Progress in the Chemistry of Organic Natural Products 109 [1st ed. 2019] 978-3-030-12857-9, 978-3-030-12858-6

Table of contents :
Front Matter ....Pages i-v
Chemistry of the Secondary Metabolites of Termites (Edda Gössinger)....Pages 1-384
Bioactive Compounds Involved in the Life Cycle of Higher Plants (Hideyuki Shigemori)....Pages 385-413
Chemical Diversity and Biological Activity of African Propolis (Natalia Blicharska, Veronique Seidel)....Pages 415-450

Citation preview

Progress in the Chemistry of Organic Natural Products

A. Douglas Kinghorn · Heinz Falk Simon Gibbons · Jun’ichi Kobayashi Yoshinori Asakawa · Ji-Kai Liu Editors

109 Progress in the Chemistry of Organic Natural Products

Progress in the Chemistry of Organic Natural Products

Series Editors A. Douglas Kinghorn, Columbus, OH, USA Heinz Falk, Linz, Austria Simon Gibbons, London, UK Jun’ichi Kobayashi, Sapporo, Japan Yoshinori Asakawa, Tokushima, Japan Ji-Kai Liu, Wuhan, China Advisory Editors Giovanni Appendino, Novara, Italy Roberto G. S. Berlinck, São Carlos, Brazil Verena Dirsch, Wien, Austria Agnieszka Ludwiczuk, Lublin, Poland Rachel Mata, Mexico, Mexico Nicholas H. Oberlies, Greensboro, USA Deniz Tasdemir, Kiel, Germany Dirk Trauner, New York, USA Alvaro Viljoen, Pretoria, South Africa Yang Ye, Shanghai, China

The volumes of this classic series, now referred to simply as “Zechmeister” after its founder, Laszlo Zechmeister, have appeared under the Springer Imprint ever since the series’ inauguration in 1938. It is therefore not really surprising to find out that the list of contributing authors, who were awarded a Nobel Prize, is quite long: Kurt Alder, Derek H.R. Barton, George Wells Beadle, Dorothy Crowfoot-Hodgkin, Otto Diels, Hans von Euler-Chelpin, Paul Karrer, Luis Federico Leloir, Linus Pauling, Vladimir Prelog, with Walter Norman Haworth and Adolf F.J. Butenandt serving as members of the editorial board. The volumes contain contributions on various topics related to the origin, distribution, chemistry, synthesis, biochemistry, function or use of various classes of naturally occurring substances ranging from small molecules to biopolymers. Each contribution is written by a recognized authority in the field and provides a comprehensive and up-to-date review of the topic in question. Addressed to biologists, technologists, and chemists alike, the series can be used by the expert as a source of information and literature citations and by the non-expert as a means of orientation in a rapidly developing discipline. Listed in Medline. All contributions are listed in PubMed.

More information about this series at http://www.springer.com/series/10169

A. Douglas Kinghorn • Heinz Falk • Simon Gibbons • Jun’ichi Kobayashi • Yoshinori Asakawa • Ji-Kai Liu Editors

Progress in the Chemistry of Organic Natural Products Volume 109

With contributions by E. Gössinger H. Shigemori N. Blicharska
V. Seidel

Editors A. Douglas Kinghorn College of Pharmacy The Ohio State University Columbus, OH, USA

Heinz Falk Institute of Organic Chemistry Johannes Kepler University Linz, Austria

Simon Gibbons UCL School of Pharmacy University College London, Research London, UK

Jun’ichi Kobayashi Graduate School of Pharmaceutical Science Hokkaido University Fukuoka, Japan

Yoshinori Asakawa Faculty of Pharmaceutical Sciences Tokushima Bunri University Tokushima, Japan

Ji-Kai Liu School of Pharmaceutical Sciences South-Central University for Nationalities Wuhan, China

ISSN 2191-7043 ISSN 2192-4309 (electronic) Progress in the Chemistry of Organic Natural Products ISBN 978-3-030-12857-9 ISBN 978-3-030-12858-6 (eBook) https://doi.org/10.1007/978-3-030-12858-6 © Springer Nature Switzerland AG 2019 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Switzerland AG. The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Contents

Chemistry of the Secondary Metabolites of Termites . . . . . . . . . . . . . . . Edda Gössinger

1

Bioactive Compounds Involved in the Life Cycle of Higher Plants . . . . . 385 Hideyuki Shigemori Chemical Diversity and Biological Activity of African Propolis . . . . . . . 415 Natalia Blicharska and Veronique Seidel

v

Chemistry of the Secondary Metabolites of Termites Edda Gössinger

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Releaser Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.1 Trail-Following Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.2 Sex-Pairing Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.3 Syntheses of Trail-Following and Sex-Pairing Pheromones . . . . . . . . . . . . . . . . 2.1.4 Home-Marking Pheromones? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.5 Phagostimulating and Food-Marking Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.6 Aggregation Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.7 Alarm Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.8 Worker-Arresting Pheromone . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.9 Egg-Recognition Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.10 Building Pheromones? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.11 Necromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1.12 Cuticular Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Primer Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.1 Caste Differentiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.2 Queen (King) Pheromones (Reproductive-Specific Pheromones) . . . . . . . . . . . 2.2.3 Syntheses of (Presumed) Primer Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Chemical Defense . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Acetogenins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.1 The Long-Chain Contact Poisons of the Rhinotermitidae . . . . . . . . . . . . . . . . . . . 3.1.2 Macrolactones (Macrolides) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Compounds of Mixed Biosynthesis Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.1 Novel Free Ceramides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.2 Aliphatic Nitro Compounds as Defense in Termites . . . . . . . . . . . . . . . . . . . . . . . . .

5 11 13 13 23 34 84 86 86 87 93 93 93 93 94 102 103 104 107 109 117 117 121 131 131 134

E. Gössinger (*) Institute of Chemistry, University of Vienna, Vienna, Austria Mistelbach, Austria e-mail: [email protected] © Springer Nature Switzerland AG 2019 A. D. Kinghorn, H. Falk, S. Gibbons, J. Kobayashi, Y. Asakawa, J.-K. Liu (eds.), Progress in the Chemistry of Organic Natural Products, Vol. 109, https://doi.org/10.1007/978-3-030-12858-6_1

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3.3 Isoprenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3.1 Monoterpenes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3.2 Sesquiterpenes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3.3 Diterpenes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Biosynthesis of Secondary Metabolites of Termites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Biosynthesis of Pheromones . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1.1 Acetogenins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1.2 Biosynthesis of Isoprenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Biosynthesis of Components of the Frontal Gland Secretion . . . . . . . . . . . . . . . . . . . . . . . . 4.2.1 Isoprenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.2 Acetogenins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.3 Defensive Compounds with a Mixed Biosynthesis Origin . . . . . . . . . . . . . . . . . . 5 Families and Subfamilies of the Termites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Abbreviations % ee [α]D 0

00

18-crown-6 2D-NMR 8H-BINAP 9-BBN abs Ac acac AIBN aq ATP ATPB BHT Bn brsm Bu Bz CAN cat. CD CHC COLOC COSY Cp CSA Cy

Enantiomeric excess Optical rotation at λ ¼ 589 nm Minutes Seconds 1,4,7,10,13,16-Hexaoxacyclooctadecane Two-dimensional nuclear magnetic resonance 2,20 -Bis(diphenylphosphino)-5,50 ,6,60 ,7,70 ,8,80 -octahydro1,10 -binaphthalene 9-Borabicyclo[3.3.1]nonane Absolute Acetyl Acetylacetonate Azobisisobutyronitrile Aqueous Adenosine-50 -triphosphate Acetonyltriphenylphosphonium bromide 2,6-Di-t-butyl-4-methylphenol Benzyl Based on recovered starting material n-Butyl Benzoyl Cerium ammonium nitrate Catalytic Circular dichroism Cuticular hydrocarbon Correlation spectrometry of long-range coupling Correlation spectrometry Cyclopentadienyl Camphorsulfonic acid Cyclohexyl

136 136 142 197 327 327 327 330 332 332 339 340 344 345

Chemistry of the Secondary Metabolites of Termites

Cyt d DABCO dba DBN DBU DCC DDQ DEAD DET DHF DHP DIAD DIBAH diglyme DMAP DMAPO DME DMF DMP DMSO dppp EDTA EPC eq ESIMS Et ether exc FAB GLC glyme h HFP HMDS HMPA HMQC HPLC HRMS HR-TOF-MS HSQC hν IBX IMDA i-Pr

Cytochrome Day(s) 1,4-Diazabicyclo[2.2.2]octane Dibenzylidene acetone 1,5-Diazabicyclo[4.3.0]non-5-ene 1,8-Diazabicyclo[5.4.0]undec-7-ene 1,3-Dicyclohexylcarbodiimide 2,3-Dichloro-5,6-dicyano-1,4-benzoquinone Diethyl azodicarboxylate Diethyl tartrate 4,5-Dihydrofuran 3,4-Dihydro-2H-pyran Diisopropyl azodicarboxylate Diisobutylaluminum hydride Bis(2-methoxyethyl) ether 4-(Dimethylamino)pyridine 4-(Dimethylamino)pyridine oxide 1,2-Dimethoxyethane Dimethylformamide 2,2-Dimethoxypropane Dimethyl sulfoxide 1,3-Bis(diphenylphosphino)propane Ethylenediamine tetraacetic acid Enantiomerically pure compound Equivalents Electronspray ionization mass spectrum Ethyl Diethyl ether Excess Fast-atom bombardment Gas-liquid chromatography 1,2-Dimethoxyethane (¼dimethylglycol) hour Hexafluoropropan-2-ol Hexamethyldisilazane Hexamethylphosphoramide Heteronuclear multiple quantum coherence High-performance liquid chromatography High-resolution mass spectrum High-resolution time-of-flight mass spectrum Heteronuclear single-quantum correlation Irradiation with light 2-Iodoxybenzoic acid Intramolecular Diels-Alder reaction Isopropyl

3

4

IR IR-120 KHMDS LAH LDA LHMDS LIS MAD MCPBA Me MEM MOM MoOPH Mp MPTACl MS Ms MS NaDPH NBS NCS NMO NMR NOE NOESY ORD PCC PDC Ph PhCH3 PhH Piv PMB PMP PP PPTS pyr RCMT Redal® rfl rt sia TADA TBAF

E. Gössinger

Infrared (spectroscopy) Acidic ion exchange beads Potassium hexamethyldisilazide Lithium aluminum hydride Lithium diisopropylamide Lithium hexamethyldisilazide Lanthanide induced shift Methylaluminum bis-(2,6-di-t-butyl-4-methylphenoxide) m-Chloroperbenzoic acid Methyl Methoxyethoxymethyl Methoxymethyl Oxodiperoxymolybdenum-(pyridine)-(hexamethylphosphoric triamide) Melting point (+)-α-Methoxy-α-(trifluormethyl)-phenylacetylchlorid (Mosher’s reagent) Mass spectrum Methanesulfonyl Molecular sieve Nicotinamide-adenine dinucleotide phosphate N-Bromosuccinimide N-Chlorosuccinimide Morpholine N-oxide Nuclear magnetic resonance (spectrometry) Nuclear Overhauser effect Nuclear Overhauser and exchange spectroscopy Optical rotation dispersion (spectroscopy) Pyridinium chlorochromate Pyridinium dichromate Phenyl Toluene Benzene Pivaloyl p-Methoxybenzyl p-Methoxyphenyl Diphosphate Pyridinium p-toluenesulfonate Pyridine Ring-closing metathesis Sodium bis(2-methoxyethoxy)aluminum hydride Reflux Room temperature 3-Methylbut-2-yl (¼siamyl) Transannular Diels-Alder reaction Tetrabutylammonium fluoride

Chemistry of the Secondary Metabolites of Termites

TBS t-Bu TEMPO t TES Tf (OTf) TFA TFAA thexyl THF TIPS TLC TMS TPAP TPS Tr Troc Ts UV y Z Δ

5

t-Butyldimethylsilyl t-Butyl 2,2,6,6-Tetramethylpiperidine N-oxide tertiary Triethylsilyl Triflate Trifluoroacetic acid Trifluoroacetic anhydride 2,3-Dimethyl-2-butyl Tetrahydrofuran, tetrahydrofuranyl Triisopropylsilyl Thin-layer chromatography Trimethylsilyl Tetrapropyl perruthenate t-Butyldiphenylsilyl Trityl ¼ triphenylmethyl Trichloroethoxycarbonyl Tosyl ¼ p-toluenesulfonyl Ultraviolet (spectroscopy) Year Benzyloxycarbonyl High temperature

1 Introduction Termites (Isoptera), which are a suborder of the cockroaches, are the oldest eusocial insects [1–3]. It is assumed that they diverged from the Cryptocercoidae, woodfeeding cockroaches, in the Late Jurassic, predating the emergence of eusocial Hymenoptera (ants, wasps, bees) by at least 50 million years [1, 4, 5]. In the case of Hymenoptera, the order of insects with the most eusocial species, the evolution toward eusociality can be seen in several extant suborders with species ranking from solitary animals to animals with brood care within small groups up to the highest developed species with colonies of thousands of individuals and at least two castes. Contrary to extant Hymenoptera excepting the suborder of the ants, extant termites are exclusively eusocial, and even the most primitive species consist of more than one caste. The only indication of the evolution toward eusociality is found in comparison with the sister clade of the termites, the wood-feeding cockroaches, Cryptocercoidae. Symbiosis with flagellates containing cellulases allowed the common ancestor of both clades of these hemimetabolous insects feeding on wood. This had dramatic consequences. There was no shortage of food, but the symbionts had to be transferred to the offspring by feeding, which necessitated intense infant care by both parents. Since wood is poor in proteins, a prolonged development to adulthood followed, and offspring of different ages stayed with the parents (most of the symbionts are lost with each molting). It is thought that the splitting of the two

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Plate 1 The last of its kind: Mastotermes darwiniensis, the only extant species of the most primitive termite family. Pseudergates and one soldier are depicted. Photograph: CSIRO, Creative Commons 3.0

clades started with the takeover of the feeding from the parents by the older offspring. The helping behavior in turn permitted more offspring thus reducing the ability of the parents to defend their offspring, which in turn necessitated the development of specialized defenders. Indeed, in the most primitive termites, a soldier caste occurs but no sterile worker caste. The abandonment of the defense by the parents allowed again higher production of offspring necessitating a stable helper caste—the sterile workers. The median size of the population of extant termite colonies is 40,000 individuals reaching from 100 individuals with Cryptotermes piceatus to 7 million with Mastotermes darwiniensis [6] (Plate 1). Nearly 3000 species belonging to ca. 280 genera are known to date, and up to 30 new species are described per annum. These small, soft-bodied, mostly blind insects are found throughout the world in tropical forests, tropical savannas, and semideserts extending to the subtropics [7]. Some of the so-called lower termites were able to colonize even temperate woodlands and temperate rain forests. The highest density in species is found in the northwestern Congolese rain forest. It is thought that termites evolved there. The termites are divided into “lower termites,” which are the evolutionary earlier termites characterized by their flagellate endosymbionts, and “higher termites” including the Termitidae which constitute the largest family with 70% of the termite species. Engel et al. suggested another division of the termites based on extant and extinct termite families [1]. These authors distinguish between the most basal termite family Mastotermitidae, the Euisoptera, termite families that possess no frontal gland, and families possessing a frontal gland, the Neoisoptera [1]. The evolutionary more recent termites (higher termites) have lost the symbiotic protists. In the case of the basal Termitidae, the Macrotermitinae, the flagellates were substituted by ectosymbionts, the basidiomycetous Termitomyces. It is thought that the cultivation of those fungi on combs, built from the feces of the termites, necessitated that these termites had to use next to their own feces soil to build their mounds. This led to the change of the gut symbionts and later on to the ability of the Termitidae to feed on soil. Thus,

Chemistry of the Secondary Metabolites of Termites

7

extant termites feed on soil, wood (dry or damp, decaying), grass, litter, lichens, or the conidia of Termitomyces. Digestion is achieved by a large assembly of symbiotic bacteria, archaea, and, in case of the lower termites, flagellates in the hindgut. Predigestion occurs in the midgut by the termite’s own cellulases, xylanases, and proteolytic enzymes [8, 9]. Often the feces are converted by microbes externally (bacterial or fungal comb growers) and taken up again by the termites (external rumen). The breakdown of cellulose under the anaerobic conditions in the hindgut as formulated by Brune et al. [10] generates the acetate, needed by the termites for nutrition as well as carbon dioxide and hydrogen: ðC6 H12 O6 Þ þ 2H2 O ¼ 2CH3 COO þ 2Hþ þ 2CO2 þ 4H2 Hydrogen in turn is used by the symbiotic archaea for methanogenesis [11]. Since the food is generally poor in proteins, enrichment in nitrogen occurs by nitrogenfixing symbionts [12, 13]. In the case of Macrotermitinae, most of the digestion is transferred to the Termitomyces fungi and in the case of Sphaerotermes sphaerothorax to bacteria [14, 15], which allows the most efficient exploitation of their food including lignocellulose. Their feeding behavior has made around 10% of the species of termites into pests for humans by destroying wooden structures, timber, crops, and fruits [16]. A Chinese proverb even warns that a single termite hole can destroy a 1000 m soil dike. In addition to the damage to man-made structures and crops and the deterioration of rangeland [17], the release of methane into the atmosphere (2–5% of the world’s atmospheric methane [1, 18, 19]), due to cellulolysis, has to be added. These negative effects are more than counterbalanced by the ecological benefit of the activity of termites. The foraging of these detritivores aerates and improves water infiltration of the soil and increases its nitrogen and phosphorus content [20– 22]. Considering that termites constitute a large part of the animal biomass of the tropics and thus dominate (next to ants) the terrestrial ecosystem in the tropics, their ecological benefit is enormous. All termite colonies contain one pair of reproductives, queen and king, most possibly the founding pair. In some species, especially of fungus-growing termites, colony foundation by multiple male and female reproductives (pleometrosis) is known [23]. The caste system varies and is more flexible in the lower termites than in the higher developed termites (Plate 2). Lower termites have no sterile worker caste. The later instars of these hemimetabolous insects are arrested in their development. They are called pseudergates (false workers) and forage, build, feed, and groom parents and siblings and care for the eggs. At a later point in their lifetime, pseudergates may develop into soldiers or alates. As an example of the caste system of the Termitidae, the developmental pathway (Plate 2) of Nasutitermes exitiosus (Plate 3) is described [24]: winged termites (alates) swarm (synchronous nuptial flight) from established mounds. After a partner is chosen and established by shedding the wings and tandem running, the pair seeks a suitable place to burrow their nest. Approximately 60 days after the royal cell is

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Neotenic Reproductives (+ Pseudergate s.l.)

(a) Linear pathway

Pseudergate sensu lato: Larvae Egg

Larvae

Pseudergate s. l.: Nymph

Primary Reproductive

Soldier

(b) Bifurcated pathway Nymphal-Alate Line Egg

Larvae Primary Reproductives

Apterous Line

Workers

Soldier

Plate 2 Social organisation and status of workers in termites. Lower and higher termites differ in their development. The hatched larvae of lower termites develop by (several) molting in totipotent pseudergates (false workers), which may develop into reproductives (either winged queens or kings or neotenics) or into soldiers. In higher termites the hatched larvae are transformed by molting either in nymphs with wingpads and further on to reproductives, or become sterile workers, which may stay as workers or develop into soldiers. Plate courtesy of Roisin Y, Korb J (2011). pp 133–164, chapter 6, in Biology of termites: A modern synthesis. Springer Dordrecht, Heidelberg, London New York. Eds Bignell DE, Roisin Y, Lo N, p 137 (Fig. 6.1)

Chemistry of the Secondary Metabolites of Termites

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Plate 3 At the other end of the evolutionary development: Nasutitermes exitiosus. Their trailfollowing pheromone was the first termite pheromone isolated. True workers, nymphs with wingbuds, and in their midst one “small” soldier, recognizable by its snout. Photograph: Entomology, CSIRO, Creative Commons 3.0

sealed, the first hatchlings appear, which are fed by queen and king until they have molted several times. Thereafter, the offspring begins to forage and feed and tend the royal pair and the successive siblings (up to 3000 per day). Meanwhile, the queen’s abdomen will extend up to 500–1000 times. In being hemimetabolous insects, no metamorphosis occurs, and the hatchlings look like diminutive, colorless adults. Unlike the lower termites, the differentiation of the castes of the higher termites starts with the first molt. In the case of Nasutitermes exitiosus (Plate 3), this leads to so-called nymphs, which are arrested in their development to winged adults, but show wing buds and may develop into supplementary reproductives called neotenics. The main bulk of the second instar is split into smaller and larger instars. After (a) further molt(s), they split into four castes: small sterile workers, small sterile presoldiers, large sterile workers, and large sterile presoldiers. One more molt transforms the presoldiers into soldiers. The development of workers and soldiers is slow; they mature within one year, and their life span is approximately 5 years. The founding royal pair may live up to 25 years. After their death, supplementary reproductives will substitute for them. The most populous castes are those of the sterile workers. The soldiers differ in a more pronounced manner from the workers and nymphs. Their heads are enlarged, pigmented, and highly sclerotized. Whereas the workers are engaged in foraging; tending to the royal pair, the younger siblings, and the eggs; feeding soldiers; building and restoring the nest (mound) and tubes or tunnels; and partly defending, the smaller soldier caste is involved in pioneering, guarding, and defending the mound and foraging or mound- and tube-building workers. The task of the larger soldiers, which usually do not leave the nest, seems to constitute the protection of the brood and the royal pair. Termite colonies are very structured, and the caste proportion is restored rapidly after disturbance.

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These small, mostly blind insects are very vulnerable in being not or only barely sclerotized. Thus, they are easy prey even to small predators. For example, dispersing queens, which are better protected than all other castes, because they are more sclerotized and possess defensive substances at this stage [25], survive to less than 1% to found a new colony [1]. Due to their limited sclerotization, termites are very sensitive to temperature variations, light, and desiccation. Thus, to survive they need protection and fortification. The most important way of protection is through their nests. Abe distinguished between one-piece nesters, which live within wood and thus never have to leave their nests to forage; a small group of intermediate nesters, for example, the most basal termites Mastotermes darwiniensis (Plate 1); and separatepiece nesters, where nest and food source are separate. Most primitive termites are wood feeders, and thus they live within wood and are protected by the wood. With the development of a true worker caste and other food than wood, the nestings became more complex, spanning from rather diffuse subgeal nests with long subterraneous galleries for foraging to nests that reach high above the ground, called mounds, or carton nests in trees, connected by tunnels (tubes) with the foraging sources. Only a few species of the separate-piece nesters have developed the ability to forage without the protection of mud tunnels in the open air [26]. The most elaborate mounds are found with the fungus-cultivating Macrotermitinae. These structures, built mostly from the feces of termites, contain up to several millions of termites and their fungus combs. The mounds are separated into different chambers with thermoregulation around 30 C, constant humidity, and gas exchange and are the most complex known architectural features in the animal kingdom. Despite their admirable fortifications [27], termites have to defend themselves against many predators, especially against ants. This is the task of the soldier caste. As mentioned above, the heads of soldiers are highly sclerotized, and thus they are able to physically defend the colony. In some species, soldiers plug entry holes or small galleries with their body. In most other species, the physical defense is accomplished by the enlarged sclerotized mandibles. Gradually to this physical defense, chemical defense was added until in the highest developed termite species, defense by soldiers relies entirely on chemical weapons. Mostly from one exocrine gland, toxins, irritants, antihealants, and sticky substances are released. The diversity of compounds isolated from these glands of different families, genera, and species is enormous. Several reviews have dealt with the physical and chemical defense of termites in the past [28–32]. The chemistry of these secondary metabolites is part of the present contribution. Living in colonies of up to a few million individuals, infections are a high risk especially due to the easily penetrable cuticle of termites and to inbreeding via neotenics. Defense against microbes is achieved by the termites’ innate immune system, by antimicrobial peptides [33], and partly by antibiotically active terpenoids of the defensive secretion [34]. Against multicellular parasites, encapsulation is used. Diseased nestmates are either cannibalized or sealed off [35, 36], and cadavers are buried under the soil [37, 38]. Eusociality is defined by three traits: cooperation in caring for the young, reproductive division of labor with more or less sterile individuals working on behalf of individuals engaged in reproduction, and overlap of at least two generations of life

Chemistry of the Secondary Metabolites of Termites

11

stages capable of contributing to colony labor [39]. Advanced eusociality is defined as a superorganism, in which interindividual conflict for reproductive privilege is diminished and the worker caste is selected to maximize colony efficiency in intercolony competition [40, 41]. In such a superorganism as a termite colony, communication is all-important. Most termites are, with the exception of the alates, blind and live subterranean; thus communication seems to be restricted to vibration, touching with antennas, body contact, smell (olfactory), as well as taste (contact chemoreception). Therefore, communication is mostly conducted by chemicals. Wilson distinguished within the intraspecific chemical communication between pheromones that trigger an immediate behavioral response—the releaser pheromones—and pheromones that initiate long-lasting physiological changes, the primer pheromones. One surprising fact of the chemical communication system of termites is the parsimony of chemicals used. Pheromonal parsimony, or the use of one compound (pheromone) for different interactions, may have been a driving force in expanding the horizons of eusociality as Blum suggests [42]. The cryptic lifestyle of termites, their smallness, the difficulty of rearing these animals in the laboratory as well as the semiochemical parsimony have made the search, isolation, and structure determination of the compounds secreted in picograms from the relatively few (exocrine) glands and the designation of the biological activity very difficult. Despite these disadvantages, contemporary isolation techniques, especially solid-phase microextraction (SPME) [43–45], modern spectrometric methods, and synthesis have permitted the identification of volatile pheromones of more than 100 species. The chemistry of the pheromones of termites is a further part of this contribution. The small size of the termites, their cryptic lifestyle, their feeding behavior, and the difficulty in maintaining termites in the laboratory have impeded the study of the biochemical pathways of their secondary metabolites particularly. Relevant information unraveled completes the present contribution.

2 Pheromones Eusocial insect organizations are based on communication and information transfer with semiochemicals being at the center [46]. It is tempting to look for pheromones of termites by analogy to those of the much more investigated pheromones of the hymenopterans, but caution is necessary due to the large evolutionary distance between the homometabolic hymenopterans and the heterometabolic Blattodea. Whereas the ants, which resemble superficially the termites, being also, with the exception of the reproductive caste, flightless, eusocial insects, have a multitude of exocrine glands (>60), morphological data reveal the presence of 20 different exocrine structures in termites [47] (and literature cited therein, [48, 49]). The scarcity of glands necessitates the use of a relatively small number of semiochemicals for more than one interaction. This so-called pheromonal parsimony [42] as well as the fact that these mostly subterranean living creatures are, with the exception of the alates, blind, thus missing an important sense most hymenopterans

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use extensively, raises the question as to how is it possible that termites are able to maintain ordered communities with up to several million members. Their communication seems to be reduced to vibration [50–55], contact (antennation), smell (olfactory), and taste (contact chemoreception). The investigation of the pheromones of the termites is hampered due to the difficulties in rearing the animals in the laboratory and thus to observe and investigate the behavior of such subterranean, light-sensitive organisms. To these impediments are added the small size of the animals themselves and thus of their glands, which secrete the semiochemicals in picogram quantities. Also, the chemical components of these small and few glands have to perform a multitude of semiochemical tasks. Depending on concentration and in combination with further components of the same or other glands, different activities may be initiated. Modern isolation techniques like solid-phase microextraction (SPME) [43–45], electroantennograms (EAG) [56–59], spectrometric methods, and synthesis have permitted the identification of volatile pheromones of ca. 100 species. In contrast, the contact pheromones, with the exception of the cuticular hydrocarbons, are mostly unknown [60]. The structure and activity of primer pheromones of termites and thus of caste differentiation and developmental retardation are with few exceptions left to speculation. Of the 20 distinct exocrine glands known to date, the contents and function(s) of their secretion are known only in part for six glands: (a) The labial or salivary gland [61], which exists in all species and castes. Labial gland secretions serve various functions during nest construction, colony defense (especially in the basal termites that possess no frontal glands (Euisoptera and Mastotermes darwiniensis (Plate 1)), colony hygiene, and aggregation at gnawing sides. (b) The sternal gland is found in all castes and in most termite families. It secretes the trail-following pheromone, which in many families is also a sexual pheromone. (c) The posterior sternal gland delivers the sex pheromone in Macrotermitinae [62]. (d) The tergal gland is found in the last tergites of alates and secretes sexual pheromones. In 2012, Costa-Leonardo detected tergal glands also in the soldiers of the subfamily Syntermitinae [63, 64]. (e) The frontal gland, the best investigated exocrine gland in termites, is found in the head of all Neoisoptera ( Termitidae, Serritermitidae, and Rhinotermitidae) as an unpaired gland. It varies considerably between families and genera, and it is well developed only in soldiers. In most taxa this gland is restricted to the head; in Rhinotermitidae it extends into the abdomen. Its contents, irritants, glues, or contact poisons are secreted from a frontal pore, the fontanelle. The secretion serves the defense but may have other functions as well. For example, it may regulate caste determination, has fungistatic and bacteriostatic activities, and induces alarm and aggregation. Of the other glands very little is known; the contents of the secretion as well as the activities these secretions initiate are not known as yet. For example, the

Chemistry of the Secondary Metabolites of Termites

13

recently discovered clypeal gland is only found in the reproductive caste and may contain queen or king pheromones [49]. (f) The cuticle, which some researchers see as the largest exocrine gland, contains a mixture of chemicals. The hydrocarbons are thought to effect nestmate recognition and protection against dehydration. The differences in the hydrocarbons (straight-chain and methyl-substituted hydrocarbons) within the colony may influence caste differentiation. The cuticle also contains antibiotics and proteinaceous compounds; the function of these compounds is still unknown.

2.1

Releaser Pheromones

Termites have few, highly conserved releaser pheromones used to determine a certain behavior, as, e.g., for trail-laying, sex-pairing, home-marking (?), foodmarking, phage stimulation, aggregation, alarm, worker-arresting, egg-recognition, etc. These are found in most termite families.

2.1.1

Trail-Following Pheromones

The trail-laying behavior of termites [62–76] has been described several times since 1911 [77–79] (and literature cited therein). However, the source and nature of the secretion remained unknown until Lüscher and Müller investigated the trail-laying behavior of Zootermopsis nevadensis [77]. By partially covering ventral body parts of the insects, they found the sternal gland as the source of the biologically active secretion. Further they determined that the active compounds are soluble in ether. Setting the animals on a wire gauze above an artificial trail of sternal gland secretion initiated trail-following, thus demonstrating the volatile nature of the secretion. For the mostly blind termites, chemical communication when foraging is essential. This is especially true for separate-piece nesters. However, trail-laying pheromones are also found in one-piece nesters, probably to guide nestmates to breaches in the nest or to mark their nest. Some termite species use additional cues for guidance. Hodotermitinae have compound eyes and thus are the only termite workers able to use optical signs [80]. Rickli and Leuthold demonstrated that Trinervitermes geminatus uses magnetic guidance [81]. Evans et al. list several species that support the chemical cues by vibrational signals [51–53]. Pheromone trails may well be templates of the construction of tunnels and galleries [27]. Kaib et al. described the search for food in the subterranean termite Reticulitermes flavipes (formerly R. santonensis) [82, 83]. A few pioneer workers explore the territory in every direction laying a dotted trail; when food is discovered, they return to the nest laying a continuous trail. This trail is followed within seconds by other workers. Soldiers are less sensitive and thus follow when a larger group of workers explores the trail laid. Consequently, galleries or tunnels are built for the main group of foragers. The chemistry and biology of trail-following communication have been described for around 70 species from all families of termites [69]. So far only nine

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O

OH 1 ((3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol)

6 ((4R*,6S*)-dimethyldodecanal)

OH 7 ((4R*,6S*)-dimethylundecanol)

2 ((–)-(R)-neocembrene)

OH 3 ((3Z,6Z)-dodeca-3,6-dien-1-ol) 8 (trinervitatriene) OH 4 ((3Z)-dodec-3-en-1-ol)

OH

O 9 ((10Z,13Z)-nonadeca-10,13-dien-2-one)

5 ((E)-2,6,10-trimethylundeca-5,9-dien-1-ol)

Scheme 1 Trail-following pheromones of termites

active compounds are known [72, 84]. Most likely these pheromones are derived from the sex pheromones of the alates [85]. The first structure determined of a trail-following pheromone was (3Z,6Z,8E)dodeca-3,6,8-trien-1-ol (1) (Scheme 1), which is found in most families of termites [70, 72, 73, 75, 84, 86–88]. For this unsaturated primary alcohol, the threshold for trail-following in Rhinotermes flavipes is 0.01 pg/cm trail, whereas stereoisomers and derivatives are only slightly active, more than seven orders of magnitude less than 1 [87, 88]. Later on, 1 was detected as the general trail-following pheromone of the Reticulitermitinae and found in other genera of Rhinotermitidae also. Saran et al. found for Reticulitermes hesperus that the pheromone stimulus lasted around 48 h and that an unbelievable 0.05 fg/cm of 1 stimulated the termites [66]! In the secretion of the sternal gland of Coptotermes formosanus (Rhinotermitidae), 1 is the main compound accompanied by small amounts of the isomeric (3Z,6E,8E)-dodeca-3,6,8trien-1-ol, but its function is unknown as yet [89, 90]. Compound 1 is thought to be the general pheromone for orientation, whereas recruitment is initiated by speciesspecific pheromones. The monocyclic diterpene neocembrene (cembrene A, (E)-6-cembrene, (1E,5E,9E,12R)-1,5,9-trimethyl-12-(1-methylethenyl)cyclotetradeca-1,5,9-triene) (2) was the first termite pheromone isolated. In this contribution the name neocembrene is used and the numbering follows the one introduced by Moore: (1R,3E,7E,11E)-3,7,11-trimethyl-1-(1-methylethenyl)cyclotetradeca-3,7,11-triene. This numbering was used consistently not only for neocembrene itself but also for the polycyclic diterpenes derived from neocembrene up to the most recent

Chemistry of the Secondary Metabolites of Termites

15

syntheses of the tetracyclic diterpenes of termites. Moore chose an abundant Australian Nasutitermes species for his investigation and demonstrated that the isolated pure compound was a trail-following pheromone [91, 92]. Much later, 2 was also identified as trail pheromone of the lower termite subfamily, Prorhinotermitinae [93]. The electroantennogram of the gas chromatogram (GC/EAD) of the sternal gland secretion showed next to neocembrene (2) a small peak at the retention time of (3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol (1), which could explain the difference in attractivity of pure neocembrene and the sternal gland extracts for Prorhinotermes sp. [94]. More than 30 years passed until the next trail-following pheromones were characterized. (Z,Z,E)-Dodeca-3,6,8-trien-1-ol (1), (3Z,6Z )-dodeca-3,6-dien-1-ol (3), and (3Z)-dodec-3-en-1-ol (4) are the respective main compounds of the sternal glands of two Macrotermitinae [95, 96]. (3Z )-Dodec-3-en-1-ol (4) was found in several more Macrotermitinae shortly afterward [97]. Later on, 4 was found as the main trailfollowing pheromone of the family of the Euisopteran Kalotermitidae [98]. In the last two decades, investigation of lower termites led to the characterization of the norsesquiterpene alcohol (E)-2,6,10-trimethylundeca-5,9-dien-1-ol (5) as a trailfollowing pheromone (Mastotermes darwiniensis (Plate 1)) [99], syn-4,6dimethyldodecanal (6) as the main trail-following pheromone of Zootermopsis species (Archotermopsidae) [100, 101], and syn-4,6-dimethylundecan-1-ol (7) as the main trail-following pheromone of Hodotermopsis sjöstedti (Archotermopsidae) [102]. Although many termite species follow trails scented with 1, the mixture of compounds secreted from the sternal gland is species specific. The explanation for this phenomenon is that 1 is the general orientation signal that is supplemented by species-specific components. Possibly cuticular hydrocarbons (CHCs), which are not only species specific but also colony specific, enhance the attractivity of the trail pheromone for nestmates. One further cyclic diterpene, the tricyclic (11E)-trinervita1(14),2,11-triene (8), was detected in glands of a Nasutitermes species, and it is assumed to have trail-following activity [103, 104]. Recently, a new unusual trailfollowing pheromone, (10Z,13Z )-nonadeca-10,13-dienone (9), was isolated from the sternal gland of Glossotermes oculatus (Serritermitidae) [69]. This compound has a surprisingly high boiling point although it should be active only for a short time. (10Z,13Z)-Nonadeca-10,13-dienone (9) has not been detected in any other termite family. Only this single compound was detected and confirmed by synthesis [69]. The threshold for detection of trail-following pheromones depends on caste and age. Depending on the species, older workers or soldiers are the pioneers and have the lowest threshold for the trail-following pheromone. After food has been detected, the pheromone trail is reinforced, which leads to mass recruitment. Wen et al. were able to demonstrate that the ratio of compounds 3 and 4 varied depending on the behavior of the forager [71]. The open-field foraging Macrotermitinae Odontotermes formosanus was chosen, and the ratio of the trail pheromones was determined [71]. These examinations revealed that 4 induced exclusively orientation, whereas 3 is responsible for orientation as well as recruitment. The fact that the ratio of pheromone components is variable depending on searching for food or recruiting was detected in eusocial insects for the first time.

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Isolation and Structure Determination of Trail-Following Pheromones A few years after Butenandt et al. isolated and characterized the first insect pheromone bombycol ((10E,12Z )-hexadeca-10,12-dien-1-ol) [105–107], Moore published the isolation of the first (trail-following) pheromone of a termite [91]. Moore’s task was even more difficult than Butenandt’s. The rearing of the domesticated silkworm Bombyx mori was relatively easy, and the larger size of the insect yielded larger amounts of secretion from its glands. In contrast, Moore had to deal with small, cryptic living, wild animals. Thus, capture and rearing of the animals as well as the isolation, structure determination, and bioassays conducted required much more effort. Moore chose the abundant, relatively easy to rear Australian termite Nasutitermes exitiosus (Termitidae, Plate 3) for his investigations. He started his research by extracting whole animals in alcohol and after evaporation of the solvent heated the residue to higher temperatures. This procedure led to the isolation of ellagic acid and isomers, which may well be part of the digested food of the termite workers [108]. In his next attempt, the termites were homogenized in ice water with 0.1% quinol as antioxidant. Steam distillation resulted in volatile, oily monoterpenes (α-pinene, β-pinene, and β-phellandrene) [109]. To detect the so far evasive trail-following pheromone(s), he then extracted homogenized termites in light petroleum [91]. The extract was treated with aqueous base, and the unsaponifiable fraction was chromatographed on alumina. The active fraction was vacuum distilled with neicosane as carrier and chromatographed subsequently on silica gel. The two compounds obtained were separated by preparative GC. The molecular weight as well as the fragmentation pattern of the mass spectra of the pure compound pointed to a diterpene. The structure was narrowed down by means of IR and UV spectra and microhydrogenation followed by mass spectrometry to a monocyclic diterpene with four isolated double bonds, but the amount of the pure compound was insufficient for a complete structure elucidation. Moore continued the structure elucidation by developing a microozonization method [110] and by looking for more abundant sources of the compound in plants guided by bioassays. The ozonization products (Scheme 2) and the 1H NMR spectra of 2 pointed to one disubstituted and three trisubstituted double bonds. These data led the authors to the erroneous assumption that the compound is a substituted cyclohexene. To prove this assumption, the possible substituted cyclohexenes were synthesized by methods developed for carotene syntheses [92] (Scheme 3). None of the synthesized compounds showed significant trail-following in termites. Therefore, guided by biosynthesis reasoning, the authors concluded that geranylgeranyl pyrophosphate, the open-chain precursor of cyclic diterpenes, closes to the 14-membered ring of the cembrenes [111–116]. However, the compound was

Chemistry of the Secondary Metabolites of Termites

O3, AcOEt

17

O

O 2

O

H O

+

+

O H

O

2

Scheme 2 Ozonization of neocembrene (2)

1)

O

O

O

PPh3

O

CH3S(O)CH3,

O

NaH, 45°C; rt 2) TsOH, (CH3)2CO, rt

O

O 1)

5:1

O

PPh3

2) TsOH, (CH3)2CO, rt 3) CH2=PPh3

CH3S(O)CH3, NaH, 45°C; rt

+

1)

O

PPh3

CH3S(O)CH3, NaH, 45°C; rt

2) TsOH, (CH3)2CO, rt 3) CH2=PPh3

1

+

:

+

2

Scheme 3 Synthesis of substituted cyclohexenes

not identical with cembrene [117]. An improved isolation from Nasutitermes exitiosus (Plate 3) permitted further experiments. Isomerization of the purified trail-following pheromone with strong base and subsequent microozonization pointed to the structure of neocembrene (2) (Scheme 4). The structure determination was corroborated by comparison with a plant product isolated and characterized at that time [118]. Although the positions of the double bonds were ascertained, their configuration had to be determined by

18

E. Gössinger - + CH3S(O)CH2 Na , CH3S(O)CH3

2

isoneocembrene

O3, AcOEt

O

O 2

O

+

O

+

O O

Scheme 4 Isomerization of neocembrene (2) and subsequent degradation

synthesis [119–122]. Six years and the cooperation of Australian and Japanese researchers [92, 123] were necessary to characterize the monocyclic diterpene as neocembrene (2). Resolution of synthetic racemic neocembrene indicated that both enantiomers are biologically active [124]. Kato, Moore et al. by synthesizing two additional stereoisomers of neocembrene demonstrated that only neocembrene (2), with all double bonds (E)-configured, showed trail-following activity in N. exitiosus [124]. Traniello et al. found that 2 effects only orientation in termites (N. corniger ¼ N. costalis) but no recruitment and it is not as active as the secretion of the sternal gland [125]. Indeed, in more recent years, several examples of two-component trail-following pheromones with neocembrene (2) as one of the components have been described [93, 94]. Since Moore’s first isolation of neocembrene (2), this monocyclic diterpene has been isolated as trail-following pheromone from Nasutitermes species [104, 117, 125, 126] but, surprisingly, also from a subfamily of lower termites, the Prorhinotermitinae [93, 94]. Neocembrene was perceived as sex pheromone of some Nasutitermitinae [103], and it was found in the frontal glands of soldiers of some species of the Cubitermitinae [127, 128]. Prestwich was able to gather enough neocembrene from the defensive secretion of Cubitermes glebae to determine its absolute configuration as ()-(R)-neocembrene (2) [128]. The same absolute configuration of neocembrene (2) and of its polycyclic derivatives was found in the secretion of all investigated termite species independent of family, caste, or gland. As mentioned above, neocembrene was isolated from plants [118, 129] but also from animals (soft corals [130], Pharaoh’s ant [131], and the Chinese alligator [132]). While Moore, Kato et al. were involved in the structure determination of neocembrene (2), Coppel et al. investigated the trail pheromone(s) of a termite species of the family Rhinotermitidae [86, 87]. The observation that wood decayed

Chemistry of the Secondary Metabolites of Termites

19

by the fungus Lenzites trabea (Gloephyllum trabeum) attracts termites [88] initiated Coppel’s search for the trail-following pheromone of Reticulitermes virginicus, a species especially attracted. For the isolation 385 g of termites and 15 kg of decaying wood were homogenized, extracted with n-pentane, and after evaporation codistilled with mineral oil. Chromatography on Florisil and further separation by preparative GC led to pure compounds. The respective compounds obtained from both sources had the same Rf value. Comparison of the spectra of the 100 μg pure compound derived from the decaying wood with the imperfect spectra of the 1 μg of pure compound derived from the termites showed no deviations. The bioassays corresponded qualitatively and quantitatively. Thus, the identity of the two compounds was accepted. The absorption at 234 μm in the UV spectrum pointed to a conjugated diene. The 1H NMR spectrum revealed signals corresponding to one methyl group; three methylene groups, two of them allylic methylene groups and one diallylic methylene group; and a primary alcohol group (Table 1). The mass spectrum with the molecular peak at 180 mu and the IR spectrum confirmed the primary alcohol group. Thus, the researchers concluded that the compound is a ndodecatrien-1-ol. Microhydrogenation produced the known n-dodecan-1-ol. The most likely structure fitting those data was a dodecatrienol with Δ3,4- and Δ8,9double bonds. The position of the third double bond as well as the configuration of the double bonds was determined by synthesis of several isomers of dodecatrienol. Bioassays showed that the most active compound (>0.1 pg for 10 cm trail) was (3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol (1), which has identical spectra to the natural

Table 1 NMR spectrometric data of the abundant trail-following pheromone (3Z,6Z,8E)-dodeca3,6,8-trien-1-ol (1) Ref. [133] 13

Structure, Nr. OH (3Z,6Z,8E)-Dodecatrien-1-ol (1)

Position 1 2 3 4 5 6 7 8 9 10 11 12 OH

H NMR (CDCl3, δ/ppm, J/Hz) 3.65 q J1,2 ¼ 6 2.37 dt J2,3 ¼ 7, J2,1 ¼ 6 5.41 dtt J3,4 ¼ 11, J3,2 ¼ 7, J3,5 ¼ 1 5.56 dtt J4,3 ¼ 11, J4,5 ¼ 7, J4,2 ¼ 1 2.95 t J5,4 ¼ J5,6 ¼ 7 5.25 br dt J6,7 ¼ 11, J6,5 ¼ 7 5.98 br t J7,6 ¼ J7,8 ¼ 11 6.32 ddq J8,9 ¼ 15, J8,7 ¼ 11, J8,10 ¼ J8,6 ¼ 1 5.69 dt J9,8 ¼ 15, J9,10 ¼ 7 2.09 q J10,9 ¼ J10,11 ¼ 7 1.42 sext J11,10 ¼ J11,12 ¼ 7 0.91 t J12,11 ¼ 7 1.68 t JOH,1 ¼ 6 1

C NMR (δ/ppm) 62.2 30.8 125.7 131 26.1 127.1 129.0 125.4 135.3 34.9 22.5 13.8

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product. Thus, (3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol (1) was determined as the first known structure of a termite (trail-following) pheromone. Despite the rapid improvement of separation techniques and spectrometric methods in the following 30 years and the knowledge of the lipophilic nature of these pheromones, only 1 was isolated from several termite species in more than one family [70, 74, 76, 89, 90, 134–136]. As an example of the search for trail-following pheromones at that time, Tokoro’s investigation of Coptotermes formosanus may be described [89, 90]. Around 200,000 pseudergates (561 g), reared on red pine in a darkened laboratory at 28 C and 80% humidity for 5 years, were separated in 10 portions and fed with filter paper for 5 days to substitute for intestinal material and prevent the isolation of dietary materials. Then the termites were soaked in nhexane for 3 days. After filtration the portions were combined, and the solvent evaporated yielding 51 g material. Fractionating by chromatography on silica gel with n-hexane/ethyl acetate mixtures led to 350 mg active material, which was further purified on silver nitrate-impregnated silica gel. Two further separation steps consisted of HPLC and GC on a nonpolar and then on a polar column. Each fraction was tested by the following simple bioassay: the dissolved fraction was streaked along a cycle (d ¼ 4.7 cm) on filter paper in a Petri dish covered with a red-colored lid. When at least a third part of the test animals stayed on the cycle for more than 2 min, “basic activity” was assumed. With these fractions the activity was further investigated by dilution steps. The pure active compound was analyzed by capillary GC combined with MS and FTIR [134]. A small molecular peak and several characteristic fragmentation ions, the UV spectrum, and several microreactions revealed the structure as identical with 1. This was confirmed by comparison of the spectra with the spectra of synthetic 1, those of synthetic stereoisomers, and those of synthetic (3Z,6Z )-dodeca-3,6-dien-1-ol (3). In 1990, Pawliszyn introduced “solid-phase microextraction” (SPME) as a new isolation technique [43], which allowed the extraction of small body parts without the use of solvents [44, 45, 137]. Primarily this method, along with the introduction of excision of body parts (e.g., glands) with microscissors under a stereomicroscope, the detection of bioactivity by electroantennogram (EAG), as well as the use of computer-assisted statistical analyses has facilitated and accelerated the discovery of new trail-following pheromones and led to the confirmation that in several termite species, more than one substance is involved in trail-following. In 2001, an additional trail-following substance was characterized as (Z )-dodec3-en-1-ol (4) [95]. Bordereau et al. chose the fungus-growing termite Macrotermes annandalei not only to employ these new methods but also to compare them with the established techniques. Whole bodies of workers or excised glands were immersed in solvents. No significant differences in the biological activity were detected when changing the solvent (n-pentane, n-hexane, or dichloromethane) or between extracts of the whole animals and of the sternal glands. The extraction by SPME was carried out under a stereomicroscope by rubbing the surface of the exposed sternal gland with a polydimethylsiloxane/divinylbenzene fiber, which then was inserted in the injection port of a gas chromatograph. The n-hexane extracts of whole animals as well as of excised glands were purified by chromatography on silica gel, and the

Chemistry of the Secondary Metabolites of Termites

21

fractions were tested by bioassays. On the filter paper a Y was drawn. At the stem and one branch of the Y, a small part of a fraction was drawn with a syringe, and at the other branch, pure solvent was drawn. The time and distance the test animal traveled were measured. The active fractions were further purified by preparative GC on capillary columns. Characterization was carried out by GC/MS. Electron-impact as well as chemical-ionization mass spectra were investigated. The peaks of the capillary gas chromatogram were compared with standardized retention indices [138–141]. The mass spectrometric data revealed a dodecen-1-ol. The position of the double bond was elucidated by the addition of dimethyl disulfide to the unsaturated primary alcohol and analysis of the fragmentation patterns of the mass spectra of the obtained adduct. The stereochemistry of the double bond was established by synthesis [95]. Comparison of the gas chromatogram of an extract of the sternal gland surface and an intertergal membrane surface by SPME showed that four compounds were specific to the gland surface including (3Z )-dodec-3-en-1-ol (4). In a similar way, the sternal gland extract of another Macrotermitinae, Ancistrotermes pakistanicus, and SPME of the sternal gland surface permitted the characterization of (3Z,6Z )-dodeca-3,6-dien-1-ol (3) [96], which was confirmed by synthesis. The use of electroantennography (EAG) [56] coupled with gas chromatography (EAD) [57] allowed the detection of a two-component trail-following pheromone for the first time [94]. The trail-following pheromone of Prorhinotermes simplex had been investigated earlier [93], and neocembrene (2) was the single substance that was bioactive. Even by the more sensitive detection technique of two-dimensional gas chromatography coupled with time-of-flight mass spectrometry (GCxGC/TOFMS), no other bioactive gland-specific compound could be detected. Using EAD revealed a second bioactive substance in the extract of the sternal gland of pseudergates of P. simplex. The retention indices and MS fragmentation pattern pointed to (3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol (1) as a second active component. Experiments with neocembrene (2) and synthetic 1 showed that Prorhinotermes simplex preferred a mixture of 2 to 1 (101 ng/cm:108 ng/cm) to pure 2 and even to a trail made of sternal gland extract. In the same manner, the trail-following pheromone of several Nasutitermitinae was investigated and found to consist of neocembrene (2) as main compound and dodecatrienol 1 as minor component [104]. In several of the Nasutitermitinae investigated, trinervitatriene 8, found earlier as a sex pheromone [103], supplemented the gland extracts. The short lifetime of laboratory-reared Nasutitermitinae prevented the delineation of the function of this diterpene [104]. Basal termites possess sternal glands and are able to follow sternal gland extracts although the function of the trail pheromones of these one-piece nesters is disputed. More recently the structures of trail-following pheromones of basal termites have been elucidated [99, 101, 102]. The evolutionary oldest extant termite species Mastotermes darwiniensis (Mastotermitidae, Plate 1) is not a one-piece nester and possesses contrary to all other termites not one but three sternal glands. Whole pseudergates and their excised sternal glands were immersed in solvents differing in polarity. Additionally, a

22

E. Gössinger

polydimethylsiloxane/divinylbenzene fiber was applied for the SPME by rubbing the exposed sternal glands. Extracts of sternal glands were fractioned by preparative GC and the fractions tested in trail-following bioassays. The same isolation techniques were employed for the isolation of the trail-following pheromone(s) of the basal one-piece nesters Porotermes adamsoni and Stolotermes victoriensis. Comparing the GC/MS of the SPME of all three species revealed that they used a common trail-following pheromone. The accumulated SPME extracts of two of the species were fractioned by preparative GC. Surprisingly, the mass spectra of the bioactive compound suggested a terpenoid structure. The authors concluded, based on the retention indices, the molecular peak, and the fragmentation pattern, that the compound is the norsesquiterpene, 2,6,10-trimethylundeca-5,9-dien-1-ol (5). Syntheses of both possible stereoisomers established not only the suggested structure but also its stereochemistry as (E)-2,6,10-trimethyl-5,9-undecadien-1-ol (5). The same techniques (SPME, GC/MS with electron ionization as well as chemical ionization, retention indices, FTIR, “T-maze” bioassay, and syntheses) were employed for the search of the trail and sex pheromones of the basal termites exemplified by two Zootermopsis species [101] and one Hodotermopsinae [102]. Infrared spectra, the molecular peak, and the fragmentation pattern in the mass spectra sufficed to determine that pseudergates of Hodotermopsis sjöstedti use (4R*,6S*)-dimethylundecan-1-ol (7) as trail-following pheromone. The stereochemistry was determined by synthesis [102]. In the same way, the trail pheromone of the pseudergates of Zootermopsis nevadensis and Z. angusticollis was characterized as (4R*,6S*)-4,6-dimethyldodecanal (6) [101]. After isolation and characterization of the trail-following pheromones of around 60 species and finding only few bioactive hydrocarbons and primary alcohols of moderate boiling point over a period of nearly 60 years, it was surprising when the sternal glands of a species of the little-investigated family Serritermitidae yielded an unsaturated long-chain methyl ketone. The excised glands of Glossotermes oculatus were extracted in n-hexane, and the complex mixture of compounds was fractionated by chromatography on silica gel [69]. The bioactive fractions were further separated by preparative GC. Two-dimensional GC coupled with TOF-MS was used to characterize the substance as nonadecadien-2-one (9). The positions of the double bonds of 9 were confirmed (Scheme 5) by the fragmentation pattern in the mass spectra of the dimethyl disulfide adducts 10. This method was developed by Vincenti et al. for linear nonconjugated dienes [142, 143]. Iodine converted dimethyl disulfide into the methylthioiodide, which added to the double bond forming the thiiranium salt i. The generated iodide attacked a further dimethyl disulfide. The formed methylthiolate attacked the charged thioepoxide of i, and methylthioiodide formed the next charged thioepoxide ii. Intramolecular attack of the sulfide opened the thioepoxide by simultaneous formation of charged thietane. According to Vincenti et al., no five-membered ring was observed. Iodide transformed iii into 10. Electron ionization occurs preferentially at the electron-rich sulfur atoms, thus yielding characteristic fragments. The stereochemistry of the double bonds was determined by the facile syntheses of 9.

Chemistry of the Secondary Metabolites of Termites

23 S

(CH3S)2, CH3(CH2)3

CH3(CH2)3

(CH2)6COCH3 I2, ether, rt

9

(CH2)6COCH3 S

S 10

CH3SI

S

(CH2)6COCH3

CH3(CH2)3 S

CH3(CH2)3

+ CH3SSCH3

I

I (CH2)6COCH3 S

S i

iii CH3SI I S CH3(CH2)3

(CH2)6COCH3 S

S ii

Scheme 5 Addition of dimethyl disulfide and iodine to skipped double bonds

The much-improved analytical techniques allowed Wen, Ji, and Sillam-Dusses to demonstrate with Odontotermes formosanus (Macrotermitinae) the dependence of the trail pheromone composition on caste as well as on behavioral context within a caste [71]. Solid-phase microextraction-GC, SPME-GC/MS, GC/EAD, retention indices, synthetic reference compounds, Y-shaped trail-following bioassays, the recorded video files, and statistical analysis showed that workers use two components, 3 and 4, as trail pheromones, whereas soldiers use only one component, namely, 4. The proportion of the two components in the secretion of the sternal gland of workers depends on the activity of the animal. Heavy work, such as brood care and wood gnawing, needs a higher proportion of 3 than trail-laying or construction of fungus gardens. Older termite workers having the largest sternal glands serve as the pioneers. A hungry worker is able to follow a trail made by 1 fg/cm! The authors found no synergistic effects between the two pheromone components. The bioassays indicated that 4 is responsible for orientation and 3 for orientation and recruitment. Trail-following activity increases with the number of workers having laid a trail, but longer exposure to 3 makes the antenna of workers insensitive and leads to chaotic behavior at the food side.

2.1.2

Sex-Pairing Pheromones

Although most of the behavior of termites [62–64] is initiated by chemical stimuli, present knowledge of sexual pheromones is still scanty. Due to the cryptic lifestyle

24

E. Gössinger

Plate 4 Pheromones and chemical ecology of dispersal and foraging in termites. (a)–(f) Calling females exposing their tergal and/or sternal glands to lure males with their sexual pheromones. (g) shows a female between callings. (h)–(j) After meeting, male and female plug their wings and decide by tandem running, with the male antennating and licking, if they fit together and look for a suitable place to found a new colony. Plate courtesy of Bordereau C, Pasteels JM (2011) Biology of termites: a modern synthesis, eds Bignell DE, Roisin Y, Lo N. Springer Dordrecht, Heidelberg, London New York, pp 279–320, chapter 11, p 282 (Fig. 11.1)

Chemistry of the Secondary Metabolites of Termites

25

of these animals and the fact that synchronous swarming occurs once a year or even less, the search for sexual pheromones is extremely difficult. Thus, only 8 compounds have been characterized from 30 species [144, 145]. In many species only one compound seems to be responsible for attracting a sexual partner. However, this does not explain why in several cases investigated, nestmates as well as related species with the same sex-pairing pheromone are not accepted as sexual partners. In many species the sex-pairing pheromone is identical with the trail-following pheromone, but is secreted in much larger quantities. It is suspected that further compounds act synergistically. Swarming seldom exceeds a few hundred meters. The landed imagoes show calling behavior. In Euisoptera females as well as males are involved in signaling, whereas in Neoisoptera only the females raise their abdomen to expose the sternal, posterior sternal, or/and the tergal glands, secreting long-distance sex pheromones to attract a partner (up to 20 cm in most species). After a suitable partner has been accepted, tandem running starts. The leading female secretes chemical stimuli (short-distance or contact pheromones), and the male keeps close contact, antennating and licking the abdomen of the female (Plate 4). Leuthold demonstrated the existence of those short-range pheromones by varnishing the last segments of females eliminating the tandem running [146]. No chemical compound has been characterized as contact pheromone. At some time prior or within this nuptial promenade, the pair sheds its wings. Tandem running finishes when a suitable site is found, where the pair burrows the nest. The first structure built is the royal cell. Contrary to most other insects, mating does not occur immediately. Again nothing is known about mating pheromones as yet, although the royal pair stays together in the royal cell and mating does occur within their whole life span. The first demonstration of a sex-pairing pheromone was published in 1971 [147], and the first attempt to characterize the structure of the sexual pheromone of a termite (Reticulitermes flavipes) was reported by Clement [148] 18 years later. However, the compound, n-tetradecyl propionate, is only effective in extra-physiological concentrations [149]. Thus, we owe most of the pioneering work to Bordereau and his research group. In 1991, investigating the sternal glands of swarming alates of the fungus-growing termite Pseudacanthotermes spiniger (Termitidae), the researchers discovered that the biologically active compound they had isolated was identical with the trailfollowing pheromone (3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol (1) [67]. Two years later they proved that the related Pseudacanthotermes militaris used the same sex-pairing pheromone 1 [70] (Scheme 6). Investigating Reticulitermes santonensis (Rhinotermitidae), they demonstrated that 1 is the sex-pairing pheromone of more than one termite family [135, 150].

26

E. Gössinger

O

OH 1 ((3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol)

2 ((–)-(R)-neocembrene)

6 ((4R*,6S*)-dimethyldodecanal)

8 (trinervitatriene)

O OH 3 ((3Z,6Z)-dodeca-3,6-dien-1-ol)

OH 4 ((3Z)-dodec-3-en-1-ol)

11 ((4R*,6S*)-dimethylundecanal)

O 12 ((E)-2,6,10-trimethylundeca-5,9-dienal)

Scheme 6 Structures of sex-pairing pheromones

In the 1990s, solid-phase microextraction (SPME) was developed permitting solvent-free extraction of small body parts [43–45], thus accelerating the search for sexual pheromones of termites considerably. This technique was used for the first time when the alates of the abundant, neotropical, mandibulated, nasute termite Cornitermes bequaerti (Termitidae) were examined. Bioassays of the extracts of female exocrine glands showed that only the contents of the tergal glands but not those of the sternal gland attracted male termites. However, while the source of the secretion attracting the males was not the sternal gland, the unsaturated primary alcohol (3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol (1) emerged as active compound [149]. The high parsimony of termite pheromones was again demonstrated by this research group when isolating the next pheromone (3Z,6Z )-dodeca-3,6-dien-1-ol (3) [96]. Alates and workers of Ancistrotermes pakistanicus (Termitidae) secrete the unsaturated primary alcohol 3 from their respective glands, which serve as sex-pairing pheromone of the female alate and as trail-following pheromone of workers [96]. The question of the reproductive isolation of closely related species nesting within a short distance and having slightly overlapping dispersal flights caused Bordereau et al. to investigate the sympatric fungus-growing termites Macrotermes annandalei and M. barneyi (Termitidae) [151]. In the course of this investigation, the research group detected a new gland of fungus-growing termites, the sex-specific posterior sternal gland [151]. The sex pheromones of the posterior sternal glands of the two investigated Macrotermitinae are polar substances, different from the trailfollowing pheromones. So far their structures are unknown.

Chemistry of the Secondary Metabolites of Termites

27

From the tergal gland of alates of the Rhinotermitidae species Coptotermes formosanus, Bland et al. isolated a single compound, occurring exclusively in the glands of the females, namely, trilinoleoyl glycerol (trilinolein) (13) (see Scheme 9). The singular occurrence of trilinolein (13) in female tergal glands and the fact that C. formosanus female alates show no calling behavior led the authors to the conclusion that trilinolein (13) is a sexual contact pheromone [152]. Other researchers assume that 13 is a nuptial gift. Behavioral tests were inconclusive, but EAG showed unequivocal signals. The structurally complex (11E)-trinervita-1(14),2,11-triene (8) was isolated as the main compound from the tergal glands of female alates of a neotropical Nasutitermitinae [103]. So far, the tricyclic diterpene 8 is the most complex sexual pheromone of termites. Even this compound shows the parsimony of termite pheromones, because it is structurally closely related to the oxygenated trinervitanes of the frontal glands of soldiers of Nasutitermes species, and it was found as possible trail-following pheromone in Nasutitermes species [104]. The investigation of Prorhinotermes simplex (Prorhinotermitinae) is an example of the difficulties the search for the sex pheromones bears [153]. Females of this species expose their tergal glands when calling, thus their tergal glands were dissected and homogenized in chloroform. The filtered solutes were tested by GC/MS, but no clear picture emerged. Bioassays showed that males were attracted to dissected tergal glands and gland extract. Gas chromatography/EAD with male antennae as detector showed that one of the small peaks of the GC of the extract caused a strong signal from the antenna. As the retention index corresponded with that of dodecatrienol 1, bioassays with synthetic 1 as well as with a second known sex-pairing pheromone, dodecadienol 3, were performed. Only 1 induced strong attraction in reproductive males of P. simplex. Thus, dodecatrienol 1 is most likely part of the sex-pairing pheromone of P. simplex [153]. However, the very small amounts prevented isolation. Thus, it has not been proven to be a content of tergal gland secretion. To expand knowledge about pheromones of termites especially looking at the evolution of sex and trail pheromones, Bordereau et al. investigated the two related, sympatric, basal termite species Zootermopsis nevadensis and Z. angusticollis (Archotermopsidae) [101]. With these dampwood one-piece nesters, both sexes show calling behavior. In both species, the female sex pheromone could be identified as the norsesquiterpene aldehyde (E)-2,6,10-trimethylundeca-5,9-dienal (12) (5–10 ng/termite), resembling closely the trail pheromone (E)-2,6,10trimethylundeca-5,9-dienol (5) of the most basal termite Mastotermes darwiniensis (Mastotermitidae) (Plate 1). In contrast to the female sex pheromone, the male sex pheromone syn-4,6-dimethyldodecanal (6) belongs to the fatty acid metabolites. Thus, both species have the same sex and trail pheromones and inhabit the same area, and their dispersal flights overlap slightly. Despite these facts, hybridization has not been observed [101]. Closely related pheromones were found in Hodotermopsis sjöstedti (Archotermopsidae), a basal dampwood termite of South Asia [102]. Investigation of the sex-specific compounds secreted from the sternal gland revealed that the main

28

E. Gössinger

component of the female sex pheromone is the same as that of Zootermopsis nevadensis, (E)-2,6,10-trimethylundeca-5,9-dienal (12), whereas the male sex pheromone is syn-4,6-dimethylundecanal (11). Synthesis of the isomeric mixture permitted assignment of the structure [102]. How reproductive isolation is maintained when sympatric termite species with identical sex pheromones have overlapping dispersal flights is an open question. To shed light on this question, Bordereau et al. investigated the sex pheromone(s) of three neotropical Syntermitinae [154]. As with all higher termites, only the females call. In Cornitermes bequaerti, C. cumulans, and C. silvestri, the sex pheromones are secreted by the tergal glands. C. bequaerti has been investigated earlier, and only one compound could be identified, (3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol (1) [149]. When looking at the secretion of the tergal glands of the related termite C. cumulans, the authors found as main compound nerolidol (14) and (3Z,6Z,8E)-dodeca-3,6,8-trien1-ol (1) (4:1), and when examining the secretion of the tergal glands of the termite C. silvestri, 1 was the main compound accompanied by nerolidol (14) and (3Z )dodec-3-en-1-ol (4) (Scheme 7). Bioassays showed that nerolidol (14) is no sex attractant. The initial tests seemed to indicate that males of all three species were attracted equally to the secretion of female tergal glands. Only when given the chance to choose between the different sex pairing pheromone of Cornitermes bequaerti OH 1 ((3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol)

sex pairing pheromone of Cornitermes tumulans

OH

HO

+

1 ((3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol)

14 ((E)-nerolidol) (no sex attractant)

sex pairing pheromone of Cornitermes silvestris

OH

HO

+

1 ((3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol)

+

14 ((E)-nerolidol) (no sex attractant) OH

4 ((3Z)-dodec-3-en-1-ol)

Scheme 7 Differences in the sex-pairing pheromones of members of the genus Cornitermes

Chemistry of the Secondary Metabolites of Termites

29

extracts or mixtures of the components, males chose exclusively the composition of the conspecific females. Thus, these experiments demonstrated that sex pheromones are at least partly responsible for reproductive isolation. Interestingly, the authors found that when the females were touched by male antennae of the other species, females fled or reacted agressively. Hence, the influence of the antenna of male reproductives contributes to the long-standing riddle of the reproductive isolation. Very recently, the tergal gland secretions of dealated queens of additional three sympatric Syntermitinae were investigated [145]. The secretion of Embiratermes neotenicus contained as main compounds equal amounts of (3Z,6Z,8E)-dodeca3,6,8-trien-1-ol (1) and (3Z)-dodec-3-en-1-ol (4). The secretion of the two Sylvestritermes species contained as main compound (3Z,6Z )-dodeca-3,6-dien-1-ol (3), and one of the species contained additionally (3Z )-dodec-3-en-1-ol (4) (3:4 ¼ 6:1). The researchers noted that (3Z)-dodec-3-en-1-ol (4) had only a negligible effect on males and thus may not be a reason for the reproductive isolation. Since differing amounts of nerolidol (14) are thought to be the main reason for the reproductive isolation of the sympatric species of the genus Cornitermes, the researchers of this recent publication looked for nerolidol (14) too. They did not find 14 in the tergal gland secretion, but isolated it from the headspace and several parts of the queen’s body! The use of 14 by the queen is unclear [145]. The investigation of the pheromones of the subfamily Psammotermitinae (family Rhinotermitidae) has been expected to contribute to the assignation of Psammotermes within the Rhinotermitidae, a polyphyletic family. No calling behavior of either sex could be observed, but tandem running occurred. The sternal and tergal glands of females of the sand termite P. hybostoma were investigated [68]. Analytical data and sex attraction bioassays of the secretion of both glands revealed dodecatrienol 1 as the pheromone strongly attractive to males of P. hybostoma. The fungus-growing Termitidae Odontotermes formosanus is one of the most harmful termite species in Southeast Asia, causing enormous damage in plantations, forests, and dikes [155]. To learn about their sex pheromones (for possible use in traps or mating disruption), Wen et al. examined the secretion of the sternal gland as well as the headspace of dealated females [156]. They detected that a response of the male alates was caused by a mixture of (3Z,6Z )-dodeca-3,6-dien-1-ol (3) and (3Z)dodecen-1-ol (4) (~40:1) but only 3 is the short-distance attractant for the male (de) alate. Small amounts of one stereoisomer of 3 and 4, respectively, were also isolated, but they initiated no biological activity. Compounds 1, 3, and 4 are known as sex pheromones of other members of the Termitidae subfamilies Macrotermitinae and Syntermitinae [62, 67, 75, 96, 154]. Indications that females of the subfamily Nasutitermitinae may use cyclic diterpenes as sex-pairing pheromones [103, 126] motivated the research group of the Université de Bourgogne to examine the secretion of the sternal as well as tergal glands of females of further Nasutitermes species [62–64]. Indeed, the compounds of the tergal gland secretion of those females differed structurally profoundly from those of the other Termitidae. Whereas the sternal glands contained dodecatrienol

30 Scheme 8 Main sexual pheromones of inquiline Ancistrotermes dimorphus

E. Gössinger

OH 3 ((3Z,6Z)-dodeca-3,6-dien-1-ol)

OH 15 ((3E,6Z)-dodeca-3,6-dien-1-ol)

OH 16 ((6Z)-dodec-6-en-1-ol)

1 and neocembrene (2) in small amounts, the secretion of the tergal glands consisted of large amounts of trinervitatriene 8 and neocembrene (2) in varying proportions from 92% of 8 in Nasutitermes ephratae to pure neocembrene (2) in N. voeltzkowi [63, 64]. Nasutitermes voeltzkowi is a good example to illustrate the large difference in the amount of the same pheromone, neocembrene (2), to signal trail-following in workers (50 pg/individual sternal gland) and sex-pairing in females (106 pg/individual tergal gland). In recent years, the inquilines, termites that cohabitate with host termites of a different species, have become of special interest. The inquilines inhabit no longer used parts of the host nests [84], feed on the nest linings [157], and invade only when the host nest is large enough, so that patrolling by hosts is sparse [158, 159]. These data point to a nonparasitic cohabitation by mutual avoidance. Wen et al. investigated the exocrine secretion of alates of Ancistrotermes dimorphus (Macrotermitinae), a facultative inquiline cohabiting with Macrotermes annandalei and Odontotermes formosanus, respectively [75]. The secretion of the sternal glands of dealates of A. dimorphus yielded as main compound, active in bioassays, (3Z,6Z)dodeca-3,6-dien-1-ol (3), accompanied by very small amounts of (3Z)-dodec-3-en1-ol (4), which showed a signal in the EAG, and the inactive compounds (3E,6Z )dodeca-3,6-dien-1-ol (15) and (6Z )-dodec-6-en-1-ol (16) [75] (Scheme 8). The ratio of these caste-specific compounds was 100:0.67:0.42:0.33. It should be noted that the active compounds are the same as those of one of the facultative hosts (O. formosanus) [156]. Experiments with mixtures of 4 and 3 showed that with higher proportions of 4, the biological activity is inhibited [75].

Isolation and Structure Determination of Sexual Pheromones Alates (150,000) of the fungus-growing termite Pseudacanthotermes spiniger were collected, when swarming [67]. Two extraction techniques were used. First, whole dealated animals were washed in doubly distilled n-pentane. Second, the hypertrophied sternal glands were excised with microscissors from cold anesthetized dealates under a stereomicroscope. The glands after removal of adjacent tissues were

Chemistry of the Secondary Metabolites of Termites

31

allowed to soak in doubly distilled n-pentane. Gas chromatography using the wash of whole animals as well as the extracts of the sternal glands showed only two peaks, which were ten times larger in extracts from females than from males. The solutes were fractionated by preparative GC. Bioassays were made with extracts and with the isolated compound, respectively, in amounts equivalent to one individual. Two dealated males were placed in a Petri dish with two filter papers. One was soaked with the extract, and the other was the control paper soaked with n-pentane. Assessed were not only which paper was favored but also the time the male termites licked and palpated the paper. The main compound, which was analyzed by GC/MS, microhydrogenation, Fourier transform infrared spectroscopy (FTIR), and 1H NMR spectrometry, turned out to be (3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol (1), also known as a trail pheromone of non-reproductives of several termite species. Females of Cornitermes bequaerti were collected when swarming. They were dealated, anesthetized by cooling, and their exposed tergal gland was rubbed gently by a fiber for SPME. The fiber was desorbed immediately in the injection port of a gas chromatograph coupled with a mass spectrometer, and the GC/MS analysis showed nine peaks. The most volatile compound had the same retention time and mass spectrum as (3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol (1) [149]. A comparison of the content of the tergal gland secretion of Cornitermes bequaerti, dodecatrienol 1 [149], with the content(s) of the tergal glands of C. cumulans and C. silvestri was accomplished by SPME of the surface of the gland followed by GC and GC/MS with synthetic 1 as well as (E)- and (Z )-nerolidol as standards. The tergal gland of the related C. cumulans contained as main compound nerolidol (14) and (3Z,6Z,8E)dodecatrien-1-ol (1) (4:1), and the secretion of the tergal glands of C. silvestri contained (3Z,6Z,8E)-dodecatrien-1-ol (1) as the main compound accompanied by nerolidol and (3Z )-dodecen-1-ol (4) [154]. (3Z,6Z )-Dodeca-3,6-dien-1-ol (3) was detected by immersing alates and workers of Ancistrotermes pakistanicus, respectively, in n-hexane as well as by SPME of the sternal glands. Subsequent GC/MS, FTIR, and synthesis [96, 133] confirmed 3 as sex pheromone of the female alate and trail pheromone of the workers. The only compound specific to female alates of Coptotermes formosanus was trilinolein (13) [152]. The excised tergal glands were, after removal of the fat bodies, extracted by n-hexane. Subsequent HPLC/MS pointed to trilinolein (13) as the main compound (Scheme 9). This assumption was confirmed by basic methanolysis of 13 yielding glycerol and a single doubly unsaturated fatty acid methyl ester. The positions of the double bonds were established by the fragmentation pattern of the mass spectrum of the dimethyl disulfide adduct of methyl linoleate (17) [142, 143]. Comparison of the purified compound with synthetic trilinolein (13) confirmed the structure. Although no response was elicited in bioassays, electrophysiological tests showed different effects with female and male antennae and maxillary palps [152]. Solid-phase microextraction of the tergal glands of female alates of two neotropical Nasutitermes species followed by GC/MS showed the same single prominent peak for each species [103]. Larger quantities of the secretion were gained by

32

E. Gössinger O O

NaOCH3, CH3OH, ether, 5', rt; aq HOAc

O O

O O 13

O

OH +

OH

HO

O 17 (methyl (9Z,12Z)-octadeca-8,12-dienoate)

glycerol

Scheme 9 Methanolysis of trilinolein (13)

H

H

H

H

H

H

H

H H

H 8

Scheme 10 NOE interactions of trinervitatriene 8

dealating females (~13,000 individuals) and submerging them in n-hexane. Preparative TLC was used for isolation. The mass spectrum pointed to a polycyclic diterpene. One- and two-dimensional NMR spectrometry and IR spectrometry revealed a trinervita-1(14),2,11-triene skeleton. Nuclear Overhauser effects permitted the determination of the configuration of the Δ11,12-double bond and the relative configuration of the stereogenic centers (Scheme 10). Comparison with ab initio calculations of the NMR spectra confirmed the assumed, complex structure of the tricyclic diterpene (4R,7S,8R,11E,15S,16S)-trinervita-1(14),2,11-triene (8). The absolute configuration was assigned tentatively according to the known hydroxylated trinervitanes found in the frontal glands of Nasutitermitinae as defensive compounds. The female sex pheromone (E)-2,6,10-trimethylundeca-5,9-dienal (12) and the male sex pheromone syn-4,6-dimethyldodecanal (6) of two Zootermopsis species were isolated using the SPME technique [101]. The hypertrophied sternal glands of female and male alates, respectively, were rubbed with a polydimethylsiloxane/ divinylbenzene-type fiber, which then was inserted in a gas chromatograph injector port. Gas chromatography/MS data were used for preliminary structure determination, followed by FTIR, online microhydrogenation fused to GC/MS [160], and

Chemistry of the Secondary Metabolites of Termites

33

syntheses to confirm the assumed structures. The female sex-pairing pheromone (E)2,6,10-trimethylundeca-5,9-dienal (12) (5–10 ng/termite) was characterized by its close resemblance to the trail pheromone of Mastotermes darwiniensis (Plate 1), (E)2,6,10-trimethylundeca-5,9-dienol (5), and the structure was confirmed by synthesis according to Kulesza and Gora [161]. The male sex pheromone 4,6-dimethyldodecanal (2–5 ng/termite) was determined mainly by GC/MS and IR. Consecutive synthesis of the syn-4,6-dimethyldodecanal (6) confirmed the stereochemistry [100]. The closely related sex-pairing pheromones of Hodotermopsis sjöstedti were isolated by SPME of the sternal gland of females and males, respectively [102]. Gas chromatography/MS, FTIR, and comparison with the spectra of identical and closely related compounds revealed the main component of the female sex pheromone as the known norsesquiterpene, (E)-2,6,10-trimethylundeca-5,9-dienal (12). The male sex pheromone was identical with synthetic syn-4,6dimethylundecanal (11). Sternal and tergal glands of females of the sand termite Psammotermes hybostoma were excised and extracted with n-hexane [68]. Concentrated extracts were used for GC/EAD and GC/MS. The data were compared with those of a solution of standard n-alkanes and (3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol (1). Additionally, sex attraction bioassays were conducted. Gas chromatography/EAD of the tergal glands as well as of the sternal glands of females showed a single significant antennal response. Co-injection with standard hydrocarbons showed the Rt value of the antennal response to be similar or identical with that of dodecatrienol 1. Analysis of GC/MS data confirmed that the active compound was indeed 1. Furthermore, bioassays showed that males were strongly attracted to 1 at 10 pg/paper. The glands of dealated females of Odontotermes formosanus were excised, and SPME was carried out with the gland secretion, with the n-hexane extract of the glands and as headspace SPME. Subsequently, the compounds extracted by the three different methods were analyzed by GC/MS or GC/EAG. Synthetic (3Z)-dodec-3en-1-ol (4) and (3Z,6Z )-dodeca-3,6-dien-1-ol (3) [96] served as standards. The results were supported by short- as well as long-range bioassays [156]. The response of the male alates was caused by a mixture of 3 and 4 (~40:1). (3Z,6Z )-Dodeca-3,6dien-1-ol (3) is the short-distance attractant for the male (de)alate. The sex pheromones secreted from the sternal glands of Ancistrotermes dimorphus were isolated by dissecting the gland, removing the fat bodies, and extracting by SPME as well as by immersing in n-hexane. As a control, a small part of the tergal cuticle was also examined. The authors found no CHCs at the cuticle of this inquiline, which may explain the ability to invade the nests of host termites without being recognized as alien. This finding was unexpected because cohabiting or parasitic insects usually have CHCs very similar to those of their hosts. The extracts of the glands examined by GC and GC/MS contained (3Z,6Z )-dodeca3,6-dien-1-ol (3) as main component (0.01–1 ng), which was characterized by comparison with synthetic 3 and the fragmentation pattern of the dimethyl disulfide adduct of 3 [75]. Examination with male antennae as detector (EAD) revealed a

34

E. Gössinger

second active compound, dodecenol 4, in very small amounts. Examination of the glands by SPME and GC/MS revealed two further compounds. The two inactive compounds, which could be characterized as dodecen-1-ol and dodecadien-1-ol by GC/MS, were treated with dimethyl disulfide and iodine. The fragmentation pattern of the mass spectra of these adducts revealed that the two unsaturated C12 alcohols were (3E,6Z )-dodeca-3,6-dien-1-ol (15) and (6Z )-dodec-6-en-1-ol (16). Their structures were confirmed by syntheses and comparison of several synthetic isomers. The thorough examination of the alates of Macrotermes annandalei and M. barneyi, caught by light trapping within their dispersal flight, led to the discovery of a new exocrine gland of these Termitidae, the posterior sternal gland [151]. Not only were all three abdominal exocrine glands (sternal, posterior sternal, and tergal gland) excised; they were separately extracted with solvents of different polarities (n-hexane, dichloromethane, and water). No responses of male (de)alates were detected with the contents of the sternal glands and the n-hexane extracts of the posterior sternal and of the tergal gland. However, the dichloromethane and water extracts of the two latter gland types led to excitement and prolonged licking by dealated males of the same species. Although neither the long-distance nor the shortdistance sex-pairing pheromones of the two species could be characterized, the investigation demonstrated the important influence of the pheromones on reproductive isolation. It also emphasized the shortcoming of the usually employed extraction methods including SPME combined with GC/MS due to the higher polarity of the short-distance sex-pairing pheromones.

2.1.3

Syntheses of Trail-Following and Sex-Pairing Pheromones

Most of these releaser pheromones are structurally simple open-chain alcohols or their oxidation products. The intention of the first syntheses was the confirmation of the structures elucidated by spectrometric data, especially of the configuration of the double bonds or of the relative configuration of the methyl substituents. The following syntheses were aimed at easy access of enough material for behavioral studies and testing the possible use in pest management. This implies that several of these syntheses differ only by small variations, with the exception being the two cyclic diterpenes, for which their more complex structures posed a challenge to synthesis chemists. Neocembrene (2) has been synthesized several times, but the trinervitatriene 8 has not succumbed to synthesis as yet. The first structure of a termite pheromone, determined in 1969, was dodeca-3,6,8trien-1-ol (1) [86]. Tai et al. established the configuration of the three double bonds by synthesizing three of the possible diastereomers and comparing the trailfollowing behavior of the termite workers using the secretion of the sternal gland and the separated synthetic diastereomers, respectively. The result showed that the specific stereochemistry of the double bonds is essential. Tai et al. used coppercatalyzed coupling of an alkyne with propargyl bromide and subsequent semihydrogenation with the Lindlar catalyst to generate the skipped (Z )-diene of (3Z,6Z,8)-dodeca-3,6,8-trienol (1). The double bond at C-8 was obtained by

Chemistry of the Secondary Metabolites of Termites

35

dehydration of the homoallylic alcohol leading to a mixture of (E)- and (Z )-configured diastereomers [73]. Only (3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol (1) was highly active. This first synthesis of 1 was achieved in six steps and 0.8% overall yield. The troublesome separation of the diastereomeric dodecatrienols motivated several research groups to design stereoselective syntheses of 1. Yamamoto et al. applied as key steps recently developed methods within their own research group, inclusive of a stereoselective formation of 1,3-dienes by addition of allyltitanium compounds to aldehydes [161, 162] and the stereoselective deconjugative isomerization of unsaturated esters [163]. Although the synthesis necessitated 11 steps, the overall yield could be improved to 13% [163, 164]. The most cited synthesis of 1 is that of Argenti et al. They followed partly Tai’s pathway but introduced the intermediate enyne stereoselectively, improved the copper(I)-catalyzed coupling considerably, and achieved high stereoselectivity by substituting the semihydrogenation with the Lindlar catalyst by reduction of the alkynes with hydroboranes [133]. This slightly convergent synthesis furnished 1 in 11 steps and 19% overall yield. The synthesis design of Tai and Argenti was used in the syntheses of Correa et al. (eight steps, 7% overall yield) [165] and Saran et al. (seven steps, 9% overall yield) [66]. The intention of the synthesis efforts of Eya’s group was to prepare all eight possible diastereomeric dodeca-3,6,8-trien-1-ols [166]. Thus, it was more important to have as few as possible starting materials and reactions to achieve the syntheses of all eight possible diastereomers than high yields. The synthesis of the actual trailfollowing pheromone 1, which was accompanied by the diastereomeric (3Z,6E,8E)-dodeca-3,6,8-trien-1-ol (18), featuring semihydrogenation of an alkyne and Wittig condensation with an unsaturated aldehyde as key steps. (3Z,6Z )-Dodeca-3,6-dien-1-ol (3), a sex and trail pheromone of the Macrotermitinae Ancistrotermes pakistanicus [96], was synthesized many times even prior to its discovery as termite pheromone. It was prepared as an analogue to (3Z,6Z,8E)-dodeca-3,6,8-trien-1-ol (1) [87, 88, 133] and found to be very attractive to termites of several families. (3Z,6Z )-Dodeca-3,6-dien-1-ol (3) is also an ideal building block for the arachidonic acid derivatives, and as a small molecule, it was very suitable to test new methodologies to form skipped double bonds. Coupling of alkynes and subsequent partial hydrogenation was the most often used methodology [87, 88, 96, 133, 160–164, 166–178]. Several of these publications are but very small variants of earlier work; detailed descriptions of each of these syntheses would be repetitive. Examples of the different variations are presented in detail. One of the drawbacks of these related synthesis pathways is the tedious separation of the targets from the stereoisomeric by-products. Some authors have used propargylic compounds as starting materials and elongated the chain by Wittig condensation [173, 179–184]. Vasilev’s synthesis is presented [181] because it deviates from the obvious pathway; it is highly stereoselective and suitable for upscaling. The synthesis of Rokach et al. is included because it furnishes a labeled dodeca-3,6-dien-1-ol [172]. Another way to avoid formation of stereoisomers was chosen by Prestwich et al. [168]. A cyclohexa-1,4diene derivative was cleaved selectively forming the (Z )-configured unsaturated open-chain aldehyde. Wittig condensation was utilized for the preparation of the

36

E. Gössinger

second double bond. The most recent synthesis by Hutzinger et al. [185] is an elaborate attempt to avoid any stereoisomeric by-products in the preparation of the skipped double bond. The synthesis of (Z )-dodec-3-en-1-ol (4) is trivial. Thus, only one synthesis is presented [133]. Two very simple syntheses verified the structure of (Z,Z )-nonadeca-10,13-dien2-one (9). The structure determination of cis-4,6-dimethyldodecanal (6), the male sex pheromone of two Zootermopsis species, which turned out to be also the trailfollowing pheromone of the pseudergates, depended on spectral data [101]. However the relative configuration had to be assigned by syntheses. Braekman et al. started by synthesizing a mixture of cis- and trans-4,6-dimethyldodecanal (6 and 19) [100] in a convergent synthesis with a sterically crowded Wittig condensation as key step. Since the resulting mixture of the stereoisomeric aldehydes, obtained in seven steps and 27% overall yield, was inseparable, the authors decided to synthesize stereoselectively the cis-4,6-dimethyldodecanal (6) relying on the predictable stereochemistry of the six-membered ring and using the Baeyer-Villiger reaction to cleave the substituted cyclohexanone. Pure 6 was obtained in ten steps and 15.5% overall yield [100]. The homologous cis-4,6-dimethylundecanal (11) was detected as the male sex pheromone of Hodotermopsis sjöstedti and the corresponding alcohol cis-4,6dimethylundecanol (7) as trail-following pheromone of the pseudergates of the same species. Braekman et al. used the same pathway as for the mixture of 6 and 19. In this case, the mixture was separable, and the relative configuration could be determined by comparison of TLC Rf values. The alcohols 7 and 20 were obtained in five steps and 32% overall yield and the corresponding aldehydes 11 and 21 in six steps and 24% overall yield [100]. So far only two sesquiterpenoids have been recorded as trail and/or sex pheromones: the norsesquiterpene aldehyde (5E)-2,6,10-trimethylundeca-5,9-dienal (12), the female sex pheromone of Archotermopsidae species [101, 102], and the corresponding alcohol, (5E)-2,6,10-trimethylundeca-5,9-dienol (5), the trailfollowing pheromone of Mastotermes darwiniensis (Plate 1) and of species of the family Stolotermitidae [99]. 2,6,10-Trimethylundeca-5,9-dienal (12) and (5E)2,6,10-trimethylundeca-5,9-dienol (5) are also present in essential oils of plants [186]. To establish the structure and especially the configuration of the central double bond of 5 and 12, Bordereau et al. [99] followed the synthesis pathway of Kulesza and Gora [161], which started with geranylacetone, and they reached 12 by a Darzens reaction and consecutive saponification and decarboxylation. This racemic synthesis needed four steps (28% overall yield). The same reaction sequence with nerylacetone led to the stereoisomer with a (Z )-configured double bond at C-5 allowing the designation of the natural product as 12. Several more syntheses of 12 and 5 have been published as building blocks of syntheses of more complex natural products [187–193]. Of these syntheses, Carreira’s enantioselective synthesis is described, which started with homogeranyl iodide and used pseudoephedrine according to Myers [194] to induce chirality, furnishing in two steps and 84% overall yield the enantiopure alcohol 5 and in three steps and 60% overall yield the enantiopure aldehyde 12.

Chemistry of the Secondary Metabolites of Termites

37

The first isolated pheromone of a termite was the monocyclic diterpene, neocembrene (2) [91]. However completion of the structure determination necessitated total synthesis too. Thus, 6 years passed until neocembrene was established as (3E,7E,11E,1R)-4,8,12-trimethyl-1-(1-methylethenyl)cyclotetradeca-3,7,11-triene. Cyclotetradecanes are on the border of medium- to large-sized rings. Usually, transannular and torsional strain aggravate the ring closure of 1,14-disubstituted chains. An additional handicap for planning and synthesizing those rings are their several conformations in a small range of energy; due to the Curtin-Hammett principle, the prediction of the stereochemistry is difficult. Unsaturation decreases the transannular and torsional strain as well as the free rotations in the open chain, thus decreasing the free enthalpy of the transition state. Accordingly, cyclization to neocembrene with its three endocyclic double bonds concurs favorably with dimerization and leads to a relatively strain-free ring as is demonstrated by the following described syntheses. Kato’s discovery that electrophilic olefin cyclization of the open-chain (E)-configured diterpenes as well as open-chain (E)-configured sesterterpenes led exclusively to the respective 14-membered rings also points to the small ring strain [124]. The low-energy conformations of neocembrene have been examined by Kato [195] and by Setzer [196]. Setzer et al. calculated that the conformation with the lowest energy has all three double bonds of the cyclotetradecatriene in a perpendicular position to the ring plane. The isopropenyl substituent and the methyl group at C-12 are in pseudoequatorial positions, whereas the methyl groups at C-4 and C-8 are in pseudoaxial positions above and below the ring plane, respectively [196]. Setzer’s research was aimed to understand the cytotoxic activity of neocembrene, that is, the conformation necessary for docking of neocembrene at the colchicine binding site of tubulin. Kodama’s and Kato’s syntheses determined the (E) configuration of all three endocyclic double bonds of neocembrene. These two syntheses were followed by ten further syntheses of neocembrene, although neocembrene is not of commercial value. However, its structure was determined at a time of intense investigation of cyclization methods, of the stereochemistry of the cyclization, and of the reactivity and stereochemistry of medium- to large-sized rings in vitro and by calculation (e.g., [197–199]). An example of this trend was the first synthesis of a cembrene by Dauben et al. in the year 1976 [200], which was an additional stimulus for the synthesis of neocembrene. Comparison of these syntheses shows that eight different ring closure methods were utilized and five different bonds of neocembrene (2) were formed by cyclization, thus contributing to the overall knowledge of the chemistry of medium- to large-sized rings. Kodama et al. started with geranylgeraniol and closed the cyclotetradecatriene at the C-1–C-2 bond by SN2 reaction of a sulfur-stabilized carbanion attacking an epoxide (67%). The synthesis needed seven steps (6% overall yield). Kato, Kitahara et al., who were involved in the structure elucidation with Australian researchers, started their first successful neocembrene synthesis with geranylgeranoic acid. Their cyclization step was the intramolecular olefinic Friedel-Crafts reaction of the acid chloride at the C-1–C-2 bond (72%). This racemate synthesis needed four steps (overall yield 23%). The authors used an intermediate allyl alcohol to separate the

38

E. Gössinger

enantiomers by esterification with a chiral auxiliary. In a second attempt, these researchers constructed the open-chain precursor starting with two units of geraniol in several steps. Intramolecular exchange of bromide by a sulfone-stabilized carbanion was used to close the ring at the C-9–C-10 bond (60%). Including the construction of the open-chain precursor, at least ten steps were needed to complete the synthesis (overall yield 99% ee) (–)-498

1) LAH, ether, rt, 98% 2)TEMPO, KBr,

aq AcOH, CH3OH, rt, 56% (2 steps)

502

O

OAc

Mucor michei lipase, i-Pr2O, 7 d, rt

2)

1) separation

Scheme 77 Asymmetric synthesis of (+)-ancistrofuran ((+)-275) using lipase as chiral catalyst

2) Bu3SnH, AIBN, PhCH3, rfl, 60% (2 steps)

MOMO

2) MOMCl, i-Pr2NEt, CH2Cl2, rt, 92%

MOMO

498

499

O 59% (66% ee)

+

OH

34% (96% ee)

O

OAc

1) CH3Li, CeCl3, THF, –78°C to rt, 89%

Mucor michei lipase, i-Pr2O, 48 h, rt

OAc

1) PhOCSCl, DMAP, CH2Cl2, rt

(+)-501

498

O

OH

172 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

173

subsequent lactonization to give (+)-501. Utilizing the methylcerium compound allowed selective addition of the methyl group to the keto group of (+)-501. The tertiary alcohol was protected as methoxymethyl ether affording 502. The lactone was now opened by reduction to the diol with lithium aluminum hydride. To achieve the desired chain elongation, the primary alcohol was selectively oxidized to the aldehyde 503 by sodium hypochlorite mediated by TEMPO. Cyanohydrin 504 was formed using potassium cyanide in acetic acid. Both secondary alcohols of 504 were removed by esterification to the respective thionocarbonates and radical reduction with tributylstannane. Addition of 3-lithiofuran to the generated cyanide 505 led to the known furan derivative 481 in enantiomerically pure form, which was converted into (+)-ancistrofuran (275) according to Baker’s second ancistrofuran synthesis [345, 346]. This EPC synthesis led in 14 steps and 3% overall yield to (+)-275.

Asymmetric Synthesis of ()-Ancistrofuran by McErlean et al. As in Saito’s racemate synthesis, the most recent synthesis featured the biomimetic polyolefin cyclization as the key step [351] (Scheme 78). (+)-9-Hydroxydendrolasin (+)-(486) was prepared starting with geranyl chloride (484), which was converted into the barium organic compound to ascertain complete retention of the alkene geometry [592–594] when adding the organyl group to furan-3-carbaldehyde (506). The resulting racemic 9-hydroxydendrolasin (486) was oxidized to 9-ketodendrolasin, which was reduced enantioselectively to (+)-9-hydroxydendrolasin (486) in over 90% enantiomeric excess using Noyori’s ruthenium catalyst (507) [595]. One of the aims of this synthesis was to introduce a newly developed chiral binolphosphoramidite catalyst for the enantioselective brominative polyolefin cyclization. In 2007, Ishihara et al., looking for an enantioselective catalyzed bromination, reasoned that phosphor (III) nucleophiles with their higher HOMO should be better catalysts for nucleophilic attack at bromine than amines. Therefore, they designed chiral monodentate phosphor (III)-containing complex-forming compounds, which indeed proved especially suited for chiral bromination [596]. McErlean et al. examined and improved the phosphoramidite catalysts [597, 598]. To test the advantage of the new catalyst (S)-TCPT (508), the researchers performed a polyolefin cyclization with racemic as well as with enantiomerically pure 9-hydroxydendrolasin (486). In both cases, the two possible stereoisomeric cyclization products were formed in the same ratio. When (+)-486 was used, the major compound was (+)-luzofuran (509), originally isolated from the red alga Laurencia luzonensis [599]. This stereochemical result was explained by the authors due to the strong inductive effect of the stereogenic center of 486, but this is contrary to Saito’s electrophilic olefin cyclization of 486 [348]. No cyclization occurred if the reaction was performed without the catalyst. The minor cyclization product (+)-epi-luzofuran (510) was debrominated with activated magnesium furnishing ()-ancistrofuran (275) in five steps and 2.5% overall yield.

O

H

Cl

Cl

(S)-TCBT:

O

X+

O 506

Cl

H

Cl

Cl

36%

4:1

(+)-486

Cl

PO N N N N

508

O

O

OH

THF, –78°C, 58%

N N N

O

H

OH

O

H

O

X+

507

Li, naphthalene; MgCl2,THF, rt;

Cl

510, THF, –78°C; –65°C, 86%

Ru N H2

N

Ts

Br

0.2 eq 508, 4 Å MS, CH2Cl2; NBS, –78°C

C10H11 (CH3)3N+Br-, AcOEt, rt, 79% (>90% ee)

2) 507, HCO2Na, H2O,

510 ((+)-epi-luzofuran)

O

1) Dess-Martin ox., NaHCO3, 0°C to 15°C, 92%

Noyori cat (= (S,S)-RuTsDPEN) :

O

486

OH

H

O

OH

O

(–)-275 ((–)-ancistrofuran)

O

(+)-486

Scheme 78 Asymmetric synthesis of ()-ancistrofuran (()-275) via (+)-9-hydroxydendrolasin (486) using Noyori’s catalyst

509 ((+)-luzofuran)

Br

0.2 eq 508, 4 Å MS, CH2Cl2; NBS, –78°C

484

Cl

Ph-Ph Li+, BaI2, THF, rt; 484, THF, –78°C;

174 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

175

Ancistrodial As indicated above, the defensive secretion of the minor soldiers of the fungusgrowing Macrotermitinae Ancistrotermes cavithorax consists mainly of the monocyclic dialdehyde ancistrodial (276). Baker et al. established the structure of ancistrodial (276) by synthesis [343] as shown in the following paragraph.

Synthesis of Racemic Ancistrodial by Baker et al. The synthesis [343] (Scheme 79) opened with the starting material ()-γ-cyclohomocitral (458), which was prepared from 2,2-dimethylcyclohexanone according to Julia et al. [600]. Stobbe reaction of 458 with diethyl succinate led to the monoester 511, which was reduced with lithium aluminum hydride followed by reoxidation to the desired dialdehyde ancistrodial (276) in four steps and 26% overall yield and its (Z )diastereomer. However, the configuration of the double bond was only assigned tentatively. Vidari et al. were able to confirm the tentative assignation in their asymmetric synthesis of 276 [354].

CO2Et

O

CO2Et

O

OEt

t-BuOK, 68%

OH

O

1) LAH, ether 2) CrO3⋅2pyr,

H

CH2Cl2, 50% (2 steps)

H

O 458

511

Scheme 79 Synthesis of the diastereomers of ancistrodial (276)

O (Z):(E) = 1:6 276

176

E. Gössinger

Asymmetric Synthesis of Ancistrodial by Vidari et al. As in Baker’s synthesis, γ-cyclohomocitral (458) was the decisive intermediate, but now as an enantiomerically pure compound with the stereogenic center in the (R)configuration (Scheme 80). Methylmagnesium bromide was added to acetylcyclopropane (512), and the resulting tertiary alcohol was treated with hydrogen iodide furnishing the openchain homoallylic iodide (513). This iodide was coupled with 2-lithio-4,5dihydrofuran according to Boeckman et al. [601] at low temperature. The generated dihydrofuran derivative 514 was cleaved with (trimethylsilyl)methylmagnesium chloride catalyzed by the nickel(II) complex 515 [602, 603]. This method permitted access to the open-chain compound 516 with mainly retention of the double-bond configuration ((Z)/(E) ~ 10:1). To introduce chirality, the primary alcohol of 516 was protected as a pivalate to enable selective addition to the sterically less encumbered of the two trisubstituted double bonds by removing the directing effect of the homoallylic alcohol. Enantioselective Sharpless dihydroxylation [604, 605] afforded diol 517 with the newly formed stereogenic center in the (S)-configuration. Only a very small amount of the regioisomeric diol was observed. To enable the planned electrophilic olefin cyclization, the 1,2-diol was converted into the oxirane via the monomesylate and intramolecular substitution by the hydroxylate. When the oxirane 518 was treated with a strong Lewis acid, the substituted γ-cyclohomogeraniol derivative 519 was produced. At this point, the small amounts of trans-substituted γ-homocyclogeraniol generated from the E-configured double bond were separated by chromatography. The obsolete secondary hydroxy group of 519 was reductively removed by esterification with phenyl thionochloroformate and treatment with tributylstannane under radical conditions. The pivalate 520 was reductively cleaved by lithium triethylborohydride (superhydride) to alcohol (R)-521, and successive Ley oxidation afforded the enantiomerically pure (R)-γ-cyclohomocitral (R)-(458). Wittig condensation with the stable 3-triphenylphosphoranylidene butyrolactone (475) [606] led exclusively to compound 522 with the E-configured double bond. Reduction of the lactone moiety with diisobutylaluminum hydride furnished the diol, which was reoxidized to the dialdehyde. Thus, enantiopure ancistrodial (276) was obtained in 3% overall yield within 16 steps. This compound proved to be identical with ancistrodial (276), hence establishing its double-bond configuration. The amounts of ancistrodial (276) isolated from the minor soldiers of Ancistrotermes cavithorax were too small to compare with the enantiomerically pure synthetic ()ancistrodial (276).

2) HI, 0°C, 79%

O

522

PivO

517

OH

513

O

Li

75% (3 steps) HO

O (–)-276 ((–)-(R)-ancistrodial)

O

514

PivO

O

LA

(R)-521

2) K2CO3, CH3OH, 0°C, 82% (2 steps)

LiBHEt3, THF, rfl,

OPiv

O

1) MsCl, pyr, CH2Cl2, –30°C

THF, –30°C to rt,

Si (CH3)3

2) (COCl)2, DMSO, (C2H5)3N, CH2Cl2, –60°C, 65%

1) DIBAH, THF, –10°C to 0°C, 72%

520

I

Scheme 80 Asymmetric synthesis of ()-ancistrodial (()-276)

O

2) Bu3SnH, AIBN, THF, rfl

1) PhOCSCl, pyr, CH2Cl2, rt

2) AD-mix α, CH3SO2NH2, t-BuOH/H2O, 0°C, 66%

HO

1) CH3MgBr, ether, rfl, 83%

1) PivCl, N(CH3)3, CH2Cl2, rt, 75%

512

O

O

CH2Cl2, rt, 75%

N

(C3H7)4N+RuO4- , O , 4 Å MS,

518

Si(CH3)3

PhH, rfl, 66% (2 steps)

O

515

PPh2

Ni2+ 2Cl-

PPh2

(R)-458

83% 2) separation

=

Ph3P 475

O

O

519

dpppNiCl2

THF, 50°C, 90%

PivO

HO

OH

Si(CH3)3

main compound 516

1) BF3•OEt2, CH2Cl2, –78°C,

(CH3)3SiCH2MgCl, dpppNiCl2 (515),

Chemistry of the Secondary Metabolites of Termites 177

178

E. Gössinger

EPC Synthesis of Ancistrodial by Mori et al. Again, (R)-γ-cyclohomocitral (R)-(458) was the key intermediate. In Mori’s case [355], the chirality was introduced by kinetic racemate resolution (Scheme 81). Racemic γ-cyclohomocitral (458) was prepared in seven steps according to Kawanobe [607], starting with 2-methylcyclohexanone by methylation, formylation, and Claisen rearrangement as key steps. The following reduction to the primary alcohol ()-521 permitted lipase-catalyzed esterification. Unfortunately, several cycles of esterification to the acetate 523, chromatographic separation, and saponification were necessary to gain ()-521 in 98% enantiomeric excess. Swern oxidation generated enantiomerically pure (R)-γ-cyclohomocitral (R)-(458), which reacted by Wittig condensation with dimethyl triphenylphosphoranylidenesuccinate (524) to give the diester 525. Reduction with lithium aluminum hydride and reoxidation with Dess-Martin periodinane led to enantiomerically pure ()-ancistrodial (276) in 7.5% overall yield starting with racemic γ-cyclohomocitral (458). 8 steps

OAc , lipase AK, MS 4 Å, 0° to 4°C, n-hexane

OH

O (±)-521

1) separation 2) NaOH, aq CH3OH,

+ OH partially resolved 521

OAc partially resolved 523

the 3 steps sequence was twice repeated 16%

Ph3P

1) (COCl)2, DMSO, NEt3, CH2Cl2, OH

–60 to 0°C, 89%

(–)-521 (98% ee)

O

CO2CH3 CO2CH3

PhH, rfl, 12h, 62%

458

1) LAH, ether, rt, 2 h, 93%

O OCH3 OCH3 525

524

O

O H

2) Dess Martin ox., pyr/CH2Cl2, rt, 15', 92%

H (–)-276

Scheme 81 Stereoselective EPC synthesis of ()-ancistrodial (()-276)

O

Chemistry of the Secondary Metabolites of Termites

179

Evuncifer Ether  4,11-Epoxy-cis-eudesmane Baker et al. isolated this tricyclic ether 278 (evuncifer ether, 4,11-epoxy-ciseudesmane) and determined its structure [359]. Prestwich et al. identified evuncifer ether (278) in the defensive secretion of Amitermes messinae and the South American termite Amitermes excellens [353, 360], Braekman et al. in Amitermes arboreus [303], and Scheffrahn in Amitermes wheeleri [373, 581], Amitermes emersoni, Amitermes coachellae [334], Amitermes minimus [361], and Amitermes desertorum [292].

EPC Synthesis of the Evuncifer Ether by Baker et al. Commercially available (R)-carvone (526) was the starting material of Baker’s synthesis [358] depicted in Scheme 82. Hydrogenation of (R)-carvone (526) with zinc under basic conditions generated 2,3-dihydrocarvone, which was hydroxylated by mercury(II) acetate and subsequent treatment with sodium borohydride. The cyclic ketone 527 was converted into the eudesmenone (528) using a variant of the Robinson annulation. Strong acid afforded dehydration to the enone α-epi-carissone (529). The now obsolete keto group was reduced furnishing the allyl alcohol. Its acetate was treated with lithium in liquid ammonia yielding epi-γ-eudesmol (530), which earlier had been synthesized by Marshall et al. using a slightly different route [578]. epi-γ-Eudesmol (530) yielded the tricyclic ethers ()-evuncifer ether (278) and dihydroagarofuran (531) by HO

O

1) Zn, NaOH, C2H5OH/H2O O

O Cl

2) Hg(OAc)2,THF, H2O; NaBH4, NaOH, CH3OH

O

OH

526

528

527

HCl conc, C2H5OH

OH

NaH, THF

OH O

OH

1) LAH, THF 2) Ac2O, pyr 3) Li/NH3 liq 530

529

Hg(OAc)2, THF, H2O;

O

O

+

NaBH4, NaOH, CH3OH, 80% 278 ((–)-evuncifer ether)

3:1

531

Scheme 82 EPC synthesis of ()-evuncifer ether (()-278) and dihydroagarofuran (531)

180

E. Gössinger

mercuration and successive reduction with sodium borohydride in a 3:1 ratio. Separation of the two compounds completed this EPC synthesis in seven steps. The identity of the optical rotation of synthetic and natural 278 established the absolute configuration of the evuncifer ether from the defense secretion of Amitermes soldiers. Later an identical synthesis was published [608].

Synthesis of Racemic Evuncifer Ether by Wijnberg et al. Wijnberg, de Groot et al. developed a general method for the synthesis [356] of all possible stereoisomers of 10-methyldecalin-3,6-dione [609], which they then further developed into syntheses of several naturally occurring eudesmane derivatives [356, 610–612]. Among others, these authors completed the racemate syntheses of the termite defensive compounds amiteol (365), intermedeol (414), neointermedeol (415), and evuncifer ether (278). The protocol for the synthesis of the common intermediates starts with the product 532 of the Robinson annulation of methyl vinyl ketone to 2-methylcyclohexanone (Scheme 83). The bicyclic enone 532 was converted into the dienyl acetate according to Chowdhury et al. [613, 614] and epoxidized. The conditions chosen for the epoxidation led immediately to hydrolysis of the enyl acetate and consecutive cleavage of the epoxide by E1cB reaction furnishing a mixture of the two epimeric unsaturated alcohols 533 and 534. Concentrated hydrobromic acid transformed the enolones via enol formation into the 1-methyldecalin-4,7-diones 535 and 536 (trans/cis ¼ 2:1). At this point, the synthesis pathway of the trans-eudesmane derivatives intermedeol (414) and neointermedeol (415), compounds isolated from termites and from plants, branched off. To convert the trans-decalindione 535 into the epimeric cisdecalindione 536, the investigators reasoned that increasing steric hindrance might change the equilibrium toward the cis-decalin. Indeed, when the mixture of the decalindiones was transformed into the diketal, the cis-decalin derivative 537 prevailed. Mild acidic hydrolysis led to cis-decalindione 536. Selective transketalization afforded the cyclic monoketal 538, the common intermediate for the syntheses of cis-eudesmanes isolated from termites. Methylmagnesium iodide attacked the keto group of 538 mainly from the convex face leading to the tertiary alcohol 539. Acidic hydrolysis of the cyclic ketal liberated the keto group. The keto group of hydroxyketone 540 was converted into the ethylidene group of 541 by Wittig condensation. anti-Markovnikov hydration by borane addition and subsequent oxidation furnished the diols 542. Oxidation led to the crystalline lactol 543b. Methylation using lithium cyano(methyl)cuprate and the strong Lewis acid boron trifluoride etherate completed the synthesis of racemic evuncifer ether (278) in 12 steps and 8% overall yield.

537

O

O

DMSO, rt, 91%

NaH, DMSO; Ph3P+Et Br-,

70% (2 steps)

(CH3)2CO/H2O, PPTS, rt,

O

2) MCPBA, dioxane, buffer, rt, 75% O

543b

O

OH

O

O

BF3•OEt2, 3', 64%

CH3Li, CuCN, 0°C; –78°C, 543,

H2O2, NaOH, THF/H2O, rt; rfl; 79%

O

538

BH3•THF, THF, rt; rfl;

85%

HBr conc, ether, rt,

(CH2OH)2 cat, TsOH, CH2Cl2, rt, 69%

541 ((E):(Z) = 1:1)

OH

O

O

533 OHax (57%), 534 OHeq (18%)

536

HO

Scheme 83 Synthesis of evuncifer ether (278) from octalinone 532

540

OH

main conformation

O

O

O

532

O

1) Ac2O, TMSCl, NaI, 0°C, 80%

O

O

278

rt

+ 2:1

CH3MgI, ether,

OH

542

OH

O

535

O 536

90%

PDC, pyr, rt,

539 + 10% epimer

OH

O

O

O

O

78% (2 steps)

(CH3)2CO/H2O, HCl, rt,

543a

OH

O

rt, 6 d

HC(OCH3)3, CH3OH, H2SO4,

Chemistry of the Secondary Metabolites of Termites 181

182

E. Gössinger

Serendipitous Synthesis of Racemic Evuncifer Ether by Kodama et al. Kodama developed a general cyclization method suited for the syntheses of mediumand large-sized rings [357]. After testing this method successfully by closing the unsaturated cyclotetradecane by intramolecular attack of a sulfur-stabilized carbanion at an oxirane [119], they extended these experiments for synthesizing the unsaturated cyclodecane system [120, 122, 615]. Kodama et al. chose as starting material phenyl farnesyl sulfide (544) and used van Tamelen’s selective epoxidation to introduce the terminal epoxide 545 (Scheme 84). n-Butyllithium at low temperature generated the allyl carbanion, which attacked the epoxide yielding (2E,6E)- and (2Z,6E)-hedycaryol phenyl sulfide (546:547 ¼ 7:5). When 547 was dissolved in methyl iodide at room temperature over a prolonged period, the authors were surprised to find neither the elimination product nor the cadinene derivative (the expected product arising via the methylsulfonium ion, followed by carbonium ion formation and cyclization). The product they isolated still contained the thiophenyl moiety, but had cyclized to the eudesmol derivative 549 and its dehydration product 548 as well as to the tricyclic ether 550. The researchers reasoned that within the long reaction time (64 h), hydrogen iodide had formed and initiated an electrophilic olefin cyclization. Indeed, when the authors treated the eudesmol derivative 549 with hydrogen iodide, it was converted into 548 and 550. Desulfuration of the tricyclic ether 550 resulted in racemic evuncifer ether (278).

548

H+

SPh

SPh

OH

HI

2) Na2CO3, CH3OH, 54% (2 steps)

549

HI

HO

–78°C, 60%

O

BuLi, DABCO, THF, N2,

548:549:550 = 5:9:3

SPh

OH

SPh

OH

SPh 545

O

Scheme 84 Synthesis of evuncifer ether (278) via transannular cyclization

544

SPh

1) NBS, THF, H2 O

550

SPh

SPh

546

Raney Ni

SPh

OH

O

7:5

+

278

OH 547

PhS

CH3I, rt, 64 h, 87%

Chemistry of the Secondary Metabolites of Termites 183

184

E. Gössinger

Syntheses of 5β,7β,10β-Eudesmanes The first example of the small group of naturally occurring 5β,7β,10β-eudesmanes was isolated from a plant in the year 1956 [616], and for the next 26 years, only three further compounds were obtained from plant origin [617–619]. Thus, it was surprising when Prestwich et al. reported three new all cis-eudesmanes from the termite Amitermes excellens [360]. Whereas the structures of the two unsaturated hydrocarbons were only given tentatively, the structure of the alcohol amiteol (365) was elucidated by spectrometric data including NOE and ascertained by total syntheses by Wijnberg, de Groot et al. A lengthy racemate synthesis [356] was followed by an EPC synthesis starting with ()-(α)-santonin (551). Ando et al. also used ()-(α)santonin (551) synthesizing 5β,7β,10β-eudesma-3,11-diene (366), thus confirming its structure [364].

The First Synthesis of Racemic Amiteol by Wijnberg et al. Wijnberg, de Groot et al. used their protocol for the first synthesis of 10-methyldecalin-3,6-diones for the initial synthesis of racemic amiteol (¼ 5β,7β,10β-eudesm-11-en-4α-ol) (365) [356]. The synthesis follows the pathway of the synthesis of evuncifer ether (Scheme 83) to the very late intermediate 543 (Scheme 85). Treatment by strong base for 1 min epimerized the stereogenic center at C-7 of the unstable lactol 543 partially. After separation, the keto group of the epimerized compound 552 was converted into the methylene moiety by diiodomethane, zinc, and titanium tetrachloride [620, 621] resulting in racemic amiteol (365) in 13 steps and 5.5% overall yield starting with the bicyclic enone 532.

Scheme 85 Synthesis of amiteol (365)

OH OH

O

O

t-BuOK, DMSO, rt, 1', 59% + 25% 543

543

OH

O

Zn, CH2I2, TiCl 4, THF, rt;

OH

rfl; rt, 74% 552

365

Chemistry of the Secondary Metabolites of Termites

185

EPC Synthesis of Amiteol by Wijnberg, de Groot et al. Wijnberg, de Groot et al. designed an effective partial synthesis by starting with chiral, commercially available ()-α-santonin (551) [356]. The first reductions followed known procedures. The lactone was reductively cleaved by lithium in liquid ammonia, and simultaneously the cross-conjugated double bond was hydrogenated. The resulting carboxylic acid was esterified to the methyl ester 553 using trimethylsilyl chloride in methanol [622, 623]. The removal of the keto group of 553 by the Huang-Minlon variant of the Wolff-Kishner reaction led to a mixture of products, having none or very little of the desired cis-octalin skeleton [624]. Fortunately, reduction of the tosylhydrazone with catecholborane led in good yield to the desired octalin derivative 554 [625]. Epoxidation by dimethyldioxirane led to a 4:1 mixture generated by attack from the convex face (epoxide 555) and the concave face (epoxide 556) (Scheme 86). Reductive cleavage of the mixture of epoxides by lithium aluminum hydride was accompanied by reduction of the carboxyl ester. The resulting epimeric diols 557 and 558 were separated, and the minor component 558 was esterified selectively by mesyl chloride. Exchange of the mesylate 559 by iodide to give 560 and elimination by base treatment furnished enantiopure (+)-amiteol (365) in nine steps and 6% overall yield.

O

O

O

555

2) TMSCl, CH3OH, rt, 86% (2 steps)

O

4:1

O

+ O

O

O

553

556

O

NHNH2

O

O BH

HO

rfl, 90%

LAH, THF,

NaOAc, CH3OH, 79% (2 steps)

2)

365 ((+)-amiteol)

O

S

O

O BF3•OEt2,THF

Scheme 86 EPC synthesis of (+)-amiteol ((+)-365) from ()-α-santonin (551)

NaHCO3, H2O, CH2Cl2, 76%

O , (CH3)2CO, O 18-crown-6,

551 ((–)-α-santonin)

O

O

1) Li/ NH3 liq, THF; to rt

1)

OH

58% (3 steps)

t-BuOK, t-BuOH, rfl

O

O

557

X

3.5:1

+

559 (X = MsO) 560 (X = I)

HO

1) MsCl, pyr, 40°C

OH

554

O

OH 558

HO

2) NaI, (CH3)2CO, rfl

O

186 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

187

EPC Synthesis of (+)-Eudesma-3,11-diene by Ando et al. Naya and Prestwich isolated eudesma-3,11-diene (366) and elucidated its structure mainly by 1H NMR spectrometry [360]. To confirm the relative configuration of 366, Ando et al. conducted the following partial synthesis shown in Scheme 87 [364]. Strong acid-assisted epimerization via conjugate dienol formation brought the C– O single bond of the lactone moiety of ()-(α)-santonin (551) in an axial position and thus parallel to the π-orbital of the neighboring double bond, the prerequisite for the easy reductive cleavage of the C–O bond of lactone 561 by zinc in acetic acid. Catalyzed by palladium on charcoal under basic conditions, the dienone 562 was hydrogenated to a 3:1 mixture of the cis-decalinone derivative 564 and the respective trans-derivative 563 [623]. After separation of the stereoisomers and esterification of acid 564 with diazomethane to give the methyl ester 565, reduction by lithium aluminum hydride led to a mixture of the epimeric diols 566 and 567. The diols were separated and transformed separately into the mesylates, which were eliminated by treatment with lithium bromide and lithium carbonate at high temperatures leading to the same product (+)-eudesma-3,11-diene (366). The 1H NMR spectrum of the synthetic eudesmane derivative 366, produced in seven steps and 12% overall yield, was identical with that of the natural product. No synthesis has been reported of the isomeric (5S*,7R*,10R*)-eudesma-3 (14),11-diene (367) also found in Amitermes species, to confirm the assumed structure. HCl gas, DMF,

Zn, AcOH, CH3OH,

O O

O

47%

O

O

O

562

O

H2 (1 atm) Pd/C, KOH, C2H5OH, O

CH2N2, ether, rt;

+ O

OH

O

OH

561

551 ((–)-α-santonin)

rt, 53%

85%

O

O OH

563

separation, 64% 564

OH O LAH, ether, rfl, 100%

OH

+

O OCH3 565

OH

OH 566

567

1) MsCl, pyr, rt 2) Li2CO3, LiBr, DMF, 150°C, 87% (2 steps) 366 ((+)-5ßH-eudesma-3,11-diene)

Scheme 87 Synthesis of (+)-5βH-eudesma-3,11-diene (366) from ()-α-santonin (551)

188

E. Gössinger

Syntheses of 8-Epicaparrapi Oxide Of the two minor cyclic sesquiterpene ethers 364 and 395 isolated from A. evuncifer [293], 395 has long been sought after as a fragrance and several syntheses have been designed [626, 627]. Mostly those syntheses yielded as main compound the epimeric 8-epicaparrapi oxide (364). Caparrapi oxide (395) has been isolated from plants, but the only natural source of 8-epicaparrapi oxide (364) is A. evuncifer. Thus, the syntheses will be presented, although the purpose was either the synthesis of the fragrant caparrapi oxide (395) or investigations of the potential of the acid-sensitive nerolidol (371) for biomimetic olefin cyclization.

The First Synthesis of ()-8-Epicaparrapi Oxide and () Carparrapi Oxide by Lombardi et al. Lombardi and Cookson [626, 627] used 7,8-dihydro-β-ionone (568) as starting material (Scheme 88). This compound is prepared easily from β-ionone by partial hydrogenation [628] or by cyclization of geranylacetone [629]. Addition of sodium acetylide provided the tertiary propargyl alcohol 569. Treatment of 569 with tin tetrachloride in dichloromethane at 20 C yielded in 46% a 5:1 mixture of the cyclic ethers 570 and 571 next to small amounts of the corresponding cis-fused cyclic ethers. The authors tested also the isomeric 7,8-dihydroionones as starting materials. However, in the case of α- and γ-7,8-dihydroionone, cyclization led to a muchincreased fraction of the cis-fused cyclic ethers. The separated trans-fused ethers were partially hydrogenated yielding mainly 8-epicaparrapi oxide (364) (33% overall yield) and caparrapi oxide (395). NaC2H, PhCH3, 85% 568

HO

O

569

SnCl4, CH2Cl2, O –20°C, 46% 570 H2, 1 atm, Lindlar cat., EtOH, rt, ~ 100%

O

364

O

+ 5:1

571 H2, 1 atm, Lindlar cat., EtOH, rt,

O

395

Scheme 88 Synthesis of 8-epicaparrapi oxide (364) and caparrapi oxide (395)

Chemistry of the Secondary Metabolites of Termites

189

EPC Synthesis of (R)-8-Epicaparrapi Oxide and (S)-Carparrapi Oxide by Barrero et al. Chiral 8-epicaparrapi oxide (364) was obtained by debromination of naturally occurring 3-bromo-8-epicaparrapi oxide (456) [580]. Barrero et al. chose the degradation of ()-sclareol (572), a volatile component of a Mediterranean sage, to obtain enantiopure 8-epicaparrapi oxide (364) and caparrapi oxide (395) (Scheme 89) [630, 631]. ()-Sclareol (572) was treated with a mixture of osmium tetroxide and sodium periodate. In a one-pot reaction, dihydroxylation by osmium tetroxide was followed by cleavage of the vicinal triol yielding the methyl ketone, which cyclized to the lactol and by dehydration furnished the cyclic enol ether i, which was immediately dihydroxylated and fragmented yielding the bicyclic ester aldehyde 573. The aldehyde 573 was converted into the silyl enol ether, which was treated with ozone. The generated aldehyde 574 was reduced to the diol by lithium aluminum hydride, and the primary alcohol was selectively esterified by tosyl chloride. Base treatment of the 3-hydroxy tosylate 575 led to Grob fragmentation. The resulting olefin 576 was converted into the aldehyde by ozonolysis. Subsequent Baeyer-Villiger reaction using meta-chloroperbenzoic acid provided the formate 577. Vinylmagnesium bromide transformed the keto group of 577 in the tertiary allyl alcohol, and successive treatment with potassium hydroxide in methanol saponified the ester. The epimeric diols 578 and 579 had been synthesized before in racemic form, but the attempted cyclization with strong acid in a polar solvent led to very poor yields of the desired bicyclic ethers 364 and 395 [627]. Thus, Barrero et al. aware of this result, chose a two-step method to circumvent the strong acid [630, 631]. To support palladium allyl formation, the tertiary allyl alcohol was esterified selectively to give acetates 580 and 581. Intramolecular attack by the second hydroxy group at the palladium allyl complex led in good yield to a 1:2 mixture of ()-364 and ()-395. Later on, cerium ammonium nitrate (CAN) was found to be the reagent of choice [632]. Diols 578 and 579 treated with CAN in acetonitrile led in 94% yield to 395 and 364. The authors assume that the high yield of the cyclization of the acid-sensitive tertiary allyl alcohols 578 and 579 and the necessity of equivalent amounts of CAN point to a radical reaction with this one-electron oxidant. The biosynthesis of neither 364 nor 395 has been investigated previously, but the most likely biogenesis is an electrophilic olefin cyclization of nerolidol (371) (Scheme 90). The acid sensitivity of the tertiary allylic alcohol nerolidol (371) is a considerable challenge for a biomimetic synthesis. Indeed, very early on the acid-catalyzed olefin cyclization of farnesol (370) and nerolidol (371) was investigated. However, no ether formation was detected when nerolidol was treated with formic acid in various concentrations and temperatures [633, 634], nor were the use of the Lewis acids, aluminum trichloride [634] or boron trifluoride etherate, successful [635]. Therefore, Lewis acids with low-lying LUMO were examined for the cyclization. No reaction was detected using mercury(II) trifluoroacetate [636, 637] and only moderate yields by bromination [638].

+

575

OH

579

OH

CAN, CH3CN, rt, 94%

TsO

i

O

576

2:1

+

Ac2O, DMAP, Et3N, THF, rfl

(–)-395

OH

NaH, DME, rfl, 95%

O

O

O OAc

+ 581

OH

577

O

O

2) O3, CH2Cl2, –78°C; Ph3P, –78°C to rt, 93%

1) TBSCl, NaH, THF, –78°C, 99%

PdCl2(CH3CN)2, THF, rt, 75% (2 steps)

AcO

2) MCPBA, CH2Cl2, rt, 95%

1) O3, CH2Cl2, –78°C; Ph3P, –78°C to rt, 75%

580

OH

(–)-364

O

573

Scheme 89 EPC synthesis of 8-epicaparrapi oxide (364) and caparrapi oxide (395) from ()-sclareol (572)

578

KOH, CH3OH, rt, 90%

HO

2) TsCl, DMAP, pyr

OH

OAc

i-PrOH, 45°C, 75%

OsO4, NaIO4,

1) LAH, THF, rt, 95%

C2H3MgBr, Et2O, 0°C to rt;

574

O

572

OH

OH

OAc

O

190 E. Gössinger

Chemistry of the Secondary Metabolites of Termites Scheme 90 Proposed biosynthesis of 8-epicaparrapi oxide (364) and caparrapi oxide (395)

191

HO

O

+

H

371 ((S)-nerolidol)

(–)-395

OH

O

H+ 371 ((S)-nerolidol)

(+)-364

Synthesis of ()-8-Epicaparrapi Oxide and () Carparrapi Oxide by Kato et al. Kato et al. tried to solve the problem by using 2,4,4,6-tetrabromocyclohexa-2,5dienone (TBCO) (582) for the electrophilic olefin cyclization of nerolidol (371) in nitromethane. Compound 582 releases positively charged bromine slowly, and the generated phenol is only moderately acidic (Scheme 91) [638]. 3-Bromocaparrapi oxide (583) and 3-bromo-8-epicaparrapi oxide (456) were formed in moderate yields in a 1:6 ratio at room temperature due to the smaller van der Waal’s radius of the vinyl group compared to the methyl group. After separation, reductive removal of the bromine by tributyltin hydride completed Kato’s synthesis of racemic 8-epicaparrapi oxide ()-(364) in two steps and 30% overall yield.

HO

O

Br

583

TBCO, CH3NO2,

+

HO

1 : 6

+

rt, 35%

Bu3SnH

O

Br

or LAH, dioxane, rfl

456

(±)-nerolidol (371)

O Br

Br

O Br (±)-364

Br

582 (TBCO)

Scheme 91 Biomimetic synthesis of 8-epicaparrapi oxide (364) and caparrapi oxide (395) via 3-bromo-8-epicaparrapi oxide (456)

192

E. Gössinger

Synthesis of ()-8-Epicaparrapi Oxide and ()-Carparrapi Oxide by Kametani et al. These investigators argued that the terminal trisubstituted double bond is less acidsensitive than a tertiary allyl alcohol and converting the trisubstituted double bond into a vicinal hydroxy selenide might alter the acid attack by support of the selenide in abstracting the vicinal hydroxy group [639]. Therefore, nerolidol (371) was epoxidized by peracid, and the epoxide 584 was attacked by in situ-prepared sodium phenylselenide (Scheme 92). The resulting vicinal hydroxy selenides 585 and 586 were then treated with trifluoroacetic acid. Disappointingly, the cyclization led only in 21% yield to a 1:1 mixture of 3βphenylselenocaparrapi oxide (588) and its C-8-epimer 587. The seleno group was removed by tributylstannane, and the troublesome separation of 364 and 395 was solved by selective bromination. The equatorial vinyl group of 395 added bromine much faster than the axial vinyl group of 364, thus permitting easy separation.

MCPBA, CH2Cl2, HO

(PhSe)2, HO

NaHCO3, rt

NaBH4; 584 O 584

371

HO

PhSe

HO

PhSe

+

OH

OH 585

586 TFA, CH2Cl2, 0°C, 21%

+

O

PhSe

1:1

587

O

PhSe

588

Bu3SnH

O

364

+

O 395

Scheme 92 Biomimetic synthesis of 8-epicaparrapi oxide (364) and caparrapi oxide (395) via epoxynerolidol (584)

Chemistry of the Secondary Metabolites of Termites

193

Synthesis of ()-8-Epicaparrapi Oxide by Zefirov at al. Zefirov et al. were the first research group using the Brønsted acid-activated olefin cyclization successfully to synthesize ()-8-epicaparrapi oxide (364). However, unusual reaction conditions and many trials were necessary [640]. Racemic nerolidol (371) was treated with super acid in a mixture of fluorosulfonylchloride and dichloromethane at 110 C (Scheme 93). No ether formation was detected with higher acid concentration, but decreasing the ratio of acid to nerolidol (371) to 5:1 yielded the desired 8-epicaparrapi oxide (364) in 80% yield. It should be noted that at the very low temperature used, the small differences in steric hindrance of the two conformations of nerolidol (371) in the presumed transition state sufficed to produce only 364. HSO3F, CH2Cl2, SO2ClF, HO

+

(R)-371

O

(–)-364

OH –110°C, 80%

(S)-371

+

O

(+)-364

Scheme 93 Biomimetic synthesis of 8-epicaparrapi oxide (364) catalyzed by a Brønsted acid

194

E. Gössinger

EPC Synthesis of (R)-8-Epicaparrapi Oxide and (S)-Carparrapi Oxide by Yamamoto et al. Yamamoto’s long-standing research in developing catalysts for the biomimetic electrophilic olefin cyclization of isoprenes and isoprenoids led him and his research group to investigate the difficult cyclization of nerolidol (371) [641–643]. The research group had developed Lewis acid-assisted chiral Brønsted acids (LBAs) and chose the chiral tin tetrachloride complex (S)-LBA (589) containing a phenolic proton, as shown in Scheme 94. They observed that ()-nerolidol (371) cyclized at 78 C with high enantioselectivity for the matched pair but much lesser enantioselectivity in case of the mismatched pair (Scheme 95). Despite the low temperature, the acid sensitivity of the tertiary allyl alcohol prevented acceptable yields. Thus, the investigators tried to reduce the acid sensitivity by supplanting the vinyl group by its product of hydration. To attain the 1,3-diol 593, they chose (E,E)-farnesol (370) as starting material. Using the Sharpless-Katsuki protocol, they converted the terminal allyl alcohol into the chiral epoxide 590 (Scheme 96). Reduction with sodium bis(2-methoxyethoxy)aluminum hydride (Redal®) furnished the desired chiral 1,3-diol 591 in excellent yield and enantiomeric excess. This diol turned out to be a stronger complex-forming reagent for tin (IV) than the chiral complex (S)-LBA (589). Thus, the primary alcohol had to be protected. The electron-withdrawing ability of the silyl group and even of the aliphatic acyl group did not suffice to attain the catalyzed cyclization. The additional complexing ability of an aromatic group was necessary to attain the cyclization. Best results were achieved with the phenylacetate 592 and a 1:1 mixture of dichloromethane and propyl chloride as solvent at 80 C. Using the (S)-configured tertiary alcohol 592 and (S)-LBA ((S)-589), (S)-8-epicaparrapi oxide phenylacetate 593 was generated in 98% enantiomeric excess. Saponification of the ester 593 by lithium hydroxide, dehydration according to Grieco via selenide 594 [644], and cis elimination after oxidation to the selenoxide afforded (+)-8-epicaparrapi oxide (S)-364 (98% ee) and ca 10% of 395 (27% ee). The use of (R)-LBA ((R)-589) and the (S)-configured tertiary alcohol 592 generated 395 in good yield and excellent enantiomeric excess.

F

O

SnCl4 = (S)-LBA

O H

OCH3 589

Scheme 94 Structure of the chiral Brønsted acid catalyst (589)

Chemistry of the Secondary Metabolites of Termites

HO

195 (R)-589, PhCH3, –78°C,

HO

+

364:395 ~ 2.5:1, 13% (R)-371

(S)-371

O

O

+

(S)-364

(R)-364

O

O

+

(R)-395

(S)-395

(R)-364:(R)-395 = >99:45% (3 steps)

1) KOH, C2H5OH/H2O rfl

742

O

–78°C to rt

3:1

+

LDA, THF, -78°C; TBSCl, THF/HMPA,

CO2TBS

O

O

OH

OH

749

OAc

746 CO2CH3

OTBS

2) AcOH, THF, H2O

1) (C2H5)2AlCN, PhCH3, –20°C to rt, 82%

739

238 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

239

secotrinervita-7,11-diene (733), a defense substance of Nasutitermes princeps [707], was the ring closure of the 1,4-trans-bridged (annulated) six-membered ring by Dieckmann condensation [702, 706]. To achieve this goal, the isopropenyl group had to be removed, a problem the authors had solved in an earlier synthesis [710]. The chloride of 134 was exchanged by acetate in acetic acid with zinc oxide as catalyst in 71% yield. Treatment of the acetate with lithium hydroxide in aqueous dioxane at higher temperature led to saponification and in consequence to retroaldol reaction. This retroaldol reaction was accompanied by deconjugation and (Z )/(E) isomerization of the conjugated double bond. Recycling of the isomerized products increased the yield of conjugated ketone 737 to a respectable 70%. The following reduction to the alcohol and acetylation yielded the monocyclic allylic acetate 738. To construct the quaternary carbon center via Claisen-Ireland rearrangement, the ester enolate was prepared by deprotonation with strong base at low temperature and stabilization as a silyl enol ester. Increasing the temperature led to the desired rearrangement. The silyl ester 739 was converted into the methyl ester 740 by acidic hydrolysis and esterification with diazomethane. Attempts to epoxidize the triene 740 selectively failed. Thus, the ester was reduced with lithium aluminum hydride, and directed epoxidation according to Sharpless et al. with t-butyl hydroperoxide catalyzed by vanadium acetylacetonide led to a mixture of products when conducted at room temperature [669, 670]. Utilizing refluxing benzene in the procedure, only the two epimeric epoxides 741 were obtained. Despite the supposed directing effect of the primary hydroxy group, equal amounts of both epimers were found, which refers to two equally populated conformations. The two epimeric epoxides could be separated, and consequently the relative configurations were determined by butanolide formation. For the planned synthesis of secotrinervitane, the primary alcohols of the mixture of epoxides 741 were protected, and subsequently the epoxides cleaved. The allylic alcohols obtained were oxidized to the unsaturated ketone 742. This allowed the functionalization by 1,4-addition. With Nagata’s reagent a 1:1 mixture of the epimeric cyanides 743 was obtained in good yields [711]. After deprotection, the primary alcohol was oxidized to the acid by Jones reagent and esterified by diazomethane yielding esters 744. Surprisingly, the consecutive reduction of the keto group of 744 with lithium aluminum tri-t-butoxy hydride led to a single alcohol in high yield that spontaneously cyclized to γ-lactones 745. The next step, basic hydrolysis of the lactone as well as the nitrile, led to a mixture of diacids, all possible γ-lactones and starting material. After esterification with diazomethane, the complex mixture was separated from the desired diester 746 and recycled repeatedly. Despite the highly basic conditions, no epimerization at the nitrile and consequently the ester could be detected, and thus improvement of the yield of 746 above 50% was not possible. The secondary alcohol of 746 was protected as a silyl ether before the Dieckmann condensation could be obtained in surprisingly high yield utilizing potassium t-butylate as base. Again, no epimerization in the α-position to the carbonyl group of β-keto ester 747 could be detected although the trans-bridged cyclohexanone is more strained than the corresponding cis-bridged cyclohexanone. The silyl ether of 747 was hydrolyzed followed by decarboxylation with sodium chloride in refluxing dimethyl sulfoxide.

240

E. Gössinger

The hydroxy group of the bicyclic keto alcohol 748 was acetylated, and the keto group was transformed into the tertiary alcohol by attack of methyllithium, which led exclusively to the undesired α-alcohol 749. By using Yamamoto’s very bulky catalyst methylaluminum bis-(2,6-di-t-butyl-4-methylphenoxide) (MAD) [712], the epimeric β-alcohol 733 became the major product, which was in its analytical data indistinguishable from natural 3α-acetoxy-15β-hydroxy-7,16-secotrinervita-7,11diene (733). This synthesis required 25 steps and the overall yield was less than 1%. In the same year, Kato et al. published their elegant and short biomimetic synthesis of ()-secotrinerviten-2β,3α-diol (724) as a communication [701]. The synthesis was presented again as full paper [703] supplemented by the synthesis approaches toward the stereoisomers of 724.

Biomimetic Synthesis of (1R*,2R*,3R*,4S*,7E,11E)-7,16-Secotrinervita-7,11,15 (17)-triene-2,3-diol As shown in Scheme 122, the starting material was 2-neocembrenone 135 [228, 667, 668], which was reduced by diisobutylaluminum hydride at 78 C. To epoxidize the generated neocembrenol regio- and stereoselectively, the researchers used the Sharpless protocol [669, 670] and obtained the desired epoxide 643 [667, 668] in 76% yield. The attempt to cleave the hydroxy epoxide led to mixtures of products. Protection of the hydroxy group as acetate 750 and consecutive treatment with boron trifluoride etherate in ether opened the epoxide, and the generated tertiary carbocation attacked the isopropenyl group. Proton abstraction completed the cyclization forming ()-2β-acetoxysecotrinerviten-3α-ol (751) in 82% yield. Removing the acetyl group by reduction with lithium aluminum hydride furnished racemic secotrinerviten-2β,3α-diol (724), thus supporting the biosynthesis hypothesis impressively.

1) DIBAH, –78°C 2) t-BuO2H,

Ac2O, DMAP,

O

H

OH VO(acac)2, benzene, 5°C to rt, 76%

pyr, CH2Cl2, rt

OAc

O

O

643

135

BF3•OEt2, –20°C,

750

LAH, ether, 0°C, 93%

82% (2 steps) OAc OH 751

Scheme 122 Biomimetic synthesis of secotrinervitane 724

OH OH 724

LA

Chemistry of the Secondary Metabolites of Termites

241

The Trinervitanes The tricyclic trinervitanes in constituting around 70 compounds (Table 11) are the most abundant of the polycyclic diterpenes derived from neocembrene. They are found in the secretion of the frontal gland of the soldiers of most Nasutitermitinae, the most highly developed termites. Soldiers squirt the secretion from the nasus (nozzle) of their pear-shaped heads. This secretion solidifies immediately thus immobilizing smaller enemies especially ants. Prestwich et al. selected soldiers of Trinervitanus gratiosus and T. bettonianus and extracted the crushed heads with an ether/n-hexane mixture [316, 715, 722]. After filtration and evaporation, chromatography of the residue on Florisil with an ethyl acetate/benzene mixture separated the monoterpenes from the cyclic diterpenes. The fraction containing the diterpenes crystallized on slow evaporation from ether [715], permitting structure elucidation by X-ray analysis [729]. This was especially fortunate as the complex structure contained unexpected features, being an ansa-compound with a tight seven-membered handle (bridge) spanned over a cishydrindene, with a double bond at the bridgehead position. The 1H NMR spectrum showed a highly complex signal pattern, which would not have permitted structure elucidation by NMR spectrometry at the given time, as the few assignments of signals in the published spectra (Table 12) of trinervi-2β,3α,9α-triol 9-O-acetate (763) demonstrate [715, 722]. The absolute configuration was determined by circular dichroism measurements of a praseodymium complex [722], a method developed by Nakanishi et al. [730]. Several more trinervitanes in small amounts (725, 752, 761, 782, 803) were isolated from the mother liquor of the above-mentioned crystallization, and their structures were determined by comparison of their spectra with those of 763, especially the 1H NMR and 13C NMR data. The same absolute configuration was assumed. Indeed, in any case where the absolute configuration of a newly described trinervitane was examined, either by Nakanishi’s method [311, 722] or by shift differences of the 1H NMR signals of the diastereomeric esters of trinervitane alcohols with chiral acids [311, 713] or kinetically by a modification of Horeau’s method [265], the absolute configuration was the same as found in neocembrene (2) and in the trinervitane 763. After this first successful isolation and structure determination of a polycyclic diterpene, the number of characterized polycyclic diterpenes from termites has risen to 87 compounds, 69 of them trinervitanes. Their isolation procedures used are similar to that used by Prestwich. Soldiers or their heads were crushed and extracted with n-hexane or dichloromethane, although in a few cases in mixtures of less polar solvents, several in dichloromethane/ methanol mixtures, and one case with methanol. One author “milked” the soldiers with a capillary, circumventing the extraction. Separation from mono- or much less frequently from sesquiterpenes was achieved by filtration through an adsorbent or by short-path distillation. The further purifications were accomplished by chromatography on Florisil, silica gel, or alumina. In several cases, repeated chromatographic steps including HPLC were necessary due to a larger number of similarly substituted trinervitanes. For example, Lacessititermes laborator ejects a secretion containing

242

E. Gössinger

Table 11 Trinervitanes Name (Nr.), formula, structure (3S,4S,7R,12S,16S)Trinervita-1(15),8(19)dien-3-ol (752) C20H32O

Termite genera Bulbitermes, Grallatotermes, Hospitalitermes, Lacessititermes, Nasutitermes, Subulitermes, Trinervitermes, Velocitermes

Isolation, notes CH2Cl2 extract of crushed termite soldiers; RP-HPLC [336]

Grallatotermes, Hospitalitermes, Nasutitermes, Subulitermes, Trinervitermes

n-Hexane extract of soldiers; LC on Florisil Structure was assigned originally to the compound isolated from T. species [337, 715] but was revised to compound 752 [713]

HO

(4S*,7R*,9S*,12R*, 16S*)-Trinervita-1(15), 8(19)-dien-9-ol (753) C20H32O OH

(4S*,7R*,9S*,12R*,16- Nasutitermes S*)-Trinervita-1(15), 8(19)-dien-9-yl acetate (754) C22H34O2

CH2Cl2 extract of soldiers; LC on silica gel

Structure determination Mp, [α]D, IR, MS, 1H and 13C NMR, absolute configuration determined by a modified Horeau method, NMR shift difference of Mosher’s ester [311] and of the praseodymium complex [311] Mp, IR, UV, and 1 H and 13C NMR Caution is advised because in most cases, the structure was determined by comparison with a compound isolated from T. bettonianus [715] which was revised to compound 752 in 1990 [713] Mp, MS, IR, and comparison with acetylated compound 753 [716]

Ref. [32, 111, 265, 306, 309–311, 315, 323, 324, 326, 336, 713, 714]

[32, 295, 296, 300, 302, 306, 314, 316, 317, 335, 337, 376, 700, 705, 708, 709, 715–717]

[32, 306, 700, 709]

OAc

(continued)

Chemistry of the Secondary Metabolites of Termites

243

Table 11 (continued) Name (Nr.), formula, structure Termite genera (2S,3R,4S,7R,12S,16S)- Nasutitermes, Trinervita-1(15),8(19)- Velocitermes diene-2,3-diol (755) C20H32O2

Isolation, notes CH2Cl2 extract of crushed termite soldiers; RP-HPLC [336]

HO OH

(2R,3R,4S,7R,12S,16S)-Trinervita-1(15), 8(19)-diene-2,3-diol (725) C20H32O2

Bulbitermes, Grallatotermes, Hospitalitermes, Longipeditermes, Nasutitermes, Subulitermes, Trinervitermes

CH2Cl2 extract of crushed termite soldiers; HPLC Easily oxidized to compound 764 Most abundant trinervitane

Lacessititermes, Nasutitermes

n-Hexane extract of crushed soldier heads

HO OH

(2R*,3R*,4S*,7R*, 12S*,16S*)-2Hydroxytrinervita1(15),8(19)-dien-3-yl acetate (756) C22H34O3

Structure determination Mp, [α]D, IR, 1 H, 13C, and 2D NMR Structure confirmed by racemate synthesis [718] Absolute configuration determined by CD of the Pr complex Mp, [α]D, MS, IR, Raman, 1 H, and 13C NMR Structure confirmed by racemate synthesis [718] Absolute configuration determined by CD of the praseodymium complex Mp depends on the solvent of crystallization (CH3OH or nhexane) Mp, MS, IR, and 1 H and 13C NMR

Ref. [32, 111, 295, 300, 302, 309, 311, 324, 336, 705, 719]

[32, 111, 295, 300, 302, 304, 305, 308– 317, 323, 326, 335, 336, 353, 376, 700, 708, 709, 714–717, 719]

[32, 111, 716]

AcO OH

(continued)

244

E. Gössinger

Table 11 (continued) Name (Nr.), formula, structure (2R*,3R*,4S*,7R*, 12S*,16S*)-3Hydroxytrinervita1(15),8(19)-dien-2-yl acetate (757) C22H34O3

Structure determination Mp, [α]D, MS, IR, and 1H and 13 C NMR

Termite genera Bulbitermes, Hospitalitermes, Lacessititermes, Longipeditermes, Nasutitermes

Isolation, notes CH2Cl2 extract of crushed termite soldiers; HPLC [336]

Ref. [32, 111, 300, 304, 305, 310, 315, 317, 323, 326, 335–337, 714]

Nasutitermes

Soldiers extracted with CH2Cl2; repeated LC on silica gel

Mp, MS, IR, and 1 H and 13C NMR

[32, 111, 295, 300, 700, 716]

(2S,3S,4S,7R,12S,16S)- Nasutitermes, Trinervita-1(15),8(19)- Subulitermes, Velocitermes diene-2,3-diol (759) C20H32O2

CH2Cl2 extract of crushed termite soldiers, HPLC Active against methicillinresistant bacteria

Mp, [α]D, IR, MS, and 1H NMR Absolute configuration determined by CD of the Pr complex

[32, 111, 295, 296, 300, 302, 309, 311, 322, 324, 336, 376, 705, 719]

HO OAc

(2R*,3R*,4S*,7R*, 12S*,16S*)-Trinervita1(15),8(19)-diene-2,3diyl diacetate (758) C24H36O4

AcO OAc

HO OH

(continued)

Chemistry of the Secondary Metabolites of Termites

245

Table 11 (continued) Name (Nr.), formula, structure (2R,3R,4S,7R,9S, 12S,16S)-2,3Dihydroxytrinervita1(15),8(19)-dien-9-yl acetate (760) C22H34O4

Structure determination MS, IR, 1H and 13 C NMR, acetylation

Termite genera Nasutitermes

Isolation, notes CH2Cl2, LC on silica gel

Ref. [111, 720]

Bulbitermes, Hospitalitermes, Lacessititermes, Longipeditermes, Nasutitermes

n-Hexane extract of crushed soldier heads

MS, IR, 1H and 13 C NMR, NOE, and 2D-NMR

[32, 111, 304, 305, 310, 315– 317, 323, 714, 715]

Bulbitermes, Hospitalitermes, Lacessititermes, Longipeditermes, Nasutitermes

n-Hexane extract of crushed soldier heads; LC on Florisil

Mp, [α]D, MS, IR, 1H and 13C NMR, NOE, and 2D-NMR

[32, 111, 295, 296, 300, 304, 305, 310, 315, 317, 323, 335, 700, 708, 714, 716, 720]

OAc

HO OH

(2R*,3R*,4S*,7R*, 9R*,12S*,16S*)-9Hydroxytrinervita1(15),8(19)-diene-2,3diyl diacetate (761) C24H36O5 OH

AcO OAc

(2R*,3R*,4S*,7R*, 9R*,12S*,16S*)Trinervita-1(15),8(19)diene-2,3,9-triyl triacetate (762) C26H38O6 OAc

AcO OAc

(continued)

246

E. Gössinger

Table 11 (continued) Name (Nr.), formula, structure (2R*,3R*,4S*,7R*, 9R*,12S*,16S*)-2,3Dihydroxytrinervita1(15),8(19)-dien-9-yl acetate (763) C22H34O4

Termite genera Bulbitermes, Grallatotermes, Hospitalitermes, Lacessititermes, Longipeditermes, Nasutitermes, Trinervitermes

Isolation, notes n-Hexane/ether extracts of soldier heads; LC on Florisil

Bulbitermes, Grallatotermes, Hospitalitermes, Lacessititermes, Longipeditermes, Nasutitermes, Trinervitermes

CH2Cl2 extract of crushed termite soldiers; RP-HPLC [336] Possibly produced by the isolation of the oxygensensitive compound 725 Active against methicillinresistant Staphylococcus aureus

Nasutitermes

CH2Cl2 extract of soldiers and “milking” of the frontal gland secretion; LC on silica gel several times

OAc

Structure determination MS, IR, [α]D, 1H and 13C NMR, and X-ray analysis; absolute configuration determined by CD of the Pr complex

HO

Ref. [32, 111, 300, 304, 305, 308, 310, 311, 313–315, 317, 323, 335, 337, 353, 700, 704, 708, 714, 715, 717, 720, 721, 722]

OH

(3R,4S,7R,12S,16S)-3Hydroxytrinervita1(15),8(19)-dien-2-one (764) C20H30O2

HO O

(3R*,4S*,7R*,9R*, 2S*,16S)-3,9Dihydroxytrinervita1(15),8(19)-dien-2-one (765) C20H30O3 OH

Mp, [α]D, MS, IR, UV, 1H, 13C, and DEPT-135 NMR, NOESY, quantum mechanical calculations, and partial synthesis by MnO2 oxidation of compound 725 Configuration of C-3 in the compound isolated from Grallotermes africanus soldiers not determined [314] [α]D, MS, IR, UV, and 1H NMR

[32, 111, 300, 302, 304, 305, 308–314, 316, 317, 323, 326, 335, 336, 700, 708, 714, 717, 719]

[111, 708]

HO O

(continued)

Chemistry of the Secondary Metabolites of Termites

247

Table 11 (continued) Name (Nr.), formula, structure (3R*,4S*,7R*,9R*, 12S*,16S)-2-Oxo-3hydroxytrinervita1(15),8(19)-dien-9-yl acetate (766) C22H32O4

Termite genera Nasutitermes, Trinervitermes

Isolation, notes CH2Cl2 extract of soldiers; repeated LC on silica gel

Structure determination [α]D, MS, IR, UV, and 1H NMR

Ref. [32, 111, 309, 700, 708, 717]

OAc

HO O

(2R*,3R*,4S*,7R*, Nasutitermitinae 12S*,13R*,16S*)Trinervita-1(15),8(19)diene-2,3,13-triol (767) C20H32O3

HO

Not isolated from termites [296], but its easy oxidation to compound 770 may have prevented its characterization [719]

Data obtained by basic hydrolysis of compound 768 [716]: MS and IR

[296, 306]

n-Hexane extract of crushed soldier heads

Mp, MS, IR, 1H and 13C NMR, and hydrolysis to the triol [716, 719]

[111, 296, 300, 309, 716]

OH OH

(2R*,3R*,4S*,7R*, 12S*,13R*,16S*)Trinervita-1(15),8(19)diene-2,3,13-triyl triacetate (768) C26H38O6

AcO

Nasutitermes

OAc OAc

(continued)

248

E. Gössinger

Table 11 (continued) Name (Nr.), formula, structure (2R*,3R*,4S*,7R*, 12S*,14S*,16S*)Trinervita-1(15),8(19)diene-2,3,14-triyl triacetate (769) C26H38O6

Structure determination Mp, MS, IR, 1H and 13C NMR, COSY, and NOE

Termite genera Nasutitermes

Isolation, notes CH2Cl2 extract of crushed soldiers; LC on silica gel and TLC with AgNO3coated silica gel C-14 epimer mentioned as a compound of a Nasutitermes species [723]

Ref. [32, 111, 309]

Nasutitermes

CH2Cl2 extract of crushed soldiers; LC on silica gel and preparative TLC on silica gel

MS, IR, and 1H and 13C NMR

[32, 111, 296, 719]

Bulbitermes, Hospitalitermes, Lacessititermes, Longipeditermes, Nasutitermes

CH2Cl2 extract of soldiers; LC on silica gel or HPLC

Mp, MS, IR, 1H and 13C NMR, and CD

[32, 111, 305, 310, 315, 317, 323, 335, 714, 716, 719]

AcO OAc OAc

(3R*,4S*,7R*,12S*, 13R*,16S*)-3,13Dihydroxytrinervita1(15),8(19)-dien-2-one (770) C20H30O3

HO

OH O

(2R*,3R*,4S*,7R, 12S*,16S*)-13-Oxotrinervita-1(15),8(19)diene-2,3-diyl diacetate (771) C24H34O5

AcO

O OAc

(continued)

Chemistry of the Secondary Metabolites of Termites

249

Table 11 (continued) Name (Nr.), formula, structure (2R*,3R*,4S*,7R*, 9R*,12S*,13R*,16S*)Trinervita-1(15),8(19)diene-2,3,9,13-tetrol (772) C20H32O4

Isolation, notes CH2Cl2 extract of soldiers and “milking” of the frontal gland secretion; LC on silica gel several times, GLC, and TLC

Lacessititermes

n-Hexane extract of soldiers; repeated HPLC

MS and 1H NMR; structure tentatively assigned

[32, 714]

Lacessititermes

n-Hexane extract of crushed soldiers; repeated HPLC

MS and 1 H-NMR; structure tentatively assigned

[32, 714]

OH

HO

Structure determination [α]D, MS, IR, and 1H NMR; stereochemistry only tentatively given in [708]

Termite genera Nasutitermes

Ref. [32, 111, 708]

OH OH

(2R*,3R*,4S*,7R*, 9R*,12S*,14S*,16S*)9-Hydroxytrinervita1(15),8(19)-diene2,3,14-triyl triacetate (773) C26H38O7 OH

AcO OAc OAc

(2R*,3R*,4S*,7R*, 9R*,12S*,14S*,16S*)9-Hydroxytrinervita1(15),8(19)-diene2,3,9,14-tetrayl tetraacetate (774) C28H40O8 OAc

AcO OAc OAc

(continued)

250

E. Gössinger

Table 11 (continued) Name (Nr.), formula, structure (2R*,3R*,4S*,7R*, 9S*,12S*,14S*,16S*)9-Hydroxytrinervita1(15),8(19)-diene2,3,14-triyl triacetate (775) C26H38O7

Structure determination MS and 1H NMR; structure characterized only tentatively

Termite genera Lacessititermes

Isolation, notes n-Hexane extract of crushed soldiers; repeated HPLC

Ref. [32, 714]

Lacessititermes, Nasutitermes

n-Hexane extract of crushed soldiers; repeated HPLC

[α]D, MS, IR, 1 H, and 13C NMR Stereochemistry at C-9 and C-13 tentatively assigned in [708]

[32, 111, 700, 708, 714, 720]

Nasutitermes

CH2Cl2 extract of soldiers; LC on silica gel

MS, IR, 1H, and 13 C NMR Stereochemistry at C-9 and C-13 tentatively given in [708]

[32, 111, 708, 720]

OH

AcO OAc OAc

(2R*,3R*,4S*,7R*, 9R*,12S*,13R*,16S*)Trinervita-1(15),8(19)diene-2,3,9,13-tetrayl tetraacetate (776) C28H40O8 OAc

OAc

AcO OAc

(2R*,3R*,4S*,7R*, 9R*,12S*,13S*,16S*)Trinervita-1(15),8(19)diene-2,3,9,13-tetrayl tetraacetate (777) C28H40O8 OAc

AcO

OAc OAc

(continued)

Chemistry of the Secondary Metabolites of Termites

251

Table 11 (continued) Name (Nr.), formula, structure Termite genera (2R*,3R*,4S*,7R*, Nasutitermes 9R*,12S*,14S*,16S*)2,3Dihydroxytrinervita1(15),8(19)-diene-9,14diyl diacetate (778) C24H36O6

Isolation, notes CH3OH/CH2Cl2 extract; repeated LC on silica gel

Structure determination HREIMS, IR, UV,1H and 13C NMR, COSY, HMQC, HMBC, and NOESY

Ref. [326]

OAc

HO OH

OAc

(2R*,3R*,4S*,7R*, 9R*,12S*,14S*,16S*)3-Hydroxytrinervita-1 (15),8(19)-diene2,9,14-triyl triacetate (779) C26H38O7

Nasutitermes

CH3OH/CH2Cl2 extract; repeated LC on silica gel

HREIMS, IR, UV, 1H and 13C NMR, COSY, HMQC, HMBC, and NOESY

[326]

Nasutitermes

CH3OH/CH2Cl2 extract of crushed soldiers; repeated LC on silica gel

HREIMS, IR, UV, 1H and 13C NMR, COSY, HMQC, HMBC, and NOESY

[326]

OAc

HO OAc OAc

(2R*,3R*,4S*,7R*,9E,12S*,14S*,16S*)2-Hydroxytrinervita1(15),8(19),9-triene3,14-diyl diacetate (780) C24H34O5

AcO OH

OAc

(continued)

252

E. Gössinger

Table 11 (continued) Name (Nr.), formula, structure (2R*,3R*,4S*,7R*, 9E,12S*,14S*, 16S*)2,3Dihydroxytrinervita1(15),8(19),9-trien-14yl acetate (781) C22H32O4

Structure determination HREIMS, 1H and 13C NMR, HMQC, HMBC, NOESY, and comparison with the spectra of compound 780 Acetylation of compound 781 yielded 780

Termite genera Nasutitermes

Isolation, notes CH3OH/CH2Cl2 extract of crushed soldiers; repeated LC on silica gel

Ref. [32, 111, 326]

Trinervitermes

n-Hexane/ether extracts of soldier heads; LC on Florisil

MS, Raman IR, 1 H, and 13C NMR

[32, 111, 308, 715]

Nasutitermes

CH2Cl2 extracts of soldiers; LC on silica gel, GC, and TLC

MS, IR, 1H, and 13 C NMR

[32, 111, 709]

HO OH

OAc

(2R*,3R*,4S*,7R*, 12S*,16S*)-2,3Dihydroxytrinervita1(15),8(19)-dien-17-yl acetate (782) C22H34O4 AcO

HO OH

(2R*,3R*,4S*,7R*, 12S*,16S*)-2,3Dihydroxytrinervita1(15),8(19)-dien-20-yl acetate (783) C22H34O4

OAc

HO OH

(continued)

Chemistry of the Secondary Metabolites of Termites

253

Table 11 (continued) Name (Nr.), formula, structure (3S*,4S*,5R*,7R*, 12S*,16S*)-3,5Dihydroxytrinervita1(15),8(19)-diene (784) C20H32O2

Termite genera Nasutitermes

Isolation, notes CH3OH/water extracts of powdered termites (mainly soldiers); HPLC Active against Gram-positive B. subtilis

Structure determination ESMS, EIMS, 1 H and 13C NMR, COSY, APT, HMQC, NOESY, and GOESY

Ref. [724]

HO

HO

Nasutitermes (3S*,4S*,5R*,7R*, 12S*,16S*)-3,5,18Trihydroxytrinervita1(15),8(19)-diene (785) C20H32O3

HO HO HO

(3S*,4S*,5R*,7R*, 12S*,16S*)-3,18Dihydroxytrinervita1(15),8(19)-dien-5-yl acetate (786) C22H34O4

Nasutitermes

CH3OH/water extracts of powdered termites (mainly soldiers); HPLC Active against Gram-positive bacteria [e.g., B. subtilis (MIC  50 μg/ cm3)]

CH3OH/water extracts of powdered termites (mainly soldiers) with sonication; HPLC

ESMS, HREIMS, 1H and 13C NMR, COSY, APT, HMQC, NOESY, and GOESY Absolute configuration assumed because all tested trinervitanes to date have the same absolute configuration EIMS, 1H and 13 C NMR, COSY, APT, HMQC, NOESY, GOESY, and comparison with partially acetylated triol (785)

[724]

[724]

AcO HO HO

(continued)

254

E. Gössinger

Table 11 (continued) Name (Nr.), formula, structure (2R*,3R*,4S*,7R*, 9R*,11R*,12R*,16S*)Trinervita-1(15),8(19)diene-2,3,9,11-tetrayl tetraacetate (787) C28H40O8

Structure determination MS, IR, 1H, and 13 C NMR

Ref. [32, 111, 714, 720]

CH2Cl2 extract of soldiers; LC on silica gel

MS, IR, 1H, and 13 C NMR

[32, 111, 720]

CH3OH/CH2Cl2 extract; repeated LC on silica gel

HREIMS, IR, 1 H and 13C NMR, COSY, HMQC, HMBC, and NOESY

[326]

Termite genera Lacessititermes, Nasutitermes

Isolation, notes CH2Cl2 extract of soldiers; LC on silica gel and repeated HPLC

Nasutitermes

OAc OAc

AcO OAc

(2R*,3R*,4S*,7R*, 9R*,11S*,12R*,16S*)Trinervita-1(15),8(19)diene-2,3,9,11-tetrayl tetraacetate (788) C28H40O8 OAc OAc

AcO OAc

(2R*,3R*,4S*,7S*, Nasutitermes 11S*,12R*,14S*,16S*)11,14Dihydroxytrinervita1(15),8(19)-diene-2,3diyl diacetate (789) C24H36O6

OH

AcO OAc OH

(continued)

Chemistry of the Secondary Metabolites of Termites

255

Table 11 (continued) Name (Nr.), formula, structure (2R*,3R*,4S*,7R*, 11R*,12S*,13R*,16S*)-11,14-Trinervita1(15),8(19)-dien2,3,11,13-tetrayl tetraacetate (790) C28H40O8

Termite genera Lacessititermes

Structure determination MS and 1H NMR

Ref. [32, 714]

CH2Cl2 extracts of soldiers; LC on silica gel, GC, and TLC

MS, IR, and 1H NMR

[32, 111, 709]

CH2Cl2 extract of crashed soldier heads and soldiers; LC on silica gel

Mp, [α]D, IR, MS, 1H NMR, and X-ray analysis

[32, 111, 322]

Isolation, notes CH2Cl2 extract of soldiers; LC on silica gel and repeated HPLC

OAc

AcO

OAc OAc

Nasutitermes (2R*,3R*,4S*,7R*, 9S*,12R*,16S*)-2,3Dihydroxytrinervita1(15),8(19)-diene-9,20diyl diacetate (791) C24H36O6 OAc OAc

HO OH

(2S*,3S*,4S*,7R*, 8S*,12S*,16S*)-3,8Epoxytrinervit-1(15)en-2-ol (792) C20H32O2

Nasutitermes

O

OH

(continued)

256

E. Gössinger

Table 11 (continued) Name (Nr.), formula, structure Termite genera (2S*,3S*,4S*,7R*, Nasutitermes 128R*,9S*,12S*,16S*)3,8-Epoxytrinervit1(15)-ene-2,9-diol (793) C20H32O3

Isolation, notes CH3OH extract of crushed soldiers; LC on silica gel

OH

Structure determination HREIMS, 1H and 13C NMR, COSY, HMQC, HMBC, NOESY, and comparison with the spectra of compound 792

Ref. [32, 326]

O

OH

(2R*,3R*,4S*,7S*, 8R*,9R*,12S*,14S*, 16S*)-17-Acetoxy8,19-epoxy-7hydroxytrinervit-1(15)ene-2,3,9,14-tetrayl tetrapropionate (794) C35H52O12 HO

Hospitalitermes

n-Hexane extract of crushed soldier heads; LC on silica gel, followed by HPLC

MS, 1H and 13C NMR, COSY, and ROESY

[32, 315, 725]

Hospitalitermes

n-Hexane extract of crushed soldier heads; LC on silica gel, followed by HPLC

MS, 1H and 13C NMR, COSY, and ROESY

[32, 315, 725]

O O CEt 2 AcO

EtCO2 O2CEt

EtCO2

(2R*,3R*,4S*,7S*, 8S*,9R*,12S*,14R*, 16S*)-17-Acetoxy8,19-epoxy-7hydroxytrinervit-1(15)ene-2,3,9,14-tetrayl tetrapropionate (795) C35H52O12 O O2CEt HO

AcO

EtCO2 EtCO2

O2CEt

(continued)

Chemistry of the Secondary Metabolites of Termites

257

Table 11 (continued) Name (Nr.), formula, structure (2R*,3R*,4S*,7S*, 8S*,9S*,12S*,14S*, 16S*)-17-Acetoxy8,19-epoxy-7hydroxytrinervit-1(15)ene-2,3,9,14-tetrayl tetrapropionate (796) C35H52O12

Termite genera Hospitalitermes

Isolation, notes n-Hexane extract of crushed soldier heads; LC on silica gel, followed by HPLC

Structure determination MS, 1H and 13C NMR, COSY, ROESY, and NOESY

Ref. [32, 315, 725]

O O2CEt HO

AcO

EtCO2 O2CEt

EtCO2

Hospitalitermes (2R*,3R*,4S*,7S*, (marked intraspe8S*,9R*,12S*,14S*, cific variations) 16S*)-8,19-Epoxy-7hydroxytrinervit-1(15)ene-2,3,9,14,17-pentayl pentapropionate (797) C35H52O12

n-Hexane extract of crushed soldier heads; LC on silica gel, followed by HPLC

MS, 1H and 13C NMR, COSY, ROESY, and NOESY

[32, 111, 315, 725– 727]

n-Hexane extract of crushed soldier heads; LC on silica gel, followed by HPLC

MS, 1H and 13C NMR, COSY, and NOESY; configuration at C-17 not determined

[32, 111, 315, 725, 727]

O O CEt 2

HO

EtCO2

EtCO2 O2CEt

EtCO2

(2R*,3R*,4S*,7S*, 8S*,9R*,12S*,14S*, 16S*)-17-Acetoxy8,19-epoxy-7-hydroxy17-methyltrinervit-1 (15)-ene-2,3,9,14tetrayl tetrapropionate (798) C35H52O12

Hospitalitermes (marked intraspecific variations)

O O CEt 2

HO

AcO

EtCO2 EtCO2

O2CEt

(continued)

258

E. Gössinger

Table 11 (continued) Name (Nr.), formula, structure Termite genera (2R*,3R*,4S*,7S*, Hospitalitermes 8S*,9R*,12S*,14S*, 16S*)-17-Acetoxy8,19-epoxy-7-hydroxy17-methyl-2,3,14tripropanoyloxytrinervit-1(15)-en-9yl-(2-propanoyloxy) acetate (799) C38H56O14 HO

O O RO

Isolation, notes n-Hexane extract of crushed soldier head; LC on silica gel, followed by HPLC

Structure determination MS, 1H and 13C NMR, COSY, ROESY, and NOESY. Configuration at C-17 not determined

Ref. [32, 725]

OR O

RO RO

RO

R ¼ COC2H5 Hospitalitermes (2R*,3R*,4S*,7S*, 8S*,9R*,12S*,14S*, 16S*)-8,19-Epoxy-7hydroxy-17-methyl2,3,9,14tetrapropanoyloxytrinervit-1(15)-en17-yl-(2-propanoyloxy) acetate (800) C38H56O14 HO

n-Hexane extract of crushed soldier heads; LC on silica gel, followed by HPLC

MS, FAB MS,1H and 13C NMR, and comparison with the spectra of compound 798. Configuration at C-17 not determined

[315]

O OR O

OR

O

RO RO

RO

R ¼ COC2H5 (continued)

Chemistry of the Secondary Metabolites of Termites

259

Table 11 (continued) Name (Nr.), formula, structure (3S,4S,7R,8Z,12S,16S)Trinervita-1(15),8-dien3-ol (801) C20H32O

Isolation, notes Ethanol extract of crushed soldiers; HPLC

Grallatotermes, Hospitalitermes, Longipeditermes, Nasutitermes, Subulitermes, Trinervitermes

CH2Cl2 extract of crushed soldier heads and soldiers; LC on silica gel as well as HPLC

Bulbitermes, Grallatotermes, Hospitalitermes, Lacessititermes, Longipeditermes, Nasutitermes, Subulitermes, Trinervitermes

n-Hexane/ether extracts of soldier heads; LC on Florisil CH2Cl2 extract of crashed soldier heads and soldiers; LC on silica gel

IR, Raman, MS, 1 H, and 13 C NMR

[32, 111, 295, 300, 304, 305, 309, 310, 313, 314, 316, 317, 322, 323, 335, 337, 376, 700, 714, 715, 717, 724]

Nasutitermes

CH3OH/CH2Cl2 extract; LC on silica gel, repeated HPLC

HREIMS, 1 H, and 13 C NMR

[32, 111, 295]

HO

(2S,4S,7R,8Z,12S,16S)Trinervita-1(15),8-dien3-ol (802) C20H32O

OH

(2R*,3R*,4S*,7R*, 8Z,12S*,16S*)Trinervita-1(15),8diene-2,3-diol (803) C20H32O2

Structure determination Mp, MS, IR, 1 H and 13 C NMR, NOE, and derivatization (Mp) Absolute configuration was determined by a modified Horeau method (NMR) Revised structure of a trinervitane of T. gratiosus [713] [α]D, IR, MS, and 1H NMR; 802 assigned as minor component of the frontal gland secretion of T. gratiosus [337] later revised to compound 801 [713]

Termite genera Bulbitermes, Grallatotermes, Hospitalitermes, Lacessititermes, Nasutitermes, Subulitermes, Trinervitermes

HO

Ref. [32, 111, 265, 309, 310, 315, 713, 714]

[32, 300, 304, 305, 312, 313, 317, 322, 323, 335, 337, 376]

OH

(2S*,3S*,4S*,7R*, 8Z,12S*,16S*)Trinervita-1(15),8diene-2,3-diol (804) C20H32O2

HO OH

(continued)

260

E. Gössinger

Table 11 (continued) Name (Nr.), formula, structure (2S*,3R*,4S*,7R*, 8Z,12S*,16S*)Trinervita-1(15),8diene-2,3-diol (805) C20H32O2

Structure determination MS, 1H and 13 C NMR, COSY, and ROESY

Termite genera Hospitalitermes

Isolation, notes n-Hexane extract of crushed soldier heads; LC on silica gel, followed by HPLC

Ref. [315]

Hospitalitermes, Nasutitermes, Trinervitermes

CH2Cl2 extract of crushed soldiers; LC on silica gel, preparative TLC, caution: probably an artifact, formed by work up

MS, IR, and 1 H NMR

[32, 311]

Nasutitermes

CH2Cl2 extract of crushed termite soldiers; HPLC

MS, IR, and 1 H NMR

[32, 111, 311, 312]

HO OH

(3R*,4S*,7R*,8Z, 12S*,16S*)-3Hydroxytrinervita1(15),8-dien-2-one (806) C20H30O2

HO O

(3S*,4S*,7R*,8Z, 12S*,16S*)-3Hydroxytrinervita1(15),8-dien-2-one (807) C20H30O2

HO O

(continued)

Chemistry of the Secondary Metabolites of Termites

261

Table 11 (continued) Name (Nr.), formula, structure (3S*,4S*,7S*,8R*, 11E,15S*,16S*)Trinervita-1,11-dien-3ol (808) C20H32O

Termite genera Trinervitermes

Isolation, notes CH2Cl2 extract of soldiers; several LC on silica gel

Nasutitermes

CH2Cl2 extract of crushed soldiers; GC/MS; HPLC Active against Gram-positive bacteria, e.g., methicillinresistant Staphylococcus aureus (MIC  63 μg/ cm3] and against nonfilamentous fungi, e.g., Candida albicans (MIC  63 μg/ cm3) n-Hexane extract of crushed soldier heads; HPLC

HO

(2R,3R,4S,7R,8R, 11E,16S)-Trinervita1(15),11-diene-2,3-diol (809) C20H32O2

HO OH

(1R*,3S*,4S*,7S*, 8R*,11E,13R*,16S*)Trinervita-11(12), 15(17)-diene-3,13-diyl diacetate (810) C24H36O4

AcO

Bulbitermes, Nasutitermes

Structure determination Mp, HRMS, [α]D, IR, and 1 H NMR; oxidation to the unsaturated ketone by MnO2. Acetylation followed by epoxidation led to a crystallizable epoxide enabling X-ray analysis HRMS, [α]D, 1H and 13C NMR, 2D NMR including NOESY, and several computational calculations to confirm the structure and its conformation and configuration

MS, IR, 1H, and 13 C NMR

Ref. [32, 111, 717, 728]

[729]

[32, 310, 323]

OAc

(continued)

262

E. Gössinger

Table 11 (continued) Name (Nr.), formula, structure Termite genera (1R*,3S*,4S*,7R*, Cortaritermes 8S*,9S*,11E,13R*, 16S*)-Trinervita11,15(17)-diene-3,9,13triyl triacetate (811) C26H38O6

Isolation, notes CH2Cl2 extract of crushed soldiers, repeated HPLC

Structure determination HRMS, IR, 1 H and 13 C-NMR, and hydrolysis to the triol (Mp)

Ref. [32, 111, 295]

OAc

OAc

AcO

(1R*,3S*,4S*,7R*, 8S*,9S*,11E,13R*, 16S*)-9-Acetoxy-13hydroxytrinervita11,15(17)-dien-3-yl propionate (812) C25H38O5

Nasutitermes

CH2Cl2 extract of soldiers and “milking” of the frontal gland secretion; repeated LC on silica gel

MS, [α]D, IR, and 1H NMR

[32, 111, 708]

CH2Cl2 extract of crushed soldiers; repeated HPLC

HRMS, IR, 1 H and 13 C-NMR, and hydrolysis to the triol (Mp)

[32, 111, 295]

OAc

OH

EtCO2

(1R*,3S*,4S*,7R*,8S*, Cortaritermes 9S*,11E,13R*,16S*)Diacetoxytrinervita11,15(17)-dienyl propionate (813) C27H40O6 OR

RO

OR

R ¼ 1 COC2H5, 2 Ac (continued)

Chemistry of the Secondary Metabolites of Termites

263

Table 11 (continued) Name (Nr.), formula, structure (1R*,3S*,4S*,7R*, 8S*,9S*,11E,13R*, 16S*)-9Acetoxytrinervita11,15(17)-diene-3,13diyl dipropionate (814) C28H42O6

Structure determination HRMS, [α]D, IR, 1 H, 13C, and LIS 1 H NMR

Termite genera Cortaritermes, Nasutitermes

Isolation, notes n-Hexane extract of crushed soldier heads; LC on Florisil

Ref. [32, 111, 295, 313, 323, 708, 721]

Nasutitermes

CH2Cl2 extract of soldiers and “milking” of the frontal gland secretion; LC on silica gel several times

MS, [α]D, IR, and 1H NMR

[32, 111, 708]

n-Hexane extract of crushed soldier heads; LC on Florisil, HPLC

HRMS, [α]D, IR, 1 H and 13 C-NMR, and hydrolysis which led to the triol, a crystalline product, allowing structure determination by X-ray analysis [721]

[32, 111, 313, 323, 708, 721]

OAc

EtCO2

O2CEt

(1R*,3S*,4S*,7R*, 8S*,9S*,11E,13R*, 16S*)-13Hydroxytrinervita11,15(17)-diene-3,9diyl dipropionate (815) C27H40O5 O2CEt

OH

EtCO2

(1R*,3S*,4S*,7R*, Nasutitermes, 8S*,9S*,11E,13R*, Leucopitermes 16S*)-Trinervita11,15(17)-diene-3,9,13triyl tripropionate (816) C29H44O6 O2CEt

EtCO2

O2CEt

(continued)

264

E. Gössinger

Table 11 (continued) Name (Nr.), formula, structure Termite genera (1S*,3S*,4S*,7R*, Nasutitermes 8S*,9S*,11S*,12S*, 13R*,16S*)-9-Acetoxy13-hydroxy-11,12epoxytrinervit-15(17)en-3-yl propionate (817) C25H38O6

Isolation, notes CH2Cl2 extract of soldiers and “milking” of the frontal gland secretion; repeated LC on silica gel

Structure determination HRMS, [α]D, IR, 1 H, and 13 C-NMR

Ref. [32, 111, 708]

OAc

O

OH

EtCO2

(1S*,3S*,4S*,7R*, Nasutitermes 8S*,9S*,11S*,12S*, 13R*,16S*)-9-Acetoxy11,12-epoxytrinervit15(17)-ene-3,13-diyl dipropionate (818) C27H42O7

CH2Cl2 extract of soldiers and “milking” of the frontal gland secretion; several LC

HRMS, [α]D, IR, 1 H, and 13 C NMR

[32, 111, 708]

CH2Cl2 extract of soldiers and “milking” of the frontal gland secretion; LC on silica gel several times

HRMS, [α]D, IR, 1 H, 13C NMR

[32, 111, 708]

OAc O

O2CEt

EtCO2

(1S*,3S*,4S*,7R*, 16S*,9S*,11S*,12S*, 13R*,16S*)-11,12Epoxytrinervit-15(17)ene-3,9,13-triyl tripropionate (819) C28H44O7

Nasutitermes

O2CEt

O

EtCO2

O2CEt

no less than 15 trinervitanes from its frontal gland [714]. Analytical separations were carried out by GC, mostly on capillary columns. Several researchers determined standardized retention times (Kovats indices) to accelerate the search for the composition of the frontal gland secretion of further Nasutitermitinae species by

Chemistry of the Secondary Metabolites of Termites

265

Table 12 NMR spectrometric data of the trinervitane 763, Ref. [715] 13

Structure, Nr. 19 8 OAc

H 7

9

11

17 1

6 16 15

3

OH 18

763

13

12 14

OH

5 4

20

10

2

Position 1 2 3 4 5α 5β 6 6 7 8 9 10 11 12 13 14 15 16 17 18 19 19 20 CH3CO CH3CO

H NMR (CDCl3, δ/ppm, J/Hz) – 3.86 dq; J2,3 ¼ 9, J2,17 ¼ 2 4.02 d – 2.13 d; br 2.35 ddd; J5,5 ¼ 13, J5,6 ¼ 12, J5,6 ¼ 6 2.13 d; br 2.60 m; br 3.09 ddd; J7,6 ¼ 12, J7,6 ¼ 10, J7,16 ¼ 13 – 5.56 t; br J9,10 ¼ 6 nd nd nd nd nd – nd nd 0.99 s 5.16 d; br J19,19 ¼ 1.5 Hz 5.22 d; J19,19 ¼ 1.5 Hz 0.93 d; J20,12 ¼ 7 Hz – nd 1

C NMR (δ/ppm) 134.6 s 73.5 d 72.3 d 46.7 s 30.3 t 24.8 t 51.7 d 151.3 s 69.6 d 32.2 t 32.2 t 27.7 d 32.6 t 36.4 t 127.9 s 57.6 d 22.8 q 21.6 q 117.5 t 20.3 q 171.0 s 21.4 q

nd not detected

dereplication. If the fragmentation pattern in the mass spectra pointed to a cyclic diterpene, the further structure determination was based on comparison of the new diterpene with spectra of known trinervitanes. As the NMR spectra of 763 show (Table 12), the assignments were difficult, and some early structure determinations had to be revised after the improvement of NMR techniques [713]. Lanthanideinduced shifts (LIS), nuclear Overhauser effects (NOE), and several 2D NMR techniques were used. Four of the known trinervitanes were characterized by X-ray structure analysis, whereas in other cases the spectrometric data had to be supplemented by chemical reactions. Acylation, hydrolysis, oxidation, and reduction were used (see Table 11). One of the disadvantages of the early structure determinations was the fast oxidation in air of trinervitenes containing the allylic 2βhydroxy group to the conjugated enone due to the nearly perfect overlapping of the front orbitals of the Δ1,15-double bond and the hydrogen bond at C-2. This

266

E. Gössinger

Table 13 NMR spectrometric data of the trinervitane 725, Ref. [336] Structure, Nr. 19 11

8

H 7

9 16 15

14

1 5 3

4 18

OH

757

OH 2

13

12

17

6

20

10

Position 1 2 3 4 5a 5b 6 6 7 8 9a 9b 10a 10b 11a 11b 12 13a 13b 14a 14b 15 16 17 18 19a 19b 20

H NMR (CDCl3, δ/ppm, J/Hz) – 3.94 d J2,3 ¼ 8.9 Hz 3.80 d J3,2 ¼ 8.9 Hz – 2.04 m 1.17 m 1.78 m 1.65 m 3.14 ddd J7,6 ¼ 8.3, J7,6 ¼ 11.2, J7,16 ¼ 12.0 – 1.80 m 2.11 m 1.49 m 1.70 m 1.15 m 1.15 m 1.66 m 1.09 m 1.67 m 2.20 m 2.26 m – 2.36 d J16,7 ¼ 12 Hz 1.73 d J ¼ 1.8 Hz 0.98 s 4.92 dd J19,19 ¼ 2 Hz J ¼ 2Hz 4.76 dd J19,19 ¼ 2 Hz J ¼ 2Hz 0.92 d J20,12 ¼ 7 Hz 1

13

C NMR (δ/ppm) 132.3 73.5 d 72.9 d 46.5 s 36.5 t 36.5 t 29.7 t 29.7 t 53.0 d 153.1 s 27.2 t 27.2 t 25.2 t 25.2 t 34.6 t 34.6 t 27.5 d 32.4 t 32.4 t 24.4 t 24.4 t 130.0 s 58.1 d 21.9 q 20.4 q 112.0 t 22.8 q

disadvantage was turned into an advantage by using it to determine the relative configuration of the C-2 hydroxy group of trinervit-1(15)-enes. The trinervitane most often detected in the subfamily Nasutitermitinae is 2β,3αdihydroxytrinervita-1(15),8(19)-diene (725). Its NMR spectrometric data are presented as a typical example of the improvement of the methods and their interpretation when compared with the first NMR spectrum of a trinervitane (see Tables 12 and 13). Despite their abundance in higher termites and their early characterization, only one total synthesis of a trinervitane has been reported. This synthesis by Kato et al. and even more the unsuccessful attempts demonstrate the severe difficulties the cyclization of the strained 11-membered ring posed and how difficult the prediction of the stereochemistry of this complex strained ring system is.

Chemistry of the Secondary Metabolites of Termites

267

Synthesis of ()-Trinervita-1(15),8(19)-diene-2β,3α-diol and ()-Trinervita1(15),8(19)-diene-2α,3α-diol by Kato et al. Kato et al. developed the biomimetic synthesis of racemic trinervitanes. They recorded the progress of their synthesis efforts in several publications [718, 731– 734]. The synthesis started with the racemic secotrinervitane derivative 751, which Kato et al. had synthesized in a biomimetic manner starting with geranylgeranoic acid via neocembrene [701, 703]. The first aim was the selective epoxidation of the exocyclic double bond (Scheme 123). The investigators chose the Sharpless protocol [239, 240] to direct the epoxidation to the exocyclic double bond. To achieve this goal, the secondary alcohol had to be protected as a methoxymethyl ether; afterwards the acetate was saponified. The liberated alcohol 820 was inverted in high yield by oxidation and hydride reduction. The axial homoallylic alcohol 821 directed the epoxidation with titanium tetraisopropoxide and t-butyl hydroperoxide toward the desired double bond. After protection of the axial alcohol as methoxymethyl ether 822, the epoxide was isomerized to the primary allylic alcohol 823 with aluminum triisopropoxide. To use the primary alcohol or its derivative for Lewis acid-catalyzed olefinic cyclization, the protecting groups of the secondary alcohols had to be exchanged for Lewis acid-stable protective groups. Hydrolysis of the methoxymethyl ethers with aqueous hydrochloric acid, temporary protection of the primary alcohol as tbutyldimethylsilyl ether 824, protection of the vicinal alcohols as cyclic carbonate, and removal of the silyl group with fluoride set the stage for the acid-catalyzed olefin cyclization. However, the allylic alcohol 825 as well as its oxidation product, the unsaturated aldehyde, proved too unstable for the necessary high acidity. Thus, the hydroxy group was exchanged for chloride using mesyl chloride and lithium chloride under basic conditions. To facilitate the cyclization, strain was reduced by saponification of the cyclic carbonate. The chloride of the diol 826 was abstracted by silver perchlorate, and the allylic cation generated attacked the Δ7,8-double bond. At 20 C the main product was diol 829 accompanied by very small amounts of the isomer with the desired exocyclic Δ8,19-double bond but with the hydrogen at C-7 in the unwanted β-position. The reaction proved temperature and solvent dependent. When the reaction was performed at room temperature, a second cyclization followed leading to the kempane ring system 830. This second cyclization is due to the acid generated, as the following experiment demonstrated. When the trinervitatrienediol 829 was treated with t-butyl chloride and silver perchlorate, the generated perchloric acid attacked the exocyclic double bond, and the newly formed tertiary carbocation attacked the Δ11,12-double bond. To improve the yield of the cyclization, the vicinal diol of 826 was protected, and the solvent was changed. The best reaction conditions found were protection of the vicinal diol as diacetate 827 and conducting the reaction at room temperature with silver perchlorate in ether containing one equivalent of pyridine to scavenge the generated perchloric acid. Saponification of the generated tricyclic diacetate 828 furnished the trinervitatrienediol 829 in very good yield. The next task in the synthesis toward

OAc

1) HCl, CH3OH, 90%

2) KOH, CH3OH, 99%

ether, rt, 93%

AgClO4, 1 eq pyr,

OH

OAc OAc 828

824

OH

OTBS

820

N O

N

N

PhH, 92%

N

2) NaBH4, CH3OH, –15°C, 92%

saponification

2) TBAF, THF, 99%

1)

OH OMOM

1) PCC, NaOAc, 4 Å MS, CH2Cl2, 90%

Scheme 123 Synthesis of secotrinervitane 829 and trinervitane 830

827

OAc OAc

Cl

2) TBSCl, imidazole, OMOM DMF, 90% OMOM 823

HO

751

OH

1) MOMCl, i-Pr2NEt, CH2Cl2, 94%

OH

O

OH 829

825 O

O

OH

AgClO4, THF, –20°C, 68%

2) KOH, H2O, glyme, 98%

1) MsCl, Et3N, LiCl, CH2Cl2, 94%

2) MOMCl, i-Pr2NEt, CH2Cl2, 94%

AgClO4, t-BuCl, –20°C, 86%

OMOM OH 821

1) t-BuO2H, Ti(i-PrO)4, CH2Cl2, 0°C, 54%

OH

OH

OH

830

66%

OH

OH

100%

Ac2O, DMAP, pyr, rt,

AgClO4, THF, 20°C, 50%

826

OH

Cl

OMOM OMOM 822

O

Al(Oi-Pr)3, toluene, rfl,

268 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

269

naturally occurring trinervitanes was the manipulation of the double bonds (Scheme 124). With Adams catalyst the exposed Δ11,12-double bond of 829 was hydrogenated regioselectively. Obviously, at least two main conformations exist, because the face selectivity was not high (α:β ¼ 7:3) and separation of the diastereomers proved impossible. To shift the balance of the conformations of trinervitane 829 and hopefully allow separation after the hydrogenation, the researchers increased the strain by forming the cyclic carbonate 831 using 1,10 -carbonyldiimidazole in benzene. Indeed, the ratio of the two diastereomeric dihydrotrinervitadienes 832 and 833 shifted to 1:6 in favor of the desired diastereomer 833. Saponification of the carbonates was the first step in the effort to shift the exocyclic double bond into the cyclohexane. The equatorial hydroxy group of the vicinal diol 833 was protected as a methoxymethyl ether. The starting material 833, the other monomethoxymethyl ether, and the dimethoxymethyl ether were recycled to increase the yield to 74%. Then, the axial hydroxy group was oxidized. The aspired shift of the deconjugated enone to the conjugated enone 834 proved difficult. Finally, by refluxing the deconjugated enone in toluene with DBU as base, the conjugated enone 834 was obtained in very good yield within 7 days. Conjugation and the increased sterical hindrance deactivated the shifted double bond enough to tackle the Δ7,8-double bond of 834. Hydroboration and subsequent oxidation led exclusively to the hydroxy group attached to the cyclopentane. Therefore, the authors aimed to introduce the hydroxy group at the C-8 position by epoxidation and consecutive isomerization of the epoxide to the allylic alcohol. They were successful in epoxidizing the isolated double bond from the convex α-face, but difficulties with the consecutive basic isomerization to the allylic alcohol led to the following reaction sequence. Exchange of the acid-sensitive methoxymethyl group by acetate was followed by epoxidation with m-chloroperbenzoic acid, which led quantitatively to the α-epoxide 835. After several trials with diverse Lewis acids, it was found that trimethylsilyl chloride was mild enough not to destroy the unstable functionalities at the six-membered ring but still strong enough to isomerize the epoxide to a mixture of the two allylic alcohols 836 and 837 in a 2:3 ratio in 69% yield. The allylic alcohol 837 was hydrogenated in high yield using Adams catalyst and hydrogen at normal pressure. The tertiary alcohol 838 was dehydrated with thionyl chloride furnishing a mixture of three double-bond isomers 839, 840, and 841. This mixture was separated on silver nitrate-impregnated silica gel supplying the desired trinervitane with an exocyclic double bond (841) in 34% yield. Reduction of the acetate as well as the keto group with lithium aluminum hydride led to the two naturally occurring trinervitadienediols 725 and 755 in 56% and 44% yield, respectively, and thus completing the first synthesis of 725, the most abundant trinervitane found in many genera and species of the subfamily Nasutitermitinae, and 755, which occurs in the secretion of the frontal glands of several Nasutitermes species and in Velocitermes velox. The completion of the synthesis starting with neocembrenone, which itself had to be synthesized, required 31 steps and yielded less than 1% of 725 and 755. It should be noted that Kato’s protocol could be used to synthesize other naturally occurring trinervitanes with small variations from 725 and 755. For

838

OAc

834

O

O

N

N

SOCl2, N2, CH2Cl2, rt

pyr, CH2Cl2, rt, 100% 3) MCPBA, CH2Cl2, 0°C, 100%

1) HCl, CH3OH, rt, 85% 2) AcCl, DMAP,

831 O

O

O

O

OAc

OH

H

725

52% 839

OH

O

O

4:5

+

+

69%

755

OH

H

11% 840

OAc

OH

O

HO

OH

832

OH

TMSCl, N2, –10°C, CH2Cl2,

2) KOH, CH3OH, rt, 99%

1) H2, 1 atm, PtO2, CH2Cl2, rt, 99%

835

OAc

Scheme 124 Synthesis of the trinervitanes 725 and 755

HO

OH

OMOM O

829

OH

N

benzene, rt, 92%

N

836

OAc

+

O

1:6

+

34% 841

OAc

H

2 : 3

+

O

O

2) LAH, THF, –20°C, 88%

rt, 96%

H2 (1 atm), PtO2, CH3OH,

CH2Cl2, 91% 3) DBU, toluene, rfl, 7 d, 89%

1) separation SiO2/AgNO3

837

OAc

OH

HO

833

OH

1) MOMCl, EtNi-Pr2, 74% 2) PCC, NaOAc,

270 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

271

example, saponification of the acetate 840 would yield the naturally occurring enone 806 and acetate 841 naturally occurring enone 764. Due to the difficulties mentioned above, three further attempts to synthesize naturally occurring trinervitanes failed.

Synthesis of (1S,2R,12S)-2-Hydroxy-11,12-dihydroneocembrene by Kato et al. To transform their synthesis of racemic trinervitane into an EPC synthesis, Kato et al. started with the commercially available chiral monoterpene R-citronellol (843) (Scheme 125) [735]. The alcohol was exchanged by bromide via the tosylate. The bromide was converted into the Grignard reagent 844, which was expected to couple with a suitably protected 8-hydroxygeraniol. To obtain this compound, geranyl acetate (845) was oxidized regioselectively to 8-hydroxygeranyl acetate (626) according to Sharpless with t-butyl hydroperoxide, catalytic amounts of selenium dioxide, and acid [232]. Several trials were necessary to decide the terminal functionalities of this geraniol derivative enabling the desired coupling reaction with Grignard reagent 844. As coupling reagent, in situ-prepared dialkylcuprate was chosen due to its preference for the SN2 versus the SN20 reaction [736]. Experiments exchanging the terminal hydroxy group of the partially protected 8-hydroxygeraniol by chloride and coupling the chloride with the Grignard reagent 844 led to a 1:1 mixture of SN2 and SN20 reaction products. To improve this result, esters instead of chloride were tested as terminal group, and the bulky pivalate led to the best results. As protecting group of the other terminal hydroxy of 8-hydroxygeraniol, the tetrahydropyranyl ether was more advantageous than the silyl ether. Thus, 8-hydroxygeranyl acetate (626) was esterified to the pivalate followed by selective saponification of the acetate. The generated primary allylic alcohol was protected as a tetrahydropyranyl ether. To this allylic ester 846 in tetrahydrofuran and dilithium tetrachlorocuprate was added very slowly the Grignard compound 844 at low temperatures. Under these conditions, the coupling product, 11,12-dihydrogeranylgeranyl tetrahydropyranyl ether (847), was obtained in over 90% yield. The next steps, hydrolysis of the tetrahydropyranyl ether, oxidation of the primary alcohol to the corresponding acid via the aldehyde and Pinnick oxidation, and subsequent acid chloride formation with thionyl chloride, set the stage to the intramolecular olefinic Friedel-Crafts reaction of 848. Due to the exchange of two sp2-centers for sp3-centers, the yield of this cyclization to the dihydroneocembrene derivative 849 was lower than that of the analogous cyclization in the original neocembrene synthesis of Kato et al. [123, 124]. Also, the higher torsional strain and thus changed conformations of the cyclization product 849 necessitated changes from the planned pathway. It proved not possible to adopt the conditions of the hydrogen chloride elimination from the neocembrene synthesis. Only when the following reaction conditions, silica gel and potassium carbonate in n-hexane at room temperature

Cl

78%

K2CO3, SiO2, n-hexane, rt,

846

844

Scheme 125 EPC synthesis of 2-hydroxy-11,12-dihydroneocembrene (842)

849

848

PivO

O

–78°C, 56% (2 steps)

SnCl4, CH2Cl2,

DMAP, CH2Cl2 , rt 3) K2CO3, CH3OH, rt 4) DHP, TsOH, CH2Cl2, rt, 71% (3 steps)

OAc

2 , CH2Cl2, OH rt, 49% 2) PivCl, pyr,

CO H

1) t-BuOOH, SeO2,

3) Mg, THF, N2, rt

O

Cl

845 (geranyl acetate)

843 ((R)-citronellol)

OH

1) TsCl, pyr, 0°C 2) LiBr, acetone, 56°C, 83% (2 steps)

O 850

92%

Li2CuCl4, THF, –78°C to rt,

OTHP

MgBr

–78°C, 78%

BuLi, DIBAH, N2, n-hexane, toluene,

847

OTHP

842

OH

3) aq NaClO2, NaH2PO4, CH3CN, DMSO, 0°C to rt, 98% 4) SOCl2, pyr, PhH, 0°C

1) TsOH, CH3OH, rt, 98% 2) Dess-Martin ox., CH2Cl2, pyr, rt, 90%

272 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

273

for 2 days, were applied was it possible to reduce the percentage of the tetrasubstituted conjugated Δ1,15-double-bond formation and to obtain the desired elimination to the isopropenyl group of dihydroneocembrene 850 in 78% yield. The synthesis efforts were concluded by the reduction of the keto group. Again, the reaction conditions had to be varied because diisobutylaluminum hydride led to a mixture of cis- and trans-alcohol. Abolishing the Lewis acidic character of the reagent by addition of equivalent amounts of butyllithium successfully changed the ratio furnishing the cis-alcohol 842 in 78% yield. No further development of this synthesis has been reported.

Synthesis of 12-Benzyloxy-12-desmethyl-1(15),8-trinervitadiene and 12-Benzyloxy12-desmethyl-1(15),8(19)-trinervitadiene by Dauben et al. After having succeeded in synthesizing kempane-2 [737], Dauben et al. planned the synthesis of a trinervitane [738]. Their emphasis in this synthesis attempt was to introduce the double bonds in the correct position to circumvent Kato’s difficulties in shifting the double bonds to the correct positions in their synthesis of racemic trinervitane. The key steps consisted of a Robinson annulation to construct the cishydrindene and a McMurry coupling to close the 11-membered ring (Scheme 126). Commercially available cyclotene 853 was converted into the enol acetate by acetic acid anhydride under reflux. Consecutive reduction with hydrogen and palladium on charcoal led to the monoketone 854. Protection of the keto group as a cyclic ketal was followed by transesterification. The liberated alcohol was oxidized by Swern’s reagent. Robinson annulation of ketone 855 with ethyl vinyl ketone led in 78% yield to hydrindenone 856. After hydrogenation of the double bond of the conjugated enone, the saturated ketone was transformed into the vinyl triflate 857 with phenylditrifluoromethanesulfonimide and magnesium diisopropylamide as base. With this step the first double bond was correctly positioned and was suitably substituted for a Stille coupling [739]. Allyltributylstannane in the presence of an excess of lithium bromide and tetrakis(triphenylphosphine)palladium in tetrahydrofuran at reflux led in nearly quantitative yield to the allyl-substituted hydrindene 858. Selective hydroboration by 9-borabicyclononane and consecutive oxidation led to the primary alcohol, which in turn was oxidized to the aldehyde 859 to enable the chain extension with lithium propyltetrahydropyranyl ether. The newly generated epimeric secondary alcohols were protected as benzyl ethers 860 prior to the acidic hydrolysis of the cyclic ketal, which also partly hydrolyzed the tetrahydropyranyl ether. The keto group was transformed into the methylene group with dibromomethane, zinc, and titanium tetrachloride [740]. To prepare for the difficult cyclization of the 11-membered ring, the protective group of the primary alcohol of

O

864

861

O

OBn

O

OBn

858

O

OAc

2) (COCl)2, DMSO, Et3N, CH2Cl2

1) 9-BBN,THF, 0°C to rt; H2O2, NaOH

11%

TiCl3(DME)1.5, Zn/Cu, DME, rfl,

DMAP, CH2Cl2, 86% (4 steps)

O

859

851

862

O

855

O

O

TBSO

3) (COCl)2, DMSO, Et3N, CH2Cl2 , 73% (5 steps)

1) (CH2OH)2, TsOH, PhH, rfl 2) NaOCH3, CH3OH

1) TsOH, CH3OH 2) TBSCl, Et3N,

854

O

Scheme 126 Synthesis of desmethyltrinervitane 852

O

Pd(PPh3)4, THF, rfl, 95%

THPO

1) Ac2O, rfl

2) H2, Pd/C, AcOEt

SnBu3

LiBr exc,

853

O

OBn

OBn

O

+

O

HO

O

856

852

2) (COCl)2, DMSO, Et3N, THF; CH3Li, 48% (3 steps)

1) 9-BBN, THF, 0°C to rt; H2O2, NaOH

2) BnBr, NaH, Bu4NI, THF, 60% (4 steps)

,

O

Li

ether, –60°C

1) THPO

CDCl3, H

O

0°C to rfl, 78%

CH3OH,

O CH3ONa,

863

TBSO

860

THPO

OBn

OBn

2) BrMgN(i-Pr)2, ether, HMPA; PhNTf2, 81% 857

OTf

1) (CO2H)2/SiO2, CH2Cl2

O

CH2Cl2, 56% (2 steps)

1) TBAF, THF 2) PCC/Al2O3,

2) CH2Br2/Zn, TiCl 4, THF, –30°C to rt

1) H2, Pd/C, AcOEt, K2CO3, 69% O

OBn

O

O

274 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

275

861 was exchanged for the silyl group prior to the regio- and stereoselective hydroboration of the methylene group of the hydrindene derivative 862 by 9-borabicyclononane. Consecutive oxidation to the primary alcohol and further oxidation by Swern’s reagent to the aldehyde allowed chain extension by methyllithium to the 1-substituted ethanol 863, with the correct relative configuration at the ring junction. Then, the silyl ether was cleaved and both alcohols were oxidized simultaneously. The keto aldehyde 864 was subjected to McMurry conditions affording after HPLC separation in 11% yield the 11-membered ring system 851 with the newly formed (Z )-configured double bond. This inseparable mixture of epimeric benzyl ethers proved unstable under slightly acidic conditions yielding slowly the less strained trinervitane with the exocyclic double bond 852.

Synthesis of a Substituted Hydrindene as Intermediate of a Planned Trinervitane Synthesis by Yadav et al. The starting material of this synthesis approach [741], geraniol, was converted into the chiral epoxide 865 according to Marshall et al. (Scheme 127) [742]. To transform the epoxy chloride 866 into the secondary propargyl alcohol, the investigators used a protocol that they had developed earlier [743]. When 866 was added to freshly prepared lithium amide in ammonia containing catalytic amounts of ferric nitrate, the chiral propargyl alcohol was obtained. Subsequently, the newly formed alcohol was protected as silyl ether 867 to permit deprotonation of the alkyne moiety by butyllithium followed by addition of formaldehyde. The primary alcohol was protected as 2-methoxyethoxymethyl ether 868 prior to the selective deprotection of the terminal silyl ether with camphorsulfonic acid in a methanol/dichloromethane solution. This set the stage for a Julia-Kocienski olefination [744–747]. The deprotected primary alcohol was converted into the tetrazole thioether 869 by Mitsunobu reaction with 1-phenyltetrazole-5-thiol. After oxidation to the sulfone 870 with ammonium molybdate and hydrogen peroxide, the sulfone was deprotonated by strong base, and the chiral aldehyde 871 was added yielding stereoselectively the conjugated (E,E)-diene. Acidic hydrolysis with methanolic hydrochloric acid set the two propargylic alcohols free enabling the facile reduction of the alkyne 872. The synthesis of the chiral aldehyde 871 (Scheme 128) started with 1,6-hexanediol (876), which was converted into the oxazolidinone derivative 877 [748].

867

N

N

Ph N

S

873

N

O

O

OTES

OBn

870

OH

OMEM

865

THF, 82%

IBX, DMSO, NaHCO3 ,

OTPS

2) MEMCl, i-Pr2NEt, CH2Cl2, 0°C to rt, 90%

1) BuLi, –78°C to rt; CH2O, 0°C to rt, 89% TPSO

TPSO

Scheme 127 EPC synthesis of the substituted hydrindene 875

CH2Cl2, rt 3) TBAF, 0°C, 5´, 68% (2 steps)

1) Red-Al®, ether, 89% 2) TESCl, NEt3,

0°C to rt, 70%

(NH4)2MoO4 H2O2, EtOH,

TPSO

OTPS

OH

OTPS

OH

OBn

+

868

O

O

874

O

871

OMEM

TPSO

1) KHMDS, THF, –78°C

SH, THF, 80%

N N

Ph N

866

N

2) NaBH4, CH3OH, 70% (2 steps)

1) BHT cat., toluene, 180°C

2) HCl, CH3OH, 55% (2 steps)

OTPS

OBn

Ph

N N N N

2) DEAD, Ph3P,

1) CSA, CH3OH, CH2Cl2, rt, 90%

CCl4, rfl, 90%

Ph3P, NaHCO3,

S

OBn

O Cl

872

OBn

875

HO

OH

869

OTPS

OH

OTES

OMEM

2) TPSCl, imidazole, CH2Cl2, 90% (2 steps)

1) LiNH2, NH3, Fe(NO3)3 cat., –78°C

276 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

277

O

O

[749]

[748]

HO

OBn

N

OH Bn

OBn

O

O

876

877

878

OBn

[750] EtO

CH3OH, 0°C O

OBn

NaBH4, NiCl2•6H2O,

879

1) LAH, THF, 0°C, 80% (2 steps) OBn 2) IBX, DMSO, THF, 91%

EtO O

880

O

871

Scheme 128 Asymmetric synthesis of the precurser 871 of hydrindene 875

Evans alkylation and conversion of the generated oxazolidinone derivative into the aldehyde 878 followed the protocol of Shioiri et al. [749]. Chain elongation of 878 by Horner-Emmons reaction was described by Dutta [750]. The resulting unsaturated ester 879 was hydrogenated by sodium borohydride and nickel chloride [751]. The ester group of 880 was converted into the aldehyde 871 via the alcohol by lithium aluminum hydride and reoxidation. Returning to the propargylic diol 872, Redal® was used to reduce the triple bond to the (E)-configured double bond. The secondary alcohol was protected as a silyl ether, which was accomplished by silylation of both the primary and the secondary alcohol and deprotection of the primary alcohol with tetrabutylammmonium fluoride at 0 C and a very short reaction time. The primary alcohol 873 was oxidized to the aldehyde 874 by 2-iodoxybenzoic acid (IBX). The following intramolecular DielsAlder reaction was conducted at surprisingly high temperature, and the product obtained 881 (see Scheme 129) was treated with sodium borohydride to reduce the aldehyde moiety. However, the reduced Diels-Alder adduct 875 the researchers depicted is not compatible with the Woodward-Hoffman rules.

278

E. Gössinger

RO

R

R exo add.

O

H

according to Yadav:

H

OTES

OTES

OTES

O

881

874

R O

R endo add.

OTES

O

R=

OBn

H OTES

874

Scheme 129 Stereochemical studies of the IMDA reaction

Although isomerization of the double bonds at high temperature is possible, a single stereoisomer would be very unusual. A two-step mechanism cannot be ruled out. The researchers did not explain this discrepancy. They based their structure determination on NMR spectrometric data especially nuclear NOEs and quantum mechanical energy minimization studies.

The Tetracyclic Diterpenes Table 14 contains the tetracyclic diterpenes encountered in termites.

Chemistry of the Secondary Metabolites of Termites

279

Table 14 The tetracyclic diterpenes Name (Nr.), formula, structure (1R,3S,4S,8Z,11R,12S, 14S,15R,16R)Kempa-6,8-diene3,14-diyl diacetate (Kempene-1) (882) C24H34O4

Isolation, notes

Bulbitermes, Havilanditermes, Nasutitermes

n-Hexane extract of crushed soldier heads; LC on Florisil, followed by HPLC Relative configuration at C-3 of the compound isolated from the frontal gland of N. kempae and N. infuscatus was not determined [300, 752]

MS, IR, UV, CD, 1 H, 13C, and LIS NMR, and EPC synthesis [753]

[32, 111, 305, 306, 310, 323, 341, 752]

Bulbitermes, Nasutitermes, Havilanditermes

n-Hexane extract of crushed soldier heads; HPLC

MS, 1H and 13C NMR, and EPC synthesis [753]

[32, 111, 305, 306, 310, 323, 341]

Nasutitermes

Mentioned in a review as naturally occurring kempane [353]; no further data were given

Bulbitermes, Nasutitermes

n-Hexane extract of crushed soldier heads; LC on Florisil, followed by HPLC Easily oxidized in air

AcO OAc

(1R*,3R*,4S*,8Z,11R*,12S*,14S*,15R*,16R*)-Kempa6,8-diene-3,14-diyl diacetate (Epikempene-1) (883) C24H34O4

Structure determination

Termite genera

Ref.

AcO OAc

(1R*,4S*,8Z,11R*,12S*, 14S*,15R*,16R*)14-Hydroxykempa6,8-dien-3-one (884) C20H28O2

[111, 353]

O OH

(1R,4S,8Z,11R,12S,14S,15R,16R)-3Oxokempa-6,8-dien14-yl acetate (Kempene-2) (726) C22H30O3

MS, IR, UV, 1H, 13 C, and LIS NMR, X-ray analysis, CD, racemate synthesis, [737] and EPC synthesis [753]

[32, 111, 306, 310, 323, 341, 752]

O OAc

(continued)

280

E. Gössinger

Table 14 (continued) Name (Nr.), formula, structure (1R,3R,4S,7S,8Z,11R,12S, 15R,16S)-3Hydroxykemp-8-en6-one (885) C20H30O2

Structure determination

Termite genera

Isolation, notes

Nasutitermes

n-Hexane/CH2Cl2 extract of crushed soldier heads; LC on Florisil, followed by HPLC

MS, IR, 1H and 13 C NMR, CD, and X-ray analysis of its 4-bromobenzoate derivative

[32, 111, 306, 376, 754, 755]

Ref.

Nasutitermes

n-Hexane/CH2Cl2 extract of crushed soldier heads; LC on Florisil, followed by HPLC

MS, IR, 1H-, 13 C NMR

[32, 111, 306, 376, 754, 755]

Grallatotermes, Nasutitermes

n-Hexane/CH2Cl2 extracts of soldier heads, LC on Florisil, followed by HPLC, The structure determined by spectroscopic data [300] was revised by X-ray structure analysis to rippertenol (827) [759] CH2Cl2 extract of soldiers; repeated HPLC

MS, IR, 1H and 13 C NMR, LIS NMR

[32, 300, 306, 314]

MS, IR, 1H NMR; the very small amounts prevented further investigation (e.g., 13C NMR); structure only tentatively given

[32, 111, 295, 306]

O

HO

(1R*,2R*,3S*,4S*,7S*,8Z, 11R*,12S*,15R*,16S*)-3-Hydroxy-6oxokempa-8-dien-2yl acetate (886) C22H30O4 O

HO OAc

(1R*,3S*,4S*,8S*,11R*,12S*,15R*)-Kemp-7(16)-en3-ol (887) C20H32O

HO

(1R*,4S*,7R*,9S*,11R*,12S*,15R*,16R*)-Kemp8(19)-en-9-ol (888) C20H32O OH

Nasutitermes

(continued)

Chemistry of the Secondary Metabolites of Termites

281

Table 14 (continued) Name (Nr.), formula, structure

Termite genera

(1R*,3S*,4S*,6S*,7S*,8R*,15S*,16S*)3,6-Diacetoxy-10oxo-kemp-11-en-20oic acid (889) C24H32O7 AcO

Isolation, notes

Nasutitermes CH2Cl2 extract of crushed soldiers; repeated LC on silica gel and preparative TLC [705]

Structure determination

Ref.

MS, IR, UV, 1H and 13C NMR, CD, and esterification with CH2N2 and X-ray analysis of the ester Structure revision in [756]

[32, 111, 705, 756]

O OH O

AcO

(1R*,3R*,4S*,11R*,12S*, 15R*,16S*)-6,19Epoxykempa6,8(19)-dien-3-yl acetate (Rojofuran) (728) C22H30O3

Nasutitermes

n-Hexane extract of soldier heads; flash chromatography Highly unstable in air

HRMS, 1H and 13 C NMR, and derivatization (adduct of maleic anhydride)

[32, 111, 757]

Bulbitermes, Grallatotermes, Hospitalitermes, Lacessititermes, Longipeditermes, Nasutitermes

CH2Cl2 extract of crushed termite soldiers; HPLC [336] Active against methicillinresistant Staphylococcus aureus [336]

[32, 111, 265, 302, 304–306, 309, 312, 314, 317, 323, 326, 335, 336, 376, 714]

Longipeditermes

n-Hexane extracts of crushed heads of soldiers; HPLC Found only in one species L. longipes and within that species only in specific nests

MS, [α]D, IR, 1H, 13 C, and LIS NMR, racemate synthesis [758], X-ray analysis of 3α-acetoxy-15,16epoxy-rippertane, and determination of the absolute configuration by CD of the ketone (obtained by oxidation) and by a modified method of Horeau MS, IR, 1H and 13 C NMR, COSY, and NOESY

O

AcO

(1S,3S,4S,7S,8R,11S,12S)-Rippert-15en-3-ol (727) C20H32O

HO

(1R*,3S*,4S*,7S*,8S*,11R*,12R*,16S*)-15(17)Longipen-3-ol (729) C20H32O

HO

[32, 111, 304, 312, 704]

282

E. Gössinger

The Kempanes Prestwich, who isolated the first polycyclic diterpene, the tricyclic trinervitane 760, from the frontal gland secretion of a higher termite, detected shortly afterward the first tetracyclic diterpenes 726 and 882 from the defense secretion of a related species. Five thousand soldiers of Nasutitermes kempae were taken from their arboreal carton nest [752]. Their crushed heads were extracted with n-hexane and after evaporation of the solvent, chromatography on Florisil, followed by HPLC, yielded the known trinervitane 760 and two related compounds. Their mass spectra pointed to oxygenated tetracyclic diterpenes. One of the substances crystallized and its X-ray structure analysis revealed the compact dome-shaped structure of keto acetate 726, which was named kempane-2. Compound 726 is unexpectedly unstable against oxygen, which may be caused by the strain of the molecule. The strain leads to a twisted conjugated diene (20 out of plane) that causes a rather low intensity of the absorption at 245 nm (ε ¼ 6500 mol1 dm3 cm1) in the UV spectrum. The structure of the second tetracycle kempane-1 (882) was determined by comparison of its spectroscopic data with those of kempane-2 (726). Neither the mass 1H, 13C, and LIS NMR spectra nor the IR and UV spectra allowed the stereochemistry to be determined at C-3. Thus, kempane-2 (726) was reduced with lithium aluminum hydride and esterified, which led to two diacetates 882 and 883. Again, the small amounts after the difficult separation prevented this determination. However, the absolute configuration could be determined by the positive helicity of the twisted diene chromophore [353]. The out-of-plane twist of the diene is increased in compound 883, as indicated by the surprisingly strong decrease of the absorption of the chromophore (ε ¼ 85 mol1 dm3 cm1). The stereochemistry at C-3 was determined when the same three kempadienes, 726, 882, and 883, were isolated from the secretion of the soldiers of Bulbitermes singaporensis [341]. Here, the separation of 882 and 883 was achieved by HPLC using four columns in series. Microcell 1H NMR spectrometry allowed the tentative assignment. Forty years later, the assignments were confirmed by synthesis [753]. The first kempenes with a keto group at C-6 were isolated from the defense secretion of Nasutitermes octopilis [754]. The structure determination of 885 and 886 depended mainly on the X-ray structure analysis of the p-bromobenzoate ester of 885. The β-positioned hydroxy group at C-3 of the β,γ-enone 885 points into the concave face of the dome-shaped molecule; thus the esterification proved difficult (pyridine, 4-bromobenzoyl chloride, 55 C, 5 days). The authors feared that the prolonged time and slightly elevated temperature under basic conditions might cause epimerization at C-7 or even isomerization to the conjugated enone, which did not occur, but led to wrong conclusions about the stability of the corresponding enone, thus hampering synthesis projects [760, 761]. The structure of 886 was determined from its spectrometric data and the comparison with the analogous values of 885. As one last example of the structure determination of a kempane, the only known polycyclic diterpene acid may be mentioned. Soldier heads of Nasutitermes corniger (¼ costalis) were extracted with dichloromethane [705]. The last fraction of

Chemistry of the Secondary Metabolites of Termites

283

chromatography on silica gel was further purified by preparative thin-layer chromatography. The lowest zone eluted was esterified by diazomethane. The mass spectrum pointed to two acetoxy, one methoxycarbonyl, and one keto group attached to a tetracyclic diterpene. 1H NMR spectrometry confirmed the kempane structure, but the IR and UV spectra were misleading. The strong twist of the chromophore, the unsaturated γ-keto ester with a torsion angle of 87 between the plane of the keto group and the double bond, led to the signal of a saturated ketone in the IR spectrum. The UV spectrum was characterized by the maximum at λ ¼ 223.5 nm and a very weak shoulder at λ ¼ 300 nm (ε ¼ 8 mol–1 dm3 cm–1!) [705]. Fortunately, ester 889 could be crystallized. The X-ray structure analysis revealed a kempene skeleton, where the C-7–H was epimeric to C-7–H of the kempenones 885 and 886 [756], and a pattern of substituents deviating strongly from that of the so far known kempanes. Syntheses of Kempanes The very compact dome-like shape of these tetracyclic diterpenes consisting of one cyclopentane, two cyclohexanes, and one cycloheptane is a veritable challenge to the organic chemist. Indeed, at least seven research groups were attracted to this challenge, and the rather early success of Dauben’s racemate synthesis of kempene-2 (726) was an additional stimulus [737]. However, the following five attempts [761–769] demonstrated the difficulties to estimate the steric and electronic effects governing the planned reactions. Although all five research groups were able to synthesize the kempane skeleton, none succeeded in synthesizing a naturally occurring kempane. Twenty years passed until Metz et al. succeeded in the EPC syntheses of kempene-1 (882), epi-kempene-1 (883), and kempene-2 (726) [753, 770]. Five of the seven groups used the Diels-Alder reaction as one of the key steps of their syntheses. In three cases (Dauben, Burnell, and Metz) [737, 753, 762–766, 770], the intermolecular Diels-Alder reaction with 2,6-dimethylbenzo-1,4quinone was chosen, thereby taking advantage of the ease of addition of quinones, which ensured high yields and exclusive endo-addition at relatively low temperatures even with trisubstituted double bonds. Additionally, the carbonyl groups of the generated unsymmetrically substituted cyclic enedione are regio- and stereoselectively attacked by nucleophiles [771, 772]. An interesting use of the Diels-Alder reaction was demonstrated by Hong utilizing fulvene in an IMDA reaction achieving the construction of three of the four rings of the kempanes in one step, as did Deslongchamps when using the transannular Diels-Alder reaction. Paquette used the Robinson annulation to synthesize the decalinone portion. For the construction of the further rings, intramolecular aldol reaction, an intramolecular Dieckmann condensation, and intramolecular Prins reaction as well as a McMurry reaction and metathesis were used. Remarkably, Metz constructed the five- as well as the seven-membered ring by tandem metathesis, whereas Paquette used Trost’s pentannulation method. Kato’s synthesis deviated from these synthesis designs, by continuing with the biomimetic syntheses of the neocembrene derivatives starting with neocembrenone and closing the rings stepwise by electrophilic olefin cyclization. Recently, Metz et al. presented the asymmetric synthesis of a naturally occurring 6-keto-kempenes [770]. Starting with the enantiomerically pure WielandMiescher ketone, this research group chose again the tandem metathesis as a key

284

E. Gössinger

step, which yielded a kempadienol. After confronting several stereochemical problems, the conversion into the naturally occurring 6-ketokemp-8-en-3-ol (885) was achieved. Total Synthesis of ()-Kempene-2 by Dauben et al. ()-Kempene-2 (¼ (1R*,4S*,8Z,11R*,12S*,14S*,15R*,16R*)-3-Oxokempa-6,8-dien-14-yl acetate) (726) was synthesized by Dauben et al. [737] The key steps of the synthesis plan were two intermolecular Diels-Alder reactions to implement one of the cyclohexanes and then the cyclopentene by successive ring contraction. The seven-membered ring was closed using the McMurry reaction (Scheme 130). The synthesis started with a Lewis acid-catalyzed Diels-Alder reaction of isoprene and 2,6-dimethylbenzo-1,4-quinone (890). The mixture of regioisomeric enediones (ratio not reported) was reduced to the regioisomeric diones with zinc in acetic acid simultaneously epimerizing the cis-octalindione to the more stable transoctalindione 891 in a disappointing 13% yield. Later on Metz et al. proved the reduction with zinc to be responsible for the low overall yield [753]. The less sterically hindered keto group was reduced stereoselectively in good yield, and the generated alcohol was protected as benzyl ether 892 to allow the introduction of a side chain by attacking the remaining keto group with [methoxy(trimethylsilyl) methyl]lithium. A Peterson olefination resulted in the methyl enol ether that was converted into the aldehyde by formic acid. Wittig condensation with methylene (triphenyl)phosphanylide completed the introduction of the vinyl group. The vinyl group of 893 was attacked selectively by disiamylborane followed by hydrogen peroxide under basic conditions. The primary alcohol formed was protected as benzyl ether 894. Using the less crowded more reactive borane-tetrahydrofuran complex and consecutive oxidation, the endocyclic double bond was converted regioselectively to the secondary alcohol that was oxidized to the ketone 895 by Swern’s reagent. Dehydrogenation to the enone 896 was achieved by bromination with pyridinium hydrobromide perbromide and successive elimination by lithium bromide and lithium carbonate in dimethylformamide. More than one quarter of the ketone 895 had been dibrominated yielding after debromination the bromo enone 897, which could be further debrominated reductively by tributylstannane increasing the yield of 896 to 72%. Then, annulation by Lewis acid-catalyzed Diels-Alder reaction of 896 with isoprene led stereospecifically, with high facial selectivity due to the allylic, angular methyl group, to a mixture of two regioisomers in a 2.6:1 ratio. After separation from the minor regioisomer by HPLC, the vicinal diol 898 was formed by osmium tetroxide. Subsequent hydrogenolysis removed the benzyl groups, and the vicinal diol was cleaved by sodium periodate. Acid treatment of the newly formed keto aldehyde 899 led to the intramolecular aldol reaction and thus to the cyclopentene. Under these acidic conditions, the generated exocyclic methyl ketone reacted with the primary alcohol yielding the tetracyclic oxepene 900. This allowed the protection of the remaining alcohol as an acetate. The acidic hydrolysis of the oxepene afforded the hydroxy ketone, which was oxidized by pyridinium chlorochromate to the keto aldehyde 901. Coupling of the dicarbonyl by low-valent

,

O

O

O

899

OH

80°C, 61%

TsOH, PhH,

Scheme 130 First synthesis of kempene-2 (726)

2) NaIO4, dioxane/H2O, 81%

1) H2, Pd/C, EtOAc, rt, 88%

OH

895

OBn

O

2) LiBr, Li2CO3, DMF, 120°C, 60% + 23% (2 steps)

2) NaH, BnBr, Bu4N+I-, THF, 83%

1) -Selectride, THF, –78°C to 0°C; NaOH, H2O2, 82%

2) (COCl)2, DMSO, CH2Cl2, 90%

OBn

891

O

O

1) pyr.HBr.Br 2, THF, 7–8°C

O

2) Zn, AcOH, rfl, 13% (2 steps)

BF3.OEt2,

1)

1) BH3.THF, 0°C; NaOH, H2O2, 77%

890

O

O

O

900

3) PCC/Al2O3 , n-hexane/ CH2Cl2, 57%

1) Ac2O, DMAP, pyr, 66% 2) HCl, EtOH, 80°C, 68%

PhH, rfl, 53%

O

O Br OBn OBn Bu3SnH, AIBN, 897

+

OBn

OBn

OH

O

896

892

2) HCO2H, EtOH, 73% (2 steps) 3) CH3PPh3I, t-BuOK THF, 0°C to rt, 78%

1) CH3OCH2TMS, sec-BuLi, THF, –60°C to –20°C; 892, –40°C

OBn

O

, EtAlCl2,

PhCH3, 80°C, 66% 2) HPLC

1)

901

OAc

O

O

rfl, 32%

TiCl 3(DME)1.5, Zn/Cu, DME,

HO

O

O

HO

2) NaH, BnBr, Bu4N+I-, THF, 98%

3) OsO4 cat, (CH3)3NO, acetone/H2O, 95%

893

OBn

1) HB(sia)2, THF, 40°C; H2O2, NaOH, 0°C, 82%

898

726

OAc

OBn

OBn

894

OBn

OBn

Chemistry of the Secondary Metabolites of Termites 285

286

E. Gössinger

titanium according to McMurry [773] yielded in 32% the racemic kempene-2 (726), thus completing the synthesis in 23 steps and 0.05% overall yield. Asymmetric Syntheses of Kempene-2, Kempene-1, and 3-epi-Kempene-1 by Metz and Schubert Exactly 20 years passed since Dauben published the racemate synthesis of kempene-2 (726) before the next total synthesis of kempene-2 (726) was presented by Metz and Schubert [753]. Metz et al. adopted Dauben’s construction of the substituted octalinone for their own synthesis plan. Slight variations improved the yield, and, most importantly, the synthesis was transformed to an asymmetric one by using a chiral catalyst for the initial Diels-Alder reaction. The synthesis plan of Metz deviates from that of Dauben in the annulation of the cyclopentene and cycloheptane. Whereas Dauben used a Diels-Alder reaction and subsequent ring contraction for the cyclopentene formation and McMurry coupling to close the seven-membered ring, Metz solved this problem elegantly by ring-closing domino metatheses [774, 775], thus considerably improving the yield compared to Dauben’s synthesis. The authors used the chiral oxazaborolidine-aluminum bromide complex, which was introduced by Corey et al. as an excellent catalyst for Diels-Alder reactions with quinones [776–778]. Indeed, 2,6-dimethylbenzo-1,4-quinone (890) was added to isoprene at 78 C quantitatively and with high enantioselectivity (Scheme 131). The enedione moiety of the enantiopure Diels-Alder adduct 902 was reduced to the dione moiety with zinc in acetic acid, which simultaneously isomerized the cisoctalindione to the trans-octalindione. Following Dauben’s synthesis the less sterically hindered ketone was reduced regio- and stereoselectively by L-Selectride®, and the alcohol formed was protected as methoxymethyl ether 903. The remaining keto group of 903 was used to introduce the first side chain. Peterson olefination with [methoxy(trimethylsilyl)methyl]lithium yielded the methoxymethylene group. Successively, the enol ether 904 was cleaved, and the resulting aldehyde was transformed to the homologous methyl enol ether 905 via Wittig olefination. Acidic methanol transformed the enol ether into the dimethyl acetal and cleaved simultaneously the methoxymethyl ether. The alcohol formed was now protected as silyl ether 906. To introduce the two further side chains for the planned tandem dienyne metathesis, the double bond of the octalin 906 was hydroborated, the generated borane oxidatively removed, and the resulting alcohol oxidized to the ketone 907. Dehydrogenation to the enone 908 was accomplished by silyl ether formation under thermodynamic conditions and subsequent oxidation with 2,3-dichloro-5,6dicyanobenzo-1,4-quinone (DDQ) [761, 779]. At this point, Metz et al. left Dauben’s synthesis pathway. After transacetalization with acidic acetone, the resulting aldehyde was converted chemoselectively into the trisubstituted double bond by Wittig olefination at 0 C. The introduction of a higher substituted double bond was essential to ensure the sequence of the planned tandem ring-closing metathesis [775]. Subsequently, the propargyl unit was attached by 1,4-addition to the enone moiety of 908. To achieve this goal, the researchers used trimethyl (propargyl)aluminate prepared from propenyl bromide via propargyl lithium [780] and trimethylalane. When added to enone 909 and t-butyldimethylsilyl triflate [781],

905

OMOM

O

cat.,

~ 2:1

AcO

883

OAc

, BuLi,

OTBS

O

913

OTBS

911

O

OTBS

1) TBAF, THF, rfl O

O

2) DDQ, PhH, rt, 67% (2 steps)

1) TMSI, HMDS, THF, rt

MesN NMes 5 mol % Cl Ru CH2Cl2, Ph Cl PCy3 rfl, 92%

I 3) CH3Li, 0°C; , HMPA, –20°C, 92%

O

O

O

726

OAc

90%

OTBS 912

OTBS

O

908

O

2) CH3OCH2PPh3I, LiHMDS; THF, 0°C to rt, 78%

1) TFA, CH2Cl2, rt, 69% (2 steps)

CH3Ph, rfl,

1:1.3

+

OMOM 904

2) Ac2O, pyr, DMAP, CH2Cl2, 91% (2 steps)

910

OTBS

907

OTBS

O

1) HCl, THF, rt 2) TMSI, HMDS,THF, rt, 97% (2 steps)

O

O

903, –78°C to –60°C; KH, –60°C to rt

CH3OCH2TMS, sec-BuLi, THF, –60°C to –23°C;

3) Ac2O, pyr, DMAP, CH2Cl2, 100% (2 steps)

TBSO

2) Dess-Martin ox, CH2Cl2, rt, 91%

1) BH3•S(CH3)2, 0°C; NaOH, H2O2, 83%

903

OMOM

O

1) TBAF, THF, rfl 2) LAH, THF, 0°C, 86%

(CH3)3Al; 909, TBSOTf, 84%

THF, –78°C;

Br

906

O

3) MOMCl, i-Pr2NEt, CH2Cl2, rfl, 96%

1) Zn, AcOH, rfl, 31% 2) L-Selectride®, THF, –78°, 90%

Scheme 131 Asymmetric syntheses of the kempenes 726, 882, and 883

882 OAc

+

909

OTBS

2) TBSOTf, 2,6-lutidine, CH2Cl2, 0°C, 95%

2) Ph3P+i-Pr Br-, BuLi, O THF; 0°C, 90%

AcO

O

O

902

1) TsOH, CH3OH, rfl, 87%

, CH2Cl2, –78°C, 94% ee, 98%

Br3Al

Ph Ph H O N B

1) PPTS, (CH3)2CO, rfl, 96%

890

O

O

Chemistry of the Secondary Metabolites of Termites 287

288

E. Gössinger

the preferred axial attack led stereoselectively to the silyl enol ether of the highly substituted octaline 910. To introduce the third side chain by α-alkylation, the voluminous silyl enol ether was cleaved chemoselectively by acid treatment at room temperature and exchanged for the easily cleavable trimethylsilyl enol ether. Allylation was achieved by removal of the trimethylsilyl group with methyllithium and addition of allyl iodide to the in situ-formed enolate. The reaction conditions led to a mixture of O- and C-allylation in favor of the O-allylation product 912 to the desired C-alkylation product 911 in a 1.3:1 ratio. The allyl enol ether 912 underwent Claisen rearrangement when heated at 110 C. Surprisingly, alkylation and rearrangement led with high stereoselectivity to the same stereoisomer 911. Most probably, both the ionic and the concerted reactions occur with the cyclohexene in a twist or boat conformation to avoid strong sterical hindrance by the angular methyl group and the axial propargyl group, respectively. Ketone 911 contained all stereogenic centers of the target molecule in the correct configuration and was now converted into the tetracyclic kempene derivative 913 by the domino metathesis reaction (Scheme 132). Due to the different degrees of substitution in the two double bonds, the sequence of the ring formation was predictable [775]. Addition of only 5 mol% of Grubbs II catalyst sufficed to obtain exclusively the desired tetracycle 913 in high yield. After exchange of the silyl group by the acetyl group, the synthesis of enantiomerically pure kempene-2 (726) was completed in 23 steps and 3.2% overall yield. Removal of the silyl group of 913 and subsequent reduction of the ketone at C-3 with lithium aluminum hydride furnished a mixture of diols. After separation of this mixture on Ph LxRu LxRu

LxRu

Ph

+

O

O OTBS 911

LxRu

O

OTBS

OTBS

LxRu

LxRu

O OTBS

913

O OTBS

+ O

Ph

O

RuLx

MesN NMes Cl Ru Ph Cl PCy3

OTBS

Scheme 132 Mechanism of the domino metathesis reaction

OTBS

LxRu

Ph

Chemistry of the Secondary Metabolites of Termites

289

silica gel, acetylation yielded two further naturally occurring defensive substances of termite soldiers kempene-1 (882) and 3-epi-kempene-1 (883) in enantiomerically pure form. Asymmetric Syntheses of (1R,3R,4S,7S,11R,12S,15R,16S)-3-Hydroxykemp-8en-6-one by Metz et al. Again, the key step of this synthesis was the domino metathesis [770]. Since the target molecule (1R,3R,4S,7S,11R,12S,15R,16S)-3hydroxykemp-8-en-6-one (885) contains no hydroxy group at C-14, the investigators abandoned the construction of the substituted trans-octalinone according to Dauben. Instead they chose the chiral Wieland-Miescher ketone (914) as starting material, prepared by stepwise Robinson annulation with the axial chiral organocatalyst (915) (Scheme 133) [782]. To differentiate between the two keto groups, the unsaturated ketone was selectively protected as dithiolane. To ascertain monoalkylation, the remaining ketone was converted into the sodium enoxytriethyl borate by deprotonation with sodium hexamethyldisilazane and addition of triethylborane [783, 784]. Subsequent addition of methyl iodide yielded the epimeric monomethylated products 916 and 917. Base treatment of the mixture formed the enolate. The subsequent protonation using methanol at very low temperature led by axial attack to octalinone 917, which was purified by crystallization. The following additions of the side chains, necessary for the domino metathesis, follow the pathway developed by Metz et al. within the kempene-2 synthesis. The small deviations the researchers introduced led to very high yields of most of the individual steps. The exception is the first enol ether formation. Here the researchers used instead of the Peterson elimination the WittigHorner reaction, which is a suitable method for the reaction with sterically hindered ketones [785, 786]. Addition of deprotonated (methoxymethyl)diphenylphosphine oxide at low temperature yielded after addition of methanol the adduct 918 in 44% yield and starting material 917. Treatment of the isolated adduct 918 with base furnished the enol ether 919. The further elongation steps were hydrolysis of the enol ether and Wittig condensation of the formed aldehyde by (methoxymethyl) triphenylphosponium chloride and base. The enol ether was again hydrolyzed to the corresponding aldehyde 920. A further Wittig condensation with isopropyltriphenylphosphonium iodide completed the construction of the prenyl side chain. Mild hydrolytic cleavage of the dithiolane, assisted by copper(II), afforded the substituted octalinone 921. For the following methylation, the method via sodium enoxytriethyl borate was used. Although the configuration of the newly formed stereogenic center was lost in the following steps, it proved necessary to convert the epimeric mixture into the slightly more stable epimer 922 by enolate formation with base and addition of methanol at very low temperatures. Only the epimer 922 yielded exclusively the trans-octalinone by Birch reduction [787]. To attach the two remaining side chains, the trans-decalinone had to be dehydrogenated to the trans-octalinone 923. Mukaiyama’s mild dehydrogenation method was chosen [788]. The decalinone was converted into the trimethylsilyl enol ether. At low temperature addition of methyllithium formed the enolate, which added to N-t-butyl phenylsulfinimidoyl chloride. The generated unstable adduct immediately

922

,

S

919

O

915

3) CH3Li,THF, 0°C; Ph O S Nt-Bu Cl –78°C, 91% (2 steps)

1) Li, NH3, t-BuOH, –78°C, 80% 2) TMSI, HMDS, CH3CN, rt

S

NH

O S O O

NH NH

OCH3 Cl

923

Li

S S

2) aq HF, CH3CN, CH2Cl2, rt; aq HCl, rt, 77% (2 steps)

TBSOTf, THF, –100°C to –80°C

1)

NaHMDS, THF, –50 to 0°C; 919, rt 3) TFA, CH2Cl2, H2O, rt, 95% (2 steps)

2) Ph3P

Al

–78°C; BEt3, –78°C; CH3I, –78°C to rt, 916:917 = 3:1, 92%

1) (CH2SH)2, CSA, AcOH, rt, 98% 2) NaHMDS, THF,

1) TFA, CH2Cl2, H2O, 0°C to rt, 85%

914

O

Scheme 133 Asymmetric synthesis of kempane 885, part 1

O

86 %

O 1) exc. NEt3 , rt

2) cat (2 mol%) , O PhCO2H, rt, 7 d, 94%, (2 steps), 94% ee

NaH, DMF, 0°C to rt,

cat:

O

O S S 917

LiHMDS, THF, –78 to 0°C;

916

+

924

O

O

I , HMPA,, –30°C to 0°C, 83% (2 steps)

1) TMSI, HMDS, CH3CN, rt 2) CH3Li, THF, 0°C;

2) Cu(BF4)2xH 2O, CH3CN, CH2Cl2, rt, 74% (2 steps)

1) Ph3P I BuLi, THF, –7°C to 0°C; 920, –7 to 0°C

CH3OH, –100°C; recryst., 74%

920

S

O

S

O

O

O

925

921

S

918

O

P(O)Ph2

1) NaHMDS, THF, –78°C; BEt3, –78°C; CH3I, –78°C to rt

S

HO

O PhCH3, Δ, 100%

+ 926

2) LiHMDS, THF, –78°C to 0°C; CH3OH, –100°C, 98% (2 steps) ,

917, –78°C; CH3OH, –100°C, 918 (44%) and 917 (56%)

O Ph P OCH3 Ph BuLi, THF, –78°C;

290 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

291

underwent cis elimination to the octalinone 923 in very high yield. As in their former synthesis, the propynyl unit was attached by 1,4-addition of trimethyl(propargyl) aluminate, and the enolate was trapped as t-butyldimethylsilyl enol ether. The relatively stable silyl group was exchanged by reaction with aqueous hydrofluoric acid to the ketone 924 and resilylation with trimethylsilyl iodide. Allyl iodide was added to the in situ-generated enolate furnishing a mixture of the C- and O-allylated products 925 and 926, which was heated in a sealed tube converting the enol ether 926 by Claisen rearrangement to 925 in high yields. Domino metathesis of 925 with surprisingly low amounts of Grubbs II catalyst led to kempadienone 927 (Scheme 134). Reduction of 927 by L-Selectride® led exclusively to the undesired alcohol 928, with the hydroxy group at the convex side of the dome-shaped molecule. Fortunately, the decalinone 925 provided the desired alcohol 929 by reduction with L-Selectride®. Domino metathesis of this alcohol 929 furnished the kempadienol 930 in excellent yield. Racemic 930 had been synthesized by Kato et al. in their biomimetic approach toward naturally occurring kempanes [767]. Contrary to their expectations, Metz et al. found that the double bond of the cycloheptene moiety of kempadienol 930 is more reactive than that of the cyclopentene moiety. Thus, the double bond of the cycloheptene had to be protected. After protection of the hydroxy group of 930 as a silyl ether, the double bond of the cycloheptene moiety was dihydroxylated by osmium tetroxide. As expected, the approach of the voluminous reagent occurred from the convex side of the molecule. Many trials were necessary to achieve the attack of an oxygenation reagent at the sterically hindered double bond of the generated vicinal dihydroxy compound 931. The researchers succeeded when using in situ-prepared bis (trifluoromethyl)dioxirane. This reagent attacked the double bond of 931 exclusively from the concave side resulting in epoxide 932 in surprisingly high yield. To remove the now unnecessary protection of the cycloheptene double bond, the vicinal diol was converted into the cyclic thionocarbonate 933 by 1,10 -thiocarbonyldiimidazole catalyzed by 4-(dimethylamino)pyridine. Subsequent reductive removal of the cyclic thionocarbonate by the phosphor(III) reagent 934 was followed by treatment of the intermediate unsaturated epoxide with silica gel in a n-pentane/ether mixture, which led to the β,γ-enone 935 with the undesired configuration at C-7 due to the 1,2-hydrogen shift within the transformation of the epoxide to the ketone. The hope that by liberating the alcohol under relatively mild acidic conditions would also form the unsaturated enol, followed by intramolecular protonation to the desired β,γ-enone 885, was not realized, because the more stable α,β-enone 936 was formed. As the research groups of Paquette et al. and Deslongchamps et al., who synthesized racemic 936 earlier [761, 769], experienced, all their trials to isomerize 937 to the desired β,γ-enone 885 proved futile. Metz et al. surmounted this difficult problem by adding phenylthiolate in 1,4-fashion to the α,β-enone 936 at slightly elevated temperatures, which provided the saturated ketone 937 with the correct configuration at C-7. Sulfoxide formation by aqueous hydrogen peroxide in 1,1,1,3,3,3hexafluoroisopropanol was succeeded by cis elimination at 100 C with trimethylphosphite added as a scavenger of the generated phenylsulfenic acid. The resulting mixture of the two β,γ-enones 938 and 885 was obtained in 96% yield. The

929

-Selectride®, THF, –78°C, rt, 92%

925

t-Bu

939

t-Bu

N N Co O O

t-Bu

t-Bu

TBSO

O

933

O

S

CH2Cl2, rfl, 93%

O

MesN NMes 3 mol % Cl Ru Ph Cl PCy3

CH2Cl2, rfl, 97%

HO

Ph P N

OH OH

O

HO

O

885

+ 55:41

O

HO 939 cat., PhSiH3, acetone, rt, 64%

935

938

RhCl3•3H2O, EiOH, 70°C, 64%

aq HF, CH3CN, CH2Cl2, rt, 99%

936

O

932

OH OH

939 cat., PhSiH3, PhSTs, EtOH, rt, 82%

HO

O

937

SPh

exc PhSH, LiOH•H2O, THF, 60°C, 86%

TBSO

1) (CF3)2CHOH, H2O2 aq, rt 2) P(OCH3)3, PhCH3, 100°C, 96%

HO

O

glyme, CH3CN, H2O, 0°C, 91% 931

928

2) OsO4, 3,5-lutidine, THF, PhCH3, –78°C, 99% TBSO

HO

1) TBSOTf, 2,6-lutidine, CH2Cl2, 0°C, 98%

rt, 73%

F3C O F3C O NaHCO3, Na2EDTA,

934

THF, 57°C;

N

930

927

-Selectride®, THF, –78°C,

SiO2, C5H12/ ether, rt, 70% TBSO

O

Scheme 134 Asymmetric synthesis of kempane 885, part 2

cat:

(imid)2CS, DMAP, CH2Cl2, rt, 99%

HO

O

MesN NMes 3 mol %, Cl Ru Ph Cl PCy3

292 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

293

desired compound 885 showed the same circular dichroism as the natural product, thus confirming Prestwich’s assumption [754]. The authors developed several ways of isomerizing the exocyclic double bond of the β,γ-enone 938. Rhodium trichloride in ethanol converted 938 into the α,β-enone 936. A recently developed mild addition of sulfur compounds to the unactivated double bound catalyzed by the cobalt (II) catalyst 939 [789] led back to the phenyl sulfide 937. The active catalyst formed from the cobalt complex 939 with phenylsilane is thought to be a cobalt hydride complex, which delivers the hydride to the less substituted carbon of the double bond and enables the addition of the thiophenyltosylate (S-phenyl 4-toluenesulfonothioate) [790] affording the phenyl sulfide 937. The assumption of the intermediate cobalt hydride led the authors to try direct isomerization of 938 with cobalt complex 939 and phenylsilane in acetone. Indeed, this reaction provided the best recycling variant raising the yield of the target molecule 885 to 78%. The synthesis was completed in 31 steps and 1.8% overall yield. Its progress was supported by X-ray analyses of several intermediate compounds. As discussed in the following paragraphs, four other research groups attempted to synthesize naturally occurring kempanes. Attempts Toward the Total Synthesis of Naturally Occurring Kempanes: Total Syntheses of (1R*,3S*,4S*,11R*,12S*,15R*,16S*)-3-Hydroxykemp-7-en-6-one, (1R*,3S*,4S*,7S*,11R*,12S*,15R*,16S*)-3-Hydroxykemp-8-en-6-one, and (1R*,3R*,4S*,11R*,12S*,15R*,16S*)-3-Hydroxykemp-7-en-6-one by Paquette et al. One year after Dauben’s successful synthesis, Paquette et al. published their synthesis efforts toward the racemic kempanes [761]. Key steps included Robinson annulation to construct the octalinone, Trost’s palladiumcatalyzed five-ring annulation [791], and an aldol reaction to close the sevenmembered ring (Scheme 135). 2-Methylcyclohexa-1,3-dione was transformed into the bulky monoenol ether with t-butanol and acid to ensure the selective methylation at the C-6 position. 1,2-Addition of 3-methylbut-3-enylmagnesium bromide (943) to the resulting substituted enone 942 was followed by acidic workup leading to enone 944 in 72% overall yield. Enone 944 was reduced by lithium in ammonia. The generated enolate permitted regioselective Robinson annulation. To attain higher yields, Stork’s variant of the Robinson annulation was performed [792, 793]. After removal of ammonia, 3-(trimethylsilyl)but-3-en-2-one was added to the ether solution of the enolate at 78 C. The neighboring substituents effected stereoselective addition; the newly formed enolate cyclized immediately to hemiketal 945. Treatment of the hemiketal with potassium hydroxide completed the annulation to the substituted octalinone 946. Reduction with dissolved metal yielded the trans-decalinone 947. Introduction of the methyl group and annulation of the five-membered ring were planned next. Several trials were necessary to find an acceptable solution. Introduction of the methyl group prior to the annulation step proved unsatisfactory. Thus, the researchers examined the efficiency of the phenylthiocarbonyl and the methoxycarbonyl group as a replacement of the methyl group. The modified decalinones were dehydrogenated. Stepwise annulation by copper-catalyzed

HO

947

951

943

MgBr

O

O

2) O3, pyr, CH2Cl2, –78°C; (CH3)2S, 60%

1) TBSOTf, Et3N, THF, rt, 94%

2) DDQ, THF, Et3N, 77%

O

O

THF, rfl; aq HCl, 84%

1) (CH3O)2CO, NaH, DME, rfl, 81%

942

O

Scheme 135 Synthesis of the kempanes 941 and 942

2) LiEt3BH, THF, 0°C to rt; NaOH, H2O2, rt, 94%

1) TsCl, DMAP, Et3N, CH2Cl2, 63% + 11% 950

–78°C, 80%

O

2) LDA,THF, i-BuO –78°C; CH3I, 86% (2 steps)

1) iBuOH, TsOH, benzene, rfl

Li, NH3, t-BuOH,

O

O

O

TBSO

948

944

O

, ether,

952

O

P(OEt)3, THF, rfl, 98% O

O O

TMS O

949

OH 945

2) HF, CH3CN, rt, 87%

1) K2CO3, CH3OH, rt, 59%

OAc,

Pd(OAc)2,

TMS

–78°C to rt,

TMS

Li, NH3, t-BuOH, –78°C;

HO

O

941

90%

LAH, THF, –78°C to rt,

5:1

+

42% (3 steps)

KOH, CH3OH, H2O, rfl,

HO

O

HO

HO

O

942

950

946

294 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

295

1,4-addition of but-3-enylmagnesium bromide, followed by oxidative cleavage of the terminal double bond, and intramolecular aldol reaction was discarded due to low yields. Therefore, the researchers examined annulation by trimethylenemethane analogues [791]. Lewis acid-catalyzed addition of [2-(chloromethyl)allyl]silane [794] failed, but Trost’s palladium-catalyzed pentannulation of 2-[(trimethylsilyl) methyl]allyl acetate [791] proved successful. Trost had demonstrated that electronwithdrawing groups at the double bond are advantageous for this annulation method; thus, the following reaction sequence was chosen: Claisen condensation of decalinone 947 with dimethyl carbonate afforded regioselectively the β-ketoester, which was dehydrogenated by 2,3-dichloro-5,6-dicyanobenzo-1,4-quinone to the octalinone derivative 948. 2-[(Trimethylsilyl)methyl]allyl acetate, palladium acetate, and triethylphosphite were added to 948 in tetrahydrofuran, and the mixture was kept under reflux. Due to the rigid conformation of the octalinone, the annulation occurred with high stereoselectivity yielding nearly quantitatively the tricyclic compound 949. The next steps served the conversion of the methoxycarbonyl group into the methyl group. Lithium aluminum hydride reduced the carboxyl group and the keto group. The high stereoselectivity of the reduction of the ketone is noteworthy as the reagent approached from the more hindered face to preserve the chair conformation and no directing effect of the carboxyl group or its reduction products was observed. Selective tosylation of the primary alcohol of diol 950 proved difficult due to its neopentyl position. Despite slow addition of tosyl chloride at low temperature, traces of the ditosylate were detected next to the 6:1 mixture of the desired monotosylate and starting material. Reduction to the methyl group by an excess of lithium triethylborohydride was followed by protection of the secondary alcohol of 951 as a silyl ether, which proved necessary to achieve acceptable yields within the following ozonolysis of the two methylene groups. The diketone 952 was the prerequisite for the cyclization to the seven-membered ring by aldol reaction with potassium carbonate in methanol. The tetracyclic kempenones obtained consisted of a 5:1 mixture of the conjugated enone 941 and its deconjugated isomer 942, the epimeric alcohol of the naturally occurring hydroxykempenone 885. Prestwich had reported that the naturally occurring deconjugated enone 885 could not be isomerized to the conjugated enone under standard conditions [795]. Also, according to force field calculations, the deconjugated enone 885 is the more stable compound of the two isomers [737]. Thus, the researchers assumed facile isomerization of the conjugated enone 941. However, they found “no set of conditions to accomplish their interconversion.” Later on, Taber using semiempirical calculations revealed that the conjugated enone 936 is the more stable compound [760]. Since Paquette et al. assumed that the isomerization is only kinetically prevented, they hoped that epimerization of the alcohol might change the conformation slightly and thus facilitate the desired double-bond isomerization. Starting with the tricyclic alcohol 950, epimerization via Mitsunobu reaction as well as via tosylation failed. Therefore, oxidation and consecutive reduction were tested. To benefit from the directing effect of a neighboring group, diol 950 was chosen as starting material (Scheme 136). To oxidize the secondary alcohol selectively, ammonium molybdate was used [796]. The resulting ketone was reduced with sodium triacetoxyborohydride [797] to

296

E. Gössinger

1) (NH4)6Mo7O24, K2CO3, H2O2, Bu4N+Cl-,

HO HO 950

1) DMP, PPTS, acetone, rt, 95%

HO HO

73% + 18% 950 2) NaBH(AcO)3, THF, rt, 94%

953

2) O3, CH3OH, –78°C; (CH3)2S, 86%

O O

O 1) K2CO3, CH3OH, rfl, 91%

O

2) PPTS, CH3OH, rt, 100%

O 954

1) CS2, DBN, DMF, rt; CH3I,

HO HO 955

2) (TMS)3SiH, AIBN, PhH, rfl, 38% (2 steps)

O

HO 936

Scheme 136 Synthesis of kempane 936

the epimeric diol 953. Conversion of the hydroxymethyl group into the methyl group as used with diol 950 (Scheme 135) by tosylation and reduction failed with the epimeric alcohol 953 due to Grob fragmentation [798]. Therefore, diol 953 was protected as a cyclic ketal prior to ozonolysis. Aldol reaction of the generated diketone 954 yielded the conjugated enone exclusively. Deconjugation could not be achieved, neither with this compound nor with the diol 955, obtained by acid treatment, nor with 3-hydroxykemp-7-en-6-one 936, obtained by selective dehydroxylation of 955 via esterification of the primary alcohol to the methyl xanthate and radical reduction with tris(trimethylsilyl)silane and 2,20 -azobisisobutyronitrile. Total Synthesis of (1R*,3R*,4S*,8Z,11R*,12S*,15R*,16R*)-Kempa-6,8-dien-3ol by Kato et al. Kato et al. developed a biomimetic synthesis protocol for mono-, di-, and tricyclic diterpenes starting with open-chain geranylgeranoic acid. Within their synthesis of the dihydroxytrinervitane 725, they observed that the electrophilic olefin cyclization of bicyclic 17-chlorosecotrinervitane (826) could be steered to the tricyclic trinervitanes or to the tetracyclic kempanes (Scheme 123) depending on the reaction conditions [732, 734]. With the construction of the skeleton of the kempanes solved, the researchers turned their attention to the suitable functionalization of the tetracyclic dienediol (830). Preliminary tests showed that the two hydroxy groups as well as the two tetrasubstituted double bonds of 830 differed enough in their reactivity to allow selective transformations. Thus encouraged, the researchers started their planned synthesis of 14-acetoxykempa-6,8-dien-3one (kempene-2 (726)) [767], one of the defensive compounds of nasute soldier secretion [6, 754]. Their first task was to upgrade the yield of the transannular cyclization of the secotrinervitane to kempane (Scheme 123). The use of triol (957) turned out to be more advantageous than that of 17-chlorosecotrinervitane (826), under equal reaction conditions, i.e. perchloric acid generated in situ from tbutyl chloride and silver perchlorate (Scheme 137).

OR

O

O

O

O

H

964

O

H

960

O

OH

O

5:1

+

O

O

O

LiBr, DMF, 105°C, 100%

2) t-BuCl, AgClO4, THF, rt, 49%

1) HCl, CH3OH, 90%

Scheme 137 Synthesis of kempane 967

O

Cl

957 (R = H)

823 (R = MOM)

RO

OH

965

O

H

OH

O

O

830

OH

OH

961

O

N O

N

N

CH2Cl2, –40°C 98%

MCPBA, NaHCO3,

2) NaOCH3, CH3OH, 0°C, 89% (2 steps)

1) Dess-Martin ox., rt

O

PhH, rt, 93%

N

O

O

O 958

O

O

O

H

962

966

O

O

O

O

O

O

O

O

O

O

963

O

959

O

O

O

H

CH3I, THF, 95% 3) Ph3SnH, AIBN, PhCH3, 110°C, 61%

1) NaBH4, CH3OH, –30°C, 88% 2) CS2, NaH;

4:1

+

with recycling 94%

SnCl4, t-BuCl, ether, rt, 50%

Cl

O

O

O

O

967

O

H

EtOH, 77%

H2, Pd/C, NaHCO3,

rt, 50%, with recycling, 85%

PDC, t-BuOOH, PhH, Celite,

Chemistry of the Secondary Metabolites of Termites 297

298

E. Gössinger

Although the selective protection of the C-3 hydroxy group is possible, protection of the diol 830 as cyclic carbonate 958 was preferred, because the additional ring led to a change in the conformation, which proved advantageous at a later step of the planned synthesis. To manipulate the Δ11,12-double bond of the cyclic carbonate 958, the more reactive Δ7,8-double bond had to be protected. Epoxidation led to a mixture of products, but hydrogen chloride addition by the in situ-prepared, bulky hydrogen pentachlorostannate in ether at room temperature led to the single 8-chlorokempane derivative 959. To deactivate the Δ11,12-double bond and enable trans addition of hydrogen as well as acquire a handle for the functionalization of the C-13 methylene group, oxidation to the enone 960 was envisaged. Regioselective allyl oxidation was attained by the action of pyridinium dichromate and t-butyl hydroperoxide [799]. Next, the Δ7,8-double bond was regenerated by treatment of the chloride 960 with lithium bromide in dimethylformamide at elevated temperature. Selective epoxidation of dienone 961 with 3-chloroperbenzoic acid at 40 C led to a mixture of the two stereoisomeric epoxides 962 and 963 with the α-epoxide 962 as the main product. The trans hydrogenation proved difficult because the enone was either inert or led to mixtures, for example, when dissolved metals were used. Hydride reduction catalyzed by copper salts reduced only the carbonyl group. Even hydrogenation with Adams catalyst led only to reduction of the carbonyl group. When palladium on charcoal was used as catalyst, the conjugated enone moiety of α-epoxide 962 was reduced to the 5:1 mixture of the saturated α-alcohols 964 and 965, whereas the enone group of β-epoxide 963 was inert against hydrogenation due to stronger steric hindrance. As expected, 964, generated by the cis-addition of hydrogen, was the main product. To convert 964 into the desired product 966 with the methyl group in an equatorial position, the saturated alcohols 964 and 965 were oxidized by Dess-Martin periodinane and subsequently treated with base, which led to the single ketone 966. Deoxygenation to 967 was achieved by reduction to the alcohol, esterification to methyl dithiocarbonate, and radical reduction with triphenylstannane. The next steps were devoted to the differentiation of the two alcohols at C-2 and C-3 (Scheme 138). Thus, the carbonate 967 was saponified, but unequal to the diol 957, the newly formed diol could not be protected selectively. The authors overcame this problem by intramolecular transesterification. The diol was treated with pivaloyl anhydride leading to a 1:1 mixture of the two possible monopivalates 968 and 969. The unchanged hydroxy groups were oxidized to the ketones 970 and 971, and subsequent treatment with base led to the sterically least strained keto pivalate 972 (Scheme 139). The now unwanted ketone was removed by reduction to the alcohol, esterification to the methyl dithiocarbonate, and radical reduction. To transform the epoxide 973 into the conjugated diene 974, the authors developed a new method by treating the epoxide with trimethylsilyl chloride at 0 C, which resulted in regioselective, quantitative diene formation. The authors concluded their efforts by removing the pivaloyl protecting group of diene 974 reductively yielding the alcohol kempa-6,8dien-3β-ol (956). This end product resembles the naturally occurring kempanes

Chemistry of the Secondary Metabolites of Termites

1) KOH, CH3OH, 98%

H

O

299

+

2) (Piv)2O, NaH, THF, 0°C, 94% O

OPiv

O 967

O

1:1

H

O

H

O

+

88% OPiv

PivO O

NaOCH3, CH3OH, rt, 83%

OPiv 971

1:1

1) NaBH4, CH3OH, –30°C to rt, 87% 2) CS2, NaH, THF, 0°C to rt;

O

OPiv

969

O

O

970

972

OH

OH

968

Dess-Martin ox., pyr, CH2Cl2, rt,

H

O

H

O

CH3I, THF, rt, 98% 3) Ph3SnH, AIBN, PhCH3, 110°C, 100%

O

(CH3)3SiCl, THF, 0°C to

PivO

rt, 100%

973

LAH, THF, PivO

rt, 100% HO

974

956

Scheme 138 Synthesis of kempadienol 956

closely, and it would not be surprising if this molecule were detected in the secretion of the frontal gland of a yet to investigate termite species. Burnell’s Several Attempts Toward the Kempenes As Dauben et al. and later Metz et al., Burnell chose [762–766] the intermolecular Diels-Alder reaction with 2,6-dimethylbenzo-1,4-quinone (890) as key step. The diene was chosen carefully. It incorporated the cyclopentane unit and, as part of the seven-membered ring of the desired kempane skeleton, a γ-lactone. This cis-oxadiquinanone moiety ascertained excellent facial selectivity in the Diels-Alder reaction. To construct the sevenmembered ring, three variants were considered: intramolecular aldol reaction, intramolecular Dieckmann reaction, and finally metathesis. The construction of cisoxadiquinanone 979 followed the protocol of Corey et al. closely [800], which offered the advantage of a convenient racemate resolution [614] for future EPC syntheses of kempanes.

300

E. Gössinger

O

O O

O

971 B

O

-

O

O

O

O

O

O

O O

O O

O

HB

O

O O O

972

Scheme 139 Base-catalyzed equilibration of 971 and 972

The starting material, 3-methylcyclohex-2-enone 975 (Scheme 140), was transformed into the conjugated cyclohexadiene 976 via the tosylhydrazone and base treatment according to Shapiro [801]. Chemo- and regioselective [2+2]addition with in situ-prepared dichloroketene was followed by reductive removal of the obsolete chlorine atoms by zinc in slightly acidic methanol [802]. Regioselective Baeyer-Villiger reaction of the bicyclic cyclobutanone 977 was attained by hydrogen peroxide in acetic acid. Ring contraction of the cyclohexene moiety of lactone 978 to the cyclopentene 979 was achieved by ozonolysis and consecutive aldol reaction under acidic conditions [762, 763]. Enol ether formation of the methyl ketone of 979 by t-butyldimethylsilyl triflate completed the synthesis of diene 980. As expected, the cycloaddition with 2,6-dimethylbenzo-1,4-quinone (890) furnished the tetracyclic enedione 981 regio-, stereo-, and facial-selectively with only 6% of the regioisomer. The researchers found that the regioselectivity of this Diels-Alder reaction is independent of further substituents at the lactone moiety but influenced by the enol-protecting group. The predicted differentiation of the two carbonyl groups of the enedione 981 was

985

O

CO2Et

TBSO

O

O

O

2) MEMCl, DIPEA, CH2Cl2, rfl, 92% MEMO

O

986

O

2) KF, CH3OH, rt, 93%

Li 1) EtO THF, –78°C, 82%

1) LiAl(t-BuO)3H, THF, 0°C, 78%

981

O

2) Zn, NH4Cl, CH3OH, rt

2) CH3Li, Et2O, 0°C

976

1) CHCl2COCl, Et3N, C5H12, rt

1) TsNHNH 2, HCl, THF, rt

Scheme 140 Synthesis of kampane lactone 988

O

O

O

PhCH3, rfl, 80%

O 890

O

975

O

O

O

HO

O

O O

OEt

7:1

+ O

O

O

2) PCC, CH2Cl2, 74% (2 steps)

CO2Et 1) Li, NH , –50°C; 3 986, dioxane, ether

982

O

977

27% (5 steps)

H2O2, AcOH, 0°C

O

O

O

CO2Et

rfl, 84%

Zn, AcOH,

2) HCl, THF, rfl, 42% (2 steps)

1) O3, CH2Cl2, –78°C; (CH3)2S

987

O

OH

OEt

978

MEMO

983

O

O

984

O

O

2) t-BuOK, PhH, rfl, 61%

980

TBSO

O

O

988

O

OH

O

rfl, 64%

HCl, CH3OH,

MEMO

O

CO2Et

CH2Cl2, 74%

TBSOTf, Et3N,

1) L-Selectride, THF, –78°C, 91%

O

O

O 979

O

O

Chemistry of the Secondary Metabolites of Termites 301

302

E. Gössinger

observed despite the rather voluminous oxadiquinanone entity, when lithium ethoxyacetylide was added. Deprotection of the enol ether of 981 by fluoride led to cyclic hemiketal formation verifying the stereochemical prediction. This cyclization was faster than the equilibration of the cis-octalinone to the more stable trans-octalinone 983, leading to a 7:1 mixture of 982:983. The mixture was treated with zinc in acetic acid, which removed the tertiary alcohol and the ether reductively, and simultaneously converted the ethyl ethynyl ether moiety into the ethyl acetate unit of 984. To isomerize the deconjugated keto ester 984 to the conjugated ketone with the ester side chain in the thermodynamically more favorable equatorial position as well as the epimerization to the desired trans-octalinone 985 required some experimentation, and was finally solved by treatment of 984 with hydrochloric acid in methanol under reflux yielding the oxygen-sensitive diketone 985. The voluminous rather weakly reductive lithium tri-t-butoxyaluminum hydride converted the saturated ketone of 985 into the axial secondary alcohol that was protected as the (2-methoxyethoxy) methyl ether 986. The dissolved metal reduction led stereoselectively to the saturated ketone 987, which was unstable under the reaction conditions and reduced to the secondary alcohol that was reoxidized by pyridinium chlorochromate. Stereoselective reduction of ketone 987 was achieved with L-Selectride® at 78 C. Dieckmann condensation with potassium t-butoxide in benzene at reflux afforded the pentacyclic β-keto lactone 988 in 61% yield. The yield of the Dieckmann condensation was increased when the hydroxy group at C-14 (kempane numbering) was protected as a methoxymethyl ether to prevent the partial lactonization of the ester group with the hydroxy group [762–764]. Introduction of the methyl group at C-4 (kempane numbering) and cleavage of the γ-lactone, the necessary steps to accomplish the synthesis of naturally occurring kempanes, seemed a straightforward task. However, so far, all efforts of Burnell’s research group proved futile; nonetheless these efforts are worth mentioning, because their partially completed syntheses demonstrate the difficulties, the intense labor, and the perseverance total syntheses of complex molecules necessitate. Some of the more successful studies are shown below. Efforts to use β-keto lactone 988 were thwarted by the inability to open the lactone or its reduction products, the lactols [763, 764]. Thus, the authors decided to open the lactone prior to the Diels-Alder reaction (Scheme 141). To open the bicyclic lactone 979 [762, 763] reductively, the exocyclic ketone had to be protected as cyclic ketal. The diol, produced by lithium aluminum hydride reduction, was protected as the stable bismethyl ether 989 by Williamson etherification. After regeneration of the ketone by acidic hydrolysis, the synthesis followed the originally devised pathway. The silyl enol ether formation was followed by the Diels-Alder reaction of 990 with 2,6-dimethylbenzo-1,4-quinone (890), and subsequently lithium ethoxyacetylide was added. The researchers found that the opening of the lactone and thus the higher flexibility of the cyclopentene part of the diene did not influence the stereochemistry of the Diels-Alder reaction to tricycle 991 nor that of the addition of the organo lithium compound to the enedione. Fluoride removed the silyl group of enol ether 992, which led immediately to the two

Chemistry of the Secondary Metabolites of Termites

303

O

O

O

O 1) (CH2OH)2, H+, PhH, rfl, 72%

O

O

2) LAH, Et2O, rt, 80% 3) NaH, CH3I, THF, rt, 81%

O

1) PPTS, acetone, H2O, rfl, 90% 2) TBSOTf, Et3N, CH2Cl2, 0°C, 91%

O

O

O 890 PhCH3, rfl, 86%

TMSO

O

979

990

989

O

O

O

OEt

O

O

Li

EtO

OH

THF, –78°C to 0°C, 80%

TBSO

KF, CH3OH, rt, 95%

TBSO O

O

991

O

992

OEt

O

O

O

OEt

O

O

CO2Et

Zn, AcOH,

+

O

O

HO

O

O 993

OH 3:2

994

O

rfl, 84% O 995

O

CO2Et

TsOH, PhCH3, rfl, 68% O 996

O

Scheme 141 Synthesis of tetracycle 996

hemiketals 993 and 994. Zinc in acetic acid cleaved the ether bridge reductively and simultaneously transformed the ethynyl ether moiety into the ethyl acetate unit of 995. Unfortunately, all attempts to isomerize the deconjugated enone 995 into the conjugated enone and simultaneously epimerize the cis-octalindione to the transoctalindione by acid treatment were accompanied by transformation of the methyl ethers to the tetrahydrofuran 996. The tricyclic enolether 991 was further used to introduce the methyl group at C-4 (kempane numbering). The investigators had learned that direct methylation, which would have secured the region as well as the stereochemistry, failed due to steric

304

E. Gössinger

O

O

O

O

O

O

CH2I2, Et2Zn, PhCH3, rt, 88%

TBSO

EtO

THF, –78°C to 0°C, 94%

TBSO

O

O

991

Li,

997

O O

O

OEt

O

O OH

+ TBSO

OH

TBSO O

OEt 998

3:5

999

Scheme 142 Addition of lithium acetylide to the 2-en-1,4-dione moiety of 997

hindrance and high oxygen sensitivity [763]. Experiments with model compounds showed that cyclopropanation of the monoenolether of the octalintrione is feasible. Modified Simmons-Smith conditions converted 991 stereoselectively in 88% yield to the cyclopropane 997 (Scheme 142). The following addition of the acetylide was no longer regioselective affording in excellent yields the two regioisomers 998 and 999. As a consequence, the researchers decided to implant the methyl group at a very late stage of their kempane synthesis. After these unexpected results, the authors decided to design the Diels-Alder diene without a side chain at the cyclopentene moiety (Scheme 143). The synthesis started with malonic diacetal 1000, which was transformed to the mono-1,3-dithiane by propanedithiol with boron trifluoride etherate. The substituted dithiane was deprotonated, and the anion replaced the primary iodide of the protected 3-keto butyl iodide yielding in 45% the protected 3,6-heptadional 1001. Acidic hydrolysis of its acetal groups by refluxing aqueous acid initiated the aldol reaction leading to the protected acetylcyclopentenone. Silyl enol ether formation by t-butyldimethylsilyl triflate completed the synthesis of diene 1002. Again, Diels-Alder reaction with 2,6-dimethylbenzo-1,4-quinone (890) and subsequent addition of lithium ethoxyacetylide proceeded with high regio- and stereoselectivity, leading via enedione 1003 to ethynyl ether 1004. Cleavage of the silyl enol ether by fluoride led to the cyclic hemiketal 1005, which in turn was reductively cleaved by zinc in acetic acid that also transformed the ethynyl ether into the ester 1006. Isomerization of the cyclic double bond of ester enone 1006 under acidic conditions led to the conjugated enone and transformed simultaneously the cis-octalindione in trans-octalindione 1007. To enable the reduction of the enone with dissolved metal, the protecting dithiane group had to be exchanged. According to Stork, using bis(trifluoroacetoxy)

S

O

O

I

2) Li, NH3, –55°C, 71%

O

S

HO

S

O

1) MOMCl, DIPEA, CH2C2, rt, 68%

rt, 78%

KF, CH3OH,

, 0°C, 84%

CO2Et

OH

OEt

O O

2) BuLi, THF, –20°C;

1) CH2(CH2SH)2, BF3•Et2O, CH3Cl, rfl, 54%

Scheme 143 Synthesis of intermediate 1010

1008

O

HO

O

1004

1000

O

O

TBSO

S

O

O

O O

1009

O

O

OEt

S

MOMO

O

1005

O

1001

S

O

CO2Et

rfl, 88%

Zn, AcOH,

O

S

1006

THF, 78°C, 74%

S

1002

TBSO

S

L-Selectride,

S

2) TBSOTf, Et3N, CH2Cl2, 0°C, 100%

1) aq HCl, THF, rfl, 85%

O

OH

CO2Et

rfl, 63%

TsOH, PhCH3,

1010

O

MOMO

O

CO2Et

PhCH3, rfl, 88%

O 890

O S

S

O

S

O

1003

1007

TBSO

S

CO2Et

O

O Li

2) L-Selectride, THF, 78°C, 83%

1) PhI(OCOCF3)2, (CH2OH)2, CH3CN, rt, 56%

THF, –78°C to 0°C, 71%

EtO

Chemistry of the Secondary Metabolites of Termites 305

306

E. Gössinger

iodobenzene in an 1,2-ethanediole-acetonitrile mixture permitted the exchange in one step [803]. Subsequently, the saturated ketone was selectively reduced by LSelectride® to the axial alcohol 1008, which was protected as methoxymethyl ether. Next, lithium in liquid ammonia reduced the enone to the saturated ketone 1009 with the methyl group in equatorial position. Again, L-Selectride® was used to reduce the saturated ketone stereoselectively to the axial alcohol 1010. In their most recent publication, Burnell et al. reexamined the bicyclic lactone 979 (Scheme 144) [766]. After protection of the exocyclic ketone of 979 as 4,4-dimethyl1,3-dioxane, the lactone was reduced by lithium aluminum hydride to diol 1011. The primary alcohol was protected as a silyl ether before the secondary alcohol was converted into the methyl ether by Williamson synthesis. Regeneration of the primary alcohol 1012 using fluoride was followed by oxidation to the aldehyde by Dess-Martin periodinane and methylenation by Eschenmoser’s reagent [804]. Then, the aldehyde 1013 was reduced to the methyl group in three steps. Reduction to the primary allylic alcohol according to Luche was succeeded by esterification to the methyl xanthate 1014 and radical reduction with tributylstannane attaining the isopropenyl group in 67% overall yield. After acidic hydrolysis of the 1,3-dioxane, the regenerated ketone was transformed to the silyl enol ether 1015. Here, the DielsAlder reaction with 2,6-dimethylbenzo-1,4-quinone (890) proceeded again regio-, stereo-, and facial-selectively. In case of the addition reaction to the enedione 1016, a different situation arose, because the substituent at the cyclopentane and the allylic metal were more voluminous as in the earlier addition reactions. Indeed, allylmagnesium bromide added exclusively at the keto group at C-14 (kempane numbering). As Burnell et al. had discovered earlier, sodium borohydride reduced mainly the carbonyl at C-14, and this regioselectivity was enhanced when Luche conditions were used [805]. Thus, the undesired regiochemistry of the Grignard addition could be circumvented by reducing selectively the keto group at C-14 according to Luche resulting in the epimeric monoalcohols 1017. When an excess of allylic Grignard reagent was added, a slow reaction occurred affording the addition products in around 30% yield next to unreacted material. To be better able to pursue their new key step, i.e., the ring closure of the seven-membered ring by metathesis, the epimeric alcohols were oxidized by Dess-Martin periodinane affording enone 1018. The metathesis proceeded with 3 mol% of a Grubbs’ catalyst II [806] and furnished the desmethylkempane 1019 in 52% yield with the required trisubstituted double bond in the seven-membered ring. EPC Synthesis of Methyl (1R,2S,3R,4S,11R,12S,14S,15R,16S)-4-Desmethyl-2chloro-3-hydroxykempa-6,8-dien-3-yl Carboxylate by Hong et al. Hong et al. chose fulvenes, very versatile templates in concerted as well as in ionic reactions, to construct polycyclic systems. One of the polycyclic ring systems they thought to be accessible by an IMDA reaction with fulvene was the kempane skeleton. It has been demonstrated that fulvenes react as dienes when treated with electron-rich dienes but as dienophiles when treated with electron-poor dienes [807]. The scope of this statement was tested with a model compound, before the researchers turned their

MgBr

ether, 0°C

O

OCS2CH3

O

1) Bu3SnH, AIBN, PhH, rfl, 96% 2) PPTS, CH3OH,

1011

O

HO

OH

TBSO

O

1018

H2O, rt, 99% 3) TBSOTf, Et3N, CH2Cl2, 0°C, 100%

2) Dess-Martin ox., NaHCO3, CH2Cl2, rt, 28% (2 steps)

1)

1014

O

O

979

OH, H+

2) LAH, THF, rt, 93%

1) HO

O

O

O

O 890

O

1012

O

O

OH

N-Mes

PhCH3, rfl, 80%

d-benzene, rfl, 52%

Ph Cl Ru Cl PCy3

Mes-N

1015

TBSO

O

CH3I, THF, rt, 91% 3) TBAF, THF, rt, 100%

1) TBSCl, imidazole, CH2Cl2, rt, 94% 2) NaH, HMPA,

Scheme 144 Synthesis of kempane derivative 1019

O

O

O

TBSO

O

1019

TBSO

O

O

O

1016

O

O

2) CH2=N(CH3)2+ I-, Et3N, CH2Cl2, rt, 60% (2 steps)

1) Dess-Martin ox., NaHCO3, CH2Cl2, rt

O

CH3OH, 0°C, 80%

NaBH4, CeCl3•7H2O,

1013

O

O

O

TBSO

O

1017

OH

O

2) NaH, CS2, THF, rt; CH3I, rt, 77%

1) NaBH4, CeCl3•7H2O, EtOH, 0°C, 91%

Chemistry of the Secondary Metabolites of Termites 307

308

E. Gössinger

O HO

O

1) BH3•S(CH3)2, THF, 0°C, 84%

O O

O

O

O

, THF,

O O

O

2) PCC, CH2Cl2, 81%

1021

HFP, rt, 73%

1022

N H

Ph

2.5:1

1023 O

Ph

O O EtO P EtO

O

O

O

O

OEt ,

O

O

, pyrrolidine, O

LDA, THF, –50°C, 69%, HPLC separation

O

CH3OH, 0°C, 85% O 1024

PhCH3, rfl, 71%

O H

O

H

O

1025

O

O

1) DIBAH, THF, –78°C, 79%

O O

2) Dess-Martin ox., CH2Cl2, 90% 1026

TiCl 2(i-PrO) 2, CH2Cl2, 0°C, 82%

Cl

H

1027

HO

O O

O 1020

O

Cl

OH

Scheme 145 Synthesis of kempane derivative 1020

attention to the EPC synthesis of the tetracyclic skeleton of the kempanes [768] (Scheme 145). Commercially available (R)-3-methylglutarate monomethyl ester (1021) was reduced chemoselectively with the borane dimethyl sulfide adduct to the ester alcohol, which in turn was oxidized by pyridinium chlorochromate to the ester aldehyde 1022. Chain elongation was achieved by Michael addition. To improve the yield of the desired diastereomer, the addition of methyl vinyl ketone was supported by the organocatalyst (R)-2-diphenylmethylpyrrolidine in hexafluoroisopropan-2-ol (HFIP). The 2.5:1 mixture of diastereomeric aldehydes 1023 was transformed to the dienones by Horner-Emmons reaction with triethyl-4-phosphonocrotonate. After separation of the diastereomeric mixture by HPLC, the diester 1024 reacted with cyclopentadiene and pyrrolidine to the fulvene derivative 1025 in good yield. Heating led to the tricyclic diester 1026 by endo-selective IMDA reaction in 71% yield. To close the fourth ring, the diester 1026 was regioselectively reduced. Due to the position of one ester at the concave face of the tricycle, diisobutylaluminum hydride reduced exclusively the ester at the side chain. The resulting alcohol was reoxidized to the aldehyde

Chemistry of the Secondary Metabolites of Termites

309

1027 by Dess-Martin periodinane. Cyclization was achieved by Prins reaction catalyzed by dichloro titanium diisopropanolate. Although this is a short and elegant elaboration of the skeleton of the kempanes, the introduction of the missing angular methyl group at position C-4 (kempane numbering) of tetracycle 1020 is not an easy task, as Burnell’s studies demonstrated. Syntheses of (1R*,3R*,4S*,11R*,12S*,15R*,16S*)-3-Hydroxykemp-7-en-6-one (1028) and (1R*,4S*,11R*,12S*,15R*,16S*)-Kemp-7-en-3,6-dione (1029) by Deslongchamps et al. Deslongchamps’ thorough investigation of transannular Diels-Alder reactions (TADA) as means of constructing complex polycyclic natural products [642] led him and his co-workers to try their hand at the especially difficult task of synthesizing the tetracyclic kempanes [769]. Their argument was that DielsAlder reactions with substituted (E,Z)-dienes usually react poorly or not at all, whereas in its transannular mode, high selectivity and good yields are to be expected. Thus, this key step was assigned to afford stereoselectively the tricyclic subunit with trans-syn-cis geometry at the ring junctions of the 5-6-6-membered ring system. Intramolecular aldol reaction was chosen to complete the tetracyclic kempane skeleton by closing the seven-membered ring. This last cyclization was planned although Paquette et al., who had used it in their synthesis of kempenes, were unable to deconjugate the resulting conjugated enone [761]. To construct the 13-membered ring necessary for the TADA reaction, a slightly convergent but lengthy pathway was chosen (Scheme 146). 4-Methyltetrahydro-2H-pyran-2-one (1030) [808] reacted stereoselectively with allyl bromide under basic conditions to the trans-disubstituted tetrahydropyranone 1031. This lactone was opened by O-methyl-hydroxylamine catalyzed by trimethylalane. The primary alcohol of the resulting Weinreb amide was exchanged by chloride using triphenylphosphine and N-chlorosuccinimide. Reduction of the hydroxamic acid methyl ester 1032 by diisobutylaluminum hydride furnished aldehyde 1033, which was converted into the terminal alkyne using the Corey-Fuchs protocol [809]. Carboxylation of the alkyne was achieved by methyl chloroformate after deprotonation by butyllithium. 1,4-Addition of dimethylcopperlithium to the alkyne ester 1034 led selectively to the conjugated unsaturated ester with (Z )configuration, which was reduced to the allyl alcohol 1035. After protection of the hydroxy group as t-butyldimethylsilyl ether, regioselective hydroboration with consecutive oxidation yielded the primary alcohol 1036, which was protected as triisopropylsilyl ether prior to exchange of the chloride by cyanide catalyzed by a crown ether. Cyanide 1037 was reduced to the aldehyde using diisobutylaluminum hydride followed by Wittig condensation with methylenetriphenylphosphanylylid. Selective deprotection of the allyl silyl ether 1038 was attained by pyridinium tosylate in ethanol, and the resulting alcohol was transformed into the bromide 1039 via the mesylate. Subunit 1039 was thus prepared in 16 steps and 44% overall yield. Chain elongation of the allyl bromide 1039 was achieved by exchange of bromide by malonic ester 1040, which was synthesized according to Larock starting with propynol in three steps [810]. To prepare for the ring-closing metathesis, the Stille reaction was used to introduce a further vinyl group to the disubstituted

O

97%

1043

OH

Cl

1040

CO2CH3

1036

1044

N O

O

H3CO2C

I

OH

1042

CH3O2C

OTPS

H3CO2C

CO2CH3

2) Ph3PCH3+I–, BuLi, THF, rt, 90%

1) DIBAH, PhCH3, –78°C, 94%

2) BuLi, ClCO2CH3, THF, –78°C, 95%

1) Dess-Martin ox., CH2Cl2, 78%

CN

Cl

1) CBr4, PPh3, CH2Cl2, 0°C, 100%

2) Ph3PCH3+I–, BuLi, THF, 85%

1037

DMF, 50°C, 100%

CO2CH3

CO2CH3 CO2CH3

OTPS

OTBS

1033

O

C2H3SnBu3, PPh3, Pd2(dba)3,

2) KCN, 18-crown-6, CH3CN, rt, 98%

1041

OH

–78°C, 95%

DIBAH, THF,

1) TPSCl, (CH3)3N, DMAP, CH2Cl2, rt, 100%

Cl

OTPS

Cl

1032

OTBS

CO2CH3

CO2CH3

NaH, THF, DMF 92%

I

2) 9-BBN, THF; NaOH, H2O2, THF, 0°C, 94%

1) TBSCl, imidazole, CH2Cl2, rt, 85% OH

2) PPh3, NCS, THF, 0°C to rt, 92%

1) CH3NHOCH3•HCl, (CH3)3Al, CH2Cl2, 0°C to rt, 95%

Et3N, PhCH3, 180°C, 93% H3CO2C

1039

Br

1035

OH

O

1031

O

Scheme 146 Synthesis of kempane derivative 1050 via transannular Diels-Alder reaction, part 1

H3CO2C

COOCH3

2) MsCl, Et3N, LiBr, –45°C to 0°C, 99%

1) PPTS, EtOH, rt, 94%

2) LAH, THF, 0°C, 95%

OTPS

Br ,

LDA, THF, –78°C;

1) CH3Li, CuI, THF, –50°C, 98%

1030

O

1038

1045

CO2CH3

2) RCMT, toluene, rfl, 74%

1) TsOH, CH3OH, 93%

OTPS

OCH3 Cl

OTBS

1034

O

310 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

311

malonic ester 1041. After deprotection of the silyl ether of the resulting tetraene 1042, cyclization to the 13-membered cyclic triene 1043 was accomplished by metathesis. This cyclization of a medium-sized ring in good yield (74%) demonstrates the power and high versatility of metathesis with later-generation Grubbs’ catalysts, in this case the Hoveyda variant [496]. Heating this monocyclic (E,Z,Z)triene 1043 to 180 C in toluene and triethylamine as base afforded in 93% yield exclusively the tricycle 1044 with trans-syn-cis geometry. The next steps were devoted to the elongation of the side chain and functional group transformations. Thus, the primary alcohol of 1044 was oxidized by Dess-Martin periodinane to the aldehyde and converted into the terminal double bond by Wittig condensation. Decarboxylation of malonic ester 1045 with cyanide at higher temperatures led to the monoester, which was converted into the hydroxamic ester 1046 with N,Odimethyl hydroxylamine (Scheme 147). Addition of methyllithium transformed the hydroxamic ester into the acetyl group of compound 1047, which under BaeyerVilliger reaction conditions not only converted the acetyl group into the acetate but also epoxidized the two double bonds yielding diepoxide 1048. Treatment of 1048 with lithium aluminum hydride removed the acetyl group and cleaved selectively the terminal epoxide. Oxidation with Dess-Martin periodinane transformed the two newly generated secondary alcohols into the diketone 1049 necessary for the intramolecular aldol reaction with potassium carbonate as base. As in Paquette’s synthesis, the seven-membered ring was obtained in good yield forming the kempane skeleton 1050 containing the undesired conjugated enone. The epoxide of compound 1050 was inert to a variety of reductive reagents due to the two angular

CO2CH3

O 1) NaCN, DMF, H2O, 120°C, 85%

H3CO2C

O

CH3Li, THF, 0°C, 89%

N

2) CH3NHOCH3•HCl, BuLi, THF, 0°C, 65% 1045

1046

O

O

O O

MCPBA, NaHCO3, CH2Cl2,

O

O 1048

1047

O 1) LAH, ether

K2CO3, CH3OH, O rfl, 70%

O

2) Dess-Martin ox., NaHCO3, CH2Cl2, 35% (3 steps)

O

O

1049

1050

Scheme 147 Synthesis of kempane derivative 1050 via transannular Diels-Alder reaction, part 2

312

E. Gössinger

methyl groups on the convex face and the inaccessibility of the epoxide by Lewis acids due to its position on the concave face. Thus, the synthesis plan was changed allowing reductive cleavage of the epoxide on the tricyclic precursor. The tricyclic malonic ester 1044 was decarboxylated using cyanide at higher temperature resulting in two epimeric monoesters, which were epoxidized by mchloroperbenzoic acid followed by the protection of the primary alcohol as the triphenylmethyl ether (Scheme 148). After separation of the epimeric esters 1051 and 1052, 1051 was reduced with superhydride yielding the diol 1053. The relative configuration of the carbinol groups and the regioselectivity of the reductive cleavage of the epoxide could be determined by oxidation of 1053 with oxygen catalyzed by rhodium(II). Under these conditions, the primary alcohol was oxidized to the aldehyde, which cyclized to the lactol and consequently was oxidized to the lactone 1054. The structure of 1054 was determined by X-ray structure analysis. Monoester 1052 with the ester group positioned at the convex face of the tricycle was reduced by superhydride to the primary alcohol and consequently converted into the phenyl selenide by phenylselenyl cyanide and tributylphosphine in pyridine. Protection of the secondary alcohol, derived by reduction of the epoxide, as silyl ether 1055 was followed by oxidation of the selenide to the selenoxide and immediate thermal elimination yielding the tricyclic compound with the exocyclic methylene group 1056. The more difficult accessibility of the primary alcohol of 1053, positioned at the concave side of the tricycle, enforced a small detour to attain 1056. Esterification of the carbinol was followed by protection of the secondary alcohol as a silyl ether. The ester was reductively cleaved, and the carbinol was converted into the methylene group as described above by the modified Grieco protocol [789, 811]. The methylene group of 1056 was transformed to the keto group by ozonolysis that removed simultaneously the trityl group (Scheme 149). The liberated primary alcohol of 1058 was oxidized to the aldehyde, which was converted into the methyl ketone 1059 via addition of methyl Grignard reagent and oxidation of the generated secondary alcohol. Contrary to the above-described aldol reaction to tetracycle 1050, aldol reaction with 1059 under a variety of basic conditions failed. Removal of the silyl group prior to the aldol reaction led in low yield to the desired tetracycle 1028. Oxidation of the secondary alcohol of 1060 changed the conformation favorably resulting in cyclization of the diketone 1061 by aldol reaction to the kempenedione 1029 in 64% yield. No efforts were described to convert this kempane into a naturally occurring product of the termite defense secretion.

O

1054

1044

OTr

1) (Ph3P)2RhCl2, O2, PhH

HO OH

OTr

1053

OTr

1.4:1

+

H3CO2C

2) TIPSOTf, lutidine, CH2Cl2, rt, 97%

1) PivCl, pyr, rt, 92%

superhydride, THF, rfl, 75%

1051

rt, 76% 3) TrCl, Et3N, DMAP, CH2Cl2, rt, 88%

2) DMAP, CH2Cl2, rt , 88% (2 steps)

OH

O

CO2CH3

1) NaCN, DMF, H2O, 120°C, 81% 2) MCPBA, CH2Cl2,

OTr

1057

OTIPS

OPiv

OTr

Ar = o-nitrophenyl

1052

O

Scheme 148 Synthesis of kempane derivatives 1028 and 1029 via transannular Diels-Alder reaction, part 1

O

H3CO2C

CO2CH3

ArSe

1055

OTIPS

pyr, THF, rt 3) MCPBA, pyr, CH2Cl2, rt, 83% (2 steps)

1) LAH, THF, rt, 100% 2) ArSeCN, Bu3P,

1056

OTIPS

MCPBA, pyr, CH2Cl2, rt, 63% (3 steps)

2) ArSeCN, Bu3P, pyr, THF, rt 3) TIPSOTf, pyr, CH2Cl2, rt

1) superhydride, THF, rfl, 92%

OTr

OTr

Chemistry of the Secondary Metabolites of Termites 313

314

E. Gössinger

OTIPS O3, CH3OH, CH2Cl2, –78°C;

1056

(CH3)2S, rt, 85%

OTr

OH

O

O

1) Dess-Martin ox., CH2Cl2, rt, 80% 2) CH3MgBr, THF, rt, 89%

OTIPS

3) Dess-Martin ox., CH2Cl2, rt, 90%

OH

1058

KOH, CH3OH, rt, 21%

TBAF, THF,

O

OTIPS

O

1059

O OH

OH O

rt, 68% O

1060

1028

Dess-Martin ox., CH2Cl2, rt, 56%

O O

KOH, CH3OH, rt, 64%

O

O

O 1029

1061

Scheme 149 Synthesis of kempane derivatives 1028 and 1029 via transannular Diels-Alder reaction, part 2

Ripperten-3-ol When the diterpenes of the defense secretions of Nasutitermes rippertii and N. ephratae were examined, several trinervitanes were isolated along with a tetracyclic diterpene. Due to the spectrometric data, the authors assumed a kempen-3-ol with a tetrasubstituted double bond [300]. To confirm the proposed structure (Scheme 150), they attempted to produce a crystallizable derivative. Acetylation led to a crystalline acetate 1062, but no useful single crystal could be produced. Therefore, the authors tried to epoxidize this compound. Despite the two neighboring methyl groups, a surprisingly facile reaction with m-chloroperbenzoic acid yielded a single epoxide 1063. Its X-ray analysis revealed a new structure type, the rippertanes. So far, ripperten-3-ol (¼ (1S,3S,4S,7S,8R,11S,12S)-rippert-15-en-3ol) (727), which later was isolated from several genera of the subfamily Nasutitermitinae, is the only compound with this skeleton. OAc OH

Ac2O, C6H14,

OAc

rt, 727

O MCPBA, buffer, pH = 7, CH2Cl2, rt

1062

Scheme 150 Derivatization of rippertenol (727)

1063

Chemistry of the Secondary Metabolites of Termites

315

Total Synthesis of ()-Rippertenol (727) by Snyder et al. Around the same time as Metz et al. succeeded in ending the 20-year period of unsuccessful synthesis approaches toward the kempanes, Snyder et al. presented the first total synthesis of the structurally related rippertenol (727) [758] (Scheme 151). To construct the 5-6-6membered tricyclic dienone, they started with the protected cyclohexenonol 1066 and annulated the two additional rings by successive intramolecular aldol reactions. The following implementation of the seven-membered ring consisted of an intermolecular, inverse Diels-Alder reaction, which enabled the stereoselective introduction of three stereogenic centers in the correct relative configuration to the three existing in the tricyclic starting material. Ring expansion with silyldiazomethane led to the rippertene skeleton, and functional group transformation completed the rippertenol synthesis. Thus, these researchers had been able to use the single stereogenic center of the cyclohexenonol to introduce stereoselectively the further six stereocenters in the correct relative configuration. Although Snyder’s synthesis is a racemate targeted one, their synthesis plan could be easily transformed into an EPC synthesis because their starting material has been separated into its enantiomers [812]. Snyder et al. varied Piers’ protocol of synthesizing the substituted hydrindenone 1069 [812] (Scheme 151). Birch reduction of 3-methylanisole (1064) led to the dienyl methyl ether, which was hydrolyzed under mild acidic conditions to the unconjugated 3-methylcyclohex-3-enone. Treatment by m-chloroperoxybenzoic acid yielded epoxide 1065, which was cleaved using acetic anhydride and base. The resulting acetate was saponified to attach a more stable protecting group. The t-butyldiphenylsilyl ether 1066 was chosen for this purpose. Learning from the difficulties Metz et al. had when trying to introduce the angular methyl groups at a late stage, Snyder et al. decided to attach the methyl groups at an early stage of the synthesis. Thus by α-alkylation, the methyl group was attached using strong base and methyl iodide. Next, the authors returned to Piers’ protocol to introduce stereoselectively the three-membered side chain via 1,4-addition of the copper organic compound 1067 derived from 2-(bromoethyl)-1,3-dioxane obtaining selectively the tetrasubstituted cyclohexanone 1068. Remarkably, Snyder et al. found that the nature of the protecting group at the side chain is quite important. Switching to the corresponding dioxolane (slightly more basic due to ring strain) led to a mixture of stereoisomers and reduced yields. Again, the predictability of the stereochemistry of the cyclohexane was used to introduce the second side chain stereoselectively. This was achieved utilizing 1-chlorobutan-3-one under acidic conditions in refluxing benzene. Under these conditions not only the side chain was attached but also the 1,3-dioxane was cleaved starting the intramolecular aldol reaction resulting in the desired hydrindenone derivative 1069. Whereas the five-membered ring was formed, more drastic conditions had to be used to effect the second aldol reaction. Here, strong basic conditions led to the aldol, which was dehydrated via mesylation under basic conditions. With the tricyclic conjugated dienone 1070, the stage was set for the decisive inverse Diels-Alder reaction. Although Lewis acid catalysis in inverse Diels-Alder reactions was only to some extent examined, the researchers decided to

1073

O

O

1065

O

O

O

TPSO

OTPS 1070

CH2Cl2, –78 to –50°C, 21%, (71% brsm)

BF3•OEt2, TMSCHN2,

2) MsCl, Et3N, CH2Cl2, 0°C, 31% (3 steps)

1) t-BuOK, THF, t-BuOH rt

H2O, rt, 83% (2 steps) 3) MCPBA, CH2Cl2, 0°C

1) Li, NH3, Et2O, t-BuOH, –33°C 2) (CO2H)2, CH3OH,

Scheme 151 Synthesis of racemic rippertenol (727)

TPSO

1069

OTPS

O

1064

O

1074

O

BF3•OEt2, CH2Cl2, –78°C, 49%

O

O

2) Na2CO3, CH3OH, rt 3) TPSCl, imidazole, DMF, rt, 74% (2 steps)

1) Ac2O, i-PrNEt 2, DMAP, CH2Cl2, rt, 81% (2 steps)

DMAP, THF, 70°C 3) Bu3SnH, AIBN, PhCH3, 100°C, 46% (2 steps)

2) O

O MgBr

1) LDA, CH3I, –78°Cto rt, 88%

TPSO

O

O

1075

2) aq HCl, THF, rt, 99% TPSO

1) Zn, CH2Br2, TiCl4, THF, rt, 48%

1067 CuBr•S(CH3)2, THF, –78 to –50°C, 86%

O

1071

O

1) NaBH4, CH3OH, THF, 0°C to rt, 93% 2) N N N N S

TPSO

1066

OTPS

O O

TBAF, THF, 45°C, 93%

1072

O

OTPS 1068

O

O

HO

727

(Ph3P)3RhCl, H2, PhH, rt, 98%

TsOH, PhH, rfl

Cl

316 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

317

use boron trifluoride as Lewis acid and the relatively stable ketene acetal, 2-ethylidene-1,3-dioxane [813]. Under these conditions, the reaction occurred at 78 C and thus with very high face selectivity and mainly exo addition (exo:endo ¼ 2.6:1). It should be noted that no reaction occurred without the catalyst, which raises the question if indeed a Diels-Alder reaction had occurred or if in two ionic reactions (vinylogous Michael reaction and Mukaiyama aldol), the ring was closed. The main adduct 1071 could be crystallized, and thus the structure was established by X-ray analysis. Then, conditions had to be found to convert the keto group of the pentacyclic adduct 1071 stereoselectively into the methyl group via methylenation and subsequent hydrogenation, without cleavage of the peri-positioned cyclic ketal or isomerization of the double bond. Wittig methylenation failed due to the E1cb reaction. The modified Lombardo procedure [620, 621] with methylene bromide, zinc, and titanium tetrachloride in tetrahydrofuran at room temperature proved mild enough leading in 48% yield to the methylene group. Subsequently, aqueous hydrochloric acid at room temperature hydrolyzed the dioxane without shift of the double bonds yielding tetracycle 1072. Whereas heterogeneously catalyzed hydrogenation of the methylene group led to a mixture of epimers, the homogenous Wilkinson’s catalyst led exclusively to the expected attack of hydrogen at the convex face of the molecule. Ketone 1073 was used to expand the cyclohexenone moiety of the tetracycle, which proved the most difficult step in this synthesis. Finally, the authors had to be content with a 21% yield using (trimethylsilyl)diazomethane and boron trifluoride etherate at low temperature. Fortunately, the starting material could be recovered. With this reaction, the correct skeleton 1074 of rippertene was reached, and functional group transformations completed the synthesis. The reduction of the keto group of the cycloheptenone 1074 to the methylene group in 1075 was achieved by sodium borohydride reduction, esterification to the thiocarbamate, and radical reduction with stannane. Cleavage of silyl ether 1075 with fluoride completed the synthesis of rippertenol 727 in 20 steps and 0.22% overall yield. Three synthesis attempts toward rippertenol by Metz et al. led to 1-desmethyl3α-hydroxyrippert-15-ene-3,5-butanolide (1076) [814], (3S,6R,7S,20 S)-3(50 -methylhex-50 -en-20 -yl)-6,10-dimethylbicyclo[5.3.0]dec-10(1)-en-2-one, (3R,6R,7S,20 S)-3-(50 -methylhex-50 -en-20 -yl)-6,10-dimethylbicyclo[5.3.0]dec-10(1)en-2-one (1088) [815], and 4-desmethyl-3α-hydroxy-15-rippertene (1103) [816]. Partial EPC Synthesis of 1-Desmethyl-3α-hydroxyrippert-15-ene-3,5butanolide by Metz et al. In 1993, Metz, Bertels, and Fröhlich began their synthesis efforts toward the rippertanes with an interesting partial EPC synthesis starting with commercially available ()-α-santonin (551). The key steps were the photochemical rearrangement of the unsaturated decalindienone to the hydroazulenone, intramolecular vinylogous aldol reaction, and intramolecular Diels-Alder reaction [814] (Scheme 152). ()-α-Santonin (551) was converted into ()-6-epi-α-santonin by acid treatment and further to ()-6-epi-β-santonin (1077) by strong base epimerizing the methyl group at the butanolide subunit. The following photoisomerization, a reaction well known from steroid chemistry but also described for ()-α-santonin (551) [817], led to the hydroazulene derivative 1078 in 33% yield. After elimination of the acetate

1) H2/Pd/C, C2H5OH,

551

O

O

O

O

1085

1081

6:1

H3CO

O

2) t-BuOK, PhCH3, 20°C, 69%

O O

O

Bu4NI, KOH, rt, 94%

2) HC CCH2Br,

1) LAH, Et2O, –78°C to rt, 99% O

O

1086

1082

O

17°C, 33%

CH2Cl2, 20°C, 90% 3) Ph3P=CHCO2Me, toluene, rfl, 76% H3CO

1) LAH, ether, 20°C, 99% 2) TPAP, NMO,

1077

O

hν, HOAc,

Scheme 152 Synthesis of 1-desmethylrippertenol derivative 1076

3) RhCl3•3H2O, EtOH, rfl, 68% (3 steps)

1) MsCl, Et3N, CH2Cl2, 0°C 2) LiBr, DMF, rfl

2 N NaOH, 20°C 2) CH2N2, CH3OH 0°C, 72% (2 steps)

O

1) aq HCl, DMF, 100°C, 77% 0°C, 94%

H2SO4 conc,

O

20°C, 98% 3) TPAP, NMO, CH2Cl2, 20°C, 83%

1083

O

1087

1) H2, Pd/C, AcOEt, 20°C, 99% O 2) LAH, Et2O,

O

t-BuOK, t-BuOH, rfl

1078

O

OAc

O

O

rfl, 69%

2% KOH, CH3OH,

2N HCl, 60°C, 99%

2) TPAP, NMO, CH2Cl2, rt, 37% (3 steps)

1) TsOH, H2O, THF, t-BuOH, 20°C

1079

O

O

O

O

CrCl2, HOAc, O

1084

1076

HO

1080

HO O

318 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

319

using concentrated sulfuric acid, partial hydrogenation of the conjugated dienone with palladium on charcoal as catalyst occurred exclusively by attack from the convex face of the tricyclic lactone 1079. To circumvent the undesired face selectivity, the butanolide of 1079 was cleaved reductively according to Büchi et al. [818] by chromium(II) chloride prior to hydrogenation. Attack at the bicyclic dienone 1080 by hydrogen catalyzed by palladium under basic conditions resulted chemoand stereoselectively in the desired stereoisomeric, bicyclic enone, which was esterified by diazomethane to the bicyclic enone ester 1081. Side-chain elongation in preparation for the vinylogous aldol reaction was achieved within six steps in 55% yield. The carbonyl groups of 1081 were reduced by lithium aluminum hydride and reoxidized to the keto aldehyde with perruthenate. The following Wittig condensation led to the unsaturated ester 1082, which was hydrogenated, and the saturated ester was converted into the desired aldehyde by reduction and reoxidation as described above. The generated aldehyde 1083 was treated with potassium hydroxide in methanol under reflux furnishing the tricyclic alcohol 1084 in 69% yield. Dehydration was achieved by mesylation and successive treatment with lithium bromide in dimethylformamide under reflux. Under these conditions, also the hydrogen at the ring junction was inverted. The resulting mixture of conjugated and deconjugated dienones was treated with rhodium chloride thereby improving the yield of the conjugated dienone 1085. This set the stage for the last key step, the IMDA reaction. Lithium aluminum hydride attacked ketone 1085 at 78 C exclusively from the convex face that was established by X-ray analysis. Williamson synthesis of the generated alcohol with propargyl bromide furnished the propargyl ether 1086. To improve reactivity and stereoselectivity of the planned IMDA reaction, the propargyl ether was isomerized to the allenyl ether by base treatment according to Kanematsu [819], which initiated immediate cyclization to the pentacyclic dihydrofuran 1087. The dihydrofuran was transformed into the γ-lactone by addition of water under acidic conditions and subsequent oxidation [819]. The pentacyclic lactone 1076 was based on the tetracyclic rippertene skeleton with the correct configuration at all stereogenic centers. The flaw of this EPC synthesis, however, was the missing angular methyl group at position C-1 (kempane numbering). EPC Synthesis of (3S,6R,7S,20 S)-3-(50 -Methylhex-50 -en-20 -yl)-6,10dimethylbicyclo[5.3.0]dec-10(1)-en-2-one and (3R,6R,7S,20 S)-3-(50 -Methylhex50 -en-20 -yl)-6,10-dimethylbicyclo[5.3.0]dec-10(1)-en-2-one by Kreutzer and Metz With the information gathered in the partial synthesis of (1)-desmethyl3α-hydroxy-15-rippertene derivative 1076, Kreutzer and Metz [815] decided to initiate an enantiopure total synthesis of 3α-hydroxy-15-rippertene (727). The intention of this approach was to start with ()-isopulegol, followed by ring expansion to the cycloheptene part of rippertene, attachment of side chains, and intramolecular aldol reaction resulting in the octahydroazulene subunit. The isopropenyl group of the isopulegol was considered to serve for building the five-membered side chain with a terminal double bond. This terminal double bond and a vinyl group at the cyclopentene were the prerequisites for the planned tandem Heck reaction, thought

320

E. Gössinger

to annulate the two cyclohexanes thus completing the construction of the rippertene skeleton. In fact, the researchers succeeded in the construction of the octahydroazulene subunit but abandoned their elegant plan involving the tandem Heck reaction. The first steps of the synthesis of the octahydroazulene subunit were devoted to the transformation of the isopropenyl group following Knochel’s protocol [820] (Scheme 153). The hydroxy group of ()-isopulegol (1090) was protected as a silyl ether prior to the stereoselective hydroboration with diethyl borane. The boron of the resulting borane 1091 was exchanged for zinc by diethyl zinc, which consequently was transmetalated by the cuprous cyanide-lithium bromide complex. Coupling of the copper organic compound with metallyl bromide furnished the desired five-membered side chain with the terminal double bond of 1092. Ring expansion was achieved starting with deprotection and subsequent oxidation of the secondary alcohol resulting in ketone 1093. Ring expansion via methylene insertion was accomplished with (trimethylsilyl)diazomethane catalyzed by Lewis acid [821]. The regioselectivity of this reaction proved highly dependent on the type of Lewis acid. Whereas trimethylalane led in equal parts to insertion in the bond from the carbonyl to the tertiary carbon atom and thus to cycloheptanone 1094 and in the bond to the secondary carbon atom to give cycloheptanone 1095, boron trifluoride etherate yielded a 1:5 mixture of 1094:1095. The desired cycloheptanone 1095 was dehydrogenated according to a modified protocol of Saegusa via the silyl enol ether and oxidation with oxygen and palladium acetate as catalyst [822, 823]. With cycloheptenone 1096, the stage was set for the annulation of the cyclopentene. Treatment of 1096 with 3-(tbutyldimethylsilyloxy)propylmagnesium bromide (1097) and cuprous iodide resulted in stereoselective addition and was followed by cleavage of the silyl ether and oxidation of the resulting primary alcohol with Dess-Martin periodinane. The usual methods to effect aldol reaction failed with aldehyde 1098. When potassium hydroxide and crown ether in benzene under reflux were used, cyclization by intramolecular aldol reaction was successful but was accompanied by partial epimerization of the side chain in α-position to the keto group. Yields of the bicyclic enone 1099 were improved by separation and epimerization of the undesired bicyclic enone 1100 under identical reaction conditions. To incorporate the necessary angular methyl group of rippertene, the researchers repeated the reaction sequence exchanging the Grignard reagent 1097 by Grignard reagent 1101. Converting the magnesium organic compound 1101 into the corresponding copper organic compound permitted 1,4-addition to cycloheptenone 1096. The following deprotection of the keto group proved difficult. Acid treatment failed, but transacetalization to acetone catalyzed by complexed palladium chloride liberated the ketone 1102 [824]. For the aldol reaction with the two ketones, potassium hydroxide in methanol sufficed to produce the octahydroazulenes (3S,6R,7S,20 S)-3(50 -methylhex-50 -en-20 -yl)-6,10-dimethylbicyclo[5.3.0]dec-10(1)-en-2-one (1088) and (3R,6R,7S,20 S)-3-(50 -methylhex-50 -en-20 -yl)-6,10-dimethylbicyclo[5.3.0]dec-10 (1)-en-2-one (1089) in high yield. Separation of the epimers proved facile, and the undesired epimer 1089 was recycled.

OH

O

O

O

MgBr

1101

to rt, 92%, 85% de 2) PdCl2(PhCN)2, acetone, rt, 73%

CuI, THF, –20°C

1)

1098

O

O

LA: (CH3)3Al, –78°C to rt or BF3•OEt2, –78°C

2) aq HCl, THF, rt

1) LA, CH2Cl2, TMSCHN2

2) Et2BH, Et2O, 0°C to rt

1102

O

KOH, 18-crown-6, PhH, 80°C, 55%

1094 45% 15%

O

1091

OTMS

BEt2

Scheme 153 EPC synthesis of the octahydroazulen-8-one derivative 1088

1096

O

2) TBAF, THF, rt, 95% 3) Dess-Martin ox., CH2Cl2, rt, 86%

1097 MgBr, CuI, THF, –20°C to rt, 86%, 93% de

1) TBSO

1093

O

1090 ((–)-isopulegol)

1) TMSCl, imidazole, DMAP, DMF, rt, 99%

1099

O

+

1.7:1

1088

O

1100

1.2 : 1 KOH, CH3OH, 65°C, 91% (1.2:1)

+

O

2) Pd(OAc)2, O2, DMSO, rt, 86% (2 steps)

1) LDA, THF, –78°C; TMSCl, –78°C to rt

1092

OTMS

KOH, 18-crown-6, PhH, 80°C, 80% (1.3:1)

1095 40% 75%

KOH, CH3OH, 65°C, 94%

+

O

2) Br , THF, –78°C to rt, 79%, >98% de (3 steps)

1) Et2Zn, 0°C; CuCN•2LiBr, THF, –78 to 0°C

O

1096

O

1089

2) PCC, CH2Cl2, rt, 99% (2 steps)

1) aq HCl, THF, rt

Chemistry of the Secondary Metabolites of Termites 321

322

E. Gössinger

EPC Synthesis of 4-Desmethyl-3α-hydroxy-15-rippertene by Metz and Hennig In this synthesis, Metz and Hennig [816] combined the experiences gathered in previous synthesis studies [814, 815]. The synthesis started with ()isopulegol (1090) and followed the synthesis pathway described above to the isomeric cycloheptanones 1094 and 1095 [815]. The side chain attached at the cycloheptanones was used to close the cyclohexene by intramolecular aldol reaction, which circumvented the difficulty of introducing the angular methyl group at C-1 of the rippertene at a late stage. As in the above synthesis approaches, the annulation of the cyclopentene was achieved by side-chain attachment and subsequent intramolecular aldol reaction. Analogous to the partial synthesis, the second cyclohexane subunit was introduced by intramolecular Diels-Alder reaction [814] (Scheme 154). The mixture of cycloheptanones 1094 and 1095 was separated. The methylene group of 1094 was converted into the ketone by dihydroxylation with potassium osmate and immediate fragmentation by periodate resulting in diketone 1104. The isomeric cycloheptanone 1095 was converted into the same diketone 1104. The keto group transposition followed a known protocol. To prevent epimerization of the side chain, kinetic deprotonation with lithium hexamethyldisilazane at low temperature was used to attach benzaldehyde by aldol reaction. After dehydration to the enone via the mesylate and base treatment, the keto group was reduced by lithium aluminum hydride to alcohol 1105, which was esterified by ethyl chloroformate, and the allylic carbonate was removed by hydrogen transfer according to Tsuji [825], thus preventing the reduction of the double bonds. The double bonds of cycloheptane 1106 were cleaved oxidatively via dihydroxylation and immediate treatment of sodium periodate attaining the desired diketone 1104. Base-catalyzed intramolecular aldol reaction furnished the homooctalinone 1107. The subsequent annulation of the cyclopentene started with allylation in an α-position to the ketone using kinetic deprotonation followed by addition of allyl bromide. Modified Wacker reaction with benzoquinone as oxidant and palladium chloride as catalyst yielded the diketone 1108 [826]. The intramolecular aldol reaction was accomplished in 64% yield by base treatment supported by microwave. As described above [814], the keto group of tricyclic dienone 1109 was reduced to the alcohol and etherified with propargyl bromide. Basic isomerization of the alkyne 1110 to the allene and subsequent IMDA reaction to the tetracycle 1111 occurred in very good yield when supported by microwave at elevated temperature (Scheme 154). The dihydrofuran subunit of the pentacycle 1111 was hydrated under acidic conditions, and the resulting hemiacetal was oxidized by Ley’s reagent to the γ-lactone 1112 (Scheme 155). Next, Vedejs’ reagent (MoOPH, oxodiperoxymolybdenum-(pyridine)(hexamethylphosphoric triamide)) was used to introduce the hydroxy group in α-position to the carbonyl, the prerequisite of the removal of the now unnecessary carboxyl group. Subsequently, the hydroxylated lactone was saponified and reduced with lithium aluminum hydride. Triol 1113 was treated with sodium periodate, and the resulting keto alcohol was reduced stereoselectively due to neighboring group direction by a modified protocol of Evans [797, 827]. The newly formed alcohol of diol 1114 was regioselectively protected as a methoxymethyl ether due to its position at the convex

1095

40°C, 64%

O

1109

OH

81%

1105

O

Bu4NI, aq KOH, PhCH3, rt, 91% (2 steps)

1) LAH, ether, –78 to 0°C 2) HC≡CCH2Br,

t-BuOK, t-BuOH, THF, 65°C,

Ph

Scheme 154 EPC synthesis of 3-desmethylrippertenol derivative 1111

1108

1107

dimethylacetamide, H2O, 35°C, 85%

O

3) DBU, CH2Cl2, rt 4) LAH, Et2O, –78 to 0°C, 74% (3 steps)

t-BuOK, t-BuOH, THF, microwave,

Br , 0°C, 84% 2) PdCl2, O O,

1) LiHMDS, THF, 0°C;

O O

O

1) LiHMDS, THF, –78°C; PhCHO, –78°C, 95% 2) MsCl, Et3N, CH2Cl2, 0°C

O

1110

1104

O

H2O, 0°C to rt, 93%

Ph

150°C, 83%

t-BuOK, t-BuOH, THF, microwave,

H2O, 0°C to rt, 93%

K2OsO2(OH)4, NaIO4, pyr, THF,

OsO4, NaIO4, pyr, THF,

2) Pd2dba3, Ph3P, Et3N, HCO2H, THF, 75°C, 98%

1) BuLi, THF, –78°C; ClCO2Et, –78°C, 96%

O

O

1106

1111

1094

Chemistry of the Secondary Metabolites of Termites 323

324

E. Gössinger

1) TsOH, H2O, THF, t-BuOH, rt, 80%

O

1111

O

2) TPAP, NMO, MS 4 Å, CH2Cl2, rt, O 100%

1) NaIO4, THF, H2O, rt, 98% 2) (CH3)4NBH(OAc)3, HOAc, CH3CN, THF, –20°C, 68%

1) LiHMDS, THF, –78°C; MoOPH, –78°C, 85%

HO HO

HO

2) aq KOH, THF, rt 3) LAH, THF, rfl, HO HO 89% (2 steps) 1112

1113

1) MOMCl, i-Pr2NEt, Bu4NI, CH2Cl2, 0°C to rt, 57%

S

2) BuLi, THF, 0°C; CS2, 0°C; CH3I, 0°C

MOMO

1114

S O

1115

1) Bu3SnH, AIBN, PhCH3, rfl, 69% (2 steps) 2) aq HCl, THF, 50°C, 91%

HO 1103

Scheme 155 EPC synthesis of 3-desmethylrippertenol 1103

face of the tetracycle, thus permitting the removal of the remaining hydroxy group by esterification to the xanthate 1115 and radical reduction with stannane. The synthesis was completed by liberating the remaining alcohol under acidic conditions yielding 4-desmethyl-3α-hydroxy-15-rippertene 1103.

Longipenol Only one source is known for longipenol (¼ (1R*,3S*,4S*,7S*,8S*,11R*,12R*,16S*)15(17)-longipen-3-ol) (729), the frontal gland secretion of minor and major soldiers of the Southeast Asian, free-ranging termite Longipeditermes longipes [29, 304, 305, 704]. The elucidation of the complex structure of this spirocyclic compound was solved by spectrometric data, especially 2D NMR (COSY, NOESY), which also allowed a determination of its conformation [704] (Scheme 156). The total synthesis of longipenol represents an even more formidable challenge for the chemist compared to the other tetracyclic diterpenes of termites. Eight stereogenic centers, one of them a spiro center, the eight-membered ring fused to a hydrindane moiety, and too few functionalities complicate this task. Thus, it is not surprising that no synthesis of longipenol exists. Only one tentative approach toward longipenol was reported [828]. Mehta et al. constructed the tricyclic subunit of the

Chemistry of the Secondary Metabolites of Termites

325

Scheme 156 NOE interactions in longipenol (729)

H3C H

H3C H

H H H H

H

H H OH

H3C

H

H

H

longipane skeleton consisting of an eight-membered, a six-membered, and a fivemembered ring. They chose the triquinane as starting material, which was synthesized by a protocol the researchers had developed previously [829]. Fragmentation led to the 5,8-membered ring system [830], and intramolecular Mukaiyama aldol reaction with the side chain afforded the additional cyclohexane.

Synthesis of 12-Methoxytricyclo[7.2.2.04,11]trideca-6,10-dione by Mehta et al. The preparation of a tricyclic intermediate (1116) of the planned synthesis of longipenol (729) [828] is illustrated in Scheme 157. 1) (EtO)2POCH2CO2Et, NaH, THF, rt, 75% O

2) (CH2SH)2, TsOH, PhCH3, rfl, 80%

O

1) LAH, ether, 90% 2) PCC, MS 4Å,

Na, NH3, ether, 55%

CO2Et

CO2Et

1117

CH2Cl2, 70% 3) CH3OH, (CH3O)3CH, PPTS, 80%

S

S

1118

1119 O

O

RuO2, NaIO4, CCl4, CH3CN, H2O, 75%

+

O

O O

O

O

1120

1121

O

O

O

2) TiCl4, CH2Cl2, 35%, (2 steps)

O

+

O O

1122

1:4

1) LiHMDS, TMSCl, THF

O

O O O

1123

PPTS, 80% O

O

O

CH3OH, (CH3O)3CH,

1116

O 3:4

1124

Scheme 157 Synthesis of bicyclo[6.3.0]undecadiones and 12-methoxytricyclo[7.2.2.03,7]trideca2,8-dione (1116)

326

E. Gössinger

Horner-Emmons condensation was applied to introduce the side chain at triquinane 1117. Under the chosen reaction conditions, triethyl acetoxyphosphonate was added chemoselectively to the saturated ketone, yielding a mixture of the unsaturated (E)- and (Z)-configured ester. Subsequently, the remaining keto group was removed via the cyclic dithioketal 1118 and consecutive reduction by sodium in ammonia, which simultaneously reduced the electron-poor double bond of the unsaturated esters. The mixture of the saturated esters 1119 was transformed into the aldehydes by reduction to the alcohols with lithium aluminum hydride and subsequent reoxidation with pyridinium chlorochromate. The unstable aldehydes were protected as dimethyl acetals 1120 prior to oxidative fragmentation with ruthenium dioxide and sodium periodate [831]. At this point, the diastereomeric diketones 1121 and 1122 (1:4) were separated. The thermodynamically more favorable compound 1122 predominated. Although the minor compound 1121 would furnish the desired cis-hydrindanone subunit by aldol reaction, the researchers studied the aldol reaction with the major product 1122. To achieve this goal, selective protection of the less hindered ketone of 1122 as dimethyl ketal 1123 was followed by silyl enol formation that underwent Mukaiyama aldol reaction with titanium tetrachloride in dichloromethane yielding the tricyclic diketone 1116 along with larger amounts of the bicyclic compound 1124 generated by deprotection of the carbonyl groups. Due to the low yields of those first steps in their planned synthesis, the authors decided to abandon this project.

Rojofuran Four hundred soldier heads of a Venezuelan Nasutitermes species were extracted with n-hexane. The main compound, rojofuran (¼ (1R*,3R*,4S*,11R*,12S*,15R*,16S*)6,19-epoxykempa-6,8(19)-dien-3-yl acetate) (728), was stable as long as it remained in the crude extract. After flash chromatography, 10 mg of a strained and therefore unstable compound was isolated. The high-resolution mass spectrum provided the molecular mass and thus the molecular formula. 1H and 13C NMR spectra in deuterobenzene in a sealed tube revealed a pentacyclic diterpene containing one acetate moiety and 13C NMR signals resembling those of a trisubstituted furan ring of a sesquiterpene [832]. More information could be attained, when rojofuran was converted into its maleic acid anhydride adduct 1125 and from the two oxidation products obtained by exposing rojofuran to air (Scheme 158). The easy decomposition of trisubstituted furans by oxygen has been explored by Woodward et al. [833]. In one of the two decomposition products, the proton of the furan moiety was replaced by an aldehyde proton, which corresponded with an 1,4-addition of oxygen to the furan and disproportionation of the intermediate ozonide 1126 to keto aldehyde 1127. The second oxidation product was the unsaturated lactone 1128. The stereochemistry was tentatively given in accordance with the stereochemistry of the kempanes. No synthesis approach toward rojofuran has been published.

Chemistry of the Secondary Metabolites of Termites

327

O O O

O O

O

air AcO

AcO

AcO 728

1126

O

O

1127

O

O O O

O O

O

AcO

AcO 1125

1128

Scheme 158 Reactions of rojofuran (728)

4 Biosynthesis of Secondary Metabolites of Termites 4.1 4.1.1

Biosynthesis of Pheromones Acetogenins

Very little is known about the biosynthesis of the pheromones of termites. It is assumed that the biosyntheses of the trail and sex pheromones 1, 3, 4, 6, 7, 9, and 11 follow the fatty acid pathway (note that no evidence of the polyketide pathway has been found in insects [455, 834]). Hanus et al. assume that the unsaturated dodecan1-ols 1, 3, and 4 are biosynthesized by three cycles of β-oxidation of oleate or linoleate followed by reduction of the carboxyl moiety, giving rise to 4 and 3, respectively. A subsequent stereospecific desaturation at the single bond C-8–C-9 of 3 leads to (3Z,6Z,8E)-dodecatrien-1-ol (1) [145]. Tokoro et al. noted that the amount of the trail-following pheromone (3Z,6Z,8E)dodeca-3,6,8-trien-1-ol (1) of Reticulitermes speratus isolated from whole workers was smaller than expected according to the trail-following behavior with artificial trails scented with 1. They assumed that the pheromone is stored as wax esters [74, 76]. To confirm the assumption, the n-hexane extract of workers was chromatographed, and the inactive fractions were hydrolyzed with methanolic potassium hydroxide. Indeed, two of the thus-treated fractions showed bioactivity, and 1 could be isolated in larger quantities than was originally found as a free alcohol. Neither the acids esterified with 1 nor the side of storage of the wax esters

328

E. Gössinger

was determined. These investigators assume that wax esters are precursors of the pheromone. Recently, comparable wax esters of the trail-following pheromones 3 and 4 were isolated from the frontal gland of soldiers of inquilines (cohabitating termites), and around 50 fatty acids bound to the pheromones 3 and 4 were recognized [264]. Regarding the biosynthesis of the cuticular hydrocarbons (CHCs), the genome of the lower termite Zootermopsis nevadensis has been sequenced [474]. Comparison with the genomes of solitary insects revealed an expansion of the genes involved in the biosynthesis of the CHCs. Only the biosynthesis of the CHCs of Zootermopsis species was examined in detail [412, 413, 835] and compared with the biosyntheses of CHCs of several insect orders [404, 407]. These hydrocarbons are synthesized in oenocytes and are transported by the carrier protein lipophorin within the hemolymph to the cuticle. Despite their transport in the hemolymph, the composition of the CHCs differs from that of the hydrocarbons of the hemolymph in Zootermopsis [413]. As Scheme 159 shows, the long-chain saturated and unsaturated CHCs follow the fatty acid pathway until the whole chain length is reached. Next, the enzyme-bound fatty acid thioester is cleaved reductively to the aldehyde. At this point the biosynthesis of the hydrocarbons of insects deviates from the biosynthesis of hydrocarbons of plants, algae, and vertebrates. Whereas the latter decarbonylate the aldehyde without use of oxygen and cofactors, the long-chain aldehydes of insects decarboxylate assisted by a cytochrome P450 enzyme that requires NADPH and oxygen [836, 837]. In social insects, the methyl-substituted CHCs are important as part of the recognition of castes and colonies (nestmates). Here Zootermopsis uses a different pathway to methyl-substituted malonate than other insects (e.g., Blattella germanica, which belongs to the same order as termites) [455]. Instead of branched amino acids, Zootermopsis uses succinate as starting material, which is converted into methyl-substituted malonate by vitamin B12 as cofactor. The nitrogen-poor nutrition of termites may be responsible for this deviation, avoiding nitrogen loss. Blomquist has assumed that gut symbionts supply this precursor for the branched CHCs [835]. By analogy to other insects, early insertion of the methyl-substituted malonate has been assumed. Nothing is known about the stereochemistry of the biosynthesis of branched CHCs.

O

4

SCoA

Scheme 159 Biosynthesis of cuticular hydrocarbons (CHCs) of termites

(CH2)7

(CH2)7

(CH2)7

(CH2)7

OH

SCoA

CH3(CH2)23CH3

O

Cyt P450, NADPH, O2

NADPH, O2

desaturase

O

O

–CO2

Cyt P450, NADPH, O2

CH3(CH2)24CHO

reductase

SCoA

SCoA

O

OH

O

(CH2)7

SCoA

OH

SCoA

OH

O

O

CH3(CH2)24COSCoAc

O

8

SCoA

(CH2)7

O

O

(CH2)14

O

(CH2)15

O

(CH2)15

–CO2

O

H

SCoA

SCoA OH

O

SCoA

O SCoA

O OH SCoA

(CH2)7

O

reductase

O

4

Vit. B12

O

10 SCoA OH

O

O

O

H

SCoA

(CH2)19CH3

–CO2

(CH2)20

Cyt P450, NADPH, O2

reductase

(CH2)20

Chemistry of the Secondary Metabolites of Termites 329

330

4.1.2

E. Gössinger

Biosynthesis of Isoprenoids

Like all animals, insects biosynthesize isoprenoids by the mevalonate pathway [838]. However, it has been known since the late 1950s that insects are unable to synthesize polycyclic triterpenes and in consequence steroids [839], by lacking the enzyme squalene synthase, which catalyzes the complicated coupling of two farnesyl diphosphate units to squalene. Thus, insects have to sequester precursors of their molting hormones, the ecdysones, with their food. At least as important, insects are unable to synthesize cholesterol, which is the key regulator of the mevalonate pathway in vertebrates [838, 840]. Instead, insects have developed an alternative branch of the mevalonate pathway, which leads to the juvenile hormones (JH). This pathway has been investigated in detail in Drosophila and more relevant in cockroaches, the phylogenetically closest relatives of termites (Scheme 160). Farnesyl diphosphate (1129) is hydrolyzed, and the resulting primary alcohol 370 is oxidized via the aldehyde 1130 to the acid 1131. Methyl transferase then esterifies the acid to ester 1132, and enantio- and chemoselective epoxidation furnishes JH III (260), the only JH found in termites. Juvenile hormones are general developmental hormones not only involved in the process of molting, but they regulate embryonic development and induce the synthesis of the hemolymph protein vitellogenin as well as pheromone production. In eusocial insects JH influences caste formation and caste homeostasis [841]. Several researchers have suggested that JH III (260) is also a primer pheromone in termites [842]. In addition, JHs are at least to some extent involved in the regulation of the mevalonate pathway in insects [838]. The biosynthesis of the pheromones 5 and 12 of the basal termites and 2 and 8 of the Termitidae also follows the mevalonate pathway. Two of the sex and trail pheromones are open-chain isoprenoids. Neither the location of the enzymes of their biosynthesis nor the biosynthesis itself has been investigated. That is also the case for the cyclic diterpenes neocembrene (2) and trinervitatriene 8 found in the sternal and/or tergal glands. However, neocembrene (2) has also been identified in the frontal gland of some Termitidae, where its biosynthesis has been investigated. The alarm pheromones, with exception of benzoquinone, are also part of the defensive secretion of the frontal gland of Neoisoptera.

O O

O

juvenile hormone epoxidase

farnesyl dehydrogenase

Scheme 160 Biosynthesis of juvenile hormone III (JHIII, 260)

1132

1130

1129

OPP

farnesyl diphosphate pyrophosphatase

O

260

1131

370

O

O

O

OH

OH

juvenile hormone methyl transferase

farnesyl oxidase

Chemistry of the Secondary Metabolites of Termites 331

332

4.2

E. Gössinger

Biosynthesis of Components of the Frontal Gland Secretion

The frontal gland is, with few exceptions, restricted to termite soldiers of the Neoisoptera. The investigation of the biosynthesis of the secondary metabolites of termite soldiers is especially difficult because soldiers are unable to feed themselves but are fed by workers. Thus, the labeled precursors may be either fed to the workers, or injected into the abdomen of soldiers, or mixed with soldier homogenate. Despite these additional difficulties, the first studies were initiated in the late 1970s. In those early investigations, the labeled compounds were fed to the workers; thus the incorporation into the defensive substances was too weak for a decisive statement.

4.2.1

Isoprenoids

Diterpenes Prestwich and his group developed a method to inject the labeled sodium mevalonate (1133) in the abdomen of soldiers and thus proved that the isoprenoids in the frontal gland secretion were synthesized by the soldiers [755, 759]. Hojo et al. have investigated the genes involved in the synthesis of the diterpenes in Nasutitermitinae [843] and Reticulitermitinae [844] and found genes encoding the enzymes of the mevalonate pathway in the cytosol of class 1 cells surrounding the frontal gland reservoir (Scheme 161). In Nasutitermes takasagoensis, Hojo et al. identified seven genes encoding the following four enzymes associated with the diterpene syntheses: 3-hydroxy-3-methylglutaryl coenzyme A synthase (HMGS), 3-hydroxy-3methylglutaryl coenzyme A reductase (HMGR), farnesyl diphosphate synthase (FPPS), and geranylgeranyl diphosphate synthase (GGPPS) [845]. In Reticulitermes speratus they identified geranylgeranyl diphosphate synthase (GGPPS). Hydrolysis of the geranylgeranyl diphosphate produced leads to the characteristic defensive structure of the Heterotermitinae, geranyllinalool (186). The significant higher expression of these genes in the gland cells confirmed Prestwich’s suggestion that the (di)terpenes are synthesized in the frontal glands of soldiers. The chemical structures of diterpenes in Rhinotermitidae (open-chain diterpenes) and Termitidae (polycyclic diterpenes) differ substantially. Hojo suggested that the two suborders acquired the ability to synthesize diterpenes independently. Prestwich et al. injected Nasutitermes octopilis soldiers with labeled sodium acetates, 2-14C-()-mevalonate (2-14C -1133) and 2-14C-R-mevalonate (2-14C–R1133). After 24 h, the animals were decapitated, and the heads were extracted with nhexane/ethyl acetate. After purification by chromatography, the incorporation of labeled precursors was measured (Scheme 162). The single major monoterpene terpinolene and the three major diterpenes (one trinervitane 725 and two kempanes 885 and 886) were found to have incorporated the labeled compounds. Of the

cyclase

O

SCoA

OH

OPP

OP

O

FPPS

OPP

phosphomevalonate kinase

O

2 ((–)-(R)-neocembrene)

OH

acetoacetylthiolase

O

SCoA

HO

SCoA

O

Scheme 161 Biosynthesis of neocembrene in termites

HO

O

O

O HO

OH O

OPP

mevalonatediphosphate decarboxylase

O

polycyclic diterpenes

OH

HMGS

OPP

SCoA OH

O

OPP GGPPS

SCoA

OPP

HMGR HO

O OH

OPP

OH

OPP

geranyl diphosphate synthase

OPP

mevalonate kinase

Chemistry of the Secondary Metabolites of Termites 333

334

E. Gössinger

OH
HO




O-Na+

O










OPP

4,8,12,16-14C-1134

2-14C-1133 (sodium 2-14C- mevalonate)

14

(4,8,12,16- C-geranylgeranyl diphosphate)











PPO

H



H

HO




O H








OH 5,9,13,16-14C-725 (5,9,13,16-14C-2b,3a-dihydroxytrinervita-8,1(15)-diene)

+ HO

O H



H

H




H

+ HO


H




H




AcO H

5,9,13,16-14C-885 (5,9,13,16-14C-3ß-hydroxykemp-8-

5,9,13,16-14C-886 (5,9,13,16-14C-2ß-acetoxy-3ß-hydroxy-

en-5-one)

kemp-8-en-5-one)

Scheme 162 Proof of the mevalonate pathway for biosynthesis of polycyclic diterpenes by radiocarbon labeling

labeled starting materials, racemic mevalonate was incorporated more effectively than labeled acetate. The assumed natural precursor, (R)-mevalonate, was incorporated more effectively than racemic mevalonate [755]. These results [755] and the isolation of many structurally related polycyclic diterpenes afterward led to the proposition of the following biosynthesis pathways [31, 304, 353, 704, 721, 755, 759] (Scheme 163). The most simple way of cyclization of the universal diterpenoid precursor (E,E, E)-geranylgeranyl diphosphate (1134) is the Mg2+ assisted removal of the diphosphate and attack of the generating carbocation at the Δ14,15-double bond. Loss of the vicinal proton affords neocembrene (2) (Scheme 163). This cyclization has been investigated in detail in plants [846] and bacteria [847]. The cyclases found differ profoundly between plants and bacteria and allow no conclusion on the corresponding cyclases in animals. The further transannular cyclizations leading to the specific polycyclic diterpenes of termites have not been investigated in detail. It is assumed that the cyclizations occur stepwise, because structural examples of every transannular cyclization step following the neocembrene formation can be found in the defensive secretion of species of the Nasutitermitinae. Presumably, selective epoxidation of the monocyclic neocembrene (2) leads to 3,4-epoxyneocembrene

H





HO

H




H

H

+

H

HO iv (trinervitane)

vi

H

H




H H


O


H







E=H




H


H

H

iii


H







E = OR




v (trinervitane)

HO

HO




H

E


599 (3,4-epoxyneocembrene)

E(+)

729 (longipenol)

HO

H+

2 (neocembrene)

E(+)







Scheme 163 Proposed biosynthesis of the polycyclic diterpenes according to labeling studies

727 (3a-hydroxy15-rippertene)

HO

1134 (geranylgeranyldiphosphate)

PPO







HO





i













E(+)

HO

HO

H










E





H+

E = OR

H ii

H







H







H

OH







HO

HO




H

kempane

ii




H





H+

733 (secotrinervitane)

AcO




730 (secotrinervitane)

HO







Chemistry of the Secondary Metabolites of Termites 335

336

E. Gössinger

(599), found in the defensive secretion of Nasutitermes surinamensis [326]. Electrophilic epoxide cleavage followed by addition of the generated tertiary cation to the isopropenyl double bond and consecutive proton abstraction or addition of water furnishes the secotrinervitanes. It is not known if proton attack or epoxidation followed by electrophilic cleavage propagate the following transannular cyclizations. Electrophilic attack at the Δ7,8 double bond of secotrinervitane (ii) and subsequent proton abstraction lead to the most abundant polycyclic diterpene system the tricyclic trinervitanes (iv,v). The intermediate tertiary ion (iii) may attack the Δ11,12-double bond yielding the kempane system. The tetracyclic kempane system may also be formed by electrophilic attack of a proton at the Δ11,12-double bond of the secotrinervitane (ii). The generated carbocation attacks at the Δ15,16-double bond, and the newly formed cation attacks the Δ7,8-double bond. Subsequent proton abstraction leads to the kempanes. Proton attack at the Δ11,12-double bond of trinervitane (iv) leads to a secondary carbocation, which immediately attacks the Δ1,15-double bond. Either tertiary carbocation can be formed. If the new carbocation is formed at C-15, subsequent proton abstraction leads to longipenol (729). If the carbocation is formed at C-1, proton abstraction yields the kempane-type cation (vi). Meerwein rearrangement by 1,2 methyl shift with considerable decrease in strain furnishes 3α-hydroxy-15-rippertenol (727). The studies of Prestwich et al. with Nasutitermes rippertii showed that labeled mevalonate was incorporated in the trinervitanes as well as in the rippertanes found in the frontal gland secretion [759]. Kato’s biomimetic syntheses of the cyclic diterpenes of termites suggested the hypotheses by Prestwich [304]. There are caveats to these proposed biosynthesis pathways. First, most of the naturally occurring trinervitanes have no Δ11,12-double bond. Second, Prestwich failed to incorporate labeled neocembrene, which was prepared from neocembrene in the manner shown in Scheme 164 [848]. Selective anti-Markovnikov hydration of the isopropenyl moiety of neocembrene (2) with 9-borabicyclo[3.3.1]nonane (9-BBN) and subsequent oxidation led to the primary alcohol 1135, which was oxidized with pyridinium chlorochromate, and the aldehyde 1136 was then reduced with tritium-enriched sodium borohydride. Mild dehydration was achieved by conversion of the labeled primary alcohol [16-3H]1135 into the selenide by o-nitrophenyl selenocyanate and tributylphosphine, and the subsequent oxidation to the selenoxide was followed by cis elimination. After purification by silica gel chromatography and chromatography on a silver nitrateimpregnated silica gel column, the labeled neocembrene ([16-3H]-2) was applied topically as well as to homogenates of soldiers of Nasutitermes corniger, but no incorporation into the polycyclic diterpenes was observed [848]. Two further monocyclic diterpenes have been isolated from the defensive secretions of the frontal gland of termite soldiers, (3Z )-neocembrene (598) and cubitene (597), which possesses contrary to the cembrenes a 12-membered ring. Their biosyntheses have not been investigated. Thus, it is not known if (3Z)-neocembrene (598) is a cyclization product of geranylgeranyl diphosphate (1134) and consecutive (E)/(Z )-isomerization of the Δ3-double bond or if geranylneryl diphosphate (which has been detected as intermediate of the biosynthesis of dolichol) has been cyclized. Several hypotheses have been developed for the biosynthesis of cubitene (597) [652] (Scheme 165).

Chemistry of the Secondary Metabolites of Termites

337

H 1) 9-BBN, THF, rfl

OH

2) 30% aq H2O2, NaOH, C2H5OH, 20°C, 62% 2

1135

H O

PCC, CH2Cl2,

[3H]-NaBH4, C2H5OH, rt, ~60%

rt, 81%

1136

H

H

H* OH

1) o-NO2PhSeCN, Bu3P, THF, rt

H*

2) 30% aq H2O2, rt, 64% (2 steps)

[16-3H]-1135

[16-3H]-2

Scheme 164 Synthesis of 3H-labeled neocembrene

Prestwich’s first proposal was cyclization of the irregular acyclic diterpenoid diphosphate (i) [651]. This hypothesis had the advantage that it explained the biosynthesis of the accompanying bicyclic diterpene cubugene (600), because the intermediate tertiary carbocation (ii) yields cubitene (597) by proton abstraction, whereas addition to the second isopropenyl moiety furnishes the bicyclic skeleton of cubugene (iii). A 1,4-hydride shift and proton abstraction would complete the cubugene (600) synthesis. The finding that tobacco plants contain, along with cembrene-1,3-diol, the open-chain diterpene aldehyde, which could be formed by fragmentation of the diol, inspired Prestwich to formulate an analogous fragmentation of cembra-2,7,11,15-tetraene-4,6-diol (1137) and subsequent recyclization of the open-chain aldehyde (iv) to the 12-membered cubitene (597). A third hypothesis was inspired by cembrenones accompanied by cubitenones found in the soft coral Eunicea calyculata [657, 658]. Fenical et al. were intrigued by the similar pattern of functionalities in both structural types, the cembrenenones and cubitenones. Since photochemical 1,3-acyl shifts were known and even used in synthesis [849–851], the authors suggested that light might be the reason of the ring contraction. Indeed, when they irradiated (1E,3E,11E)-cembra-1,3,11-trien-6-one, they isolated along with other compounds the same products as found in this soft coral. It was suggested that this may also be the mechanism of the ring contraction of neocembrenones (v) to cubitenones (vi) in termites. Whereas this special soft coral dwells in shallow waters where sunlight can reach it, termites avoid sunlight. Thus, none of these hypotheses is convincing.

i

ii

iii

Scheme 165 Hypotheses of the biosynthesis of cubitene

600 (cubugene)

PPO

vi

597 (cubitene)

O

hν

v

iv

O

O

2

1137

OH H+

OH

338 E. Gössinger

Chemistry of the Secondary Metabolites of Termites

339

Of the two remaining bicyclic diterpenes found in Termitidae, cubugene (600) is thought as a by-product of the synthesis of cubitene (597), or it is its successor. No studies of the biosynthesis of (1S,6R,7R,11S)-()-biflora-4,10(19),15-triene (600) are known, although it deviates from the biosynthesis concept that cyclization of geranylgeranyl diphosphate (1134) to the 14-membered ring is the first step in the biosynthesis of the cyclic diterpenoids of the Termitidae. Prestwich suggested electrophilic ring closure of 1134 to the respective cyclodecadiene followed by transannular attack forming the cadinene core of 602 [128]. Alternatively, an electrophilic olefin cyclization cascade forming successively the two six membered rings of the cadinene core seems possible.

Biosynthesis of Mono- and Sesquiterpenes Mono- and sesquiterpenes found in the frontal glands of Termitidae and Rhinotermitidae soldiers have been shown to have high enantiomeric purity and mostly show the antipodal absolute configuration compared with the same compounds biosynthesized by plants [278, 289, 290, 297, 342]. These findings as well as Prestwich’s investigation with labeled mevalonate, which resulted in labeled mono-, sesqui-, and diterpenes in the defensive secretion were the first strong indications that the terpenoids are synthesized in the frontal glands of soldiers, especially as Prestwich could demonstrate that termite workers do not incorporate labeled acetates and mevalonate in terpenes, which discredited the hypothesis the terpenes of the frontal gland may be synthesized by endosymbionts [755].

4.2.2

Acetogenins

In the frontal glands of Neoisoptera, hydrocarbons differing from the CHCs have been found, and it is thought that they act as antihealants, when inserted in the wounds produced by the mandibles of the termite. Other long-chain compounds, alcohols, aldehydes, and ketones, were also found mostly in the higher developed Rhinotermitidae. Their biological activity is unclear. Long-chain vinyl ketones and keto aldehydes, found mostly in Schedorhinotermitinae, Rhinotermitinae, and Acorhinotermitinae, are contact poisons. All these compounds are thought to be synthesized according to the fatty acid pathway [538]. No detailed studies have been conducted. To resolve the question of the detoxification by the worker termites of the same species, Prestwich et al. synthesized labeled tetradeca-1,13-dien-3-one (286) (Scheme 166). Undec-10-enoic acid (1138) was reduced to the corresponding primary alcohol 283, which was esterified with tosyl chloride. Exchange of the tosylate 1139 by labeled sodium cyanide was followed by reduction to the imine by diisobutylaluminum hydride and hydrolysis to the labeled aldehyde 1140. Addition of vinylmagnesium bromide led to the unsaturated secondary alcohol, and oxidation with activated manganese dioxide furnished the desired 3-[14C]-tetradeca-1,13-dien-

340

E. Gössinger

1) NaAlH2(OCH2CH2OCH3)2, PhH, rfl, 90%

O (CH2)8

OH

1138

MgBr THF, 0°C, 87%

* O

(CH2)9

2) MnO2, n-hexane, rt, 76% [1-14C]-1140

OTs

(CH2)8 2) TsCl, NEt3, CH2Cl2, 0°C, 5 d; lactic acid, 0°C,1 d, 95%

1) Na14CN, DMSO, rt, 95% 2) DIBAH, PhH, rt, 70%

1139

O (CH2)9 * [3-14C]-282

O (CH2)9 * [3-14C]-1141

Scheme 166

14

C-3 labeling of tetradeca-1,13-dien-3-one and tetradeca-13-en-3-one

3-one ([3-14C]-282). Topical application of [3-14C]-282 to workers of Schedorhinotermes lamanianus for 24 h led to conversion of the labeled vinyl ketone into labeled acetate and to the labeled 3-[14C]-tetradec-13-en-3-one ([3-14C]-1141). This result was determined by homogenizing the workers, extraction of the homogenate with methanol followed by reversed-phase HPLC chromatography with different solvents and capillary gas chromatography and comparison with unlabeled reference substances. The ethyl ketone 1141 is much less toxic than the corresponding vinyl ketone 282. The detoxification by substrate-specific alkene reductases was unprecedented. Facile adduct formation via Michael addition of thiols, for example, glutathione, had been expected. Synthesis of these adducts and comparison with the extract of workers treated with [3-14C]-282 disproved the expected adduct formation [538]. Whereas long-chain fatty acids were found in the frontal glands of several Rhinotermitidae, macrolactones were detected in the frontal glands of two lower Nasutitermitinae genera and in the labial gland of a Macrotermitinae species. The sequence of the oxidation and the cyclization within the biosynthesis pathway of these macrolides have not been elucidated as yet.

4.2.3

Defensive Compounds with a Mixed Biosynthesis Origin

Unsaturated nitro compounds were isolated from Prorhinotermitinae. Originally Spanton and Prestwich hypothesized that the main defensive compound of the frontal gland of the Prorhinotermitinae, (E)-1-nitropentadec-1-ene (346), is biosynthesized starting with palmitic acid. Topically applied labeled palmitic acid failed to produce labeled defensive secretion in the frontal glands of the termites. Thus, the authors presented an alternative pathway involving myristic acid and

Chemistry of the Secondary Metabolites of Termites

341

serine [538]. Jirosova et al. confirmed the sphinganine-like biosynthesis, using labeled starting materials [514] (Scheme 167). Administration of 14C-2-glycine as well as L-14C-2-serine and 14C-1-tetradecanoic acid (myristic acid) to soldiers of Prorhinotermes simplex led to a remarkable increase in radioactivity of 346. Especially the labeled amino acids showed very good incorporation rates, whereas the labeled fatty acid was incorporated to a lesser extent. When the authors used 14C-1glycine or L-14C-1-serine, no incorporation was detected. These observations inspired the authors to formulate three possible biosynthesis pathways. Of those three pathways, they favored the pathway starting with glycine, because it is the energetically most advantageous, avoiding the oxidation and consequent decarboxylation step. Myristic acid adds to glycine, and the intermediate activated 3-keto acid is decarboxylated. Oxidation step(s) to the nitro compound [562, 852], reduction of the keto group, and elimination of water complete the proposed biosynthesis (Scheme 167). To decide between those pathways, the researchers undertook a very thorough investigation including metabolomics analysis, tracking age-related dynamics in the occurrence of putative intermediates, incubation of putative labeled intermediates, RNA sequencing of soldiers and workers to identify enzymes involved in the biosynthesis of nitroalkenes, and identifying genes expressing the initial steps of the biosynthesis of nitroalkenes [516, 853]. By means of these diverse techniques, the researchers were able to exclude the pathway via phosphorylation (see Scheme 167) and identify the enzymes of the first two steps of the biosynthesis, which correspond with those of the ceramide biosynthesis (Scheme 168). Tetradecanoic acid adds to serine, and the resulting β-keto acid decarboxylates to 1-hydroxy-2-aminohexadecan-3-one (1142). This first intermediate is already formed in the frontal gland of the late stage of the presoldiers (ps) of P. simplex. After reduction, the diol 1143 accumulated in the presoldiers immediately prior to the molting to soldiers (1 day). Oxidation to acid 1144 occurs in the freshly molted soldier (less than 24 h after ecdysis ¼ 0 days). A further oxidation step leads to β-keto acid 1145; the researchers could not exclude that part of 1145 is synthesized directly by oxidation of 1142. Decarboxylation to 1-aminopentadecan-2-one (1146) was also detected in freshly molted soldiers. The six-electron oxidation of the amino group of 1146 to the nitro group of 1-nitropentadecan-2-one 1147, which appears 3 days after molting, starts with oxidation to the hydroxylamine and proceeds to the oxime, which is in equilibrium with the nitroso compound, and ends as a nitro compound. Enzymes and cofactors are unknown. The most likely precursor of nitroalkene 346, the nitroalcohol i, has not been characterized. Seven days after ecdysis to the soldier, the optimal amount of 346 is reached. The pathway favored originally by the researchers starting with glycine and tetradecanoic acid could not be excluded, due to the appearance of 1-aminopentadecan-2-ol (1148) 1 day after molting to the soldier. The researchers offer an alternative explanation: 1148 may act as an inhibitor of the ceramide pathway [854], thus increasing the pool of sphinganine 1142 for the nitroalkene production. The authors investigated also the origin of the tetradecanoic acid. In fat bodies palmitic and stearic acid abound and very small amounts of tetradecanoic acid could be detected. Thus, synthesis of this acid in the frontal gland is assumed supported by transcriptomic data [516].

H2 N

–C*O2

C*OOH

hydroxymethyltransferase

C*OOH

O

(CH2)12 OH

–C*O2

(CH2)12 OH

O

H2N

OH

O

OH (CH2)12

(CH2)12

H2N

Scheme 167 Proposed biosynthesis of 1-nitropentadec-1-ene

H2N

OH

H2N

OH

O

(CH2)12

–CO2

H2N

O

H2N

O

OH

OH

(CH2)12

oxydase

O

OH

(CH2)12

ATP

O2N O

H2N

O (CH2)12

(CH2)12

OPO3

OH

reductase

O2N

O

O2N

O 2N

–CO2, –PO4H2-

oxydase

2-

(CH2)12

OH

(CH2)12

(CH2)12

OPO3

OH

342 E. Gössinger

O

OH

COOH

–CO2

(CH2)12 OH

O

–CO2

O

(CH2)12

O

(CH2)12

H2N

O

OH 1148 (1 d)

H2N

(CH2)12

(CH2)12

1147 (3 d)

O2N

OH

OH (CH2)12

1143 (–1 d)

H2N

O

oxydase

reductase

3-ketodihydrosphingosine

1146

(CH2)12

1146 (0 d)

H2N

1142 (ps, –1 d)

H2N

OH

Scheme 168 Biosynthesis of 1-nitropentadec-1-ene with age-related occurrence of intermediates

H2N

–CO2

(CH2)12 OH

serine palmitoyl transferase

(CH2)12

COOH

1145 (0 d, 7 d)

H2N

O

H2N

OH

O

O 2N

ceramides

i

OH

(CH2)12

OH

OH

346 (7 d)

O2N

(CH2)12

1144 (0 d)

H2N

O

(CH2)12

Chemistry of the Secondary Metabolites of Termites 343

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E. Gössinger

Scheme 169 Detoxification of nitroalkenes by reduction

* (CH2)12

NO2

346

* (CH2)12

NO2

347

As mentioned above, Spanton and Prestwich examined the detoxification of this contact poison by applying labeled 346 (preparation see Scheme 67) topically on the workers (pseudergates) of the Prorhinotermitinae and found that no conjugates were formed, but substrate-specific oxidoreductases hydrogenated the nitroalkene 346 to the nitroalkane 347 (Scheme 169).

5 Families and Subfamilies of the Termites Families and subfamilies of extant termites are listed below according to Engel, Grimaldi, and Krishna [1].

Family MASTOTERMITIDAE

Subfamily Mastotermes darwiniensis

EUISOPTERA Hodotermitidae Archotermopside Stolotermitidae

Stolotermitinae Porotermitinae

Kalotermitidae

NEOISOPTERA Stylotermitidae Rhinotermitidae

Coptotermitinae Heterotermitinae Prorhinotermitinae

Psammotermitinae Termitogetoninae Rhinotermitinae

Sphaerotermitinae Macrotermitinae Foraminitermitinae Syntermitinae

Nasutitermitinae Apicotermitinae Cubitermitinae Termitinae

Serritermitidae Termitidae

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Acknowledgment The author thanks the editors, especially Prof. Falk for his help and patience and Prof. Kinghorn for trying to improve my meager English. Many thanks to Dr. F. Wuggenig for reading through the manuscript. Last but not least I have to thank the team of the Fernleihe of OEZBPH; without their help in providing copies of hard to attain publications, this work could never have been done.

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384

E. Gössinger Edda Gössinger was born in Asparn/Zaya, Austria, in 1942. She completed her studies in chemistry at the University of Vienna with a Ph.D. thesis on the isolation and structure determination of natural products. She then became a postdoctoral fellow at ETH-Zürich, where she assisted in the partial synthesis of a steroid alkaloid and later on worked on a total synthesis of her own design. With this synthesis, she moved to NIH in Bethesda, Maryland, USA, where she continued to work on the total syntheses of natural products. She concluded her stay in the United States by getting acquainted with high-pressure chemistry at the State University of New York at Stony Brook. On returning to Austria, she joined the Institute of Organic Chemistry at the University of Vienna. There she received the Venia Legendi in 1982, which enabled her to lecture on the planning of syntheses and diverse themes on natural product chemistry. Her research interests focused mainly on the total synthesis of natural products, with excursions into chemical methodology and mechanistic aspects. She retired from her academic position in 2006.

Bioactive Compounds Involved in the Life Cycle of Higher Plants Hideyuki Shigemori

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Allelopathy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Phototropism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Cholodny–Went Theory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Repetition of the Theory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Bruinsma–Hasegawa Theory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Apical Dominance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Thigmonastic Movement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Nyctinastic Movement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Flowering . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Senescence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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1 Introduction Unlike animals, higher plants are sessile and permanently attached to the substrate. Therefore, to adeptly respond to changes in the environment, they exhibit various life phenomena such as allelopathy, phototropism, apical dominance, nyctinastic movement, flowering, and senescence. Such phenomena have been observed in detail, and hypotheses regarding their mechanisms have been proposed over time. In the course of such research, plant hormones that act as bioactive compounds related to these life phenomena have been discovered and their mechanisms revealed. With the recent advancements in molecular biology, these mechanisms have been explained in terms of the genes associated with these plant hormones. However, it has been found recently that these phenomena cannot be explained by H. Shigemori (*) Faculty of Life and Environmental Sciences, University of Tsukuba, Tsukuba, Ibaraki, Japan e-mail: [email protected] © Springer Nature Switzerland AG 2019 A. D. Kinghorn, H. Falk, S. Gibbons, J. Kobayashi, Y. Asakawa, J.-K. Liu (eds.), Progress in the Chemistry of Organic Natural Products, Vol. 109, https://doi.org/10.1007/978-3-030-12858-6_2

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plant hormones alone and that new bioactive compounds may also be involved in these processes. Meanwhile, doubts about conventionally accepted theories have arisen based on the findings of recent research studies. This contribution describes the studies conducted to elucidate the structures of bioactive compounds other than plant hormones that are involved in the lifecycle of plants and the mechanisms behind various life phenomena that utilize these compounds [1–3].

2 Allelopathy In the autumn in Japan, goldenrod (Solidago altissima) can be seen growing en masse by the roadside and on vacant land. The growth and development of other plants is suppressed within and around these areas because of chemical compounds emitted by goldenrod. The phenomenon involving the release of chemical compounds by plants that result in the inhibition of growth of other plant species is called allelopathy. This word was derived from the Greek words “αλληλων” (mutual) and “παθoζ” (sense). In the book “Der Einfluss einer Pflanze auf die andere: Allelopathie”, published in 1937 by the plant ecologist Hans Molisch, allelopathy was first defined as “a phenomenon in which a chemical compound secreted and released from a microbe or plant affects other plants in some manner” [4]. This phenomenon has been ascertained in the pre-Christian era, and the Greek philosopher Theophrastus (ca. 300 BC) wrote that chickpeas (Cicer arietinum), unlike other closely related plants of the Leguminosae family, “wear down soil” [5]. In a seminal chemical study conducted in 1928, the compound juglone (5-hydroxynaphthoquinone) (1) (Fig. 1) was isolated from the black walnut (Juglans nigra) and identified as an allelopathic compound. Juglone (1) is produced by a glycoside of 1,4,5-trihydroxynaphthalene, which is contained in the leaves and fruits of the black walnut, released into the soil through the action of rain or from fallen leaves and fruits, and then hydrolyzed by microbes and oxidized by the oxygen in the air. The mechanism of action of 1 (Fig. 1) thus involves the suppression of growth of surrounding plants [6]. Studies on allelopathy in goldenrod have also been conducted. Among the vegetation in the suburbs of Japan, ragweed (Ambrosia artemisiifolia) grows first, and after 2 years, daisies (Erigeron annuus) grow instead of ragweed. After 3 years, the daisies decline and the goldenrod becomes the dominant species, and these plants, after several more years, are replaced by the Japanese pampas grass (Miscanthus sinensis). It has been reported that the polyacetylene compounds produced by these Asteraceae species act as allelopathic compounds and contribute to such plant succession, which includes 2-(Z )dehydromatricaria ester (2) [7]. Other examples of recently discovered allelopathic compounds are shown in Fig. 1. The allelopathic compounds camphor (3) and 1,8-cineole (4) have been isolated from Salvia leucophylla Greene [8], nordihydroguaiaretic acid (5) from creosote bush (Larrea tridentata J.M. Coult.) [9], and patulin (6) from fungal-contaminated wheat (Triticum aestivum L.) [10]. Scopoletin (7), isolated from oat plants (Avena sativa L.), hinders mitosis of the roots of Timothy-grass (Phleum pratense L.) [11] and suppresses the oxidation

Bioactive Compounds Involved in the Life Cycle of Higher Plants OH

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O O CO2Me

O 1 (juglone)

3 (camphor)

2 ((Z)-dehydromatricaria ester)

O

HO

O OH

HO

O

O

OH 4 (1,8-cineole)

OH

6 (patulin)

5 (nordihydroguaiaretic acid)

OH

HO O

CO2H

HO

HO

O

O

NH2

HO

S O

7 (scopoletin)

9 (zeylanoxide A)

8 (L-DOPA)

OH

HO

O

OH HO O

O

O

OH OH

HO OH

OH

O OH 11 ((–)-lariciresinol)

10 (syringin) O

H N C N

NH

H 12 (cyanamide)

13 (caprolactam)

H2N

COOH

14 (6-aminohexanoic acid)

Fig. 1 Inhibitory allelochemicals

of indole-3-acetic acid (IAA) [12]. Numerous other allelopathic compounds that inhibit the growth of other plants have been isolated and their structure determined, including 3,4-dihydroxyphenylalanine (L-DOPA) (8) from mucuna (Stizolobium deeringianum Bort) [13]; zeylanoxide A (9) from Sphenoclea zeylanica Gaertn. [14]; L-tryptophan, syringin (10), and ( )-lariciresinol (11) from mesquite (Prosopis juliflora DC.) [15, 16]; and cyanamide (12) from hairy vetch (Vicia villosa Roth) [17]. Cyanamide (12) is a component of the fertilizer calcium cyanamide, but the fact that it is also obtained from unfertilized plants suggests that it is biosynthesized in plants. Since 12 has powerful growth-inhibiting activity, it is currently used for weed control in idle fields, abandoned areas, and fruit orchards that use hairy vetch [18]. As a result of mixed planting of buckwheat (Fagopyrum esculentum Moench)

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seeds with various other plants, species-selective growth-inhibiting or growthstimulating activity occurs in other plants, which we recently discovered. Isolation and analysis of the active compound in cress (Lepidium sativum L.) using the exudate of buckwheat seeds with growth-inhibiting activity as an indicator showed that it is caprolactam (13) [19]. Our research on the correlation between structure and activity to determine the activity expression site of 13 showed that this cyclic lactam structure plays an important role in growth suppression [20]. On the other hand, 6-aminohexanoic acid (14) has been identified as the active compound in the exudates of buckwheat seeds, with growth-stimulating activity against cockscomb (Celosia cristata (L.) Kuntze). On the basis of these findings, we believe that this activated allelopathic compound varies depending on how buckwheat seeds recognize other plants. As previously described, allelopathy is a phenomenon involving the release of a chemical compounds by one plant to suppress the growth of surrounding plants, thus, facilitating its own growth. Conversely, there are also cases in which a chemical compound released by one plant stimulates the growth of the surrounding plants. We refer to the former as “inhibitory allelopathy” and the latter as “stimulatory allelopathy.” Nearly all the compounds discovered thus far act in an inhibitory manner on the growth of other plants, but we have ascertained, under certain conditions, that stimulatory allelopathy definitely exists. When we planted cress (Lepidium sativum L.), burdock (Arctium lappa L.), and oat (Avena sativa L.) seeds, mixed with other plants, we observed that these markedly stimulated the growth of the other plants, thus demonstrating that the germinated seeds release stimulatory allelopathic compounds to their immediate environment. During the course of the study, we isolated a growth-stimulating compound from the exudate of cress seeds and found that it is a disaccharide consisting of rhamnose and deoxyuronic acid, and we named it lepidimoide (15), after the plant’s Latin name (Fig. 2) [21]. The Fig. 2 Stimulatory allelochemicals

OH

O

HO HO

O HO O

O OH

OH 15 (lepidimoide) O

O O

O

OH OH

O HO

HO

O O 16 (arctigenin)

O O 17 (arctigenic acid)

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greening of the buds was stimulated at the same time as shoot growth. This compound also has a chlorophyll synthesis-stimulating action judging from an increased amount of chlorophyll in the plant treated with this compound. Furthermore, multifaceted effects have been seen, such as the stimulated synthesis of 5-aminolevulinic acid, which is a rate-liming step of the chlorophyll synthesis system; increased leaf surface area, plant length, and fresh and dry weights; less time to flower bud formation; and an increased number of seeds in thale cress [22]. In addition, from burdock seed exudate, we isolated and identified arctigenin (16) and arctigenic acid (17) (Fig. 2) as stimulatory allelopathic compounds that exhibited the same or greater degree of growth-stimulating activity as that by lepidimoide (15) (Fig. 2) [23].

3 Phototropism The phenomenon involving the curvature of a plant’s stem toward the source of light (blue light) is called phototropism. A well-known hypothesis on the mechanism of phototropism often described in high school textbooks and many specialized books is the Cholodny–Went theory that was proposed in 1937 [24], although several problems relating to this concept have recently emerged.

3.1

Cholodny–Went Theory

In the book The Power of Movement in Plants (1880), English naturalist Charles Darwin, who is very widely recognized for his theory of evolution, point out that the tip of a plant serves as its main sensory region for light, based on the observation that stem curvature was not observed in irradiated plants with the bud tips cut off and covered using tin foil (Fig. 3) [25]. After this, Boysen-Jensen et al. discovered that plants curved when mica sheets were inserted into the tips of buds parallel to the direction of light but did not curve when inserted perpendicularly, thus surmising that curving occurs because of a stimulating factor produced in the tips moving horizontally (1926) (Fig. 4) [26]. Two years later, F.W. Went measured the amount

Fig. 3 Darwin’s experiment

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light A mica sheet inserted curvature on a parallel with light

light A mica sheet inserted on a vertical with light

no curvature

Fig. 4 Boysen-Jensen’s experiment

Fig. 5 Went’s experiment

Fig. 6 Cholodny–Went Theory

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of stimulating factor in the tips of buds that had been light irradiated in an Avena curvature test and hypothesized that curvature occurred based on the amount of the stimulating factor reaching the illuminated side and shaded side (Fig. 5) [27]. This stimulating factor was named auxin and was later identified as indole-3-acetic acid (IAA) [28]. According to Cholodny, Went’s theory could also be explained by gravitropism, facilitating the development of the Cholodny–Went theory, which states that phototropism is induced by the horizontal movement of auxin from the illuminated side to the shaded side (Fig. 6) [24].

3.2

Repetition of the Theory

More than 80 years have elapsed since the proposal of the Cholodny–Went theory, and experiments conducted in relation to this theory have further generated doubts on its concept. When Firn and Hasegawa et al. independently conducted Darwin’s experiment, they ascertained that plants undergo stem curvature in response to irradiation even when the tips of their buds were cut off and covered, although the degree of curvature was somewhat reduced (Fig. 7) [29, 30]. Furthermore, experiments conducted by Boysen-Jensen et al. showed that when mica sheets were inserted into the tips of buds, plant curvature occurred either parallel or perpendicular to the light (Fig. 8). It was also observed that curvature does not readily occur when mica sheets were inserted randomly (Fig. 8) [31], judging from the fact that the portions with the inserted mica sheets were greatly bent back to the outside in a photograph in the study (Fig. 4). The Went experiment showed a higher IAA level in the agar of the illuminated-side tissue compared to the shaded-side, as shown by the results of the Avena curvature test. Gas chromatography and high-performance liquid chromatography analysis of IAA levels in the respective agar fragments showed almost no differences between the two (Fig. 9) [32], indicating that auxin

Fig. 7 Repetition of Darwin’s experiment

392

H. Shigemori

Fig. 8 Repetition of Boysen-Jensen’s experiment. Left: a mica sheet was inserted on a parallel plane with light, right: a mica sheet was inserted on a vertical plane with light. Upper: normal; lower: damage owing to crude splitting

Fig. 9 Repetition of Went’s experiment

activity was reduced in the illuminated-side tissue by irradiation, but the amount of auxin was unchanged. This can be explained if one considers that a compound that suppresses the activity of auxin was produced and diffused in the agar fragment of the illuminated-side tissue. These three traditional experiments, thus, are related to doubtful results, which facilitated in the development of the Cholodny–Went theory.

Bioactive Compounds Involved in the Life Cycle of Higher Plants

3.3

393

Bruinsma–Hasegawa Theory

In contrast to the Cholodny–Went theory, the “growth-suppressing substance theory” or the Bruinsma–Hasegawa theory (1990), postulates that a growth-suppressing compound produced in the light-side tissue suppresses the movement of auxin, which is distributed uniformly on the illuminated side and shaded side, thus, resulting in the inhibition of growth of the illuminated-side tissue and the occurrence of stem curvature (Fig. 10) [33]. Hasegawa et al. ascertained that the amount of raphanusanin (18) and 4-methylthio-3-butenyl isothiocyanate (4-MTBI) (19) isolated from radish (Raphanus sativus) was higher in the illuminated-side tissue because of irradiation, resulting in growth suppression of the hypocotyl and the induction of curvature (Fig. 11) [34]. On the other hand, 2,4-dihydroxy-7-methoxy-2H-1,4-benzoxazin-3 (4H )-one (DIMBOA) (20) [35] and 6-methoxy-2-benzoxazolinone (MBOA) (21)

Fig. 10 Bruinsma–Hasegawa Theory

light myrosinase

S-glc

S 4-MTBG (inactive)

N

OSHO3

H+

C

S

19 (4-MTBI) (active) S

N H

N

S

S

-H+ N H

S

S

18 (raphanusanin) (active)

Fig. 11 Formation of radish phototropism-regulating compounds

394

H. Shigemori light O

O

O-Glc

N

O H2O

OH

O

b -glucosidase

OH

N

O

OH

glucose

22 (DIMBOA-Glc) (inactive)

O

20 (DIMBOA) (active)

O

O O N H 21 (MBOA) (active)

Fig. 12 Formation of maize phototropism-regulating compounds

light S-Glc NOSO3

N C S

myrosinase

N H

N H

CO2H N H indole-3-acetic acid

CN

nitrilase N H

23 (indole-3-acetonitrile) (active)

Fig. 13 Formation of Arabidopsis and cabbage phototropism-regulating compounds

[36] were isolated in wheat and corn (Zea mays L.); their levels increased with light irradiation and exhibited antiauxin activity. Further, in the illuminated-side tissue immediately before phototropism occurs, the amount of the inactive precursor molecule (DIMBOA-Glc) (22) decreases, and then the amounts of 20 and 21 increase (Fig. 12). Furthermore, indole-3-acetonitrile (23) has been shown to be a photo-induced growth-suppressing compound in both cabbage (Brassica oleracea L.) [37] and Arabidopsis thaliana Schur [38] (Fig. 13). It has also been shown that the level of 23 increased in wild-type A. thaliana after irradiation with blue light, whereas this did not change in the mutant strain, nph3-101, which does not elicit phototropism. Other light-induced growth-suppressing compounds include uridine (24) in oat plants and 8-epixanthatin (25) [39] and helian (26) [40] in sunflower (Helianthus annuus L.) (Fig. 14). These findings suggest the existence of species-specific plant compounds that control phototropism.

Bioactive Compounds Involved in the Life Cycle of Higher Plants Fig. 14 Phototropismregulating compounds from sunflower and oat

395

O HN HO

O

O N

O

O O

OH OH 24 (uridine) (oat)

HO

25 (8-epixanthatin) (sunflower)

HO HO

OH O

O

OH

26 (helian) (sunflower)

Studies using radish (dicots) and maize (monocots) showed that the activity of hydrolytic enzymes related to the production of light-induced growth-suppressing compounds was markedly increased in the illuminated-side tissues after short-term light irradiation (myrosinase in radish [41], β-glucosidase in maize [42]) (Figs. 11 and 12). Investigations using immunofluorescence microscopy demonstrated that raphanusanin (18) suppressed changes in the orientation of microtubules in the cell walls, resulting in growth inhibition [43], whereas DIMBOA (20) hardens cell walls and inhibits cell elongation [44]. It has also been shown that MBOA (21) suppresses the binding of auxin to auxin-binding proteins and inhibits the expression of the SAUR gene of auxin early response genes [45]. Since 18 and 21 act by eliminating apical dominance in garden pea plants, they may also act as polar auxin transport inhibitors. These findings on growth-suppressing compounds can serve as an additional mechanism responsible for the decreased growth of light-side tissues other than the mechanism involving the horizontal movement of auxin. These enumerated findings show that short-term blue light irradiation increases the expression of hydrolyzing enzymes in illuminated-side tissues, and active lightinduced growth-suppression compounds are produced from inactive glycosides. At the molecular level, light-induced growth-suppressing compounds bind to auxinbinding proteins and then downregulate the expression and enzyme activity induced by auxin. At the same time, at the cellular level, these inhibit cell elongation by suppressing changes in microtubule orientation induced by auxins, thereby causing growth suppression of illuminated-side tissues. During this time, the growth rate of shaded-side tissues does not change because the amount of light-induced growthsuppressing compound does not increase. As a result, a deviation in growth occurs between the two tissues, resulting in the stem curvature response (Fig. 15) [46].

396

H. Shigemori Plant seedling Illuminated side Blue light

Shaded side

Phototropin activation

Phosphorylation of membrane proteins

Local enzyme activation (b-glucosidase, myrosinase, etc.)

Inactive precursors (ex. DIMBOA-Glc, MTBG)

Active growth inhibitors (ex. DIMBOA, MTBI)

Endogenous auxin

Auxin binding protein(s)

Auxin binding protein(s)

Inhibited cell elongation

Cell elongation

Inhibited growth promotion

Growth promotion

Differential flank growth Phototropic curvature

Fig. 15 A mechanism of phototropism in higher plants

4 Apical Dominance The phenomenon involving the suppression of growth and differentiation of lateral buds, because of the presence of an apical bud, is called apical dominance and is generally considered a typical example of plant–organ correlation. By using fava beans (Vicia faba L.), Thimann and Skoog demonstrated in 1933 that cutting off the apical buds to enhance the growth of lateral buds is mainly because of the decrease in the plant hormone auxin produced by the apical buds [47]. On the other hand, another plant hormone, cytokinin, enhances growth of the lateral buds. These findings indicate that apical dominance is regulated by a balance between auxins, which are supplied by the apical buds, and cytokinin, which is supplied by the roots. However, it has also been shown that the lateral buds grow in the absence of apical buds and after direct exposure to auxins. It is possible that the contribution of auxin to apical dominance is neither simple nor direct. Additionally, a study showing that the lateral buds grow even after apical bud excision and before the time required for cytokinin to be supplied contradicts the fact that the mechanism of apical dominance

Bioactive Compounds Involved in the Life Cycle of Higher Plants I

I

O

397

OH O

COOH

N H

I 27 (2,3,5-triiodobenzoic acid)

28 (N-1-naphthylphthalamic acid)

O

CHO

N H

O

O OH

29 (indole-3-aldehyde)

O

O

30 (strigol)

Fig. 16 Bioactive compounds involved in apical dominance

is simply the balance between auxin and cytokinin. Thus, Hasegawa et al. administered the auxin activity-inhibiting compounds raphanusanin (18) and MBOA (21) discovered from radish and maize to intact apical buds, internodes, or lateral buds of garden pea plants (Pisum sativum L.), and examined the effect of those compounds on lateral bud growth. They also conducted the same experiment by using the polar auxin transport inhibitors 2,3,5-triiodobenzoic acid (27) and N-1naphthylphthalamic acid (28) (Fig. 16). The results showed that lateral bud growth occurred when 18 and 21 were provided to garden pea apical buds, internodes, or lateral buds. On the other hand, lateral bud growth was only observed when 27 and 28 were provided to intact garden pea apical buds and internodes. When provided directly to lateral buds, the lateral buds did not grow and apical dominance was not eliminated [48]. Thus, we attempted to find a lateral bud growth-suppressing compound other than an auxin. In our search for a compound with components other than those in auxin, we discovered indole-3-aldehyde (29) suppresses growth of lateral buds [49]. We confirmed that lateral bud growth is suppressed when 29 is directly administered to garden pea plants whose apical buds have been cut off. We believe that when apical buds are present, auxin supplied downward from apical buds induces the production of 29 while moving through the internodes, and this migrates into the lateral buds and suppresses lateral bud growth (Figs. 16 and 17). Extensive research on branching of shoots has been conducted using mutants. From A. thaliana, Leyser et al. discovered the mutants more axillary growth 1–4 (max1–4) that show excessive shoot branching. Additionally, garden pea ramosus (rms) and rice tillering dwarf (d) spontaneous mutants are involved in shoot branching [50]. After this, the RMS, MAX, and D genes were cloned, and it was shown that carotenoid cleavage dioxygenases CCD7 and CCD8 and F-box protein contribute to excessive branching in these mutants [51]. From the presumed function of the causative genes, it has been hypothesized that the RMS/MAX/D gene complex produces a carotenoid-derived compound that suppresses branching in the respective

398

H. Shigemori

Fig. 17 Plausible mechanism of apical dominance

plants [51]. A recent study involving rice d mutants has shown that strigolactones induce the specific effect [52], and similar results were obtained using garden pea rms mutants [53]. Strigol (30) (Fig. 16), a type of strigolactone, was first isolated from cotton root exudate [54] and is known to stimulate seed germination in the root parasitic weeds Striga and Orobanche, which belong to the Orobanchaceae family. Current research focuses on the association between parasitism and branching or apical dominance in plants.

5 Thigmonastic Movement The instantaneous movement of the leaves of mimosa (Mimosa pudica) when touched, which involves closing and drooping of the leaf joint, is called thigmonastic movement. This movement also occurs with temperature changes (i.e., when a flame is held close to a leaf). In this case, the leaves close and droop by using the pulvini on the leaf joints of the lobules, accessory lobes, and primary lobes as pivots. Motor cells are present in these pulvinus parts, and opening and closing of the leaves occurs by contraction of the motor cells. A decrease in turgor at this time occurs only in the motor cells on the lower side of the pulvinus, allowing a flexible response to the deformation of the cells because of decreased turgor. A recent study showed that actin cytoskeletons are present around these motor cells. Additionally, by using an experiment involving an anti-phosphotyrosine antibody and a tyrosine phosphatase inhibitor, it has been demonstrated that the quick movements of mimosa are regulated by phosphorylation–dephosphorylation of tyrosine residues in actin [55]. It has also been found that when a mimosa leaf is touched, an impulse is transmitted across the cells surrounding the fibrovascular bundles, resulting in the closure of leaves in succession. It is believed that this impulse is produced by an

Bioactive Compounds Involved in the Life Cycle of Higher Plants H

OH KOOC

COOK

31 (potassium L-malate)

HOOC

COO2+ •Mg COO-

32 (magnesium (E)-aconitate)

399

Me2NH2+•X– 33 (dimethylammonium salt)

Fig. 18 Stimulatory compounds of Mimosa pudica L.

electrical current carried by chlorine ions, and the ion channels on the cell membranes receive contact stimulation [56]. Chemical compounds may be present when this impulse is transmitted, and research on this topic has been conducted over a number of years. In 1998, S. Yamamura et al. isolated compounds that transmit stimulation from a mimosa leaf extract and conducted an activity test by using mimosa leaves as an indicator. Instrumental analysis allowed to identify these compounds as potassium L-malate (31), magnesium (E)-aconitate (32), and a dimethylammonium salt (33) (Fig. 18). Each individual compound did not elicit the action of closing leaves, but the combination of 31, 32, and 33 at extremely low concentrations of 10 8 to 10 9 M results in a thigmonastic movement [57].

6 Nyctinastic Movement The phenomenon in which leaves close at night and open in the morning is observed in many Leguminosae family plants, including mimosa. This phenomenon was known before the Christian era, and there are records of it from the time of Alexander the Great. It was reported by a French scientist in the seventeenth century that mimosa brought into a cave exhibited nyctinastic movement for several days in the pitch darkness. This suggests that the nyctinastic movement is not regulated by light but is controlled by the mimosa’s biological clock (see Plate 1). Later, in The Power of Movement in Plants (1880) [25], Darwin compiled records of scientific observation of nyctinastic movement in various plants. Early in the twentieth century, it was proposed that chemical compounds induced nyctinastic movement, although the various attempts to identify these compounds were not successful. In 1983, Schildknecht isolated K-PLMF1 (plant leaf-movement factor) (34) (Fig. 19) from plants that exhibit nyctinastic movement, and since he found it in plants exhibiting a variety of nyctinastic movements, he named it “turgorin,” a plant hormone related to nyctinastic movement [58]. Some textbooks and reference books of that time also described “turgorin” as a plant hormone. However, Yamamura et al. found that 34 exists in plants as a potassium salt, and although they examined its activity as a potassium salt, they found that it exhibits almost no activity. Thus, the search for the true active compound had begun, and potassium chelidonate (35) was discovered to cause leaves to close (nyctinastic compound) in cassia (Cassia mimosoides L.), which exhibits nyctinastic movement (Plate 1; Fig. 20) [59]. Studies on the correlation between structure and activity showed that the nature of this compound in being a potassium salt was responsible

400

H. Shigemori

'XULQJWKHGD\

$WQLJKW

Cassia mimosoides L.

Cassia occidentalis L.

Lespedeza cuneata G. Don

Plate 1 Nyctinastic movements of Cassia mimosoides L., C. occidentalis L., and Lespedeza cuneata G. Don

HO HO

OSO3H O O OH HO

OH

CO2H

34 (K-PLMF1; turgorin)

Fig. 19 Structure of K-PLMF1 (turgorin) of Schildknecht

for leaf closure. After this, during the search for a nyctinastic compound in another plant, Lespedeza cuneata G. Don (Plate 1), it was accidentally discovered that there is a fraction in which the leaves remain open even at night. This became the impetus for research to find compounds that cause leaves to open (leaf-opening compounds), resulting in the discovery of potassium lespedezate (36) (Fig. 20) [60]. In a subsequent study on the action in Cassia by using the nyctinastic compound potassium chelidonate (35) and the leaf-opening compound potassium lespedezate (36), an antagonistic action involving the opening and closing of leaves was discovered and was controlled by the two compounds at a concentration of 10 6 M in a 1:1 ratio [61]. Compounds having both activities were later isolated from various plants and their structures determined. As of the present time, leaf-closing and leaf-opening compounds 37–44 have been isolated from Mimosa pudica, Cassia mimosoides,

Bioactive Compounds Involved in the Life Cycle of Higher Plants

Leaf-Closing Compounds OH O

O HO HO KOOC

O

HO HO O

OH O

O

OH

OH

OH

COOK

401

O

COOK

O 35 (potassium chelidonate) (Cassia mimosoides L., Cassia occidentalis L.)

COOK

(Phyllanthus urinaria L.)

O

O

HO HO

OH OH KOOC

38 (phyllanthurinolactone)

37 (potassium 5-O-b -glucopyranoside gentisate) (Mimosa pudica L.)

O

OH

KOOC

OH OH 39 (potassium D-idarate)

40 (potassium b -D-glucopyranpsyl 11-hydroxyjasmonate) (Albizzia julibrissin Durazz.)

(Lespedeza cuneata G. Don)

Leaf-Opening Compounds

HO HO

OH O

OH O

HO HO

OH NH3 O

NH2

OH Ca2+

O

OH COOK

–

OH

COO–

OOC 2

41 (calcium 4-O-b -D-glucopyranosyl (Z)-4-coumarate) (Cassia mimosoides L.)

36 (potassium lespedezate) (Lespedeza cuneata G. Don)

42 (phyllurine) (Phyllanthus urinaria L.) OH

O

O

OH OH N

N

N NH2 OH NH2 43 (mimopudine)

(Mimosa pudica L.)

H3N

H N NH2

O N H

44 ((Z)-4-coumaroylagmatine) (Albizzia julibrissin Durazz.)

Fig. 20 Five pairs of leaf-movement factors. Each pair was isolated from the same nyctinastic plant

402

H. Shigemori

Phyllanthus urinaria L., L. cuneata, and Albizia julibrissin Durazz., and their structures have been determined (Fig. 20) [62–67]. The ten compounds have completely different structures but are characterized by a glucose moiety in either the nyctinastic or the leaf-opening compounds in each plant. In a later study that examined changes in the concentration of nyctinastic (glucoside-type compound) and leaf-opening (non-glucoside-type compound) compounds in Phyllanthus, almost no fluctuations were observed in leaf-opening compounds throughout a 24-h period, whereas the concentration of the nyctinastic compound was lower during the day and higher at night. This shows that since glucose is eliminated from the nyctinastic compound during the day and the concentration of that nyctinastic compound decreases, whereas at night, glucose binds to it and the concentration of nyctinastic compound increases. When a fluorescent probe was attached to this compound and then introduced to slices of pulvinus and observed by fluorescence microscopy, this compound was present near motor cells, and the active compound bound specifically to motor cells. Thus, these findings suggest that glucosidase is activated under the control of a biological clock and that because of glucose separating from or binding to the glucose-binding active compound, its concentration changes and the plant’s concentration balance of compounds having opposite activity reverses from day to night, resulting in the opening and closing of leaves, i.e., nyctinastic movement (Plate 1; Fig. 21) [68]. Additionally, Ueda et al. conducted a research study to identify the target molecules of these compounds using an enantiodifferential photoaffinity labeling experiment. Currently, two proteins, namely, (1) active substance receptor protein, which participates in control of cell turgor, and (2) β-glucosidase, which produces the rhythm of nyctinastic movement, participate in the control of nyctinastic movement [69].

Fig. 21 Mechanism of chemical control of nyctinasty

Bioactive Compounds Involved in the Life Cycle of Higher Plants

403

7 Flowering Plants form flower buds based on changes in the day length and by switching from trophic growth to reproductive growth. Undifferentiated cells in apical buds transform into flower primordia, and the switch is complete when flower bud formation occurs. M. K. Chailakhyan believed that differentiation into flower primordia is regulated by a leaf compound called florigen that is transmitted to the apical buds (1937) [70]. In 1958, he presented the two-component florigen hypothesis, wherein florigen is composed of two compounds, namely, gibberellin, which is a plant hormone discovered at that time, and anthesin (hypothesized), which has never been isolated to date. Then, in 1989, it was reported that flower buds formed when an ethanol extract of freeze-dried leaves of flowering tobacco (Nicotiana tabacum) was administered to the shoot apex of short-day wild spinach (Chenopodium album var. centrorubrum), although isolation of this florigen was not successful [71]. In the meantime, by performing bioassays involving duckweed, C. F. Cleland et al. identified salicylic acid (45) from the sieve tube fluid of cocklebur (Xanthium strumarium) as a compound having flowering-inducing activity [72]. Subsequently, nicotinic acid (46), nicotinamide (47), and L-pipecolinic acid (48) were isolated as flowering-inducing compounds from gibbous duckweed (Lemna gibba) [73, 74] and phenylglyoxal (49) from immature seeds of common morning glory (Ipomoea purpurea) [75] (Fig. 22). Furthermore, norepinephrine (50) has been isolated from the supernatant fraction of the fluid from crushed duckweed (Lemna paucicostata) [76], but in a subsequent study, the α-ketol linolenic acid derivative (S,12Z,15Z )-9-hydroxy-10-oxooctadeca12,15-dienoic acid (KODA) (51) was isolated from the sediment fraction (Fig. 23) [77]. Since high flowering-inducing activity was observed when norepinephrine was present, it was mixed with 51, incubated, and then purified, and the active substance FN1 (52) was isolated (Fig. 23) [78]. It has been shown that 51 is also present in Japanese morning glory (Pharbitis nil cv. ‘Violet’) and that its endogenous levels vary according to changes in the day length. Therefore, we focused our attention on the participation of these fatty acid-related compounds in flowering. By using A. thaliana, the aerial plant parts were continuously cultivated under short-day Fig. 22 Flower-inducing compounds

CO2H

CO2H

CONH2

OH N 45 (salicylic acid)

N

46 (nicotinic acid)

CO2H

O

47 (nicotinamide) CHO

NH 48 (L-pipecolinic acid)

49 (phenylglyoxal)

OH

O

O

OH

OH O

O

O

O O O

O

O O O

O

O

HOOC

O

?

?

54 (tuberonic acid glucoside)

b-Glc O

63 (arabidopside A)

?

53 (MGDG)

Fig. 23 Biosynthesis pathways of oxylipins

OH 55 (theobroxide)

HO

HO

OH

OH OH O

HO

HO

HO

O

O

HO

O

O

O

OOH

56 (epi-JA)

HOOC

OPDA

(13S)-HPOT

linolenic acid

O

O

HO

HO

O

O O

52 (FN1)

HO O

HO

OH NH2

N H

OH

50 ((–)-norepinephrine)

HO

HO

51 (KODA)

HO

404 H. Shigemori

Bioactive Compounds Involved in the Life Cycle of Higher Plants

405

conditions, given long-day treatment for 3 days and then cultivated under long-day conditions from after initiating treatment until flower buds formed. These were subjected to extraction by methanol and analyzed by means of high-performance liquid chromatography. The results showed a decrease in several peaks after flower bud formation, which represented sn1-O-(octadecatrienoyl)-sn2-O-(hexadecatrienoyl)mono-galactosyl diglyceride (MGDG) (53) (Fig. 23) [79]. When plants receiving long-day treatment under weak light the day before the bioassay were treated with this compound, flower bud formation of the plants was observed, but not stimulated in plants that had not received long-day treatment. These results suggest that 53 plays a key role in flower bud formation in A. thaliana, acting as a precursor or substrate of a flower bud-forming compound. On the other hand, tuberonic acid glucoside (54) was discovered as a compound that induces the formation of tubers in potato (Solanum tuberosum), migrating to the flower bud portion when flowering occurred under longday conditions (Fig. 23) [80]. Theobroxide (55), which is produced in the filamentous fungus Lasiodiplodia theobromae [81] induces flowering in potato plants and Japanese morning glory, but since this compound does not migrate, it is not metabolized in the plant, activating lipoxygenase; fatty acid-related compounds such as 51 are not considered to participate in the flowering process [82]. Araki et al. recently discovered that when A. thaliana senses a change in day length, its leaves express a FLOWERING LOCUS T (FT) gene related to flower bud formation stimulation and that flower buds are formed by its protein migrating and binding to the corresponding FT protein [83]. They did not clarify, however, how the FT protein in leaves migrates to shoot apical meristems and stimulates flowering. After this, by using a transgenic plant obtained by creating a fusion protein of FT expressed specifically in phloem cells and a fluorescent reporter protein, Corbesier et al. showed that this protein migrates to the apex and also migrates a long distance between grafted plants, thus, inducing flowering in A. thaliana. These results showed that the FT protein is a flowering hormone (florigen) [84]. On the other hand, Shimamoto et al. reported that a protein coded by an FT ortholog (homologous gene), Hd3a, migrates from the leaves to shoot apical meristems in rice and induces flowering [85]. These results obtained by using a long-day A. thaliana and short-day rice have shown that the FT/Hd3a protein synthesized in leaves in response to light cycle induces flowering in angiosperms. However, it cannot be ruled out that lowmolecular-weight compounds such as fatty acid-related compounds participate in flowering.

8 Senescence Senescence in plants is associated with a decrease in the amounts of proteins, sugars, and nitrogen in plant cells, which is a response to a lower level of chlorophyll production, which ultimately results in leaf abscission and fruit production. Thimann and Sachs confirmed the presence of compounds in an extract of garden pea root that stimulate aging, such as the amino acid L-serine [86, 87]. Ueda and Kato reported on

406

H. Shigemori

Fig. 24 Senescencepromoting and inhibiting compounds

O

CO2R 57 (methyl 7-epi-jasmonate (R = Me))

O O 58 (capillarin) O

59 (capillin)

60 (capillene)

O

CO2R 61 (jasmonic acid (R = H)) 62 (methyl jasmonate (R = Me))

the remarkable aging-stimulating activity in the ethyl acetate-soluble portion of leaves and stems of wormwood (Artemisia absinthium L.), consisting of methyl 7-epi-jasmonate (57) (Fig. 24) [88]. Similarly, capillarin (58) (Fig. 24) was isolated as a chlorophyll decomposition-stimulating compound in capillary wormwood (Artemisia capillaris) [89], whereas capillin (59) and capillene (60) were identified as chlorophyll decomposition-suppressing compounds in the same plant (Fig. 24) [89]. Incidentally, because 59 and 60 in relatively high concentrations stimulate the decomposition of chlorophyll, xanthophyll, and carotenoid-based pigments, it is known that they have both activities [89, 90]. To date, the presence of jasmonic acid (61) and methyl jasmonate (62) have been determined in at least 150 families and 206 plant species, including filamentous fungi, mosses, pteridophytes, and higher plants. As related compounds, we identified novel oxylipin compounds in A. thaliana and named these as arabidopsides A–D (63–66) and F, after the plants Latin name (Fig. 25) [91–93]. These compounds are glycolipids in which the jasmonic acid biosynthesis precursors, 12-oxophytodienoic acid (OPDA) and dinoroxophytodienoic acid, are bound to galactoglycerol. When we examined their chlorophyll decomposition-stimulating activity by using leaf fragments of oat (Avena sativa), it was found that they exhibit the same degree of activity as methyl jasmonate (62) [94]. Decomposition of chlorophyll starts with the phytol moiety

Bioactive Compounds Involved in the Life Cycle of Higher Plants

HO OR O O HO OH

O O O O

HO OR O O

HO

O

OH O

63 (arabidopside A (R = H)) 64 (arabidopside C (R = a-Gal))

407

O O O

O

O O 65 (arabidopside B (R = H)) 66 (arabidopside D (R = a-Gal))

Fig. 25 Senescence-promoting compounds from Arabidopsis thaliana

being hydrolyzed. The product then moves into vacuoles through a multistep reaction and is metabolized via chlorophyll catabolites of the bile pigment type [95] into a monopyrrole and substituted maleimides [96]. The cDNA of chlorophyllase, which performs the phytol hydrolysis, was cloned and it was shown that the expression of one of the genes of chlorophyllase increases because of 62 [97]. The research conducted thus far suggests that, at least in A. thaliana, biosynthesis from linolenic acid synthesis to OPDA occurs in the chloroplasts, and glycoproteins that have OPDA such as arabidopsides are present on the membrane of chloroplasts (Fig. 23). For this reason, we believe that arabidopsides activate chlorophyllase and stimulate chlorophyll decomposition more directly than by 62.

9 Conclusion This contribution provides an overview of research on bioactive compounds involved in the lifecycle of higher plants. This year (2019) will mark Darwin’s 210th birthday, although the inexplicable and mysterious plant phenomena observed and recorded by Darwin et al. have now been elucidated based on the discovery of specific bioactive compounds. Ironically, doubt has arisen due to efforts in verifying the mechanisms behind these phenomena by using the latest technology. However, despite the identification of new errors amid recent discoveries, the information contained in current biology textbooks and reference books has not been updated. Thus, the latest research results in this particular area, must be cited. Although the current theories still warrant validation, additional investigations on the biology and chemistry should provide the necessary information to establish the finer details of these mechanisms. Acknowledgments The author wishes to thank Professor S. Yamamura, Keio University, Professor K. Hasegawa, and Associate Professor K. Yamada, University of Tsukuba, for valuable discussions and comments.

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69. Nakamura Y, Manabe Y, Inomata S, Ueda M (2010) Recent advances on bioorganic chemistry of plant metabolites controlling nyctinasty. Chem Rec 10:70 70. Chailakhyan MK (1936) Hormonal theory of plant development. Dokl Akad Nauk SSSR 3:443 71. Chailakhyan MK, Lozhnikova V, Seidlova F, Krekule J, Dudko N, Negretzky V (1989) Floral and growth responses in Chenopodium rubrum L. to an extract from flowering Nicotiana tabacum L. Planta 178:143 72. Cleland CF, Ajami A (1974) Identification of the flower-inducing factor isolated from aphid honeydew as being salicylic acid. Plant Physiol 54:904 73. Fujioka S, Yamaguchi I, Murofushi N, Takahashi N, Kaihara S, Takimoto A, Cleland CF (1986) Isolation and identification of nicotinic acid as a flower-inducing factor in Lemna. Plant Cell Physiol 27:103 74. Fujioka S, Sakurai A, Yamaguchi I, Murofushi N, Takahashi N, Sakurai S, Takimoto A (1987) Isolation and identification of L-pipecolic acid and nicotinamide as flower-inducing substances in Lemna. Plant Cell Physiol 28:995 75. Suzuki Y, Yamaguchi I, Murofushi N, Takahashi N (1988) Identification of phenylglyoxal as a flower-inducing substance of Lemna from Pharbitis purpurea. Agric Biol Chem 52:1013 76. Takimoto A, Kaihara S, Shinozaki M, Miura J (1991) Involvement of norepinephrine in the production of a flower-inducing substance in the water extract of Lemna. Plant Cell Physiol 32:283 77. Yokoyama M, Yamaguchi S, Inomata S, Komatsu K, Yoshida S, Iida T, Yokokawa Y, Yamaguchi M, Kaihara S, Takimoto A (2000) Stress-induced factor involved in flower formation of Lemna is an α-ketol derivative of linolenic acid. Plant Cell Physiol 41:110 78. Yamaguchi S, Yokoyama M, Iida T, Okai M, Tanaka O, Takimoto A (2001) Identification of a component that induces flowering of Lemna among the reaction products of α-ketol linolenic acid (FIF) and norepinephrine. Plant Cell Physiol 42:1201 79. Hisamatsu Y, Goto N, Hasegawa K, Shigemori H (2006) A glycolipid involved in flower bud formation of Arabidopsis thaliana. Bot Stud 47:45 80. Yoshihara T, Amanuma M, Tsutsumi T, Okumura Y, Matsuura H, Ichihara A (1996) Metabolism and transport of [2-14C]()-jasmonic acid in the potato plant. Plant Cell Physiol 37:586 81. Nakamori K, Matsuura H, Yoshihara T, Ichihara A, Koda Y (1994) Potato micro-tuber inducing substances from Lasiodiplodia theobromae. Phytochemistry 35:835 82. Kong F, Gao X, Nam K-H, Takahashi K, Matsuura H, Yoshihara T (2005) Theobroxide inhibits stem elongation in Pharbitis nil by regulating jasmonic acid and gibberellin biosynthesis. Plant Sci 169:721 83. Abe M, Kobayashi Y, Yamamoto S, Daimon Y, Yamaguchi A, Ikeda Y, Ichinoki H, Noguchi M, Goto K, Araki T (2005) FD, a bZIP protein mediating signals from the floral pathway integrator FT at the shoot apex. Science 309:1052 84. Corbesier L, Vincent C, Jang S, Fornara F, Fan Q, Searle I, Giakountis A, Farrona S, Gissot L, Turnbull C, Coupland G (2007) FT protein movement contributes to long-distance signaling in floral induction of Arabidopsis. Science 316:1030 85. Tamaki S, Matsuo S, Wong HL, Yokoi S, Shimamoto K (2007) Hd3a protein is a mobile flowering signal in rice. Science 316:1033 86. Thimann KV, Sachs T (1966) The role of cytokinins in the fasciation disease caused by Corynebacterium fascians. Am J Bot 53:731 87. Shibaoka H, Thimann KV (1970) Antagonisms between kinetin and amino acids: experiments on the mode of action of cytokinins. Plant Physiol 46:212 88. Ueda J, Kato J (1980) Isolation and identification of a senescence-promoting substance from wormwood (Artemisia absinthium L.). Plant Physiol 66:246 89. Ueda J, Kojima T, Nishimura M, Kato J (1984) Isolation and identification of aliphatic compounds as chlorophyll-preserving and/or degrading substances from Artemisia capillaris Thunb. Z Pflanzenphysiol 113:189 90. Ueda J, Shiotani Y, Kojima T, Kato J, Yokota T, Takahashi N (1983) Identification of hinesol as a chlorophyll-preserving substance. Plant Cell Physiol 24:873

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Hideyuki Shigemori is a Professor at the University of Tsukuba. He obtained his Ph.D. degree from Keio University, Japan, in 1990. Following a period as a researcher at the Sagami Chemical Research Institute, he became an Assistant Professor at the Faculty of Pharmaceutical Sciences, Hokkaido University, in the same year and was promoted to Associate Professor in 1999. He transferred to the University of Tsukuba in 2001 as an Associate Professor and was promoted to Professor at this university in 2009. He has published over 200 original papers as well as several book chapters and books on natural product chemistry and plant physiology.

Chemical Diversity and Biological Activity of African Propolis Natalia Blicharska and Veronique Seidel

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Chemical Diversity of Phytochemicals Isolated from African Propolis . . . . . . . . . . . . . . . . . . . 3 Biological Activity of African Propolis Extracts and Constituents . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Antimicrobial Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.1 Antibacterial Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1.2 Antifungal Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Antiparasitic and Antiprotozoal Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Anti-inflammatory Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Antioxidant Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5 Organ-Protective Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6 Antiviral and Immunomodulatory Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.7 Cytotoxic and Antitumor Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.8 Other Miscellaneous Biological Effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Conclusion and Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

415 417 428 433 433 434 434 435 435 436 437 437 438 438 440

1 Introduction Natural remedies, sourced from plants, microbes, and animal products, have for centuries played a significant role in traditional medicine and continue to represent a unique reservoir of new chemical entities for drug discovery research. Between 1981 and 2014, 50% of all small molecule-approved drugs released on the market were either directly derived from a natural product or synthetic compounds based on a natural product pharmacophore [1].

N. Blicharska · V. Seidel (*) Natural Products Drug Discovery Research Group, Strathclyde Institute of Pharmacy and Biomedical Sciences, University of Strathclyde, Glasgow, UK e-mail: [email protected] © Springer Nature Switzerland AG 2019 A. D. Kinghorn, H. Falk, S. Gibbons, J. Kobayashi, Y. Asakawa, J.-K. Liu (eds.), Progress in the Chemistry of Organic Natural Products, Vol. 109, https://doi.org/10.1007/978-3-030-12858-6_3

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Plate 1 Propolis sample. Photograph: Goldmull, Creative Commons 3.0

Propolis, also known as bee glue, is a natural substance produced by honeybees from plant secretions such as resins and sticky exudates on leaf buds and plant wounds (Plate 1). The word propolis is derived from Greek, in which πρo (pro) means “at the entrance to” and πóλις (polis) means “community” or “city.” Bees use propolis as a construction and repair material to seal gaps and smooth out internal walls in their hives and as an antiseptic coating to generally protect the hive from external contamination [2–4]. The chemical composition of propolis can be highly variable, and this is attributed to differences in plant sources, governed by factors such as climatic conditions and seasons, within the geographical locations from where it is collected [4–6]. Propolis has a long history of use as a natural remedy for a variety of conditions, and there has been in recent years a renewed interest in reinvestigating the potential of propolis for drug development with some significant advancements in the understanding of its chemistry and biological activity [7–11]. The purpose of this contribution is to report more specifically on the current body of knowledge on the chemistry and biological activity of African propolis.

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2 Chemical Diversity of Phytochemicals Isolated from African Propolis Propolis is a complex mixture composed of resins, wax, fatty acids, essential oils, pollen, sugars, enzymes, minerals, and microelements [11]. Over 500 phytochemicals have been collectively identified in propolis collected from around the world [10]. It is well understood that the phytochemical composition (and subsequent biological activity) of propolis is highly variable and largely depends on the available flora in different locations and season of collection [6]. Indeed, propolis is commonly categorized into distinct chemotypes according to its botanical origin. For example, samples collected from temperate regions tend to possess phytochemicals that are characteristic of poplar bud phenolics due to the main source of propolis in such regions being poplar trees. Thus, “poplar-type” propolis is rich in flavonoids, cinnamic acids and esters, phenolic acids and esters, and other aromatic acids [12]. On the other hand, bees collecting propolis from tropical regions have a wider array of plant sources at their disposal, and propolis from tropical regions is characterized by the presence of other types of phytochemicals such as terpenoids, lignans, stilbenes, benzophenones, and phenolic lipids [13–17]. Standard hyphenated techniques (e.g., HPLC-DAD, GC-MS, LC-MS, and LC-MS-MS) have been largely employed to chemically profile propolis samples [18–20]. However, it has to be said that in some cases the true identity of specific phytochemicals could not be conclusively confirmed using the aforementioned techniques alone [21]. For that reason, we decided to focus our literature search for this contribution solely on phytochemicals from African propolis that have been isolated through the use of various preparative chromatography techniques and characterized unambiguously by means of mass spectrometry and NMR analysis. Our search retrieved a total of 134 phytochemicals from propolis samples originating from nine African countries. The structurally diverse phytochemicals were grouped into five main chemical classes including several phenylpropanoids (Fig. 1, Table 1), flavonoids (Figs. 2 and 3, Table 2), terpenoids (Figs. 4–6, Table 3), phenolic lipids (Fig. 7, Table 4), and a range of miscellaneous compounds (Figs. 8 and 9, Table 5) [22–36].

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N. Blicharska and V. Seidel O R2

A=

OR1

R3

B=

1 (caffeic acid) 2 (prenyl caffeate) 3 (methyl caffeate) 4 (isopentyl caffeate) 5 (2-methyl-2-butenyl (E)-caffeate) 6 (3-methyl-3-butenyl-(E)-caffeate) 7 (phenethyl-(E)-caffeate, CAPE) 8 (p-coumaric acid) 9 (p-coumaric acid methyl ester) 10 (cinnamic acid) 11 (isoferulic acid)

R1

R2

R3

= = = OH R1 = A, R2 = R3 = OH R1 = Me, R2 = R3 = OH R1 = B, R2 = R3 = OH R1 = C, R2 = R3 = OH R1 = D, R2 = R3 = OH R1 = E, R2 = R3 = OH R1 = R3 = OH, R2 = H R1 = Me, R2 = H, R3 = OH R1 = R2 = R3 = H R1 = H, R2 = OH, R3 = OMe

C=

D=

E=

OH OH

HO

HO

OH O

HO O

COOH

HO

COOR1

12 (caftaric acid) R1 = H 13 (caftaric acid 1-methyl ester) R1 = Me

O

O O O

COOR1 COOH

14 ((+)-chicoric acid) R1 = H 15 ((+)-chicoric acid methyl ester) R1 = Me

Fig. 1 Structures of phenylpropanoids isolated from African propolis

Table 1 Phenylpropanoids isolated from African propolis Compound Caffeic acid (1) Prenyl caffeate (2) Methyl caffeate (3) Isopentyl caffeate (4) 2-Methyl-2-butenyl (E)-caffeate (5) 3-Methyl-3-butenyl-(E)-caffeate (6) Phenethyl-(E)-caffeate (CAPE) (7) p-Coumaric acid (8) p-Coumaric acid methyl ester (9) Cinnamic acid (10) Isoferulic acid (11) Caftaric acid (12) Caftaric acid methyl ester (13) (+)-Chicoric acid (14) (+)-Chicoric acid methyl ester (15)

Origin Algeria Algeria Algeria Algeria Algeria Algeria Algeria Algeria Algeria Algeria Algeria Egypt Algeria Algeria Algeria Algeria

Refs. [22, 23] [22] [22] [22] [15] [15] [15] [22] [22] [22] [22] [24] [23] [23] [23] [23]

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R8 R7

R1 R2

O

R3

R6 A=

R5 R4

O R1 = R3 = R5 = R6 = R8 = H, R2 = R4 = OH, R7 = OMe 16 (acacetin) 17 (quercetin) R1 = R3 = R6 = H, R2 = R4 = R5 = R7 = R8 = OH 18 (3-O-methyl quercetin) R1 = R3 = R6 = H, R2 = R4 = R7 = R8 = OH, R5 = OMe 19 (kaempferol) R1 = R3 = R6 = R8 = H, R2 = R4 = R5 = R7 = OH 20 (chrysin) R1 = R3 = R5 = R6 = R7 = R8 = H, R2 = R4 = OH 21 (tectochrysin) R1 = R3 = R5 = R6 = R7 = R8 = H, R2 = OMe, R4 = OH 22 (galangin) R1 = R3 = R6 = R7 = R8 = H, R2 = R4 = R5 = OH 23 (galangin-5-O-methyl ether) R1 = R3 = R6 = R7 = R8 = H, R2 = R5 = OH, R4 = OMe 24 (myricetin-3,7,4',5'-tetramethyl ether) R1 = R3 = H, R2 = R5 = R7 = R8 = OMe, R4 = R6 = H 25 (apigenin) R1 = R3 = R5 = R6 = R8 = H, R2 = R4 = R7 = OH 26 (pectolinarigenin) R1 = R5 = R6 = R8 = H, R2 = R4 = OH, R3 = R7 = OMe 27 (pilosin) R1 = R2 = R4 = OH, R5 = R6 = R8 = H, R3 = R7 = OMe 28 (ladanein) R1 = R5 = R6 = R8 = H, R3 = R4 = OH, R7 = R2 = OMe 29 (macarangin) R2 = R4 = R5 = R7 = OH, R1 = R6 = R8 = H, R3 = A 30 (izalpinin) R1 = R3 = R6 = R7 = R8 = H, R4 = R5 = OH, R2 = OMe 31 (pachypodol) R1 = R3 = R6 = H, R4 = R7 = OH, R2 = R5 = R8 = OMe 32 (3,3'-dimethoxy-5,7,4'-trihydroxyflavone) R1 = R3 = R6 = H, R2 = R4 = R7= OH, R5 = R8 = OMe 33 (3-methoxy-5,7,4'-trihydroxyflavone) R1 = R3 = R6 = R8 = H, R2 = R4 = R7 = OH, R5 = OMe 34 (quercetin-3,7-di-O-methyl ether) R1 = R3 = R6 = H, R4 = R7 = R8 = OH, R2 = R5 = OMe

R8 R9

R1 R2

O

R3

R6

OH C=

OH

R5 R4

35 (naringenin) 36 (6-prenylnaringenin) 37 (8-prenylnaringenin) 38 (pinocembrin) 39 (pinobanksin) 40 (pinobanksin-3-acetate) 41 (pinobanksin-3-(E )-caffeate) 42 (pinostrobin) 43 (isonymphaeol C) 44 (isonymphaeol B) 45 (isonymphaeol D) 46 (nymphaeol B) 47 (lonchocarpol A) 48 (6,8-diprenyl-aromadendrin) 49 (lespedezaflavanone C) 50 (6,8-diprenyleriodictyol) 51 (liquiritigenin)

B= R7

O

O

R1 = R3 = R5 = R6 = R8 = R9 = H, R2 = R4 = R7 = OH R1 = R5 = R6 = R8 = R9 = H, R2 = R4 = R7 = OH, R3 = B R1 = B, R2 = R4 = R7 = OH, R3 = R5 = R6 = R8 = R9 = H R1 = R3 = R5 = R6 = R7 = R8 = R9 = H, R2 = R4 = OH R1 = R3 = R6 = R7 = R8 = R9 = H, R2 = R4 = OH, R5 = OH R1 = R3 = R6 = R7 = R8 = R9 = H, R2 = R4 = OH, R5 = OAc R1 = R3 = R6 = R7 = R8 = R9 = H, R2 = R4 = OH, R5 = OC R1 = R3 = R5 = R6 = R7 = R8 = R9 = H, R2 = OMe, R4 = OH R1 = R3 = R5 = R9 = H, R2 = OMe, R4 = R7 = R8 = OH, R6 = A R1 = R3 = R5 = R9 = H, R2 = R4 = R7 = R8 = OH, R6 = A R1 = R3 = R5 = R9 = H, R2 = R7 = R8 = OH, R4 = OMe, R6 = A R1 = R3 = R5 = R6 = H, R2 = R4 = R7 = R8 = OH, R9 = A R1 = R3 = B, R2 = R4 = R7 = OH, R5 = R6 = R8 = R 9= H R1 = R3 = B, R2 = R4 = OH, R5 = OH, R6 = R7 = R8 = R9 = H R1 = R8 = B, R2 = R4 = R7 = OH, R5 = OH, R3 = R6 = R9 = H R1 = R3 = B, R2 = R4 = R7 = R8 = OH, R5= R6 = R9 = H R1 = R3 = R4 = R5 = R6 = R8 = H, R2 = R7 = OH

Fig. 2 Structures of flavonoids isolated from African propolis I

420

N. Blicharska and V. Seidel R1 R2

O

HO

O

OH

R5

3

R

R4

O

R6

O

52 (genistein) R1 = R3 = R5 = H, R2 = R4 = R6 = OH 53 (calycosin) R1 = R3 = R4 = H, R2 = R5 = OH, R6 = OMe

54 ((3S)-vestitol)

Fig. 3 Structures of flavonoids isolated from African propolis II Table 2 Flavonoids isolated from African propolis Compound Acacetin (16) Quercetin (17) 3-O-Methyl-quercetin (18) Kaempferol (19) Chrysin (20) Tectochrysin (21) Galangin (22) Galangin-5-O-methyl ether (23) Myricetin-3,7,40 ,50 -tetramethyl ether (24) Apigenin (25) Pectolinarigenin (26) Pilosin (27) Ladanein (28) Macarangin (29) Izalpinin (30) Pachypodol (31) 3,30 -Dimethoxy-5,7,40 -trihydroxyflavone (32) 3-Methoxy-5,7,40 -trihydroxyflavone (33) Quercetin-3,7-di-O-methyl ether (34) Naringenin (35) 6-Prenylnaringenin (36) 8-Prenylnaringenin (37) Pinocembrin (38)

Pinobanksin (39) Pinobanksin-3-acetate (40) Pinobanksin-3-(E)-caffeate (41) Pinostrobin (42)

Origin Algeria Algeria Algeria Algeria Algeria Egypt Algeria Egypt Algeria Egypt Egypt Algeria Tunisia Algeria Algeria Algeria Algeria Kenya Nigeria Egypt Tunisia Egypt Egypt Egypt Algeria Nigeria Nigeria Algeria Nigeria Egypt Algeria Algeria Algeria Algeria Egypt

Refs. [22] [22] [22] [15, 22] [15, 22, 25, 26] [24, 27] [22] [24, 27] [15, 22] [24] [24] [15] [28] [15, 25, 26] [25, 26] [25, 26] [25, 26] [13] [29] [24] [28] [24] [24] [24] [22] [29] [29] [22] [29] [24] [15] [15, 22] [15] [22] [24, 27] (continued)

Chemical Diversity and Biological Activity of African Propolis

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Table 2 (continued) Compound Isonymphaeol C (43) Isonymphaeol B (44) Isonymphaeol D (45) Nymphaeol B (46)

Origin Egypt Egypt Egypt Egypt Nigeria Cameroon Congo Cameroon Cameroon Congo Nigeria Algeria Nigeria Nigeria

Lonchocarpol A (47) 6,8-Diprenylaromadendrin (48) Lespedezaflavanone C (49) 6,8-Diprenyleriodictyol (50) Liquiritigenin (51) Genistein (52) Calycosin (53) (3S)-Vestitol (54)

Refs. [30] [31] [31] [31] [29] [17] [17] [17] [17] [17] [29] [22] [29] [29]

R1 OH

HO 56 (18-hydroxy-cis-clerodan-3-en-15-oic acid) R1 = COOH 57 (cistadiol) R1 = CH2OH

55 ((–)-(S)--terpineol)

OH

HO OH O 58 (isoagathotal)

R1

COOH 1

60 (cupressic acid) R = COOH 62 (torulosol) R1 = CH2OH 64 (torulosal) R1 = CHO

59 (imbricatoloic acid)

OH

OH

R1 61 (isocupressic acid) R1 = COOH 63 (agathadiol) R1 = CH2OH

HOOC 65 (totarol)

Fig. 4 Structures of terpenoids isolated from African propolis I

66 (pimaric acid)

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N. Blicharska and V. Seidel

HO

HO 68 (cycloart-12,25-dien-3-ol)

67 (3b -cycloartenol)

Fig. 4 (continued)

COOH

COOH

COOH R1

HO

O

O

69 (ambonic acid)

70 (ambolic acid)

71 (mangiferonic acid) R1 = H 72 (27-hydroxymangiferonic acid) R1 = CH2OH

COOH COOH

R1

R1 74 (3b -cycloartenol-26-oic acid) R1 = b -OH 75 (3a -cycloartenol-26-oic acid) R1 = a-OH

73 (mangiferolic acid) R1 = b -OH 76 (isomangiferolic acid) R1 = a -OH

COOMe R1 OH HO

HO 77 (methyl-3b ,27-dihydroxycycloart-24-en-26-oate)

78 (betulinaldehyde) R1 = CHO 79 (betulin) R1 = OH R2 R3

R1

R1

O 80 (lupenone)

R1

81 (lupeol) = OH 82 (lupeol acetate) R1 = OAc

83 (-amyrin) 84 (-amyrin acetate) 85 (3a-hydroxy-olean-12-en-30-ol) 86 (erythrodiol)

Fig. 5 Structures of terpenoids isolated from African propolis II

R1 = b -OH, R2 = R3 = Me R1 = b -OAc, R2 = R3 = Me R1 = a-OH, R2 = CH2OH, R3 = Me R1 = b -OH, R2 = Me, R3 = CH2OH

Chemical Diversity and Biological Activity of African Propolis

423

O

HO 87 (-amyrin)

88 (-amyrone)

HO

AcO

89 (25-cyclopropyl-3b -hydroxyurs-12-ene)

90 (pseudotaraxasterol acetate)

HO

AcO 91 (taraxasterol acetate)

92 (lanosterol)

HOOC

HO

HO 93 (3b -hydroxylanostan-9,24-dien-21-oic acid)

94 (3b -sitosterol)

HO 95 (bacchara-12,21-dien-3b -ol)

Fig. 6 Structures of terpenoids isolated from African propolis III

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Table 3 Terpenoids isolated from African propolis Compound

Origin

Refs.

(–)-(S)-α-Terpineol (55) 18-Hydroxy-cis-clerodan-3-en-15-oic acid (56) Cistadiol (57) Isoagathotal (58) Imbricatoloic acid (59) Cupressic acid (60) Isocupressic acid (61) Torulosol (62) Agathadiol (63) Torulosal (64) Totarol (65) Pimaric acid (66) 3β-Cycloartenol (67)

Cameroon Algeria Algeria Algeria Algeria Algeria Algeria Algeria Algeria Algeria Algeria Algeria Cameroon Egypt Cameroon Cameroon Nigeria Cameroon Nigeria Cameroon Cameroon Cameroon Egypt Egypt Cameroon Cameroon Cameroon Cameroon Cameroon Congo Cameroon Cameroon Cameroon Egypt Cameroon Cameroon Cameroon Cameroon Nigeria Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon

[17] [15] [15] [15] [15] [15] [15] [15] [15] [15] [22] [22] [16] [27] [32, 33] [16] [34] [16, 33] [16, 33] [34] [33] [16] [27] [27] [16] [33] [17] [33] [17, 32] [17] [16, 32] [17, 32] [16, 17] [27] [17] [17] [17, 32] [16, 17] [34] [17] [32] [17] [17] [17] [33] [33] [17]

Cycloart-12,25-dien-3β-ol (68) Ambonic acid (69) Ambolic acid (70) Mangiferonic acid (71) 27-Hydroxymangiferonic acid (72) Mangiferolic acid (73) 3β-Cycloartenol-26-oic acid (74) 3α-Cycloartenol-26-oic acid (75) Isomangiferolic acid (76) Methyl-3β,27-dihydroxycycloart-24-en-26-oate (77) Betulinaldehyde (78) Betulin (79) Lupenone (80) Lupeol (81) Lupeol acetate (82) β-Amyrin (83) β-Amyrin acetate (84) 3α-Hydroxyolean-12-en-30-ol (85) Erythrodiol (86) α-Amyrin (87) α-Amyrone (88) 25-Cyclopropyl-3β-hydroxyurs-12-ene (89) Pseudotaraxasterol acetate (90) Taraxasterol acetate (91) Lanosterol (92) 3β-Hydroxylanostan-9,24-dien-21-oic acid (93) β-Sitosterol (94) Bacchara-12,21-dien-3β-ol (95)

n R1 = H, n = 10 R1 = H, n = 13 R1 = H, n = 14 R1 = H, n = 15 R1 = H, n = 16 R1 = H, n = 18

Fig. 7 Structures of phenolic lipids isolated from African propolis

109 (5-pentadecylresorcinol) R1 = OH, n = 14 110 (5-hexadecylresorcinol) R1 = OH, n = 15 111 (5-heptadecylresorcinol) R1 = OH, n = 16

96 (3-undecylphenol) 97 (3-tetradecylphenol) 98 (3-pentadecylphenol) 99 (3-hexadecylphenol) 100 (3-heptadecylphenol) 101 (3-nonadecylphenol)

R1

OH

n1

n2

112 (5-((10’Z)-pentadecyl)-resorcinol) 113 (5-((8’Z)-heptadecyl)-resorcinol) 114 (5-((11’Z)-heptadecyl)-resorcinol) 115 (5-((12’Z)-heptadecyl)-resorcinol) 116 (5-((14’Z)-heptadecyl)-resorcinol) 117 (5-((14’Z)-nonadecyl)-resorcinol)

102 (3-((10’Z)-pentadecenyl)-phenol) 103 (3-((12’Z)-pentadecenyl)-phenol) 104 (3-((8’Z)-heptadecenyl)-phenol) 105 (3-((12’Z)-heptadecenyl)-phenol) 106 (3-((14’Z)-heptadecenyl)-phenol) 107 (3-((13’Z)-nonadecenyl)-phenol) 108 (3-((14’Z)-nonadecenyl)-phenol)

R1

OH

R1 = OH, n1 = 9, n2 = 3 R1 = OH, n1 = 7, n2 = 7 R1 = OH, n1 = 10, n2 = 4 R1 = OH, n1 = 11, n2 = 3 R1 = OH, n1 = 13, n2 = 1 R1 = OH, n1 = 13, n2 = 3

R1 = H, n1 = 9, n2 = 3 R1 = H, n1 = 11, n2 = 1 R1 = H, n1 = 7, n2 = 7 R1 = H, n1 = 11, n2 = 3 R1 = H, n1 = 13, n2 = 1 R1 = H, n1 = 12, n2 = 4 R1 = H, n1 = 13, n2 = 3

Chemical Diversity and Biological Activity of African Propolis 425

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Table 4 Phenolic lipids isolated from African propolis Compound 3-Undecylphenol (96) 3-Tetradecylphenol (97) 3-Pentadecylphenol (98) 3-Hexadecylphenol (99) 3-Heptadecylphenol (100) 3-Nonadecylphenol (101) 3-((100 Z )-Pentadecenyl)-phenol (102) 3-((120 Z )-Pentadecenyl)-phenol (103) 3-((80 Z )-Heptadecenyl)-phenol (104) 3-((120 Z )-Heptadecenyl)-phenol (105) 3-((140 Z )-Heptadecenyl)-phenol (106) 3-((130 Z)-Nonadecenyl)-phenol (107) 3-((140 Z )-Nonadecenyl)-phenol (108) 5-Pentadecylresorcinol (109) 5-Hexadecylresorcinol (110) 5-Heptadecylresorcinol (111) 5-((100 Z )-Pentadecenyl)-resorcinol (112) 5-((80 Z )-Heptadecenyl)-resorcinol (113) 5-((110 Z )-Heptadecenyl)-resorcinol (114) 5-((120 Z )-Heptadecenyl)-resorcinol (115) 5-((140 Z )-Hheptadecenyl)-resorcinol (116) 5-((140 Z )-Nonadecenyl)-resorcinol (117)

Origin Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon Cameroon

Refs. [16] [16] [16] [16] [16] [16] [16] [16] [16] [16] [16] [16] [16] [16] [16] [16] [16] [16] [16] [16] [16] [16]

OH O

OH

OH OH

HO HO

O

O O

O OH

118 (tyrosol)

119 (1'-O-eicosanyl glycerol)

O

O O

121 (medicarpin)

OH

O

O

O

O

O O

120 (deperoxidized derivative of plukenetione C)

O 122 ((+)-sesamin)

O

O HO 123 (riverinol)

Fig. 8 Structures of miscellaneous compounds isolated from African propolis I

Chemical Diversity and Biological Activity of African Propolis O

427 R1

O

O

O

O O

O

O

O

O

O O

O

125 (6-methoxydiphyllin) R1 = OH 126 (phyllamyricin C) R1 = H

124 (tetrahydrojusticidin B)

R1 OH HO

R2 A= OH

127 ((E)-resveratrol) R1 = R2 = H 128 (5-((E)-3,5-dihydroxystyryl)-3-((E)-3,7-dimethylocta-2,6-dien-1-yl)benzene-1,2-diol) R1 = OH, R2 = A

OH O

OH

HO

HO

OH

HO

O R

OH

130 (schweinfurthin A) R = OH 131 (schweinfurthin B) R = OMe

129 ((E)-5-(2-(8-hydroxy-2-methyl-2-(4-methylpent-3-en-1-yl)2H-chromen-6-yl)vinyl)-2-(3-methylbut-2-en-1-yl) benzene-1,3-diol)

Fig. 8 (continued)

O

OH R1

HO

A= OR2

O R3

132 (gerontoxanthone H) R1 = R2 = H, R3 = A 133 (6-deoxy-γ-mangostin) R1 = A, R2 = R3 = H 134 (1,7-dihydroxy-3-O-(3-methylbut-2-enyl)-8-(3-methylbut-2-enyl)xanthone) R1 = R3 = H, R2 = A

Fig. 9 Structures of miscellaneous compounds isolated from African propolis II

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Table 5 Miscellaneous compounds isolated from African propolis Compound Tyrosol (118) 10 -O-Eicosanylglycerol (119) Deperoxidized derivative of plukenetione C (120) Medicarpin (121) (+)-Sesamin (122) Riverinol (123) Tetrahydrojusticidin B (124) 6-Methoxydiphyllin (125) Phyllamyricin C (126) (E)-Resveratrol (127) 5-((E)-3,5-Dihydroxystyryl)-3-((E)-3,7-dimethylocta-2,6-dien-1yl)benzene-1,2-diol (128) (E)-5-(2-(8-Hydroxy-2-methyl-2-(4-methylpent-3-en-1-yl)-2Hchromen-6-yl)vinyl)-2-(3-methylbut-2-en-1-yl)benzene-1,3-diol (129) Schweinfurthin A (130) Schweinfurthin B (131) Gerontoxanthone H (132) 6-Deoxy-γ-mangostin (133) 1,7-Dihydroxy-3-O-(3-methylbut-2-enyl)-8-(3-methylbut-2-enyl) xanthone (134)

Origin Algeria Cameroon Cameroon Nigeria Libya Nigeria Kenya Kenya Kenya Algeria Ghana

Refs. [22] [33] [35] [29] [36] [29] [13] [13] [13] [22] [35]

Ghana

[35]

Kenya Kenya Nigeria Nigeria Nigeria

[13] [13] [34] [34] [34]

3 Biological Activity of African Propolis Extracts and Constituents Extracts of African propolis have been investigated for a range of biological activities including for their antimicrobial, antiparasitic/antiprotozoal, antiviral, antioxidant, anti-inflammatory, organ-protective, immunomodulatory, and cancerrelated inhibitory properties. Some interesting biological effects have also been reported for specific phytochemicals present in African propolis (Table 6) [37–132].

Chemical Diversity and Biological Activity of African Propolis

429

Table 6 Biological studies performed on phytochemicals from African propolis Compound Caffeic acid (1)

Phenethyl-(E)-caffeate (CAPE) (7)

p-Coumaric acid (8) Caftaric acid (12) Caftaric acid methyl ester (13) (+)-Chicoric acid (14) (+)-Chicoric acid methyl ester (15) Acacetin (16) Quercetin (17)

Kaempferol (19) Chrysin (20)

Tectochrysin (21) Galangin (22)

Myricetin-3,7,40 ,50 -tetramethyl ether (24) Apigenin (25) Pectolinarigenin (26)

Ladanein (28) Macarangin (29)

Izalpinin (30)

Tested biological activity Antimicrobial Antioxidant Antifibrinolytic, anticollagenolytic Antimicrobial Antioxidant Anti-inflammatory Antitumor Antimicrobial Antifibrinolytic Antifibrinolytic Antifibrinolytic, anticollagenolytic Antifibrinolytic, anticollagenolytic Antitumor Aromatase inhibition Antimicrobial Antioxidant Anti-inflammatory Antitumor Myeloperoxidase inhibition Antioxidant Antimicrobial Antioxidant Anti-inflammatory Antitumor Antioxidant Anti-inflammatory Antimicrobial Anti-inflammatory Antitumor Antitumor Estrogenic Anti-inflammatory Antitumor Hepatoprotective Antitumor Antiviral Antimicrobial Antiprotozoal Antioxidant Antioxidant Anti-inflammatory Antimuscarinic

Refs. [37] [13, 38– 40] [23] [41] [13, 38] [42] [43] [44] [23] [23] [23] [23] [45–47] [48] [41] [22, 38, 49] [41] [50] [22] [38] [51, 52] [27] [27] [27] [27] [53] [51, 54] [41] [55, 56] [57] [58] [59] [60, 61] [62] [63] [64] [13] [29] [13] [65, 66] [53] [66] (continued)

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Table 6 (continued) Compound Pachypodol (31)

Tested biological activity Antiviral Antitumor

6-Prenylnaringenin (36)

Tyrosinase inhibition Antitumor Antiprotozoal Antimicrobial Antiprotozoal Antioxidant Antitumor Estrogenic Antimicrobial

8-Prenylnaringenin (37)

Pinocembrin (38)

Pinobanksin (39) Pinobanksin-3-acetate (40) Pinostrobin (42)

Isonymphaeol C (43) Isonymphaeol B (44) Isonymphaeol D (45) Nymphaeol B (46)

Lonchocarpol A (47)

Liquiritigenin (51)

Genistein (52)

Antioxidant Neuroprotective Anti-inflammatory Anti-apoptotic Aromatase inhibition, estrogenic Antiprotozoal Antimicrobial Antimicrobial Antimicrobial Phosphodiesterase and acetylcholinesterase inhibition Antioxidant Antitumor Antimicrobial Antioxidant Antitumor Antimicrobial Antimicrobial Antiprotozoal Antioxidant Antitumor Antimicrobial Antiprotozoal Antioxidant Anti-inflammatory Antitumor Anti-inflammatory Xanthine oxidase inhibition Estrogenic Estrogenic

Refs. [67] [50, 68, 69] [70] [71, 72] [29] [73] [29] [74] [71, 72] [75] [37, 51, 52] [27] [76] [76] [76] [48] [29] [37, 52] [51] [52] [27] [27] [77] [30] [78, 79] [80] [31] [40, 81] [29, 82] [40, 78] [80, 82] [83] [84] [74] [85] [74] [86] [87] [88] [58] (continued)

Chemical Diversity and Biological Activity of African Propolis

431

Table 6 (continued) Compound Calycosin (53) (3S)-Vestitol (54) Cistadiol (57) Isoagathotal (58) Cupressic acid (60) Isocupressic acid (61)

Agathadiol (63) Torulosal (64) Totarol (65) Pimaric acid (66) 3β-Cycloartenol (67) Cycloart-12,25-dien-3β-ol (68) Ambonic acid (69) Mangiferonic acid (71)

Mangiferolic acid (73) 3β-Cycloartenol-26-oic acid (74)

3α-Cycloartenol-26-oic acid (75) Isomangiferolic acid (76) Methyl-3β,27-dihydroxycycloart-24-en-26oate (77) Betulin (79)

Lupenone (80)

Tested biological activity Antiprotozoal Antiprotozoal Antiprotozoal Antimicrobial Antitumor Antimicrobial, antioxidant Hepatoprotective Antimicrobial Antioxidant Antitumor Hepatoprotective Antimicrobial Antitumor Antimicrobial Antimicrobial Retinoic acid receptor activation Antiatherosclerotic Antioxidant Acetylcholinesterase inhibition Antimicrobial Antiprotozoal Antiprotozoal Antimicrobial Antioxidant Antitumor Antitumor Antioxidant Acetylcholinesterase inhibition Phosphodiesterase inhibition Antioxidant Acetylcholinesterase inhibition Antitumor Antioxidant

Refs. [29] [29, 89] [90] [52] [77] [91] [91] [52, 91] [91] [77, 92] [91] [52] [77] [93] [52] [94] [95] [27] [27] [32] [34] [34] [96] [33, 96] [96, 97] [97] [27] [27] [27] [27] [27] [97] [33]

Antitumor Immunomodulatory Anti-inflammatory Antiprotozoal Antimicrobial Antiviral Anti-inflammatory Antioxidant Antitumor

[98] [99] [100] [100] [100] [100] [101] [96] [96] (continued)

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Table 6 (continued) Compound Lupeol (81)

β-Amyrin (83) β-Amyrin acetate (84) Erythrodiol (86)

α-Amyrin (87) 25-Cyclopropyl-3β-hydroxyurs-12-ene (89) 3β-Hydroxylanostan-9,24-dien-21-oic acid (93) β-Sitosterol (94)

3-((100 Z )-Pentadecenyl)-phenol (102) 5-Pentadecylresorcinol (109)

5-Heptadecylresorcinol (111)

5-((100 Z )-Pentadecenyl)-resorcinol (112) 5-((80 Z )-Heptadecenyl)-resorcinol (113) 5-((110 Z )-Heptadecenyl)-resorcinol (114) 5-((120 Z )-Heptadecenyl)-resorcinol (115) 5-((140 Z )-Nonadecenyl)-resorcinol (116) 10 -O-Eicosanylglycerol (119) Deperoxidized derivative of plukenetione C (120) Medicarpin (121) (+)-Sesamin (122)

Tested biological activity Antiporotozoal Antimicrobial, antioxidant Antitumor Antiprotozoal Antioxidant Antimicrobial Antioxidant Antitumor Antiplatelet Anti-inflammatory Antiprotozoal Antimicrobial Antioxidant

Refs. [89] [96] [98] [89] [27] [32, 44] [102] [102, 103] [104] [101] [34] [32] [33]

Anti-inflammatory Analgesic, antiparasitic, antimutagenic Antitumor Antihyperglycemic Immunomodulatory Antifertility Antioxidant Antimicrobial Acetylcholinesterase inhibition Anti-inflammatory Antitumor DNA-cleaving activity Antiviral Anti-inflammatory Antitumor Cytochrome P450s, P-glycoprotein and pregnane X receptor inhibition Antimicrobial DNA-cleaving activity Anti-inflammatory Antitumor Antiviral Antitumor Antioxidant Antiprotozoal

[105] [106]

Antiprotozoal Antiprotozoal Anti-inflammatory

[107] [108] [109] [110] [111] [112] [113] [114] [115–117] [118] [119] [114] [120] [121] [122] [118] [114] [123] [119] [120] [33] [35] [29, 89] [36] [124, 125] (continued)

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Table 6 (continued) Compound Riverinol (123) 6-Methoxydiphyllin (125) Phyllamyricin C (126) (E)-Resveratrol (127)

5-((E)-3,5-Dihydroxystyryl)-3-((E)-3,7dimethylocta-2,6-dien-1-yl)benzene-1,2-diol (128) (E)-5-(2-(8-Hydroxy-2-methyl-2(4-methylpent-3-en-1-yl)-2H-chromen-6-yl) vinyl)-2-(3-methylbut-2-en-1-yl)benzene1,3-diol (129) Schweinfurthin A (130) Schweinfurthin B (131) Gerontoxanthone H (132) 6-Deoxy-γ-mangostin (133) 1,7-Dihydroxy-3-O-(3-methylbut-2-enyl)-8(3-methylbut-2-enyl)xanthone (134)

3.1 3.1.1

Tested biological activity Antiprotozoal Antimicrobial Antimicrobial Anti-inflammatory Cancer chemopreventive Anti-inflammatory Cardioprotective Antiobesity Antiprotozoal

Refs. [29] [13] [13] [126] [127] [128] [129] [130] [35]

Antiprotozoal

[35]

Antimicrobial Antitumor Antimicrobial Antitumor Antiprotozoal Antiprotozoal Antiprotozoal

[13] [131, 132] [13] [131] [34] [34] [34]

Antimicrobial Activity Antibacterial Activity

The current development of antibiotic resistance among bacterial pathogens is a global health threat that urgently requires the development of new drugs to fight off infections [133]. Many studies have investigated the antibacterial potential of propolis, but it is important to highlight at this point that the use of inconsistent assay methodologies and extraction techniques and the screening of chemically non-standardized samples make it difficult to compare the available data [39, 134, 135]. It has been noted, in agreement with previous observations [136], that extracts of African propolis showed potent activity against Gram-positive bacteria such as Staphylococcus aureus, Staphylococcus epidermidis, Streptococcus pyogenes, Streptococcus anginosus, Enterococcus faecalis, Bacillus subtilis, Bacillus cereus, and beta-hemolytic streptococci [13, 17, 25, 51, 134, 135, 137, 138] and rather weak

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activity against Gram-negative pathogens like Escherichia coli, Pseudomonas aeruginosa, Proteus mirabilis, Klebsiella pneumoniae, Enterobacter cloacae [13, 17, 25, 32, 39, 51, 134, 135, 137], and non- and alpha-hemolytic streptococci [25]. Interestingly, Egyptian propolis has been found to inhibit the growth of drugresistant strains of E. coli and S. aureus [139]. Studies have also reported the ability of African propolis to impair bacterial biofilm formation with Tunisian propolis displaying direct antibiofilm activity on a range of oral pathogens including a range of Enterococcus, Gemella, and Streptococcus spp. [138]. When applied to nanoparticle-treated catheters, Moroccan propolis showed a reduced adherence of methicillin-resistant Staphylococcus aureus (MRSA) strains [140]. Few studies have endeavored to identify the phytochemicals responsible for the observed antibacterial activities of African propolis or to unravel the mechanism of action by which these compounds exert their biological effects. However, it has been noted that the presence of flavonoids appears to confer some antibacterial activity to African propolis [25, 26, 141]. In particular, the observation that galangin (present in Algerian and Egyptian propolis) causes damage to bacterial cytoplasmic membranes [54] may explain the observed antibacterial effect of this type of propolis.

3.1.2

Antifungal Activity

African propolis has demonstrated variable activity against fungal pathogens including Aspergillus spp. [142–144], Candida sp. [17], Cryptococcus neoformans [51], Colletotrichum sp. and Fusarium spp. [143]. Studies investigating the antifungal effect of African propolis against Candida albicans have afforded mixed results. Good activity against C. albicans and C. neoformans has been reported for South African propolis [51]. For samples of Egyptian origin, the activity has been reported to be similar to that of ketoconazole and clotrimazole [134]. On the other hand, neither the extracts nor any of the phytochemicals isolated from Kenyan propolis have exhibited activity against C. albicans [13]. These results are likely to be explained by the different chemical compositions between South African, Kenyan, and Egyptian propolis. The latter has also been reported to inhibit the growth of Aspergillus spp. involved in the production of aflatoxins in foodstuffs [142, 144] and has showed promising antibiofilm activity against Candida spp., including drug-resistant strains, which is of particular potential interest in dental care [139, 145].

3.2

Antiparasitic and Antiprotozoal Activity

The global challenge posed by the rise in drug resistance also applies to diseases caused by parasites, including protozoa [146]. Nigerian and Libyan propolis extracts

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have demonstrated activity against Trypanosoma brucei brucei that was greater than that obtained for individually isolated phytochemicals, suggesting the presence of a synergistic effect between compounds [21, 34, 36]. Libyan propolis has also revealed varying degrees of activity against Leishmania donovani, Plasmodium falciparum, and Crithidia fasciculata [21], and the antitrypanosomal activity of Nigerian propolis has been observed against two multidrug-resistant strains. (3S)Vestitol (54), 6-prenylnaringenin (36), 8-prenylnaringenin (37), α-amyrin (87), and gerontoxanthone H (132) were identified as being responsible for these antiprotozoal effects [29, 34]. Cameroonian and Ghanaian propolis have shown activity against T. brucei brucei, with the greatest effect obtained for the deperoxidized derivative of plukenetione C (120) [35]. Egyptian propolis has demonstrated activity against Fasciola gigantica [141, 147] and Cryptosporidium spp. [148], and its combined administration with the antiparasitic drug praziquantel significantly lowered the burden of Schistosoma mansoni in infected mice [149].

3.3

Anti-inflammatory Activity

Inflammation is an important physiological response to harmful stimuli and one that is necessary for tissue repair. Chronic inflammation has been implicated in the pathogenesis of a range of diseases including neurodegeneration [150], cancer [151], cardiovascular diseases [152], and asthma [153]. The anti-inflammatory effect of Egyptian propolis has been reported in asthmatic mice and attributed to the presence of flavonoids and phenolics [154]. The selective targeting of phosphodiesterase type-4 (PDE4) is a strategy pursued in the search for novel treatments for respiratory diseases associated with inflammation such as asthma [155], and 3β-cycloartenol-26-oic acid (74), isolated from Egyptian propolis, has been shown to strongly inhibit the activity of this enzyme. The flavonoids chrysin (20) and pinostrobin (42) also reduced PDE4 activity [27]. Quercetin (17) and galangin (22) have been identified as responsible for the antiinflammatory activity of South African propolis [41]. Phenethyl-(E)-caffeate (CAPE) (7), present in Algerian propolis, has exerted antiallergic activity via a reduction in the release of inflammatory mediators such as histamine and leukotrienes [42].

3.4

Antioxidant Activity

Oxidative damage to biomolecules, such as DNA, RNA, proteins, enzymes, and lipids, is attributed to an imbalance in the formation and elimination of reactive oxygen (ROS) or nitrogen (RNS) species. This imbalance, and subsequent damage to biomolecules, plays a role in many conditions including cancer [156],

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neurodegeneration [157], diabetes, and cardiovascular diseases [158, 159]. African propolis possesses varying degrees of antioxidant capabilities depending on the presence of varying amounts of antioxidant compounds. It has been observed that propolis extracts rich in phenolic substances (e.g., flavonoids and phenolic acids) exhibit the strongest antioxidant activity [15, 38, 160–162]. This could be explained through mechanisms such as the direct scavenging of ROS, the chelation of metal ions involved in free-radical formation, and the inhibition of the activity of enzymes producing ROS [163]. Chrysin (20), tectochrysin (21), pinostrobin (42), 3β-cycloartenol (67), 3βcycloartenol-26-oic acid (74), 3α-cycloartenol-26-oic acid (75), and β-amyrin acetate (84), all isolated from Egyptian propolis, have shown radical-scavenging activity as well as xanthine oxidase inhibitory activity [27]. Mangiferonic acid (71), methyl-3β,27-dihydroxycycloart-24-en-26-oate (77), 3βhydroxylanostan-9,24-dien-21-oic acid (93), and 10 -O-eicosanylglycerol (119), isolated from Cameroonian propolis, showed radical-scavenging activity that was higher than that of their corresponding crude extracts [33].

3.5

Organ-Protective Activity

The presence of antioxidant compounds, which help counteract cell damage, has often been attributed to the observed protective activity of propolis against a range of organs [164, 165]. When administered prior to the anticancer drug doxorubicin, Algerian propolis has been found to induce cardio-, nephro-, and hepatoprotective effects [166–168]. Egyptian propolis can attenuate the testicular toxicity of doxorubicin [169]. It has also been reported that it may protect against ovarian toxicity following exposure to the pesticide methoxychlor [170] and limit cytotoxicity (on reproductive organs and the liver) and genotoxicity (chromosomal aberrations in bone marrow cells) induced by the anticancer drug cisplatin [171]. Egyptian propolis also protects against aflatoxin B1-induced hepatotoxicity [172]. Moroccan propolis showed a protective effect against ethylene glycol-induced hepatotoxicity and nephrotoxicity [173]. Tunisian propolis can limit nephrotoxicity following exposure to heavy metals [174], and Nigerian propolis has hepatoprotective and pancreatoprotective properties [175]. Pinocembrin (38), present in Egyptian, Nigerian, and Algerian propolis, when administered as a prophylactic long-term treatment to animals with induced cerebral ischemia reperfusion, showed neuroprotective activity [76].

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3.6

437

Antiviral and Immunomodulatory Activity

The antiviral effect of African propolis has been poorly investigated. African propolis extracts have only been tested against the infectious bursal disease, the reovirus, and the Newcastle disease viruses. Samples showed a reduction in infectivity mean viral titers to varying degrees, with Egyptian propolis exhibiting the highest activity against all viruses [27, 39, 176–178]. Studies investigating the immunomodulatory properties of African propolis are also limited. Egyptian propolis has been shown to strengthen the defense system of the Nile tilapia fish (Oreochromis niloticus) [179] and protect rats through immunostimulation, from the symptomatic manifestations associated with S. aureus and Pasteurella multocida infections [180–182]. In a study investigating the effect of Egyptian propolis on cutaneous warts, it was demonstrated that treated individuals showed no recurrence of plantar and common warts. This was explained partly due to the antiviral and immunomodulatory effects of this type of propolis [183].

3.7

Cytotoxic and Antitumor Activity

Cancer is a major cause of global morbidity and premature mortality, and its burden is expected to grow over the coming years [184]. Tunisian propolis has demonstrated potent dose-dependent cytotoxicity against a range of cancer cell lines [138]. Algerian propolis can synergize the antitumor effect of doxorubicin on human pancreatic cancer cells by blocking efflux pump activity, inducing cell cycle arrest, and promoting apoptosis [185]. When Algerian propolis was administered to mice with melanoma, a reduction in melanoma tumor growth and an increased survival rate was observed. A prophylactic treatment with Algerian propolis also reduced tumor growth, but with no effect on life prolongation [55, 56]. Egyptian propolis, given alone or in combination with doxorubicin, has shown antiproliferative and apoptotic effects against PC3 cancer cells that were greater than doxorubicin alone [186]. Mice treated with Egyptian propolis prior to being injected with Ehrlich ascites carcinoma (EAC) cells have been found to live longer than a control group that received no propolis. These findings were attributed to multiple mechanisms of action, including the inhibition of tumor proliferation, induction of apoptosis, and immunostimulation. In particular, it has been reported that the administration of propolis, prior to inoculation of EAC cells, arrested the cells in the S-phase and prevented further proliferation. Egyptian propolis has also been found to induce the sub-G1 apoptosis process in cancerous cells, resulting in tumor reduction [187, 188]. Phenethyl-(E)-caffeate (CAPE) (7), present in Algerian propolis, significantly increased the antiproliferative and cytotoxic effects of docetaxel and paclitaxel in

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various prostate cancer cell lines through modulation of the estrogen receptor ER-β [43].

3.8

Other Miscellaneous Biological Effects

The anti-aging, wound healing, and tissue regenerative properties of African propolis have been investigated for extracts of Algerian propolis revealing potent inhibition of stromelysin-1 (MMP-3), an enzyme involved in the proteolytic degradation of collagen and elastin fibers. Caffeic acid (1), chicoric acid (14), and chicoric acid methyl ester (15), present in Algerian propolis, were identified as the phytochemicals responsible for this effect. Algerian propolis can also inhibit human plasmin enzymes involved in the pathway leading from pro-MMP-3 to the active MMP-3 enzyme [23]. Moroccan propolis when administered to rats with Capparis spinosa honey can trigger a diuretic effect [162] and has the potential to treat and prevent kidney stones, crystaluria, and proteinuria [173]. It has also been reported that Moroccan propolis can inhibit glucosidase and amylase enzymes [189]. Cameroonian propolis has been found to exert estrogenic effects and can help alleviate hot flushes in rats [190]. Nigerian propolis has displayed antihyperglycemic and hypocholesterolemic effects by decreasing blood glucose, glycated hemoglobin (HbA1c), and very low-density lipoprotein (VLDL) levels and elevating highdensity lipoprotein (HDL) levels in diabetic rats [191].

4 Conclusion and Perspectives The use of natural products in the development of new pharmaceuticals has proven to be a well-founded and viable drug discovery strategy so far [1]. The African continent is characterized by a wide range of geographical regions and a rich diversity of ecosystems [192] where a range of different plant species can be used by bees to produce propolis and subsequently exploited by scientists to afford new potential drug templates. To date, we found that propolis from only 9 African countries (of a total of 54) has been investigated for its biological activity and/or phytochemical constituents. The samples investigated have yielded a high diversity of compounds and exhibited a range of biological properties, including antimicrobial, antiparasitic, antiinflammatory, antioxidant, organ-protective, antiviral, immunomodulatory, cancerrelated inhibitory, and other miscellaneous effects. In many cases however, little is known about whether the aforementioned effects depend upon the presence of some specific phytochemical(s) or a potentiation between different compounds that may act synergistically.

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Much of what is known about the biological activity of African propolis relies on studies testing crude or semi-fractionated extracts that have been poorly chemically standardized. The lack of quantification (i.e., determination of the nature and relative abundance of phytochemicals in a sample) also prevents the results from different studies to be compared in a meaningful way. Another issue that has limited the progression of scientific knowledge on African propolis is the use of different assay methodologies to screen samples for bioactivity. This, however, should not discourage further research on this topic because the limited studies to date have already revealed that African propolis has a tremendous potential for providing diverse biologically active chemicals that can serve as templates for the development of new drugs leads. These include 7-Odialkylaminoalkyl pectolinarigenins, like 135, a synthetic compound based on the structure of pectolinarigenin (26) (Fig. 10) that has strong antiproliferative activity against human lung carcinoma COR-l23 and A549 cancer cell lines [60], and some derivatives of nymphaeol B (46) like 136 which have, so far, showed greater potency against PC-3 prostate cancer cells than 5-fluorouracil [193] (Fig. 10). Research on African propolis must be expanded to encompass propolis samples from many different countries and geographical regions in this vast continent. This will provide a more complete picture of the diversity of propolis phytochemicals available for potential drug design and development. Further research should also aim to standardize screening methods to ensure consistency in methodologies. Altogether, this will allow for the findings of different studies to be more easily compared. Finally, research must move beyond the use of extracts in biological assays and focus on isolating phytochemicals and screening them to identify any bioactive compounds. Determining which phytochemicals are biologically active as well as unraveling their molecular targets will help to determine the mechanisms through which propolis achieves its biological effects. We believe that this contribution provides a starting point upon which further research investigating, yet unexplored, African propolis samples for the presence of new biologically active chemical entities may be based. O O

O N

O

O

O

O O

O

HO

O OH

O OH

135 (7-O-dimethylaminopropyl-pectolinarigenin)

136 (6-(3-Hydroxy-3-methylbutyl)-2'-(7-hydroxy-3,7dimethyloct-3-enyl)-3',4',5,7-tetramethoxyflavanone)

Fig. 10 Compound structures of bioactive synthetic compounds 135 and 136 based on natural product templates isolated from African propolis

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N. Blicharska and V. Seidel Natalia Blicharska graduated with a BSc (Honors) degree in Pharmacology and Environment and Health from the University of Toronto (Canada) in 2016 and obtained her MSc (Distinction) in Medicinal Natural Products and Phytochemistry from the UCL School of Pharmacy (London, UK) in 2017. During her academic career, Natalia tailored her studies to reflect her fascination for natural source drug discovery as well as to develop the didactic ground work and research skills required to excel in this field. She pursued several international research projects in order to explore this field of research, including a summer research project at the Chinese University of Hong Kong and a semester-long research project at the University of Strathclyde, the findings of which have been published in the University of Toronto’s Journal for Undergraduate Life Sciences (2016) and Scientific Reports (2018), respectively. She has been a member of the Phytochemical Society of Europe since 2017. As an aspiring young scientist, Natalia is driven to further develop her career in Pharmacognosy and Phytochemistry. Her particular research interests focus on isolating and identifying compounds of interest from natural sources that may have the potential to be developed into new drugs as well as exploring the value of medicinal natural products in contemporary medicine.

Veronique Seidel graduated with a degree in Pharmacy from the University of Lille (France) (1995) and obtained a Ph.D. in Phytochemistry from the University of Strathclyde (Glasgow, UK) (1999). Following a period as a Teaching Assistant in Pharmacognosy at the University of Lille, she worked as a Postdoctoral Researcher in Natural Products at De Montfort University (Leicester, UK) (1999–2001) and as a Postdoctoral Researcher in Microbiology at the University of London (2001–2003). She was appointed as a Lecturer (2003) and Senior Lecturer (2016) in Natural Products Drug Discovery at the University of Strathclyde. She is a Scientific Adviser for the International Foundation for Science (Food Science and Nutrition and Natural Products Programs) and serves on the editorial boards of Evidence-Based Complementary and Alternative Medicine, the Journal of Pharmaceutical Microbiology, Scientific Reports, and Phytochemistry Letters. She has held the position of Honorary Treasurer of the Phytochemical Society of Europe since 2014. Her field of research is the investigation of Nature as a source of biologically active substances, particularly the search for novel antimicrobials from medicinal plants and products from the beehive.