synthesis of necine bases (±)-isoretronecanol (100) and (±)-trachelanthamidine (101), which are contained in a variety of pyrrolizidine alkaloids, has been achieved through the combination of silylformylation and cyclohydrocarbonylation (Scheme 16) [2,59]. Silylformylation of 5-ethynyl-2-pyrrolidinone 96 with HSiMe2Ph catalyzed by Rh(acac)(CO)2 gave 97 exclusively in excellent yield. Aldehyde 97 was then reduced with NaBH4, the silyl group removed and the resulting alcohol was protected as the TBDMS ether to yield 98. Cyclohydrocarbonylation of 98 catalyzed by HRh(CO)(PPh3)3 in HC(0Et)3 afforded pyrrolizidine 99 as a mixture of diastereomers. After separation of the two diastereomers of 99, the syn isomer was converted to isoretronecanol (100) while the fran^-isomer led to trachelanthamidine (101) through removal of the protecting groups followed by reduction with LAH.

3.3.2 Ant Venom (35,5iR,85)-3.Heptyl-5-Methylpyrrolizidine The key precursor to the ant venom (35,5i?,85)-3-heptyl-5-methylpyrrolizidine (77) was synthesized through the regioselective rhodium-catalyzed hydroformylation of l-r-Boc-2ethenylpyrrolidine 102 (Scheme 17) [60]. The extremely high linear selectivity in the formation of aldehyde 103 was achieved with the Rh(acac)(CO)2-BIPHEPHOS complex as the catalyst.

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

401

Aldehyde 103 was converted to alcohol 104 through the reaction with n-heptyl magnesium bromide. The nitrogen protecting group was changed from Boc to Cbz giving intermediate 105 [61] which had been previously converted to the ant venom through deprotection and cyclization in one step.

H

CHO 97

96

ii, iii, iv

OTBDMS

72 %

i

OEt

O

L.

OTBDIVIS

98

99 syn isomer

trans isomer VI, VII

H

iT-OH

100 i. Rh(acac)(C0)2, HSii^egPh, 20 atm CO, toluene, rt; ii. NaBH4, EtOH/HjO (1:1); iii. TsH, CH3CN (2 % H2O), reflux; iv. TBDMSCI, imidazole; v. HRh(CO)(PPh3)3,110atmCO/H2(1:1), HC(0Et)3, 100°C, (syn/anti = 2/1); vi. n-Bu4NF ; vii. LiAIH4,THF, reflux.

Scheme 16

402

1. Ojima and D. M. lula

Me^''

N I COg'Bu

I COg'Bu

77%

103

102

CTH

7^15

Me^''

Me

(CH2)6Me 77

i. Rh(acac)(CO)2 , BIPHEPHOS, 5 atm CO/H2 (1:1), THF, 60°C; ii. NaBH4, EtOH, 0°C; iii. 3 N HCI, then B20C(0)CI; iv) Hg, Pcl/BaS04, MeOH Scheme 17

403

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

3.4 Synthesis of Indolizidine Alkaloids (±)-167B and (±)-209D via Carbonylation H

H

N. (CH2)2Me

(CH2)5Me

106

107

Formal syntheses of indolizidine alkaloids (±)-167B (106) and (±)-209D (107), isolated from the skin extracts of frogs belonging to the genus Dendrohates, have been achieved [60] in a manner similar to the synthesis of pyrrolizidine alkaliod 77. The key step in the synthesis of these naturally occurring alkaloids is the extremely regioselective hydroformylation. In this case, the hydroformylation of 2-propenyl-l-r-Boc-pyrrolidine (108) yielded terminal aldehyde 109 exclusively which gave the indolizidine 110 through deprotection and cyclization in the presence of KCN (equation 13). Rh(acac)(C0)2 BIPHEPHOS N C02*Bu

CO/H2 (1:1,5atm) 60°C, THF 83%

CHO

(13)

N COo^Bu

108

109 3NHCI KCN

110

Intermediate 110 can be converted to each of the four diastereomers of alkaloids 167B and 209D (Scheme 18) [62]. Reaction of the cyanoindolizidine 110 with an appropriate Grignard reagent gave the (55,9/?) isomers of 106 and 107 in good yields, while the reaction of 110 with LDA followed by the addition of an appropriate alkyl bromide gave 111 which was reduced to give the {5R,9R) isomers of 106 and 107.

404

1. Ojima and D. M. lula

H

H RMgBr

N.

EtgO, -78°C to 0°C

N.

CN 110

(5S, 9R) 106: R = C3H7 (167B) (5S, 9fl) 107: R = CgHia (209D;

i) LDA, THF -78°CloO°C ii) RBr - 78° C to 0°C

NaBH. EtOH,25°C

lir.RrrCaHy 111:R = C6Hi3

(5/?,9fl)106:R = C3H7(167B) (5f?, 9R) 107: R = CQH^^ (209D)

Scheme 18

3.5

Syntheses of Functionalized Quinolizidine Alkaloids

Quinolizidine alkaloids can be synthesized through elaboration of functionalized piperidines obtained from the cyclohydrocarbonylation of aminodienes (Section 2.5). For example, quinolizidine 116, the key intermediate of 6-epi-porantherilidine (6-epi-83), was synthesized in several steps from carbonylation product 73 (Scheme 19) [41]. Aldehyde 73 was protected as the TBDMS ether and subjected to catalytic hydrogenation to give saturated piperidine 112. A^-f-Boc-piperidine 112 was treated with ^ec-BuLi and TMEDA followed by the addition of dimethyl sulfate to afford exclusively fran5-6-methylpiperidine 113 in excellent yield. TBDMS ether 113 was treated with TBAF followed by oxidation with TPAP to afford aldehyde 114. Wittig-Homer reaction of 114 gave 115, which was treated with TFA followed by K2CO3 to afford both diastereomers of 116 through an intramolecular Michael addition. Further manipulation of the ester moiety of 116 led to 6-epi-porantherilidine (epi-83).

405

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

i CHO

80% H

COa'Bu

71

73

OTBDMS

OTBDMS

'C COg'Bu 112

113

CHO

114

COaEt

115

Me

OC(0)Ph epi-83

COgEt

116

i) Rh(acac)(C0)2 .^BIPHEPHOS, CO/Hg (1:1, 4 atm), 65°C,THF; ii) NaBH4, EtOH; ii) TBDMSCI, imidazole, THF; iv) Hg, cat. Rh(C), EtOH; v)sec-BuLi, TMEDA, Me2S04; vi) n-BuNF; vii) TPAP; vii) (H3CO)3P(0)CH2C(0)OEt, NaH; ix) a. TFA, b. KgCOg Scheme 19

I. Ojima and D. M. lula

406

4. SYNTHESES OF QUINAZOLINE ALKALOID SKELETONS THROUGH CARBONYLATIONS Quinazolines constitute a small part of the alkaloid kingdom, yet there is substantial interest in these alkaloids because of their long history of usage in folk medicines [63,64]. Examples from the four main distinct quinazolines are shown in Figure 3. Glomerine (117) is produced by the millipede Glomeris marginata [65]. Glycosmicine (118) has been isolated from the leaves of Glycosmis arborea [66]. Pyrroloquinazoline alkaloid deoxyvasicine (119), has been isolated from Adhatoda vasica [67]. Pyridoquinazoline alkaloid 6,7,8,9-tetrahydrolliy-pyrido[2,l-Z7]- quinazoline (120) has been isolated from the leaves and stems of Mackinlaya subulata [68]. Me

117

118

119

120

Figure 3 The synthesis of quinazoline skeletons via transition metal-catalyzed carbonylation dates back to 1956 when the synthesis of quinazoline 122 was reported via the cobalt-catalyzed carbonylation of azobenzene (equation 14) [69]. In 1971, quinazoline 124 was shown to be formed from the reaction of 4-methylaniline with CCI4 and CO in the presence of a chromium carbonyl catalyst (equation 15) [3,70]. A radical-based mechanism was proposed for this unique transformation [70].

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

407

CO, C02{CO)8 1

(14)

230°C, 150atm 80%

121

Cr(C0)6 CO, CCI4

123

124

A novel approach to the synthesis of quinazoline alkaloids has been developed by means of the rhodium-catalyzed hydroformylation-cyclocondensation of diaminoalkenes [71]. The hydrocarbonylation of 2-(iV-allylaminomethyl)anilines 125a and 125b catalyzed by a Rh complex resulted in the formation of quinazolines 126 and 127, respectively, in good yields (equation 16). Benzylamine 125a gave hexahydropyrroloquinazoline 126a exclusively through extremely regioselective hydroformylation in the first step, whereas 125b bearing a methallyl moiety gave a mixture of hexahydropyrroloquinazoline 126b and tetrahydroquinazoline 127b. Reaction of 2-(iV-3-butenylaminomethyl)aniline (128) gave hexahydropyridoquinazoline 129 along with a small amount of the tetrahydroquinazoline 130 in excellent total yield. Although the mechanistic study has not been reported for these unique processes, formation of hexahydroquinazoline 126b can be rationalized by a mechanism shown in Scheme 20. The first step should be the hydroformylation of the olefin moiety of 125b to give diaminoaldehyde 131. Nucleophilic attack of the proximal amino group on the aldehyde moiety should

Ojima and D. M. lula

408

result in cyclization to form hemiaminal 132. Then, the formation of the iminium ion 133 followed by the addition of the aniline nitrogen should take place to give 126b.

[Rh(OAc)2]2 PPhg *CO/H2(1:1,400psi) ethyl acetate, 80°C

126a R = H (96 % ) 126b R = M e

125a R = H 125b R = Me 127a R = H (not observed) 127b R = M e

130

(16)

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

.NHa

CHo

409

[Rh] CO/H2 CHO

125b

131

132

133 H*

Scheme 20

CONCLUSION This chapter has described the current status of synthetic methods and methodologies on the basis of transition-metal catalyzed carbonylations for the syntheses of piperidine, izidine, and quinazoline alkaloids. Although these approaches to the syntheses of alkaloids are still new and under development, the new carbonylations, cyclohydrocarbonylations, and related reactions discussed in this chapter certainly have a very high potential to serve as efficient and convenient methods for the rapid construction of core alkaloid skeletons. The further development of efficient catalyst systems, that can achieve extremely high regioselectivity in the

410

1- Ojima and D. M. lula

hydroformylation and related functionalizations under mild conditions, will surely expand the applicability of these methods. In addition to the extremely high regioselectivity, the efficient asymmetric induction in these hydrocarbonylation process will be extensively investigated in the next decade, which will make these methods and methodologies discussed here more attractive and practical to many synthetic organic chemists who are engaged in alkaloid synthesis.

6.

ACKNOWLEDGMENTS

The authors would like to thank the National Institutes of Health (NIGMS), the National Science Foundation, and Mitsubishi Chemical Corporation for their generous support of our research in the development of new methodologies for alkaloid syntheses.

7. REFERENCES 1.

LS Hegedus, Transition Metal Organometallics in Organic Synthesis; Vol. 12, Elsevier Science, Ltd.,

2.

I Ojima, Catalytic Asymmetric Synthesis; VCH: New York, 1993.

3.

HM Calquhoun, DJ Thompson, and MV Twigg, Carbonylation: Direct Synthesis of Carhonyl Compounds;

Tarrytown, 1995.

Plenum Press, New York, 1991. 4.

GB Fodor and B Colasanti, In Alkaloids: Chemical and Biological Perspectives; Vol. 3, SW Pelletier, Ed., John Wiley and Sons, New York, 1985, pp 1.

5.

GM Strunzand and JA Findlay, In The Alkaloids; Vol. 26, A Brossi, Ed., Academic Press, New York, 1986, pp89.

6.

A Numata and T Ibuka, In The Alkaloids, Vol. 31, A Brossi, Ed., Academic Press, New York, 1987, pp 193.

7.

JW Daly, HM Garraffo, and TF Spande, In The Alkaloids, Vol. 43, G. Cordell, Ed., Academic Press, San Diego, 1993, pp 185.

8.

MW Edwards, JW Daly, and CW Myers, J. Nat. Prod. 51, 1188 (1988).

9.

JW Daly, Y Nishizawa, MW Edwards, JA Waters, and RS Aronstam, Neurochemical Res. 16, 489 (1991).

10. MJ Schneider, In Alkaloids: Chemical and Biological Perspectives, Vol. 10, SW Pelletier, Ed., Elsevier Science Ltd, New York, 1996, pp 155. 11. L Marion, R Lavigne, and L Lemay, Can. J. Chem.. 29, 347 (1951). 12. P Pec and I Frebort, J. Enzyme Inhib. 4, 327 (1991). 13. WV Brown and BP Moore, Aust. J. Chem. 55, 1255 (1982).

New Approaches to the Syntheses of Piperidine, Izidine, and Quinazoline Alkaloids

411

14. JW Daly, CW Myers, and N Whittaker, 7b;cicon. 59,395(1987). 15. JW Daly and TF Spande, In Alkaloids: Chemical and Biological Perspectives, Vol. 4, S. W. Pelletier, Ed., John Wiley and Sons, New York, 1986, pp 1. 16. K Izawa, S Nishi, and S Asada, 7. Mol. Cat. 41, 135-146 (1987). 17. K Izawa, /. Syn. Org. Chem., Jpn. 46, 218-231 (1988). 18. K Izawa, Petrotech 1989,12, 100-109. 19. H Wakamatsu, J Uda, and N Yamakami, Chem. Comm. 1540 (1971). 20. I Ojima, I. and Z Zhang, Organometallics 9, 3122-3127 (1990). 21. Z Zhang and I Ojima, /. Organometallic Chem. 454, 281-289 (1993). 22. I Ojima and A Korda, Tet. Lett. 30, 6283 (1989). 23. I Ojima, A Korda, and WR Shay, J. Org. Chem. 56, 2024-2030 (1991). 24. P Beak and WK Lee, J. Org. Chem. 58, 1109 (1993). 25. T Shono, Y Matsumura, and K Tsubata, J. Am. Chem. Soc. 103, 1172 (1981). 26. T Shono, Y Matsumura, and K Inoue, /. Chem. Soc. Chem. Comm. 1169 (1983). 27. T Shono, Y Matsumura, K Tsubata, and K Uchida, J. Org. Chem. 51, 2590 (1986). 28. T Shono, Y Matsumura, O Onomura, and T Sato, J. Org. Chem. 53, 4118 (1988). 29. T Shono, T Fujita, and Y Matsumura, Chem. Lett. 81 (1991). 30. MF Robert and MF Brov/n^Phytochemistry 20, 447 (1981). 31. C Ludwig and LG Wistrand, Acta. Chem. Scand. 48, 367 (1994). 32. H Neuhofer, L Witte, M Gorunovic, FC Czygan, Pharmazie 48, 389 (1993). 33. S Leclercq, I Thirionet, F Broeders, D Daloze, R Vander Meer, and JC Braekman, Tetrahedron 50, 8465 (1994). 34. I Ojima, M Tzamarioudaki, and M Eguchi, / Org. Chem. 60, 7078-7079 (1995). 35. H Takahata, H Bandoh, and T Momose, Tetrahedron 49, 11205 (1993). 36. FJ Ritter, IBM Rotgans, E Talman, REJ Verweil, and F Stein, Experientica 29, 530 (1973). 37. TH Jones, RJ Hight, MS Blum, and HM Fales, J. Chem. Eco. 10, 1233 (1984). 38. AB Hughes, DW Rudge, Nat. Prod. Rep. 11, 135 (1994). 39. ES Vidal, and I Ojima, 214th American Chemical Society National Meeting, Las Vegas, NV 1997, September 7-11, ORGN 311. 40.

M Tzamarioudaki, Z Li, D M lula, M Eguchi, and I Ojima, 211th American

Chemical

Society

National

Meeting, New Orleans, LA 1996, March 24-28, ORGN 206. 41. DM lula, M Tzamarioudaki, and I Ojima, 214th American Chemical Society National Meeting, Las Vegas, NV 1997, September 7-11, ORGN 339. 42. DJ Robins, In The Alkaloids; Vol. 46, GA Cordell, Ed., Academic Press, San Diego, 1995,pp 1. 43. JT Wrobel, In The Alkaloids; Vol. 26, A Brossi, Ed., Academic Press, Orlando, 1985, pp 327. 44. T Hartmann, and L Witte, In Alkaloids: Chemical and Biological Perspectives; Vol. 9, SW Pelletier, Ed., John Wiley and Sons, New York, 1994, pp 155. 45. AS Howard and JP Michael In The Alkaloids, Vol. 28, A. Brossi, Ed., Academic Press, New York, 1986, pp 183.

412

I- Ojima and D. M. lula

46. AD Elbein, and RJ Molyneux, In The Alkaloids: Chemical and Biological Perspectives, Vol. 5, SW Pelletier, Ed., John Wiley and Sons, New York, 1986, pp 1. 47. H Takahata and T Momose, In The Alkaloids, Vol. 44, GA Cordell, Ed., Academic Press, San Diego, 1993, pp 189. 48. S Ohmiya, K Saito, and I Murakoshi, In The Alkaloids; Vol. 47, GA. Cordell, Ed., Academic Press, New York, 1995, pp 1. 49. DJ Robins, Nat. Prod. Rep. 8, 213 (1991). 50. DJ Robins, Fortschr. Chem. Org. Naturst. 41, 115 (1982). 51. FP Guengerich, SJ DiMari, and HP Broquist, /. Am. Chem. Soc. 95, 2055 (1973). 52. PE Gross, MA Baker, JP Carver, and JW Dennis, Clin. Cancer. Res. 1, 935-944 (1995). 53. CW Daly and CW Myers. Science 156, 970 (1967). 54. K Asres, WA Gibbons, JD Phillipson, and P Mascagni, Phytochemistry 25, 1443 (1986). 55. SR Johns, JA Lamberton, AA Sioumis, and H Suares, Aust. J. Chem. 27, 2025 (1974). 56. P Beak, A Basu, DJ Gallagher, YS Park, S Thayumanavan, Ace. Chem. Res. 29, 552 (1996). 57. I Ojima and Z Zhang, Organometallics 417, 253-276 (1991). 58. I Ojima, D Machnik, RJ Donovan, O Mneimne, Inorg. Chimica Acta 251, 299-307 (1996). 59. I Ojima, RJ Donovan, M Eguchi, WR Shay, P Ingallina, A Korda, and Q Zeng, Tetrahedron 49, 54315444 (1993). 60. GD Cuny and SL Buchwald, Synlett 519 (1995). 61. O Provot, JP Celerier, H Petit, and G Lhommett, J. Org. Chem. 57, 2163 (1992). 62. RP Polniaszek and SE Belmont, J. Org. Chem. 55, 4688 (1990). 63. S Johne, In The Alkaloids; Vol. 29, A Brossi, Ed., Academic Press, New York, 1986, pp 99. 64. MF Grundon, In Alkaloids: Chemical and Biological Perspectives; Vol. 6, SW. Pelletier, Ed., Wiley: New York, 1988, pp 339. 65. YC Meinwald, J Meinwald, and T Eisner, Science 154, 390 (1966). 66. SC Pakrashi and J Bhattacharyya, J. Sci. Ind. Res. B (India) 21, 49 (1962). 67. K Pandita, MS Bhatia, RK Thappa, SG Agarwal, KL Dhar, and CW Atal, Planta Med. 48, 81 (1983). 68. JS Fitzgerald, SR Johns, JA Lamberton, and AH Redcliffe, Aust. J. Chem. 19, 151 (1966). 69. S Murahashi, S Horiie, J. Am. Chem. Soc. 78, 4816 (1956). 70. Y Mori and J Tsuji, Tetrahedron 27, 3811 (1971). 71. EM Campi, WR Jackson, QJ McCubbin, and AE Trnacek, Aust. J. Chem. 47, 1061 (1994).

Subject Index

Note: Bold page numbers refer to tables and illustrations. There are often textual references on the same pages

abereamines 196 acetoxyacetylpyrrole 267, 268 2-Qf-acetoxyhetisane-l^, 9^, \9a triol see 2-acetylseptentriosine 6-0-acetylacosepticine 303, 304 7V-acetyladeemin 180, 181 14-0-acetylbrowniine 305, 307, 312, 314, 337 acetylcholine 99, 101, 222-3, 224 acetylcholine esterase 222-3 11-acetyl-13-cinnamoylhetisine see palmadine acetylcorymine 195, 196 2-acetyl-13-dehydro-11 -e/7/-hetisine 352, 353 1,14-di-O-acetyldelcosine 315 14-0-acetyldelcosine 305, 312, 318, 319, 337 14-0-acetyldelectine 305 6-0-acetyldelpheUne (10-deoxydeltaline) 306, 307, 318, 319, 321 1-0-acetyldelphisine 301 14-0-acetyl-l-deoxydelcosine 318, 319 14-0-acetyl-10-deoxydictyocarpine 310 14-0-acetyldictyocarpine 306, 307, 310, 318 ll-acetyl-2,13-didehydrohetisine 350, 351-2, 353 13-acetyl-2,11 -didehydrohetisine 349 6-O-acetylheteratisine 323, 324 2-acetylhetisine 350, 351 11-acetylhetisine 349, 350, 351 11-0-acetylhetisine 328 11,13-di-O-acetylhetisine 328 13-acetylhetisine 349, 350, 351 13-0-acetylhetisine 328 tri-0-acetylhetisine 330 6-0-acetyl-14-O-methylforesticine 299 8-0-acetyl-14-0-methylforesticine 299 14-acetyl-8-0-methyltalatizamine 296 8-0-acetylneohne 357 14-0-acetylneoline 300, 356, 357 14-0-acetyl-l-e/7/-neoline 301, 357

acetylnorechitamine 202, 203 14-0-acetylnudicaulidine 311, 312, 337 14-0-acetylperegrine 344 2-acetylpyrrole 239 0-acetylsamandarine 4 iV-acetylsepaconitine 299 2-acetylseptentriosine (2-a-acetoxyhetisaneIjS, 9)3, 19a triol) 331, 332 8-0-acetyltalatizamine 296 14-0-acetyltalatizamine 295 8,14-di-O-acetyltalatizamine 296 A^-acetyltryptamine 220 13-0-acetylvakhmatine 334 acoforesticine 297, 298 acoforestine 297, 298 acoforestinine 297, 299 acoforine 297, 298 aconite-type norditerpenoid alkaloids 292, 294-303, 315-17 aconitine 22, 294, 316, 317, 318, 319, 320, 356, 362 acosepticine 303, 304 acoseptridine 299, 300, 303, 304 acoseptrigine 299 acoseptriginine 299, 300 acoseptrine 303, 304 acoseptrinine 303, 304 A^-acyl amino acids 373, 375 A^-acylamino alkanals 373, 375 //-acylglycinate 386, 387 acyliminium ion 383, 387, 388 adaline 373, 374 agelasine G 253 ageliferins 250 ageline B 253 aging, cyclotryptophans and 221-2 ajabicine 294, 334-5 ajacine 305, 309, 315 ajaconine 339, 341, 342

413

414

ajacusine 305 ajadelphine 305 ajadelphinine 305, 314 ajadine 305 ajadinine 305, 306 ajanine 305, 306 akuammilane 199 akuammiline 197 aldisin 246 alkaloids see amphibian skins alkenyl-hemiamidal 394 2-alkoxypipecolate 379 6-alkoxypipecolate 386, 387 2-alkoxypiperidine 379, 380, 381-2, 384 alline 188 allodihydrohistrionicotoxin 32, 35, 37 allopumiliotoxins, see also pumiliotoxins allosedamine 383 allotetrahydrohistrionicotoxin 32 alstonamide 201 amauromine 172, 173, 181 ambiguine 298, 305, 309, 310 amidal 392 Z>-amino acid 166 aminoalcohols 267, 268 aminodienes, cyclohydrocarbonylation of 392-4 4-amino-1,6-heptadienes 393 aminolactone 243, 244 amphibian skins, alkaloids from 1-161 tabulated 113-47 see also bicyclic alkaloids; indole alkaloids; monocyclic alkaloids; pyridines; steroidal alkaloids; tricyclic alkaloids anabasamine 103-4 anabaseine 103-4 anabasine 103-4 anatoxin 101 andersobine 294, 337, 338, 339 andersonidine 311, 312, 337 andersonine 311, 312, 337 anhweidelphinine 361 anthranilate 181, 219 anthranilic acid 170, 219 4-anthranoyllapaconidine 299, 300 anthranoyllycoctonine 303, 304, 305, 306, 309 relative toxicity in mice 360, 361 ardeemin 180 asperazine 183, 184 asperlicin C 218, 219

Subject Index

asperlicin E 170, 181, 182, 218, 219 asterinins A-C 278 aszonalenin 170, 180, 218 atisane-type diterpenoid alkaloids 292 atisine 329, 330, 332 auricularine 191, 192 axinohydantoin/fuscin 246, 247 azaacenaphthylene 83, 85 azabicycl-[3.3.0] octanes see pyrrolizidines azabicycl-[4.3.0] nontanes see indoUzidines azabicycl-[4.4.0] decanes see quinolizidines azabicycl-O-5.3.0]decanes 12, 36, 76, 77, 110 azafulvenium 254, 255 azitine 336, 337 azomethine 322, 323 bacitracin 166 balfourine 295, 296 barbaline 343, 344 barbeUne 306, 307 barbinidine 307, 308 barbinine 307, 308, 361 barbisine 343, 344 monoacetate 343 batrachotoxinin A 6, 7, 9-10, 22 batrachotoxins 6, 7, 8-10, 22, 109-10, 112 batzellines A 256 B 256 C 256, 259, 260, 261 BE 10988 273, 274 iV-benzoylallylglycinate 385, 386 18-0-benzoyldavisinol 343, 344, 346 14-0-benzoyldelphonine 301, 302 6-benzoylheteratisine 323, 324, 332, 362 iV-benzoylteneraic acid 385, 386 bicoloridine 298 bicyclic alkaloids 10-77, 110 azabicycl-O-5.3.0]decanes 12, 76, 77, 110 see also decahydroquinolines; histrionicotoxins; homopumiHotoxins; indolizidines; pumiliotoxins and allopumiliotoxins; pyrrolizidines; quinolizidines bikhaconitine 295 bisditerpenoid alkaloids 300 bishaconitine 296 bishatisine 296 blacknidine 313 blacknine 313 borreverine 191, 192

Subject Index brevetoxin 22 brevianamide E 172, 173, 219 bromoaldisin 246 bromocyclotryptamines 210 4-bromohymenin 253, 254 bromonium 254, 255 bromopyrrole alkaloid 249, 250 L-6-bromotryptophan 224 browniine 305, 306, 307, 310, 312, 337 bufadienolides 2 Bugula pyrrole 268 bullatine C 295, 337 a-bungarotoxin 99 3-butenamide 375, 377, 378 2-(iV-3-butenylaminomethyl)aniline 407, 408 butyl-m-cycloheptylprodigiosin 242 butyl-indolizidines 60 butyl-pyrrolizidines 53 rra«5-2-butyl-5-pentylpyrrolidine 89 /ra«5-l-/-Boc-6-«-butylpipecolate 387, 389 2-3-(butynyl)pyrrolidine 398, 399 1 -(3-butynyl)tetrahydroisoquinoline 398, 400 cabuamine 204, 206 cabufiline 199, 202, 205 calycanthadine 189, 190 calycanthine 108, 109, 111, 214 calycosidine 192, 193 cammaconine 295, 296, 317, 318 carbonylations see transition metal catalyzed carbonylations L-3-(2-carboxy-4-pyrrolyl)-alanine 276 catalyzed carbonylations see transition metal catalyzed carbonylations ceylanicine 204, 206 ceylanine 204, 205 chaetocin 174, 185 chaetocin B and C 174, 175 chasmanine 295, 297, 302 chellespontine 336, 337 chetomin 175, 176 chetracin A 174, 175, 185 chimonanthine 108, 109, 111, 189, 190, 191, 213, 214, 219 chlorisondamine 100 10-chlorodelpheline 320, 321 10-chloro-lO-deoxydeltaline 320, 322 chloropyridyl azetidine ABT-594 101 cholesterol acyltransferase 179 chromoxymycin 274

415 clathramides 251 clathrodin 251, 253 cobalt-catalyzed processes in synthesis of 2,6-disubstituted piperidines 385-6 coccinellines 20, 83, 84-5, 109, 111, 112 columbianine 295 columbidine 296, 298 condelphine 295, 360 congeners homopumiliotoxin 28-31 pumiliotoxin 23-5 corymine 195, 196 coryzeylamine 197, 198 crassicaudine 302, 303 crassicauline A 297, 302, 303, 319 crassicausine 302, 303 crassicautine 302, 303 cuauchichicine 353, 354 cyanoindolizidine 403-4 cyclizations in peptides and proteins 220-5 cyclodipeptides 170 cyclohydrocarbonylation of aminodienes 392-4 2,6-disubstituted piperidines 384-90 syntheses 384-90 tri- and tetra-substituted piperidine alkaloid syntheses via 390-2 cycloinamide-A 208 cycloneosamandarine 5 cycloneosamandione 4 cyclopenta[Z?]quinolizidines 80, 81-2, 111 cycloprodigiosin 242 cyclosporin A 166 cyclotryptamine 165, 216 see also cyclotryptophans cyclotryptophan 165, 216 cyclization in peptides and proteins 220 senescence and aging 221 cyclotryptophans and cyclotryptamines, naturally occurring 163-236 animals 207-15 bacteria 166-8 biosynthesis 215-20 cyanobacteria 158-70 fungi 170-85 higher plants 186-207 see also cyclizations in peptides and proteins c>^c/o-(L-tryptophyl-L-phenylalanyl) 182, 184 cytisine 99

416 damirone A and B 260, 261 davisine 344, 346, 347 davisinol 343, 344, 345, 346 deacetylajadine 305, 306 14-0-deacetylambiguine 305, 309, 310 S-deacetyl-S-O-ethyl-l-fi'/^/delphisine 357 deacetylheterophylloidine 327 deacetyllappaconitine 330 N-deacetyllappaconitine 299, 300, 361-2 8-0-deacetyl-8-0-methyl-l-£77/delphisine 357 14-deacetylnudicauline 359, 360, 361 14-0-deacetylnudicauline 307, 311, 312, 313, 339 14-deacetylnudiculine 337 14-0-deacetylnudiculine 312 6-deacetylperegrine 344 15-O-deacetylvacognavine 327, 328, 343 8-deacetylyunaconitine 295 debromo-8, 8a-dihydroflustramine C 212 debromoaxinohydantoin 247 debromofiustramine B 209, 210, 212 debromohymenialdisine 247, 253, 254, 256 decahydroquinolines 11-12, 20-1, 36-7, 39, 40-5,46,47,48, 109, 110, 112 decamethonium 100 iV-deethyldelstaphinine 301 deformylcorymine 196, 197 deformylcoryzeylamine 197, 198 14-dehydrobrowniine 307, 315 dehydrobufotenine 104, 256, 257 dehydrococcinelline 84 14-dehydrodelcosine 307 1-dehydrodelphisine 301, 302, 318, 319 1,2-dehydrodelphisine 318, 320 1-dehydrodelstaphisine 301 1-dehydrodeltamine 307 13-dehydro-2,l 1-diacetylhetisine 352-3 6-dehydrodictyocarpinine 308 6,14-dehydrodictyocarpinine 308 6,7-dehydro-5,8-disubstituted indolizidine 30-1 ll-dehydrohetisine 348,349-50 13-dehydrohetisine 352 dehydrohistidine 218 8,9-dehydrohomopumiliotoxins 29, 30 14-dehydrotalatizamine 296 delajacine 305, 306 delajacirine 305, 306 delajadine 305, 306 delatisine 339, 340, 341 delavaine 311, 337

Subject Index A and B 309, 361 delbruline 314, 315 delbrusine 314 delcorine 314 delcosine 305, 307, 310, 312, 337, 360 delectine 304 delectinine 311,313,337,339 delelatine 307, 310, 311, 313, 339 delnudine-type diterpenoid alkaloids 292 delphatine 304, 305, 306, 307, 314 delpheline 294, 306, 307, 310, 312, 313, 314 318, 321, 339 delphidine 315, 316 delphinines 95, 300, 301, 305, 314-15, 316, 337, 356, 360-1 delphisine 315, 316, 318, 319, 337 epimerization of C(l)-Qf-hydroxyl group of 356, 357-9 delsemine A and B 309, 360 delsoline 305, 308, 311, 315, 360 delstaphidine 300, 301 delstaphigine 301, 302 delstaphinine 301, 302 delstaphisagnine (14-0-acetylneoline) 300 delstaphisagrine 300, 301 delstaphisine 300, 301 delstaphisinine 301 deltaline 296, 337, 339 deoxydeltalines 306, 307, 318, 319, 321 heterolytic fragmentation of 320-3 lycoctonine-type alkaloids and 305, 306, 307, 309, 312, 313 methylenation and demethylation 314, 315, 318 relative toxicity in mice 360, 361 deltamine 307, 309, 312, 318, 319, 339 deltamine-S-methyldithiocarbonate 318, 319 deltatsine 305, 310 delvestidine 303, 308, 309 delvestine 308, 309 demethoxyalstonamide 201 demethoxy-A^-demethylvincovine 205 demethylcorymine 195, 197 demethyldeformylcorymine 197 6-0-demethyldelphatine 303, 304 demethylenation of norditerpenoid alkaloids 315-17 demethylpeceyline 204 A^-demethylpleicorine 201, 202 deoxyaconine 356 deoxybrevianamide E 173

Subject Index }-deoxydelcosine 318, 319 6-deoxydelpheline (occidentaline) 306, 307, 314, 318 10-deoxydeltaline (6-0-acetyldelpheline) 306, 307, 318, 319, 321 6-deoxydeltamine 318, 319 9-deoxyhomopumiliotoxins 28, 29 deoxynojirimycin 390, 392 8-deoxyperhydrohistrionicotoxin 34-5 deoxypumiliotoxins and other pumiliotoxin congeners 23-4, 25 deoxysamanine (samane) 5 deoxyvasicine 406 depsipeptides 165 desacetyltacraline 206 A^-desethylaconitine 319, 320 A^-desethyldeltaline 361 A^-desethylmethyllcaconitine 361 A^-desethylpacifiline (pacifinine) 313 desformocorymine 196, 197 16,18-di-O-desmethyl aconitine 316, 317 16,18-di-O-desmethyldelphinine 316 16,18-di-O-desmethyI-14,16diacetyllappaconitine 317 9-desmethylhomopumiliotoxins 25, 28, 29 14,16-di-O-desmethyllappacinitine 317 8-desmethylpumiliotoxin 23, 25 desoxycabufiline 201, 202 2-despentylperhydrohistrionicotoxin 34 dethio-tetra(methylthio)chetomin 175, 176 diacetyldavisine 344, 346 6,14-diacetyldelelatine 310 2,3-diacetylhetidine 330 2,11-diacetylhetisine 349, 350, 351, 352 11,13-diacetylhetisine 350-1 8,14-diacetyltalatizamine 356 2,6-dialkylpiperidine 378, 379 2,3-dibromo-5-methoxymethylpyrrole 248 dibromoagelaspongin hydrochloride 248 dibromoisophakellin 248 dibromophakellstatin 249 dibromopyrrole 248 metabolite 251 4,5-dibromo-2-pyrrollic acid 248 dictyocarpine 296, 306, 307, 310, 312, 314, 337, 361 dictyocarpinine 307 dictyzine 294, 341, 342 6,14-didehydrodictyocarpinine 308 2,11-didehydrohetisine 348,349-50 5-6-didehydropipecolate 386, 387

417 didehydropiperidine 392 -aldehyde 394, 398 didemnins 212 digoxin 361 dihydro-)8-erythrodine 100 dihydro-j8-erythroidine 100 dihydroasitine 336, 337 dihydroaszolenalenin 180 7,8-dihydrobatrachotoxinin A 7 3-£/7/-dihydrocorymine 196, 197, 203 3-acetate 196, 197 3,17-acetate 196, 197 17-acetate 196, 197 dihydroflustramine C and A^-oxide of 210, 211 dihydrogephyrotoxin 78, 79 dihydrohistrionicotoxin 32 dihydropiperidines 392 6,10-dihydropumiUotoxin 23, 25 dihydropyridone 378 dihydro-2-pyridone 375, 377 3,4-dihydro-2-pyridone 377, 378 3,12-dihydroroquefortine 171 dihydroveatchine 355, 356 dihydroxychaetocin/melinacidin 174, 176, 177, 179 2,5-diketogluconic acid 243 diketopiperazine 170, 179, 181, 217 "dimers" 43, 44, 45 2,2-dimethoxypropane 391 dipropylquinolizidines 67 discorhabdin C 256, 257, 259, 260 dispacamides 251, 252 2,5-disubstituted decahydroquinolines 36, 110 disubstituted indoUzidines 3,5-disubstituted 12, 50, 53-4, 55-7, 58-60, 109, 110 5,8-disubstituted 12, 53, 61, 62-5, 66-7, 110 2,6-disubstituted pipecolate 379-80, 381, 382 2,6-disubstituted piperidines 92, 93-4, 95, 109, 111,383,384,385,389,392 2,5-disubstituted pyrroUdines 89, 90, 91-2, 109, 111 3,5-disubstituted pyrrolizidines 48, 49, 50, 53, 109, 110 disubstituted quinolizidines 1,4-disubstituted 12, 73-5, 76, 110 4,6-disubstituted 12, 50, 71, 72-3, 109, 110

418 diterpenoid alkaloids 292, 294 hetisine-type 326-47 rearrangement reactions 347-59 ditryptophenaline 182, 184 dopamine 101 echitamine 202, 203 echitaminic acid 203 eladine 312, 313, 339 elanine 313, 339 elasine 312, 313 elatine 312, 314, 339, 360, 361 elsasine 339 enantiopure 387 D-serine 391 epiamauromine 181 epibatidines 95, 96, 97, 98-103, 111 epiboxidine 102 epidithiodiketopiperazines 177 see also sporidesmin epilupinine esters 278 epinardins A-D 258, 259 epipolythiodioxopiperazines see leptosin erinicine 195, 196 erinine 195, 196 eripinal 196 eripine 195, 196 erysodine 100 eseramine 188, 189 ethyl-3-methylindolizidine 389, 390 0-ethylhemiamidal 394 ethyl-indolizidines 64, 67 5-ethyl-3-methylindolizidine 389 eudistomin 263-5 A 263 M 263, 264 exochomine 278, 279 falaconitine 296, 297 falconericine 296, 297 falconeridine 296, 297 falconeridinine 296, 297 falconerine 296, 297, 317 -8-0-acetate 296, 297 flexicorine 199, 200 fluorotryptamine 219 fluorotryptophan 219 jflustarine B 189, 210, 215 flustramide A and B 209, 210 flustramine A 209, 210, 211-12

Subject Index flustramine B 209, 210, 211-12 flustramine B A^-oxide 210 flustramine C 209, 210, 212 flustramine D and A^-oxide of 210, 211 flustramine E 209, 210 flustraminol A and B 209, 210 flustrarine B 210 folicanthine 189, 190 foresaconitine (vilmorrianine C) 297, 302 foresticines 297, 298, 299, 300 forestine 297, 298, 302 A^-formylkynurenine 229 formylroquefortine 171, 172 fructigenine A 172 fructigenine B/verrucofortine 172 funebradiol 243 funebral 243, 244, 245 funebrine 242, 243 3-furfuryl pyrrole-2-carboxylate 239, 240 fuscin/axinohydantoin 246, 247 gancidin 179 garryfoline 353, 354, 355 geneserine 189, 210, 215 geneseroline 189, 210, 215 gephyrotoxins 39, 57, 78, 79-80, 111 223AB 395, 396 4-geranylpyrrole-2-carboxylic acid see pyrrolostatin gigactonine 305, 308 glaucedine 307 glaucenine 306, 307 glaucerine 307 glomerine 406 glutathione 178 glycerinopyrin 239 glycosmicine 406 gramicidin S 166 grossularine 224 guanidine 246 gypsetin 178, 179 hemiamidal 374, 375, 385, 386 heptenylindolizidines 64 heptenynylindolizidines 64 3-heptyl-5-methylpyrrolizidine 395, 396, 400, 401-2 heptynylindolizidines 64 heteratisine 323, 332 -type norditerpenoid alkaloids 292, 323-6 heterophyllidine 323, 324

Subject Index

heterophylline 323, 324 heterophyllisine 323, 324 heterophylloidine 326, 327 hetidine 327 hetisine 328, 329, 330, 341, 344 acetylation of 350-1 acid-catalyzed rearrangement of 347, 348 ll-c/^z-hetisine 348 oxidation 348-9 -type diterpenoid alkaloids 326-47 hetisinone 341, 342, 344 hexahydropyrroloquinazoline 407, 408 hexahydroquinazoline 407, 408 hexahydroquinolines 43-4 hexamethonium 100 hexenyl-indolizidines 67 3-hexyl-5-methylindolizidines 60 5-hexyl-3-methylindolizidine 389, 390 5-hexyl-indolizidines 50 hexylpropylpyrrolizidines 56 himastatin 167, 168 hippocasine 84 hippodamine 84 histamine 104 histidine 218 histrionicotoxins 11, 31, 32-5, 36-9, 48, 78, 110, 111,373,374 hodgkinsine 192, 193 hokbusine A 298 homobatrachotoxin 6, 7, 9, 110 homopumiliotoxins 11, 25, 26-7, 110 congeners 28, 29-30, 31 HS3 270 hunteracine 196, 197 hydroformylation 377 hydrogen peroxide 178 hydroxyacetylcyclotryptamine 220 3-(hydroxyacetyl)indole 219 hydroxycyclotryptamin 188, 215 hydroxycyclotryptophans 218, 220, 224 hydroxydihydroborreverine 191 (2S, 3S, 4R)-hydroxyisoleucine 243 a-hydroxyisovaleric acid 168 2-hydroxymelatonin 215 6-hydroxymelatonin 215 cw, c w-4-hydroxy-2-methy 1-6nonylpiperidine 89 2-hydroxymethylpiperidine 389 hydroxy-iV-methyltrimethoxyaporphine 337 5-hydroxypiperazic acid 168 17^-hydroxysamane 5

419 hymenialdisine 247, 248, 253, 254, 256 hymenidin 245, 246, 247, 249 hymenin 246, 253, 254 hypaconitine 319, 320 ibogaine 100 idiospermuUne 190, 191 imidazole 391 imidazolide 318, 319 3-0-(imidazolylthiocarbonyl) aconitine 319, 320 3-0-(imidazoiylthiocarbonyl)yunaconitine 319 iminophosporane 264, 265 iminoquinone 199, 200 indaconitine 295, 296 indole alkaloids 104-9 chimonanthine/calycanthine 108, 109, 111, 189,190, 191,213,214,219 see also pseudophrynamines indole-3-carbaldehyde 219 indolenine 216 indolizidines from amphibian skins 10, 12, 21, 39, 53-71, 110, 112 6,7-dehydro-5,8-disubstituted 30-1 3,5-disubstituted 12, 20, 53-4, 55-7, 58-60, 109, 110 5,8-disubstituted 12, 61, 62-5, 66-7 5,6,8-trisubstituted 12, 67, 68-70, 71 from transition metal catalyzed carbonylations 372, 389, 390, 395, 398, 400, 4 0 3 ^ indolizindine 398, 399 isoatisine 327, 353, 354-5 isobatzelline A, B and D 257 isobatzelline C 257, 259, 260, 261 isoborreverines 191 isocorymine 195, 196 isocycloneosamandaridine 4, 5 isodelectine 304, 308, 309 isodelpheline 312, 339 isodihydrohistrionicotoxin 31, 32, 35 isoflustramine D 210, 211 isophakellistatin 208 isopsychotridine A-C 193, 194 "isopumiliotoxin" 28 isoretronecanol 400, 401 isotalatizidine 295 isotetrahydrohistrionicotoxin 32 izidines 110

420

hydrocarbonylation routes to 395-400 spiropyrrolizidines 85, 86-8, 89, 109, 111 see also indolizidines; pyrrolizidines; quinolizidines kapakahine B 223, 224 karakoline 344 kawaguchipeptin B 170 keramadine 251, 253 keronopsins A and B 272 11-ketohetisine 327 kobusine 344, 346 kuwaguchipeptin 168, 169 kynureninase 229 kynurenine 229 lactone 243, 245 lamellarins A-D 265, 266 E-H 266 lapaconidine 299 lapaconine 299 lappaconitine 299-300, 317, 330, 361-2 lassiocarpine 347 leptosins A-H 184, 185, 187 2I-K 185 lipofuscin 221 LL-S490/3 170, 180 ludaconitine 295 lupinine 395, 396 lusidusculine 347 lycoctonine 293, 337, 339 -type norditerpenoid alkaloids 292, 303-13, 314-15 relative toxicity in mice 360, 361 lycogarubins A-C 275 lysozyme 224 makaluvamines 258, 259 A-C 260, 261 D 258, 259, 260, 261 manzacidins 251, 253 mauritamide A 251, 252 mauritiamine 251-2 mecamylamine 100, 101 melatonin 215 melinacidin 174, 176, 177, 179 II 176 III 176, 179

Subject Index mesaconitine 319, 320, 356 18-methoxygadesine 305 a. methoxy piperidine 383 5-methoxytryptamine 263, 264 8-0-methyl-14-acetyl-1 -epi-ntoXmt 357 18-des-O-methyl aconitine 316, 317 14-0-methylblacknine (pacidine) 313 methylborreverine 191, 192 A^-methylcarbamate 383 AT-methylconiine 379, 380, 381 A^b-methylcyclotryptamine 187 methyldecahydroquinolines 39 1-0-methyldelphisine 295, 337 S-methyldithiocarbonate 307, 318, 319 methylenation of norditerpenoid alkaloids 314-15 A^-methylepiamauromine 181 0-methyleudistomin M 264 14-0-methylforesticine 299, 300 0-methylhemiamidal 380, 381 A^-methylhistrionicotoxin 35 14-0-methylisodelpheline (paciline) 313 methyllindolizidines 56, 60, 64, 66, 67, 389, 390 methyllycaconitine 100, 305, 307, 309, 311, 312, 313, 314, 337, 339, 361 8-0-methyllycaconitine 303, 304, 312, 359, 360, 360 8-0-methyl-l-e'/7/-neoline 357 iV-methylperhydrohistrionicotoxin 35 TV^-methylpiperidines 92 rr<2rt5-6-methylpiperidine 404, 405 methylpropylpyrrolizidines 56 A^-methylpseudoconhydrine 379, 380, 382 methylpyrrolinone 378 methylpyrrolizidines 51, 53 8-0-methyltalatizamine 296 8-0-methyl-14-0-veratroylpseudaconine 295 micro toxins 168 milpacamide 251, 253 minfiensine 207 mithaconitine 296, 297 monocyclic alkaloids 89-95 see also piperidines; pyrrolidines monomargine 278, 279 monomorine 54, 389, 390 I 58, 59-60 morphine 101, 108 mycobacteria 177 myrmicarins 58, 84-5 myrrhine 84

Subject Index naloxone 100 napelline 347 neodihydrohistrionicotoxin 32 neolines 295, 300, 301, 315, 316, 356, 357 neolinine 300, 301 neopyrrolomycin 239, 240 nicotine 95,98,99, 104, 111 nifedipine 100 nigrifortine/amauromine 172, 173, 181 nitrendipine 22 nitropolyzonamine 85, 86, 87, 88, 111 nonadienynylindolizidines 64, 67 nonenylindolizidines 64, 67 noranabasamine 95, 103, 104, 111 nordesoxycabufiline 201, 202 norditerpenoid alkaloids 289-326, 347-62 aconite-type 292, 294-303, 315-17 biological activity 359-62 demethylation 315-17 deoxygenation reactions 318-20 heteratisine-type 292, 323-6 heterolytic fragmentation of deltaline 320-3 isolation techniques 293 lycoctonine-type 292, 303-13, 314-15 methylenation of 314-15 NMR spectral data in structure determination 293-4 phytochemical investigations 293 rearrangement reactions 347-59 norechitamine 202, 203 A^-oxide 203 norepinephrine 101 noreripinal 196 norisocorymine 195, 196 nornicotine 101 A^^-norphysostigmine 167, 168, 188 norvincorine 200, 201 nudicauline 311, 312, 337, 339, 360 occidentaline (6-deoxydelpheline) 306, 307, 314, 318 octahydrohistrionicotoxin 31, 32, 35 octahydroquinolines 43 octenynylindolizidines 64, 67 okaramine A and B 173 oroidin 251 oroidine 245, 246, 249 oscillatoric acid 168 oscillatorin 168, 169 oxaline 216

421 19-oxoajacine 305 19-oxoanthranoyllycoctonine 305 19-oxodelphatine 305 6-oxoeladine (pacinine) 313, 339 14-0-methylblacknine (pacidine) 313 pacidine (14-0-methylblacknine) 313 pacifidine 313 pacifiline 313 pacifinine (A^-desethylpacifiline) 313 paciline (14-0-methylisodelpheline) 313 pacinine (6-oxoeladine) 313, 339 palau'amine 250 palmadine (ll-acetyl-13cinnamoylhetisine) 328, 329 palmasine 328, 329 panicutine 326, 327 peceylanine 204, 205 peceyline 204 pelankine 204 pellitorine 277 pempidine 100 penicillin 166 pentenyldecahydroquinolines 39 pentenyhndolizidines 64, 67 pentylindolizidines 56, 64, 66 peptide synthetases 212, 224, 225 peptides 208 see also cyclizations in peptides and proteins perhydro-9^-azaphenalane 84 perhydrogephyrotoxin 79 perhydrohistrionicotoxin 34—5, 38, 48, 67 phakellins and related compounds 245, 248-56, 249 phakellistatin 208 phallotoxins 221 phenoxybenzamine 100 phenyl-ethy lene-ditry ptophenaline 182 3-phenylpropanoic acid 277 phenylpyrrolyloxazoles 262 philanthrotoxin 38 phorbazoles 262 phorbazoles A-D 261, 262 physostigmine 111, 167, 168, 188, 222 A^-oxide 189 pinidine 384 cw-pipecolate 385 /ra«5-pipecolate 385 piper alkaloids 276-7 piperazinedione 185

422

piperidines 241 2,6-disubstituted 92, 93-4, 95, 109, 111, 383, 384, 385-90, 392 from amphibian skins 39, 92, 93-4, 95,109, 111 iV-r-Boc-piperidine 404, 405 synthesis via transition metal catalyzed carbonylations 372, 374, 375-94 2,6-disubstituted 384, 384-90, 392 alkaloid skeletons 392, 393-^ pipeline 373, 374 pleicorine 201, 202 polyzonimine 85, 86, 87, 88 porantherilidine 395, 396 6-epi-porantherilidine 404, 405 precoccinelline 20, 83, 84 prenyltransferase 209, 218 prodigiosin 242, 242 propenylindolizidines 64 5-propyl-indolizidies 50 6-propylpipecolic acid 380, 381, 382 propyldecahydroquinolines 39 propyleine 83, 84 propylindolizidines 64, 66 prosopinine 373, 374, 390, 391 pseudaconitine 295, 296 pseudoceratine 262 pseudophrynamines 104-5,106-7, 108-9, 111 A 104, 106, 214 pseudophrynaminol 104, 105, 106, 107-8, 213, 214 psycholeine 193 psychotridine 193, 194 pteridine 213 puberaconitidine 298 pumiliotoxin A 12-13, 15, 18-19, 21 pumiliotoxin B 12-13, 16, 19, 21, 22, 395, 396 pumiliotoxin C 39 pumihotoxins and allopumiliotoxins 12-13, 14-18, 19-22, 2S-4, 25, 110 8-deoxypumiliotoxins and other pumiliotoxin congeners 23-^, 25 pyralomycins 274 pyridine 187 pyridines 95-104 epibatidine 96, 97, 98-103 noranabasamine 95, 103, 104 pyroaconitine-type norditerpenoid alkaloids 292 pyrodelphinine 301

Subject Index

pyrro-2-carboxylic acid methy esters 246 pyrroindomycins see pyrrolomycins pyrrole alkaloids 237-87 from acidians 263-5 from higher plants 278-9 from microrganisms 273-6 minor piper alkaloids with pyrrole skeleton 276-7 miscellaneous 266-72 from molluscs 265-6 simple pyrroles 237-40 from sponges 245-62 pyrrole-2-carboxylic acid 239 pyrrolethers 272 pyrrolidines 39, 89, 90-1, 92, 109, 111, 398, 399 pyrroUndomycins A and B 271 pyrrolines 91 pyrrolizidines 12, 48, 49-52, 53, 110, 112, 372, 395, 400-2 pyrrolnitrin 240-1 pyrrololactams 245, 246-7 pyrrolomycin A-E 269, 270 pyrrolomycins and pyrroindomycins 269-72 pyrrolopiperazine 251 pyrroloquinoHnes 256-61 pyrrolosporin 275 pyrrolostatin 239, 240 pyrroxamycin 270, 271 quadrigemine A and B 192, 193 quadrigemine C 193 quinazoline alkaloids 372 skeletons synthesis through carbonylations 406, 407, 408-9 quinolines 39, 43-4, 398, 400 pyrroloquinoHnes 256-61 see also decahydroquinolines quinolizidines 67, 372, 395, 398, 399 from amphibian skins 12, 71, 72-5, 110, 112 1,4-disubstituted 12, 73-5, 76, 110 4,6-disubstituted 12, 50, 71, 72-3, 109, 110 cyclopental[/?] 80,81-2, 111 functionalized 404, 405 rausutramine 199, 200 rausutrine 199, 200 retrorsine 395, 396 rhazidigenine 206, 207, 222

423

Subject Index

rhazidine 206, 207, 217, 222 rhazinilam 278, 279 rhodium-catalyzed synthesis of 2,6-disubstituted piperidines 385-6 rhodophycan pyrroles alkaloids 267, 268 roquefortine C 170, 171, 172, 216-17, 218 D 171, 217 roseophilin 279, 280-1 rumbrin 273

steroidal alkaloids 3-10 see also batrachotoxins; samandarines stevensine 245, 246, 253, 254, 255 strictamine 205 styloguanidines 250 superoxide radical 178 surfactin 166 surfactin synthetase 225 swainsonine 395, 396

sachaconitine 295 samandaridine 4 samandarines 3, 4, 5, 109 samandarone 4 samandenone 4 samandinine 4 samane (deoxysamanine) 5 samanine 4 sarmentine 277 sarmentosine 277 Sch 52900 and 52901 177 sciodole 276 a-scorpion toxin 22 jS-scorpion toxin 22 securamine A 211, 212, 217 securamine B 211, 212, 217, 222 securamine C-G 211, 212 securine A and B 211, 222 sedamine 373, 374, 383 sepaconitine 299 sepentriosine 298 septatisine 331, 332 septentrionine 303 septentriosine 331 solenopsins A and B 94, 95, 378, 379, 384 songorine 347 spartein 395, 396 spermacoceine 191, 192 spinceamines 104 spiradine A acetate 327 spirasin XIII and XV 343 spiropyrrolizidines 85, 86-8, 89, 109, H I , 112 sporidesmin A 174, 177, 178, 218 sporidesmin B, C, E and G 177, 178 sporidesmin D 177, 178, 179 sporidesmin F, H and J 177, 179 staphisadrine 301, 302 staphisadrinine 301, 302 staurosporine 219

tacraline 206 takaosamine 305, 310, 311, 337 talatizamine 295, 296, 297, 317, 318 talatizidine 295 tambjamines and related compounds 268, 269 tangirine 323-4, 325, 326, 332 tangutisine 332, 333 tanwusine 332 tatsidine 294, 310, 311 tatsiensine 309, 310 tatsinine 310 tatsirine 294 teneraic acid 385, 386 6,7,8,9-tetrahydro-11 //-pyrido-quinazoline 406 tetrahydroatisine 354 tetrahydrohistrionicotoxin 32 tetrahydroquinazoline 407, 408 tetrahydroquinolines 43-4 tetrasubstituted piperidine alkaloid syntheses via cyclohydrocarbonylation 390-2 tetrodotoxins 2, 22 thiazohalostatins 273 2-thiosubstituted tryptophan 221 trachelanthamidine 400, 401 transition metal catalyzed carbonylations 373-412 izidine alkaloids 395-400 piperidine alkaloid synthesis 373-94 quinazoline alkaloid skeletons 406-9 2,11,13-triacetylhetisine 350-1 1,2,19-triacetylseptentriosine 331 2,3,4-tribromopyrrole 239 tricyclic_alkaloids 77-89 coccinellines 20, 83, 84^5, 109, 111, 112 cyclopental[Z>]quinolizidines 80, 81-2, 111 spiropyrrolizidines 85, 86-8, 89, 109, 111 see also gephyrotoxins triketramine 267

Subject Index

424

trisubstituted indolizidines 12, 53, 67, 68-70, 71, 110 trisubstituted piperidine alkaloid syntheses 390-2 trypargine 104 tryptamine 165, 216, 220 tryptophan 105, 165, 216, 217, 218, 219, 220 urochordamine A and B 212, 213 vacognavine 327, 328, 343, 344 vakatidine 327 vakatisine 327 vakatisinine 327 vakhmadine 329, 330 vakhmatine 329, 330, 334 vakognavine 327, 328 vatamidine 194 vatamine 194 vatine and vatine A 194 veatchine 354, 355 -type norditerpenoid alkaloids 292, 323-6 veratridine 10, 22 14-0-veratroylbikhaconine 295

veratroylpseudaconine 295, 296 verrucofortine 172 verticillin A, B and C 176, 177 vilmorrianine C (foresaconitine) 297, 302 vincarubine 200, 201 vincorane 199 vincoridine 200 vincorine 200, 202, 204, 205 derivatives 202, 203 vincovine 200, 204, 205 vincristine 193 wakayin 257, 258 WIN 64745 183, 184 WIN 64821 183, 184, 219 X-14885A 272 xenoveine 51 yunaconitine 295, 297, 302, 303, 318, 319, 356 zaliline 361

Organism Index

Bold page numbers refer to tables.

Acanthella carteri 248 Aconitum ix, 292, 293, 359, 361 A. balfourii 295 A. columbianum 360 Nutt. ssp. columbianum 295, 296, 359, 360 A. crassicaule 302 A.falconeri 296 A.ferox 295 A.forrestii 297 Stapf. var. albo-villosum 297 A. heterophyllum 323, 326, 347 A. palmatum 323, 327, 343 A, paniculatum 326 A. septentrionale 299, 303, 330, 331, 332 A. tanguticum 323, 332, 362 Acrostalagmus 176 A. cinnabarinus var. melinacidinus 176 Actinomadura A. sp. HQ24 273 A. spiralis 21A actinomycete 242 Actinosporangium vitaminophilum SF2080 269 Adalia bipunctata 373 Adelotus 115 Adhatoda vasica 406 y4. clathrodes 251 ^ . conifera 251 >4. ^w/7<2r 2 5 1

A. flabelliformis 248 ^ . longissima 251 ^ . mauritiana 251 y4. oroides 248 Y4. sceptum 249 /I. altaicum 188 ^ . anisopodium 188 y4. bidentatum 188

v4. leucocephalum 188 v4. odorum 188 y4. ramosum 188 ^ . J^«e5c^«5 188 y4. sibiricum 188 ^ . splendens 188 v4. stelleranum 188 ^ . victorialis 188 Alstonia 201-^ yl. angustifolia 202 ^ . congensis 202 ^ . deplanchei 201 v4. glaucescens 203 ^ . macrophylla 201 ^ . odontophora 201 ^ . ondulata 201 y4. pachycharra {Winchia calophylla) 202-3 ^ . pZ/n'^r/ 202, 205 ^ . plumosa 201 yl. scholar is 201 yl. sphaerocapitata 201 y4. undulifolia 202 yi. vitiensis var. wovo ebudica monachino 201-2 Amauroascus sp. 173 Amphibia 213-15 Anabasis aphylla 103 y4«a/w 84 Anisostica 84 Anthonomus 84 Aphaenogaster 103, 104 Apocynaceae 186, 195-207 Aromabates 114 Ascidiaceae 212 Ascidiacea 212-13 Ascomycete 179 Aspergillus 180-4 ^ . alliaceus 181, 219 A. fisheri var. brasiliensis 180 A.flavus 182

425

426 var. columnaris 182 A. niger 184 A. ochraceus 181 A. zonatus 180, 218 Aspidospermaa quebracho bianco 206 Aster tataricus 278 Astroclera willeyana 250 Atelopus 115 Auxarthron umbrinum 273 Avena 172 Axenellida 208 Bacillus B. marinus 213 ^. 5w/?/f/w 225, 257 Balanus amphitrite 251, 252 Batzella sp. 256 Bombacaceae 282 Borreria verticillata 193 Botrylloides sp. 213 5r>^ozofl 209-12 Bufonidae 114, 115 alkaloid derivatives listed 117, 122-4, 127-8, 131-2, 134, 136, 138, 144-5 Bufo 115 B. marinus 108, 256 Bugula dentata 268, 269 Cabucala 204-6 C. caudata 205 C. erythrocarpa var. erythrocarpa 204 Ca/v/« 84 Calycanthaceae 109, 189-90, 191 Calycanthus 189, 214 Calycodendron milnei 193, 194 Candida albicans 257 Catharanthus 188 56e also Vinca Chaetomium 174-6 C. abuense 175 C. cochlioides 176 C.funicola 176 C. globosum 176 C. minutum 174 C. nigricolor 175 C. retardatum 175-6 C. subglobosum 176 C tenuissimum 175 C. thielavioideum 174 C. umbonatum 176 C. virescens var. thielavioideum 175

Organism Index Chartella 212 Chauliognathus 84 Chelaner antarcticus 48 Chenopodiaceae 103 Chilocorus 84 Chimonanthus 189 C. fragrans 219 Ciona savignyi 111, 251 Clavelina sp. 257 Clitocybe acromalaga 276 Coccinellidae 84 Coccinella 84, 85 Coccinula 84 Coleomegilla 84 Colostethus 114 Coltricia cinnamomea 111 Compositae 292 Conium 379 Conopharyngia chippii 206 Consolida ix, 292, 293, 336, 359, 360 C. ambigua (Delphinium ajacis) 304-5, 309, 334 C. hellespontica {Delphinium hellespontica) 336, 337 C. imperialis 225 C. radiatus 224 Cornebacterium fluccumfaciens 171 Corollospora 179 C.pulchella 179 Cribochalina olemda 224 Cryptobranchus maximus 5 cyanobacteria 168-70 Cy dorana 115 Delphinium ix, 292, 293, 361 JD. <3/flcw {Consolida ambigua) 304-5, 309, 334 i). andersonii 311, 337 i). Z?arZ)^>^/ 306, 307, 343, 359 Z). brunonianum 314, 341 D. ^flvw/f 344 D. delavayi Franch. var. pogonanthum 309 £). dictocarpum 341 D. e/flf/wm 310, 312, 314, 339 var. 'black night' 313 var. Pacific Giants Mix 312, 313 />. geyeri 359 D. glaucescens 359-60 D. hellespontica {Consolida hellespontica) 336, 337

427

Organism Index D. hybridum 360 D. occidentale 306, 318, 359, 360 D. staphisagria 300, 301, 316, 326 D. tatsienense 309, 310, 341 D. tricome 359, 360 Z). vestitum 308, 309 Demospongiae 207, 208 Dendrobatidae 96, 114, 395 alkaloid derivatives listed 116-46 Dendrabates 8, 9, 10, 19, 52, 66, 72, 75, 109, 110, 111, 113, 373,403 D. auratus 20, 21, 36, 37, 47, 53, 59, 60, 77, 79, 83, 84, 85, 87, 91 D. granuliferus 11, 91 D. histrionicus 31, 35, 36, 53-4, 58, 59, 60, 78, 79, 89, 90 D. imitator 39, 54, 59, 60 D. lehmanni 28, 36, 76, 77 D. leucomelus 21, 47, 60 D. occulator 59 D. pumilio 13, 24, 28, 36, 59, 61,67, 69, 73, 77, 83, 84, 85, 91, 95 D. speciosus 36, 54, 59, 61, 77, 89, 92, 95, 373 D. tinctorius 87 D, variabilis 24, 39 Dendrodoa grossularia TIA Dendrophryniscus 115 Dictyodendrilla sp. 262 Didemnum chartaceum 266 Diplorhoptrum 47, 53, 60, 73, 91 Ectoprocta 209 Epicladia flustrae 210 Epipedobates 8, 10, 19, 36, 52, 59, 66, 72, 75, 95,98, 109, 110, 113, 114 E. anthonyi 96 E. bassleri 39, 70 E. espinosai 98 E. macero 85, 87 E. pictus 98 E. pulchripectus 70, 87 E. silverstonei 83, 84, 98 E, tricolor 13, 16, 23, 24, 71, 87, 96, 98 Eudistoma olivaceum 263 Euphorbiaceae 395 Exochomus 84 E. quadripustulatus 21S Flavobacterium marinotipycum 213 Flindersia fourn ieri 191

Flustra 209, 210-12 F.foliacea 168,210 Garryaceae 292 Garry a spp. 359 Gliocladium 177 Glomeris marginata 406 Glycosmis arborea 406 Gonioma kamassi 206 Gracilariopsis lemaneiformis 267 Halo Cynthia roretzi 213 Hedyotis auricularia 192 Heleioperus 115 Helicoverpa zea 181 Herpes simplex 263 Hippodamia 84 Hodgkinsonia frutescens 193 Hmteria 195-9, 203 H. congolana 196 H. eburnea 196 H. elliotii 196 H. umbellata 195 H. zeylanica var. africana 196, 199 Hymeniacidon, H. aldis 246 Hyrtios proteus 184 Idiospermaceae australiensis 191 Lamellaria sp. 265 Latruncula 257 Leguminosae 186, 188-9 see also Physostigma Leptosphaeria 185 Liliaceae 186, 188 see also Allium Lissodendoryx sp. 246 Loganiaceae 186, 207 Lycogala epidendrum 275 Mackinlaya subulata 406 Mammalia 215 Mantellinae 115 Mantella 10, 16, 20, 24, 28, 31, 36, 43, 46, 53, 6 0 , 6 6 , 7 3 , 8 4 , 8 7 , 9 1 , 9 5 , 109, 113 alkaloid derivatives listed 116-47 M. aurantiaca 29 M. baroni 25, 28, 71, 75 M. betsileo 41 M. crocea 29 M. laevigata 71

428 M. viridis 28 Mantidactylus 115 Megalomyrmex 48, 53, 91 Melanophryniscus 10, 36, 66, 71, 73, 87, 95, 109, 113 M. moreirae 19, 46, 59 M. stelzneri 20, 28, 46, 48, 53, 54, 59, 60, 75, 83, 84 Micrapsis 84 Micrococcus luteus 21A Microcystis 168-70 M. aeruginosa 168 Micromonsporium 275 Minyobates 8, 10, 19, 20, 36, 52, 59, 66, 72, 75,87, 109, HI, 113 M. altobueyensis 70 M. bombetes 70, 80-1 M. minutus 70 Monanchora 247 Monocarpia marginalis 278 Monomorium 48, 53, 91, 95, 395 M. pharaqnis 54, 58, 60, 389 Mortierella ramaniana 213 Musca domestica 99 Muscicapidae 9 Myobatrachidae 114, 115 alkaloid derivatives listed 134,136, 138-^2, 144, 145-6 Myrmicaria 60, 84 M. eumenoides 58 Myrrha 84 Mytilus edulis galloprovencialis 213 Nannizzia 179 iV. gypsea var. incurvata 179 Notaden 115 Oscillatoria 168 O. agardhii 168 Penicillium 170-3, 216, 217P. atramentosum 171 P. brevicompactum 172, 219 P. chrysogenum 171 P. corymbiferum 171 P. crustosum 171 P. expansum 171 P.farinosum 171 P. fructigenum 172 P. glandicola 171 P. nigricans 172

Organism Index P. oxalicum 171, 217 P. purpurrescens 171 P. roquefortii 170, 171, 217 P. simplicissimum 173 P. terrestre 171 P. verrucosum var. cyclopium 171, 172 Petchia 204 P. ceylanica 204 Phakellia 208-9 P. carrer/ 208 P.flabellata 245 P.fusca 246 P. mauritiana 249 Phorbas aff. clathrata 262 Phyllobates 3, 10, 19, 36, 52, 66, 70, 72, 87, 95, 104, 109, 110, 111, 113 P. aurotaenia 6, 8, 9, 21, 31, 59, 60 P. ^/co/or 6, 8, 59, 85 P. %MZ>rw 8, 9, 71 P. /^rnZ?//w 6, 8-9, 59, 103, 108, 214 P. v//rfl[/M5 8, 9, 24, 83 Physostigma venenosum 188-9 Piper P. nigrum 373 P. sarmentosum 276, 277 Pithomyces 177-9 P. chartarum 177, 218 Pz7(?/iw/ 3,9, 110, 112 P. dichrous 9 P.ferrugineus 9 P. kirhocephalus 9 P. nigrescans 9 Polycitor sp. 264 Polyphysia crassa 239 Po/vzofl 209 Polyzonium rosalbum 87 Poranthera corymbrosa 395 Porifera 207-9 Prianos 257 Propylaea 84 Prosopsis africana 373, 390 Psammocinia sp. 208 Pseudoceratina purpurea 262 Pseudokeronopsis rubra 272 Pseudomonas P. nautica 213 P. pyrrocinia 240 Pseudophryne 10, 20, 36, 46, 60, 66, 71, 73, 75, 84, 87, 95, 109, 110, 111, 113, 213-15 P. coriacea 22, 104, 107, 214 P. corroboree 107

429

Organism Index P. guentheri 107 P. occidentalis 107 Pseudostellaria heterophylla 239 Psycho tria 193 P. beccaroides 193 P. forsteriana 193 P. oleoides 193 P. rostrata 193 Pythium altimum 176 Quaraibea 282 Quararibea, Q. funebris 242 Ranunculaceae ix, 292, 293 Rauwolfia 199-200 i?. reflexa 199 i?. sumatrana 199, 205 Rhacophoridae 114, 115 Rhazya stricta 206 Rhizoctonia leguminicola 395 Rosaceae 292 Rubiaceae 186, 192-5 Rutaceae 191-2 Salamandridae 3 Salamandra 3, 109 S. atra 5 5. salamandra 3, 5 5. terdigitata 5 Sargassum tortile 185 Saxifragaceae 292 Securiflustra 209-10, 212 5'. securifrons 210, 212 Senecio isatideus 395 Serratia marcescens 269 Solenopsis 91, 395 S. saevissima 384 subgenus Diplorhoptrum 47, 48, 60, 95, 389 5. conjurata 389 subgenus Euophthaline 47 subgenus Solenopsis 47, 95 Spirea japonica L. var. fortunei 343 Staphylococcus aureus 170, 174, 179 Streptomyces 166-7, 168

5. antibioticus 272 5. chrestomyceticus 239 5. griseolavus 224 5. griseoviridis 279 5. hygroscopicus 168 5. pseudogriseolus 168 iS. rubropurpureus 274 5. rugisporus 271 5. sp. MI 424-38 Fl 239 A-5071 239 BE 18591 268 S46506 270 . S. staurosporeus 219 5. violaceus 239 Strychnos minfiensis 207 Stylotella S. agminata 250 S. aurantium 250 Tabernaemontana 206 r . chippii 206 Tonduzia 202 Torp^Jc? 38, 60, 66, 99 Treponema hydrodysenteriae Tricholoma sciodes 276 Trikentrion loeve 267

Til

Uperoleia 115 Verticillium 111 V. dahliae 111 V. tenerum 111 Vinca 200-1 K. mmor 200, 205 F/rg//za F. divaricata 278 F. oroboides 278 Winchia calophylla {Alstonia pachycharra) 202-3 Xenopus 99 Zyzz3^a sp. 258

This Page Intentionally Left Blank